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POSITION-DEPENDENT REGULATION OF THE NOR-1

PROMOTER IN THE AFLATOXIN GENE CLUSTER AND

lMMUNO—LOCALIZATION OF PROTEINS INVOLVED IN
AFLATOXIN BIOSYNTHESIS

presented by

Ching-Hsun Chiou

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POSITION-DEPENDENT REGULATION OF THE NOR-1 PROMOTER IN THE
AFLATOXIN GENE CLUSTER AND IMMUNO-LOCALIZATION OF PROTEINS
INVOLVED IN AFLATOXIN BIOSYNTHESIS
By

Ching-Hsun Chiou

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

ABSTRACT

POSITION-DEPENDENT REGULATION OF THE NOR-1 PROMOTER IN THE
AF LATOXIN GENE CLUSTER AND IMMUNO-LOCALIZATION OF PROTEINS
INVOLVED IN AF LATOXIN BIOSYNTHESIS

By

Ching-Hsun Chiou

Aflatoxins (AF) are highly toxic and carcinogenic fungal secondary metabolites.
AF contamination in crops including com, peanuts, cottonseed, and specific tree nuts has
caused significant agricultural economic and human health concerns throughout the
world. My research goal was to study regulation of AF biosynthesis in Aspergillus
parasiticus. As along term goal, these studies should aid efforts to discover strategies to
efficiently eliminate AF contamination of agricultural commodities and in turn to ensure
product safety and consumer health.

Biosynthesis of AF in the filamentous fungi in the genus Aspergillus utilizes a
complex pathway consisting of at least eighteen enzymes and at least one positive
regulatory factor, AflR; these genes are clustered within a 75 kb chromosomal region.
Expression of most of the AF genes in the cluster is co-regulated but the mechanisms are
not clear. The localization and interaction of these enzymes to synthesize AF at the
cellular level has yet to be determined. Therefore, two hypotheses are included in my
study. 1) Expression of genes involved in aflatoxin biosynthesis is positively regulated
within the AF gene cluster. 2) Enzymes involved in AF biosynthesis are localized within
developmental structures (e.g., conidiophores) in the colony.

The objectives of my research were to: 1) study the positional-effects on AF gene
(nor-I) expression in the gene cluster; 2) generate highly-specific antibodies against an
AF protein (anti-VBS) for immuno-localization studies; 3) develop methods to localize
AF proteins within a fungal colony; and 4) localize proteins involved in AF biosynthesis

in a time-course dependent fractionated colony.

To study positional effects on nor-I promoter function, a plasmid pAPGUSNP
containing the nor-1 promoter sequence fused to a reporter gene (uidA encoding B-
glucuronidase, GUS) plus a pyrG selectable marker was constructed. Transformation of
pAPGUSNP into CSIO (ver-I, wh-I, pyrG) resulted in homologous recombination at 5'
nor-I, 3' nor-1, or the pyrG locus. The plasmid integration site was screened by a novel
rapid DNA extraction/PCR assay and confirmed by Southern hybridization analysis. The
nor-1 promoter was positively regulated in the AF gene cluster while outside the cluster
(pyrG locus), nor-1 promoter activity decreased dramatically. The data supported the
hypothesis that expression of AF genes is cluster position-dependent.

The second hypothesis was tested by studying localization AF proteins utilizing
anti-VBS antibodies raised against recombinant Maltose Binding Protein-VBS expressed
in Escherichia coli (generated in this study), and other antibodies available in the
laboratory (anti- Nor-l , Ver-l , and OmtA). A 72 h-old colony was fractionated based on
24 h intervals to correlate the time of grth with AF protein distribution. Colony
fractions were fixed and embedded in paraffin to generate 4-5 pm sections for immuno-
fluorescence labeling. Confocal Laser Scanning Microscopy (CLSM) demonstrated that
Nor-1, Ver-l , OmtA, and VBS were evenly distributed among different cell types and not
concentrated in conidiophores, contrary to the hypothesis. These proteins were not
evenly distributed throughout a 72 h—old colony. Among three fractions, colony fraction
S2 (24 to 48 h growth) exhibited the highest abundance of Nor-1 , Ver—l , OmtA, and VBS
(analyzed by CLSM and quantitative fluorescence intensity analysis), the highest levels of
AF Bl (determined by enzyme-linked-immunosorbent-assay), and the highest numbers of
newly-developed conidiophores. These data support a spatial and temporal association

between AF protein expression/AF production and sporulation at the colony level.

ACKNOWLEDGMENTS

I would first like to thank my major professor, Dr. John Linz, for his support and
guidance throughout this study. He is a good mentor who teaches me science and many
other precious things beyond the science. I admire Dr. Linz as a good scientist and a
knowledgeable person.

Appreciation is also expressed to Dr. Karen Klomparens, Dr. James Pestka, Dr.
Gale Strasburg, and Dr. Jay Schroeder for serving on my guidance committee.

I am also especially gratefiil to Dr. Shirley Owens and others who assisted me in
the Center for Advanced Microscopy at Michigan State University.

Thanks to Li-Wei Lee, Mike Miller, Matt Rarick, and other members in the Linz
laboratory for all the help they have provided.

I also appreciate my parents, family, and friends, who always stand by me and

show me their support and encouragement.

iv

TABLE OF CONTENTS

LIST OF TABLES .................................................................................. ix
LIST OF FIGURES ........................................................................................................ x
CHAPTER 1
Literature review .............................................................................................................. 1
DELETERIOUS EFFECTS OF AFLATOXINS .................................................. 1
Background and history .............................................................................. 1
Economic impact and regulation of AF in foods ......................................... 3
Environmental factors and controlling AF contamination .......................... 4
Toxicological effects of AF ........................................................................ 5
Aflatoxin and liver cancer ............................................................................ 6
Bioactivation of AFB1 .................................................................... 6
Detoxification of AF ........................................................................ 6
AF B1-DNA adducts and AF B1 indcued p53 gene mutations ........ 6
A synergistic effect between AFB1 and hepatitis B virus
(HBV) ......................................................................................... 8
AFLATOXIN BIOSYNTHESIS .......................................................................... .9
Aflatoxin gene cluster ...................................................................... 9
Regulation of AF biosynthesis ....................................................... 9
Partially-duplicated AF gene cluster ............................................. 13
Sugar utilization gene cluster ........................................................ 14
SECONDARY METABOLISM AND FUNGAL DEVELOPMENT .............. 14
Association between secondary metabolism and fungal
development ......................................................................................... 14
G-Protein signaling pathway ..................................................................... 14
FluG .......................................................................................................... 1 5
AF bioynthesis and conidiation ............................................................... 15
LOCALIZATION OF AF ASSOCIATED PROTEINS ................................... .17
Penicillin biosynthesis as an example ........................................................ 17
Organelles possibly involved in AF biosynthesis18
Microbodies .................................................................................. 18
Woronin bodies ............................................................................. 18
Vesicles ......................................................................................... 19
Vacuoles ........................................................................................ 19
Endoplasmin reticulum (ER) ........................................................ 21
CONFOCAL LASER SCANNING MICROSCOPY (CLSM) IN THE STUDY
OF FUNGAL BIOLOGY ..................................................................................... 22
Introducion to CLSM ................................................................................ 22
Image quality enhancement by CLSM ..................................................... 23
CLSM Imaging ...................................................................................... 25
Optical sections ........................................................................... .25

Z-series .......................................................................................... 25

Phi-z sectioning ............................................................................. 26
Time-lapse and live imaging .......................................................... 26
Fluorescence imaging ................................................................... .26
CHAPTER2
A rapid DNA extraction method for screening large numbers of Aspergillus
transformants via polymerase chain reaction ............................................................. 31
ABSTRACT ......................................................................................................... 31
INTRODUCTION ............................................................................................... 3]
MATERIAL AND METHODS .......................................................................... 32
RESULTS AND DISCUSSION ........................................................................... 33
ACKNOWLEDGEMENTS ................................................................................. 4 1
CHAPTER3
Position-dependent regulation of the nor-1 promoter in the aflatoxin gene
cluster ............................................................................................................................... 42
ABSTRACT ......................................................................................................... 42
INTRODUCTION ............................................................................................... 43
MATERIALS AND METHODS ....................................................................... 45
Strains and grth conditions .................................................................. 45
Construction of pAPGUSNP ................................................................... 45
Fungal transformation ............................................................................... 48
Identification of the site of plasmid integration ........................................ 48
Rapid DNA extraction procedure for PCR analysis ..................... 48
PCR analysis ................................................................................. 48
Solid-culture GUS activity assay .............................................................. 49
Southern hybridization analysis ................................................................ 49
5' nor-1 integration ........................................................................ 51
3' nor-1 integration ........................................................................ 51
Integration at pyrG ........................................................................ 51
RESULTS .............................................................................................................. 51
Generation of pAPGUSNP ..................................................................... .51
Transformation of A. parasiticus CSIO with pAPGUSNP ...................... 56
Solid-culture GUS assay ........................................................................... 56
Determination of pAPGUSNP integration site within the
chromosome ......................................................................................... 56
Confirmation of pyrG expression in pyrG+ transformants ....................... 59
DISCUSSION ....................................................................................................... 59
ACKNOWLEDGEMENTS ................................................................................. 63
CHAPTER4
Immuno-localization of Nor-l, Ver-l, and OmtA proteins involved in aflatoxin
biosynthesis in Aspergillus parasiticus .......................................................................... 64
INTRODUCTION ............................................................................................... 64
MATERIALS AND METHODS ........................................................................ 67

vi

Fungal strains ............................................................................................ 67

Time-dependent fractionation of colonies grown on solid medium .......... 67
Preparation of paraffin-embedded fungal sections .................................... 67
Immuno-fluorescence labeling of firngal cells .......................................... .69
Confocal Laser Scanning Microscopy (CLSM) ........................................ 70
Quantitative fluorescence intensity analysis ............................................. 70
RESULTS .............................................................................................................. 71
Immuno-localization of Nor-1 and Ver-l .................................................. 71
Immuno-localization of OmtA .................................................................. 71
Quantitative fluorescence intensity analysis ............................................. 79
DISCUSSION ....................................................................................................... 85
ACKNOWLEDGEMENTS ................................................................................. 89
CHAPTERS
Distribution and sub-cellular localization of the aflatoxin enzyme versicolorin B
synthase (VBS) in time-fractionated colonies of Aspergillusparasiticus .................. 90
ABSTRACT ......................................................................................................... 90
INTRODUCTION ............................................................................................... 91
MATERIALS AND METHODS ........................................................................ 94
Fungal strains and growth conditions ....................................................... 94
Production of antibodies against native VBS ........................................... 94
Generation of MBP (Maltose Binding Protein)-VBS fusion protein ....... 95
Production and purification of PAb against MBP-VBS ........................... 95
Time-dependent fractionation of fiingal colonies ..................................... 96
Fungal protein extraction .......................................................................... 96
SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE) and
Western blot analysis ........................................................................... 97
Detection of AFB1 in colony fiactions ..................................................... 97
Thin Layer Chromatagraphy (TLC) ............................................. 97
Enzyme-Linked-Immunosorbent-Assay (ELISA) ........................ 97
Preparation of paraflin-embedded fungal sections and immuno-
fluorescence labeling ............................................................................ 98
Confocal Laser Scanning Microscopy (CLSM) ........................................ 99
Quantitative fluorescence intensity analysis ............................................. 99
Statistical analysis ................................................................................... 100
Preparation of ultra-thin fungal sections, immuno-gold labeling,
and TEM analysis ............................................................................ . 100
RESULTS ............................................................................................................ 100
Generation of PAb against native firngal VBS ......................................... 100
Expression of MBP-VBS fusion protein and production of PAb
against VBS ........................................................................................ 101
Expression of VBS in A. parasiticus grown on solid medium ............... . 106
Confocal Laser Scanning Microscopy (CLSM) of time-dependent
expression of VBS in a 72 h-old firngal colony .................................. 109
Fluorescence intensity quantification and statistical analysis ................. 109
Production of AFB1 in colony fractions ................................................. 109

vii

Sub-cellular localization of VBS ............................................................ . 109

DISCUSSION ..................................................................................................... l 16
ACIG‘IOWLEDGEMENTS ............................................................................... 124
BIBLIOGRAPHY ......................................................................................................... .125

viii

LIST OF TABLES

Table 2.1. Primer sequences and PCR cycling parameters used for analysis of fungal
genomic DNA templates ........................................................................... 34

Table 3.1. Primer sequences and PCR cycling parameters used for analysis of site of
integration using DNA templates prepared by the rapid method ............................. 50

Table 3.2. Summary of GUS activities and plasmid integration sites in A. paraciticus
CSIO transformants carrying the nor::GUS firsion plasmid pAPGUSNP (Chiou), and
NR-l harboring pAPGUSNNB (Wilson, 1998) and pNG3.0 (Miller, 2003) ............... 61

Table 4.1. Quantative fluorescence intensity analysis of A. parasiticus SU-l and AF SIO
immuno-fluorescence labeled with Nor-1, Ver-l and OmtA antibodies ..................... 86

Table 5.1. Quantitative fluorescence intensity analysis of A. parasiticus SU—l and
AF SlO labeled with polyclonal antibodies to VBS and OmtA ............................ 113

Table 5.2. Quantitative detection (ELISA) of AFB1 in colony fi'actions of SU-I cultured
on YES solid media for 72 h ..................................................................... 114

LIST OF FIGURES

Figure 1.1. Chemical structures of AFB1, AFB2, AFG], AFG2, AFM], AFM2, and

AFB 1-8, 9-epoxide ................................................................................... 2
Figure 1.2. Generally accepted pathway for sterigmatocystin and aflatoxin

biosynthesis .......................................................................................... 10
Figure 1.3. The G-protein signaling pathway involved in ST biosynthesis in A.

