2w 3*? . fixiié“; . 35— - .ii'é-Eéfinq fi,.. ’ \ i 51‘5- fifir’éxz WF-‘Qf. £‘ 'rn u... m V. r 3% ., . .. 3 ‘ ' ’ « Us « ., . “w ‘ . . . ‘ “h ‘ y , + V fififigfigfi; . ' ' . ‘ - 1 7.. ‘ - 3ofiqgu'g; M “045:; W 7, ~ ' . ' ' ' .4 ' I I ' ‘ , ’_.'_( {I r v 3‘13“ 1'? k‘ by; ' ‘ lg: K3439 . .¥F¢“;‘ ‘ ' . ' ~ '- ‘v‘ z" . - t ‘--" - - . . -- "my“ 793.391 V“? V. ‘1- -m v “.331: 'rr 8. warm, ENM I x n a -I A. :‘m Law‘— ’ ,- J :5 A ~ "I. ,1 .P‘ Jr... "r... ‘3» Y- :-£..:._-.-.,y~ ..;_.. J- $54-; 5 * ‘ J u a prxrac-gmi “é: LIBRARY Michigan State ‘ University This is to certify that the dissertation entitled Transcripitonal regulation of the Aspergillus parasiticus aflatoxin biosynthetic pathway gene nor-1 presented by Michael Joseph Miller has been accepted towards fulfillment of the requirements for the Food Science and Ph.D. degree in Environmental Toxicology 821271;) /%fl/fi Major Professor’ 5 Sighature 2.“ ZS -- 03 Date MSU is an Affirmative Action/Equal Opportunity Institution TRANSCRIPTIONAL REGULATION OF THE ASPERGILL US PARASI TIC US AF LATOXIN BIOSYNTHETIC PATHWAY GENE NOR-1 By Michael Joseph Miller 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 2003 Q X \ II ABSTRACT TRANSCRIPTIONAL REGULATION OF THE ASPERGILL US PARASI TIC US AF LATOXIN BIOSYNTHETIC PATHWAY GENE NOR-1 By Michael Joseph Miller Aflatoxin is a potent hepatocarcinogen produced predominantly by Aspergillus flavus and A. parasiticus that can contaminate several commodities including com, cotton, peanuts and certain tree nuts. Aflatoxin biosynthesis is a complex process that requires several genes that all reside in an aflatoxin gene cluster. Since it would be unpractical to investigate the regulation of all aflatoxin genes, we have chosen nor-1 to serve as a model aflatoxin gene. Nor-l catalyzes the conversion of the first stable intermediate, norsolorinic acid to averantin. An early pathway gene, such as nor-I, is an ideal target for trancriptional regulation studies. In addition, there were several tools available including an A. parasiticus nor-1 mutant strain, Nor-1 recombinant protein, polyclonal anti-Nor-l antibody and a nor-I reporter strain. The specific objectives of these studies were to: 1) demonstrate that transcriptional activation of nor-I (and presumably the other aflatoxin structural genes) is at least partly responsible for the increase in aflatoxin production under inducing conditions; 2) develop a nor-I reporter system for use in identifying cis-acting sites in the nor-1 promoter; 3) determine the role of an aflatoxin pathway regulator, AflR, in the regulation of nor-I ; and 4) identify additional possible cis-acting sites in the nor-1 promoter. To measure nor-I trancriptional activation, plasmids that contained the nor-1 promoter fused to the B-glucuronidase gene were transformed into A. parasiticus. 6 Preliminary experiments utilized A. parasiticus D8D3, a strain that carries a 3.0 kb nor-1 promoter fragment fused to GUS that integrated at the nor-1 terminator. The transcriptional activation of nor-1 mirrored the accumulation of aflatoxin, nor-1 transcript and Nor-1 protein. In addition, under culture conditions that generated the most aflatoxin, the highest GUS activities were recorded. A new nor-1 ::GUS reporter plasmid was constructed that enabled easy nor-1 promoter insertions. However, the site of integration of the nor-1 ::GUS plasmid was demonstrated to be an important consideration. Integration outside of the aflatoxin gene cluster resulted in significantly reduced nor-1 transcription. Of the three putative AflR binding sites located in the nor-1 promoter area (AflRl, AflR2 and AflR3), only AflRl is clearly necessary for nor-1 transcriptional activation. AflR2 may also be involved in nor-l transcriptional activation but the presence of the ORF 3 gene (unknown function), which is located between nor-1 and AflR2, prevents a clear conclusion. AflR3 is not necessary for nor-I transcriptional activation because deletion of AflR3 in a nor-1 ::GUS reporter strain resulted in no change in nor-l transcriptional activation. Substitution of a putative TATA box in the nor-1 promoter in the context of a larger promoter demonstrated the requirement for the TATA box in nor-l transcriptional activation. Evidence was also presented for two additional cis-acting sites in the nor-I promoter, norL and CRE. While the studies described in this dissertation have accomplished the objectives, they have also generated several new questions. Future scientists interested in aflatoxin gene regulation have several possible research avenues including: 1) identify CRE and norL binding proteins, 2) investigate the mechanisms of aflatoxin gene-cluster-dependent regulation, and 3) investigate the function of ORF3. To my parents, for all their unconditional love and support iv 3 I.“ U) i J & Ln. ._ ACKNOWLEDGMENTS I would first like to thank my major advisor, Dr. John Linz. This dissertation would not have been possible without him. Dr. Linz is not only a great scientist but also a great man. I have been very fortunate. I am also grateful to all my guidance committee members, Dr. James Pestka, Dr. Steve Triezenberg, Dr. Tom Whittam, Dr. Joseph Schroeder, Dr. Bill Helferich and Dr. Zachary Burton. They have all contributed to increasing the value of this dissertation. Several people in the Linz laboratory have also provided invaluable help including Dr. Liang, Dr. Wu, Dr. Zhou, Dr. Trail, Dr. Roze, Dr. Mahanti, Dave Wilson, Ching-Hsun Chiou and Le-Wei Lee. Specifically, I would like to thank Matt Rarick for his help in almost every aspect of this dissertation. These colleagues are also great friends that I will miss greatly. Lastly, I would like to thank all my friends and family for all their unconditional support during my graduate career. Cl TABLE OF CONTENTS LIST OF TABLES ....................................................... x LIST OF FIGURES ..................................................... xi LIST OF ABBREVIATIONS ........................................... xvii CHAPTER 1 LITERATURE REVIEW ........................................... 1 Background and Significance ........................................ 1 Aspergillus and Aflatoxin ..................................... l Toxicology and Epidemiology .................................. 3 Economic Costs of Aflatoxin Contamination ...................... 5 Aflatoxin Biosynthesis - General Review ......................... 6 nor-1 - An Aflatoxin Biosynthetic Structural Gene ............. ' ..... 7 Transcriptional Regulation of Mycotoxin Biosynthesis .................... 9 Regulation of Aflatoxin Gene Transcription ...................... 10 AflR - Discovery ..................................... 10 AflR - Function ...................................... 10 AflR - Regulation ..................................... 12 Evidence for Other Transcription Factors .................. 12 ORF3 and nor-I/pksA transcriptional activation ....... 16 AflJ .......................................... 17 Regulation of Trichothecene Gene Transcription .................. 18 Tri6 - Discovery ...................................... 18 Tri6 - Function ...................................... 19 TrilO - Discovery ..................................... 19 Regulation of Biosynthesis Gene Transcription for other Mycotoxins . . 21 Mycotoxin Gene Clusters .......................................... 22 Organization of Gene Clusters ................................. 22 Aflatoxin Gene Cluster ................................ 22 Trichothecene Gene Cluster ............................. 23 Fumonisin Gene Cluster ............................... 27 Impacts of Mycotoxin Gene Clusters ........................... 27 Identification of Gene-Cluster-Dependent Regulation ........ 27 Significance of Gene-Cluster-Dependent Regulation ......... 31 Evolution of Mycotoxin Gene Clusters .......................... 33 Origin of the Aflatoxin Gene Cluster ..................... 33 Origin and Diversity of Trichothecene Gene Clusters ......... 34 ACKNOWLEDGMENTS .......................................... 35 CHAPTER 2 Effects of nitrogen source, carbon source and zinc concentration on the regulation of the aflatoxin biosynthetic gene nor-1 from Aspergillus parasiticus ........ 36 vi 17-51 5/ .y't 5‘6 INTRODUCTION ................................................ 36 MATERIALS AND METHODS ..................................... 39 Strains and Growth Conditions ................................ 39 RNA and Protein Extraction .................................. 40 Northern Hybridization Analysis ............................... 41 Western Blot Analysis ....................................... 42 Liquid Culture GUS Assays .................................. 42 Protein Extraction for EMSA ................................. 42 nor-1 and ver-I Promoter Fragments ............................ 44 Electrophoretic Mobility Shift Assay (EMSA) .................... 44 ELISA ................................................... 45 RESULTS ...................................................... 45 Generation of anti-Nor-l antibody ............................. 45 Carbon, nitrogen and zinc affect aflatoxin production and aflatoxin gene expression .......................................... 47 Nutrient effects are in part regulated at level of transcription ......... 51 Nutrients affect DNA promoter binding by cellular proteins ......... 53 DISCUSSION ............................................... ". . . . 58 ACKNOWLEDGMENTS .......................................... 60 CHAPTER 3 Chromosomal Location Plays a Role in Regulation of Aflatoxin Gene Expression in Aspergillus parasiticus .......................................... 61 INTRODUCTION ................................................ 61 MATERIALS AND METHODS ..................................... 63 Strains and growth conditions ................................. 63 Plasmid Constructs ......................................... 64 pNele ............................................ 64 pNiaD-Al ........................................... 64 pNANG-3 ........................................... 67 pBNG3.0-3F ......................................... 67 pAPGUSNN-B ....................................... 68 Transformation and Identification of the site of plasmid integration . . . 68 Transformation ....................................... 68 Rapid DNA extraction procedure ........................ 68 3' PCR analysis ...................................... 69 5' PCR analysis ...................................... 69 Southern hybridization analysis .......................... 70 Reporter Assays ............................................ 70 Solid culture GUS Assay ............................... 70 RESULTS ...................................................... 71 Generation of transformants .................................. 71 Phenotype Screening of Transformants ......................... 71 Genotype Screening of Transformants .......................... 73 DISCUSSION ................................................... 81 ACKNOWLEDGMENTS .......................................... 84 vii CHAPTER 4 Role of AflR in nor-1 transcriptional activation in Aspergillusflavus and A. parasiticus ...................................................... 85 INTRODUCTION ................................................ 85 MATERIALS AND METHODS ..................................... 89 Strains ................................................... 89 Plasmid Construction ........................................ 89 A. parasiticus reporter plasmids ......................... 89 A. flavus reporter plasmids .............................. 89 A. flavus tAflR expression plasmid construction ............. 92 Generation and Selection of Transformants ...................... 93 Transformation ....................................... 93 Site of Integration PCR Assay (A. parasiticus) .............. 93 Southern Hybridization (A. parasiticus) ................... 94 Dot Blot Hybridization Analysis (A. flavus) ................ 94 GUS Reporter Assays ....................................... 95 Solid culture GUS Assay (A. parasiticus) .................. 95 Liquid Culture GUS Assay (A. parasiticus) ............ . . . . 95 Quantitative GUS Assay (A. flavus) ...................... 96 A. flavus tAflR (truncated AflR) binding studies .................. 97 Induction and Purification of tAflR ....................... 97 Preparation of oligo probes ............................. 98 Southwestern Blot .................................... 99 RESULTS ..................................................... 100 AflR binding to the A. flavus nor-1 promoter .................... 100 A. flavus GUS Activity Assays ............................... 100 Generation and Screening of A . parasiticus Transformants ......... 103 A. parasiticus GUS reporter assays ............................ 105 DISCUSSION .................................................. 1 12 ACKNOWLEDGMENTS ......................................... 1 17 CHAPTER 5 Identification of novel cis-acting sites in the aflatoxin biosynthetic nor-1 promoter of Aspergillus parasiticus ......................................... 1 18 TNTRODUCTION ............................................... l 18 MATERIALS AND METHODS .................................... 121 Strains and growth conditions ................................ 121 Plasmid Constructs ........................................ 121 pNANG-3 .......................................... 121 pBNG332 and pBNG332TATAmut ..................... 122 pBNG332, pBNG298, pBNG268, pBNG238 and pBNG210 . . 122 pBNGnoerut ...................................... 123 Generation and Screening of Transformants ..................... 123 Transformation ...................................... l 23 Site of Integration PCR Assay .......................... 123 Southern Hybridization ............................... 124 GUS Reporter Assays ...................................... 124 viii Solid culture GUS Assay .............................. 124 Liquid Culture GUS Assay ............................ 125 in vitro DNA Binding Assays ................................ 126 Protein Extraction ................................... 126 Probe Generation .................................... 127 Electrophoretic Mobility Shift Assay (EMSA) ............. 127 RESULTS ..................................................... 127 Generation and screening of A. parasiticus transformants .......... 127 in vitro analysis nor-l promoter elements ....................... 133 DISCUSSION .................................................. 136 ACKNOWLEDGMENTS ......................................... 143 CHAPTER 6 Future Studies .................................................. 144 LIST OF REFERENCES ................................................ 146 ix flan? LIST OF TABLES Table 2.1. Medium components used in this study ............................. 40 Table 5.1. Oligonucleotides used for electrophoretic mobility shift assays ........ 128 C Figure Figure Figure Figure Figure Figure Figure Figure Figure LIST OF FIGURES 1.1. Molecular structure of the primary aflatoxins. ........................ 2 1.2. Enzymatic activation of AF 8,. The 8,9 double bond of AF BI is epoxidated by mixed function mono-oxygenases to the reactive epoxide. The epoxide can then form adducts with cellular macromolecules including DNA. ............ 4 1.3. Genomic organization of the aflatoxin biosynthetic gene cluster in Aspergillus parasiticus. Arrowheads indicate the direction of transcription. Drawn approximately to scale. ....................................... 8 1.4. Schematic of the nor-I/pksA intergenic region in A. parasiticus. The numbers indicate the number of nucleotides included upstream from the primary transcriptional start site of nor-1. Several potential cis-acting sites are indicated including the AflR binding sites AflRl, AflR2 and AflR3 and PacCl and BrlA3. The location of an open reading frame (ORF 3) of unknown function is also shown. (Figure 1.4 adapted from Miller et al., 2003a) .................... 13 1.5. Identification of cis-acting sites in the nor-1 promoter of A. parasiticus. The numbers indicate the number of nucleotides from the primary transcriptional start site of nor-1. (Figure 1.5 adapted from Miller et al., 2003b) ............... 15 1.6. Proposed regulatory model for trichothecene biosynthesis. Solid arrows indicate positive activators while open arrows indicate inhibitory activities. Question marks indicate other proposed but unknown regulatory signals or factors. (Figure 1.6 adapted from Tag etal., 2001) ....................... 20 1.7. Schematic of the sugar utilization gene cluster in A. parasiticus. The moxY gene is at one end of the aflatoxin gene cluster. The spacer regions on either side of the sugar utilization cluster do not contain open reading frames. Map is roughly to scale. (Figure 1.7 adapted from Yu et al., 2000a) ............... 24 1.8. Genomic organization of the trichothecene biosynthetic gene cluster of F usarium sporotrichioides and F. graminearum. Arrowheads indicate the direction of transcription and the number underneath each arrow refer to the specific gene. Genes with the same number from both F usarium species are homologues. tri 7 in F. graminearum is non-functional. Map is roughly to scale. (Figure 1.8 adapted from Brown et al., 2001) ........................... 26 1.9. Schematic of the fumonisin biosynthetic gene cluster in F usarium verticillioides. Arrowheads indicate the direction of transcription. Map is roughly to scale. (Figure 1.9 adapted from Seo et al., 2001) ................ 28 Figure 2.1. Western blot analysis of the native Nor-1 protein from A. parasiticus SU-l. xi Each lane contained 10 mg protein. The primary antibody was the IgG fraction of the antiserum raised against the Nor-lc/MBP fusion protein (10 mg/ml). Lanes: 1, total crude extract (10,000 X g supematent) from the nor-1 disrupted strain A. parasiticus ANor-l cultured in YES liquid medium for 60 h; 2 and 3, total crude extract from the wild type A. parasiticus SU-l cultured in YES liquid medium for 48 h and 60 h respectively; 4, the native Nor-l protein (31 kDa) purified with anti-Nor-lc/MBP fusion protein PAb affinity column from the total crude extract of A. parasiticus SU-l cultured in YES medium for 60 h. ................. 46 Figure 2.2. Aflatoxin production by the reporter strains A. parasiticus D8D3 (A) and 14 (B) grown in GMSO, GMS, NMS and PMS (D8D3 only) to 36, 48 and 72 hours (D8D3) or 48, 60 and 72 hours (14). Aflatoxin concentration of the growth media was determined by competitive-direct ELISA using polyclonal antibodies. Aflatoxin concentrations are reported as the average of the triplicate flasks, analyzed twice, with the standard deviation represented by errors bars. Non- detectable activity is represented by ND. .............................. 48 Figure 2.3. nor-1 transcript and Nor-1 protein accumulation and aflR transcript accumulation by the reporter strain A. parasiticus D8D3. (A) nor-1 transcript accumulation assessed by Northern hybridization analysis. For the time points (36, 48 and 72 h), each replicate flask is presented (A, B and C). Top panel is RNA from GMS cultures and bottom panel is RNA from PMS cultures. The lane designated “+” has RNA from a 48 h A. parasiticus SUl culture in YES medium. Equal loading of RNA is demonstrated by ethidium bromide staining of RNA as shown in the panels marked “EtBr”. (B) Nor-l protein accumulation by Western blot analysis. One flask per time point (36, 48 and 72 h) was analyzed from PMS and GMS cultures. (C) aflR transcript accumulation assessed by Northern hybridization analysis. For the time points (36, 48 and 72 h), each replicate flask is presented (A, B and C). Top panel is RNA from GMS cultures and bottom panel is RNA from PMS cultures. The lane designated “+” has RNA from a 48 h A. parasiticus SUl culture in YES medium. Equal loading of RNA can be determined from part A in the panels marked “EtBr”. .................... 49 Figure 2.4. fl-glucuronidase (GUS) expression by the reporter strains A. parasiticus I4. Cultures were grown in GMS, GMS0 (GMS - 0) and NMS for 48, 60 and 72 h. ............................................................... 52 Figure 2.5. Map of EMSA Oligonucleotides used for electrophoretic mobility shift assay (EMSA). Three putative cis-acting sites are boxed, AflR TATA and CRE. For nor-R*, the AflR site has been mutated (TCchcagCGA to AGTttaaaCAG). For nor-TATA*, the TATA box was mutated (5'—ATATATAG-3' to 5'-GTTTAAAC- 3'). ............................................................ 54 Figure 2.6. EMSA of nor-R. Protein extracts were collected from two different A. parasiticus strains, SUl (S) and AF S10 (A). Each strain was grown in two different media, PMS (P) and GMS (G). 32 mg of protein was used per lane. The two competitors (250 fold excess) used were nor-R (R) and nor—R* (R*). ..... 55 xii Figure 2.7. nor-R competition EMSA. Protein extracts were collected from A. parasiticus SUl (S) grown in GMS (G). 32 mg of protein was used per lane. The competitors used were: nor-R (R), nor-TATA* (RT*), nor-R1 (R1), nor-R2 (R2) and CRE. ....................................................... 57 Figure 3.]. Restriction endonuclease maps of relevant plasmids. (A) pNEBl93 (New England Biolabs, Beverly, MA). This plasmid is a modification of pUC 19 that incorporates additional restriction endonuclease sites in the multiple cloning sites. These additional restriction enzymes recognize 8 bp sequences and consequently are less likely to exist in the DNA being cloned. (B) pNele. This plasmid has a Not] site that replaces a BamHI site in pNEBl93 by insertion of a Not] linker. (C) pNiaD-Al. A 7.4 kb XhoI/Sal] fragment from pSL82 (Homg etal., 1990) that contains the niaD selectable marker was cloned into the Sal] site of pNebN 1. (D) pNANG-3. In addition to carrying the niaD selectable marker, pNANG-3 carries a small part of the nor-1 coding sequence (10 amino acids) fused to the B- glucuronidase (uidA or GUS) coding sequence which is in turn fused to the 2 kb nor-1 3' terminator fragment. The small numbers of codons that were changed in the nor-l coding sequence were all acceptable based on codon usage and ' maintained the correct reading frame. The 4 kb GUS/nor-l terminator fragment was amplified by PCR using pAPGUSNN-B (Figure 3.1F) as template with primers that had Not] (5') and AscI (3') tails. Appropriate promoter pieces, amplified by PCR using primers with Not] (and Pacl if directionally cloned) can be cloned into pNANG-3. (E) pBNG3.0-3F. This plasmid contains a 3 kb PCR~ amplified nor-I promoter piece cloned into the Not] site of pNANG-3. (F) pAPGUSNN-B. Original nor-1 ::GUS reporter plasmid constructed by Wilson (Chiou et al., 2002). .............................................. 65 Figure 3.2. Experimental design. pBNG3.0-3F fungal transformants were first tested for GUS activity using a solid culture GUS assay. Selected GUS+ and GUS- transformants were then tested using a 3' and 5' nor-1 site of integration PCR assay. Lastly, 3' nor-I integration status was confirmed for selected transformants using southern hybridization analysis ...................... 72 Figure 3.3. Solid culture GUS assay. Screening of transformants was performed using a solid culture GUS assay. Transformants 1, 2, 3, 4 and 6 are GUS- while transformant 5 is GUS+. ........................................... 74 Figure 3.4. PCR analysis of site of integration in GUS+ transformants. (A) Restriction endonuclease map of pBNG3.0-3F. Only restriction endonuclease sites relevant to this study are shown. Ten GUS+ (lanes 1 to 6 and 8 to 11) and one GUS- transfonnant (lane 7) were analyzed for both 5' and 3' nor-I integration. Template DNA was prepared using a rapid boiling procedure. (B) Schematic for 5' nor-1 integration with PCR data. 5' nor-l integrants resulted in a 3.1 kb PCR fragment (lanes 1, 2, 5, 8, 11). (C) Schematic for 3' nor-I integration with PCR data. 3' nor-l integrants resulted in a 2.1 kb PCR fragment (lanes 3, 4, 6, 9, 10). All GUS+ transformants were 5' or 3' nor-1 integrants while the GUS- transformant (lane 7) was negative for both 5' (B) and 3' (C) nor-1 integration. x'ii Lanes labeled M in panels B and C represent molecular size markers ......... 75 Figure 3.5. PCR analysis of site of integration in GUS- transformants. Template DNA was prepared using a rapid boiling procedure. (A) 3' site of integration PCR assay. All 22 randomly chosen (lane numbers indicate transformant number) GUS- transformants lacked the 2.1 kb PCR fragment diagnostic for 3' nor-1 integration. Positive controls (lanes +) did contain the 2.1 kb PCR fragment. (B) 5' site of integration PCR assay. All 22 randomly chosen (same as in A) GUS- transformants lacked the 3.1 kb PCR fragment diagnostic for 5' nor-I integration. A positive control (lane +) did contain the 3.1 kb PCR fragment. All GUS- transformants were negative for both 3' (A) and 5' (B) nor-1 integration. Lanes labeled M in panels B and C represent molecular size markers. ............. 76 Figure 3.6. Southern hybridization analysis of selected GUS+ and GUS- transformants for 3' nor-I integration. (A) Confirmation of 3' nor-I integration. All 8 transformants (designated by transformant number) were GUS+ and tested positive for 3' nor-l integration with the site of integration PCR assay. All 8 transformants tested confirmed 3' nor-I integration. (B) Testing for 3'n0r41 integration. All 5 GUS- transformants (designated by transformant number) tested negative for 3' (and 5') nor-1 integration using the site of integration PCR assay. Southern hybridization analysis confirmed lack of 3' nor-I integration in these 5 GUS- tranformants. All 5 GUS+ transformants (designated by transformant number) tested positive for 5' nor-I integration with the site of integration PCR assay. Southern hybridization analysis confirmed lack of 3' nor- 1 integration in these 5 GUS+ transformants. ........................... 78 Figure 3.7. Solid culture GUS assay. Transformants 1 and 6 are 3' nor-1 integrants and are GUS positive. Transformants 2 and 3 are 5' nor-1 integrants and are GUS positive. Transformants 4 and 5 are not 3' or 5' nor-1 integrants and are GUS negative. ........................................................ 80 Figure 4.1. Restriction endonuclease maps of relevant plasmids. (A) pNANG-3. In addition to the niaD selectable marker, pNANG-3 carries a small part of the nor-1 coding sequence (10 amino acids) fused to the B-glucuronidase (GUS) coding sequence which is in turn fused to the 2 kb nor-l 3' terminator fragment. Appropriate promoter pieces, amplified by PCR using primers with Not] and PacI tails, were cloned into pNANG-3. The small number of codons that were changed in the nor-l coding sequence to create the useful Not] site were all acceptable based on codon usage and maintained the correct sense. (B) pBNG3.0-3F. This plasmid contains a 3 kb PCR amplified nor-I promoter piece cloned into the Not! site of pNANG—3. Other nor-1 promoters that were tested (1250, 1200, 664, 332P, 332, 332AflRmut, 76 and 64) were also made by PCR and were directionally cloned into the Not] and Pool sites in pNANG-3. (C) pGAP12. This plasmid contains the GUS gene connected to a 1.3 kb BamHI nor- ] promoter fragment. pGAP 12 was used as a template for PCR to make the A. flavus reporter plasmids used in this study. (D) pGAP12-138. This plasmid contains 138 bp upstream of translational start site of nor-1. ............... 90 xiv 7 Of '3"? fit Figure 4.2. Southwestern blot analysis of AflRl with purified tAflR (A. flavus) and a PP2 probe. PP2 (CCAACTCGGCCAGCGACCAACACACCACC), a 29 bp oligo from the A. flavus nor-1 promoter, contains the AflR binding site AflRl (bold). PP2MUT (CCAACQCIGCCAGCGACCAACACACCACC) is the same as PP2 except for 2 mutations (underlined) in the AflR binding site. PP2 was labeled with 32F and used as probe in all three lanes. Competitors had a 15 fold molar excess and were unlabeled. Lane 1, no competitor; lane 2, PP2 competitor; lane 3, PP2MUT competitor ....................................... 101 Figure 4.3. Schematic of the A. flavus nor-1 promoters and GUS analysis. The AflR binding site AflRl is located between residues -103 and -91. Fold induction was determined by dividing the GUS activity (nmole MU / min / mg protein) in an inductive medium (SLS) by GUS activity in a non-inductive medium (PMS). .............................................................. 102 Figure 4.4. Schematic of the nor-1 promoter region in A. parasiticus. The numbers indicate the number of nucleotides included upstream from the transcriptional start site. Several potential cis-acting sites are indicated including the AflR binding sites AflRl , AflR2 and AflR3 and PacCl and BrlA3 which were reported to be involved in pksA transcriptional regulation (Ehrlich etal., 2002). The location of an open reading frame (ORF) of unknown function is also shown. The sizes of the nor-1 promoters used in this study are also indicated ( 1250, 1200, 664, 332, 76 and 64). ............................................. 104 Figure 4.5. Southern hybridization analysis of A. parasiticus transformans with 332AflRmut (332*), 332, 76, 64 and NR-l. Each letter indicates a different isolate from the same nor-1::GUS construct. Two of the 3' integrants shown from each nor-I ::GUS reporter construct were used for solid culture and liquid culture GUS assays. .................................................... 106 Figure 4.6. Liquid culture GUS assay analysis was performed on two different 3' integrants from each nor-1 ::GUS reporter construct grown in duplicate. The GUS activity is reported as the mean pmol/min/mg d: the standard deviation. ..... 107 Figure 4.7. Solid culture GUS assay analyses were performed on different 3' integrants from each nor-1 ::GUS reporter construct on 46 h YES agar colonies. The recipient strain NR-l was added as a negative control and had no detectable activity. (A) The 332 nor-1 ::GUS transformant (includes AflRl) had detectable GUS activity while the 332AflRMUT (AflRl mutated), 76 (includes AflRl) and 64 (AflRl deleted) nor-1::GUS transformants had no detectable GUS activity after 24 h incubation with GUS substrate. (B) The 3000 (includes AflRl, AflR2 and AflR3) and 1250 (includes AflRl and AflR2) nor-1::GUS transformants had similar activity. The 1200, 664 and 332P (all 3 include AflRl only) nor-[::GUS transformants all had similar activity which was significantly less than the 3000 and 1250 nor-I ::GUS transformants. The 332 nor-1::GUS transformant (includes AflRl) had significantly less GUS activity than all other nor-1::GUS transformants shown. ............................................. 108 XV Tag 017503 Figure 4.8. Genetic map of 3' integrants for both 332 and 332P nor-1 ::GUS transformants. 3' integration with the plasmids used in the A. parasiticus study result in the niaD selectable marker being immediately upstream of the nor- ] ::GUS promoter. Included in the 7.4 kb niaD fragment is 680 bp of m'iA coding sequence. A 503 bp PCR product that included the pyrG translational stop codon and transcriptional terminator was inserted into pBNG332 between the niaD selectable marker and nor-1 promoter at the PacI site. .................. 1 1 1 Figure 5.1. Importance of the TATA box in the nor-I promoter. (A) A liquid culture GUS assay analysis was performed on two different 3' integrants from each nor- 1::GUS reporter construct (332 and 332TATAmut) grown in duplicate. GUS activity is reported as the average of 4 values in pmol/min/mg i the standard deviation. (B) Solid culture GUS assays were performed on different 3' integrants from each nor-I ::GUS reporter construct on 46 h YES agar colonies. Colonies analyzed are: A 332-l, B 332-2, C 332TATAmut-1, D 332TATAmut-2, and E NR1. .......................................................... 129 Figure 5.2. Identification of norL cis-acting site. (A) A liquid culture GUS assay was performed on two different 3' integrants from each nor-1::GUS reporter construct grown in duplicate. The GUS activity is reported as the average of 4 values in pmol/min/mg :t the standard deviation. (B) Solid culture GUS assays were performed on different 3' integrants from each nor-I ::GUS reporter construct on 46 h YES agar colonies. Colonies analyzed are: A 332-1, B 298-1, C 268-1, D 238-1, E 210-1, G 332noerut-1 and H NR1. ......................... 130 Figure 5.3. EMSA with a 206/244 oligo. 20 frnol of the 206/244 oligonucleotide was used as a probe with 32 pg of protein extract from SUl or AF S10. Norpr complex and complex A are indicated with arrows. ..................... 