nidulans ............................................................................................... 16
Figure 1.4. Diagram of the light path in a confocal instrument ............................... 24

Figure 1.5. CLSM Z-series of A. parasiticus SU-l conidiophores stained with SYTOX
green fluorescence dye .............................................................................. 26

Figure 1.6. CLSM single optical section and phi-z scan of A. parasiticus SU-l
microbodies .......................................................................................... 28

Figure 2.1. Homologus recombination of pAPGUSNP into the A. parasiticus
chromosome and primers used in integration analysis ......................................... 35

Figure 2.2. Amplification of PCR targets using the rapid DNA extraction technique. . .37

Figure 3.1. Restriction endonuclease maps of plasmids containing the nor-1::GUS
construct related to this study ..................................................................... 46

Figure 3.2. Southern hybridization analysis of site of integration in pAPGUSNP
transformants ........................................................................................ 52

Figure 3.3. Southern hybridization analysis of pAPGUSNP integrated at
pyrG ................................................................................................... 54

Figure 3.4. PCR analysis to detect site of integration in pAPGUSNP transformants. . . 57

Figure 3.5. Solid-culture GUS activity assay on CSIO transformants harboring
pAPGUSNP .......................................................................................... 58

Figure 4.1. Time-dependent fractionation of firngal colonies ................................ 68

Figure 4.2. Immuno-fluorescence confocal microscopy of Nor-1 and Ver-l in A.
parasiticus SU-l and AF SlO (cyIR knockout mutant) grown on YES agar for 72 h ........ 72

Figure 4.3. CLSM immuno-fluorescence detection of Nor-1 in A. parasiticus SU-1.. ...74

Figure 4.4. CLSM immuno-fluorescence detection of Ver—l in A. parasiticus SU-l . . . ...76

Figure 4.5. Immuno-fluorescence labeling of Ver-l antibodies in the control strain A.
parasiticus AF 810 and the wild-type strain SU-l .............................................. 78

Figure 4.6. OmtA protein localization in time-fractionated colonies of A. parasiticus SU-
1, AF S10 ((ng knockout), C 810 and LW1432 (omtA knockout) grown on YES agar for 72
h ........................................................................................................ 80

Figure 4.7. Protein localization using OmtA PAb and anti-SKL, and detection of nuclei
using SYTOX Green ................................................................................ 82

Figure 4.8. CLSM immuno-fluorescence detection of OmtA in A. parasiticus SU-l . . . .84

Figure 5.1. Western blot analysis of VBS detected by PAb raised against native firngal
VBS ......... . ........................................................................................ 102

Figure 5.2. SDS-PAGE of recombinant MBP-VBS fusion protein ........................ 103

Figure 5.3. MBP-VBS treated with Factor Xa to generate MBP (42 kDa) and VBS (52
kDa) .................................................................................................. 104

Figure 5.4. Western blot analysis of VBS produced in liquid culture using highly specific
PAb raised against MBP-VBS ................................................................... 107

Figure 5.5. Western blot analysis of VBS expressed in A. parasiticus SU-I colonies
(solid culture) and colony fractions .............................................................. 108

Figure 5.6. CLSM of VBS protein distribution in time-fractionated colonies of A.
parasiticus SU-l (S1, S2, S3) and AF 810 (R1, R2, and R3) grown on YES agar for
72h ................................................................................................... 110

Figure 5.7. Thin Layer Chromatagraphy analysis of AFB1 production in colony
fractions ............................................................................................. 111

Figure 5.8. CLSM immuno-fluorescence of VBS A. parasiticus SU-l grown in solid
culture ............................................................................................... 112

Figure 5.9. CLSM dual detection for localization of VBS and nuclei in A. parasiticus
SU-l .................................................................................................. 115

Figure 5.10. Immuno-gold labeling of VBS in A. parasiticus AF $10 and SU-l analyzed
by TEM ............................................................................................. 117

CHAPTER 1

Literature review

DELETERIOUS EFFECTS OF AFLATOXINS

Background and history. Aflatoxins (AF) are secondary metabolites mainly
produced by the filamentous fungi Aspergillus flaws and A. parasiticus (Wilson and
Payne, 1994; CAST, 2003). They are less frequently produced by A. nomius, A.
pseudotamari, A. bombycis, and one isolate of A. ochraceoroseus (Klich et al., 2000; Ito

et al., 2001; Bhatnagar et al., 2003). A. parasiticus synthesizes four major aflatoxins
including aflatoxin B1(AF B1), AFB2, AFG1, and AFG2 (Asao et al., 1963). Animals

ingest AF and produce metabolites AF M1 and AF M2 in milk (Fig. 1.1). Among these

AF molecules, AF B1 is the most toxic and carcinogenic (Eaton and Groopman, 1994;

Mclean and Dutton, 1995).

AP contamination of corn, peanuts, cotton seed, and certain tree nuts has caused
significant agricultural economic and human health problems throughout the world
(CAST, 2003). This mycotoxin first aroused public attention in the early 19608 with the
incidence of "Turkey X" disease (Blount, 1961). Large quantities (approximtely 10, 000)
of turkeys died from acute hepatotoxicity caused by contamination of AF in their peanut
feed (Siller and Ostler, 1961). Rainbow trout that received AF contaminated diets were
discovered to develop liver cancer in the United States (Sinnhuber et al., 1977). In the
19708, severe pre-harvest AF contamination was reported in the Southern and midwestem
United States (Anderson, 1975). A. flavus and AF contamination are frequently seen in
the Southern US., consistent with the observation that serious outbreaks are related to
below-average rainfall and above-average temperature (Payne, 1998). Dietary exposure

to AF has been associated with a significant incidence of human liver cancer. Studies on

experimental animals indicated that AF B1 is a potent mutagen

 

 

 

 

 

 

 

 

 

 

 

 

AF B1-8,9-epoxide I

 

40A, / ...

Figure 1.1. Chemical structures ofAFB], AFBz, AFG1, AFG2, AFM], AFMz, and
AF Bl-8, 9-epoxide.

and hepato-carcinogen (reviewed in Eaton and Groopman, 1994). Based on this

information, the International Agency for Research on Cancer (IARC) identified AF Bl as
a group I carcinogen in humans (IARC, 1993). Several studies also indicated that
infection of hepatitis B virus (HBV) causes a synergistic effect with AF B1 induced p53
mutation related to hepatocarcinogenesis (HCC) (Hsia et a1, 1992; Harris and Hollstein,
1993; Jackson and Groopman, 1999, CAST, 2003). Hepatitis C viral (HCV) infection is
the most important risk factor for HCC in western countries; however, no adequate
epidemiological data on the relative roles of HCC and AF were available (Henry et al.,
1999; Montalto et al., 2002).

Economic impact and regulation of AF in foods. Approximately 25% of food
crops are affected by mycotoxin contamination (including AF) every year throughout the
world and this results in serious health and economic concerns (Mannon and Johnson,
1985). Since 1977, documents regarding mycotoxin regulation and control have been
published by the Food and Agricultural Organization (FAO) (Boutrif, 1995). Over 50
countries developed guidelines to regulate mycotoxins. The United States set action
levels for aflatoxin under the Food, Drug and Cosmetic Act (See. 402) (Sharma and
Salunkhe, 1991; Bhatanagar et al., 2002; CAST, 2003). The US. Food and Drug

Administration (FDA) prohibits interstate commerce of feed grain containing more than
20 parts per billion (ppb, or ug/kg) AF B] and prohibits the sale of milk and eggs

containing more than 0.5 ppb AF M1. The action level for a feed crop varies for different

animals depending on their sensitivity to AP. Over the world, the maximum allowed
levels for aflatoxins in foodstuff range fiom zero tolerance to 50 ppb (Van Egrnond,
1993; Bhatanagar et al., 2002).

Contamination of agricultural commodities with AF poses a significant threat to
the food supply world-wide. In the United States, the mean economic losses of crops due
to mycotoxin contamination (including AF, fumonisins, and deoxynivalenol) are

estimated to be $932 million. The loss attributed to AF contamination on food crops

(corn and peanuts) was approximately $47 million per year and approximately $225
million per year in feed corn (CAST, 2003). Thus, prevention and control of mycotoxin
contamination is an important issue.

Environmental factors and control of AF contamination. In the field, A.
flaws and A. parasiticus are able to grow over the temperature range from 12 to 48 0C
(optimum 35 to 37 0C) and with a water activity as low as 0.80 (optimum 0.95).
However, AF production in these fungi requires a smaller range of temperature (28 to 33
0C, optimum 31°C) and water activity (0.85 to 0.97, optimum 0.90) (Hussein and Brasel,
2001; Bhatanagar et al., 2002).

Pre-harvest contamination of AF on agriculture commodities is usually related to
the environmental conditions that generate fungal growth and toxin synthesis. Several
methods have been applied to control AF at this stage, including use of fimgicides and
insecticides (insects facilitate Aspergillus infection of plants), modified irrigation
practice, and the use of atoxigenic biocompetitive strains (Cotty and Bhatnagar, 1994;
Cary et al., 2000; Bhatnagar et al., 2002). However, these procedures have limited
effects, particularly in years of drought when fungal growth and toxin synthesis are
elevated. Ongoing research is focused on alternative strategies to control AF
contamination in the field.

Time of harvest is also critical to restrain the levels of AF contamination. For
example, contaminated corn should be harvested early and a subsequent drying procedure
may help to prevent an increase in AF contamination that occurs post-harvest. Post-
harvest contamination can also be controlled by maintaining proper storage conditions
(CAST, 2003). For example, limiting the water activity of the harvest grains below 0.85
and maintaining the storage temperature below 10°C may prevent A. flavus spore
germination (Marin et al., 1998; CAST, 2003).

Toxins on contaminated products can be removed or detoxified by various

physical, chemical and biological methods (Bhatnagar et al., 2002; CAST, 2003). For

example, toxins can be removed by filtration or adsorption onto clays or activated
charcoal. Oil roasting or dry roasting peanuts (Marth and Doyle, 1979) and microwave
treatment (F arag et al., 1996) may partially destroy AF although they are heat stable. AF
in contaminated grains may be extracted by a selected solvent mixture (e.g., 95% ethanol,
90% aqueous acetone, and 80% isopropanol), yet this procedure is costly and product
safety remains a concern (CAST, 2003).

Toxicological effects of AF. The degree of AF-induced toxic effects in animals
varies, depending on species, dose, exposure duration, and nutritional status of the host
(CAST, 2003). The major organ targeted by AF is the liver. Acute aflatoxicosis often
produces hepatitis in animals and humans, and ingestion of a high dose may be lethal.
Acute hepatitis has been related to consumption of foodstuff contaminated with high
doses of AF, especially corn (Ngindu et al., 1982). Under severe conditions, 25%
mortality has been reported in outbreaks in India (Krishnamachari et al., 1975).

Sublethal doses generate chronic toxicity and low levels of chronic exposure can
result in cancer. A sensitive model to study AF toxicity is trout, and the LD50 was
estimated to be 0.5 to 1.0 mg/kg (CAST, 2003). A high incidence of hepatoma was seen
in trout fed with 80 ppb AF contaminated diet. Studies in cattle showed chronic clinical
signs including reduced weight gain, irnmuno-suppression, lower milk production, liver
damage, and lower reproduction (Bodine and Mertens, 1983). In humans, chronic dietary
exposure to AF has a high risk of generating hepatocellular carcinoma (HCC). A high
level of AF daily intake (1.4 ug)(Wild et al., 1992) and a high incidence of hepatitis B
infection were associated with HCC (Jackson and Groopman, 1999). Epidemiological
studies indicated that an increase in AF ingestion from 3 to 222 ng/kg body weight per
day corresponded to elevated liver cancer incidence (from 2 to 35 cases per 100,000

population per year) (Jackson and Groopman, 1999).

Aflatoxin and liver cancer.
Bioactivation of AFB1. AF B1 naturally produced in the fungus is not a potent
mutagen and carcinogen to humans (and other animals); however, the liver cytochrome

P450 enzymes generate highly reactive molecules during the oxidative metabolic
processes (Eaton and Groopman, 1994). Hence, the major targets for AF Bl toxicity are

hepatocytes. In vitro studies have shown that biotransformation of AF B1 requires

molecular oxygen and NADPH. The biotransforrned AFB], the AF B1-8,9-epoxides (Fig.

1.1), are potent bioreactive compounds causing mutation and carcinogenesis. The
epoxide molecules covalently bind to proteins, DNA and RNA molecules to generate
adducts. Adduct formation in turn impairs the normal molecular and cellular fimctions
for the targets (McLean and Dutton, 1995).

Detoxification of AF. Detoxification of AF B1-8,9—epoxides mainly depends on
conjugation with glutathione (GSH). Glutathione S transferase (GST) activity is therefore

an important determinant for the susceptibility of different species to the toxic effects of

AFB1 (Eaton and Gallagher, 1994). Different animal species also showed different
responses to AFBl toxicity due to variability in bioactivation and detoxification

processes. In addition to humans, rat, rainbow trout and turkey are susceptible to AF B1

induced toxicity, whereas mice, ducks and chickens showed lower toxic response
(Mclean and Dutton, 1995). It is likely that genetic variability in the expression of
cytochrome P450 enzymes may result in substantial differences between individuals in
susceptibility to the carcinogenic effects of aflatoxins. For example, glutathione S
transferase is the predominant detoxifying activity of AF B1-8,9-epoxides in mice.
However, in humans, GST has very little activity toward AF B1-8,9-epoxides, suggesting
higher human sensitivity to the genotoxic effect of AF B1 (Hussein and Brasel, 2001).
AFB1-DNA adducts and AFB1 induced p53 gene mutations. The N7-guanine
residue of DNA is a potential target for AF B1-8,9-epoxides, forming a AF Bl-N7-guanine

adduct. Adduct formation or depurination can lead to mutations in DNA during DNA

replication, mainly resulting in GC—-)TA transversions (Eaton and Groopman, 1994).
Mutation in the p53 tumor suppressor gene by AF B1 epoxide has also proposed to be
involved in hepatocarcinogenesis (Massey et al., 1995; Mclean and Dutton, 1995;
Jackson and Groopman, 1999).