135 Figure 5.4. EMSA with a CREl oligo. 20 frnol of CRE] oligonucleotide was used as probe with 32 pg of protein extract from SUl or AFSlO. CREbp complex is indicated with an arrow ............................................ 137 Figure 5.5. Location of putative TATA boxes in A. parasiticus aflatoxin biosynthesis gene promoters. The experimentally determined major transcriptional start point is shown at +1. (A) nor-1 (Trail et al., 1994) (B) avnA (Cary et al., 2000) (C) kaA (Ehrlich et al., 2002) (D) ver—I (Skory et al., 1992) (E) aflR (Ehrlich et al., 1999a). ........................................................ 138 Figure 5.6. Proposed model for nor-1 transcriptional activation. Arrows indicate positive interactions, blocked lines indicate negative interactions and lines with no block or arrow indicate unknown interaction. Definite interactions are represented by solid lines while inconclusive interactions are represented by dashed lines. .................................................... 142 xvi ‘i_m1.7lfl1 GMS PMS YES nor-1 ver-l pksA ORF3 (if/R AflRl AflR2 AflR3 C RE 1 CREbp norL Norpr EMSA nor-R LIST OF ABBREVIATIONS glucose + minimal salts medium; aflatoxin inducive peptone + minimal salts medium; aflatoxin non-inducive yeast extract + sucrose medium; aflatoxin inducive aflatoxin biosynthetic gene aflatoxin biosynthetic gene aflatoxin biosynthetic gene; polyketide synthase open reading frame 3; located between kaA and nor-1 in Aspergillus parasiticus aflatoxin biosynthetic gene; pathway transcriptional activator AflR binding site in nor-I promoter; -65 from nor-l transcriptional start site AflR binding site in nor-1 promoter; -1205 from nor-I transcriptional start site AflR binding site in nor-1 promoter; -1553 from nor—l transcriptional start site potential cis-acting site in the nor-I promoter protein that binds CRE] potential cis-acting site in the nor-1 promoter protein that binds norL electrophoretic mobility shift assay PCR amplified fragment from nor-1 promter; contains AflRl and CRE] xvii Wail I CHAPTER 1 LITERATURE REVIEW Background and Significance Aspergillus and Aflatoxin Aflatoxins are highly toxic and carcinogenic secondary metabolites of certain strains of Aspergillus parasiticus, A. flavus, A. nomius and A. psuedotamarii with only A. parasiticus and A. flavus being economically important (Council for Agricultural Science and Technology, 2003). Growth and production of aflatoxins require environmental conditions that are usually found in tropical and subtropical regions but may also be found in more temperate areas such as the United States (Dvorackova, 1990). Several different crops have been found to be contaminated with aflatoxins including com, cotton, peanuts and certain tree nuts (Council for Agricultural Science and Technology, 2003). Infection of the plant by toxigenic Aspergilli can occur before harvest or during storage after harvest (Wilson and Payne, 1994). Seventeen different compounds have been isolated and designated as aflatoxins yet the term usually refers to four metabolites ofthis group (Figure 1.1): aflatoxin B, (AFB,), B2 (AFBz), G, (AFG,) and G2 (AFGZ) (McLean and Dutton, 1995). Toxigenic A. parasiticus typically produces all four compounds while toxigenic A. flavus only produces AFB, and AF B2 (Dvorackova, 1990). AFB, is usually the most concentrated in food samples followed by AF G,, AFB2 and AFG2 (McLean and Dutton, 1995). Unfortunately, AFB, is also the most toxic, carcinogenic, and mutagenic of the four major aflatoxins (Roebuck and Maxuitenko, 1994). ' , -. llll ‘1”! E OCH3 Figure 1.1. Molecular structure of the primary aflatoxins. ",I‘ 'L" I fir. a- .. 34E; \ V" a 7&6? d VF" Toxicology and Epidemiology In 1960, an outbreak of acute hepatotoxic disease killed more than 100,000 turkeys (Blount, 1961). The etiologic agents were identified as A. flavus metabolites that were subsequently characterized and named aflatoxins (A. flvus Ms) (Asao et al. 1963). Since their discovery in the early 1960's, aflatoxin toxicity has been a subject of intense investigation. Aflatoxins, collectively, have been described as being a “quadruple threat” toxin — a potent toxin, mutagen, carcinogen and teratogen (McLean and Dutton, 1995). The order of acute and chronic toxicity is AFB, > AFG, > AFB2 > AFG2 (McLean and Dutton, 1995). The order of toxicity reflects the importance of the 8,9 double bond and also the greater potency associated with the cyclopentenone ring of the B series when compared to the lactone ring of the G series (McLean and Dutton, 1995). AFB, requires metabolic activation in order to reach its toxic and carcinogenic potential (McLean and Dutton, 1995). The 8,9 double bond in AFB, is epoxidated by mixed- function mono-oxygenases to the reactive form which can form adducts with cellular macromolecules including DNA, RNA, and proteins (Figure 1.2) (McLean and Dutton, 1995). Epidemiological data have linked aflatoxins with human hepatic cancer, primarily in under-developed countries where the elimination of contaminated products is not practiced (Reviewed in Dvorackova, 1990, Hall and Wild, 1994 and Council for Agricultural Science and Technology, 2003). Initially, many researchers noticed a striking correlation between regions with high aflatoxin contamination of food products and hepatic cancer. The prevalence of viral hepatitis and other mycotoxins in these high liver cancer areas complicated the epidemiological data and prevented epidemiologists from concluding unequivocally that AFB, by itself is a human carcinogen (Hall and 3 I”...‘ '4 M L... .035 g. (1700?. 'L/L \l O\\)\/\| 0 0 OCH3 0 0 OCH3 Figure 1.2. Enzymatic activation of AF 8,. The 8,9 double bond ofAFB, is epoxidated by mixed function mono-oxygenases to the reactive epoxide. The epoxide can then form adducts with cellular macromolecules including DNA. Wild, 1994). With the development of biomarkers for aflatoxin exposure, such as aflatoxin adducts in urine, the etiology of liver carcinomas could be further explored. The mutational spectrum of aflatoxin is dominated by one genetic change: the GC to TA transversion (Smela et al., 2001). In fact, aflatoxin exposure has been linked to a specific transversion (GC to TA) in the tumor suppressor gene, p53 (third position of codon 249) (Council for Agricultural Science and Technology, 2003). Use of specific biomarkers for aflatoxin exposure has demonstrated that AFB, and hepatitis B interact as risk factors for human liver cancer (Scholl et al., 1995). In addition, aflatoxin carcinogenicity studies clearly demonstrate the high potency of AFB, in several species of lab animals (reviewed in World Health Organization, 1979). Consequently, the elimination of human exposure to aflatoxin is a worthwhile goal. Economic Costs of Aflatoxin Contamination Due to valid health concerns, many countries have set regulatory action levels for aflatoxin in food and feed. In the United States, the action level is 20 ppb in food crops with higher action levels in feed crops (Council for Agricultural Science and Technology, 2003). Determining the precise economic cost of these action levels is not possible due to various uncertainties which include the extent and level of contamination, variability of contamination, variability of the price and quantity of the affected commodity, costs of efforts to mitigate contamination and loss in livestock value from contaminated feed (Council for Agricultural Science and Technology, 2003). Vardon et al. used a Monte Carlo computer simulation to estimate the yearly cost of aflatoxin contamination in the United States (Council for Agricultural Science and Technology, 2003). They estimated that the mean simulated potential value of crops lost because of aflatoxin contamination 5 was $47 million per year in food crops (peanut and com), S 225 million per year in feed corn, and $4 million per year in livestock cost (Council for Agricultural Science and Technology, 2003). Using a different method, Robens calculated that aflatoxin management costs the United States at least $100 million a year (Robens, 2001). While these values are estimates, it is clear that aflatoxin contamination has a significant economic impact in the United States. There are substantial economic costs to under—developed countries as well. At a conference in May, 2001, the United Nations Secretary-General Kofi Annan said, a World Bank study has calculated that the European Union regulation on aflatoxins costs Africa $750 million each year in exports of cereals, dried fruit and nuts. And what does it achieve? It may possibly save the life of one citizen of the European Union every two years Surely a more reasonable balance can be found” (Annan, 2001). The low aflatoxin action levels in many countries including the United States and the European Union have reduced aflatoxin exposure yet there are large economic costs associated. Economically feasible methods to reduce or eliminate aflatoxin exposure are clearly needed. Aflatoxin Biosynthesis - General Review A potential means to accomplish the goal of aflatoxin elimination is to elucidate the molecular mechanisms that regulate aflatoxin biosynthesis. This information is likely to generate novel approaches and targets for inhibition of aflatoxin gene expression The biosynthetic pathway for AFB, has been elucidated through the use of feeding studies, pathway mutants and inhibitors (reviewed in Bhatnagar et al., 1994). At least 16 different enzymatic steps are required for AFB, synthesis (Bhatnagar et al., 1991). A pair 6 r. 1 d of aflatoxin associated fatty acid synthases, F as-l and F as-2, together form the hexanoate starter molecule which is then acted on by a polyketide synthase (pksA) to form the first stable intermediate, the decaketide norsolorinic acid (Watanabe et al., 1996). Several enzymes responsible for the various biochemical steps have now been identified including Nor-l , Ver-l, and OmtA (Yu et al., 1995b). Different methods have been used to clone the biosynthetic genes including reverse genetics (Yu et al., 1993), subtractive hybridization (F eng et al., 1992) and complementation (Chang et al. 1992 and Skory et al. 1992). Complementation takes advantage of the many pathway mutants that are available and was used to clone the nor- ] (Chang et al. 1992), ver—I (Skory et al., 1992), aflR (Payne et al., 1993) and fas-IA (Mahanti et al., 1996) genes. Skory et al. identified one cosmid that contained both the nor-1 and ver-I genes which suggested that the genes in the pathway may be clustered (Skory et al., 1992). Mapping of this cluster identified several different transcripts that have the same timing of expression as nor-1 and ver-l and the genes that encode many of these transcripts have been subsequently identified and cloned (Trail et al., 1995b). The aflatoxin gene cluster from A. parasiticus is presented in Figure 1.3. nor-I - An Aflatoxin Biosynthetic Structural Gene Using the A. parasiticus norsolorinic acid accumulating strain B62 (niaD, nor-l), nor-I was cloned by complemention (Chang et al., 1992). Subsequently, gene disruption of nor-1 in wild type A. parasiticus resulted in the accumulation of norsolorinic acid supporting the proposed function of Nor-l protein (Trail et al., 1994). The transcriptional start site and the polyadenylation site for nor-1 were determined (Trail et al., 1994). Nucleotide sequence analysis of nor-1 revealed that it encodes an alcohol dehydrogenase 7 Figure 1.3. Genomic organization of the aflatoxin biosynthetic gene cluster in Aspergillus parasiticus. Arrowheads indicate the direction of transcription. Drawn approximately to scale. ¢fi«7- “gt-w domain consistent with its proposed function of reducing norsolorinic acid to averantin (Trail et al., 1995a). Functional analysis with recombinant Nor-l demonstrated that it is capable of converting norsolorinic acid to averantin in vitro confirming the proposed function (Zhou and Linz, 1999). At least 20 different genes are directly involved in aflatoxin biosynthesis. An understanding of how the early pathway genes such as nor-1 are regulated could be more effective in helping develop methods to reduce aflatoxin contamination. Since it would be impractical to study the transcriptional regulation of all aflatoxin genes, we chose to study nor-1 specifically. The timing of expression of nor-1 transcript and protein accumulation was consistent with the occurrence of aflatoxin (Skory et al., 1993). A reporter gene has been constructed with the nor-1 promoter fused to the B-glucuronidase gene (uidA) from Escherichia coli (Chiou et al., 2002). Prior to the work performed in this dissertation, no specific transcription factors or their binding sites have been identified in the nor-1 promoter. Transcriptional Regulation of Mycotoxin Biosynthesis Mycotoxins are secondary metabolites produced by several species of fungi. As secondary metabolites, mycotoxins are not constitutively synthesized. Rather, mycotoxin biosynthesis is a complex process that responds to developmental, environmental and nutritional cues. The primary means that fungi use to regulate mycotoxin biosynthesis appear to be transcriptional regulation of the mycotoxin biosynthetic genes. Aflatoxin and trichothecene biosynthesis are the most characterized of the mycotoxins. Information gathered about aflatoxin and trichothecene biosynthesis has been helpful for researchers studying other mycotoxin biosynthetic pathways. In the following sections, 9 A!" ’%~‘ I , 01 £10] I .700 the regulation of aflatoxin and trichothecene biosynthesis gene transcription is reviewed with brief mention of other mycotoxin pathways. Regulation of Aflatoxin Gene Transcription AflR - Discovery A. flavus strain 650 contains a mutation in the 0/72 locus (Bennett and Papa, 1988). The parental strain for A. flavus strain 650 was a norsolorinic acid accumulator while the A. flavus 650 strain did not accumulate norsolorinic acid (Bennett and Papa, 1988). Metabolite feeding studies and enzyme activity measurements of A. flavus strain 650 demonstrated that the aflatoxin biosynthetic enzymes were not present in the strain and that perhaps a mutation in a regulatory gene was responsible for the phenotype (Payne et al., 1993). The gene was identified in A. flavus by complementation and named qflR (Payne et al., 1993). Subsequently, aflR was identified in A. parasiticus (Chang et al., 1993) and A. nidulans (Yu et al., 1996). AflR - Function Several pieces of biochemical evidence indicate a regulatory role for aflR. Metabolite feeding studies and enzymatic activity measurements with A. flavus 650 (cg/IR mutant) and wild type A. flavus demonstrated that aflR is necessary for the aflatoxin biosynthetic enzyme activities to be detected (Payne et al., 1993). Later, studies demonstrated that aflR is required for aflatoxin biosynthetic gene transcript accumulation (Yu et al., 1996). Insertion of an additional copy of aflR into an A. parasiticus O- methylsterigmatocystin producing strain resulted in the overproduction of pathway intermediates including O-methylsterigmatocystin (Chang et al., 1993). In addition, 10 “balm; insertion of an additional copy of aflR in A. parasiticus resulted in the transcription of pathway genes under aflatoxin non-inducing conditions (Chang et al., 1995). Use of aflR linked with an inducible promoter in A. nidulans demonstrated that induction of ale in aflatoxin non-inducing conditions can activate genes in the biosynthetic pathway (Yu et al., 1996). Based on amino acid sequence identity, AflR belongs to a common class of fungal transcription factors called zinc binuclear cluster proteins that includes Gal4 (Woloshuk et al., 1994). While the amino-terminus of AflR contains the DNA binding domain, the carboxyl-terminus of AflR has a highly acidic region that functions as a transactivation domain (Chang et al., 1999). However, the total acidity in this region is not a major determinant in AflR transactivation (Chang et al., 1999). I DNA binding by AflR has also been investigated. Using methylation interference footprinting and electrophoretic mobility shift assays, the palindromic AflR binding site was first identified as TCGNNNNNCGA in the sth promoter in A. nidulans (Femandes et al., 1998). The consensus AflR binding cite was later defined as TCGSWNNSCGR (S = C/G, W = A/T, R = A/G) based on in vitro binding studies with recombinant AflR (Ehrlich et al., 1999b). Recombinant AflR binds to several aflatoxin biosynthetic gene promoters in vitro (Femandes et al., 1998, Ehrlich et al., 1999a , Ehrlich et al., 1999b and Miller et al., 2003a). AflR cis-acting sites have been shown to be necessary for transcriptional activation of several aflatoxin biosynthetic genes in vivo including 5th (Femandes et al., 1998), avnA (Ehrlich et al., 2002), kaA (Cary et al., 2000) and nor-1 (Miller et al., 2003a). AflR is necessary for transcriptional activation of several, if not all, aflatoxin biosynthetic genes. 11 AflR - Regulation Aflatoxin production is affected by environmental and nutritional factors such as pH, temperature, carbon source and nitrogen source (Miller et al., 2003c, Liu and Chu, 1998 and Luchese et al., 1993). Since AflR is a pathway regulator, the simplest model for these environmental and nutritional stimuli to impact aflatoxin gene transcription is directly through the aflR promoter. The timing of expression of aflR in response to these stimuli mimics the expression of aflatoxin structural genes (Miller et al., 2003c and Liu and Chu, 1998). A. parasiticus AreA (major nitrogen regulatory protein) can bind to the AflR promoter in vitro yet in vivo significance is unknown (Chang et al., 2000). A PacC (pH sensing) cis-acting site has also been identified in the aflR promoter using in vitro methods (Ehrlich et al., 1999a). A functional AflR binding site is located in the aflR promoter and is necessary for aflR transcription in vivo suggesting autoregulation (Ehrlich et al., 1999a). Additional studies on the transcriptional regulation of aflR are needed in order to understand how environmental, nutritional and even developmental stimuli affect aflatoxin biosynthesis. Evidence for Other Transcription Factors Though AflR is necessary for transcriptional activation of several, if not all, aflatoxin biosynthetic genes (Femandes et al., 1998, Ehrlich et al., 2002, Cary et al., 2000 and Miller et al., 2003a), there is evidence for the involvement of additional transcription factors. Studies with the kaA promoter (Ehrlich et al., 2002) provided evidence that both PacC (pH sensing) and BrlA (sporulation) can impact kaA transcriptional regulation through cis-acting sites in the pksA/nor—I intergenic region (Figure 1.4). However, deletion of the consensus cis-acting sites for PacC and BrlA did not affect 12 -1073 -520 +1 -1731 -1553 4205 ATG stop -65 - >-a.-C>- AflR3 AflR2 BrlA3 PacCl kaA ORF3 nor-1 Figure 1.4. Schematic of the nor-I/pksA intergenic region in A. parasiticus. The numbers indicate the number of nucleotides included upstream from the primary transcriptional start site of nor-1. Several potential cis-acting sites are indicated including the AflR binding sites AflRl, AflR2 and AflR3 and PacC l and BrlA3. The location of an open reading frame (ORF3) of unknown function is also shown. (Figure 1.4 adapted from Miller et al., 2003a) 13 nor-I transcriptional regulation in A. parasiticus under their culture conditions (Miller et al,2003a) The A. nidulans gene sth (A. parasiticus ver—I homologue) also appears to be regulated by additional transcriptional factors besides AflR. A full length .9th promoter fused to a GUS reporter resulted in only a 2-3 fold increase in activity when transformed into a wild-type strain compared to transformation into an AflR mutant strain (F emandes et al., 1998). In addition, substitution of both AflR binding sites only resulted in an approximately 5 fold reduction (not reduced to baseline expression levels) in GUS activity (Femandes et al., 1998). Due to the relatively high activity in the AflR mutant strain and the moderate decrease resulting from binding site substitution, the possibility exists that other transcriptional activators besides AflR are involved in sth transcriptional activation (Femandes et al., 1998). For avnA in A. parasiticus, substitution of the AflR binding site resulted in a 10 fold decrease in promoter activity (Cary et al., 2000). While there is no evidence of other transcriptional activators in the avnA promoter, a potential cis-acting site for a repressor of aflatoxin biosynthesis was located (Cary et al., 2000). Deletion of 78 bp upstream of an AflR binding site in an avnA::GUS reporter construct resulted in a 3 fold increase in reporter activity (Cary et al., 2000). In addition, protein extracts collected under non-aflatoxin inducing condition demonstrated specific binding to this region (Cary et al., 2000). The nor-1 gene is perhaps the most studied of all the aflatoxin structural genes. A detailed analysis of the nor-1 promoter has identified several cis-acting sites including AflRl, AflR2, a putative TATA box, CREbp and norpr (Miller et al., 2003a and Miller et al., 2003b - Figure 1.4 and 1.5). Although putative TATA boxes have been identified in aflatoxin biosynthetic promoters, the nor-1 TATA box was the first to be rigorously l4 m ‘2; Lung} "Va ., ‘0. I i»: If." -238 -210 -76 -65 -52 -47 +15 +22 —- NorL AflR TATA CRE Figure 1.5. Identification of cis-acting sites in the nor-I promoter of A. parasiticus. The numbers indicate the number of nucleotides from the primary transcriptional start site of nor-1. (Figure 1.5 adapted from Miller et al., 2003b) 15 tested for functional significance (Miller et al., 2003b). Substitution of the TATA box in the context of a larger promoter resulted in non-detectable GUS activity (Miller et al., 2003b). A novel cis-acting site (norL) was identified that is necessary for maximum nor- ] transcriptional activation in vivo. Using electrophoretic mobility shift assays (EMSA), specific protein binding to norL was demonstrated (Miller et al., 2003b). EMSA also identified another potential cis-acting site, CREl, in the nor-1 promoter (Miller et al., 2003b and Miller et al., 2003c). Both the norL binding protein and CREl binding protein appear to rely on functional AflR for maximum DNA binding (Miller et al., 2003b and Miller et al., 2003c). The transcriptional regulation of nor—1 appears to involve additional proteins besides AflR. ORF 3 and nor-I/pksA transcriptional activation Reporter studies with pksAzzGUS (Cary et al., 2000) and nor-[::GUS (Miller et al., 2003a) both demonstrated that inclusion of AflR2 (Figure 1.4) resulted in greater transcriptional activation. We propose two alternative models to explain these data: 1) AflR2 works synergistically with AflRl and AflR3 to mediate transcription of the nor-l and pksA promoters respectively; or 2) AflR2 mediates expression of ORF 3 (potentially encodes a polypetide of approximately 300 amino acid residues - Figure 1.4) directly downstream from AflR2 which directly or indirectly impacts transcription of the nor-1 and kaA promoters. Identification of a cDNA corresponding to ORF 3 in an A. parasiticus cDNA library strongly suggests it represents a functional gene. Interestingly, blast searches using ORF 3 as a query sequence have not provided solid clues regarding potential function. Model 2 allows us to make 2 related predictions regarding nor-1 and pksA promoter function. 1) Since fungal isolates carrying nor-1 ::GUS or pksA::GUS 16 '7'“. ~. I! ’51 I ‘51,) a (‘ constructs with promoter fragments that include a functional AflR2 and ORF 3 show the highest GUS expression levels, accumulation of additional ORF3 protein in strains with two copies (native plus plasmid copy) overcomes a protein threshold resulting in extreme upregulation of nor-1 and pksA promoter activity. 2) Loss of ORF3 function due to AflR2 deletion in the 2"d copy accounts for downregulation (or lack of upregulation) of both nor-1 and kaA expression. Similar results are seen with the insertion of an additional copy of AflR (Chang et al., 1993, Chang et al., 1995 and Chang et al., 2001) suggesting a possible regulatory role for ORF 3. Future experiments will determine the function of ORF3 and how AflR2 impacts pksA and nor-I transcriptional activation. AflJ The function of aflJ in aflatoxin biosynthesis is still unclear. The aflJ gene resides adjacent to aflR in the aflatoxin gene cluster with the two genes being divergently transcribed. An A. flavus aflJ knockout strain does not make aflatoxin and lacks the ability to convert several aflatoxin intermediates to aflatoxin (Meyers, et al., 1998). However, the aflJ knockout strain does accumulate several aflatoxin biosynthetic transcripts under aflatoxin conducive conditions suggesting that aflJ is not involved in the transcriptional regulation of aflatoxin biosynthesis (Meyers et al., 1998). Sequence analysis of aflJ did not reveal any enzymatic function but did identify three potential membrane-spanning domains and a putative microbody targeting signal (Meyers et al., 1998). Consequently, Meyers et al. proposed two hypotheses regarding AflJ function: 1) AH] is involved in either transmembrane transport of aflatoxin pathway intermediates through intracellular compartments; or 2) AflJ is involved in the localization of pathway enzymes to an organelle (Meyers et al., 1998). Conversely, AflJ has been shown to 17 5.... o 0 7a interact with AflR in a two-hybrid assay (Chang and Yu, 2002). Insertion of an additional copy of aflR and aflJ into A. parasiticus resulted in greater aflatoxin biosynthesis than insertion of aflR alone while insertion of aflJ alone had no affect on aflatoxin biosynthesis (Chang et al., 2001). Consequently, Chang et al. have described qflJ as being a transcriptional co-activator (Chang et al., 2001). More work is needed to clearly define the activity of AflJ. Regulation of Trichothecene Gene Transcrigtion Tri6 - Discovery Following the identification of three trichothecene biosynthesis genes, T rli5 (Hohn and Desjardins, 1992), Tri4 (Hohn et al., 1995) and Tri3 (McCormick et al., 1996), it was realized that they all resided in a 9-kb region and that the trichothecene biosynthesis genes might be clustered (Hohn et al., 1993b). The location of aflR within the aflatoxin biosynthetic gene cluster (Payne et al., 1993) suggested the possiblity that other mycotoxin gene clusters, including trichothecene, may be regulated by a gene present within the cluster. While the discovery of aflR relied on complementation of a regulatory mutant (Payne et al., 1993), the initial identification of Tri6 as a putative trichothecene biosynthesis transcriptional activator relied on additional sequencing of the trichothecene gene cluster (Proctor et al., 1995). Found immediately upstream of T ri5 , Tri6 is an Open reading frame of 2 1 7 amino acids with regions similar to CyszHis2 zinc finger proteins (Proctor et al., 1995). The CyszHis2 zinc finger is a common motif for transcription factors including BrlA from A. nidulans (Adams et al., 1988). 18 Tri6 - Function Several lines of evidence, in addition to sequence data, demonstrate that Tri6 is a transcriptional activator for trichothecene biosynthesis. Tri6 expression mirrored the expression of other trichothecene genes including Tri3 and T ri4 (Proctor et al., 1995). Disruption of T ri6 resulted in a strain with greatly reduced trichothecene production, trichothecene enzyme activities and trichothecene biosynthetic transcript steady-states (Proctor et al., 1995). Tri6 functioned as a transcriptional activator in Saccharomyces cerevisiae when fused to the DNA binding domain of Gal4 (Proctor et al., 1995). The Tri6 binding site was identified in F usarium sporotrichioides as YNAGGCC using electrophoretic mobility shift assays with Tri6 produced in vitro (Hohn et al., 1999). The Tri6 binding site was confirmed in vivo using F. sporotrichioides Tri4 reporter strains (Hohn et al., 1999). All of the trichothecene biosynthetic genes identified in the F. .s'porotrichioides and F. graminearum clusters have the Tri6 binding site located in their promoter regions except for Tri 10 (Brown et al., 2001 and Hohn et al., 1999). Tri10 - Discovery Located upstream of Tri5 in the trichothecene gene cluster (Figure 1.6) , Tril 0 expression mirrored the timing of several trichothecene genes including T ri6, Tri5 and Tri4 (Tag et al., 2001). A TrilO disruption strain accumulated significantly less trichothecene biosynthetic transcripts (Tag et al., 2001). Conversely, transformants that had increased T ril 0 expression resulted in significantly increased trichothecene gene expression (Tag et al., 2001). As noted above, TriIO lacks a consensus Tri6 binding site (Brown et al., 2001) and is significantly upregulated in the Tri6 disruptant strain (Tag et al., 2001). Tag et al. postulated that T ri10 is not positively regulated by Tri6 but instead 19 arm -.- 7 7 l fl l Tri cluster genes T-2 Toxin /' ? —> TrilO—> "ll“ri6\A U Tri101 Self-protection / ? ? Figure 1.6. Proposed regulatory model for trichothecene biosynthesis. Solid arrows indicate positive activators while open arrows indicate inhibitory activities. Question marks indicate other proposed but unknown regulatory signals or factors. (Figure 1.6 adapted from Tag et al., 2001) 20 THE 4.- " ,- Q‘fflnf may be negatively regulated by unknown mechanisms when Tri6 is present (Tag et al., 2001). The impact of Tri10 extends to genes involved in the primary metabolic steps that generate trichothecene precursors including Fpps (Tag et al., 2001). While Tri10 lacks any known transcription factor motif, its function is clear as an essential regulator in trichothecene gene regulation (Tag et al., 2001). Future investigations are focused on determining the precise mode of action of Tri10 and on further identification of the regulatory circuits defined by Tri 1 0 (Tag et al., 2001). Regulation of Biosynthesis Gene Transcription for other Mycotoxins Although most information regarding mycotoxin gene regulation is centered on aflatoxin and trichothecene biosynthesis, other mycotoxin related transcription factors have been identified. In fact, information about aflatoxin and trichothecene biosynthesis pathways have aided in the analysis of different mycotoxin pathways. For example, a Tri6 homolog has been identified (MRTri6) in the distantly related trichothecene pathway of Myrothecium roridum (Hohn et al., 1999). Dothistroma pini synthesizes dothistromin, a difuranoanthraquinone toxin similar to aflatoxin, with several analogous genes (Bradshaw et al., 2002). Bradshaw et al. hope to potentially identify the dothistromin pathway regulator by using A. parasiticus aflR as a probe and/or by sequencing the entire dothistromin gene cluster (Bradshaw et al., 2002). Sequencing of the paxilline biosynthesis cluster in Penicillium paxilli found two potential transcription factors, pcch and paxS (Young et al., 2001). Like aflR, both paxR and paxS contain the uniquely fungal Zn(II)2Cys6 binuclear cluster DNA binding motifs (Young et al., 2001). It is currently unknown the exact function of paxR and paxS in paxilline biosynthesis. 