The p53 gene is a tumor suppressor gene that regulates cell growth. The
biological functions of P53 are involved in control of cell cycle, DNA replication and
repair, cell differentiation and apoptosis (Harris and Hollstein, 1993; Shen et al., 1996).
Mutations in the p53 tumor suppressor gene were frequently detected in HCC patients
(Greenblatt et al., 1994). For example, codon 249 in exon VII of p53 has been identified
as a mutational hot-spot in HCC but not in other tumor types (Yang, et al., 1997; Jackson
and Groopman, 1999). A point mutation in the third base of codon 249 of p53 results in a
G—)T transversion (AGG—-> AGT) and a change in amino acid sequence from Arg to Ser.
This mutation has been fi'equently associated with high levels of AF B1 exposure in HCC
patients.

The mutation of the p53 tumor suppressor gene at codon 247-250 (nucleotide
14067-14075) induced by AF B1 has been studied (Aguilar et al., 1993). AFB] was
activated by rat liver rrricrosomes and mutagenesis at codons 247-250 of p53 gene was
analyzed in human hepatocarcinoma cells, HepG2, by restriction fragment length
polymorphism/polymerase chain reaction (RFLP/PCR). The results indicated that AF B1
induces the G—-)T transversion at the third position of codon 249 with relatively high
frequency. However, cells exhibiting different bioactivation and detoxifying systems
may influence the results. Mace et al. (1997) investigated AFB 1 -induced DNA adduct
formation and p53 mutations in human liver cells expressing various cytochrome p450

isozymes. The results suggested that CYP1A2 is the most effective isoforrn in the

activation of AF B1 to cytotoxic products and CYP3A4 for genotoxic metabolite

formation. Cells containing CYP1A2 and CYP3A4 also showed a higher level of AF B1-

DNA adduct formation and increased levels of p53 mutations (Mace, et al., 1997).

A synergistic effect between AFB1 and hepatitis B virus (HBV). Exposure to

AFB] alone has been verified as an important risk factor for development of human

HCC. A high prevalence of hepatitis B virus (HBV) infection frequently occurred along
with a high level of aflatoxin contamination in many regions of the world. HBV infection
and AF B] exposure have been suggested to have synergistic effects on tumor formation
as well as p53 gene mutation, (Lunn et al., 1997; Yang, et al., 1997). This was supported
by a study using a transgenic mouse model that expresses hepatitis B surface Ag (HBsAg)
and genetic inbred p53 null mice to determine the association of AF B] , HBV and p53
expression on liver tumor formation (Ghebranious and Sell, 1998). The results showed
that AF B] induced a higher level of tumor formation than HBV infection alone. A
synergistic effect between AFB] and HBV infection increased HCC. The highest risk
group for liver cancer formation was generated with combinations of AF B] ingestion,
HBV expression and p53 mutation.

Epidemiological studies also suggest that high levels of AF B] exposure and HBV
infection may have synergistic effects on HCC induction. Shen and Ong (1996)
summarized the results from more than thirty reports from 1991 to 1995 with more than

1500 human HCC samples that were examined for p5 3 mutation with respect to different

AFB] exposure levels. It was found that the frequency of codon 249 mutations in the p53

tumor suppressor gene directly parallels the environmental AFB] exposure level. Lasky

and Mahden (1997) performed similar research by retrieving twenty studies that provided
analytical data in p53 mutations in HCC patients. The analysis suggested that high levels
of AF exposure (such as Southern Africa and China) were associated with the hot-spot
mutation (coden 249) in p53 in a high proportion (approximately 50%) of HCC patients.
On the contrary, codon 249 mutations were not detected in areas with low AF exposure

such as Japan, Europe and the USA (Jackson and Groopman, 1999).

 

AFLATOXIN BIOSYNTHESIS

Aflatoxin gene cluster. Biosynthesis of AF in A. flavus and A. parasiticus from
the starter unit, acetate, requires at least eighteen enzymatic activities (Woloshuck and
Prieto, 1998; Payne and Brown, 1998; Minto and Townsend, 1998; Motomura et al.,
1999; Cary et al., 2000; Bhatnagar et al., 2003). A. nidulans possesses a similar
biosynthetic pathway to synthesize the secondary metabolite Sterigrnatocystin (ST), a
precursor of AF, but this species lacks the final enzymatic activities (OmtA and OrdA) to
convert ST to AF (Brown, et al., 1996). Many of the ST/AF genes have been identified
and they were found clustered within a 75 kb chromosomal DNA region (Trail et al,
1995a; Woloshuck & Prieto, 1998; Payne & Brown, 1998; Bhatnagar et al., 2003)(Figure
1.2). These genes include aflR, a pathway specific regulatory factor of this gene cluster
(Chang et al., 1993; Woloshuck et al, 1994), and many structural genes including pksA
(Chang et al., 1995; Feng and Leonard, 1995),fas1,fas2 (Mahanti et al., 1996; Trail et
al., 1995), nor-1 (Chang et al, 1992 ), avnA (Yu et al., 1997), avf-I (Prieto et al., 1996),
estA (Yu et al., 2002), vbs (Silva et al., 1996), ver-I (Skory et al., 1992), omtA (Yu et al.,
1993) and ordA (Yu et al., 1998).

Expression of the AF genes appears to be co-regulated within the gene cluster.
Most of the structural genes are activated under AF producing conditions and the timing
of their expression is similar (Skory et al., 1993; Trail et al., 1995b; Brown et al., 1996).
However at least one AF gene, estA (encodes an esterase activity that converts versiconal
hemiacetal [VHA] to versiconal [VAL]), was recently reported to have a different
expression pattern compared to other AF genes (Yu et al., 2002).

Regulation of AF biosynthesis. An important positive regulator of the AF
biosynthetic pathway is AflR. This GAL4-type transcription factor contains a Cy86Zn2-
type DNA binding domain and is required for biosynthesis of AF (Chang et al., 1995b;
Ehrlich et al.,1999a; Ehrlich, et al., 2003). Many of the structural genes involved in AF

biosynthesis such as nor-I , ver—I, and omtA, as well as aflR itself, contain the

Figure 1.2. Generally accepted pathway for sterigmatocystin and aflatoxin biosynthesis.
(Adatpted from Bhatnagar et al., 2003)

10

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

1 Dehydrogenase "0’3 I
2 Monooxygenase cypA I
3 AF Transporter aflT I
4 Polyketide
synthase pksA I I
avnA I
5 Reductase —:Inor-1 15 Monooxgenase I
A 16 Desaturase It verB _
' L
6 Eggfhggigz fas-z 17 Dehydrogenase 3"“
7 Fatty acid 1 f 1 18 Me transferase I omtB
S nthase-1 35'
y 19 Me transferase II omtA I
8 Transcription A aflFLI —
Factor 20 Monooxgenase I ordA
9 Co-activator V aflJ
- 21 Ver B synthase I Vbs
10 Dehydrogenase v adhA I I-
"' 22 Monooxgenase I cpr I
11 Esterase II estA I _
12 A I-alcohol "
de ydrogenase ‘I norA I 23 Monooxgenase I
13 Dehydrogenase ver-1 24 Monooxgenase

 

14 Monooxgenase I “AI

 

 

 

NADH oxidase

Hexose
transporter

  

Glucosidase

Transcrp. Factor “m

 

Figure 1.2

palindromic consensus AflR binding site (5'-TCGN5CGR-3') in their promoter region
and require AflR for expression (Ehrlich et al., 1999b). In addition to aflR, an adjacent
gene aflJ possibly plays a role in regulation of AF biosynthesis. aflR and ale are
divergently transcribed; both genes share part of the promoter regions (Meyer et al.,
1998). Disruption of ale down-regulated AF production but the transcripts of other
structural genes including pksA, nor-1, ver-I, and omtA were still present (Meyer et al.,
1998). AflJ protein did not show any known sequence identity to other enzymes, yet its
peptide sequence contains three putative membrane spanning domains and a peroxisomal
targeting sequence, PTSl (Meyer et al., 1998). Thus, AflJ was suggested to be involved
in transport of AF proteins to specific targets (Meyer et al., 1998). However, the actual
function of AflJ has not been identified.

It is not known why AF genes are clustered but the organization and coordination
of expression of AF genes appears to be important (Trail et al., 1995b; Brown et al.,
1996). In AF producing Aspergillus spp., the transcription factor AflR is an important
positive regulator but AflR alone is not sufficient to regulate AF biosynthesis. Some
Aspergillus spp., e.g. A. sojae and A. oryzae used in fermentation of foods, contain
homologues of AF biosynthesis genes including aflR but they are not aflatoxigenic (Trail,
19953). A 28-amino acid zinc finger binding domain located at the C-terminus of AflR
(CTSCASSK-VRCTKEKPACARCIERGLAC) was present in A. flavus, A. parasiticus,
A. sojae, and some strains of A. oryzae (Chang et al., 1995b). aflR and other structural
genes such as nor-1 and ver-I were present in some A. sojae and A. oryzae strains but
they do not produce AF (Whatson et al., 1999). Transcripts of the aflR homologue in the
non-AF producing A. oryzae were also not detected under AF inducing conditions by
reverse-transcription polymerase chain reaction (RT-PCR) (Kusumoto et al., 1998).
Absence of AF biosynthesis in koji mold (A. sojae) was later confirmed to be caused by
defects in aflR expression, resulting in lack of expression of AF related genes

(Matsushima et al., 2001). In addition, some mutant strains contain an incomplete gene

12

cluster affecting gene expression and ST/AF production, and implicating a loss of
functional regulation in these species (Cary, et al., 1999; Butchko, et al., 1999). These
data also suggested that in addition to AF LR, other mechanisms appear to regulate
biosynthesis of AF.

The cluster effects for expression of AF genes play an important role in AF
biosynthesis. Significance of AF genes organized in a cluster was investigated as one
major objective in my study (using nor-1 gene as an example, Chapter 3). The results
confirmed that the AF gene cluster exerts a positive position-dependent regulatory
influence on the nor-1 promoter (Chiou et al., 2002).

Partially-duplicated AF gene cluster. A second, partially-duplicated AF cluster
was discovered in some strains of A. parasiticus (Liang et al., 1996; Chang and Yu,
2002). This partial AF gene cluster contains homologs of aflR-aflJ-adhA—estA -norA-ver1
plus omtB, yet is missing verA-avnA -verB-ava, omtA, ordA, and vbs present in the
complete AF cluster in A. parasiticus strain 56775 (F ig. 1.2) (Chang and Yu, 2002). The
truncated AF gene cluster contained several mutations and did not seem to be functional.
Several structural genes were found to be defective, including pre-termination of verIB
(Liang et al., 1996) and norA2, point mutations in adhAZ, and a large deletion in omtB
(Chang and Yu, 2002).

Mutations were also found in the second copy aflR (aflRZ), and a specific point
mutation in nucleotide residue 896 (with a T to A substitution) can be used to
discriminate between aflRI and aflRZ (Cary et al., 2002). However, this mutation did not
result in a pre-terminated gene product. AflR2 was not able to activate transcription in a
yeast two hybrid system (Cary et al., 2002), and it could not complement the loss of
functional activity of the primary AflRl (Chang and Yu, 2002). An A. parasiticus aflR
knockout mutant AF 810, was disrupted in aflRI but not aflRZ. This strain did not
produce AF, any precursors in the pathway, or transcripts of aflRZ and other AF genes

such as nor-I , ver-I , and omtA analyzed by Northern hybridization analysis. However,

13

using RT-PCR, the transcripts of aflRZ and these AF genes were detected at low levels
(Cary et al., 2002). The aflRI knockout strain AF S10 was used as a negative control in
my AF protein localization studies (Chapter 4 and 5).

Sugar-utilization gene cluster. Downstream of the AF gene cluster following a
5 kb spacer region, a sugar utilization gene cluster consisting of 4 genes (sugR, hxtA,
gch, and nadA) was identified (Yu et al., 2000). Expression of hxtA, which encodes a
hexose-transport protein, followed the same timing of expression under AF inducing
conditions (glucose minimum salt medium) as AF genes such as omtA (Yu et al., 2002).
This sugar utilization cluster, or the hexose-transporter thA specifically, may play a role
in regulation of AF biosynthesis, since simple sugars, e.g. glucose, were found to induce

AF biosynthesis (Buchanan and Lewis, 1984; Skory et al. 1993; Miller, 2003).

SECONDARY METABOLISM AND FUNGAL DEVELOPMENT

Association between secondary metabolism and fungal development. In
Aspergillus as well as many other microorganisms, the production of secondary
metabolites is frequently associated with developmental processes, e.g., sporulation
(Bennett, 1983; Horinouchi et al., 1994, Guzman-De-Pena and Ruiz-Herrera, 1997;
Adam and Yu, 1998; Adams et al., 1998; Calvo et al., 2002). The hypothesis is best
supported by the gram-positive bacterial genus Streptomyces in which morphological
differentiation resembles development of filamentous fungi. In Streptomyces griseus,
secondary metabolism and formation of aerial mycelium, a structure required for
subsequent generation of spores, are both triggered by a 'y-butyrolactone containing
molecule, A-factor (Horinouchi and Beppu, 1994; Kato et al., 2002).