21 Mycotoxin Gene Clusters While common in prokaryotes, the linkage of functionally related genes is relatively rare in higher eukaryotes. In filamentous fungi, however, the clustering of genes is a common feature for several metabolic pathways. Examples of gene clusters in secondary metabolism include aflatoxin (Trail et al., 19953), trichothecene (Brown et al., 2001), penicillin (Diez et al., 1990), fumonisin (Seo et al., 2001), dothistromin (Bradshaw et al., 2002), paxilline (Young et al., 2001) and several others. Several nutrient utilization pathways are also clustered in filamentous fungi including ethanol ( F illinger and F elenbok, 1996) and nitrate (J ohnstone et al., 1990) utilization in A. nidulans. The physical linkage of related genes suggests two possible hypotheses: l) linkage of metabolic pathway genes provides a means to regulate pathway gene transcription; and 2) linkage provides a means for genetic transfer of entire pathways between species. Both of these hypotheses are discussed below. Organization of Gene Clusters Aflatoxin Gene Cluster Analysis of aflatoxin pathway mutants first established the likely clustering of at least some of the aflatoxin biosynthetic genes (reviewed in Bennett and Papa, 1988). Subsequently, the first two aflatoxin genes identified, nor-1 and vcr-I, were found to localize to the same A. parasiticus cosmid (Trail et al., 1995b). The aflatoxin gene cluster has been mapped in A. parasiticus (Trail et al., 1995b), A. flavus (Yu et al., 1995a) and A. nidulans (Brown et al., 1996). The timing of transcript accumulation ofall the cluster genes is consistent with their involvement in aflatoxin biosynthesis (Trail et al., 1995b and Brown et al., 1996). In fact, many of the genes have been studied in detail 22 ”"211 I flap “3;..537efiw I a»: _. and been shown to be involved in aflatoxin biosynthesis. Interestingly, the aflatoxin gene clusters of these three related organisms are not organized identically. For example, the distance between aflR and ver-I is 32 kb in A. nidulans but 8 kb in A. parasiticus and A. flavus. The maintenance of gene clusters despite changes in gene order suggests that gene clustering has functional significance for the fungi. A 5 kb spacer region is located at one end of the aflatoxin gene cluster (Yu et al., 2000a and Yu et al., 2000b). The spacer region contains no open reading frames (ORFs) and has a sugar utilization cluster located on the other side of the spacer region (Figure 1.7 — Yu et al., 2000a and Yu et al., 2000b). Aflatoxin production is closely linked to carbon source with simple sugars like glucose and sucrose able to induce aflatoxin biosynthesis (Buchanan and Lewis, 1984). Consequently, the localization of this sugar utilization cluster near the aflatoxin cluster suggests a regulatory connection between the two clusters. However, of the four genes in the sugar cluster, only hxtA (proposed hexose transporter protein) expression was shown to be concurrent with aflatoxin pathway cluster genes (Yu et al., 2000a). It is unknown if there are any functional AflR cis-acting sites in the sugar cluster gene promoters. In addition, the border at the other end of the aflatoxin gene cluster has not yet been defined. Trichothecene Gene Cluster Two overlapping cosmid clones were able to complement different trichothecene mutants suggesting that the trichothecene genes were clustered in F usarium sporotrichioides (Hohn et al., 1993b). A 23 kb trichothecene gene cluster has been sequenced for both F. sporotrichioides and F. graminearum and contains 12 genes (Brown et al., 2001). All of the 12 clustered genes studied so far have been shown to be 23 mfi. ‘. 7.. 77:5. :1 I 5 kb spacer I I | I 4 kb spacer moxY nadA hxtA gch sugR Figure 1.7. Schematic of the sugar utilization gene cluster in A. parasiticus. The moxY gene is at one end of the aflatoxin gene cluster. The spacer regions on either side of the sugar utilization cluster do not contain open reading frames. Map is roughly to scale. (Figure 1.7 adapted from Yu et al., 2000a) 24 involved in trichothecene biosynthesis (Figure 1.8) (reviewed in Brown et al., 2001). Yet, the 12 genes in the identified trichothecene gene cluster are insufficient to account for all known trichothecene structures (Brown et al., 2001) leading to two hypotheses: 1) additional trichothecene biosynthesis genes are located beyond the flanking sequence of tri8 and tri12; and 2) all trichothecene genes are not located within the gene cluster. Since most, if not all, of the genes necessary for aflatoxin biosynthesis are located within the aflatoxin gene cluster, it would be reasonable to sequence beyond tri8 and tri12 to potentially identify additional trichothecene genes. In addition, it will be interesting to determine if there is an extended spacer region that forms a border of the trichothecene gene cluster as there is with the aflatoxin gene cluster. tri101 exists outside of the 23 kb cluster in both F. sporotrichioides (McCormick et al., 1999) and F. graminearum (Kimura et al., 1998a) and is the only known trichothecene gene to exist outside of the cluster. The genes on either side of tri 101 (UTP-ammonia ligase and phosphate permease) are not involved in trichothecene biosynthesis and tril 01 is at least 35 kb from either end of the identified trichothecene cluster in F. graminearum (Kimura et al., 1998b). TrilOl is a 3-O-acetyltransferase that is required for T-2 production in F. sporotrichioides (McCormick et al., 1999). Since the enzymatic products of Tri101 are less toxic than the substrates, it was originally proposed that the purpose of TrilOl was to protect the fungus (Kimura et al., 1998a). Expression of tri I 01 in yeast (Kimura et al., 1998a and McCormick et al., 1999) and plants (Muhitch et al., 2000) resulted in increased tolerance for trichothecenes. Yet, a F. sporotrichioides tri] 01 disruptant could both germinate and grow in the presence of trichothecenes suggesting that tri101 is not an essential self-defense mechanism for F. sporotrichioides (McCormick et al., 1999). The evolution oftrilOI is discussed in later sections. 25 F. sporotrichiOideS W 8 7 3 46 510911 12 F. graminearum 87346 51091112 Figure 1.8. Genomic organization of the trichothecene biosynthetic gene cluster of F usarium sporotrichioides and F. graminearum. Arrowheads indicate the direction of transcription and the number underneath each arrow refer to the specific gene. Genes with the same number from both F usarium species are homologues. tri7 in F. graminearum is non-functional. Map is roughly to scale. (Figure 1.8 adapted from Brown et al., 2001) 26 F umonisin Gene Cluster The identification of the fumonisin gene cluster utilized allele tests with three different F usarium monoliforme fumonisin mutants (Desjardins et al., 1996). Subsequently, a polyketide synthase gene, fum5 , was isolated and shown to be required for fumonisin biosynthesis (Proctor et al., 1999). Based on the structure of fumonisin B1, at least 8 different enzymatic steps are needed for fumonisin biosynthesis (Seo et al., 2001). Sequencing downstream from fum5 identified four additional ORFs,fum6, -7, -8 and -9, whose expression is correlated with fumonisin production (Figure 1.9 — Seo et al., 2001). Gene disruption analysis of fum6 and fum8 revealed that they are necessary for fumonisin biosynthesis (Seo et al., 2001). Nucleotide sequence analysis in the DNA regions flanking fum5 and fum9 may identify additional fumonisin biosynthetic genes. Based on aflatoxin and trichothecene gene clusters, it is reasonable to expect that a transcriptional activator specific for fumonisin biosynthesis located within the fumonisin gene cluster will be found with the additional sequencing. Impacts of Mycotoxin Gene Clusters Identification of Gene-Cluster-Dependent Regulation Preliminary evidence for a role for clustering in aflatoxin gene regulation was reported by Liang et al. (Liang et al., 1997). The promoter of the aflatoxin biosynthesis structural gene ver-I was fused to uidA (encodes B-glucuronidase [GUS]) to generate a reporter plasmid (pHD6.6) that contained niaD (encodes nitrate reductase) as a selectable marker (Liang et al., 1997). pHD6.6 integrated predominantly at the ver—I or niaD locus via homologous recombination (Liang et al., 1997). Single—copy integration of pHD6.6 at niaD resulted in a 500-fold reduction in ver-I promoter activity when compared with 27 a -. I.“ .41 fum5 fum6 firm 7 fum8 film9 w Figure 1.9. Schematic of the fumonisin biosynthetic gene cluster in F usarium verticillioides. Arrowheads indicate the direction of transcription. Map is roughly to scale. (Figure 1.9 adapted from Seo et al., 2001) 28 .. .J-V’ single-copy integration at the ver-I locus; however, the temporal pattern of expression appeared to be similar at both loci (Liang et al., 1997). One explanation for reduced ver- 1 promoter function at the niaD locus is that location in the aflatoxin cluster results in positive, position-dependent regulation of ver—I exression. An alternative hypothesis is that the expression of ver-I integrated at niaD is negatively influenced by niaD regulation. In the absence of preferred nitrogen sources and in the presence of nitrate, niaD is expressed. Under the rich growth conditions tested by Liang et al. (Liang et al., 1997), niaD may have been repressed, and therefore the lack of ver-l expression at the niaD site could have resulted from niaD-dependent regulation. Subsequently, the promoter of the aflatoxin biosynthesis gene nor-1 was fused to GUS to generate a reporter plasmid, pAPGUSNN-B, containing niaD as a selectable marker (Chiou et al., 2002). Transformants with pAPGUSNN-B integrated at the niaD locus had no detectable GUS activity while nor-I integrants had GUS activity (Chiou et al., 2002). In addition, the cloned nor—I promoter functioned similarly to the native nor-I promoter when it integrated at the nor-1 locus (Chiou et al., 2002). Because niaD dependent regulation could account for the absence of expression at niaD for both nor- ] ::GUS (Chiou et al., 2002) and ver-IzzGUS (Liang et al., 1997), a third chromosomal location was analyzed using pAPGUSNP, which contained nor-1 ::GUS plus pyrG (encodes OMP decarboxylase) as a selectable marker (Chiou et al., 2002). GUS expression was detectable only when pAPGUSNP integrated at nor-1 and was not detectable at pyrG, even under growth conditions that required pyrG expression (Chiou et al., 2002). While the mechanism is unknown, nor-l and perhaps other aflatoxin biosynthetic genes are susceptible to aflatoxin gene-cluster-dependent regulation. 29 The genome of some strains of A. parasiticus includes a partial duplication of the aflatoxin gene cluster (Chang and Yu, 2002). The region from aflR to ver-I plus omtB is duplicated in A. parasiticus SUI (Chang and Yu, 2002). It is unknown why the genes between ver-I and omtB are not in the duplicated region. All of the duplicated genes appear to have mutations in them that would make them non-functional except for aflR-Z and aflJ-Z (Chang and Yu, 2002). Northern and RT-PCR analyses of RNA indicated that qflR-Z is expressed at much lower levels than aflR-I (Cary et al., 2002). Nucleotide sequence analysis upstream of the aflR-Z translational start codon revealed that the AflR binding site was intact and that there were few base changes (2%) compared to the corresponding region of aflR-I (Cary et al., 2002). Cluster-dependent regulation may explain the poor expression of aflR-Z (Cary et al., 2002 and Chang and Yu, 2002). It is currently unclear if the other genes in the duplicated region are expressed at similar levels to their cluster counterparts. Reporter constructs that can integrate into the aflatoxin gene cluster and the duplicated region may provide valuable insight into the mechanism of cluster-dependent regulation. Trichothecene biosynthesis genes are also clustered (Hohn et al., 1993b) and there is some evidence for position-dependent regulation with the pathway regulator tri6 (Chen et al., 2000). Introduction of a plasmid containing a tri5::GUS fusion with a functional tri6 resulted in 50 to 100 fold more GUS activity with tri5 integration compared to ectopic integration (Chen et al., 2000). However, there were no differences in GUS activity between tri5 and ectopic integration of a tri5::GUS plamid without a functional tri6 (Chen et al., 2000). Integration of a tri4::GUS fusion into the M4 locus resulted in 2 to 5 fold more GUS activity than ectopic integration (Hohn et al., 1999). Additional 30 experiments are necessary to clarify the significance of cluster-dependent regulation in trichothecene biosynthesis. It is currently unknown if cluster-dependent regulation occurs with other mycotoxin gene clusters like fumonisin. However, cluster-dependent regulation appears to be less significant with trichothecene biosynthesis than with aflatoxin biosynthesis. Except for aflatoxin biosynthesis, the description of mycotoxin gene-cluster-dependent regulation needs further study. While the construction of plasmids with mycotoxin promoters fused to reporter genes is useful in identifying cis-acting sites in the mycotoxin promoter, they also provide a means to determine differences between mycotoxin cluster integration and ectopic integration. Significance of Gene-Cluster-Dependent Regulation The targeting of reporter constructs to specific chromosomal locations for promoter analysis has been suggested by other investigators studying expression of fungal genes (Hamer and Timberlake, 1987 and Timberlake and Marshal, 1988 and van Gorcom et al., 1986). The gene-cluster-dependent regulation seen with the aflatoxin gene cluster emphasizes the importance of determining the site of integration for mycotoxin reporter plasmids. In addition, screening transformants by genotype (site of integration) rather than phenotype (reporter activity) will prevent possible sampling bias. A rapid method for screening the site of integration for A. parasiticus has been described (Chiou et al., 2002) that makes screening by genotype feasible. Although the mechanisms of position-dependent gene expression have not been fully elucidated in filamentous fungi, it has been hypothesized that enhancer elements may be responsible for the position-dependent effect (Kinsey and Rambosek, 1984). 31 Studies performed on the SpoCl gene cluster in A. nidulans showed that clustered genes can be coordinately expressed during development and that placement of cluster genes at ectopic chromosomal locations results in the loss of that coordination (Miller et al., 1987 and Timberlake and Barnard, 1981). The hypothesis that positive cis-acting factors have regional control over the transcription of aflatoxin genes is reasonable, since removal of aflatoxin reporter fusions from the aflatoxin gene cluster results in reduced GUS expression. More recently, studies with the mammalian [i-globin gene identified a region named a locus control region (LCR) (Grosveld et al., 1987). The B-globin LCR was identified as a region that was necessary to confer positional independence of a [i-globin transgene (Grosveld et al., 1987). Subsequently, several additional LCRs have been identified (reviewed in Li et al., 2002). We hypothesize that aflatoxin gene expression is influenced by a locus control region. However, a locus control region has not been identified in fungal gene clusters. If a LCR does influence the regulation of nor-1 transcription, it is located at least 3 kb upstream from the transcription initiation site in the 5' nor-I region, at least 1.8 kb downstream of the transcription termination site in the 3' region of nor-1, or within the nor-1 coding region. Because similar position-dependent expression is observed with the ver-I ::GUS reporter construct (Liang et al., 1997), it is possible that the same LCR element is influencing the regulation of both nor-1 and ver-I . Comparison of steady state levels of mRNA transcripts from genes present in the aflatoxin gene cluster and the duplicated region (ie ver-IA vs. ver—IB) may help narrow the search for the LCR(s). It will be interesting to determine if the duplicated region includes the LCR(s) necessary for cluster-dependent regulation. In addition, the 5 kb spacer region at one border of the aflatoxin gene cluster is a tempting place to look for 32 possible boundary elements. Boundary elements appear to provide a functional boundary for both accessible and inaccessible chromatin (Labrador and Corces, 2002). The possible boundary element in the spacer region would prevent the LCRs from affecting transcriptional activation of genes in the sugar cluster or beyond. While solid evidence for cluster-dependent regulation has only been shown for aflatoxin biosynthesis, it is reasonable to predict that several other mycotoxin gene clusters will also be subject to cluster-dependent regulation. Consequently, the proper use of reporter plasmids to study mycotoxin gene transcriptional regulation must include the identification of the site of integration of the plasmids to ensure proper data interpretation. Evolution of Mycotoxin Gene Clusters Origin of the Aflatoxin Gene Cluster The occurrence of fungal pathway gene clusters may result from horizontal gene transfer from prokaryotes where clustering of pathway genes is common (Hohn and Keller, 1997). Perhaps the strongest case for the horizontal gene transfer of an entire secondary metabolic pathway from bacteria to fungi is with penicillin biosynthesis (Aharonwitz et al., 1992 and Weigel et al., 1988). The G+C content of the penicillin gene cluster is more similar to Streptomyces than the producing fungi and the fungal penicillin genes lack introns which is generally a characteristic of bacteria (Hohn and Keller, 1997). Horizontal gene transfer is unlikely with the aflatoxin biosynthetic pathway due to the presence of introns and similar G+C content inside and outside of the aflatoxin cluster (Brown et al., 1996). Yet the changes in gene order within the aflatoxin gene cluster of different Aspergilli suggests that there is selective pressure to keep the 33 nu... , d. a m. 5175 0 aflatoxin genes clustered. Dothistroma pini, a fungal pine pathogen, produces the mycotoxin dothistromin that is structurally similar to versicolorin A, an intermediate in aflatoxin biosynthesis (Bradshaw et al., 2002). Four genes identified in the D. pini dothistromin gene cluster all have high similarity with known aflatoxin biosynthesis genes (Bradshaw et al., 2002). Detailed analysis of the dothistromin and aflatoxin gene clusters may help explain the evolutionary history of the aflatoxin gene cluster. Origin and Diversity of Trichothecene Gene Clusters The current literature provides few clues for the origin of the trichothecene gene cluster. The ten trichothecene genes identified so far in the cluster (Tri-3 through Tri-12) contain a total of 37 introns (Brown et al., 2001) suggesting that horizontal gene transfer from prokaryotes is unlikely. While there are many different trichothecenes that are produced by various fungi, F usarium species appear to be the primary source of trichothecenes in agricultural products (Hohn et al., 1999). The mechanism for how different Fusarium species produce different trichothecenes is unknown. The T-2 gene cluster in F usarium sporotrichioides and the deoxynivalenol gene cluster in F. graminearum are very similar (see Figure 1.8) in that the 23 kb region included 12 homologous genes (Brown et al., 2001). However, Tri7 (acetylates the oxygen on O4) is non-functional in F. graminearum (Brown et al., 2001) suggesting a possible mechanism for generating trichothecene structural diversity. The origin of the non-cluster tri101 gene may be different than the clustered trichothecene genes (Kimura et al., 1998c). Kimura et al. reasoned that a translocation event was unlikely to explain the location of tri101 because it is flanked by essential primary metabolism genes (Kimura et al., 1998b). In addition, homologues of tril 01 were found in Saccharomyccs cerevisiae and 34 '1'." 1‘ anLE 2:3}— . . . 131341 Schizosaccharomyces pombe (Kimura et al., 1998b). Consequently, Kimura et al. reasoned that it is feasible that trichothecene producers have acquired tri101 through horizontal gene transfer (Kimura et al., 1998b). ACKNOWLEDGMENTS Two sections from Chapter 1 (Transcriptional Regulation of Mycotoxin Biosynthesis and Mycotoxin Gene Clusters) will appear in the chapter titled “Fungal Genetics and Mycotoxin Biosynthesis” in Food Biotechnology. The chapter will be co- authored with Dr. John Linz who has written other sections of the chapter. The I publication date for Food Biotechnology hasn't been set but we expect 2004. 35 'hq’ l i INt L‘ -| r f‘ .175 0 CHAPTER 2 Effects of nitrogen source, carbon source and zinc concentration on the regulation of the aflatoxin biosynthetic gene nor-I from Aspergillus parasiticus INTRODUCTION Aflatoxins are highly toxic and carcinogenic secondary metabolites of certain strains of Aspergillus parasiticus, A. flavus and A. nomius (Wilson and Payne, 1994). Several different crops have been found to be contaminated with aflatoxins including peanuts, cottonseed, corn, pistachios, walnuts, and almonds (Dvorackova, 1990). Epidemiological data have linked aflatoxins with human hepatic cancer, primarily in countries where the elimination of contaminated products is not practiced (Reviewed in Dvorackova, 1990 and Hall and Wild, 1994). Aflatoxin carcinogenicity studies clearly demonstrate its high potency in various laboratory animals (reviewed in World Health Organization, 1979). Due to the animal and human toxicology data, aflatoxin susceptible crops are monitored for aflatoxin contamination in the United States with an action level of 20 ppb for food for human consumption, 0.5 ppb for milk and 20—300 ppb for most animal feeds (Council for Agricultural Science and Technology, 2003). Due to health and economic concerns, the elimination of aflatoxin from the food chain is desirable. One approach to achieve the goal of aflatoxin elimination has been to elucidate the biosynthesis of aflatoxin at the molecular level. Aflatoxin biosynthesis is a complex process that requires at least 16 different enzymatic steps (Bhatnagar et al., 1994). The aflatoxin biosynthesis genes reside in a 70 kb cluster and appear to be co-regulated (Trail 36 § (Q\>‘ ,3 et al., 1995b). Located within the aflatoxin gene cluster are the genes nor-1, ver-l and aflR. Nor-l catalyzes the conversion of the first stable aflatoxin biosynthesis intemediate, norsolorinic acid, to averantin (Zhou and Linz, 1999). The nor-1 gene is physically located between pksA and fas2, two of the first three (fasl is the other) structural genes in aflatoxin biosynthesis (Trail et al., 1995b). Ver-l is associated with the conversion of versicolorin A to sterigmatocystin, a late step in aflatoxin biosynthesis (Skory et al., 1992). The ver-I gene is physically located at the other end of the aflatoxin gene cluster from nor-I . Analysis of nor-1 and ver-I transcriptional activation provides a good cross-section for the aflatoxin biosynthesis structural genes. The accumulation of nor-1 and ver-l transcripts were shown to be coregulated (Skory et al., 1993) along with several other aflatoxin biosynthesis genes (Trail et al., 1995b). Analysis of the regulatory mutant A. flavus strain 650 (Bennett and Papa, 1988) first identified aflR as a pathway regulator (Payne et al., 1993). Subsequently, aflR has been identified in A. parasiticus (Chang et al., 1993) and A. nidulans (Yu et al., 1996). While AflR has been clearly implicated in the regulation of specific aflatoxin biosynthetic genes (Cary etal., 2000, F emandes et al., 1998, Ehrlich et al., 2002 and Miller et al., 2003a), it is unclear whether AflR alone is sufficient to stimulate transcription of all aflatoxin biosynthetic genes. Environmental and nutritional factors such as temperature, pH, carbon source, nitrogen source and zinc concentration all strongly influence aflatoxin accumulation in laboratory media (reviewed in Luchese and Harrigan, 1993). Defined media such as Reddy’s, GMS (glucose minimal salts) and PMS (peptone minimal salts, semi-defined) were designed to investigate the effects of metals, nitrogen source and carbon source on aflatoxin production. Several researchers have demonstrated that glucose as a sole 37 carbon source supports aflatoxin production while peptone does not (Buchanan and Stahl, 1984, Abdollahi and Buchanan, 1981a and Abdollahi and Buchanan, 1981b). Nutritional shift experiments with transcriptional and translational inhibitors indicated that glucose stimulates de novo synthesis of aflatoxin biosynthesis transcripts and proteins (Abdollahi and Buchanan, 1981b). Various nitrogen sources including amino acids, nitrate and ammonium have been extensively studied in Aspergillus parasiticus growth and aflatoxin formation (reviewed in Luchese and Harrigan, 1993). While reports vary with what nitrogen source results in the most aflatoxin production, it is clear that ammonium is significantly more stimulatory than nitrate with ammonium nitrate (N H4NO3) intermediate. Nitrate as the sole nitrogen source has been shown to inhibit de novo synthesis of aflatoxin biosynthesis enzymes (Kachholz and Demain, 1983). Metal ions also can affect aflatoxin biosynthesis as well. In particular, omission of zinc in the culture medium results in non-detectable levels of aflatoxin (Coupland and Niehaus, 1987 and Bennett et al., 1979). In addition, researchers have shown a correlation between zinc concentration in corn and their aflatoxin concentration (Failla et al., 1986). Nutritional factors such as carbon source, nitrogen source and zinc concentration provide a useful tool in studying the molecular mechanisms of aflatoxin biosynthesis gene regulation. Our hypothesis is that these nutritional factors affect transcriptional regulation of aflatoxin biosynthetic genes. The purpose of this study was to use nutritional factors to study the regulation of the aflatoxin biosynthesis genes nor-1, ver-I and aflR in batch cultures using the nor-1::GUS reporter strain D8D3 and the ver-l ::GUS reporter strain 14. The nor-l ::GUS and ver-1::GUS strains permit the study of transcriptional activation of nor-l and ver-I, respectively, in these various media. GMS (glucose minimal salts) 38 TH \ 57m supported aflatoxin biosynthesis, nor-1, ver-I, and aflR transcript accumulation, Nor-1 and Ver-l protein accumulation and nor-1 ::GUS and ver-I ::GUS activity. PMS (bacto- peptone minimal salts - D8D3 only) and NMS (glucose minimal salts with nitrate), supported low to non-detectable aflatoxin biosynthesis, nor-l, ver-l, and aflR transcript accumulation, Nor-1 and Ver-l protein accumulation and nor-1::GUS and ver-l ::GUS activity. Aflatoxin biosynthesis with GMS0 (glucose minimal salts with no Zinc added) was similar to GMS (glucose minimal salts) at 24 hours but aflatoxin production, transcript accumulation, protein accumulation and GUS activity decreased thereafter. Using a 160 base pair nor-1 promoter fragment as probe (nor-R), electrophoretic mobility shift assays (EMSA) with A. parasiticus SUI (wild type) and AF S10 (AflR knock-out) PMS extracts both contained one specific complex of similar mobility that did not bind to the AflR binding site (TCGnnnnnCGA) that is included in nor-R. GMS protein extracts from A. parasiticus AF SlO produced no specific shifted complexes with nor-R. A. parasiticus SUI GMS protein extracts produce a specific complex with nor-R (different mobility than the specific shifts seen with the PMS extracts) that also does not involve the AflR binding site. Competition experiments localized the potential cis—acting site in nor-R to a region designated CREI. This work is a comprehensive analysis of the use of nutritional components as tools to study aflatoxin gene transcriptional regulation. MATERIALS AND METHODS Strains and Growth Conditions A. parasiticus D8D3 contains a nor-1::GUS (GUS = uidA from Escherichia coli) translational fusion at the 3' region of nor-1 (Chiou et al., 2002). A. parasricus Isolate 4 39 lfi'n’s AI-HL] (14) contains the ver—I ::GUS translational fusion at the 3' region of ver-l (Liang et al., 1997). A. parasiticus SUI is an wild type aflatoxin producing strain. A. parasiticus AF S10 is an aflR knock-out strain (Cary et al., 2002). GMS and PMS media were made as described by Buchanan with the exception of the changes described in Table 2.1 (Buchanan and Lewis, 1984). For liquid culture experiments, 100 mL of medium in a 250 mL flask with 5, 6-mm, glass beads were inoculated with 2 X 106 spores and incubated at 29°C in the dark with shaking at 150 rpm. D8D3 and 14 were grown in triplicate for each time point. The time points for D8D3 were 36, 48 and 72 h. 14 grows slower than D8D3 and there was not enough mycelia for analysis at 36 h. Consequently, the time points for 14 were 48, 60 and 72 h. YES medium (2% yeast extract, 6% sucrose; pH=5.8) was used for Nor-1 :Maltose binding protein studies. Table 2.1. Medium components used in this study. mediuma carggn source nitrgggn gource Zing ggngentration PMS peptone ammonium (+ peptone) 10 mm NMS glucose nitrate 10 mm GMS0 glucose ammonium 0 mm GMS glucose ammonium 10 mm " Base medium was the same for all as previously described (Buchanan and Lewis, 1984). RNA and Protein Extraction Liquid cultures of A. parasiticus D8D3 and 14 were filtered at the indicated time points through miracloth (Calbiochem, La Jolla CA), the mycelia was then frozen with liquid nitrogen and stored at -80°C. For RNA extraction, approximately 200 mg of 40 frozen mycelia was macerated with a mortar and pestle. TriZOL (Gibco BRL, Grand Island NY) was used to extract total RNA from the ground mycelia following manufacturers instructions. For protein extraction, approximately 200 mg of frozen mycelia was macerated with a mortar and pestle. The ground mycelia was suspended in 0.5 ml of GUS lysis buffer (50 mM NaHzPO4, 10 mM EDTA, 0.1% Triton X-100, 0.1% SDS, 10 mM B-mercaptoethanol, pH 7.0), vortexed for 15 seconds and centrifuged for 10 minutes at 10,000 X g. The supematent was carefully removed and placed in a new tube and stored on ice. The protein concentration was determined using the BioRad (BioRad, Hercules CA) protein dye reagent following manufacturer’s instructions. Northern Hybridization Analysis Northern analysis was performed as described in Current Protocols in Molecular Biology (Ausubel et al., 2003) with 15 pg of total RNA loaded per lane and electrophoresed in a 1% agarose gel with 0.4 M formaldehyde. The resolved RNA was transferred by capillary action to a Nytran nylon membrane (Schleicher & Schuell, Keene NH) and immobilized by UV crosslinking in a Stratalinker (Stratagene, La Jolla CA). nor-1 and ver-l DNA probes were generated by PCR using the nor-1 cDNA plasmid pQE3l (Zhou and Linz, 1999) and the cosmid norA (Trail et al., 1995b) respectively. The primer pairs for each reaction were: nor-1, 5'-GCGACACGAACCCAG-3', 3'-CGTC CCAAAACGACC-S'; ver-I, 5'-AGCGCGGAGCCAAAG-3', 3'-CGGGCGACATCCAC AG-5'. The PCR products were gel purified using the QiaexII Gel Extraction Kit (Qiagen, Santa Clarita CA). Approximately 120 ng of each DNA fragment was radio labeled by the random primed method (BMB, Indianapolis IN) using 50 pCi [32P]-dCTP (NEN, Boston MA). The probes were hybridized to the immobilized RNA for 16 hours 41 at 65°C. The membranes were washed twice at room temperature for 15 minutes under low stringency (2X SSC, 0.1% SDS) followed by a high stringency wash at 65°C (0.2X SSC, 0.1% SDS). Signals were detected using a Phosphorimager F X (BioRad, Hercules CA). Western Blot Analysis Western blot analyses were performed by standard procedures (Ausebel et al., 2003). Thirty pg of total protein were resolved by electrophoresis through a 12% acrylamide gel using a Miniprotean II electrophoresis cell (BioRad, Hercules CA). The proteins were electroblotted to PVDF membrane and probed with anti-Nor-l or anti-Ver— l (Liang et al., 1997) as primary antibody. Alkaline phosphatase labeled rabbit anti-IgG (Sigma, St. Louis MO) was used as a secondary antibody. Colorimetric detection of the enzyme linked secondary antibody was carried out using BCIP/NBT tablets (Amresco, Solon OH). Duplicate gels were stained with Coomassie Brilliant Blue R—250 to evaluate the composition and condition of the protein extracts and to verify equal loading. Liquid Culture GUS Assays Liquid culture GUS assays were performed with 1 mg of fresh protein extract as described by Miller (Miller et al., 2003a). GUS activities are reported as the mean of the triplicate flasks, analyzed twice with the standard deviation represented by errors bars. Protein Extraction for EMSA Cellular protein was extracted from cultures of A. parasiticus SUI using modifications of the methods of Peters and Perez-Esteban (Peters and Caddick, 1994 and 42 "_ “u. LtTIQa“ wan " w" Perez-Esteban et al., 1993). 1 liter flasks containing 10, 6mm glass beads and 500 mL medium (GMS or PMS) were inoculated with 1 X 108 spores. The cultures were incubated for 48 hours at 29°C in the dark with shaking at 150 rpm. The mycelia were filtered through miracloth (Calbiochem, La Jolla CA), washed with cold, sterile water, frozen with liquid nitrogen and stored at -80°C. Frozen mycelia were ground using mortar and pestle with liquid nitrogen and transferred to a tared 125 ml flask. 