G-Protein signaling pathway. In A. nidulans, coordinated regulation of asexual
sporulation and biosynthesis of the mycotoxin sterigmatocystin (ST) has been
demonstrated (Adams and Yu, 1998; Adams et al., 1998). The balance between

vegetative growth, sporulation, and ST production is regulated by two antagonistic

14

factors, F adA (Fluffy Autolytic Dominant), and F le (Fluffy, Low _b_rlA; brlA is a
conidial vesicle specific gene) involved in a G-protein signaling pathway (Yu et al., 1996;
Hicks et al, 1997)(Figure 1.3). The fadA gene encodes the Ga subunit of a heterotrimeric
G protein and activation of this gene facilitates cell proliferation and vegetative growth,
while flbA encodes a RGS (Regulator of G protein signaling) domain that antagonizes the
function of F adA. Inactivation of the F adA G protein signaling pathway is required for
both ST biosynthesis and asexual sporulation (Hicks et al, 1997). Conversely,
developmental activation requires at least partial inactivation of fadA. Dominant
activating mutations of fadA as well as loss of function mutations offlbA resulted in
inhibition of both conidiation and ST production. Conversely, overexpression offlbA or
a dominant interfereing mutation of fadA leads to precocious ST gene expression and ST
accumulation, as well as unscheduled conidiation (Hicks et al, 1997).

FluG. Another factor F luG, acting up-stream of this G-protein linked pathway, is
associated with synthesis of an extracellular, low molecular weight diffusible factor
leading to asexual conidiation and ST production (Lee and Adams, 1994). Loss of
function mutations in fluG resulted in a complete block in ST production and sporulation,
similar phenotypes to loss of function mutations in brlA and dominant activation
mutation in fadA (Hicks, et al., 1997). The FluG-dependent asexual sporulation pathway
seems to be in competition with the vegetative growth pathway promoted by FadA.
Whereas F adA negatively regulates ST biosynthesis, F luG is believed to promote ST
production via inactivation of FadA signaling (D'Souza et al., 2001). The detailed
mechanisms of this G-protein signaling pathway observed in Aspergillus are under
investigation.

AF biosynthesis and conidiation. Secondary metabolites and conidiation are
usually observed when cells enter the stationary phase of grth (Betina, 1995). In A.

flavus and A. parasiticus, AF was detected in developmental structures including conidia

15

Carbon Stress

 

 

 

 

 

 

 

Nitrogen Stress
\
.- .1.“ s . ,1-“7‘ . . ,g \ .. _. _-
T
fle- IlbD- fIbB\
2"
Receptor? brIA -—> Development
/
0+5» .. .L
Extracellular T""';’I"" """"""""" '
FluG signal ' ’ .
“1:le ;
D —. *zzFadi-GTP.” Colony proliferation
and Aytolysls .v
Growthslgnal GDP
? ———->afIR—> ST production
. J
k. A ,

 

Figure 1.3. The G-protein signaling pathway involved in ST biosynthesis in A. nidulans.
(Adapted from Adams et al., 1998)

16

and sclerotia (Wicklow and Schotwell, 1983). A. parasiticus with a defect in fluP, a gene
that encodes a large polyfunctional enzyme with sequence similarity to polyketide
synthetases, did not develop conidiospores and AF production was reduced 3 to 5 fold
compared to the wild type (Zhou et al., 1999). Production of ST/AF in Aspergillus spp.
typically accompanies conidiation but the converse is not always true because ST/AF can
be synthesized in liquid and submerged shake culture where conidiation does not usually
occur (Kale et al. 1994). However, Guzman-de-Pena et al. (1998) indicated a lower level
production of AF in liquid culture compared to AF production fi'om the same mass of A.
parasiticus grown on solid media with conidiation. Also, chemicals that suppress
conidiation such as fluoroacetic acid (ReiB, 1982) and diaminbutanone, an ornithine
decarboxylase inhibitor (Guzman-de-Pena et al., 1998) reduced production of AF in A.

parasiticus.

LOCALIZATION OF PROTEINS INVOLVED IN AF BIOSYNTHESIS

One of the goals of my study was to determine where AF proteins are synthesized
within a cell and to understand how these enzymatic activities associate with each other
in order to make the final AF molecules.

Penicillin biosynthesis as an example. In Penicillium chrysogenum,
biosynthesis of the secondary metabolite penicillin is similar to ST/AF biosynthesis in
Aspergillus because penicillin biosynthetic genes are also organized in a cluster. Three
gene products, ACVS (aminoadipyl-cysteinyl-valine tripeptide synthetase), IPNS
(isopenicillin N-synthetase), and AT (acyl CoA: 6-iso-penicillin-N-acyltransferase)
involved in penicillin biosynthesis have been isolated and localized (Muller et al., 1991).
ACVS was thought to be associated with a Golgi-like organelle (Kurylowicz et al., 1987)
and vacuoles (Lenderfeld et al., 1993), but later was confirmed to be localized in the
cytoplasm (Van der Lende et al., 2002). The second enzyme IPNS was also found to be a
cytosolic protein (Van der Kamp et al., 1999; Van der Lende et al., 2002). By irnmuno-

17

transmission electron microscopy, only the final enzyme activity, AT, was found
associated with microbodies (aka. peroxisomes) (Muller et al., 1995).

Organelles possibly involved in AF biosynthesis. This section reviews several
organelles that may be associated with AF biosynthesis in A. parasiticus.

l) Microbodies. Microbodies (aka. peroxisomes) are found universally in fungi,
but the number, shape and function are highly variable (Markham, 1995). Even within a
species, generation of microbodies may be affected by various factors including age,
developmental stage, and nutrient source. Microbodies contain a variety of enzyme
activities and are potentially involved in a wide range of degradative and biosynthetic
activities (Carson and Cooney, 1990). There are two types of microbodies with different
enzyme activities. The peroxisome contains catalases and enzymes involved in fatty acid
B-oxidation, while the glyoxysome houses the glyoxylate cycle and also enzymes
involved in fatty acid B-oxidation (Keller et al., 1991). Microbodies are common in
fungal spores, where they are associated with lipid inclusions (Maxewell et al., 1977).

Two types of Beroxisomal Targeting Sequence (PTS) have been identified. PTSl
is a carboxyl-terminal tri-peptide sequence Ser—Lys-Leu (SKL) and conserved variants
(S/A/(CXK/R/HXL/MXGould et al., 1987). The second matrix protein targeting signal,
PTSZ, is an N-terminal nonapeptide motif with the consensus sequence (R/K)(LN/I)X5-
(H/Q)(L/A)(Subramani, 1996). However, many peroxisomal enzymes have neither a
PTSl nor a PTSZ motif (Waterham and Cregg, 1999). None of the AF proteins contains
the conserved PTSl or PTSZ located at the C-terminus or N-terminus, respectively.
However, the PTSl sequence was detected in AflJ, OmtA and OrdA, but not at the C-
terrninus (predicted by PSORT version 6.4).

2) Woronin bodies. Woronin bodies are highly refractile particles seen in over
50 species of filamentous fimgi (Markam and Collinge, 1987; Jedd and Chua, 2000).
This organelle is bounded by a single bilayer membrane and the major component is

protein (Head, 1989). The Woronin body has a spherical to oval shape and the size is

18

approximately 0.1-0.75 pm, slightly larger than the septa] pore (Markham, 1995;
Markham and Collinge, 1987; Jedd and Chua, 2000).

Woronin bodies are usually located in the septa and have been suggested to
function by plugging the damage within the septa] pore in response to hyphal injury
(Colling and Markham, 1985; Tenney et al.,- 2000). This function indicates that the
Woronin body is capable of moving within hyphal cells, implying a possible role in
transport of AF. The phytotoxin prehelminthosporol has been found localized in
Woronin bodies in Bioplaris sorokiniana (Akesson, et al., 1996).

The Woronin body was suggested to be derived fiom peroxisomes in A. nidulans
(V alenciano et al., 1998). A gene encoding a major Woronin body protein, hex], was
recently identified in Neurospora crassa (Tenney et al., 2000; Jedd and Chua, 2000).
The Hexl protein contains the C-terminal PTSI sequence, suggesting a relationship
between the woronin body and the peroxisome. In Neurospora crassa , Woronin bodies
are associated with catalase-negative peroxisomes (J edd and Chua, 2000).

3) Vesicles. Vesicles are small but they are important and very abundant in
filamentous fimgi (Markam, 1995). Vesicle sizes vary with diameters ranging from
approximately 30 nm to 400 nm (Markam, 1995). A high density of vesicles is usually
clustered in the hyphal apex; they are involved in transport of various structural proteins,
enzymes, and other cellular components including lipids and polysaccharides (Markam,
1995). The vesicles are able to migrate through the septal pore and the movement is
associated with microtubule components (Hoch et al, 1987). There are many types of
vesicles such as wall vesicles, microvesicles, and chitosomes, with respect to their
particular functions.

4) Vacuoles. Vacuoles are the largest organelles in eukaryotic microorganisms.
The large vacuoles of fungi have many features similar to animal lysosomes. Fungal
vacuoles, similar to higher plant vacuoles, are acidic compartments containing hydrolases

and lysosomal activities (Klinlosky, 1997; Ashford, 1998); the acidic environment is

19

regulated by a vacuolar ATPase (Wada and Anraku, 1994). The functions of vacuoles are
associated with intracellular homeostasis, storage of metabolites, and degradation of
proteins (Thrumm, 2000). This organelle is also involved in intracellular protein
transport such as endocytosis, protein secretion, and intracellular protein localization
(Wendland et al., 1998). The structure and dynamic activities of fungal vacuoles indicate
that this organelle has the capacity to interact and transport material over long hyphal
distances (Ashfor, 1998).

Under the microscope, the vacuole appears as a large, hemi-spherical hollow-like,
fluid -filled organelle surrounded by a single membrane (Markham, 1995; Ohsumi et al.,
2001; Ohneda et al., 2002). Vacuoles are often observed in aged cells and they are not
usually present in the active apical tip regions (Markham, 1995). Some large fungal
vacuoles may contain small membrane-bound vesicles, which are derived from fusion of
multivesicular bodies with large vacuoles (Hoch and Staples, 1983).

Over 40 vacuolar protein-sorting (vps) and vacuolar-morphology (vam) associated
genes have been identified in Saccharomyces cerevisiae (Raymond et al., 1992). One
vacuole resident protein, carboxypeptidase Y (CPY), has been used as a vacuole marker
protein (Johnson et al., 1987). In A. nidulans, cypA that encodes CPY (Ohsumi et al.,
2001), and vpsA involved in vacuolar biogenesis (Tarutani et al., 2001) have been cloned
and characterized.

Klinonsky (1998) reviewed vacuole-related functions and suggested that of all the
organelles, the vacuole may be the most adaptive compartment as a delivery mechanism.
In fed-batch fermentation of P. chrysogenum, during the switch fiom the rapid growth to
the production phase as nutrients became limited, a high degree of mycelial fragmentation
was seen along with heavy vacuolation (Paul et al., 1994), implicating an association of
vacuole biogenesis and secondary metabolism. In addition, immunogold-transmission
electron microscopy localized OmtA, an enzyme involved in a late stage AF biosynthesis,

in the cytoplasm as well as in the vacuole in cells located near the substrate surface of the

20

colony (Lee et al., manuscript submitted). In this regard, the vacuole is hypothesized to
play a role in AF biosynthesis.

5) Endoplasmic reticulum (ER). The ER is a continuous membrane system, yet
various domains are present in this single organelle that perform different functions
(Markhem, 1995; Voeltz et al., 2003). The ER appears to make physical contact with the
membranes of many organelles such as nuclei (nuclear envelopes) and vacuoles, and
dynamic contact with cytoplasmic vesicles via the continuous traffic system (Beckette et
al., 1974; Wedlich-SOlender et al., 2002). This organelle is involved in protein folding
and several post-translational modifications in the ER lumen, and translocation of
proteins (e.g. secretory proteins) across the ER membrane (V oletz et al., 2003; Ellgaard
and Helenius, 2003).

N-glycosylation of proteins along the secretory pathway is an important protein
quality control mechanism. This proof reading system serves beyond the levels of
transcription and translation to ensure native conformation of newly synthesized proteins
and the fidelity of cellular functions (Ellgaard and Helenius, 2003; Roth, 2002; Helenius
and Aebi, 2001; Parodi, 2000). After proteins reach their native tertiary and quaternary
structure, they subsequently translocate to their final cellular destination. In general, the
proteins involved in the secretory pathway are targeted to the plasma membrane, vesicles,
or extracellar space for secretion (Roth, 2002; Ellgaard and Helenius, 2003). As part of
the ER quality control, misfolded proteins are translocated fiom the ER to the cytoplasm
and subjected to an ER-associated degradation (ERAD) process and subsequently
destroyed by the 26S proteosome (Kostova and Wolf, 2003).

According to protein sub-cellular localization prediction programs listed in the
Expasy molecular server (http://us.expasy.org), proteins targeted to the ER usually require
an N-terminal hydrophobic signal peptide and N-glycosylation site(s) (Asn-X-Ser/Thr)
(Signal IP; Target P; NetGlyc 1.0; Ausubel et al., 2003). BR resident proteins firrther

require an ER-retention signal, KDEL, within the C-terminal peptide sequence (Teasdale

21

and Jackson, 1996; Harter and Wieland, 1996). Among AF proteins, only VBS was
confirmed to be N-glycosylated (Silva et al., 1996) and appeared to be modified in the ER
(see Chapter 5). Based on protein sequence prediction, only one other protein, OrdA
involved in the final step of AF biosynthesis, contains a putative N-terminal signal
peptide and one N-glycosylation site, suggesting OrdA may be associated with the ER
secretory pathway like VBS (ExPasy molecular server). Both proteins were not predicted
to be ER resident proteins. Association of AF proteins along the ER secretory pathway
may be significant to their protein functions, and possibly play a role in regulation of AF

biosynthesis.