5 ml of lysis buffer (25 mM Hepes-KOH (pH 7.5), 50 mM KCI, 5 mM MgC12, 0.1 mM EDTA, 10% glycerol, 0.5 mM DTT and 1 mM PMSF) per gram of ground mycelia was added to the flask. In addition, 1 ml of protease inhibitor cocktail (Sigma, St. Louis MO - product # P8215) was added to the flask per gram of mycelia. After stirring on ice for 15 minutes, saturated ammonium sulfate was slowly added to a final concentration of 10%. The suspension was then stirred on ice for 15 minutes and then set idle for 15 minutes on ice. Cell debris was then pelleted at 100,000 x g (30 minute spin at 4°C). The volume of the supematent was then determined using a graduated cylinder and transferred to a 50 m1 flask. Then solid ammonium sulfate was added slowly over 1.5 hours to raise the concentration from 10% to 70%. The ammonium sulfate addition was conducted while stirring on ice. After all ammonium sulfate was added, the flask was incubated for 30 minutes on ice without stirring. The protein was pelleted at 10,000 X g (20 minutes at 4°C). The pellet was resuspended in dialysis buffer (15% glycerol, 15 mM Hepes-KOH [pH 7.9], 100 mM KCl, 1 mM EDTA, 2 mM DTT, 0.5 mM PMSF and protease inhibitor cocktail [1 ml per 20 grams of mycelia]) and dialyzed twice in 2 liters dialysis buffer using a 10K slide-a-lyzer (Pierce, Rockford IL). Protein concentration was determined using the BioRad (BioRad, Hercules CA) protein dye reagent following manufacturers instructions. The dialyzed solution was then aliquoted and stored at -80°C. 43 r" nor-1 and ver-I Promoter Fragments The 5' boundary of the ver-I promoter region was delineated by the poly A site of the gene immediately upstream, norA. The 3' boundary was delineated 25 basepairs downstream of the translational start site. Similarly, the 5' boundary of the nor-l promoter region was delineated by the poly A site of the gene immediately upstream - ORF 3 (Miller et al., 2003a). The 3' boundary was delineated 34 basepairs downstream for the transcriptional start site. The promoter regions were divided into three subfragments designated R (right), M (middle) and L (left). Overlapping PCR primers were designed for each fragment. The primers for each ver-I subfragment carried an additional 8 bases which included a BamHI site. The PCR fragments were cut with BamHI, gel purified (Qiagen, Santa Clarita CA), and labeled using [32P]-dCTP (NEN, Boston MA) for a fill-in reaction (Ausebel et al., 2003). The nor-1 promoter PCR subfragments were gel purified (Qiagen, Santa Clarita CA) and end-labeled with [33P]- ATP (NEN, Boston MA) using Ready-to-Go Kinase (Pharmacia, Piscataway NJ). Both nor-R and ver-R contain the TATA box, transcriptional and translational start sites as well as the putative AflR binding site TCGnnnnnCGA (Femandes et al., 1998). Electrophoretic Mobility Shift Assay (EMSA) EMSA was performed essentially as described in Current Protocols in Molecular Biology (Ausebel et al., 2003). Five percent acrylamide (80:1 acrylamidezbis- acrylamide) non-denaturing gels were used. 20 fmol of nor-1 and ver-I promoter probes were incubated for 15 minutes at 30°C with 2 pg dIdC, 7.5 pg BSA and competitor (if desired) with 32 mg protein extract (added last). The entire binding reaction volume was 44 Pr... .31 6 .,;?€( HWML. If 25 pl which consisted of 20 pl of dialysis buffer in order to keep the glycerol concentration of the binding reaction greater than 10%. ELISA Enzyme-Linked Immunosorbent Assays were conducted by the method of Pestka (Pestka et al., 1980) using polyclonal antibodies provided by Neogen Corporation (Neogen, Lansing MI). Aflatoxin concentrations are reported as the mean of the triplicate flasks, analyzed twice with the standard deviation represented by errors bars. RESULTS Generation of anti—Nor-l antibody An A. parasiticus nor-1 knock-out strain accumulated norsolorinic acid suggesting that Nor-1 catalyzes the conversion of norsolorinic acid to averantin (Chang et al., 1992). Subsequently, recombinant Nor-l was shown to be able to convert norsolorinic acid to averantin in the presence of NADPH (Zhou and Linz, 1999). Purified recombinant Nor-1:Maltose Binding Protein (Zhou and Linz, 1999) was used as antigen for antibody production. The IgG fraction was purified and used for Western blot analysis (Figure 2.1). In YES shake cultures, A. parasiticus SU-l (wild type) had two major bands at 31 and 28 kDa at both 48 and 60 hours (lanes 2 and 3, respectively). Meanwhile, the Nor-l disrupted strain A. parasiticus ANor-l did not have either band at 60 hours (lane 1). An A. parasiticus SU-l extract that had been purified using an anti- Nor-l affinity column resulted in a single 31 kDa band. The prediced size of Nor-l is also 31 kDa. While the identity of the 28 kDa protein is unknown, these observations 45 '37.?er I l 2 3 4 ill! WWW l lllllllllllllllllllll lllllll ”,l lllllllll “llllillllllllllllllllllllllll . , lllllll llllllllllllll lll lll . llllllllllllll ;. lllllllllllll llllll ll lllllllllllllll iii'llum lllllllllllllll l llll ", ,, ,x'l r "1 lllllllllllllllll .‘ ‘ — _ lllllll . ~ ~ N°r 1 ‘ l‘llll ullllllllll.H . . lllllllllllllllllllllllllll I""llllulll'lllllllll‘,lll,,t, .1. ill "1' 'llll'lll'lll'llllllll "mm“, , llivu-‘ll. ' l i 1""l‘ 'ulil-llllllllulmlih llllllll ill lllllll Figure 2.1. Western blot analysis of the native Nor—l protein from A. parasiticus SU-l. Each lane contained 10 mg protein. The primary antibody was the IgG fraction of the antiserum raised against the Nor-lc/MBP firsion protein (10 mg/ml). Lanes: 1, total crude extract (10,000 X g supematent) from the nor-l disrupted strain A. parasiticus ANor—l cultured in YES liquid medium for 60 h; 2 and 3, total crude extract from the wild type A. parasiticus SU-l cultured in YES liquid medium for 48 h and 60 h respectively; 4, the native Nor-1 protein (31 kDa) purified with anti-Nor-lc/MBP fusion protein PAb affinity column from the total crude extract of A . parasiticus SU-l cultured in YES medium for 60 h. 46 suggest that the 31 kDa protein is the native Nor-1 protein encoded by the nor-l gene. In addition, the Nor-1 antibody is specific for Western blot analysis for native Nor-1 protein. Carbon, nitrogen and zinc affect aflatoxin production and aflatoxin gene expression The effects of zinc, carbon and nitrogen source on aflatoxin synthesis have been previously reported (reviewed in Luchese and Harrigan, 1993). To verify previously observed effects of these specific nutrients, aflatoxin synthesis was measured in the nor- ] ::GUS reporter strain D8D3 (Figure 2.2A) and the ver-I ::GUS reporter strain 14 (Figure 2.28) separately, in each of the following media: PMS (D8D3 only), NMS, GMSO, and GMS. Consistent with previous studies, D8D3 batch cultures in PMS did not produce detectable amounts of aflatoxin at any time point tested. Aflatoxin levels in NMS batch cultures were either non-detectable (D8D3 all three time points and 14 at 48 hours) or detectable but at greatly reduced concentrations compared to GMS shake cultures. In GMS0 batch cultures of D8D3 and 14, a low constant level of aflatoxin was present at all three time points tested. To examine the nutrient effects on accumulation of aflatoxin biosynthesis transcripts and proteins in D8D3 and 14, Northern and Western analysis were performed. Consistent with the data on aflatoxin synthesis, native Nor-1 protein and nor-I transcript accumulated in GMS cultures at all three time points for both D8D3 (Figure 2.3A,B) and 14 (data not shown). Native Nor-1 protein and nor-1 transcript accumulation in PMS cultures was undetectable for D8D3 (Figure 2.3A,B). With D8D3 and I4, NMS cultures did not contain detectable amounts of nor-l transcript or Nor-1 protein except for a light signal for nor-1 transcript at 72 hours with D8D3 in NMS (data not shown). Native 47 "‘3, t‘. r '56 r)! '.,.. 500 900 —----—- , ‘ L_; GMSO 800 _ 1:1 GMS0 l m GMS pm GMS ' l 400 ‘ \\\\r NMS 7 700 ‘ kw NMS i r 1 2529529 PMS 3 ¢ 7 o, 600 ¢ , E 300 / T E 7 ' E -‘ E 500 . ' l f-i l '5 400 ' g 200 -- . g . / i g; ,y if 300 ‘ I J :é I ' 100 ~- l F? / 200 I , ¢ 7 . % E? 100 4 L7 ,7: . / O ' ré: ND git—3'0 ND . o . . k/ND ' [.m‘L' _ 36 48 72 48 60 72 Time (Hours) Time (Hours) Figure 2.2. Aflatoxin production by the reporter strains A. parasiticus D8D3 (A) and 14 (B) grown in GMS,), GMS, NMS and PMS (D8D3 only) to 36, 48 and 72 hours (D8D3) or 48, 60 and 72 hours (14). Aflatoxin concentration of the growth media was determined by competitive-direct ELISA using polyclonal antibodies. Aflatoxin concentrations are reported as the average of the triplicate flasks, analyzed twice, with the standard deviation represented by errors bars. Non-detectable activity is represented by ND. 48 a7§( film“ 1 A" Figure 2.3. nor-l transcript and Nor-l protein accumulation and q/IR transcript accumulation by the reporter strain A. parasiticus D8D3. (A) nor-1 transcript accumulation assessed by Northern hybridization analysis. For the time points (36, 48 and 72 h), each replicate flask is presented (A, B and C). Top panel is RNA from GMS cultures and bottom panel is RNA from PMS cultures. The lane designated “+” has RNA from a 48 h A. parasiticus SUl culture in YES medium. Equal loading of RNA is demonstrated by ethidium bromide staining of RNA as shown in the panels marked “EtBr”. (B) Nor-l protein accumulation by Western blot analysis. One flask per time point (36, 48 and 72 h) was analyzed from PMS and GMS cultures. (C) aflR transcript accumulation assessed by Northern hybridization analysis. For the time points (36, 48 and 72 h), each replicate flask is presented (A, B and C). Top panel is RNA from GMS cultures and bottom panel is RNA from PMS cultures. The lane designated “+” has RNA from a 48 h A. parasiticus SUI culture in YES medium. Equal loading of RNA can be determined from part A in the panels marked “EtBr”. 49 36 48 72 ,ABCABCABC + GMS m IN llxi' 1133 H t 5.55 W 13931 Elli- - '1 , L. lwul‘p !.r.rn.rIMR.-&J._‘.»ru.- -v n v‘ nun: !. . . , 7, -_. ._. - w- v 5. ~~-;——i- -.. 4.": in ‘wvn—zr I . ...u-~‘ 4... --vv- ' PMS W '.~-' 1'7 - ., " .. :-N "-.f’ “ Wow-m", Mr" 1: r-t . PMS GMS 36 48 72 36 48 72 36 48 72 ABCABCABC + GMS ""“P in” I If”. ‘5'“ . ' “3'? r' if ,1 , . M tftu. 'LX't‘ PMS ' I 3 ' ‘ 1M." WIIJ Figure 2.3 50 m1 , (1756‘ u“: 'n' ;afi-‘L'Lm 5‘ Nor-1 protein and nor-l transcript accumulated at the 36 hour time point for GMS0 but were non-detectable at 48 and 72 hours with D8D3 and 14 (data not shown). Analysis of total protein from GMS0 on SDS-PAGE revealed that proteins at 48 and 72 hours displayed extensive proteolysis which was much less severe at 24 hours (data not shown). Native Ver-l protein and ver-I transcript levels in 14 followed a very similar pattern as native Nor-l protein and nor-I transcript accumulation in D8D3 (data not shown). AflR is a transcription factor that is required for aflatoxin biosynthesis gene transcription. To examine the nutrient effects on accumulation of aflR transcripts in D8D3 and 14, Northern hybridization analyses were performed (Figure 2.3C). aflR transcript accumulated to greater levels at all three time points for D8D3 and 14‘ (data not shown) in GMS cultures compared to PMS cultures (Figure 2.3C). aflR transcript accumulation in NMS and GMSO in D8D3 and 14 was non-detectable except for the first time point with GMSO. Transcript steady states of the aflatoxin transcription factor qflR coincides nor-l and ver-I transcript and protein accumulation. Nutrient effects are in part regulated at level of transcription The previous results suggested that rates of transcript accumulation in part mediate changes in aflatoxin synthesis, but do not clarify if accumulation occurs via changes in transcription rate or transcript stability. Reporter assays with D8D3 (data not shown) and I4 (Figure 2.4) were used to analyze if zinc concentration, carbon source and nitrogen source affect transcriptional regulation of the aflatoxin genes nor-1 and ver-l . GUS activity of 14 GMS cultures was 9 fold higher than 14 GMS0 cultures at 72 h (Figure 2.4). I4 GMS cultures had 60 fold higher GUS activity than 14 NMS cultures (Figure 2.4). Results with D8D3 activities with GMS, GMSO and NMS were similar to 14 (data 51 '7'? 07 “1750' ’5: 700 g [:1 GMS .E 600 _— GMS-0 g NMS i D E 500 a .g, E 400 - «S E 300 ~ 3‘ ‘6 < 100 a a) _ 48 6O 72 Time (Hours) Figure 2.4. B-glucuronidase (GUS) expression by the reporter strains A. parasiticus I4. Cultures were grown in GMS, GMS0 (GMS - O) and NMS for 48, 60 and 72 h. 52 not shown). In addition, the GUS activity of D8D3 PMS cultures was non-detectable (data not shown). While we cannot eliminate transcript stability as a factor, transcriptional activation of nor-1 and ver-I is at least partially responsible for the differences in aflatoxin production under the conditions tested. Nutrients affect DNA promoter binding by cellular proteins In order to determine if DNA binding proteins mediate nor-1 promoter activity, electrophoretic mobility shift assays (EMSA) were performed with A. parasiticus SUI and AFS l 0 protein extracts. The DNA probes were generated from the promoter region of nor-1 (Figure 2.5). To test for specificity, unlabeled Oligonucleotides were used as competitors (250 fold molar excess) for protein binding. The promoter fragment nor-R, which contains an AfiR binding site, produced two shifted complexes with both A. parasiticus SUl (wild type) and A. parasiticus AFSlO (AflR knock-out) extracts from PMS cultures. Of these two shifted complexes, the faster migrating complex appears to be specific. The promoter fragment nor-R* is the same as nor-R except the AflR binding site was altered (TCchcagCGA to AGTttaaaCAG). Since nor-R and nor-R* are both effective competitors for the specific complex, the AflR cis-acting site is not responsible for the shifted complex. In addition, since the specific shifted complex appears in AFSlO extracts, the protein responsible for the shifted complex appears to be AflR independent. The nor-R oligonucleotide produced three shifted complexes with AFSIO extracts from GMS medium (Figure 2.6). nor—R and nor-R* did not compete for any of these three shifted complexes. Consequently, all three complexes are non—specific. The nor-R oligonucleotide produced two shifted complexes with SUI extracts from GMS medium. Of the two shifted complexes, the intensity of the faster migrating complex was reduced 53 93:: pi] r @700 nor-R nor-R* -l 17 AflR -52 nor-R1 nor-R2 -I 17 AflR . ‘ +55 nor-TATA* ' Figure 2.5. Map of EMSA Oligonucleotides used for electrophoretic mobility shift assay (EMSA). Three putative cis-acting sites are boxed, AflR TATA and CRE. For nor-R*, the AflR site has been mutated (TCchcagCGA to AGTttaaaCAG). For nor-TATA*, the TATA box was mutated (5'-ATATATAG-3' to 5'-GTTTAAAC-3'). 54 J at no; 35m f} strain - S S S A A A S S S A A A medium - P P P P P P G G G G G G competitor - - R* - R R* - R R* — R R* Figure 2.6. EMSA of nor—R. Protein extracts were collected from two different A. parasiticus strains, SUI (S) and AFSlO (A). Each strain was grown in two different media, PMS (P) and GMS (G). 32 mg of protein was used per lane. The two competitors (250 fold excess) used were nor-R (R) and nor-R* (R*). 55 in the presence of both nor-R and nor-R* as competitors. Since nor-R and nor-R* are both effective competitors for the specific complex, the AflR cis-acting site is not responsible for the shifted complex. The protein responsible for the specific shifted complex with SUI GMS extracts is dependent on functional AflR for activity because the AFS IO (AflR knock-out strain) extract lacked this shifted complex. In addition, the mobility of the specific shifted complex appears different than the specific shifted complex seen with the PMS extracts (both SUI and AFSIO) suggesting that different proteins may be responsible for the specific complexes. In order to localize the specific DNA binding site in the SUI GMS extracts, additional competition experiments were performed (Figure 2.7) with various I oligonucleotide competitors (Figure 2.5). For example, nor-R was divided into two overlapping Oligonucleotides, nor-R1 and nor-R2. Since nor-R2 was an effective competitor while nor-R1 was not, additional competitors were designed to test potential sites in nor-R2. nor-TATA* is the same as nor—R except the TATA box has been substituted (5'-ATATATAG-3’ to 5'-GTTTAAAC-3'). nor-TATA* is an effective competitor suggesting that the TATA box is not responsible for the shifted complex. Lastly, an oligonucleotide that overlaps the translational start point, CRE, was an effective competitor. The specific complex seen with A. parasiticus SUI protein from GMS cultures bound to the C REl region. The CREI binding protein is likely a nor-1 transcripitonal activator. 56 I. f) “:31 fit I strain - S S S S I I medium - G G G G GI I competitor - - R RT* R1 R2 CREI It" in 4w“ ”13‘qu klilyll I! I Figure 2.7. nor—R competition EMSA. Protein extracts were collected from A. parasiticus SUI (S) grown in GMS (G). 32 mg of protein was used per lane. The competitors used were: nor—R (R), nor-TATA* (RT*), nor-R1 (R1), nor-R2 (R2) and CRE. 57 DISCUSSION The effects of environmental and nutritional factors such as temperature, pH, carbon source, nitrogen source and zinc concentration on aflatoxin production have been previously studied (reviewed in Luchese and Harrigan, 1993). However, nutritional factors such as carbon source, nitrogen source and zinc concentration provide useful tools in studying the molecular mechanisms of aflatoxin biosynthesis gene regulation. In addition, the creation of a nor-1 ::GUS (D8D3) strain, a ver-I ::GUS strain (14) and an q/IR knock-out strain (AF S 10) permitted the study of these nutritional factors and aflatoxin gene transcriptional regulation. We presented a comprehensive analysis of the use of nutritional components as tools to study aflatoxin gene transcriptional regulation. NMS and PMS cultures produced low or non-detectable levels of nor-I and ver-I transcript, Nor-1 and Ver-l protein and aflatoxin. However, GMS cultures produced significant levels of nor-1 and ver—I transcript, Nor-I and Ver-l protein and aflatoxin. GUS reporter activities confirmed that the increase in nor-1 and ver-I transcript steady states in GMS was due, at least in part, to transcriptional activation. However, we can not rule out transcript stability because the nor-I ::GUS and ver-I ::GUS transcripts may also be subject to the same transcipt stability process. aflR transcript steady state was also higher in GMS cultures. Insertion of an additional copy of aflR results in the upregulation of aflatoxin biosynthesis (Chang et al., 1995). In addition, aflR transcript and AflR protein accumulation in various nutritional and environmental conditions have been shown to be consistant with aflatoxin production, in that culture conditions with higher aflR steady states also have higher concentrations of aflatoxin (Liu et al., 1999). GMS (glucose, ammonium and IOmM zinc) induces aflatoxin gene transcription, 58 “1,1321 .0 “14‘ possibly via the upregulation of aflR. GMS0 cultures appear similar to GMS cultures at the earliest time point. However, at the later time points, the amount of aflatoxin in the culture medium, nor-1 and ver-I transcript and protein steady states and nor-1 and ver-I transcriptional activation either stayed the same or decreased. Analysis of the protein extracts from the GMS0 cultures on SDS-PAGE gels revealed that the protein extract was subject to extensive proteolysis. Repeated attempts to extract GMS0 cultures revealed the same proteolysis. The omission of zinc from the culture medium not only results in a decrease in aflatoxin production but also in growth (Bennett et al., 1979). Consequently, we hypothesize that once the growing culture runs out of zinc, the fungal cells lyse‘ and the proteins degrade. If true, the lack of transcriptional activation in GMSO cultures is not due to a specific repression of aflatoxin biosynthesis. Since GMS cultures induce nor-1 transcriptional activation, protein extracts from GMS cultures are a logical place to look for transcriptional activators specific for aflatoxin. Using a 160 base pair nor-1 promoter fragment as probe (nor-R), electrophoretic mobility shift assays (EMSA) with A. parasiticus SUI (wild type) GMS extracts produced a single specific shifted complex. Using various competitors for the specific nor-R shifted complex allowed us to localize the protein binding site to a region that we designated CREI. The protein that binds to CREI appears to be dependent on AflR because the A. parasiticus AFSlO (aflR knock-out) lacks this shifted complex. A limitation with EMSA is that it is an in vitro assay. However, preliminary experiments indicate that the CREl site has functional significance in vivo as well (Dr. Ludmila Roze, personal communication). We did not detect AflR binding to nor-R consistent with previous studies with recombinant AflR (Ehrlich et al., 1999b). However, this AflR 59 PM, (2;) a ( binding site was shown to be necessary for nor-1 transcriptional activation (Miller et al., 2003a). It is unknown why our A. parasiticus SUI (wild type) protein extract lacks AflR activity. AflR is necessary for nor-1 transcriptional activation (Miller et al., 2003a). In addition, forced expression of AflR using an inducible vector results in aflatoxin production even in non-aflatoxin inducing conditions (Flaherty and Payne, 1997). Nutrients such as nitrogen source may affect aflatoxin production by modulating aflR expression. AreA, a global nitrogen regulatory protein, has been shown to bind to the q/IR promoter (Chang et al., 2000). The number of AreA binding sites in the aflR/aflJ intergenic region may reflect differences between different strains and the amount of aflatoxin they make in response to different nitrogen sources (Ehrlich and Cotty, 2002). Additional experiments are needed to elucidate the mechanisms for transcriptional activation of aflatoxin biosynthetic genes in response to nitrogen and carbon sources. ACKNOWLEDGMENTS Dr. Renqing Zhou generated the anti-Nor-I polyclonal antibody and provided Figure 2.1. All other experiments described in Chapter 2 were performed by Matt Rarick and Michael Miller. Chapter 2 will be submitted for publication in the near future. 60 .dJL‘ . 1 . ‘3‘.W l -2 CHAPTER 3 Chromosomal Location Plays a Role in Regulation of Aflatoxin Gene Expression in Aspergillus parasiticus INTRODUCTION Aflatoxins are highly toxic secondary metabolites produced predominantly by the fungi Aspergillus/laws, A. parasiticus and A. nomius (Council for Agricultural Science and Technology, 2003). These toxins frequently contaminate food and feed crops, including peanuts, corn, cottonseed, and tree nuts, and generate a large health and economic impact in the United States and in other countries (reviewed in Council for Agricultural Science and Technology, 2003). To help eliminate aflatoxin contamination of food and feed, we have studied the mechanisms that regulate aflatoxin gene expression. This knowledge will be used to design novel, effective control strategies to reduce aflatoxin contamination of crops in the field and during storage. Aflatoxin biosynthesis is a complex process that requires at least 16 different enzymatic steps (Bhatnagar et al., 1994) encoded by up to 25 individual genes (Keller and Adams, 1995). These genes are clustered in A. flavus, A. parasiticus and A. nidulans (A. nidulans synthesizes the penultimate pathway intermediate, sterigmatocystin). It was previously hypothesized that clustering of aflatoxin genes may allow coregulation in response to environmental cues, although no conclusive data were reported (Trail et al., 1995b). Preliminary evidence for a role for clustering in aflatoxin gene regulation was 61 ”H L1,) at reported by Liang et al. (Liang et al., 1997). The promoter of the aflatoxin biosynthesis structural gene ver—I was fused to uidA (encodes D-glucuronidase [GUS]) (Jefferson, 1989) to generate a reporter plasmid, pHD6.6, containing niaD (encodes nitrate reductase) as a selectable marker. pHD6.6 was transformed into A. parasiticus NR1 (niaD), resulting in integration predominantly at the ver-I or niaD locus via homologous recombination. Single-copy integration of pHD6.6 at niaD resulted in a 500-fold reduction in ver-l promoter activity when compared with single-c0py integration at the ver-I locus; however, the temporal pattern of expression appeared to be similar at both loci. One explanation for reduced ver-I promoter function at the niaD locus is that the location in the aflatoxin cluster results in positive, position-dependent regulation of ver-I exression. An alternative hypothesis is that the expression of ver-l integrated at niaD is negatively influenced by niaD regulation. In the absence of preferred nitrogen sources and in the presence of nitrate, niaD is expressed. Under the rich growth conditions tested by Liang et al. (Liang et al., 1997), m'aD may have been repressed, and therefore the lack of ver-l expression at the niaD site could have resulted from niaD-dependent regulation. Subsequently, the promoter of the aflatoxin biosynthesis gene nor-l was fused to GUS to generate a reporter plasmid, pAPGUSNN-B, containing niaD as a selectable marker (Chiou et al., 2002). Transformants with pAPGUSNN-B integrated at the niaD locus had no detectable GUS activity while nor-1 integrants had GUS activity (Chiou et al., 2002). In addition, GUS transcript and protein accumulation correlated with nor-1 transcript and protein accumulation in terms of timing and amount (C hiou et al., 2002). Because niaD dependent regulation could account for the absence of expression at niaD for both nor-1::GUS (Chiou et al., 2002) and ver-121GUS (Liang et al., 1997), a third chromosomal location was analyzed using pAPGUSNP, which contained nor-1::GUS 62 plus pyrG (encodes OMP decarboxylase) as a selectable marker. GUS expression was detectable only when pAPGUSNP integrated at nor-1 and was not detectable at pyrG, even under growth conditions that required pyrG expression. Our hypothesis was that the aflatoxin biosynthesis genes are subject to gene- cluster-dependent regulation. To test our hypothesis, we wanted (i) to generate a new nor—I ::GUS reporter construct (pNANG-3) that enables easy promoter replacement and (ii) to validate the positional-dependent regulation by correlating the site of integration with solid culture GUS activity for pBNG3.0 transformants. We generated a series of plasmids that enable a nor-1 promoter to be easily amplified by PCR and ligated into a nor-1::GUS reporter plasmid containing niaD as a selectable marker (pNANG-3). Using a solid culture GUS assay, we screened pBNG3.0 (3.0 kb nor-1 promoter fragment ligated into pNANG-3) transformants for GUS activity. GUS+ transformants were tested by a rapid PCR site of integration assay which identified all 21 GUS+ transformants as either 5' nor-I (12) or 3' nor-1 (9) integrants. Conversely, all 22 GUS- transformants were negative for 5' nor-] and 3' nor-1 integration. Southern hybridization analysis was used to confirm 3' nor-I integration of all 8 preliminary 3' nor-1 integrants tested. These results confirm the utility of the nor-1::GUS reporter plasmid for analyzing nor-1 promoter function and the positional-dependent regulation previously reported (Liang et al., 1997 and Chiou et al., 2002). MATERIALS AND METHODS Strains and growth conditions Escherichia coli DHSOL F’c [F ’ endAI 115de 7 (rk- mk-) supE44 thi-I recA gvrA 63 L (NaI') AreIAI (lacZ YA argF)u,69:(m80 AlacZ M15)] (Invitrogen, Carlsbad, CA) was used to amplify plasmid DNA using standard procedures (Ausubel et al., 2003). Aspergillus parasiticus NR1 (niaD) was used as the recipient strain for all fungal transformations (Horng et al., 1990). To measure solid culture GUS activity, YES agar (2% yeast extract, 6% sucrose; pH=5.8) plates were used. Plasmid Constructs (i) pNele (Figure 3.18). pNEBl93 (Figure 3.1A - New England Biolabs, Beverly, MA) was digested with BamHI and the protruding ends were filled in by Klenow fragment. A Not] linker (5'-AGCGGCCQCT-3') was then ligated onto the blunt ends. The DNA was then digested with Not], agarose gel purified using QIEX II (Qiagen, Valencia, CA) and then re-ligated. Bacterial transformants were screened by digesting mini-prep DNA with Not]. The resulting plasmid (pNele) no longer has a BamHI site and can no longer be used for blue/white screening. (ii) pNiaD-Al (Figure 3.1C). A 7.4 kb XhoI/SaII fragment from pSL82 (Horng et al., 1990) was agarose gel purified using QIEX Il (Qiagen, Valencia, CA) and then ligated into the Sal] site of pNele. Bacterial transformants were screened using the colony hybridization technique (Ausubel et al., 2003). The XhoI/Sal] 7.4 kb fragment was radioactively labeled using the random primed labeling kit (Roche Applied Science, Indianapolis, IN) and used as probe. Only 6 of 600 clones were positive in the colony hybridization. The positive clones were further analyzed by digesting plasmid DNA with Psi]. Plasmids in the “A” orientation result in 4.2 and 5.9 kb DNA fragments while plasmids in the “B” orientation result in 3.2 and 6.9 kb fragments. Functionality of the 64 Figure 3.1. Restriction endonuclease maps of relevant plasmids. (A) pNEBl93 (New England Biolabs, Beverly, MA). This plasmid is a modification oprCl9 that incorporates additional restriction endonuclease sites in the multiple cloning sites. These additional restriction enzymes recognize 8 bp sequences and consequently are less likely to exist in the DNA being cloned. (B’) pNebN 1. This plasmid has a Not] site that replaces a BamHI site in pNEBl93 by insertion ofa .N’otl linker. (C) pNiaD-Al. A 7.4 kb X/mI/Sall fragment from pSL82 (Homg et al., 1990) that contains the niaD selectable marker was cloned into the Sal] site oprebN l. (D) pNANG-3. In addition to carrying the niaD selectable marker. pNANG-3 carries a small part of the nor-I coding sequence (10 amino acids) fused to the D-glucuronidase (m’dA or GUS) coding sequence which is in turn fused to the 2 kb nor-1 3' temiinator fragment. The small numbers of codons that were changed in the nor-l coding sequence were all acceptable based on codon usage and maintained the correct reading frame. The 4 kb GUS/nor-l terminator fragment was amplified by PCR using pAPGUSNN—B (Figure 3. 1 F) as template with primers that had Not] (5') and Ascl (3') tails. Appropriate promoter pieces, amplified by PCR using primers with N011 (and Pacl if directionally cloned) can be cloned into pNANG-3. (E) pBNG3.0-3 F. This plasmid contains a 3 kb PC R-amplified nor-1 promoter piece cloned into the Not] site of pNANG—3. (F) pAPGUSNN-B. Original nor-[::GUS reporter plasmid constructed by Wilson (Chiou et al., 2002). 65 pNEBl93 2700 bp AscI 418 Not] 426 PacI 432 Pstl 7860 SalI 784' pNiaD—Al 10100 bp Pstl 3648 E Scall6400 Pstl 14612 Sall 14600 Ascl 418 C131 1218 C131 2118 amp 3' nor-1 pBNG3.0 GUS 16900 bp Not14226 ScaI 4326 Scal 5126 niaD 5 nor-l Pstl 6226 Pstl 6426 Notl 7226 Pstl 10400 Figure 3.1 66 AscI418 D Ascl 418 amp \ ' nor-1 pNANG-3 13900 bp Pstl 11612 Sal] 11600 GUS Notl 4226 Pacl 4236 Pstl 7400 F Pstl 1000 Sea] 2100 Sea] 2900 ' 3 00 Pstl 13800 “mu" 0 pAPGUSNNB 17000 bp 3’ nor-l EeoRI 4800 Clal 5000 Amp C121] 5930 SC81 8300 niaD gene was tested by performing a fungal transformation with pNiaD-Al, pNiaD-BZ and the positive control pSL82 (Homg et al., 1990). The transformation frequency for pNiaD-Al, pNiaD-BZ and pSL82 was 17 colonies/pg, 19.2 colonies/pg and 17.6 colonies/pg respectively. (iii.) pNANG—3 (Figure 3. l D). To insert the uidA gene (GUS), the reporter construct pAPGUSNN-B (Chiou et al., 2002) was utilized as a template for PCR. Primers were designed to amplify a 4 kb fragment that contained the nor-I terminator region, GUS coding sequence and the nor-1 coding sequence upstream of the fusion point. The primers contained restriction sites (underlined residues) to facilitate cloning: the 5' primer (5'-TAGCGGCCGCGATCAAGAGAAGCTCTATACG-3') contained Not] and the 3' primer (5'-TTGGCGCGCCCTCGATGATGATGCTCTG-3') contained Ase]. Since the Not] site was located within the coding sequence, care was taken to maintain the correct reading frame. The 4 kb Natl/Ase] PCR product was ligated into the Not] and AscI sites of pNiaD-Al. Bacterial transformants were screened by colony hybridization with the Natl/Ase] PCR product as probe (same procedure as with pNiaD-Al). More than 90% of transformants were shown to carry the insert. Selected positives were then screened by restriction enzyme analysis with Pstl (expected sizes: 4.2 and 9.7 kb). Because PCR mediated mutations in the Natl/Ase] fragment may have occured, four different positive clones (pNANG-1, -2, -3 and -4) were saved. (iv.) pBNG3.0-3F (Figure 3. l E). A 3 kb nor-I promoter piece was amplified by PCR with pAPGUSNN-B as template and appropriate primers with Not] tails (5'- TCGCGGCCGCTAAGTGATCCATTCATTATGTC-3' and 5'-TTGCGGCCGCTCCTT GTCTCTGTACTGATAAA-3'). In this way, the nor-1 promoter could be easily removed and replaced with other putative promoters to test for function. Both pNANG 67 n .2 :fig‘bl'fi'r.‘ (pNANG-1, -2, -3 and -4) and the PCR fragment were digested with Not] and then gel purified using QIEXII (Qiagen, Carlsbad, CA). The promoter insert was then ligated into all four vectors. Bacterial transformants were screened by colony hybridization (see pNiaD-Al) with the nor-1 promoter PCR fragment used as probe. Orientation of the promoter insert was determined by digesting plasmid DNA with Pstl with the'proper orientation resulting in 8.6, 4.2, 4.0 and 0.2 kb fragments. The nucleotide sequence from pBNG3.0-1, -2, -3 and -4 was analyzed from the nor-1::GUS fusion site to 322 bp upstream from the nor-1 transcriptional start site and confirmed the correct nucleotide sequence in this reporter construct. pBNG3.0-3F was used for fungal transformations. (v.) pAPGUSNN-B (Figure 3.1F). The pAPGUSNN-B (Chiou et al., 2002) nor- ] ::GUS fusion construct was derived from pAPGUSN (Chiou et al., 2002) by insertion of a 7.4 kb SalI/XhOI fragment carrying niaD from pSL82 (Horng et al., 1990). The nor-l promoter region consists of a 3 kb HindIII DNA restriction fragment carrying flanking sequences, promoter, and the first 21 amino acid residues of Nor-l fused in frame to uidA from E. 0011' (Jefferson 1989). The nor-l terminator is a 1.8 kb EcoRI/Sall genomic DNA fragment containing the last six amino acids of Nor- l , the transcription terminator and flanking sequences. Transformation and Identification of the site of plasmid integration (i) Transformation. Transformation of A. parasiticus protoplasts was performed as described by Homg et a1. (1990). 10 pg of pBNG3.0-3F was transformed into 2.7 X 107 NR-l (niaD) protoplasts. Selection of niaD+ colonies resulted in 276 total transformants (transformation frequency = 1.0 X 10‘6 CF U/protoplast/ug plasmid). (ii) Rapid DNA extraction procedure. A. parasiticus transformants were 68 inoculated onto Czapek-Dox (Difco, Detroit, MI) agar plates and incubated for 3-4 days in the dark at 29°C. A sterile pipette tip was used to scrape some mycelia from the surface of the agar which was deposited in a 1.5 mL screw cap tube containing 100 uL TE (10 mM Tris-Cl, 1 mM EDTA, pH = 8). The sample was then boiled for 5 minutes and centrifuged for 10 minutes at 15,000 X g. The resulting supematent was used as a template for PCR. (iii) 3' PCR analysis. The primers for 3' integration were IL 102 (5'-CGCAAGG TGAGGGTTCGAACCGAGG-3') and JL 103 (5'-CCGCAGCAGGGAGGCAAACAAT GAA-3'). Each 50 uL PCR contained: 2.5 uL crude template, 1X reaction buffer for Pfu Turbo (Stratagene, La Jolla, CA), 200 11M dNTP, 1 mM MgC12, 50 pmol IL 102, 50 pmol JL103, 2.5 units Pfu Turbo (Stratagene, La Jolla, CA) and water. Reaction conditions included an initial denaturation step at 95°C for 3 minutes followed by 35 cycles; each cycle consisted of 95°C for 1 minute, 68°C for 1 minute and 72°C for 3 minutes. The reaction was terminated by incubation at 72°C for 10 minutes. Following PCR, 20 p.L of reaction mixture was loaded on a 1% agarose gel. After electrophoresis, the occurrence of a 2.1 kb DNA fragment demonstrated a positive reaction for 3' integration. (iv) 5' PCR analysis. The primers for 5' integration were I L99 (5'-TTTCACGGG TTGGGGTTTCTACAGG-3') and JLIOO (5'-GACGGGCAACCTCTTTACAAACATC- 3'). Each 50 11L PCR contained: 2.5 uL crude template, 1X reaction buffer, 200 11M dNTP, 1 mM MgC12, 50 pmol JL 99, 50 pmol JL100, 2.5 units Pfu Turbo (Stratagene, La Jolla, CA) and water. Reaction conditions included an initial denaturation step at 95°C for 3 minutes followed by 35 cycles; each cycle consisted of 95°C for 1 minute, 62°C for 1 minute and 72°C for 5 minutes. The reaction was terminated by incubation at 72°C for 69 10 minutes. Following PCR, 20 1.1L of reaction mixture was loaded on a 1% agarose gel. A 3.1 kb DNA fragment demonstrated a positive reaction for 5' integration. (v) Southern hybridization analysis. Genomic DNA was purified from A. parasiticus cultures shaken for 48 h in 100 ml YES (2% yeast extract, 6% sucrose and pH=5 .8) liquid medium at 29°C with five, 6 mm glass beads (Skory et al., 1993). The DNA was electrophoresed under standard conditions (Ausubel etal., 2003). Southern hybridization analysis was performed according to standard procedures (Ausubel etal., 2003). 2.5 u g of genomic DNA was digested with Seal and probed with a 900 bp C la] fragment isolated from the nor-1 terminator region of pAPGUSNN-B (Chiou etal., 2002). Digestion of DNA from the recipient strain NR-l generates a 3.0 kb DNA restriction fragment while a 3' nor-1 integrant results in 3.7 and 4.0 kb DNA restriction fragments. Reporter Assays (i) Solid culture GUS Assay. Autoclaved Nytran SPC (Schleicher & Schuell, Keene, NH) membranes were first placed on top of YES (2% yeast extract, 6% sucrose, 1.5% agar and pH=5.8) agar plates. Transformants were then transferred via sterile toothpick to the Nytran membrane. After incubation for 46 h at 29°C in the dark, the Nytran filters were removed, frozen in liquid nitrogen and thawed at room temperature (2 repetitions) and then incubated for 2 h with GUS substrate solution that includes 0.04% of the colorimetric GUS substrate (X-Glu: 5-bromo-4-chloro-3-indoly1 B-D-glucuronide) in GUS reaction buffer (60 mM NaZHPO4, 40 mM NaHzPO4, 10 mM KCl, 1 mM MgSO,, 0.07% B—mercaptoethanol) and visually evaluated. 70 TH C ya RESULTS Generation of transformants Previously, nor-l ::GUS reporter strains generated using pAPGUSNN-B (Figure 3. 1 F) demonstrated the usefulness and validity of the GUS reporter system (Chiou et al., 2002). A drawback with pAPGUSNN-B was the inability to replace the 3.0 kb nor-1 promoter with a different nor-l promoter. Consequently, a new nor-1 ::GUS reporter plasmid was designed that utilized unique restriction enzyme sites to facilitate the independent cloning of a nor-1 promoter. A 3.0 kb nor-1 promoter fragment was amplified with PCR using primers that had N011 tails. The promoter fragment was then cloned into the Not] site in pNANG-3 (Figure 3.1D) resulting in pBNG3.0-3F (Figure 3. l E). Using a similar cloning strategy, several different nor—1 promoter fragments have been fused to GUS and been analyzed (Miller eta1., 2003a and Miller et al., 2003b). Figure 3.2 outlines the procedure used to generate, screen and test pBNG3.0-3F transformants. Ten ug of pBNG3.0-3F was transformed into 2.7 X 107 NR-l (niaD) protoplasts. Selection of m'aD+ colonies resulted in 276 total transformants (transformation frequency = 1.0 X 10" CFU/protoplast/ug plasmid). The transformation frequency was similar to previously reported frequencies with the related pAPGUSNN-B plasmid (Chiou et al., 2002). Phenotype Screening of Transformants Colonies grown for 42-48 h on YES agar were used for solid culture GUS analysis. GUS positive fungal colonies convert the GUS substrate X-glu (5-bromo—4- chloro-3-indolyl B-D-glucuronide) to a blue product. After 1-2 h, the blue color is 71 Fungal Transformation pBNG3.0-3F tong DNA with 27000000 protoplasts 127 total transformants I Solid Culture GUS Assay p I ' 1 ” i I GUS + GUS - 29 (24.8%) ' 98 (752%) PC R Integration Assay ——-> I 1 fl i ' ' l 9121 - 3' integration 12/21 - 5‘ integration 0/22 - 5' integration 0/22 - 3' integration Southem Blot Analysis —-> I I I I . I I I I _ IIIIII 8/8 - confirmed 3' integration ; 5/5 confirmed not 3‘ integration 5/5 - confirmed not 3’ integration 5/5 conSIstant With 5' integration 2/5 - confirmed niaD gene replacement 3/5 — consustant WIII'I niaD integration Figure 3.2. Experimental design. pBNG3.0-3F fungal transformants were first tested for GUS activity using a solid culture GUS assay. Selected GUS+ and GUS- transformants were then tested using a 3' and 5' nor-1 site of integration PCR assay. Lastly, 3' nor-l integration status was confirmed for selected transformants using southern hybridization analysis 72 clearly evident for GUS positive colonies (Figure 3.3). 1 17 niaD+ transformants were initially tested for GUS activity by the solid culture assay. Of the 117 transformants tested, 29 were GUS positive colonies (24.8 %). In Figure 3.3, transformant 5 was GUS positive while transformants l, 2, 3, 4 and 6 were GUS negative. Genotype Screening of Transformants In order to determine if the site of integration of pBNG3.0-3F is a factor in GUS activity, the chromosomal location of nor-I ::GUS in 21 GUS positive and 22 GUS negative transformants was investigated. Each nor-1 ::GUS reporter construct could theoretically integrate into the A. parasiticus chromosome by homologous recombination at three independent sites: niaD, 5' nor-1 (nor-1 promoter) and 3' nor-1 (nor-I terminator). Screening for clones in which integration occurred at the 3' nor-I locus is essential for two reasons: 1) niaD integrants have been shown to have lower transcriptional activity than aflatoxin cluster integrants (Liang et al., 1997 and Chiou et al., 2002); and 2) 5' integrants result in the chromosomal nor-I promoter fused to the GUS gene and the plasmid nor-1 promoter fused to the chromosomal nor-1 gene. A rapid site of integration PCR assay (Chiou et al., 2002) was initially used to screen these transformants. With the 3' nor-I PCR assay, a 2.1 kb fragment is diagnostic for 3' nor-1 integration while a 3.1 kb fragment is diagnostic for 5' nor-I integration with the 5' nor-1 PCR assay. All 21 GUS positive transformants were positive for 5' (12) or 3' (9) nor-1 integration with the PCR assay (Figure 3.4). Conversely, 22 randomly selected GUS minus transformants were tested for site of integration using the rapid PCR assay and were all negative for 3' (Figure 3.5A) and 5' nor-I (Figure 3.5B) integration. 73 ....:..13~a,... 11111111 I1 1“ iIIIIII111 11 1116 111111 . 11111111I11I1'1I'11ii1 iIIIIIIIIII III II ‘ .1i III ‘ 111111 ‘11111' ’11 ‘IIII III, I . [.3 ‘ “ 111.1 I 5Iii111IIumIiIiIIIIiI . . 3 III IIIIII ' 1111111111 1111111,: I ; 1'1 .III IiiiIiIIIIIIIIIIII‘. “ 11111 . 1:" I III“ Figure 3.3. Solid culture GUS assay. Screening of transformants was performed using a solid culture GUS assay. Transformants 1, 2, 3, 4 and 6 are GUS— while transformant 5 is GUS+. 74 A Seal 16400 Asc1418 Clal 1218 Pstl 14612 C1a12118 pBNG3.0 16900 bp NotI 4226 Scal 4326 Seal 5126 Pstl 6226 Pstl 10400 Pstl 6426 B Notl 7226 C ll.\l345(i78.\191011 llkl}45678.\1‘)111|| 31kt)“ Figure 3.4. PC R analysis of site of integration in GUS+ transformants. (A) Restriction endonuclease map of pBNG3.0-3F. Only restriction endonuclease sites relevant to this study are shown. Ten GUS+ (lanes 1 to 6 and 8 to I 1) and one GUS- transformant (lane 7) were analyzed for both 5' and 3' nor-1 integration. Template DNA was prepared using a rapid boiling procedure. (B) Schematic for 5' nor-1 integration with PCR data. 5' nor-I integrants resulted in a 3.1 kb PCR fragment (lanes 1, 2, 5, 8, 11). (C) Schematic for 3' nor-1 integration with PCR data. 3' nor—1 integrants resulted in a 2.1 kb PCR fragment (lanes 3, 4, 6, 9, 10). Lanes labeled M in panels B and C represent DNA ladder. 75 A 3 9140M 5 932811634107M71458 753531 M2944101216996M 8 t + 3 9140M 5 932811634107M71458 753$31M2944101216996M 8 .3.1kb Figure 3.5. PCR analysis of site of integration in GUS- transformants. Template DNA was prepared using a rapid boiling procedure. (A) 3' site of integration PCR assay. A11 22 randomly chosen (lane numbers indicate transformant number) GUS- transformants lacked the 2.1 kb PCR fragment diagnostic for 3' nor-1 integration. Positive controls (lanes +) did contain the 2.1 kb PCR fragment. (B) 5' site of integration PCR assay. All 22 randomly chosen (same as in A) GUS- transformants lacked the 3.1 kb PCR fragment diagnostic for 5' nor-l integration. A positive control (lane +) did contain the 3.1 kb PCR fragment. All GUS- transformants were negative for both 3' (A) and 5' (B) nor-1 integration. Lanes labeled M in panels B and C represent molecular size markers. 76 ‘£!'*" While the PCR assay could identify 3' nor-I integrants, it could not identify multiple integrants (multiple 3' nor-I integrations and/or additional non-3' nor-1 integrations). Consequently, Southern hybridization analysis was used to confirm PCR integration assay results for selected 3' nor-I (Figure 3.6A), 5' nor-1 (Figure 3.68) and non nor-1 (Figure 3.6B) integrants. Disappearance of a 3.0 kb DNA fragment in the recipient strain, NR1, and the appearance of 3.7 and 4.0 DNA fragments is diagnostic for 3' nor-I integration. A11 PCR site of integration assay 3' nor-1 integrants tested (8) were confirmed for 3' nor-1 integration by Southern hybridization analysis (Figure 3.6A). While the Southern hybridization scheme utilized can not distinguish between 5' nor-I, niaD and heterologous integration, it can confirm that these transformants are not 3' nor- ] integrants. All GUS+, 5' nor-I integrants analyzed were confirmed not to be 3' nor-1 integrants (Figure 3.68). All GUS negative transformants (also negative for 5' and 3' nor-l integration with the PCR assay) were also confirmed not to be 3' nor-1 integrants (Figure 3.6B). In addition, two of these transformants (3 and 93) are likely products of double—crossover homologous recombination at niaD (gene replacement) because they contain only the 3.0 kb band seen in the recipient strain, NR1 (Figure 3.6B). Solid culture GUS assay with 2 3' nor-l integrants, 2 5' nor-I integrants and 2 presumed niaD integrants is shown in Figure 3.7. The 3' and 5' nor-1 integrants are GUS positive while the presumed niaD integrants are GUS negative. These data confirmed the data reported above for pAPGUSNN-B; ie integration at nor-1 results in normal nor-I promoter activity while integration elsewhere results in undetectable levels of nor-l promoter activity. In addition, the rapid site of integration PCR assay is a useful tool to identify the valuable 3' nor-1 integrants necessary for future promoter studies. 77 a} I Figure 3.6. Southern hybridization analysis of selected GUS+ and GUS- transformants for 3' nor-1 integration. (A) Confirmation of3' nor-l integration. All 8 transformants (designated by transfomiant number) were GUS+ and tested positive for 3' nor-I integration with the site of integration PCR assay. All 8 transformants tested confirmed 3' nor-I integration. (B) Testing for 3' nor-l integration. All 5 GUS- transformants (designated by transformant number) tested negative for 3' (and 5') nor-1 integration using the site of integration PCR assay. Southem hybridization analysis confirmed lack of 3' nor-] integration in these 5 GUS- tranformants. All 5 GUS+ transformants (designated by transformant number) tested positive for 5' nor-1 integration with the site ofintegration PCR assay. Southern hybridization analysis confirmed lack of 3' nor-1 integration in these 5 GUS+ transfomiants. 78 GUS+ NR114 19 32 43 67 68 81 78 4.0 ———-> 3.0——> GUS- GUS+ NR13 9140 5 93 4.5 ——> 3.0——> Figure 3.6 79 2 22 23 26 41 H 'lllllllau ' mollll“ 1"“ Mi: will . l Ii IIII‘IIIIIIIII I , ‘ . . . i" ill I "‘1 “I iI'I ' ' ii . III III .IIl'l6I iiW . ill“ M I "III IllI II 'IIIWIliIIIl . it i ill """"“ " I I?" ' III IIIIIIIII , . . i ii l ‘ . ii . III" llIH IIIIl 'I I IIiiI * lIlI Iilll ' 1'V'iiiil il'ili 'II ll 11".“: ' llIM lll'illlll I II, . liiii IIIIIIIIIIIIIIIIl ,. 'i'IIIIIIIII " . I IIl ' " ,.III|IllllIll ‘ lII' lllllllllll "'“Hi "'iIIII-....IIIII ;.I Figure 3.7. Solid culture GUS assay. Transformants l and 6 are 3' nor-I integrants and are GUS positive. Transformants 2 and 3 are 5' nor-1 integrants and are GUS positive. Transformants 4 and 5 are not 3' or 5' nor-1 integrants and are GUS negative. 80 mama“ . f] DISCUSSION The goals of this study were (i) to generate a new nor-1::GUS reporter construct (pNANG-3) that enables easy promoter replacement and (ii) validate the positional- dependent regulation effect by correlating the site of integration with solid culture GUS activity for pBNG3.0 transformants. Previously, nor-I ::GUS reporter strains generated using pAPGUSNN—B (Figure 3.1) demonstrated the usefulness and validity of the GUS reporter system (Chiou et al., 2002). A drawback with pAPGUSNN-B was the inability to replace the 3.0 kb nor-1 promoter with different nor-1 promoters. The new construct, pNANG-3, will permit analysis of multiple nor-1 promoter deletions and substitutions in order to identify cis-acting sites in the nor-1 promoter (Miller et al., 2003a and Miller et al., 2003b). Our experiments validated the positional effect by demonstrating that all 21 GUS+ transformants analyzed were either 3' or 5' nor-1 integrants while all 22 GUS- transformants tested were not 3' or 5' nor-1 integrants. These data confirmed the preliminary observation by Liang et al. (Liang et al., 1997) that chromosome location plays a role in regulation of aflatoxin gene expression. The targeting of reporter constructs to specific chromosomal locations for promoter analysis has been suggested by other investigators studying expression of fungal genes (Hamer and Timberlake, 1987 and Timberlake and Marshal, 1988 and van Gorcom et al., 1986). Although the mechanisms of position-dependent gene expression have not been fully elucidated in filamentous fungi, it has been hypothesized that enhancer elements may be responsible for the position-dependent effect (Kinsey and Rambosek, 1984). The hypothesis that positive cis-acting factors have regional control over the transcription of aflatoxin genes 81 nan-£19m 7‘; I is reasonable, since removal of aflatoxin reporter fusions from the aflatoxin gene cluster results in reduced GUS expression. An alternative explanation for the lack of GUS activity at the m‘aD loci is that a transcriptionally inactive chromatin structure exists at this site. We think that such inactivation is unlikely for two reasons. First, NR1 transformed with pGAPN2B (l3- tubulinzzGUS) expressed GUS activity at similar levels at 24, 48, and 72 h when integrated at the niaD locus indicating that transcriptional activation can occur at the niaD locus in YES agar (Chiou et al., 2002). Second, in transformants carrying pAPGUSNP (nor-1 ::GUS with pyrG as selectable marker), the nor-1 promoter carried on this plasmid was not active at the pyrG locus even though growth on YES medium I required expression of the pyrG gene (Chiou et al., 2002). In addition, several pAPGUSNP transformants had no detectable GUS activity even when they carried multiple copies of plasmid as long as no copies of the plasmid integrated at 5' or 3' nor-I (Chiou et al., 2002). Based on previous data (Liang et al., 1997 and Chiou et al., 2002) and the data presented here, we hypothesize that aflatoxin gene expression is influenced by an locus control element. The clustering, or close linking, of genes involved in the same secondary metabolic pathway is a common theme in the filamentous fungi (Keller and Hohn, 1997). However, cis—acting elements that regulate many genes simultaneously in fungal gene clusters have not been identified. If locus control region does influence the regulation of nor-I transcription, it is located at least 3 kb upstream from the transcription initiation site in the 5' nor-I region, at least 1.8 kb downstream of the transcription termination site in the 3' region of nor-l, or within the nor-l coding region. Because of similar position- dependent expression observed with the ver-I ::GUS reporter construct (Liang et al., 82 1997), it is also possible that the same locus control region is influencing the regulation of both nor-1 and ver-l . Studies performed on the SpoCl gene cluster in A. nidulans showed that clustered genes can be coordinately expressed during development and that placement of cluster genes at ectopic chromosomal locations results in the loss of that coordination (Miller et al., 1987 and Timberlake and Barnard, 1981). The physical linkage of aflatoxin biosynthesis genes also may have a regulatory role. Trichothecene biosynthesis genes are also clustered (Hohn et al., 1993b) and there is some evidence for positional-dependent regulation with the pathway regulator Tri6 (Chen et al., 2000). Introduction of a plasmid containing a Tri5::GUS fusion with a functional Tri6 results in 50-100X more GUS activity with Tri5 integration compared to ectopic integration (Chen et al., 2000). Studies designed to establish such a phenomenon and to determine the nature of the regulatory mechansim may eventually lead to a broader understanding of the expression of secondary metabolism genes and the ability to manipulate its expression in microorganisms. We utilized a rapid procedure to prepare DNA templates from fungal colonies that are sufficiently pure for the site of integration PCR assay. This development was critical for current and future studies on the regulation of the nor-1 and ver-I promoters because detection of the correct site of integration in the genome is essential to accurately measure environmental influences on the aflatoxin gene promoters. In this regard, the speed and effectiveness of the rapid assay make screening of large numbers of fungal transformants practical. In addition, screening of transformants can be based on genotype (site of integration) rather than phenotype (GUS activity) which is important for nor-l ::GUS constructs that have reduced GUS activity. 83 ACKNOWLEDGMENTS The work in Chapter 3 was a contribution to a publication by Ching-Hsun Chiou (Chiou CH, Miller MJ, Wilson DL, Trail F and Linz JE. 2002. Chromosomal location plays a role in regulation of aflatoxin gene expression in Aspergillus parasiticus. Applied and Environmental Microbiology. 68(1): 306-315). 84 21.3 CHAPTER 4 Role of AflR in nor-1 transcriptional activation in Aspergillus/laws and A. parasiticus INTRODUCTION Aflatoxins, mycotoxins produced predominantly by Aspergillus parasiticus and Aflavus, frequently contaminate several economically important crops including com, cotton, peanuts and tree nuts (Wilson and Payne, 1994). Aflatoxin Bl (AF B1), the most abundant of the aflatoxins, is also the most toxic and carcinogenic (McLean and Dutton, 1995). Animal studies have shown aflatoxin to be a potent hepatocarcinogen and human epidemiological data have linked aflatoxin exposure with liver cancer (reviewed in Hall and Wild, 1994 and Dvorackova, 1990). As a result, crops susceptible to aflatoxin contamination are monitored for aflatoxin concentrations in the United States and throughout the world imparting a huge economic cost to growers and marketers of commodities. Consequently, our long-term goal is to reduce or eliminate aflatoxin from the food chain. To accomplish this goal, we are focused on elucidating the molecular mechanisms that regulate aflatoxin biosynthesis. This information is likely to generate novel approaches and targets for inhibition of aflatoxin gene expression. Aflatoxin biosynthesis is a complex process that requires at least 16 different enzymatic steps (Bhatnagar et al., 1994). The genes involved in aflatoxin biosynthesis reside in a 70 kb cluster and appear to be co-regulated (Trail et al., 1995b). Analysis of the regulatory mutant A. flavus strain 650 (Bennett and Papa, 1988) first identified AflR as a pathway 85 "I" “VJ regulator (Payne et al., 1993). Subsequently, aflR homologs were identified in A. parasiticus (Chang et al., 1993) and A. nidulans (Yu et al., 1996). Several pieces of evidence demonstrate a key regulatory role for AflR in aflatoxin biosynthesis: 1) based on amino acid sequence identity, AflR belongs to a common class of fungal transcription factors called zinc binuclear cluster proteins (Woloshuk et al., 1994); 2) aflR null mutants do not produce detectable aflatoxin biosynthetic enzymatic activities (Payne et al., 1993) nor the aflatoxin biosynthetic transcripts stcS, stc V, sth, stcT (Yu et al., 1996); 3) increased transcription of aflR is correlated with increased aflatoxin production (Chang et al., 1993, Yu et al., 1996, Flaherty and Payne, 1997); 4) Recombinant AflR binds to several aflatoxin biosynthetic gene promoters in vitro (F emandes et al., 1998, Ehrlich et al., 1999a and Ehrlich et al., 1999b); and 5) AflR cis-acting sites have been shown to be necessary for transcriptional activation for three aflatoxin biosynthetic genes in vivo including sth (Femandes et al., 1998), avnA (Ehrlich et al., 2002) and pksA (Cary et al., 2000). Although it is clear that AflR is a key regulator of aflatoxin synthesis, it is not clear if AflR is the only specific transcription factor needed for transcriptional activation of all aflatoxin biosynthesis genes and if all consensus AflR cis-acting sites in the aflatoxin cluster are functionally significant. To address these questions, we focused attention on expression of the nor-1 gene that catalyzes the conversion of the first stable aflatoxin biosynthesis intermediate, norsolorinic acid, to averantin (Trail et al., 1994 and Zhou and Linz, 1999). The nor-1 promoter includes a consensus AflR binding site (TCGnnnnnCGR) located at -75 to -64 bp (AflRl) from the primary nor-I transcriptional start site (+1); additional upstream consensus AflR binding sites are located at -1213 (AflR2) and -1563 (AflR3). Genes upstream from nor-1 include an open reading frame of unknown function (ORF 3: 86 r. translational start at -1073) and the divergently transcribed pksA (translational start at - 1731) (Trail et al. 1995, Chang et al. 1996). A. parasiticus nuclear extracts and recombinant AflR were unable to bind to a DNA fragment containing AflRl in vitro (Ehrlich et al., 1999b) suggesting that AflRl might not be functionally relevant. Directed mutations in AflRl did not affect transcription of the divergently transcribed pksA in vivo (Ehrlich et al., 2002) suggesting AflRl may not be important for expression of kaA (effects of AflRl mutation on nor-1 or ORF 3 expression were not reported). However, using the same mutagenesis approach, both AflR2 and AflR3 were shown to be necessary for pksA transcription but effects of AflR2 and AflR3 mutation on nor-1 and ORF3 expression were not reported (Ehrlich et al., 2002). Furthermore, alteration of additional cis-acting sites by directed mutagenesis (BrlA and PacC), significantly altered pksA transcription indicating that AflR may not be the only transcriptional regulator for pksA (Ehrlich et al., 2002). In contrast, other experimental data suggest that AflR alone is necessary for transcriptional activation of the middle pathway gene avnA through a single consensus cis-acting site (Cary et al., 2000). An additional potential AflR binding site (TCGnnnnnCGR) in the avnA promoter, when mutated, did not alter avnA transcription in vivo and recombinant AflR protein was unable to bind to this non-functional binding site in vitro (Cary et al., 2000). The sth promoter has three consensus AflR binding sites within 800 bp of the translational start point (Femandes, et al., 1998). Directed mutations in the three AflR sites revealed that the most distal site, -762, had no affect on sth transcription while AflR consensus binding sites at -81 and -168 both appeared to be functional in viva (Femandes et al., 1998). The two functional AflR binding sites were not additive because .9th promoters with only the -81 site or the —168 site were indistinguishable from strains that had both 87 AflR binding sites (Femandes et al., 1998). The objective of the current study was to clarify a role for AflR and the consensus AflR binding sites AflRl, 2 and 3 in expression of nor-1 in the predominant aflatoxigenic species A. flavus and A. parasiticus. More specifically, we wanted to: I) analyze AflR binding to AflRl in the nor-I promoter in A. flavus in vitro, 2) determine if AflRl is necessary for nor-l transcriptional activation in A. parasiticus and A. flavus in vivo, 3) determine if AflR2 and AflR3 are necessary for nor-I transcriptional activation in A. parasiticus in vivo and 4) determine if AflR is the only transcriptional activator necessary for nor-I expression in A. parasiticus. Although the relevant AflR consensus sites were identified previously based on sequence analysis (Ehrlich et al., 2002), this was the. first functional analysis of these sites with respect to nor-1 promoter function in A. parasiticus and A. flavus. We showed that recombinant A. flavus AflR bound to an oligonucleotide containing AflRl using the Southwestern blot procedure. Additionally, deletion analysis of the nor-1 promoter in nor-1 ::GUS reporter strains showed that AflRl is necessary for nor-1 transcriptional activation in A. flavus in vivo. Using analogous nor-I ::GUS reporter strains in A. parasiticus containing nor-I promoter deletions and substitutions, we demonstrated that AflRl and possibly AflR2 and additional cis-acting site(s) are necessary for nor-1 transcriptional activation in vivo. We also tentatively identified a novel gene (ORF3) that may function to directly or indirectly activate nor-1 transcription in A. parasiticus. 88 MATERIALS AND METHODS Strains Aspergillus parasiticus NR-l (niaD; [Homg et al., 1990] and A. flavus 656-2 (white, aflR, pyrG, leu; [Payne et al., 1993]) were used as recipient strains for transformations. Aspergillus/lavas NRRL 3357 (National Center for Agricultural Utilization Research, Peoria IL) is a wild type strain. Plasmid Construction A. parasiticus reporter plasmids The plasmids pNANG-1 and pBNG-3.0 have been previously described (Chiou et al., 2002). Additional nor-I promoter pieces were generated by PCR using pAPGUSNN- B as template with primers with Not] (3') and Pacl (5') tails. The PCR products were ligated into the Not] and Fuel sites in pNANG-3 resulting in pBNG-n (where n designates the size of the promoter in base pairs). Plasmid maps for pNANG-3 and pBNG-3.0 are shown in Figure 4.1. A. flavus reporter plasmids GAP4 contains a promoterless E. coli uidA (GUS) gene (Woloshuk and Payne, 1994). A BamHI site was introduced at the nor-I translational start site by site directed mutagenesis (Kunkel, 1985). Then a 1.3 kb BamHI fragment containing the nor-1 promoter was ligated into the BamHI site in GAP4 resulting in GAP12. The nor-1::GUS reporter plasmids GAP12-138, GAP12-103 and GAP12-9l were created by PCR using GAP12 as a template. The downstream primer for all three constructs was the M1 3F 89 tn} 3&3 Figure 4.1. Restriction endonuclease maps of relevant plasmids. (A) pNANG-3. In addition to the niaD selectable marker. pNANG—3 carries a small part ofthe nor-l coding sequence (10 amino acids) fused to the B—glucuronidase (GUS) coding sequence which is in turn fused to the 2 kb nor-l 3' terminator fragment. Appropriate promoter pieces, amplified by PCR using primers with Null and Par-l tails, were cloned into pNANG-3. The small number ofcodons that were changed in the nor-l coding sequence to create the useful Not] site were all acceptable based on codon usage and maintained the correct sense. (B) pBNG3.0-3F. This plasmid contains a 3 kb PCR amplified nor-I promoter piece cloned into the Not] site of pNANG-3. Other nor-I promoters that were tested (1250, 1200, 664, 3321’, 332, 332AflRmut. 