CONFOCAL LASER SCANNING MICROSCOPY (CLSM) IN THE STUDY OF
FUNGAL BIOLOGY

Introduction to CLSM. Microscopy serves as an useful tool to conduct studies
in cell biology including analyses of cell dynamics, morphogenesis, cytochemistry, fungal
spore structure and development (Spear et al., 1999). Fungal cellular organelles can also
be visualized; however, the resolution of light microscopy has limitations (theoretical
maximum resolution is 0.2 pm) in traditional light microscopes (Paddock, 2000). In
conjunction with fluorescence-based techniques such as immuno-fluorescence labeling as
well as the recent application of the green fluorescent protein (OF P), fluorescence tagged
cellular targets can be tracked by fluorescence microscopy. In the past, conventional epi-
fluorescence microscopy has been widely used. Confocal laser scanning microscopy
(CLSM) provides a wide range of advantages over traditional light microscopy (and epi-
fluorescence microscopy) although CLSM is based on the fundamental principles used to
light microscopes.

The first stage-scanning confocal microscope was invented in mid-1957 by
Minsky for imaging of neural networks in unstained brain cells (Minsky, 1961; 1988).

Modern confocal microscopes are based on the configuration of this prototype (with the

22

use of a pinhole) and an enhanced model became commercially available in 1987 after
air-cooled lasers and PC computers and software became prevalent (Amos et al., 1987).

The characteristic feature of CLSM is the ability to generate "confocal images"
that eliminate the interference of out-of-focus light captured by the detector. This
confocal effect is achieved by use of a confocal pinhole positioned between the specimen
and the detector in the light path (Figure 1.4). CLSM utilizes one or more focused beams
of light illuminated by the laser (light amplification by stimulated gmission of radiation)
which scans across the sample generating "optical sections" at various focal planes.
Thus, contrast and resolution are enhanced on each single optical section. This feature is
particularly beneficial for viewing thick samples since CSLM is capable of scanning
through specimens (up to hundreds of microns in thickness) instead of mechanical
sectioning of samples and acquiring serial optical sections (Z-sections) (Whallon, 1998).

Image quality enhancement by CLSM. Since CLSM utilizes the basic
configuration of the light microscope, the instrument resolution per se, that is influenced
by the wavelength of light, the objective lens, and properties of the specimen itself, is not
improved drastically (Paddock, 2001). However, the final image resolution of modern
CLSM is improved by the following factors. First, stable multiple wavelength lasers
instead of conventional tungsten-halogen and/or mercury lamps are used to generate
bright point sources of light to scan the samples. Typically air-cooled lasers are used
including the helium-neon laser emitting at 543 nm and 633 nm and the argon-ion laser
emitting at 488 nm and 514 nm. The laser spot size affects instrument resolution; similar
to scanning electron microscopy, the smaller the beam spot, the greater the resolution

(Czymmeck et al., 1994). The spot size is also affected by the wavelength of light and
nmneric aperture (NA) of the objective lens (Dresolution= 0.61X [Wavelength]/

NAobjective)- For example, the spot size for the 488 nm laser with a high NA (1.4) lens

is approximately 250 nm (Czymmeck et al., 1994). In addition, a laser focused to a small

23

Beam Splitter

L
I aser CollectingLens

p
To
A M Mom...
T p
| .

/. Objective Lens Confocal Unit

With Pinhole
Focal Plane
Outof-Focus Area

—Path 01 ln‘Focus Light

 

Figure 1.4. Diagram of the light path in a confocal instrument. (Adapted from Czymmek
et al., 1994)

24

point (as equipped in the CLSM models; Zeiss 210 and Zeiss LSM5 PASCAL, Carl-Zeiss
Inc., Germany) generates better resolution than a laser focused through a slit (as equipped
in Meridian Insight®, Meridian Instrument, Inc., Okemos, Mich.). Theoretical resolution
of CLSM under 488 nm excitation with a 1.4 NA oil objective was estimated to be 160
nm (Brelje et al., 1993) and approximately 130 run under ideal conditions (Carlsson and
Aslund, 1987).

Second, image detection in modern confocal microscopes utilizes sensitive and
low noise detectors (e.g., photomutiplier tube (PMT) or a digitized charge-coupled device
(CCD) camera, particularly the cooled-charged CCD with reduced noise read out. In
addition, advanced CLSM is equipped with fast microcomputers with image processing
capability and elegant software to analyze images. High quality output images can be
processed directly in a computer imaging system or recorded on hard copy devices
(Poddock, 2001). Advanced image analysis programs also allow CLSM digitized images
to be processed further, for example, reconstruction of three-dimensional (3-D) images
from Z-stacks (Czymmek et al., 1994), or pixel analysis to quantify fluorescence intensity
(Spear et al., 1999; Chapter 5 in this dissertation).

CLSM imaging. CLSM is capable of generating many types of microscopic
images. General light microscopy applications, such as phase contrast, polarization, and
differential interference contrast (DIC), can be conducted via CLSM with enhanced
image qualities. However, an important feature of CLSM is detection of fluorescence.
The characteristic functions are described as follows.

1) Optical sections. The basic unit of confocal microscopy is the single optical
section (confocal image). The contrast of an optical section is enhanced more intensely
than non-confocal images generated by epi-fluorescence microscopy.

2) Z-series. For one specific field of interest, several optical sections can be
collected as a series to generate a-Z-stack. The CLSM is commonly equipped with a

stepping motor to control the movement of the stage with a pre-determined distance (step

25

size). Using a macro program, CLSM can acquire images from the top (or from the
bottom) of a specimen, scan through the sample with a pre-set step size, and acquire as
well as store the whole series of optical sections on the hard drive to generate a Z-series
(Z-stack). The Z-series can be further processed into a 3-D presentation, stereo images,
or viewed as a movie depending on the program capabilities. A representative Z-series of
A. parasiticus SU-l condiophores stained with SYTOX green nuclear staining dye
(Molecular Probes, Eugene, OR) is shown in Fig. 1.5.

3) Phi-z sectioning. Optical sectioning is not limited to the horizontal focal plane
but also can be collected in the vertical plane to generate a phi-z or x-z section. Phi-z
sectioning is accomplished by scanning the laser across a defined single line. It allows
signals to be detected fi'om another angle of view. For example, the microbodies of A.
parasiticus SU-l immuno-labeled with anti-SKL antibodies followed by secondary
antibodies conjugated with Alexa 488 probe are displayed in a single optical section (Fig.
1.6, A). Analysis by phi-z scanning indicated that several rrricrobodies are located inside
the hyphae (Fig. 1.6, B).

4) Time-lapse and live imaging. Since CLSM is capable of scanning through the
sample without mechanical sectioning, live cells may be studied. This application may be
somewhat difficult to accomplish, for example, if the experiment requires fixation of the
cell, or if the target signals are not stable under illumination by the laser. In addition, the
position of the specimen has to be maintained under the microscope. Nevertheless, time-
lapse imaging allows visualization of dynamic events at a rate close to realtime or video-
rate imaging (Whallon, 1998).

5) Fluorescence imaging. CLSM is a powerful tool for fluorescence imaging.
The argon-ion laser emits at 488 nm and generates optimum excitation spectra for many
commonly used fluorescence probes, including FITC (fluorescein isothiocyanate (Abs/Em

490/523 nm), Alexa 488 (Abs/Em 495/519 nm), the red-shift green fluorescent protein

26

 

Figure 1.5. CLSM Z-series of A. parasiticus SU-l conidiophores stained with SYTOX
green fluorescence dye. Fungal thin sections (4 pm) derived from colony fraction 82 (24-
48 h growth) of a 72 h-old SU-l colony grown on YES agar were stained with SYTOX
green dye (Molecular Probes). (l) - (10): Ten optical sections with step size (2 section
interval) 750 nm. (A) An extended focus image of this CLSM Z-stack. (B) A bright field
image in the middle of the Z-stack. Bars, 25 um.

27

Phi-z scan _>

 

Figure 1.6. CLSM single optical section and phi-z scan of A. parasiticus SU-l
microbodies. Fungal thin sections (4 pm) derived from colony fraction 32 (24-48 h
growth) of a 72 h-old SU-l colony grown on YES agar were immuno-labeled with anti-
SKL antibodies (Zymed) followed by secondary antibody conjugated with Alexa 488
probe (Molecular Probes). CLSM was performed using a Zeiss 210 confocal scanning
microscope (Carl Zeiss Inc.). (A) Single optical section. (B) Phi-z scan. The arrows

indicate the location of microbodies inside the cell. Bar, 5 pm.

28

variant, EGF P (A bs/Em 488/507 nm), and a nuclear staining dye, SYTOX green (Abs/Em
504/523 nm). CLSM can also perform multiple fluorescence detection, for example, dual
channel detection of green and red fluorescence on the same image (Chapter 5, Fig. 5.9).
This is particularly useful if these fluorophores have minimum overlapping emission
spectra to avoid cross-talking signals. Mixed krypton-argon gas lasers are popular for
multiple-wavelength confocal microscopy because they emit at three well-separated
wavelengths (488 and 568 and 647 nm) allowing detection of green to far-red
fluorophores (e.g. F ITC; Lissarnine rhodarnine B, Abs/Em 570/590 nm; Cy5, Abs/Em
649/666 nm)(Ausubel et al., 2003). However, some fluorophores excite at shorter waves
(e.g., DAPI [4,'6-diamidino-2-phenylindale, Abs/Em 458/461 nm] nuclear stain, and
CMAC [7-amino-4-chloromethylcoumarin, Abs/Em 353/466 nm] vacuolar stain);
detection of these probes requires CLSM equipped with expensive UV lasers.
Photobleaching may cause significant problems for fluorescence detection,
particularly when samples are illuminated with high intensity lasers. It is a quenching
phenomenon that causes fading of fluorescence signals, yet the mechanism is not fully
understood. One important class of bleaching is photodynarnics and this involves
interaction of fluorophores with light and oxygen. The amount of photodamage depends
on the concentration of molecular oxygen and the distance between the fluorophore,
molecular oxygen and other cellular constituents (Herman, 1998). Protection against
photobleaching can be achieved by reducing excitation energy or exposure time;
however, this will also reduce the yield of the signal. Other methods to prevent
photobleaching include deoxygenation of solutions; use of singlet oxygen quenchers such
as histidine, diphenylisobenzofuran, or crocetin (a water soluble acratenoid); or antifade
reagents such as n-propyl-gallate. Several antifade mounting media are also
commercially available (e.g. Vectashield [Vector] and Prolong antifade mounting media

[Molecular Probes]).

29

In summary, CLSM is capable of generating higher contrast and resolution images
compared to traditional light microscopy by reducing background interference
significantly. Advanced image detection systems and analysis programs render this
instrument more capable of analyzing cellular structures or other aspects in detail. In this
dissertation, CLSM is an important tool to study localization of proteins involved in AF

biosynthesis in the filamentous fungus A. parasiticus (Chapter 4 and 5).

30

CHAPTERZ

A rapid DNA extraction method for screening large numbers of Aspergillus
transformants via polymerase chain reaction

ABSTRACT

The efficiency of screening fungal transformants using available methods,
particularly in large quantity, has been a problem in fungal molecular biology. A rapid
DNA extraction method was developed for screening Aspergillus transformants via
polymerase chain reaction (PCR). Fungal cells were collected directly from colonies on a
culture plate into microcentrifuge tubes containing TE buffer (10 mM Tris-HCl, 1.0 mM
EDTA, pH 8.0). The cell suspension was boiled for five minutes to release nucleic acids.
After a brief centrifugation step (12,000 g for 5 min), fungal cell extracts (supernatant)
were obtained. PCR of diluted fungal extracts with appropriate primers resulted in
amplicons up to 2.5 kb with Taq DNA polymerase and up to 3.5 kb with PfiiTurbo DNA
polymerase. The procedure was successfully utilized for amplification of native fungal
genes as well as identification of the integration site of plasmid sequences after

homologous recombination.

INTRODUCTION

Fungal transformation provides an important tool for the study of fungal biology.
However, limited numbers of transformants can be screened conveniently due to labor-
intensive assay procedures. The presence of the fungal cell wall limits the efficiency of
genomic DNA isolation. Disruption of the fungal cell wall to release nucleic acids is
usually accomplished by the laborious procedure of grinding under liquid nitrogen. DNA
purification also commonly includes proteolytic enzyme digestion, phenol/chloroform
extraction, and precipitation of DNA (Ausubel et al., 2003; Borgia et al., 1994). The

quantity and quality of the genomic DNA isolated by these procedures tends to vary

31

significantly. A high content of polysaccharides and nucleases represent significant
obstacles for obtaining high quality fungal genomic DNA. Hence, my goal was to
develop a rapid, inexpensive, and efficient DNA extraction method for screening large
numbers of fungal transformants.

I present a simple procedure to obtain crude DNA templates directly from fungal
colonies that was successfully used to amplify endogenous fungal genes as well as to
detect the integration site of plasmid sequences afier homologous recombination.
Twenty-four fungal colonies can be easily assayed in a three to four hour period including

time for PCR and electrophoretic analysis.

MATERIAL AND METHODS

Aspergillus parasiticus CS10 (wh-l , ver-l pyrG) (Skory et al., 1990) was
transformed with plasmid pAPGUSNP (Fig. 2.1, A) and the transformants were selected
on Czapek-Dox medium (CZ medium, Difco) for ability to synthesize uridine
monophosphate. As soon as the prototrophic transformants sporulated (approximately 2
days), spores were transferred onto duplicate CZ plates with a toothpick. After three days
(or as soon as colonies could be observed), colonies were analyzed. A sterile 200 pl
plastic pipette tip (Lee and Cooper, 1995) was used to transfer a portion of the fungal
colony (including conidiospores and mycelia) into a microcentrifuge tube containing 100
pl TE buffer (10 mM Tris-HCl, 1.0 mM EDTA, pH 8.0). This process was repeated up to
three times to ensure that sufficient fungal cells were transferred. Tubes were heated in a
boiling water bath for 5 min and placed on ice. The samples were centrifuged at 12,000 g
for 5 min at 40C. The supernatant (firngal extract) containing crude DNA templates was
transferred into a new microcentrifuge tube. The extract was further diluted with TE
buffer (undiluted, 1/5, 1/20, 1/ 100). In most assays, 2.5 pl of the 1:5 dilution resulted in

good amplification in a 50 pl PCR reaction. The remainder of the fungal extract was

32

stored at -80°C and could be successfully analyzed by PCR for at least three months.
Diluted samples stored at 4°C could be analyzed by PCR for at least one week.

PCR analysis was conducted on DNA extracted from A. parasiticus transformants
and the recipient strain, CS10. Four primer pairs were used in the assay (Table 2.1).
Primer pair J L136 and JL137 produced a 1.4 kb exon H fiagment from the fungal gene
encoding versicolorin B synthase (vbs) (Silva et al., 1996). Three primer pairs, JL99 and
JL100, JL102 and JL103, JL221 and JL222 were designed to determine the integration
site of pAPGUSNPyrG in the 5' region of nor-1, in the 3' region of nor-I or at the pyrG
locus repectively. One primer from each pair targeted a sequence located within the
fungal genome. The other primer targeted one site within the plasmid sequence (Figure
2.1).

A 50 pl PCR reaction mixture consisted of 2.5 pl of the diluted fungal extract, 20
mM Tris-HCl pH 8.8, 10 mM KC], 10 mM (NH4)2SO4, 0.1% Triton X-100, 0.1 mg/ml

BSA, 0.5 - 1.0 pM of each primer, 200 pM each dNTP, 2.0 - 3.0 mM MgC12 and either

2.5 units of Taq thermal stable DNA polymerase (5 U/pl, Gibco BRL) or 2.5 units of
PfuTurbo DNA polymerase (2.5 U/ pl, Stratagene). PCR cycling parameters were
adjusted depending on primer design and the target size (Table 2.1). PCR was performed
in a Perken-Elmer GeneAmpli System 9600 thermal cycler. All PCR reactions were
initialized by a denaturation step at 95°C for 3 min followed by 35 cycles of denaturation,
annealing and extension. Reactions were terminated with an extension reaction at 72°C
for 10 min. PCR products (10 p] of reaction) were observed by electrophoresis on a 1%

agarose gel using standard procedures (Ausubel et al., 2003).

RESULTS AND DISCUSSION
DNA was extracted by the boiling procedure from seven representative
transformants and the recipient strain, CS10 (Figure 2.2). The DNA was analyzed by PCR
with appropriate primer pairs. A 1.4 kb fragment was generated from all eight

33

Table 2.1. Primer sequences and PCR cycling parameters used for analysis of fungal

genomic DNA templates.

 

Primer Target Primer Sequence

PCR
cycling parameters

 

Endogenous vbs ExonII

JL136 1.4 kb 5' AGG crc GAA AGG CGC ATA CGA 3'
JL137 5' GT0 GTC TAC TGC CCA Gcc ATC 3'

Integration of pAPGUSNP, 3' region of nor-l gene
JL102 2.1 kb 5' CGC AAG GTG AGG GIT CGA ACC GAG G 3'

JL103 5' CCG CAG CAG GGA GGc AAA CAA TGA A 3'

Integration of pAPGUSNP, 5' region of nor-1 gene

JL 99 3.2 kb 5'TITCACGGGTI‘GGGGTI‘TCTACAGG3'
JL100 5'GACGGGGAACCTCTI‘TACAAACATC3'

Integration of pAPGUSNP at pyrG

JL221 3.5 kb 5' CCA GAA AAr GCG AAG GTA AGT GCT TC3'
JL222 5' GCG ACA CGG AAA TGT TGA ATA crc AT3'

95°C 1', 66°C 1', 72°C 2'

95°C 1', 65°C 1', 72°C 3'

95°C 1', 62°C 1', 72°C 5'

95°C 1', 55°C 1', 72°C 5'

 

34

Figure 2.1. Homologus recombination of pAPGUSNP into the A. parasiticus
chromosome and primers used in integration analysis. (A) Plasmid map of pAPGUSNP.
(B) pAPGUSNP integrated within the 5' region of the nor-1 gene generates a 3.2 kb PCR
fiagment using primers J L99 and J L100. (C) pAPGUSNP integrated within the 3' region
of the nor-1 gene generates a 2.1 kb PCR fragment using primers JL102 and JL103. (D)
pAPGUSNP integrated at the pyrG locus generates a 3.5 kb PCR fiagment using primers
JL221 and JL222. (E) Amplification of endogenous vbs gene fi'agment in A. parasiticus
using primers JL136 and JL137.

35

  
     
   
 
     

 

 

Plasmid
Xhol 10600
al 2100
Sac19600
pA PGUSNP Sca|2900
12100 bp
Scal8300
3' nor-1
Chl 5000
C|a|5900 Sphl 5200
Genome
D B C
X 5' X X 3'
I:
genomic pyrG genomb nor-1 vbs
E
-1 JL13
5'nor-1 genome no; genogne 6'>‘.JL137
W~~”~— _ _ _ .
1.4 kb
JL100> ‘JL99
3.2 kb
nor-1 genome
C 5' 3'
_ _ _ _ s\\\\\‘
vbs
1.4 kb
JL103WL102
2.1kb
pyrG genome
Mx//zxx/xw K\\\\V
vbs
1.4 kb

JL221, ‘JL222
3.5 kb
Figure 2.1

36

1081012345672. ACS101234567¢X174

3.5kb —>
2.1kb —>

 

Figure 2.2. Amplification of PCR targets using the rapid DNA extraction technique.

The assay was performed on eight A. parasiticus colonies, including the host strain, CS10,
and seven representive transformants. (A) Integration analysis using various primer sets.
Cell extracts of C S10, and transformants 100, 149, 164 were PCR amplifed with primers
JL221 and JL222. A 3.5 kb target fragment was produced in transformants 100, 149, 164
Canes 1-3) but not the host strain CS10. PCR of cell extracts from 57 and 104 (Lanes 4-
5) with primers JL102 and JL103 produced a 2.1 kb fragment. PCR of cell extracts from
transformants 17 and 24 with primers JL99 and JL100 generated a 3.2 kb fragment (Lanes
6-7). All DNA extracts produced the endogenous 1.4 kb vbs fragment with primers JL136
and JL137 (shown in B).

37

strains fi'om the endogenous vbs gene (Figure 2.2, B). Using the same fungal extracts, the
three sets of primer pairs for integration analysis successfully determined the integration
sites of pAPGUSNP within the genome in the transformants (Figure 2.2, A). With primer
pair JL221 and JL222, PCR amplification of cell extracts from transformants 100, 149
and 164 (Lanes 1-3) produced a 3.5 kb PCR product, indicating that the plasmid
integrated at the chromosomal pyrG locus while PCR amplification of cell extracts from
the host strain CS10 did not generate this fiagment. PCR amplification of cell extracts
fiom transfonnants 57 and 104 (Lanes 4 and 5) produced a 2.1 kb fiagment with primer
pair JL102 and JL103, indicating integration of pAPGUSNP within the 3' region of the
nor-I gene while cell extracts of transformants 17 and 24 (Lanes 6 and 7) produced a 3.2
kb fiagment indicating integration within the 5' region of the nor-1 gene. The integration
location of pAPGUSNP in each strain was confirmed by Southern hybridization analysis
(data shown in Chapter 3). After demonstrating the utility of this DNA extraction
procedure, over 500 transformants were screened. The assay correctly identified the site
of integration in over 80% of the transformants.

Several genomic DNA extraction methods for plant tissues and fungi were
developed to simplify the extraction procedure and to allow large numbers of samples to
be analyzed. However, these methods still have some drawbacks. For example, some
procedures applied to fungi (Cenis et al. 1992) and plant tissues (Steiner et al. 1995)
required mechanical grinding of tissues for isolation of DNAs. Alternatively, cell
disruption was achieved by temperature extremes in Podospora anserina (Lecellier and
Gael 1994) or by alkaline lysis in Neurospora and Aspergillus to prepare DNA for dot
blot analysis (Min et a]. 1995). DNA extracted from whole colonies of yeast cells treated
with NaOH was used directly in PCR (Wang et al. 1996), but the method could not
reliably amplify targets greater than 600 bp (Steiner et a]. 1995).

The fungal cell wall can also be digested by specific enzymes to facilitate release

of nucleic acids. For example, yeast cells were treated with zymolase (Ling et al. 1995)

38

and the filamentous firngus Aspergillus niger was treated with Novozyme 234 (Zeijl et al.
1998). Both assays could amplify targets up to 3 kb. However, these methods utilized
costly enzymes, which require additional time for preparation and cell wall digestion.

A rapid spore PCR assay that used conidiospores ofAspergiIlusfizmigatus directly
fiom culture plates for screening gene disruption events has been reported (Aufauvre-
Brown et a]. 1993). Low numbers of Magnaporthe. grisea spores (104-10°) were also
used directly in PCR without extraction of DNA (Xu and Hamer 1995). Spores from five
strains of filamentous fungi were treated with microwave irradiation to facilitate release
of DNA into the supernatants (Ferreira and Glass 1996). However, it is important to note
that these direct spore PCR methods could only amplify PCR targets smaller than 1 kb.
Attempts to produce a 1.4 kb PCR fragment using the direct PCR methods described
above were not successful.

The fungal DNA extraction procedure presented here is simple and effective.
Only a small quantity of the firngal cells must be acquired directly from the culture plates
eliminating the need for extended culture. Fungal colonies grown for 48 h to 72 h were
easily analyzed. Alternatively, colonies could be replicated onto several assay plates
reducing the growth time to less than 36 hours. I observed that too much material could
introduce a high level of contaminants for PCR, reducing effectiveness. This problem
was overcome in most cases by increasing the dilution of the sample prior to PCR
analysis.

The initial version of our rapid DNA extraction utilized sonication to release
DNA. Sonication has been used successfully with other cell types, e.g. bacterial cells
(Ausubel et al. 2003). In our laboratory, conidiospores of Aspergillus parasiticis SU-l
(2x107 spores/ml) were disrupted by sonication (output control level 4, 10 pulse x 4
cycles, SONIFIER® cell disruptor W—350, Branson Sonic Power Co.). An increase in
optical density at 260 nm (A260= 0.34) suggested the release of nucleic acids into the

lysates. High temperature treatment of cell lysates was attempted to inactivate cellular

39

nucleases and other PCR inhibitory factors. However, when the efficiency of sonication,
boiling, and a combination of the two to release DNA was tested (A260= 0.701, 0.962,
0.989 respectively), the boiling process by itself resulted in the effective release of nucleic
acids. Both procedures provided DNA templates for PCR amplification (data not shown).
However, the boiling method was nearly as effective and did not require expensive
equipment.

Boiled fimgal lysates may still contain PCR inhibitory factors which cannot be
heat inactivated. This problem can be minimized by diluting the extracts with TE buffer
to the point where sufficient DNA templates remain while minimizing the level of
inhibitors. In our hands, a 2.5 pl aliquot from the 1:5 dilution of the fungal extract worked
well in the 50 pl PCR reaction. Dilution of the original cell extract of transformants
generated fi'om other fungal species may have to be adjusted appropriately.

Success of PCR amplification also depended on the reaction conditions (e.g.,
optimum MgC12 concentration, proper annealing temperature and extension time). I
observed that the activities of specific thermal stable DNA polymerases played an
important role in determining success of amplifying long target fragments. T aq
polymerase (Gibco BRL) inconsistently amplified targets up to 2.5 kb using this DNA
extraction procedure. Addition of PCR adjuncts and stablizers such as BSA (10-100
pl/ml) and DMSO (1-10%) (Stratagene manual) had limited effects on enhancement of
product yield. However, PfirTurbo polymerase (Stratagene) efficiently amplified target
fiagrnents up to 3.5 kb. Longer targets were not tested in this assay.

Conditions for sample storage also play a role in determining success of the
analysis. In our hands, fungal colonies on plates could be stored at 4°C for at least a
week before analysis. Once samples were collected into TE buffer, fungal extracts
(supernatants) could be used for up to one week of storage at 4°C and up to three months
of storage at -80°C. However, the sooner the extracts were used, the better the results.

Preparation of fungal extracts for only 10 to 12 samples at one time is recommended.

40

This appears to reduce degradation of nucleic acids which occurs during sample
preparation.

This rapid DNA extraction method for PCR analysis provides an easy,
inexpensive, convenient and efficient assay procedure for screening a large number of
Aspergillus transformants and may be applicable to a wide variety of other fungal genera.
This rapid procedure to prepare DNA templates fi'om fungal colonies provides sufficient
purity for PCR analysis of the site of plasmid integration into the A. parasiticus genome
(Chiou et al., 2002). This development was critical for current and future studies on the
regulation of the AF promoters (such as nor-1 and ver-I) 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 makes screening of large numbers of fungal transformants practical.

ACKNOWLEDGEMENTS
Special thanks to David Wilson, a former student working in the laboratory, for
providing the primers to analyze the plasmid integration sites at nor-1 by PCR.
This rapid DNA isolation/PCR method is published as part of the data in a
manuscript "Chiou, C. H., M. 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".