76 and 64) were also made by PCR and were directionally cloned into the Natl and Pucl sites in pNANG-3. (C) pGAP12. This plasmid contains the GUS gene connected to a 1.3 kb BamHl nor-1 promoter fragment. pGAP12 was used as a template for PC R to make the A. fluvus reporter plasmids used in this study. (D) pGAP12-138. This plasmid contains 138 bp upstream of translational start site ofnor-l. 9O A B Ascl 418 Sea] 16400 A861418 Pstl 14612 Cla11218 Cla12118 Pstl 11612 Sall 11600 pNANG pBNG3.0 GUS GUS 13900 bp 16900 bp Notl 4226 ScaI 4326 N0" 4226 niaD 5' "0“ Seal 5126 Pacl 4236 Pstl 10400 P28113226 S Pstl 7400 NotI 7226 Nch 183 l Pvull 306 Ndel 183 ‘ EcoRl 396 Pm" 306 Sacl 402 EcoRl 396 Sacl 824 ‘ Sacl 402 lacZ \ amp 3' 3W" Sacl 824 GAP12-138 5071bp GUS PW” 4155 Hindlll 399 SP“ 3988 'Smal 2659 Pstl 3982 33mm 2664 Pvu112993 Sall 397 Bglll 2914 Hindlll 2832 Xbal 3970 Sphl zszq BamHl 3964 Pstl 2820 Sall 2814 BamHl 2664 Figure 4.1 91 primer. The upstream primers were as follows: GTTGGCATACCATCAAATGC for GAP12—138, CCAACTCGGCCAGCGACCAACACACCACC for GAP12-103 and GCGACCAACACACCACC for GAP12-91. The PCR product was cloned into the pCRII vector from the Invitrogen TA Cloning Kit (Invitrogen, Carlsbad CA). Plasmids were named GAP12—n where n designates the number of bases upstream from the nor-l translational start site included in the vector. Figure 4.1 includes a map of GAP12 and GAP12-138. A. flavus tAflR expression plasmid construction The open reading frame (ORF) of aflR was amplified from the cosmid B9X2 (F laherty, et al., 1995) by PCR with primers that also contained restriction enzyme sites for ease of cloning the product in-frame into the expression vector, pET30c (N ovagen, Madison WI). The upper (5') primer (5'-GCGGATCCAAATGGTGACCATATCTCCC 93') contained a restriction site for the endonuclease BamHI (bolded) and 20 nucleotides of aflR sequence (underlined), beginning at the start codon of the open reading frame. The lower (3') primer (5 '-GCGAAGCTTATAATGTCGGAGGACACGGCG-3') contained a restriction site for the endonuclease HindIII (bolded) and 21 nucleotides of q/IR sequence (underlined) from the 3' end of the ORF up to, but not including, the stop codon. The resulting PCR product was 1330 bp in length (the aflR open reading frame is 131 1 bp in length). This product was cloned into the pCRII vector from Invitrogen (San Diego, CA) and the resulting plasmid was subsequently digested with BamHI and HindIII restriction enzymes for isolation of a 1323 bp fragment. This fragment was then cloned into the pET30c expression vector at the BamHI and HindlII sites. The resulting expression construct, pET30c-AFLR, contains a 1503 bp ORF, encoding 501 amino 92 acids. The predicted protein product has a molecular mass of 53,940 Daltons. The aflR ORF makes up 437 of the 501 amino acids. The other 64 amino acids make up the histidine tags at the N-terminal and C-terminal ends and an S-tag and enterokinase cleavage site at the N-terminal end of the recombinant AFLR protein. Because of solubility problems with the full-length AFLR recombinant protein, a truncated AF LR protein that contained the DNA binding domain was desired. To produce the pET30c-tAF LR construct for overexpression of a truncated AF LR protein, the pET3OC-AF LR construct was digested with XhoI and HinDIII and then religated. This allowed retention of 665 bp of the original 1323-bp fragment that was cloned into the pET30c vector. This 665-bp fragment contains sequence for the N-terminal histidine tag, the S-tag, the enterokinase cleavage site, and the 5’ end of the aflR ORF including the entire AFLR zinc binuclear cluster region, but the 3’ end of the aflR ORF was deleted. Generation and Selection of Transformants Transformation Transformation of A. parasiticus protoplasts was performed as described by Homg et al. (1990). 2-4 pg of DNA was used with approximately 107 protoplasts resulting in approximately 100 transformants. Transformation of A. flavus was as described by Woloshuk (Woloshuk etal., 1989). Site of Integration PCR Assay (A. parasiticus) A rapid DNA extraction procedure and PCR analysis were performed as 93 described by Chiou (Chiou et al., 2002). Following PCR, a 2.0 kb DNA fragment is diagnostic for 3' integration. Southern Hybridization (A. parasiticus) Genomic DNA was purified from A. parasiticus cultures shaken for 48 h in 100 ml YES (2% yeast extract, 6% sucrose and pH=5.8) liquid medium at 29°C with five, 6 mm glass beads (Skory et al., 1993) and subjected to standard agarose gel electrophoresis (Ausubel et al., 2003). Southern hybridization analysis was performed according to standard procedures (Ausubel et al., 2003). 2.5 pg of genomic DNA was digested with Seal and probed with a 900 bp ClaI fragment isolated from the nor-1 terminator region of pAPGUSNN-B (Chiou et al., 2002). Digestion of DNA from the recipient strain NR-l generates a 3.0 kb restriction fragment while a 3' integrant results in 3.7 and 4.0 kb restriction fragments. Dot Blot Hybridization Analysis (A. flavus) Co-transformants were first screened for uracil prototrophy and then screened for presence of the reporter plasmid by dot blot hybridization of genomic DNA with a GUS gene probe. Genomic DNA from cotransformants was isolated as previously described (Woloshuk, et al., 1989). Each genomic DNA sample was boiled in a total volume of 0.5 ml (final concentration = 0.4 M NaOH and 10 mM EDTA) for 10 minutes. The samples were applied to Zeta probe nylon membranes (BioRad, Melville NY) on a dot blot apparatus (Schleicher and Schuell, Keene NH) per manufacturers’ instructions. The loaded membranes were rinsed in 2X SSPE (0.18 M NaCl, 10 mM NaPO4 (pH 7.7), 1 mM EDTA), air dried, and then cross-linked. The cross-linked membranes were probed 94 «=1 .2311 fiLM’nm with a 2.0 kb Kpnl fragment of the E. coli uidA gene from GAP13 (F laherty et al., 1995) for selection of primary cotransformants containing the GUS gene. GUS Reporter Assays Solid culture GUS Assay (A. parasiticus) Autoclaved Nytran SPC (Schleicher & Schuell, Keene NH) membranes were first placed on top of YES (2% yeast extract, 6% sucrose, 1.5% agar and pH=5.8) agar plates. Transformants were then transferred via sterile toothpick to the Nytran membrane. After incubation for 46 h at 29"C in the dark, the Nytran filters were removed, frozen in liquid nitrogen and thawed at room temperature (2 repetitions) and then incubated for up to 24 h with GUS substrate solution that includes 0.04% of the colorimetric GUS substrate (X- Glu: 5-bromo—4—chloro-3-indolyl B-D-glucuronide) in GUS reaction buffer (60 mM NaZHPO4, 40 mM NaHZPO4, 10 mM KCl, 1 mM MgSO,, 0.07% D-mercaptoethanol) and visually evaluated. Liquid Culture GUS Assay (A. parasiticus) Two confirmed A. parasiticus 3' transformants containing reporter constructs were grown in duplicate for 48 h at 29°C in the dark with shaking in 100 ml of GMS (Buchanan and Lewis, 1984) liquid medium in a 250 ml flask with five 6 mm glass beads. Mycelia were collected by filtration through Miracloth (Calbiochem, La Jolla CA) and ground in a mortar with a pestle under liquid N2. Approximately 200 mg of ground mycelia was transferred to a 1.5 mL microcentrafuge tube and kept on ice. After all the samples had been collected, 500 mL of GUS lysis buffer (50 mM NaHzPO4 [pH=7], 10 mM EDTA, 0.1% Triton X-100, 0.1% SDS, 0.07% B-mercaptoethanol and 25 95 b4 mg/ml PMSF [phenylmethylsuflonyl fluoride]) was added. The samples were vortexed for 15 s and then centrifuged for 10 min at 10,000 X g at 4°C. The supematent was withdrawn and placed in a new tube. The protein concentration was determined by Bio- Rad protein assay (Bio-Rad, Hercules CA). GUS activity was determined with 100 pg of each sample incubated at 30°C with 145 pg of the GUS substrate 4-methylumbelliferyl B-D—glucoside (MUG) in a final volume of 200 pl. At 0, 10 and 30 min, 800 pl of stop solution (200 mM NazCO3) was added. After stopping the GUS reaction, 200 pl of sample was loaded into a microtiter plate which was analyzed using a Cytofluor II fluorometer (Biosearch Co., Bedford MA). Excitation and emission wavelengths were 360 nm and 460 nm, respectively. To evaluate the fluorometer results, a standard curve of 4-Methylumbelliferone (10 nM to 600 nM) in GUS lysis buffer was also read in the fluorometer. GUS activity was reported as pmols 4-methylumbelliferone / minute / mg protein. Quantitative GUS Assay (A. flavus) Growth of selected transformants for quantitative analysis of GUS activity was carried out in 24-well plates (Becton Dickinson & Company, Lincoln Park, NJ) in media conducive (sucrose low salts; SLS) and non-conducive (peptone mineral salts; PMS) to aflatoxin production and expression of pathway genes, as previously described (Flaherty, et al., 1995). Conidia ( lxlO°/well) were suspended initially in 1 ml of PMS and allowed to grow for 3 d at 28°C. After this initial growth period, three replicate mycelial mats of each transformant were collected (0 h time point), while three additional replicate mycelial mats of each transformant were resuspended in 1 ml of either PMS or SLS medium and allowed to continue growth at 28°C. These additional replicates were 96 harvested 24 h after resuspension (24 h PMS and 24 h SLS timepoints). The harvested mycelial mats were placed in 1.5 ml disposable snap-cap tubes, quick-frozen with liquid N, and placed at —80°C until protein extraction could be carried out. Cultures for each time point (0 h, 24 h PMS, and 24 h SLS) were grown in separate 24-well plates, and after collection of the mycelial mats at each timepoint, the plates were placed at —20°C to preserve the culture filtrates for aflatoxin analysis. The GUS activity of cotransformants was measured quantitatively using the procedures of Jefferson (Jefferson, 1987) for analyzing GUS activity in plant tissue as modified by Flaherty (F laherty et al., 1995) for analyzing GUS activity in A. flavus tissue. One exception was that 50 pl of each extracted fungal protein sample was added to 500 p1 of GUS substrate buffer (1 mM 4- methyl-umbelliferyl B-D-glucuronide in extraction buffer) which had been pre-incubated at 37°C. After 1 h incubation at 37°C, 100 pl of assay buffer (extracted protein sample + substrate buffer) was removed and added to 900 pl of stop buffer (0.2 M NazCO3). Relative fluorescence was subsequently measured using a Hoefer TK-lOO fluoremeter following the manufacturer’s instructions. GUS activity was reported as nmole MU / min / mg protein. A. flavus tAflR (truncated AflR) binding studies Induction and Purification of tAflR A truncated aflR gene product, tAF LR, was expressed in a bacterial system for production and purification of recombinant tAFLR protein for use in DNA binding studies. Single colonies of PET30c-tAflR were used to inoculate LB-Kan (30pg/ml, 500 ml LB). The culture was grown at room temperature for 4 h followed by 37°C for 3 h with shaking at 200 rpm. When the OD“, was approximately 0.9, IPTG (isopropyl-Q-D- 97 thiogalactopyranoside) was added to a final concentration of 0.4mM. Cultures were allowed to grow for an additional 2-3 h. Cells were collected by centrifugation and frozen for subsequent purification. A pellet from 250ml of culture was used for purification on a Ni-NTA agarose column (Qiagen, Valencia CA) according to manufacturers specifications with modifications. Briefly, the pellet was resuspended in 4 ml Buffer B (8M urea, 0.1M NaHzPO4, 0.01M TRIS/HCL, pH 8.0). Cells were partially lysed by gentle homogenization and centrifuged to obtain a cleared lysate (10,000 x g for 10 min.). The Ni-NTA column was pre-equilabrated with Buffer B and 1.8 ml cleared lysate was added to the column. The column was washed with 9 ml Buffer C (8M urea, 0.1M NaHZPO4, 0.01M TRIS/HCL, pH 6.3). After washing, the protein was eluted with 9 ml Buffer E (8M urea, 0.1M NaHZPO4, 0.01M TRIS/HCL. PH 4.5). A final elution step was performed with 3 ml of 0.75 x Buffer E with 250mM imidazole. The final 6 ml of the eluted fractions were dialyzed in 500ml dialysis buffer ( lOmM TRIS, pH7.6, 200mM KCL, lmM EDTA, SmM DTT and 10% glycerol) with decreasing concentrations of urea : 6M, 3M, 2M, 0.5M and 0M.. Protein concentration was determined by the Bradford method. Preparation of oligo probes PP2 was used a probe while PP2 and PP2MUT were used as competitors for Southwestern blot analysis. PP2 (CCAACTCGGCCAGCGACCAACACACCACC) sequence is from the A. flavus nor-1 promoter and includes AflRl , an AflR consensus binding site (TCGnnnnnCGA). PP2MUT (CCAACQCIGCCAGCGACCAACACAC CACC) is the same as PP2 except for two mutations (underlined residues) in the consensus AflR binding site. Double stranded Oligonucleotides were used for labeling 98 and cold competition experiments. A typical end—labeling reaction used 1 pmole double stranded 011 go with 8-10 units polynucleotide kinase (Promega, Madison WI) and 1.5 pl gamma ATP (3000 Ci/mmol, 10 mCi/ml) (New England Nuclear, Boston MA). The reaction was incubated at 37°C for 10 minutes and stopped with the addition of 2 pl 0.5 M EDTA. Oligonucleotides were purified with Centn' Spin 20 columns (Princeton Seperations, Adelphia NJ) according to manufacturers instructions. Southwestern Blot Southwestern blots were performed essentially as described by Tully and Cidlowski (Tully and Cidlowski, 1993). Briefly, 2.8 pg of tAflR per lane was fractionated on SDS-PAGE, incubated for 2 x 1h washes in renaturation buffer (50mM NaCl, lOmM TRIS-HCI at pH 7.5, 20mM EDTA, 0.1mM dithiothreitol and 4M urea) and electro-blotted (mini tank system from BioRad, 84volts and initially 294 milliamps, 1-1.5 h) onto nitrocellulose. Nitrocellulose filters were blotted overnight in 200ml blocking buffer (5% (w/v) nonfat dry milk, SOmM NaCl, IOmM TRSl-HCl, pH 7.5, and lmM EDTA). Vertical strips, containing a molecular weight marker lane and a tAflR lane, were cut from the nitrocellulose filter. Filters were placed into heat-scalable plastic bags and incubated for 1.5 h at room temperature with 1 ml blocking buffer or 1 ml blocking buffer supplemented with 5 pmole competitor DNA. 250 ml blocking buffer containing 2 x 10° cpm PP2 (approx. 0.34 pmole for a specific activity of 1 x108 cpm/mg) was added to each bag and incubated at room temperature for 1.5h. Filters were removed from the bag and washed in 50 m1 blocking buffer for 30 min, rinsed briefly in TBS-T, allowed to dry and autoradiography was performed. 99 RESULTS AflR binding to the A. flavus nor-1 promoter Southwestern blot analysis was performed to test the ability of a truncated AflR (tAflR) to bind to an oligonucleotide containing an AflR consensus binding site (AflRl, Figure 4.2). 32P labeled oligonucleotide PP2 (contains AflRl binding site from the A. flavus nor-1 promoter) bound to tAflR (lane 1). To demonstrate specificity, competition experiments were performed. A 15 fold molar excess of unlabeled PP2 competed with labeled PP2 (lane 2) while a 15 fold excess of PP2MUT (carries two mutations in the consensus AflR binding site) competed marginally with labeled PP2 for tAflR binding (lane 3). These data confirm that AflR protein can bind to AflRl in a specific manner. A. flavus GUS Activity Assays Aspergillusflavus 656-2 was co-transformed with plasmid pAF-l containing the pyr4 gene for uracil biosynthesis (Payne et al., 1993) and GAP12-n (n = size of nor-1 promoter). Dot blot analysis was used to screen the pyr4+ transformants for presence of the GUS gene (data not shown). Five transformants of each deletion construct that contained a single copy of GUS were used for quantitative analysis. Each transformant was grown for 3 days in PMS (aflatoxin non-supportive medium) and then transferred to either PMS or SLS (aflatoxin supportive medium) for l d. Three different A. flavus strains (see Figure 4.3) were used for GUS analysis. GAP12-91 (lacks AflRl) had no detectable GUS activity in either PMS (non aflatoxin supportive medium) or SLS (aflatoxin supportive medium). GAP12-103 and GAP12-138 both contain AflRl and had GUS activity in PMS and SLS. However, a 4.5 fold induction (activity in SLS divided 100 208 _ 129 — 86 '— 32.8— 18.1— 7.4 — Figure 4.2. Southwestern blot analysis of AflRl with purified tAflR (A. flavus) and a PP2 probe. PP2 (CC AACTCGGCCAGCGACCAACACACCACC), a 29 bp oligo from the A. flavus nor-1 promoter, contains the AflR binding site AflRl (bold). PP2MUT (CCAACQCIGCCAGCGACCAACACACCACC) is the same as PP2 except for 2 mutations (underlined) in the AflR binding site. PP2 was labeled with 32P and used as probe in all three lanes. Competitors had a 15 fold molar excess and were unlabeled. Lane 1, no competitor; lane 2, PP2 competitor; lane 3, PP2MUT competitor 101 L r... 3...’ .r 1.. ”l Ki Fold PMS SL Induction GAP 12-138 1 2.2 9.9 4.5 AFLR GAP 12-103 m 1 3.5 9.9 2.8 GAP 12-91 I I ND. ND. - Figure 4.3. Schematic of the A. flavus nor-1 promoters and GUS analysis. The AflR binding site AflRl is located between residues ~103 and -91. Fold induction was determined by dividing the GUS activity (nmole MU / min / mg protein) in an inductive medium (SLS) by GUS activity in a non-inductive medium (PMS). 102 by activity in PMS) was observed for GAP 12-138 and a 2.8 fold induction was observed for GAP 12-103 confirming that AflRl is necessary for nor-1 transcriptional activation in A. flavus in vivo. Generation and Screening of A. parasiticus Transformants In order to determine whether AflR is necessary for nor-1 transcriptional activation in A. parasiticus, several nor-I ::GUS reporter plasmids were constructed (Figure 4.4). We defined the upstream border of the nor-1 promoter as the polyadenylation site for ORF 3, located 332 bp upstream from the primary transcriptional start site of nor-I . Deletion analysis generated clones with 332, 76 and 64 bp promoter fragments (AflRl located between -76 and -64) carried on nor-1 ::GUS constructs. GUS activity in transformants that carry these plasmids was analyzed to determine functionality of AflRl. In addition, a transformant carrying a substitution of AflRl (TCchcagCGR to AGTttaaaCAG) in the context of the 332 bp promoter was also analyzed. Clones carrying larger nor-I promoter fragments (3000, 1250, 1200 and 664 bp) were analyzed to determine if any additional upstream cis-acting sites including AflR2 and AflR3 were involved in nor-I transcriptional activation. Each nor-1::GUS reporter construct could theoretically integrate into the A. parasiticus chromosome by homologous recombination at three independent sites: niaD, 5' nor-l (nor-I promoter) and 3' nor-l (nor-1 terminator). Screening for clones in which integration occurred at the 3' nor-I locus was essential for two reasons: 1) niaD integrants have been shown to have lower transcriptional activity than aflatoxin cluster integrants (Liang et al., 1997 and Chiou et al., 2002); and 2) 5' integrants result in the chromosomal nor-I promoter fused to the GUS gene and the plasmid nor—1 promoter 103 -1073 -520 +1 -1731 -1553 -1205 ATG stop -65 43...:— m AflR3 AflR2 BrlA3 PacCl pksA ORF3 nor-I Figure 4.4. Schematic of the nor-I promoter region in A. parasiticus. The numbers indicate the number of nucleotides included upstream from the transcriptional start site. Several potential cis-acting sites are indicated including the AflR binding sites AflRl, AflR2 and AflR3 and PacCl and BrlA3 which were reported to be involved in pksA transcriptional regulation (Ehrlich et al., 2002). The location of an open reading frame (ORF) of unknown function is also shown. The sizes of the nor-l promoters used in this study are also indicated (1250, 1200, 664, 332, 76 and 64). 104 fused to the chromosomal nor-I gene. Transformants from the nor-1 ::GUS reporter constructs were screened initially by a rapid PCR assay specific for nor-1, 3' integration (Chiou et al., 2002). A 3' nor-I integrant generates a 2.1 kb DNA fragment in the rapid PCR integration assay regardless of the size of the nor-I promoter in the reporter plasmid. The percentage of transformants that had a 2.1 kb band with the rapid assay varied from experiment to experiment ranging from 2 to 10% (data not shown). While the PCR assay could identifyI 3' integrants, it could not identify multiple integrants (multiple 3' integrations and/or additional non-3' integrations). Consequently, Southern hybridization analysis was used to confirm PCR integration assay results for each reporter construct. Disappearance of a 3.0 kb DNA fragment in the recipient strain, NR- 1, and the appearance of 3.7 and 4.0 DNA fragments was diagnostic for 3' nor-1 integration (Figure 4.5). Two confirmed 3' nor-1 integrants for each nor-I ::GUS reporter construct were used for solid culture and liquid culture GUS reporter analysis. A. parasiticus GUS reporter assays The nor-1::GUS transformants were tested for GUS activity using liquid culture (Figure 4.6) and solid culture assays (Figure 4.7). Liquid culture measurement of GUS reporter activity was reported as the average GUS activity in 2 independent transformants per nor-1 ::GUS reporter construct analyzed in duplicate after growth for 48 h in GMS liquid shake cultures. The solid culture GUS activity assay was performed on 48 h colonies grown in duplicate on YES solid medium. In order to determine if AflRl was necessary for nor-I transcriptional activation, fungal isolates carrying the 332, 332AflRmut, 76 and 64 bp promoter fragments in nor- ] ::GUS constructs were tested for GUS activity using liquid culture (Figure 4.6) and solid culture (Figure 4.7A) activity assays. A. parasiticus isolates carrying the 105 A-) Figure 4.5. Southern hybridization analysis of A. parasiticus transformans with 332AflRmut (332*), 332, 76, 64 and NR-l. Each letter indicates a different isolate from the same nor-I ::GUS construct. Two of the 3' integrants shown from each nor-l ::GUS reporter construct were used for solid culture and liquid culture GUS assays. 106 Sample 3000 H AflR3 AflR2 '> :2 E 1250 - AflR2 AflRl 1200 664 332P 332 332AflRmut NR-l b 1:: ——> a AflRl Hui AflRl ———-— AflRl —-E.T.i AflR] AflRl GUS Activity (pmol/min/mg) 168i115 429 :t 442 17.2 :t 8.1 16.0:l:4.0 21.1 i 14.5 5.4i 2.2 ND ND Figure 4.6. Liquid culture GUS assay analysis was performed on two different 3' integrants from each nor-l ::GUS reporter construct grown in duplicate. The GUS activity is reported as the mean pmol/min/mg :t the standard deviation. 107 '“~», 3000 332 w I 125 .. 332Aflanut IIIIIIII IIIII'IIII . , iIlIIlllllllllil ‘iNRl . ‘ 1 7° .3 I» 1111200 1‘ ,1. ,IlIII. wllIIIIlIlIlll“ I ll‘ NR1 64 . ' "iii... , ""1884 " ,1 I" Figure 4.7. Solid culture GUS assay analyses were performed on different 3' integrants from each nor-1: :GUS reporter construct on 46 h YES agar colonies. The recipient strain NR-l was added as a negative control and had no detectable activity. (A) The 332 nor- 1::GUS transformant (includes AflRl) had detectable GUS activity while the 332AflRMUT (AflRl mutated), 76 (includes AflRl) and 64 (AflRl deleted) nor-1::GUS transformants had no detectable GUS activity after 24 h incubation with GUS substrate. (B) The 3000 (includes AflRl , AflR2 and AflR3) and 1250 (includes AflRl and AflR2) nor-[::GUS transformants had similar activity. The 1200, 664 and 332P (all 3 include AflRl only) nor-l ::GUS transformants all had similar activity which was significantly less than the 3000 and 1250 nor-[::GUS transformants. The 332 nor-[::GUS transformant (includes AflRl) had significantly less GUS activity than all other nor- 1: :GUS transformants shown. 108 332 nor-1 ::GUS construct (includes AflRl) converted 5.4 pmol/min/mg while the isolates carrying the 332AflRmut nor-1::GUS transformant (AflRl mutated) had no detectable GUS activity (Figure 4.6). In agreement with these liquid culture data, isolates carrying the 332 nor-1::GUS construct (includes AflRl) showed clearly detectable GUS activity (blue color within colonies) while the 332AflRmut (AflRl mutated), 76 (includes AflRl) and 64 (AflRl deleted) nor-l ::GUS transformants displayed no detectable GUS activity with the solid culture assay (Figure 4.7A). These data confirm that AflRl is necessary for nor-1 transcriptional activation and that an additional cis-acting site may be located between -76 and -332. A. parasiticus isolates carrying nor-1 ::GUS with either a 1250 bp nor-1 promoter fragment (includes AflRl and AflR2) or a 1200 bp promoter fragment (includes AflRl only) were analyzed to determine if AflR2 is necessary for nor-l transcriptional activation. Deletion of AflR2 resulted in at least a 10 fold reduction in liquid culture GUS activity (Figure 4.6). Fungal isolates carrying the 1250 bp promoter in the nor- ] ::GUS construct clearly demonstrated more activity than the 1200 bp promoter in the solid culture GUS assay as well (Figure 4.7B). These data strongly suggest that the presence of AflR2 in the nor-1::GUS construct results in greater nor-1 expression. The role of AflR3 in nor-1 transcriptional activation was analyzed using two nor- ] ::GUS fusion constructs; one carrying a 3000 bp nor-1 promoter fragment and the other a 1250 bp promoter fragment (Figure 4.6 and 4.7B). Fungal isolates carrying nor- 1::GUS constructs with the 3000 bp fragment (includes AflRl, AflR2 and AflR3) and the 1250 bp fragment (includes AflRl and AflR2) driving expression of nor-1::GUS had similar activities in both the liquid culture and solid culture GUS assays suggesting that AflR3 is not involved in nor-I transcriptional activation and that no additional cis-acting 109 sites for nor-1 transcriptional activation are located between -l250 and —3000. Fungal isolates carrying nor-l ::GUS constructs with 1200, 664, 332P and 332 promoter fragments were analyzed to determine if there were any cis-acting sites located between -1200 and -332 in the nor-1 promoter. Isolates carrying nor-I ::GUS constructs with the 1200 and 664 promoter fragments (both include AflRl only) had similar liquid culture (Figure 4.6) and solid culture (Figure 4.7B) activities which were greater than activity measured for isolates carrying the 332 promoter fragment. We hypothesized that the increased activity in these larger (1200 and 664 bp) promoter fragments was due either to the presence of a cis-acting site located between -332 and -664 or possibly due to negative effects of plasmid-derived sequences located adjacent to and upstream from the 332 bp promoter fragment. To distinguish between these possibilities, fungal isolates carrying nor-1 ::GUS with 332 and 332? promoter fragments were analyzed. The 332P promoter fragment has a 503 bp PCR product that includes the pyrG translational stop codon and transcriptional terminator inserted into pBNG332 between the niaD selectable marker and nor-1 promoter at the Fuel site (Figure 4.8). Fungal isolates carrying the 332P, nor-l ::GUS construct (has AflRl and pyrG 3') displayed a clear increase in GUS activities compared to isolates carrying the 332, nor-1::GUS construct in both the liquid culture (Figure 4.6) and solid culture GUS assays (Figure 4.7B). In addition, isolates canying the 332P, nor-l ::GUS construct had similar GUS activity to the isolates carrying the 664 and 1200 nor-1::GUS constructs as measured in both liquid culture (Figure 4.6) and solid culture (Figure 4.7B) assays indicating that the difference in activity between 332 and 664 is likely not due to a specific cis-acting site in that region. 110 PacI 332 _—l.:," . . ——I niaD niiA 332 bp GUS 3’ nor-1 5’ nor-I PacI PacI 332P l——I I. I ,r: 4 niaD niiA 3’ pyrG 332 bp GUS 3’ nor-1 5’ nor-I Figure 4.8. Genetic map of 3' integrants for both 332 and 332P nor-1 ::GUS transformants. 3' integration with the plasmids used in the A. parasiticus study result in the 11100 selectable marker being immediately upstream of the nor-1::GUS promoter. Included in the 7.4 kb niaD fragment is 680 bp of niiA coding sequence. A 503 bp PCR product that included the pyrG translational stop codon and transcriptional terminator was inserted into pBNG332 between the niaD selectable marker and nor-1 promoter at the P061 Site. 111 fl. 4.; .90mb'a.‘ DISCUSSION Since its discovery in A. flaws in 1993 (Payne et al., 1993), AflR frequently has been described as “the” aflatoxin biosynthesis pathway regulator. Experiments with AflR null mutants and AflR inducible expression strains along with AflR-DNA binding studies have, at least in part, supported this designation (reviewed in introduction). However, electrophoretic mobility shifi assays with A. parasiticus recombinant AflR and fungal protein extracts suggested that A. parasiticus AflR either does not bind AflRl in the nor-I promoter or has a much lower affinity for AflRl compared to consensus AflR binding sites in several other aflatoxin biosynthetic promoters (Ehrlich et al., 1999b) casting doubt on the functional significance of AflRl. In contrast to these observations, we demonstrated that recombinant A. flavus tAflR binds to AflRl in vitro. Because the consensus AflRl sequence is identical in A. parasiticus and A. flavus and because A. parasiticus AflR complements a AflR null mutant in A. flavus (Chang et al., 1993), we hypothesize that native AflR binds to the AflRl site in the nor-I promoter in both species in vivo. We tested this hypothesis by analyzing the effects of deletion and replacement mutations on nor-I promoter function using nor-1 ::GUS reporter constructs. In previous studies, three aflatoxin biosynthetic gene promoters have been studied in some detail: avnA with two consensus AflR binding sites (A. parasiticus; Cary et al., 2000), kaA (A. parasiticus; Ehrlich et al., 2002) and sth (A. nidulans; F emandes et al., 1998) with three consensus AflR binding sites each. With avnA, only one of the AflR binding sites was functional. The most distal AflR binding site in kaA (AflRl) and sth had no impact on transcriptional activation of their respective genes. The other two AflR binding sites in the kaA promoter, AflR2 and AflR3, are both needed for maximal kaA 112 transcription. With sth, only one of the two functional AflR binding sites is necessary for maximal sth transcription. In our study, deletion of AflRl in the A. parasiticus or A. flavus nor-1 promoter greatly decreased GUS activity. In addition, substitution of only nucleotide residues in AflRl in the 332 base pair A. parasiticus nor-1 promoter also resulted in a significant decrease in GUS activity. These data confirm that AflRl is necessary for nor-1 transcriptional activation in A. flavus and A. parasiticus. It is currently unknown if the difference in GUS activity between the 138 and 103 nor- ] ::GUS transformants in A. flavus is biologically significant. Comparison of GUS activity data between A. parasiticus isolates carrying nor- ] ::GUS with the 1250 and 3000 bp promoter fragments suggests that there are no key cis- acting sites located in that region that are necessary for nor-1 transcriptional activation supporting the conclusion that AflR3 is non-functional with respect to nor-I expression. Others have shown that AflR3 is clearly required for pksA expression (Ehrlich et al., 2002). Perhaps the most surprising observation in our study was that deletion of 50 nucleotide residues containing the AflR2 site in the 1250 nor-1 ::GUS promoter fragment resulted in at least a 10 fold reduction in GUS activity; deletion of AflRl then reduced GUS activity to non-detectable levels. These data allow us to conclude that AflRl is necessary for full nor-1 expression (332 bp nor-I ::GUS activity defined as full activity). The data also strongly suggest that AflR2 is necessary for greater nor-I expression. We propose two alternative models to explain these data: 1) AflR2 works synergistically with AflRl to mediate transcription of the nor-1 promoter; or 2) AflR2 mediates expression of ORF 3 (potentially encodes a polypetide of approximately 300 amino acid residues - Figure 4.4) directly downstream from AflR2 which directly or indirectly impacts transcription of the nor-1 promoter. Identification of a cDNA 113 corresponding to ORF 3 in an A. parasiticus cDNA library (a generous gift from Dr. Jeff Cary, USDA) strongly suggests it represents a functional gene. Interestingly, blast searches using ORF3 as a query sequence have not provided solid clues regarding potential function. Model 2 allows us to make 2 related predictions regarding nor-I promoter function. 1) Since fungal isolates carrying nor-1 ::GUS constructs with the 3000 and 1250 bp promoter fragments carry two copies of ORF 3 with AflR2 and show the highest GUS expression levels, accumulation of additional ORF3 protein in strains with two copies overcomes a protein threshold resulting in extreme upregulation of nor-1 promoter activity. 