41

CHAPTER3

Position-dependent regulation of the nor-I promoter in the
aflatoxin gene cluster

ABSTRACT

Expression of most of the AF genes within the 75 kb gene cluster in Aspergillus
spp. appears to be co-regulated but the mechanism is not yet clear. I hypothesized that
regulation of AF genes is cluster position-dependent; i.e., AF genes are positively
regulated within the AF gene cluster. In this study, the nor-1 gene that encodes
norsolorinic acid reductase, an early AF pathway enzymatic activity, was used as a study
model. Promoter activity of nor-1 was compared within the AF gene cluster (nor-1
locus) and outside the cluster (pyrG locus). A plasmid pAPGUSNP containing the nor-1
promoter sequence fused to a reporter gene (uidA, B-glucuronidase, GUS) plus a
selectable marker (pyrG) was constructed to study positional effects on nor-1 promoter
function. Transformation of pAPGUSNP into C810 (ver-I, wh-I, pyrG) resulted in
homologous recombination at 5' nor-1, 3' nor-1, or the pyrG locus. The plasrrrid
integration site was screened by a rapid DNA extraction/PCR assay and confirmed by
Southern hybridization analysis. GUS reporter activity was analyzed by a solid-culture
GUS assay. Among 178 transformants analyzed, 42 transformants showed positive GUS
activity (24.0%). Based on a rapid PCR assay, random selection of 14 GUS+
transformants indicated that plasmid integration occurred within the nor-1 gene (11
within 5' nor-1 and 3 within 3' nor-1). Southern hybridization analysis confirmed that of
10 GUST isolates analyzed, they all harbored pAPGUSNP within the nor-1 locus (7 at 5'
nor-1; 2 at 3' nor-1; 1 showed multiple integration at both sites). None of the 11
transformants that harbored the norzzGUS fusion at the pyrG locus expressed detectable

GUS activity. The results agree with previous observations that reduced ver-I and nor-1

42

promoter activity is expressed at niaD (encodes nitrate reductase). These data support

our hypothesis that expression of AF genes is cluster position-dependent.

INTRODUCTION

Many of the genes involved in aflatoxin (AF) biosynthesis in Aspergillus spp.
have been identified and they were found clustered within a 75 kb chromosomal DNA
region (Trail et al., 1995; Payne and Brown. 1998; Woloshuck and Prieto, 1998.).
Expression of these genes appeared to be coordinately regulated although details of the
regulatory mechanism are not yet clear.

The nor-1 gene in the filamentous fungus Aspergillus parasiticus encodes a
ketoreductase activity that converts norsolorinic acid (NOR), the first stable intermediate
in the AF biosynthetic pathway, to averantin (AVN) (Trail et al., 1994; Zhou and Linz,
1999). The nor-1 gene is expressed under AF inducing conditions, concurrent with many
other AF genes (Skory et al., 1993; Zhou and Linz, 1999). The timing of nor-I
expression and protein accumulation was consistent with the accumulation of AF (Skory
et al., 1993).

Previously, Liang et al. (1997) reported preliminary evidence that the organization
of AF genes in a cluster may play a role in regulation of AF biosynthesis. The ver-I
promoter, which drives expression of a gene encoding a ketoreductase activity involved
in a middle step of AF biosynthesis, was fused to uidA (encodes B-glucuronidase; GUS)
to generate a reporter plasmid, pHD 6.6, containing niaD (encodes nitrate reductase) as a
selectable marker. pHD6.6 was transformed into A. parasiticus NR—l (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-I
promoter activity when compared with single copy integration at the ver-I locus. Similar
results have been shown for two norzzGUS constructs located at niaD and the nor-1 locus

(Wilson, 1998; Chiou et al., 2002; Miller, 2003). GUS reporter activity was detected at

43

high levels when the nor-1 promoter was positioned at the nor-1 region within the AF
gene cluster but was not detected at niaD. However, in A. parasiticus, niaD expression is
under the control of nitrogen catabolite repression mediated by areA and the pathway
specific induction of nirA. In the presence of nitrate, niaD is positively regulated. On the
other hand, nitrate has been shown to down regulate AF biosynthesis (Payne and Brown.
1998). Lack of ver-I and nor-1 expression at niaD could have resulted from niaD-
dependent regulation.

In this study, nor-1 promoter activity expressed at another chromosomal locus,
pyrG, was investigated. The pyrG gene encodes orotidine monophosphate (OMP)
decarboxylase that converts OMP to uridine monophophate (UMP), a compound essential
for cell growth (Skory et al., 1993). Unlike niaD, expression of pyrG has not been
reported to be associated with regulation of AF biosynthesis. In this study, a plasmid
(pAPGUSNP) containing four major DNA fiagrnents was constructed. This plasmid
includes the 5' nor-I fragment (upstream promoter sequences of the nor-1 gene), a GUS
reporter (obtained fiom the Escherichia coli uidA gene which encodes B-glucuronidase),
the 3' nor-I fragment (the nor-I terminator and down stream sequences), and a selectable
marker pyrG. Transformation of pAPGUSNP into A. parasiticus CS 10 (ver-I , wh-I ,
pyrG), an uridine auxotroph, resulted in homologous recombination of pAPGUSNP into
5 ' nor-1, 3' nor-1 or pyrG locus. The plasmid integration site was screened by a rapid
DNA extraction/PCR assay and confirmed by Southern hybridization analysis. GUS
activity was analyzed by a solid culture GUS assay. The results showed that the nor-1
gene was positively regulated only when the promoter was positioned at its native
localization (nor-1). When the nor-1 promoter was placed outside the cluster (at pyrG),
expression of nor-I was reduced dramatically. This result was in agreement with our
previous studies on positional effects at nor-1 and niaD (Wilson, 1998; Miller, 2003).
These data provide strong support for the hypothesis that the AF gene cluster exerts a

positive position-dependent regulatory influence on the nor-1 promoter.

44

MATERIALS AND METHODS

Strains and growth conditions. Escherichia coli DHSa F'e [F'/endAI hstI 7
(r1; mk')supE44thi-1 recAI gyrA (Nalr)Are1A1 (lacZYA argF)u169: (m80 AlacZ M15)
(Gibco BRL, Life Technologies Inc., Rockville, MD) was used to amplify plasmid DNA
using standard procedures (Ausubel et al., 2003). A. parasiticus CSlO (wh-I, ver-I,
pyrG) (Skory et al., 1990) was derived from A. parasiticus ATCC 36537 (wh-I, ver-I) by
nitrosoguanidine mutagenesis and was used as a recipient for the reporter construct
carrying the pyrG selectable marker (pAPGUSNP). To grow the recipient strain CSlO,
YES medium or Czapek Dox medium (CZ, Difco, Detroit, MI) was supplemented with
uridine (100 pg/ml).

Construction of pAPGUSNP. Plasmid pAPGUSN (Fig. 3.1, A) contains the
nor-1::GUS backbone and was generated by Dr. Frances Trail, Department of Botany and
Plant Pathology, Michigan State University. The _5'n_0r-L fragment (3.0 kb) contains
flanking sequences, promoter, and nucleotides encoding the first 21 amino acid residues
of Nor-1. This fiagment was fused in fiarne to a 1.8 kb GUS reporter (B-glucuronidase)
derived from E. coli uidA. The 3'_n_0£1_ fragment (2.0 kb) contains nucleotide sequences
encoding the last 6 amino acids of Nor-1, the transcription terminator, and flanking
sequences. To generate pAPGUSNNP, pAPGUSN was digested with Nhel and the ends
filled-in with Klenow fiagment of DNA Poll (Roche Molecular Biochemicals,
Indianapolis, Ind.). The 2.5 kb pyrG gene was amplified by PCR with Taq DNA
Polymerase (Gibco BRL, Rockville, MD), pPG3J (Skory et al., 1990) as template, and
appropriate primers (5'-GTAGAAGTTGTTCAGTAGCTGATGG-3' and 5'-
CGAGTATCACAGTCAGGACTCCACG-3'). The resulting PCR fiagment was end-

45

Figure 3.1. Restriction endonuclease maps of plasmids containing the nor-1 ::GUS
construct related to this study. Only restriction endonuclease sites relevant to this study
are shown. (A) pAPGUSN contains the nor-I ::GUS backbone (generated by Dr. Trail,
Department of Botany and Plant Pathology, Michigan State University). (B) pAPGUSNP
contains the nor—1::GUS fusion plus the pyrG selectable marker (Chiou, this study). (C)
pAPGUSNNB contains the nor-1 ::GUS fusion plus the niaD selectable marker
(generated by Wilson, 1998). (D) pBGN3.0 contains the nor-1 ::GUS fusion plus the
niaD selectable marker (generated by Miller, 2003).

46

PSI] 800
P511 1000

    
  
     
 
 
 
 
 

A

Sea] 8300

Sea] 2 100

pAPGUSN
9600 bp

 

  
 
 
  
 
 

   

   
 
  
  
 

Sca12900
Hindlll 3000 Xhol ]
3' nor-l
Clal 5930 ‘3‘: L Xhol 10600 I 'i "H
C Ia] 5000 EcoRl 4800 pyrG Sea] 2100
Sac] 9600
I Sea] 2900
PAPGUSNP iinmd1113000
12100 bp
0 Xhol | P51] 800 Amp
Pstl 1000 GUS
. A Sea] 8300
\ Sea] 2100
5' “0“ - Hindlll 3000 -
Pstl 13800 5;: 5‘0“ 480°
‘ Cla15000
Cla15900 Sph15200
ma], pAPGUSNNB
Gus
17000 bp
/ 1' EcoRl 4300 D
/ Clal 5000
3' nor-l / /
/ / Sea] “5400 Ascl 418
/ Clal 5030 Clal 1218

Clal2118

Sea] 8300

\\

1 i
I'fI'\Noti 4226

i' Sea] 4326

7 Sea] 5126

P511 6226
P511 6426
Not] 7226

Pstl I0400

Figure 3.1

47

polished by Pfir polymerase (Stratagene, LaJolla, CA) and blunt-end ligated into
pAPGUSN to generate pAPGUSNP (Fig. 3.1, B).

Fungal transformation. Plasmid pAPGUSNP (5 -10 pg) was transformed into
protoplasts of CS10 (an uridine auxotroph) according to a method described previously
(Horng et al., 1990; Skory, et al., 1993). Uridine prototrophic transformants were
selected on CZ agar medium with 20% sucrose as osmotic stabilizer. Agar plates were
incubated in the dark at 290C for 3 to 5 days.

Identification of the site of plasmid integration. (i) Rapid DNA extraction
procedure for PCR analysis. Plasmid site of integration was screened by a rapid DNA
extraction and PCR method discussed in Chapter 2 (Chiou et al., 2002). Transforrnants
were grown until sporulation (usually 3 to 5 days) and conidiospores were transferred to
duplicate Czapek Dox agar plates using a sterile toothpick or sterile pipette tip. After 3 to
4 days (or as soon as colonies could be observed), colonies were analyzed. A sterile 200
pl plastic pipette tip was used to transfer a portion of the fungal colony (including
conidiospores and mycelia) into a microcentrifuge tube containing 100 pl TE buffer (10
mM Tris-HCI, pH 8.0, 1 mM EDTA). The process was repeated up to 3 times to ensure
that sufficient fungal cells were transferred. Tubes were heated in a boiling water bath for
5 min and placed on ice. The samples were centrifuged at 12,000 x g for 5 min at 40C.
The supernatant (fungal extract), containing DNA, was transferred to a new
microcentrifuge tube. The extract was diluted with TE buffer (undiluted, 1:5, 1:20, and
1:100). In most assays, 2.5 pl of the 1:5 dilution resulted in good amplification in a 50 pl
PCR reaction. The remainder of the fungal extract was stored at -800C and could be
analyzed by PCR for at least 3 months. Diluted samples stored at 40C could be analyzed
for at least 1 week.

(ii) PCR analysis. PCR analysis was conducted on DNA extracted from A.
parasiticus transformants and’the recipient strain A. parasiticus CS10. The primer pairs

utilized depended upon the plasmid vector used and the site of integration analyzed

48

(Table 3.1). A 50 p1 reaction consisted of 2.5 pl of the diluted fungal extract, 20 mM
Tris-HCl (pH 8.8), 10 mM KC], 10 mM (NH4)ZSO4, 0.15% Triton X-100, 0.1 mg/ml
BSA, 0.5 to 1.0 pM of each primer, 200 pM of each dNTP, 2.0 to 3.0 mM MgC12, and
either 2.5 units of Pfu Turbo polymerase (2.5 U/pl, Stratagene, LaJolla, CA) or 2.5 units
of Taq polymerase (5 U/pl) (Gibco BRL). PCR cycling depended on primer design and
the target size (Table 3.1). PCR was performed in a Gene Ampli System 9600 (Perkin-
Elmer) thermal cycler. All PCR reactions were initiated by a denaturation step at 950C
for 3 min followed by 35 cycles of denaturation, annealing, and extension. Reactions
were terminated with an extension reaction at 720C for 10 min. PCR products were
observed by electrophoresis on a 1% agarose gel using standard procedures (Ausubel et
al., 2003).

Solid-culture GUS activity assay. A solid-culture assay to analyze GUS activity
was developed based on published procedures (Kolar et al., 1991; Verdoes et al., 1994).
Transforrnants were transferred from selective medium to agar plates containing an
aflatoxin-inducing solid growth medium (YES) overlaid with sterilized circular 82 mm
Nytran Plus membranes (Schleicher and Schuell Inc., Keene, NH). Plates were
incubated in the dark at 290C for 36 to 55 h. Membranes with attached fungal colonies
were removed from the surface of the agar plates and immersed in liquid nitrogen for 30
s. Membranes were allowed to thaw and immersed again in liquid nitrogen for 30 s.
Afier thawing, membranes were incubated in GUS substrate solution (60 mM NazHPO4,
40 mM NaH2P04, 10 mM KC], 1 mM MgSO4, 0.27% B-mercaptoethanol. 0.04% X GLU
[Gold Biotechnology, St. Louis, MO]) for up to 24 h. Positive GUS activity resulted in a
blue color in the colony

Southern hybridization analysis. The plasmid integration site was confirmed by
Southern hybridization analysis. Fungal genomic DNA was purified from A. parasiticus

transformants grown in YES liquid shake culture (the recipient strain CSlO required

49

Table 3.1. Primer sequences and PCR cycling parameters used for analysis of site of

integration using DNA templates prepared by the rapid method.