2) Loss of ORF3 function due to AflR2 deletion in the 2'1d copy accounts for downregulation (or lack of upregulation) of both nor-1 and pksA expression. Similar results are seen with the insertion of an additional copy of AflR (Chang et al., 1995) suggesting a possible regulatory role for ORF3. Transcriptional read-through from ORF 3 may also explain the affect seen with AflR2. These predictions will be analyzed in follow-up studies. While it is clear that AflR is a key positive regulator of aflatoxin biosynthesis, the case for AflR being the sole regulator of all aflatoxin biosynthesis structural genes is not as strong. For example, AflJ has been putatively assigned a role as a transcriptional co- activator (Chang et al., 2001) and has been reported to directly interact with AflR (Chang and Yu, 2002). It is unknown how AflJ acts as a transcriptional co-activator. Studies with the kaA promoter (Ehrlich et al., 2002) found evidence that both PacC (pH sensing) and BrlA (sporulation) can impact kaA transcriptional regulation through cis-acting sites in the pksA/nor-I intergenic region (see F igure4.4). However, deletion of these consensus cis-acting sites for PacC and BrlA do not affect nor-l transcriptional regulation in A. parasiticus under the culture conditions tested here. 114 Solid culture GUS analysis of fungal isolates carrying nor-1 ::GUS constructs with the 332 and 76 bp promoter fragments indicate that an additional cis-acting site(s) located between —332 and -76 contributes to nor-1 transcriptional activation. Because no additional consensus AflR binding sites (TCGnnnnnCGR) can be found in this promoter region, it follows that one or more unknown transcriptional activitors(s) bind to this promoter region and influence nor-I transcriptional activation in A. parasiticus. In support of this idea, recent experiments have localized 3 additional cis-acting sites located in the 332 bp nor-1 promoter fragment (Miller and Linz, unpublished data) which influence nor-I promoter function in vivo. In addition, protein extracts have been shown to specifically bind to these cis-acting sites using electrophoretic mobility shift assays (Miller and Linz, unpublished data). However, we can not rule out the possibility that AflR binds to additional non-consensus AflR binding sites in the nor-1 promoter. Our studies with A. flavus demonstrate that AflRl is necessary for nor-1 transcriptional activation but evidence is lacking for additional transcription factor(s) binding sites. The regulation of middle and late aflatoxin biosynthetic genes is also possibly regulated by additional transcriptional factors. With a full length sth promoter fused to GUS, only a 2-3 fold increase in activity was found compared to an AflR mutant strain (F emandes et al., 1998). In addition, substitution of both AflR binding sites only resulted in an approximately 5 fold reduction (not reduced to baseline expression levels) in GUS activity (Femandes et al., 1998). Due to the relatively high activity in the AflR mutant strain and the moderate decrease resulting from binding site substitution, the possiblity exists that other transcriptional activators besides AflR are involved in sth transcriptional activation (Femandes et al., 1998). For the middle gene avnA in 115 A. parasiticus, substitution of the AflR binding site resulted in a 10 fold decrease in promoter activity (Cary et al., 2000). A possible shortcoming of many aflatoxin promoter studies is that the integration site for reporter constructs was located outside of the aflatoxin gene cluster (eg trpC for sthzzGUS and niaD for avnAzzGUS) (Femandes et al., 1998 and Cary et al., 2000). Previous studies have revealed a greater than 500 fold decrease in promoter activity with integration of ver—I ::GUS at niaD versus integration at ver-I (Liang et al., 1997). While it appeared that the timing of transcription was the same, the magnitude was severely affected (Liang et al., 1997). A similar effect has also been reported at the pyrG locus (Chiou et al., 2002). In our current work, the A. parasiticus nor-1 ::GUS reporter constructs were integrated at the nor-1 locus at which we have demonstrated “normal” timing and level of transcription as compared to the wild type nor-1 gene (Chiou et al., 2002). While it is known that insertion of aflatoxin biosynthetic genes at niaD results in a dramatic down regulation of aflatoxin gene transcription, it is unknown if insertion of niaD into the aflatoxin gene cluster can have an effect on neighboring aflatoxin gene transcription. 3' integration with the plasmids used in the A. parasiticus study result in the niaD selectable marker being immediately upstream of the nor-1::GUS promoter (Figure 4.8). Included in the 7.4 kb niaD fragment in pNANG is 680 bp of niiA coding sequence. A 503 bp PCR product that includes the pyrG translational stop codon and transcriptional terminator was inserted into pBNG332 between the niaD selectable marker and nor-l promoter at the PacI site (Figure 4.8). Fungal isolates carrying the 332P, nor-1::GUS construct (has AflRl and pyrG 3') had significantly higher GUS activity than isolates carrying the 332, nor-1::GUS construct (Figure 4.68 and 4.7). In 116 addition, isolates carrying the 332P, nor-1 ::GUS construct had similar GUS activity to the 664 and 1200 nor-I ::GUS transformants (Figure 4.68 and 4.7) indicating that the difference in activity between 332 and 664 may not be due to a specific cis-acting site in that region of the nor-1 promoter. Rather, the neighboring niaD/niiA genes appear to have a negative affect on nor-1 transcriptional activation that can be mitigated by inserting the pyrG fragment (or native sequences) as a spacer. In summary, we demonstrated recombinant A. flavus AflR binding to an oligonucleotide containing the proposed AflR binding site AflRl using the Southwestern blot procedure. Additionally, deletion analysis with nor-1 ::GUS reporter strains showed that AflRl is necessary for nor-1 transcriptional activation in A. flavus. Using nor— ‘ 1::GUS reporter strains containing nor-1 promoter deletions and substitutions, we demonstrated that in A. parasiticus, AflRl and possibly AflR2 and additional cis-acting site(s) are necessary for nor-1 transcriptional activation. We also tentatively identified a novel gene (ORF 3) that may function to directly or indirectly activate nor-I transcription in A. parasiticus. ACKNOWLEDGMENTS Chapter 4 has been submitted for publication. (Miller MJ, Brown-Jenco C, OBrian G, Payne GA and Linz JE. submitted 2003. Role of AflR in nor-1 transcriptional activation in Aspergillusflavus and A. parasiticus. Appied and Environmental Microbiology.) All experiments with A. parasiticus described in Chapter 4 were performed by Michael Miller. Michael Miller also was the primary author. 117 CHAPTER 5 Identification of novel cis-acting sites in the aflatoxin biosynthetic nor-I promoter of Aspergillus parasiticus INTRODUCTION Aflatoxin is a potent hepatocarcinogen that can contaminate several commodities including corn, cotton, peanuts and certain tree nuts (Council for Agricultural Science and Technology, 2003). Of the four species of Aspergillus that make aflatoxin, only two are of economic importance, A. flavus and A. parasiticus (Council for Agricultural Science and Technology, 2003). In areas of the world with high aflatoxin exposure, aflatoxin and hepatitis B interact as risk factors for human liver cancer (Scholl etal., 1995). Due to health concerns, several countries have established limits for aflatoxin contamination including the United States. As a result, more than $100 million is estimated to be lost in the United States each year due to aflatoxin contamination (Robens, 2001). Due to health and economic concerns, it is desirable to reduce or eliminate aflatoxin from the food chain. A potential means to accomplish this goal is to elucidate the molecular mechanisms that regulate aflatoxin biosynthesis. This information is likely to generate novel approaches and targets for inhibition of aflatoxin gene expression. Aflatoxin biosynthesis is a complex process that requires at least 16 different enzymatic steps (Bhatnagar et al., 1994). A11 identified aflatoxin structural genes reside in a 70 kb gene cluster and appear to be co-regulated (Trail et al., 1995b). In addition to several 118 structural genes, the aflatoxin gene cluster also contains at least one regulatory gene, aflR (Payne et al., 1993). Several pieces of evidence demonstrate a key regulatory role for AflR in aflatoxin biosynthesis (reviewed in Miller et al., 2003a). Although it is clear that AflR is a key regulator of aflatoxin synthesis, it is not clear if AflR is the only specific transcription factor needed for transcriptional activation of all aflatoxin biosynthesis genes. To address this question, we have focused attention on expression of the nor-1 gene (Chiou et al., 2002, Miller et al., 2003a, Miller et al., 2003b). Nor-l catalyzes the conversion of the first stable aflatoxin biosynthesis intermediate, norsolorinic acid, to averantin (Trail et al., 1994 and Zhou and Linz, 1999). Previously identified cis-acting sites in the nor-I promoter include a consensus AflR binding site (TCGnnnnnCGR) located at -75 to -64 bp (AflRl) from the primary nor-I transcriptional start site (+1); additional upstream consensus AflR binding sites are located at -1213 (AflR2) and -1563 (AflR3). Genes upstream from nor-1 include an open reading frame of unknown function (ORF 3: translational start at -1073) and the divergently transcribed pksA (translational start at -l731) (Ehrlich et al., 2002 and Miller et al., 2003a). Initial experiments with a 3.0 kb nor-1 promoter fused to the GUS (uidA) reporter gene validated the use of the GUS reporter system (Chiou et al., 2002). In addition, valuable screening techniques were developed including a rapid PCR site of integration assay and a solid culture GUS assay (Chiou et al., 2002). Subsequent analysis using these nor-I reporter constructs demonstrated that AflRl and possibly AflR2 are necessary for maximum nor-1 transcriptional activation in vivo under aflatoxin inducing conditions (Miller et al., 2003a). Preliminary experminents also identified a potential cis- acting site located between -76 and -332 in the nor-1 promoter (Miller et al., 20033). 119 Only 5 A. parasiticus aflatoxin biosynthetic genes have had their transcriptional start point experimentally determined: nor-I (Trail et al., 1994), avnA (Cary et al., 2000), kaA (Ehrlich et al., 2002), ver—I (Skory et al., 1992) and aflR (Ehrlich et al., 1999a). While putative TATA boxes have been identified in 4 of them (all but aflR) , only the TATA box in avnA has been functionally analyzed (Cary et al., 2000). Deletion of the TATA box in the avnA promoter resulted in at least a 5 fold reduction in avnA transcriptional activation (Cary et al., 2000). Unfortunately, deletion of the TATA box also resulted in deletion of a functional AflR cis-acting site and perhaps other functional cis-acting sites. Replacement of the TATA box in the context of a wild type promoter would provide direct evidence for TATA box function in any promoter of interest I including avnA. The objective of the current study was to further define cis-acting sites that are necessary for maximum nor-1 transcriptional activation in Aspergillus parasiticus under aflatoxin inducing conditions. Specifically, we wished to: I) test the in vivo function of a putative TATA box in the nor-1 promoter, 2) further refine the location of a cis-acting site previously shown to occur between nucleotide residues -76 and -332 (+1 is transcriptional start) using nor-1 reporter constructs in vivo, and 3) test for in vitro protein binding to potential cis-acting sites. Although putative TATA boxes have been identified in aflatoxin biosynthetic promoters, this was the first direct functional analysis of a TATA box in an aflatoxin gene promoter. Substitution of the TATA box in the context of a larger nor-I promoter resulted in non-detectable GUS activity. We identified a novel cis-acting site (norL) located between —210 and -238 that is necessary for maximum nor-1 transcriptional activation in vivo. Using electrophoretic mobility shift assays, we demonstrated specific protein binding to norL (N orpr) and an 120 additional site, CREl (CREbp). Both norpr and CREIbp appear to rely on functional AflR for maximum DNA binding. Lastly, we propose a model for how AflR, Norpr and CREbp may interact to govern nor-I transcriptional activation. MATERIALS AND METHODS Strains and growth conditions Escherichia coli DHSOL F’c [F ’ endAI hstI 7 (rk- mk-) supE44 thi-I recA gyrA (Nal') ArelAI (lacZ YA argF)u,69:(m80 AlacZ M15)] (Invitrogen, Carlsbad, CA) was used to amplify plasmid DNA using standard procedures (Ausubel et al., 2003). Aspergillus parasiticus NR1 (niaD) was used as the recipient strain for all fungal transformations (Homg et al., 1990). A. parasiticus SUI is a wild type strain. A. parasiticus AF 810 is an qflRI knock-out strain (Cary et al., 2002). To measure solid culture GUS activity, YES agar (2% yeast extract, 6% sucrose; pH=5.8) plates were used. To measure liquid culture GUS activity, GMS (Buchanan and Lewis, 1984) liquid media was used as previously described (Miller et al., 2003a). To extract protein for electorphoretic mobility shift assays, GMS (Buchanan and Lewis, 1984) liquid media was used. Plasmid Constructs (i) pNANG-3. The construction of pNANG-3 was previously described (Miller et al., 2003a). pNANG-3 contains the niaD selectable marker, GUS (uia'A) reporter enzyme and nor-l terminator. Appropriate promoter pieces, amplified by PCR using primers with NotI and PacI tails, can be directionally cloned into pNANG—3 resulting in pBNG-n 121 (where n designates the size of the promoter in base pairs). pAPGUSNN-B (Chiou et al., 2002) was used as a template for PCR amplification of all nor-l promoter fragments. pBNG-n will have the PCR amplified promoter driving the transcripiton of the GUS reporter gene. (ii) pBNG332 and pBNG332TATAmut. The polyadenylation site for ORF 3 (Miller et al., 2000a) is 332 bp upstream from the transcriptional start site of nor-1. A 332 nor-1 promoter piece was amplified by PCR with appropriate primers with PacI (J L267: 5'-GTTAATTAAGTCG AGCGGACATGGCCACG-3') and Not] tails (JL186: 5'-TCGCGGCCGCTAAGTGATCCATTC ATTATGTC-3'). For pBNG332TATAmut, the TATA box was mutated to a Pme] site (5'-ATATATAG-3' to 5'-GTTTAAAC-3') in the context of the 332 bp nor-1 promoter. The 332 bp nor-1 promoter was divided into two PCR fragments (Pac/Pme and Not/Pme) that joined at the TATA box. The primers used for the Pac/Pme fragment were .1 L267 and J L414 (5'-ATGTTTAA ACTGGGATAC GATCATGGGTC-3'). The primers used for the Not/Pme fragment were I L186 and IL 415 (5'-GGGTTTAAACGGCGGTG TGTTGGTCG-3'). After digestion with the appropriate restriction endonucleases, a three piece ligation was performed with pNANG-3, Pac/Pme fragment and Not/Pme fragments. The sequence of the nor-I promoter in both pBNG332 and pBNG332TATAmut was verified by sequence analysis. (iii) pBNG332, pBNG298, pBNG268, pBNG238 and pBNG210. To generate the nor-l promoter deletion series, different upstream PCR primers with PacI tails were used with the same downstream PCR primer (JL186) that contained a Not] tail. The upstream primers used were: I L267 for pBNG332, JL41 l (5'-CCTTAATTAAACTGCTATGGTG ACCTATTG-3') for pBNG298, J L412 (5'-CATTAATTAACCACATAGGCTACTCAA AAT-3') for pBNG268, JL413 (5'-GGTTAATTAAAGATCTCTGCTATTAAGTCGG-3') 122 for pBNG238 and JL302 (5'-C CCTTAATTAATAGCGTGCTGGATGCGCGAA-3') for pBNG210. (iv) pBNGnoerut. For pBNGnoerut, the -210 to -238 region was substituted (5'-AG ATCTCTGCTATTAAGTCGGTGATTAG-3' to 5'-GTATAAGAAGTTTGTGA TGGGATTCGT C-3') in the context of the 332 bp nor-1 promoter. The 332 bp nor-1 promoter was divided into two PCR fragments (Pac/210 and Not/238) that joined at -224. The primers used for the Pac/210 fragment were JL267 and JL613 (5'-CAAACTTCTTA TACGCTCATGTCAATTTTGAG-3'). The primers used for the Not/238 fragment were I L186 and IL 612 (5'-TGATGGGATTCGTCCGTGCTGGATGCGC-3'). JL612 and I L613 do not include a restriction endonuclease tail. After digestion with the apprOpriate restriction endonucleases, a three piece ligation was performed with pNANG-3, Pac/210 fragment and Not/238 fragment. The sequence of the nor-I promoter in pBNG332noerut was verified by sequence analysis. Generation and Screening of Transformants Transformation Transformation of A. parasiticus protoplasts was performed as described by Homg et a1. (1990). 2-4 pg of DNA was used with 107 protoplasts resulting in approximately 100 transformants. Site of Integration PCR Assay A rapid DNA extraction procedure and PCR analysis were performed as described by Chiou (Chiou et al., 2002). Following PCR, a 2.0 kb DNA fragment is diagnostic for 3' integration. 123 Southern Hybridization Genomic DNA was purified from A. parasiticus cultures shaken for 48 h in 100 m1 YES (2% yeast extract, 6% sucrose and pH=5.8) liquid medium at 29°C with five, 6 mm glass beads (Skory et al., 1993) and subjected to standard agarose gel electrophoresis (Ausubel et al., 2003). Southern hybridization analysis was performed according to standard procedures (Ausubel et al., 2003). 2.5 pg of genomic DNA was digested with Seal and probed with a 900 bp ClaI fragment isolated from the nor-1 terminator region of pAPGUSNN-B (Chiou et al., 2002). Digestion of DNA from the recipient strain NR-l generates a 3.0 kb restriction fragment while a 3' integrant results in 3.7 and 4.0 kb restriction fragments. GUS Reporter Assays Solid culture GUS Assay Autoclaved Nytran SPC (Schleicher & Schuell, Keene NH) membranes were first placed on top of YES (2% yeast extract, 6% sucrose, 1.5% agar and pH=5.8) agar plates. Transformants were then transferred via sterile toothpick to the Nytran membrane. After incubation for 46 h at 29°C in the dark, the Nytran filters were removed, frozen in liquid nitrogen and thawed at room temperature (2 repetitions) and then incubated for up to 24 h with GUS substrate solution that includes 0.04% of the colorimetric GUS substrate X- Glu (5-bromo-4-chloro-3-indolyl B-D—glucuronide) in GUS reaction buffer (60 mM NazHPO4, 40 mM NaHzPO4, 10 mM KCl, 1 mM MgSO,,, 0.07% B-mercaptoethanol) and visually evaluated. 124 Liquid Culture GUS Assay Two confirmed A. parasiticus 3' transformants containing reporter constructs were grown in duplicate for 48 h at 29°C in the dark with shaking in 100 m1 of GMS (Buchanan and Lewis, 1984) liquid medium in a 250 ml flask with five 6 mm glass beads. Mycelia were collected by filtration through Miracloth (Calbiochem, La Jolla CA) and ground in a mortar with a pestle under liquid N2. Approximately 200 mg of ground mycelia was transferred to a 1.5 mL microcentrafuge tube and kept on ice. After all the samples had been collected, 500 mL of GUS lysis buffer (50 mM NaHzPO4 [pH=7], 10 mM EDTA, 0.1% Triton X-100, 0.1% SDS, 0.07% B-mercaptoethanol and 25 mg/ml PMSF [phenylmethylsuflonyl fluoride]) was added. The samples were vortexed for 15 s and then centrifuged for 10 min at 10,000 X g at 4°C. The supematent was withdrawn and placed in a new tube. The protein concentration was determined by Bio- Rad protein assay (Bio-Rad, Hercules CA). GUS activity was determined with 100 pg of each sample incubated at 30°C with 145 pg of the GUS substrate 4-methylumbelliferyl B-D-glucoside (MUG) in a final volume of 200 p1. At 0, 10 and 30 min, 800 pl of stop solution (200 mM NaZCO3) was added. After stopping the GUS reaction, 200 pl of sample was loaded into a microtiter plate which was analyzed using a Cytofluor II fluorometer (Biosearch Co., Bedford MA). Excitation and emission wavelengths were 360 nm and 460 nm, respectively. To evaluate the fluorometer results, a standard curve of 4-methy1umbelliferone (10 nM to 600 nM) in GUS lysis buffer was also read in the fluorometer. GUS activity was reported as pmols 4-methylumbelliferone / minute / mg protein. 125 in vitro DNA Binding Assays Protein Extraction Cellular protein was extracted from cultures of A. parasiticus SUI using modifications of the methods of Peters and Perez-Esteban (Peters and Caddick, 1994 and Perez-Esteban et al., 1993). 1 liter flasks containing 10 6mm glass beads and 500 mL medium (GMSIO or PMSm) was inoculated with 1 X 108 spores. The cultures were incubated for 48 hours at 29°C in the dark with shaking at 150 rpm. The mycelia was filtered through miracloth (Calbiochem, La Jolla CA), washed with cold, sterile water, frozen with liquid nitrogen and stored at -80°C. Frozen mycelia was ground using mortar and pestle with liquid nitrogen and transferred to a tared 125 ml flask. 5 ml of lysis buffer (25 mM Hepes—KOH (pH 7.5), 50 mM KCl, 5 mM MgC12, 0.1 mM EDTA, 10% glycerol, 0.5 mM DTT and 1 mM PMSF) per gram of ground mycelia was added to the flask. In addition, 1 ml of protease inhibitor cocktail (Sigma, St. Louis MO - product # P8215) was added to the flask per gram of mycelia. After stirring on ice for 15 minutes, saturated ammonium sulfate was slowly added to a final concentration of 10%. The suspension was then stirred on ice for 15 minutes and then set idle for 15 minutes on ice. Cell debris was then pelleted at 100,000 x g (30 minute spin at 4°C) and the volume of the supematent, determined using a graduated cylinder, was transferred to a 50 ml flask. Then solid ammonium sulfate was added slowly over 1.5 hours to raise the concentration from 10% to 70%. The ammonium sulfate addition was done while stirring on ice. After all ammonium sulfate was added, the flask was incubated for 30 minutes on ice without stirring. The protein was pelleted at 10,000 X g (20 minutes at 4°C). The pellet was resuspended in dialysis buffer (15% glycerol, 15 mM Hepes—KOH (pH 7.9), 100 mM KC], 1 mM EDTA, 2 mM DTT, 0.5 mM PMSF and protease inhibitor cocktail (1 ml per 126 20 grams of mycelia) and dialyzed 2X in 2 liters dialysis buffer using a 10K slide-a-lyzer (Pierce, Rockford IL). The protein concentration was determined using the BioRad (BioRad, Hercules CA) protein dye reagent following manufacturers instructions. The dialyzed solution was then aliquoted and stored at -80°C. Probe Generation Probes were end-labeled with YEP using Ready-to-Go Kinase following the manufacturers instructions (Amersham, Piscataway NJ). The probes and competitors used are listed in table 5.1. Electrophoretic Mobility Shift Assay (EMSA) EMSA was performed essentially as described in Current Protocols in Molecular Biology (Ausebel et al., 2003). Five percent acrylamide (80:1 acrylamidezbis- acrylamide) non-denaturing gels were used. 20 fmol of nor-1 and ver—I promoter probes were incubated for 15 minutes at 30°C with 2 pg dIdC, 7.5 pg BSA and competitor (if desired) with 32 pg protein extract (added last). The entire binding reaction volume was 25 pl which consisted of 20 pl of dialysis buffer in order to keep the glycerol concentration of the binding reaction greater than 10%. RESULTS Generation and screening of A. parasiticus transformants In order to identify cis-acting sites necessary for nor-l transcriptional activation in A. parasiticus; several nor-l ::GUS reporter plasmids were constructed (Figure 5.1 and 127 Table 5.1. Oligonucleotides used for electrophoretic mobility shift assays Oligo Locationa Sequence" 206/244 -244 to - ATG AGC AGA TCT CTG CTA TTA AGT CGG TGA 205 TTA GCG TGC T 206/244 —244 to - ATG AGC GTA TAA GAA GTT TGT GAT GGG ATT mut 205 CGT CCG TGC T CRE] +4 to +31 TTC TAA GCC GTG ACA TAA TGA ACG GAT C CRElmut +4 to +31 TTC TAA GCC GTG _T_G_A TAA T GA ACG GAT C CRE2 -258 to - TAC TCA AAA TTG ACA TGA GCA GAT CTC T 231 CRE2mut -258 to - TAC TCA AAA TTG EA TGA GCA GAT CTC T 231 a Location in relationship to the nor-1 transcriptional start site. b Mutated bases are underlined, ATG is italicized in CREl/CRElmut and the possible cis-acting sites are bold in CREl and CRE2. 5.2). We defined the upstream border of the nor-1 promoter as the polyadenylation site for ORF 3 (Miller et al., 2003a), located 332 bp upstream from the primary transcriptional start site of nor-I. Transformants canying a substitution of the putative TATA box (5'- ATATATAG-3' to 5'—GTTTAAAC-3') in the context of the 332 bp nor-1 promoter were used to test TATA box function in the nor-I promoter. Deletion analysis generated clones with 332, 298, 268, 238 and 210 bp nor-1 promoter fragments carried on nor- ] ::GUS reporter constructs. GUS activity in transformants that carried these plasmids was analyzed to localize a candidate cis-acting site. In addition, transformants carrying a replacement of the norL region (-210 to -238) were also analyzed. Each nor-1 ::GUS reporter construct could theoretically integrate into the A. parasiticus chromosome by homologous recombination at three independent sites: niaD, 128 GUS Activity Sample (pmol/min/mg) 332 5.4 d: 2.2 332TATAmut ND Figure 5.1. Importance of the TATA box in the nor-1 promoter. (A) A liquid culture GUS assay analysis was performed on two different 3' integrants from each nor-1::GUS reporter construct (332 and 332TATAmut) grown in duplicate. GUS activity is reported as the average of 4 values in pmol/min/mg i the standard deviation. (B) Solid culture GUS assays were performed on different 3' integrants from each nor-1 ::GUS reporter construct on 46 h YES agar colonies. Colonies analyzed are: A 332-1, B 332-2, C 332TATAmut-l, D 332TATAmut-2, and E NR1. 129 ‘4‘ «any Figure 5.2. Identification of norL cis-acting site. (A) A liquid culture GUS assay was performed on two different 3' integrants from each nor-I ::GUS reporter construct grown in duplicate. The GUS activity is reported as the average of4 values in pmol/min/mg i the standard deviation. (B) Solid culture GUS assays were performed on different 3' integrants from each nor-l ::GUS reporter construct on 46 h YES agar colonies. Colonies analyzed are: A 332-l. B 298-1, C 268-1, D 238-1, E 210-1, G 332noerut-1 and H NR1. 130 Sample 332 298 268 238 210 332noerut Figure 5.2 norL norL norL GUS Activity (pmol/min/mg) 5.4 i 2.2 13.0i4.1 1.0i0.6 7.2 i 5.8 ND 2.1 d: 1.6 5' nor-1 (nor-I promoter) and 3' nor-l (nor-I terminator). Screening for clones in which integration occurred at the 3' nor-1 locus was essential for two reasons: 1) niaD integrants have been shown to have lower transcriptional activity than aflatoxin Cluster integrants (Liang et al., 1997 and Chiou et al., 2002); and 2) 5' integrants result in the chromosomal nor-1 promoter fused to the GUS gene and the plasmid nor-1 promoter fused to the chromosomal nor-I gene. Transformants from the nor-l ::GUS reporter constructs were screened initially by a rapid PCR assay specific for nor-1, 3' integration (Chiou et al., 2002). A 3' nor-1 integrant generates a 2.1 kb DNA fragment in the rapid PCR integration assay regardless of the size of the nor-1 promoter in the reporter plasmid. The percentage of transformants that had a 2.1 kb band with the rapid assay varied from experiment to experiment ranging from 2 to 5% (data not shown). While the PCR assay could identify 3' integrants, it could not identify multiple integrants (multiple 3' integrations and/or additional non-3' integrations). Consequently, Southern hybridization analysis was used to confirm PCR integration assay results for each reporter construct. Disappearance of a 3.0 kb DNA fragment in the recipient strain, NR1, and the appearance of 3.7 and 4.0 DNA fragments was diagnostic for 3' nor-1 integration (data not shown). Two confirmed 3' nor-1 integrants for each nor-1::GUS reporter construct were used for solid culture and liquid culture reporter analysis. In order to determine if the TATA box was necessary for nor-l transcriptional activation in vivo, fungal isolates carrying the 332 and 332TATAmut promoter fragments in nor-1::GUS constructs were tested for GUS activity using liquid culture (Figure 5.1A) and solid culture (Figure 5.1B) activity assays. A. parasiticus isolates carrying the 332 nor-1::GUS construct (includes TATA) converted 5.4 pmol/min/mg while the isolates carrying the 332TATAmut nor-1::GUS transformant (TATA mutated) had no detectable 132 GUS activity (Figure 5.1A). In agreement with these liquid culture data, isolates carrying the 332 nor-] ::GUS construct (includes TATA) showed clearly detectable GUS activity (blue color within colonies) while the 332TATAmut (TATA mutated) nor-1::GUS transformants displayed no detectable GUS activity. These data confirm that the TATA box is necessary for nor-1 transcriptional activation in vivo. Previously, a potential cis-acting site in the nor-1 promoter was localized between -332 and -76 (Miller et al., 2003a). Isolates carrying nor-[::GUS constructs with the 210 and 76 promoter fragments had no detectable liquid culture or solid culture GUS activity (data not shown). Fungal isolates carrying nor-1 ::GUS constructs with 332, 298, 268, 238 and 210 promoter fragments were analyzed to localize the potential cis-acting ‘site located between -332 and -210 in the nor-1 promoter. Isolates carrying nor-I ::GUS constructs with the 332, 298, 268 and 238 promoter fragments all had detectable liquid culture GUS activity (Figure 5.2A) and solid culture GUS activity whereas the 210 promoter fragment had reproducibly less activity (Figure 5.28). To verify the in vivo significance of the 210 to 238 region (norL), isolates carrying a nor-[::GUS construct with norL replaced in the context of the 332 bp nor-1 promoter fragment was used. Replacement of norL resulted a 2.5 fold reduction in liquid culture GUS activity (Figure 5.2A). In addition, replacement of norL also resulted in a decrease in solid culture GUS activity (Figure 5.2B). While a functional norL site is not sufficient for nor-l transcriptional activation, it is necessary for maximum nor-1 transcriptional activation in the context of the 332 promoter. in vitro analysis nor-1 promoter elements To test for protein affinity for specific DNA sites in the nor-I promoter, 133 electrophoretic mobility shift assays (EMSA) were used. The probes and competitors used for EMSA are described in Table 1. Total cellular protein used for EMSA was extracted from two different strains: SUI (wild type) and AF SIO (aflR knock-out). Extracts from each strain were prepared twice on two separate occasions. Comparison of the shifted complexes generated with each extract provided evidence regarding the role of AflR in the activity. The 206/244 oligo spans the norL cis-acting site that was identified using nor- ] ::GUS reporter strains in vivo (Figure 5 .2) and was used as a probe for EMSA (Figure 5.3). The 206/244mut oligo has the same substitution in the 210 to 238 region as the nor- ] : :GUS reporter construct 332noerut. The substitution resulted in changes in 2 l of the 28 nucleotides while maintaining the same GC ratio. With 206/244 as probe, two shifted complexes were identified with an SUI protein extract. A 250 fold excess of unlabeled 206/244 as a competitor decreased the intensity of Norpr while a 250 fold excess of unlabeled 206/244mut was a less effective competitor for Norpr. Complex A appears to be due to non-specific binding because both 206/244 and 206/244mut were effective competitors. No specific protein complexes could be identified with the AFSIO protein extract and the 206/244 probe. In addition, no specific shifted complexes were identified with 206/244mut as a probe with either SUI or AF 810 protein extracts (data not shown). Norpr binds specifically to the nor-L cis-acting site and requires AflR either directly or indirectly for binding function. A potential cis-acting site, CREI, that may be involved in the upregulation of aflatoxin synthesis in response to CAMP, has been identified in the nor-1 promoter (Dr. Ludimila Roze, personal communication). A similar site was identified upstream from 134 , SUI AFSIO Protein —> — — 206/ 206/ Competitor —> - - 206/ 244 _ 206/ 244 244 mut 244 mut Norpr —> .. Figure 5.3. EMSA with a 206/244 oligo. 20 fmol of the 206/244 oligonucleotide was used as a probe with 32 pg of protein extract from SUI or AFSIO. Norpr complex and complex A are indicated with arrows. 135 C REI and was named CRE2. Both CREI and CRE2 contain the core sequence: TGACAT a/g A. With CREl as probe, one shifted complex of similar migration was identified with both SUI and AF S10 protein extracts (Figure 5.4). We have designated the protein responsible for the complex CREbp. To test for specificity of CREbp binding, competitions were performed with 250 fold excess of unlabeled CREI, CRElmut and CRE2 oligos. For both SUI and AFSIO protein extracts, CREl was an effective competitor and CRElmut was not. With the SUI extract, CRE2 appears to only marginally compete for CREbp binding. No specific complexes were identifed with CRE lmut, CRE2, and CRE2mut probes with either SUI or AF 810 protein extracts (data not shown). CREbp binds specifically to CREI and appears to require AflR for maximum complex formation. DISCUSSION The role of TATA boxes and TATA binding protein (TBP) in expression of aflatoxin biosynthesis genes has largely been ignored. While candidate TATA boxes have been previously identified, we demonstrated that the nor-I promoter TATA box is necessary to detect GUS activity. Transformants canying the 332TATAmutn0r—1zzGUS vector had no detectable activity despite containing functional AflR, CREbp and Norpr binding sites. The nor-I TATA box needs to be present for these other cis-acting sites to be functional in terms of GUS activity. Currently, only five aflatoxin biosynthetic genes have had their transcriptional start point experimentally determined (Figure 5.5). In Figure 5.5, the sixty bases upstream from the major transcriptional start point in each of these genes are displayed. 136 - SUI AFSIO Protem —> - — CREI CREl Competitor ‘ - - CREI mut CRE2 — CRE] mut CRE2 , ,' ; n. I ~ 3.». CREbp ’ M” m ‘11th mu. m I mm“ Figure 5.4. EMSA with a CREl oligo. 20 frnol of CREI oligonucleotide was used as probe with 32 pg of protein extract from SUI or AFSIO. CREbp complex is indicated with an arrow. 137 -60 -50 -40 -30 -20 -10 +1 [\_ CACCGCCATA TATAGTGGGA TACGATCATG GGTCTTTGGT GGTTTCAACA TTTCTTGAGT A TATCTAATAT CAATTTATTA TCTTAGACCT CCTCATGCAA CGGTGCTTCC TTCTGCCAGT G CCGAGGAAAG ATTTGTTTGG TGGCCAACCA TCCATAGCTG CGTATATATG TACTACATGC C (: .ATCTCGAAGT GTAGTTTTCA AATACTGATA TAGCTTCCTA TAGCTCCCTCG GGGCGGACC T E3 GGGCCGGCTA CTCTCCCGGA GCAAGCCTTC ACCTTGTGTG TTTTCTTTCC CGCTTTCAAT T Figure 5.5. Location of putative TATA boxes in A. parasiticus aflatoxin biosynthesis gene promoters. The experimentally determined major transcriptional start point is shown at +1. (A) nor-1 (Trail et al., 1994) (B) avnA (Cary et al., 2000) (C) pksA (Ehrlich et al., 2002) (D) ver—l (Skory et al., 1992) (E) qflR (Ehrlich et al., 1999a). 138 The location of putative TATA boxes in avnA (Cary et al., 2000; Figure 5.5B), pksA (Ehrlich et al., 2002; Figure 5.5C) and ver-I (Skory et al., 1992; Figure 5.5D) are -40, -29 and —13 respectively. While the functionality of the avnA, pksA and ver-I TATA boxes is unknown, we hypothesize that the structural genes in the aflatoxin biosynthesis pathway contain functional TATA boxes. To test this hypothesis, the transcriptional start point needs to be identified in more aflatoxin genes and the TATA boxes need to be tested for function in the context of a wild type promoter. Substitution of a suspected cis-acting site in the context of a wild type promoter is a more rigorous test of function than a deletion. While the four structural genes described above have putative TATA boxes in their promoters, the first 60 bp of the aflR promoter does not contain a sequence resembling a TATA box (Ehrlich et al., 1999a; Figure 5.5E). AflR is an aflatoxin biosynthetic pathway regulator and perhaps is regulated differently than the aflatoxin structural genes. As additional aflatoxin promoters are mapped, it will be interesting to see if AflR is unique in its lack of a TATA box. The TATA binding protein (TBP) binds to the TATA box in eukaryotes. The TBP gene has been cloned from A. nidulans (Kucharski and Bartnik, 1997). The A. nidulans TBP promoter has consensus cis-acting sites for CreA (carbon repression), AreA (nitrogen repression) and AbaA (conidiophore development) (Kucharski and Bartnik, 1997). The levels of A. nidulans TBP mRNA varied several fold under diverse growth conditions that are consistant with the presence of CreA and AreA sites in the TBP promoter (Kucharski and Bartnik, 1997). Previously, a potential cis-acting site was localized between -332 and -76 bp from the nor-1 transcriptional start point (Miller et al., 2003a). Analysis of different sized\ 139 nor-1 promoters in nor-1::GUS strains further localized the potential cis-acting site to between -210 and -238. The in vivo significance of the 210 to 238 region (norL) was verified by analyzing isolates carrying a nor—1::GUS construct with norL replaced in the context of the 332 bp nor-1 promoter fragment. While a functional norL site is not sufficient for nor-1 transcriptional activation, it is necessary for maximum nor-1 transcriptional activation in the context of the 332 bp promoter. Using EMSA, we demonstrated that Norpr binds specifically to norL. In addition, Norpr is dependent on AflR for binding activity, either directly or indirectly. The identity of Norpr and whether other aflatoxin genes contain Norpr cis-acting sites is unknown. Future work is focused on further defining the cis-acting site and cloning norpr. Another possible cis-acting site was recently identified, CREl (Dr. Ludmilla Roze, personal communication). Preliminary experiments demonstrated that CREbp binding to CREl and nor-1 transcriptional activation are increased in the presence of CAMP in A. parasiticus solid cultures (Dr. Ludmilla Roze, personal communication). In addition, CAMP addition at high levels decreased protein kinase A (PKA) activity which coincided with decreased phosphorylation of CREbp (Dr. Ludmilla Roze, personal communication). We demonstrated that CREbp binds specifically to CREI in A. parasiticus liquid cultures and that AflR is necessary for maximum CREbp binding activity. The gene for CREbp has not been identified but efforts are underway to purify CREbp and in order to generate protein sequence data (Dr. Ludmilla Roze, personal communication). Interestingly, the ver—l, pksA, and avnA promoters do not have an identical match for CREl (TGACATAA). Nor-l catalyzes an early step in the pathway and may be regulated differently than other aflatoxin structural genes. In addition, 140 further analysis of CREl may define the core CREl more specifically which may help identify CREbp cis-acting sites in other aflatoxin promoters. By combining all the information regarding AflR, Norpr and CREbp, a model for nor-I transcriptional activation emerges (Figure 5.6). AflR cis-acting sites are necessary for all aflatoxin genes analyzed including nor-1 (Miller et al., 2003a), pksA (Ehrlich et al. 2002) and avnA (Cary et al., 2000). In addition, an AflR site in the aflR promoter is necessary for aflR transcription (Ehrlich eta1., 1999a). Norpr appears to require functional AflR for activity based on in vitro experiments but the mechanism is not known. Our experiments suggest that Norpr is a transcriptional activator for nor-1. Confirmation of Norpr function requires additional testing. CREbp appears to be only marginally affected by functional AflR in in vitro experiments. CREbp bound to CREl under aflatoxin inducing conditions yet there is evidence that CREbp also binds under non-inducing conditions (Dr. Ludmila Roze, personal communication). Consequently, we can not determine if CREbp is an activator and/or repressor of nor-1 transcription. Confirmation of CREbp function also requires additional testing. Protein kinase A (PKA) is activated by CAMP. Preliminary evidence suggests that phosphorylation of AflR by PKA results in reduced AflR function while phosphorylation of CREbp by PKA resulted in increased CREbp binding to CRE (Dr. Ludmila Roze, personal communication). Additional experiments are needed to clarify how phosphorylation of C REbp and AflR by PKA affect their function. This study is significant because it provides the first direct evidence for the existence of transcription factors other than AflR in aflatoxin gene expression. Future I41 9 III... CREbp Figure 5.6. Proposed model for nor-1 transcriptional activation. Arrows indicate positive interactions, blocked lines indicate negative interactions and lines with no block or arrow indicate unknown interaction. Definite interactions are represented by solid lines while inconclusive interactions are represented by dashed lines. 142 identification of the CREbp and norpr genes will help verify their role in nor-1 transcriptional activation. In addition, how CREbp and norpr are regulated and what signals they respond to will aid our understanding of nor-I trancriptional regulation. ACKNOWLEDGMENTS All experiments described in Chapter 5 were performed by Michael Miller. The future publication status of this chapter is undecided. 143 CHAPTER 6 Future Studies The studies described in this dissertation have provided a more detailed understanding of nor-1 transcriptional regulation. However, several questions about nor- ] transcriptional regulation remains. Confirmation of the functionality of CREI and norL cis-acting sites will require the identification of their respective binding proteins (CREbp and norpr respectively). Once the binding proteins are cloned, experiments could be designed to explore how CREbp, norpr and AflR all interact to regulate nor-1 transcription. In addition, we do not know if the CRE and norL cis-acting sites are unique to nor-1. Perhaps all the aflatoxin structural genes are regulated in a similar manner. However, the regulation of nor-I may have more levels of control than other aflatoxin structural genes because it catalyzes an early or critical step for aflatoxin biosynthesis. The function ofAflR2, an AflR binding site located approximately 1200 bp from nor-1 transcriptional start site, in nor-1 transcriptional activation is currently unclear because of the ORF 3 gene located between AflR2 and nor-1. A nor-l reporter plasmid that includes AflR2 in the nor-1 promoter may also include a transcriptionally competent and functional copy of ORF 3. Integration of this reporter plasmid creates an extra copy of ORF3. ORF3 has no known function and sequence analysis has not provided any clear clues. However, an increased concentration of ORF3 protein may explain the higher nor-1 transcriptional activation, especially if ORF 3 is a transcriptional activator (C REbp or norpr?). An ORF 3 knock-out would provide preliminary evidence for 144 ORF 3 function. In addition, a nor-1 reporter plasmid that includes AflR2 but has a missense mutation in ORF3 would determine if AflR binding to AflR2 can directly activate nor-1 transcription. In filamentous fungi, the clustering of genes is a common feature of several metabolic pathways including mycotoxin biosynthesis. Others have postulated that the linkage of metabolic pathway genes provides a means to coordinately regulate pathway gene transcription, possibly via enhancers. Integration of a nor-1 reporter plasmid outside of the aflatoxin gene cluster results in significantly lower nor-1 transcriptional activation than integration within the aflatoxin gene cluster. Mechanisms for gene- cluster-dependent regulation in fungi are currently unknown. The aflatoxin gene cluster is an ideal model for future studies of cluster-dependent regulation for several reasons: 1) aflatoxin cluster is well characterized, 2) some strains of A. parasiticus (including NR1) have portions of the aflatoxin cluster duplicated elsewhere in their genome, and 3) several tools are already available including ver-I and nor-1 reporter strains. Determination as to whether the duplicated region of the aflatoxin gene cluster also has cluster—dependent regulation would help localize possible enhancers. The large spacer region at one end (the other end of aflatoxin cluster is currently undefined) of the aflatoxin gene cluster is an attractive target for future studies on cluster dependent regulation as well. Mechanistic analysis of aflatoxin gene-cluster-dependent regulation may help studies with other gene clusters as well. 145 LIST OF REFERENCES Abdollahi, A. and R. L. Buchanan. 19813. Regulation of aflatoxin biosynthesis: characterization of glucose as an apparent inducer of aflatoxin production. J. Food Sci. 46: 143-146. Abdollahi, A. and R. L. Buchanan. 1981b. Regulation of aflatoxin biosynthesis: induction of aflatoxin production by various carbohydrates. J. Food Sci. 46:633-635. Adams, T. H., M. T. Boylan, and W. E. Timberlake. 1988. brlA is necessary and sufficient to direct conidiophore development in Aspergillus nidulans. Cell 54:353-362. Aharonowitz, Y., C. Cohen, and J. F. Martin. 1992. Penicillin and cephalosporin biosynthetic genes: structure, organization, regulation, and evolution. Annu. Rev. Microbiol. 46:461-495. Annan, K. Secretary General tells special event on poverty eradication, "best hope" for least developed countries would be new round of global trade negotiations. Press Release G/SM/7802 Dev/2311 . 5-14-2001. Asao, T., G. Buchi, M. M. Abdel-Kader, S. B. Chang, E. L. Wick, and G. N. Wogan. 1963. Aflatoxins B and G. J. Am. Chem. Soc. 85:1706-1707. Ausubel, F. M., R. Brent, R. E. Kingston, D. D. Moore, J. A. Seidman, J. A. Smith, and K. Struhl. 2003. Current Protocols in Molecular Biology. John Wiley & Sons, New York. Bennett, J. W. and K. E. Papa. 1988. The aflatoxigenic Aspergillus spp. Adv. Plant Pathol 62263-280. Bennett, J. W., P. L. Rubin, and P. N. Chen. 1979. Influence of trace elements and nitrogen sources on versicolorin production by a mutant strain of Aspergillus parasiticus. Mycopathologia 69: 161-166. Bhatnagar, D., T. E. Cleveland, and P. J. Cotty. 1994. Mycological aspects of aflatoxin formation, p. 327-346. In D. L. Eaton and J. D. Groopman (eds.), The Toxicology of Aflatoxins, Academic Press, San Diego. Bhatnagar, D., T. E. Cleveland, and D. G. Kingston. 1991. Enzymological evidence for separate pathways for aflatoxin B1 and B2 biosynthesis. Biochem. (Mosc). 3024343- 4350. Blount, W. P. 1961. Turkey "X" disease. J. Brit. Turkey Fed. 9:52-54. 146 Bradshaw, R. E., D. Bhatnagar, R. J. Ganley, C. J. Gillman, B. J. Monahan, and J. M. Seconi. 2002. Dothistroma pini, a forest pathogen, contains homologs of aflatoxin biosynthetic pathway genes. Appl. Environ. Microbiol. 68:2885-2892. Brown, D. W., S. P. McCormick, N. J. Alexander, R. H. Proctor, and A. E. Desjardins. 2001. A genetic and biochemical approach to study trichothecene diversity in F usarium sporotrichioides and F usarium graminearum. Fungal. Genet. Biol. 32: 121- 133. Brown, D. W., J. H. Yu, H. S. Kelkar, M. Fernandes, T. C. Nesbitt, N. P. Keller, T. H. Adams, and T. J. Leonard. 1996. Twenty-five coregulated transcripts define a sterigmatocystin gene cluster in Aspergillus nidulans. Proc. Natl. Acad. Sci. USA. 93:1418-1422. Buchanan, R. L. and D. F. Lewis. 1984. Regulation of aflatoxin biosynthesis: effect of glucose on activities of various glycolytic enzymes. Appl. Environ. Microbiol. 48:306- 310. Buchanan, R. L. and H. G. Stahl. 1984. Ability of various carbon sources to induce and support aflatoxin biosynthesis by Aspergillus parasiticus. J. Food Sci. 6:271-279. Cary, J. W., J. M. Dyer, K. C. Ehrlich, M. S. Wright, S. H. Liang, and J. E. Linz. 2002. Molecular and functional characterization of a second copy of the aflatoxin regulatory gene, aflR-Z, from Aspergillus parasiticus. Biochim. Biophys. Acta 1576:316- 323. Cary, J. W., B. G. Montalbano, and K. C. Ehrlich. 2000. Promoter elements involved in the expression of the Aspergillus parasiticus aflatoxin biosynthesis pathway gene avnA. Biochim. Biophys. Acta 1491:7-12. Chang, P. K., J. W. Bennett, and P. J. Cotty. 2001. Association of aflatoxin biosynthesis and sclerotial development in Aspergillus parasiticus. Mycopathologia 153:41-48. Chang, P. K., J. W. Cary, D. Bhatnagar, T. E. Cleveland, J. W. Bennett, J. E. Linz, C. P. Woloshuk, and G. A. Payne. 1993. Cloning of the Aspergillus parasiticus apa-Z gene associated with the regulation of aflatoxin biosynthesis. Appl. Environ. Microbiol. 59:3273-3279. Chang, P. K., K. C. Ehrlich, J. E. Linz, D. Bhatnagar, T. E. Cleveland, and J. W. Bennett. 1996. Characterization of the Aspergillus parasiticus niaD and niiA gene cluster. Curr. Genet. 30:68-75. Chang, P. K., K. C. Ehrlich, J. Yu, D. Bhatnagar, and T. E. Cleveland. 1995. Increased expression of Aspergillus parasiticus aflR, encoding a sequence-specific DNA- binding protein, relieves nitrate inhibition of aflatoxin biosynthesis. Appl. Environ. Microbiol. 61:2372-2377. I47 Chang, P. K., C. D. Skory, and J. E. Linz. 1992. Cloning of a gene associated with aflatoxin B1 biosynthesis in Aspergillus parasiticus. Curr. Genet. 21:231-233. Chang, P. K. and J. Yu. 2002. Characterization of a partial duplication of the aflatoxin gene cluster in Aspergillus parasiticus ATCC 56775. Appl. Microbiol. Biotechnol. 58:632-636. Chang, P. K., J. Yu, D. Bhatnagar, and T. E. Cleveland. 1999. The carboxy-terminal portion of the aflatoxin pathway regulatory protein AFLR of Aspergillus parasiticus activates GALl ::lacZ gene expression in Saccharomyces cerevisiae. Appl. Environ. Microbiol. 65:2508-2512. Chang, P. K., J. Yu, D. Bhatnagar, and T. E. Cleveland. 2000. Characterization of the Aspergillus parasiticus major nitrogen regulatory gene, areA. Biochim. Biophys. Acta 1491:263-266. Chen, L., S. P. McCormick, and T. M. Hohn. 2000. Altered regulation of 15- acetyldeoxynivalenol production in F usarium graminearum. Appl. Environ. Microbiol. 66:2062-2065. Chiou, C. H., M. Miller, D. L. Wilson, F. Trail, and J. E. Linz. 2002. Chromosomal Location Plays a Role in Regulation of Aflatoxin Gene Expression in Aspergillus parasiticus. Appl. Environ. Microbiol. 68:306-315. Council for Agricultural Science and Technology. 2003. Mycotoxins: Risks in Plant, Animal, and Human Systems. CAST, Ames. Coupland, K. and W. G. Niehaus, Jr. 1987. Effect of nitrogen source, Zn”, and salt concentration on kojic acid and versicolorin biosynthesis by Aspergillus parasiticus. Exp. Mycol. 11:206-213. Desjardins, A. E., R. D. Plattner, and R. H. Proctor. 1996. Linkage among genes responsible for fumonisin biosynthesis in Gibberellafujikuroi mating population A. Appl. Environ. Microbiol. 62 :2571-2576. Diez, B., S. Gutierrez, J. L. Barredo, P. van Solingen, L. H. van der Voort, and J. F. Martin. 1990. The cluster of penicillin biosynthetic genes. Identification and characterization of the pchB gene encoding the alpha-aminoadipyl- cysteinyl-valine synthetase and linkage to the pch and penDE genes. J. Biol. Chem. 265: 16358-16365. Dvorackova, I. 1990. Aflatoxin and Human Health. CRC Press, Boca Raton. Ehrlich, K. C., J. W. Cary, and B. G. Montalbano. 1999a. Characterization of the promoter for the gene encoding the aflatoxin biosynthetic pathway regulatory protein AFLR. Biochim. Biophys. Acta 1444:412-417. 148 Ehrlich, K. C., B. G. Montalbano, and J. W. Cary. 1999b. Binding of the C6-zinc cluster protein, AFLR, to the promoters of aflatoxin pathway biosynthesis genes in Aspergillus parasiticus. Gene 230:249-257. Ehrlich, K. C., B. G. Montalbano, J. W. Cary, and P. J. Cotty. 2002. Promoter elements in the aflatoxin pathway polyketide synthase gene. Biochim. Biophys. Acta 1576: 171-175. F ailla, L. J., D. Lynn, and W. G. Niehaus, Jr. 1986. Correlation of Zn2+ content with aflatoxin content of corn. Appl. Environ. Microbiol. 52:73-74. F eng, G. H., F. S. Chu, and T. J. Leonard. 1992. Molecular cloning of genes related to aflatoxin biosynthesis by differential screening. Appl. Environ. Microbiol. 58:455-460. Fernandes, M., N. P. Keller, and T. H. Adams. 1998. Sequence-specific binding by Aspergillus nidulans AflR, a C6 zinc cluster protein regulating mycotoxin biosynthesis. Mol. Microbiol. 28: 1355-1365. Fillinger, S. and B. Felenbok. 1996. A newly identified gene cluster in Aspergillus nidulans comprises five novel genes localized in the alc region that are controlled both by the specific transactivator AlcR and the general carbon-catabolite repressor CreA. Mol. Microbiol. 20:475-488. F laherty, J. E. and G. A. Payne. 1997. Overexpression of aflR leads to upregulation of pathway gene transcription and increased aflatoxin production in Aspergillusflavus. Appl. Environ. Microbiol. 63:3995-4000. F laherty, J. E., M. A. Weaver, G. A. Payne, and C. P. Woloshuk. 1995. A beta- glucuronidase reporter gene construct for monitoring aflatoxin biosynthesis in Aspergillusflavus. Appl. Environ. Microbiol. 61:2482-2486. Grosveld, F ., G.B. van Assendelft, D.R. Greaves, and G. Kolias. 1987. Position- independent, high-level expression of the beta-globin gene in transgenic mice. Cell. 51:975-985. Hall, A. J. and C. P. Wild. 1994. Epidemiology of aflatoxin-related disease, p. 233-258. In D. L. Eaton and J. D. Groopman (eds.), The Toxicology of Aflatoxins, Academic Press, San Diego. Hamer, J. E. and W. E. Timberlake. 1987. Functional organization of the Aspergillus nidulans trpC promoter. Mol. Cell Biol. 7 A22352-2359. Hohn, T. M. and A. E. Desjardins. 1992. Isolation and gene disruption of the Tox5 gene encoding trichodiene synthase in Gibberella pulicaris. Mol. Plant Microbe Interact. 5:249-256. 149 Hohn, T. M., A. E. Desjardins, and S. P. McCormick. 1993a. Analysis of Tox5 gene expression in Gibberella pulicaris strains with different trichothecene production phenotypes. Appl. Environ. Microbiol. 59:2359-2363. Hohn, T. M., A. E. Desjardins, and S. P. McCormick. 1995. The Tri4 gene of F usarium sporotrichioides encodes a cytochrome P450 monooxygenase involved in trichothecene biosynthesis. Mol. Gen. Genet. 248295-102. Hohn, T. M., R. Krishna, and R. H. Proctor. 1999. Characterization of a transcriptional activator controlling trichothecene toxin biosynthesis. Fungal. Genet. Biol. 26:224-235. Hohn, T. M., S. P. McCormick, and A. E. Desjardins. 1993b. Evidence for a gene cluster involving trichothecene-pathway biosynthetic genes in F usarium sporotrichioides. Curr. Genet. 24:291-295. Horng, J. S., P. K. Chang, J. J. Pestka, and J. E. Linz. 1990. Development of a homologous transformation system for Aspergillus parasiticus with the gene encoding nitrate reductase. Mol. Gen. Genet. 224:294-296. Jefferson, R. A. 1987. Assaying chimeric genes in plants: the GUS gene fusion system. Plant Mol. Biol. 5:387-405. Jefferson, R. A. 1989. The GUS reporter gene system. Nature 342:837-838. Johnstone, I. L., P. C. McCabe, P. Greaves, S. J. Gurr, G. E. Cole, M. A. Brow, S. E. Unkles, A. J. Clutterbuck, J. R. Kinghorn, and M. A. Innis. 1990. Isolation and characterisation of the crnA-niiA-niaD gene cluster for nitrate assimilation in Aspergillus nidulans. Gene 90: 181-192. lKachholz, T. and A. L. Demain. 1983. Nitrate repression of averufin and aflatoxin biosynthesis Aspergillus parasiticus. J. Nat. Prod. 46:499-506. Keller, N. P. and T. H. Adams. 1995. Analysis of a mycotoxin gene cluster in Aspergillus nidulans. SAAS. Bull. Biochem. Biotechnol. 8:14-21. Keller, N. P. and T. M. Hohn. 1997. Metabolic Pathway Gene Clusters in Filamentous Fungi. Fungal. Genet. Biol. 21:17-29. Kimura, M., I. Kaneko, M. Komiyama, A. Takatsuki, H. Koshino, K. Yoneyama, and I. Yamaguchi. 1998a. Trichothecene 3-O-Acetyltransferase protects both the producing organism and transformed yeast from related mycotoxins. J. Biol. Chem. 273: 1654-1661. Kimura, M., G. Matsumoto, Y. Shingu, K. Yoneyama, and I. Yamaguchi. 1998b. The mystery of the trichothecene 3-O-acetyltransferase gene. Analysis of the region around TrilOl and characterization of its homologue from F usarium sporotrichioides. 150 FEBS Lett. 435: 163-168. Kimura, M., Y. Shingu, K. Yoneyama, and I. Yamaguchi. 1998c. Features of TrilOl, the trichothecene 3-O-acetyltransferase gene, related to the self-defense mechanism in F usarium graminearum. Biosci. Biotechnol. Biochem. 62: 1033-1036. Kinsey, J. A. and J. A. Rambosek. 1984. Transformation of Neurospora crassa with the cloned am (glutamate dehydrogenase) gene. Mol. Cell. Biol. 42117-122. IS Kucharski, R. and E. Bartnik. 1997. The TBP gene from Aspergillus nidulans-structure and expression in Saccharomyces cerevisiae. Microbiology 143 ( Pt 4): 1263-1270. Kunkel, T. A. 1985. Rapid and efficient site-specific mutagenesis without phenotypic selection. Proc. Natl. Acad. Sci. USA. 82:488-492. Labrador, M. and V.G. Corces. 2002. Setting the boundaries of chromatin domains ,9 and nuclear organization. Cell. 111:151-154. Li, Q., K.R. Peterson, X. Fang, and G. Stamatoyannopoulos. 2002. Locus control regions. Blood. l00(9):3077-3086. Liang, S. H., T. S. Wu, R. Lee, F. S. Chu, and J. E. Linz. 1997. Analysis of mechanisms regulating expression of the ver-l gene, involved in aflatoxin biosynthesis. Appl. Environ. Microbiol. 63: 1058-1065. Lin, B. H., D. Bhatnagar, and F. S. Chu. 1999. Purification and characterization of 40- kDa sterigmatocystin O- methyltransferase involved in aflatoxin biosynthesis. Nat. Toxins. 7:63-69. Lin, B. H. and F. S. Chu. 1998. Regulation of aflR and its product, AflR, associated with aflatoxin biosynthesis. Appl. Environ. Microbiol. 64:3718-3723. Luchese, R. H. and W. F. Harrigan. 1993. Biosynthesis of aflatoxin--the role of nutritional factors. J. Appl. Bacteriol. 74:5-14. Mahanti, N., D. Bhatnagar, J. W. Cary, J. Joubran, and J. E. Linz. 1996. Structure and function of fas-IA, a gene encoding a putative fatty acid synthetase directly involved in aflatoxin biosynthesis in Aspergillus parasiticus. Appl. Environ. Microbiol. 62:191- 195. McCormick, S. P., N. J. Alexander, S. E. Trapp, and T. M. Hohn. 1999. Disruption of TRIlOl , the gene encoding trichothecene 3-O- acetyltransferase, from F usarium sporotrichioides. Appl. Environ. Microbiol. 65:5252-5256. McCormick, S. P., T. M. Hahn, and A. E. Desjardins. 1996. Isolation and characterization of Tri3, a gene encoding 15-0- acetyltransferase from F usarium 151 sporotrichioides. Appl. Environ. Microbiol. 62:353-359. McLean, M. and M. F. Dutton. 1995. Cellular interactionss and metabolism of aflatoxin: an update. Pharmacol. Ther. 65: 163—192. Meyers, D. M., G. Obrian, W. L. Du, D. Bhatnagar, and G. A. Payne. 1998. Characterization of aflJ, a gene required for conversion of pathway intermediates to aflatoxin. Appl. Environ. Microbiol. 64:3713-3717. 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 SpoCl gene cluster of Aspergillus nidulans. Mol. Cell Biol. 7 :427-434. Miller, M. J., C. S. Brown-Jenco, G. OBrian, G. A. Payne, and J. E. Linz. 2003a. Role of AflR in nor-1 transcriptional activation in Aspergillus/lavas and A. parasiticus. Appl. Environ. Microbiol. Submitted. Miller, M. J. and J. E. Linz. 2003b. Identification of novel cis-acting sites in the . aflatoxin biosynthetic nor-1 promoter of Aspergillus parasiticus. In preparation. Miller, M. J., M. D. Rarick, and J. E. Linz. 2003c. Nitrogen source, carbon source and zinc concentration as tools to study the regulation of the aflatoxin biosynthetic gene nor- 1 from Aspergillus parasiticus. In preparation. Muhitch, M. J., S. P. McCormick, N. J. Alexander, and T. M. Hohn. 2000. Transgenic expression of the TRIlOl or PDR5 gene increases resistance of tobacco to the phytotoxic effects of the trichothecene 4,15-diacetoxyscirpenol. Plant Sci. 157:201-207. Payne, G. A., G. J. Nystrom, D. Bhatnagar, T. E. Cleveland, and C. P. Woloshuk. 1993. Cloning of the afl-2 gene involved in aflatoxin biosynthesis from Aspergillus flavus. Appl. Environ. Microbiol. 59:156-162. Perez-Esteban, B., M. Orejas, E. Gomez-Pardo, and M. A. Penalva. 1993. Molecular characterization of a fungal secondary metabolism promoter: transcription of the Aspergillus nidulans isopenicillin N synthetase gene is modulated by upstream negative elements. Mol. Microbiol. 92881-895. Pestka, J. J., P. K. Gaur, and F. S. Chu. 1980. Quantitation of aflatoxin BI and aflatoxin Bl antibody by an enzyme- linked immunosorbent microassay. Appl. Environ. Microbiol. 40: 1027- l 03 1. Peters, D. G. and M. X. Caddick. 1994. Direct analysis of native and chimeric GATA specific DNA binding proteins from Aspergillus nidulans. Nucleic Acids Res. 22:5164- 5172. Proctor, R. H., A. E. Desjardins, R. D. Plattner, and T. M. Hohn. 1999. A polyketide synthase gene required for biosynthesis of fumonisin mycotoxins in Gibberellafujikuroi 152 mating population A. Fungal. Genet. Biol. 27: 100-1 12. Proctor, R. H., T. M. Hohn, S. P. McCormick, and A. E. Desjardins. 1995. Tri6 encodes an unusual zinc finger protein involved in regulation of trichothecene biosynthesis in F usarium sporotrichioides. Appl. Environ. Microbiol. 61: 1923-1930. Robens, J. The costs of mycotoxin management to the USA: management of aflatoxins in the United States. APSnet Feature Article , 2-8. 2001. Roebuck, B. D. and Y. Y. Maxuitenko. 1994. Biochemical Mechanisms and Biological Implications of the Toxicity of Aflatoxins as Related to Aflatoxin Carcinogenesis, p. 27- 44. In D. L. Eaton and J. D. Groopman (eds.), The Toxicology of Aflatoxins, Academic Press, San Diego. Scholl, P., S. M. Musser, T. W. Kensler, and J. D. Groopman. 1995. Molecular biomarkers for aflatoxins and their application to human liver cancer. Pharmacogenetics 5 Spec No:Sl7l-Sl76. Seo, J. A., R. H. Proctor, and R. D. Plattner. 2001. Characterization of four clustered and coregulated genes associated with fumonisin biosynthesis in F usarium verticillioides. Fungal. Genet. Biol. 34:155-165. Skory, C. D., P. K. Chang, J. Cary, and J. E. Linz. 1992. Isolation and characterization of a gene from Aspergillus parasiticus associated with the conversion of versicolorin A to sterigmatocystin in aflatoxin biosynthesis. Appl. Environ. Microbiol. 58:3527-3537. Skory, C. D., P. K. Chang, and J. E. Linz. 1993. Regulated expression of the nor-1 and ver-l genes associated with aflatoxin biosynthesis. Appl. Environ. Microbiol. 5921642- 1646. Smela, M.E., S.S. Currier, E.A. Bailey, and J.M. Essigmann. 2001. The chemistry and biology of aflatoxin B 1: from mutational spectrometry to carcinogenesis. Carcinogenesis. 22(4): 535-545. Tag, A. G., G. F. Garifullina, A. W. Peplow, C. Ake, Jr., T. D. Phillips, T. M. Hahn, and M. N. Beremand. 2001. A novel regulatory gene, TrilO, controls trichothecene toxin production and gene expression. Appl. Environ. Microbiol. 67:5294-5302. Timberlake, W. E. and E. C. Barnard. 1981. Organization of a gene cluster expressed specifically in the asexual spores of A. nidulans. Cell 26:29-37. Timberlake, W. E. and M. A. Marshall. 1988. Genetic regulation of development in Aspergillus nidulans. Trends Genet. 4: 162-169. Trail, F ., P. K. Chang, J. Cary, and J. E. Linz. 1994. Structural and functional analysis of the nor-I gene involved in the biosynthesis of aflatoxins by Aspergillus parasiticus. Appl. Environ. Microbiol. 60:4078-4085. 153 Trail, F., N. Mahanti, and J. Linz. 1995a. Molecular biology of aflatoxin biosynthesis. Microbiology 141 ( Pt 4):755-765. Trail, F ., N. Mahanti, M. Rarick, R. Mehigh, S. H. Liang, R. Zhou, and J. E. Linz. 1995b. Physical and transcriptional map of an aflatoxin gene cluster in Aspergillus parasiticus and functional disruption of a gene involved early in the aflatoxin pathway. Appl. Environ. Microbiol. 61:2665-2673. Tully, D. B. and J. A. Cidlowski. 1993. Protein blotting procedures to evaluate interactions of steroid receptors with DNA. Methods Enzymol. 218:535-551. van Gorcom, R. F., P. J. Punt, P. H. Pouwels, and C. A. van den Hondel. 1986. A system for the analysis of expression signals in Aspergillus. Gene 48:211-217. Watanabe, C. M., D. Wilson, J. E. Linz, and C. A. Townsend. 1996. Demonstration of the catalytic roles and evidence for the physical association of type I fatty acid synthases and a polyketide synthase in the biosynthesis of aflatoxin B1. Chem. Biol. 3:463-469. Weigel, B. J., S. G. Burgett, V. J. Chen, P. L. Skatrud, C. A. F rolik, S. W. Queener, and T. D. Ingolia. 1988. Cloning and expression in Escherichia coli of isopenicillin N synthetase genes from Streptomyces lipmanii and Aspergillus nidulans. J. Bacteriol. 170:3817-3826. Wilson, D. M. and G. A. Payne. 1994. Factors affecting Aspergillus flavus group infection and aflatoxin contamination of crops, p. 309-346. In D. L. Eaton and J. D. Groopman (eds.), The Toxicology of Aflatoxins, Academic Press, San Diego. Woloshuk, C. P., K. R. F outz, J. F. Brewer, D. Bhatnagar, T. E. Cleveland, and G. A. Payne. 1994. Molecular characterization of aflR, a regulatory locus for aflatoxin biosynthesis. Appl. Environ. Microbiol. 60:2408-2414. Woloshuk, C. P. and G. A. Payne. 1994. The alcohol dehydrogenase gene ath is induced in Aspergillus/lavas grown on medium conducive to aflatoxin biosynthesis. Appl. Environ. Microbiol. 60:670-676. Woloshuk, C. P., E. R. Seip, G. A. Payne, and C. R. Adkins. 1989. Genetic transformation system for the aflatoxin-producing fungus Aspergillusflavus. Appl. Environ. Microbiol. 55:86-90. World Health Organization. 1979. Mycotoxins, p. 1-127. World Health Organization, Geneva. Young, C., L. McMillan, E. Telfer, and B. Scott. 2001. Molecular cloning and genetic analysis of an indole-diterpene gene cluster from Penicillium paxilli. Mol. Microbiol. 39:754-764. Yu, J., J. W. Cary, D. Bhatnagar, T. E. Cleveland, N. P. Keller, and F. S. Chu. 1993. Cloning and characterization of a cDNA from Aspergillus parasiticus encoding an O- 154 methyltransferase involved in aflatoxin biosynthesis. Appl. Environ. Microbiol. 5923564- 3571. Yu, J., P. Chang, D. Bhatnagar, and T. E. Cleveland. 2000a. Cloning of a sugar utilization gene cluster in Aspergillus parasiticus. Biochim. Biophys. Acta 1493221 1-214. Yu, J., P. K. Chang, D. Bhatnagar, and T. E. Cleveland. 2000b. Genes encoding cytochrome P450 and monooxygenase enzymes define one end of the aflatoxin pathway gene cluster in Aspergillus parasiticus. Appl. Microbiol. Biotechnol. 53:583-590. Yu, J., P. K. Chang, J. W. Cary, M. Wright, D. Bhatnagar, T. E. Cleveland, G. A. Payne, and J. E. Linz. 1995a. Comparative mapping of aflatoxin pathway gene clusters in Aspergillus parasiticus and Aspergillus/laws. Appl. Environ. Microbiol. 61:2365- 2371. Yu, J., P. K. Chang, G. A. Payne, J. W. Cary, D. Bhatnagar, and T. E. Cleveland. 1995b. Comparison of the omtA genes encoding O-methyltransferases involved in aflatoxin biosynthesis from Aspergillus parasiticus and A. flavus. Gene 163:121-125. Yu, J. H., R. A. Butchko, M. F ernandes, N. P. Keller, T. J. Leonard, and T. H. Adams. 1996. Conservation of structure and function of the aflatoxin regulatory gene aflR from Aspergillus nidulans and A. flavus. Curr. Genet. 29:549-555. Zhou, R. and J. E. Linz. 1999. Enzymatic function of the nor-I protein in aflatoxin biosynthesis in Aspergillus parasiticus. Appl. Environ. Microbiol. 65:5639-5641. 155 Mlv'vHL‘Ah :IAIL LINV‘JEW‘SII V th‘il-Alilk a 3 1293 02365 8176