 

Primer Target Primer Sequence PCR cycling

size (kb) parameters

 

Integration in 5' region of nor-1 gene

PCR 1
IL 99 3.1 kb S'rrr CAC GGG TTG GGG TTT CTA CAG G 3' 950C 1', 620C 1', 720C 5'
JLIOO 5'GAc GGG GAA ccr crr TAC AAA CAT c3'

Integration in 3' region of nor-1 gene

PCR 2

JL102 2.1 kb 5'CGC AAG GTG AGG GTT CGA ACC GAG G3' 950C 1', 680C 1', 720C 3'
JL103 5'CCG CAG CAG GGA GGc AAA CAA TGA A3'

Integration in pyrG

PCR 3

JL221 3.5 kb 5'CCA GAA AAT GCG AAG GTA AGT GCT Tc3' 950C 1', 550C 1', 720C 5'
JL222 5'GCG ACA CGG AAA TGT TGA ATA crc AT3'

Endogenous vbs ExonII

(Control)
JL136 1.4 kb S'AGG CTC GAA AGG CGC ATA CGA3' 950C 1', 680C 1', 720C 2'
JL137 5'GTG GTC TAC TGC CCA GCC ATC3'

 

50

supplementation with 100 pg/ ml Uridine) for 48 h at 29°C with five, 6 mm glass beads
(Skory et al., 1993). Southern hybridization analysis was performed using digoxigenin
labeled probes and a non-radioactive hybridization detection procedure provided by the
manufacturer (Roche Molecular Biochemicals, Indianapolis, Ind). The restriction
enzymes and probes used to analyze the integration site are summarized in Table 3.1.
The integration scheme and resulting blots are shown in Figures 3.2 and 3.3.

(i) 5' nor-1 integration. Genomic DNA of transformants was digested with Xhol
and probed with Probe 1. DNA fi'om A. parasiticus CS10 generates a 5.8-kb band while
DNA from a 5' integrant of pAPGUSNP generates 13.0 and 3.4-kb bands (Fig. 3.2).

(ii) 3' nor-1 integration. Genomic DNA of transformants was digested with ScaI
and probed with Probe 2. DNA from A. parasiticus CSlO generates a 3.0-kb band while
DNA from a 3' integrant of pAPGUSNP generates 4.0 and 4.4-kb bands (Fig. 3.2).

(iii) Integration at pyrG. Genomic DNA of transformants was digested with
SphI and SacI, respectively, and probed with a 2.5-kb pyrG fragment generated by PCR
(Probe 3). DNA fi'om A. parasiticus CSlO digested with SphI generates a 12.6-kb band
while DNA from a pyrG integrant generates 16.0 and 8.8-kb bands. CSlO genomic DNA
digested with SacI generates a 8.1-kb band while DNA from a pyrG integrant generates
5.9 and 14.3-kb bands. (Fig. 3.3).

RESULTS
Generation of pAPGUSNP. Plasmid pAPGUSNP was constructed by blunt-end
ligation of a 2.5 kb pyrG fragment into a 9.6 kb nor-1 ::GUS backbone derived from
pPAPGUSN (Fig. 3.1, A). The rate of successful blunt-end ligation was low. The ligated
plasmids were transformed into E. coli DHSOL. Among 1200 transformants generated,
only three transformants contained the 2.5 kb pyrG gene; these were identified by DNA
dot blot analysis using digoxigenin-labeled pyrG fragment (2.5 kb) as a probe.

Restriction digestion analysis indicated only one transformant harbored the correct

51

Figure 3.2. Southern hybridization analysis of site of integration in pAPGUSNP
transformants. Restriction endonuclease map of pAPGUSNP is shown on the top. (A)
Schematic for Southern hybridization and PCR analyses of pAPGUSNP integration into
5' region of nor-1 and Southern hybridization data. A. parasiticus genomic DNA was
digested with XhoI and probed with Probe 1. Single crossover integration of pAPGUSNP
into the 5’ region of nor-1 results in two bands, 19 kb and 3.4 kb, and the disappearance
of the 5.8-kb wild-type band. (B) Schematic for Southern hybridization and PCR analysis
of pAPGUSNP integration into the 3' region of nor-1 and Southern hybridization data.
Genomic DNA was digested with Sea] and probed with Probe 2. Single crossover
integration of pAPGUSNNB into the 3’ region of nor-1 results in two bands, 4.0 kb and
4.4 kb, and the disappearance of the 3.4-kb wild-type band. Abbreviations: X = Xhol; Sc
= ScaI; C = Clal.

52

  
 
 
 
  
     
 
 
   

Xhol 10600

Sac] 9600

Scal 8300

A 5’nor—1

X ScSc X CCSc

 

— nor-l
Probe 1
'2 .. .._,.-. -..__ «..I
X 5.8 kb X X
. PCR]
I "5 "' ..
- GUS pyrG - nor-l
I I l
l 19 kb I 3.4 kbl
3.1 kb

C810 24 37 4O 60 84 17 32 57

-231 kb
'- Vl‘;"""- 9.41m
— 6.5 kb
5.8 kit—>3!
— 4.31m
-— 2.3 kb
Figure 3.2

pAPGUSNP
12100 bp

11

Seal 2100

Sea] 2900
HindIII 3000

EcoRI 4800
Cla15000

B 3’nor-1
X

X ScSc X CCSc

ENE.

r-tl'r
. ., a],
Or- 4* '1‘”
_H ..

nor-

3.4 kb—

 

CS‘IO 24 37 40 60 84 17 32 57

—23.1kb
U VI.-
. _ 9.4 kb
«7 — 6.5 kb
4.41m.> '- .. V “' .. "‘ “a.
4.0 kb—> - we - 4.3 kb
3.4kb—> “U 'w-”~~
— 2.3 kb

53

Figure 3.3. Southern hybridization analysis of pAPGUSNP integrated at pyrG.
Restriction endonuclease map of pAPGUSNP is shown on the top. Genomic DNA was
digested with Sphl (A) or SacI (B) and hybridized with the digoxigenin-labeled pyrG
probe (2.5 kb, Probe 3). With Sphl restriction endonuclease digestion (panel A), single
crossover integration of pAPGUSNP into pyrG results in two bands, 16 kb and 8.8 kb,
and the disappearance of the 12.6-kb wild-type band. With Sacl restriction endonuclease
digestion (panel B), single crossover integration of pAPGUSNP into pyrG results in two
bands, 14.3 kb and 5.9 kb, and the disappearance of the 8.1-kb wild-type band.
Abbreviations: Sp = SphI; Sa = Sad.

54

........

 
 
 
 
 
  
 

Xhol l0600

Sea] 2100

Sacl9600
Sea] 2900

HindIIl 3000

   

pAPGUSNP
12100 bp

   
   

Sea] 8300

EcoRI 4800
Clal 5000

 

 

 

 

 

 

 

1——1
A SphI 3.5 kb B $5161
(308- GUS-
l I I l
csm1oo 21121 133 135 142145154 14 95 105 57 cs101oo 21121 133 135 142149154 14 95 105 57
l
l
i
15.0 115...: ,
12.5 lib->5» ~ -' w l
8'8 kb- 14.3 115->- . I
5.1 kb-
5.9115. 1
i
i
1
Figure 3.3

55

construct. This plasmid (pAPGUSNP) carries nor-1:: GUS and pyrG oriented in opposite
directions, resulting in divergent transcription the two genes (Fig. 3.1, B).

Transformation of A. parasiticus C810 with pAPGUSNP. Transformation of
the A. parasiticus uridine-auxotrophic strain CS 10 with plasmid pAPGUSNP was
performed in three separate experiments. The pyrG+ uridine-prototrophic transformants
were selected on CZ media with 20% sucrose. In total, 178 transformants were
generated and subsequently analyzed using a solid-culture GUS activity assay.

Solid-culture GUS assay. All of the pyrG+ transformants selected on CZ
medium were analyzed for expression of GUS on AF inducing medium (YES agar plate).
Solid-culture GUS assay of these transformants (178 in total) resulted in identification of
42 GUS+ isolates (24%). Randomly selected GUS+ or GUS' isolates were subsequently
analyzed by PCR or Southern hybridization to determine the plasmid integration sites.

Determination of pAPGUSNP integration site within the chromosome.
Fourteen GUS + isolates were randomly selected and screened by a rapid PCR assay for
site of integration (Fig. 3.4). The rapid PCR assay utilized a quick and easy template
preparation procedure (see Materials and Methods) and the results were comparable to
PCR using purified genomic DNA (Chiou et al., 2002). All 14 GUS + isolates had
pAPGUSNP integrated at 5' nor-1 (11 isolates) or 3' nor-1 (3 isolates). The site of
integration of pAPGUSNP was confirmed in 4 of these isolates and 6 additional
randomly selected GUS + isolates by Southern hybridization analysis (Fig. 3.2). Of these
10 isolates, 7 had pAPGUSNP integrated at 5' nor-1 and 2 carried pAPGUSNP at 3' nor-
1. Isolate 57 was likely a multiple integrant of pAPGUSNP; these data suggested this
isolate carried one copy of pAPGUSNP at both 5' and 3' nor-1 (Fig. 3.2). In contrast, of
11 isolates with pAPGUSNP integrated at pyrG, none expressed GUS activity (Southern
hybridization analysis shown in Fig. 3.3).

56

A 1051012345673.

3.1 kb
3.5 kb —>

2.1 kb —>

 

B 108101234 567¢x174

1.4 kb

 

Figure 3.4. PCR analysis to detect site of integration in pAPGUSNP transformants.
Template DNA for 5' and 3' nor-1 and pyrG integration assays was prepared with a rapid
boiling procedure. (A) GUS+ and GUS' transformants (lanes 1 through 7) and CS10
(negative control) were analyzed for 5' nor-1, 3' nor-I , or pyrG integration. Primers
PCR3 were used to detect pyrG integration (lanes 1, 2, and 3; GUS' transformants) and
generated a 3.5-kb PCR fragment. Primers PCR2 were used to detect 3' nor-1 integration
(lanes 4 and 5; GUS+ transformants) and generated a 2.1-kb PCR fragment. Primers
PCR] were used to detect 5' nor-1 integration (lanes 6 and 7; GUS+ transformants) and
generated a 3.1-kb PCR fiagrnent. (B) DNA from all 7 transformants and the control
strain CS10 was amplified with a primer pair specific for exon III of the versicolorin B
synthase gene (positive control). A 1.4-kb PCR fragment was expected in all fungal
isolates. Lanes containing molecular size markers in panels A and B are represented by
AHindlIl digest and ¢x174 HaeIII digest. Nucleotide sequences of all primers are found in
Table 3.1.

57

YES

 

Figure 3.5. Solid-culture GUS activity assay on CSIO transformants harboring
pAPGUSNP. The recipient strain CS10 (control) and representative pyrG+ transformants
were inoculated onto Nytran membranes overlaid onto YES agar medium (panels C and
D) or YES supplemented with 5 mg uridine per plate (panels A and B). A GUS assay
was conducted on filters in panels B, C and D. Transformants carried a single copy of
pAPGUSNP integrated at 5' nor-1 (transformants 17 and 32), at 3' nor-I (transformant
37), or at pyrG (transformants 100 and 121). Transformants 24 and 60 carried multiple
copies of pAPGUSNP with at least one copy integrated at 5' nor-I . Transformant 57
carried multiple copies of pAPGUSP with at least one copy integrated at 3' nor-1.
Transformants l4 and 133 carried multiple copies of pAPGUSNP with at least one copy

at pyrG and no copies at nor-I .

58

Confirmation of pyrG expression in pyrG+ transformants. Expression of pyrG
in A. parasiticus CS 1 0 (used as the recipient) was analyzed under these aflatoxin-
inducing growth conditions (Fig. 3.5). Growth of A. parasiticus CS10 (wh-I, ver-I,
pyrG) on YES required supplementation with uridine (100 pg/ml) while transformation
of CSIO with a functional pyrG gene overcame this requirement demonstrating that a
functional copy of pyrG must be present and expressed for growth of CS10 on YES

medium.

DISCUSSION

Based on reduced ver-I promoter activity expressed at the niaD locus observed
previously (Liang et al., 1997), I hypothesized that location within the AF cluster
provided a positive regulatory influence on AF gene promoter function under aflatoxin-
inducing growth conditions. Suppression of promoter activity of another AF gene, nor-1,
at niaD was also demonstrated prior to the current study (Wilson, 1999; Miller, 2003).
However, alteration of promoter activity may be specifically affected by the niaD gene
since niaD is only expressed under non-AF inducing conditions using nitrate as the sole
nitrogen source; otherwise the gene is repressed. Reduction of ver-I and nor-1 promoter
activity at the niaD locus may be a consequence of regional control. Thus, investigation
of positional effects of AF promoter activity at an additional locus provides supporting
data for our previous observations. In this study, promoter activity of nor-1 located in the
pyrG locus and its native chromosomal location was investigated. The pyrG locus was
chosen for two reasons. 1) The selectable marker (pyrG) that encodes uridine
monophosphate decarboxylase was cloned previously (Skory et al., 1990). 2) Expression
of pyrG is not associated with regulation of AF biosynthesis, unlike niaD.

The results generated in this study were consistent with results generated in
analysis of nor-1 promoter activity within the AF gene cluster and at niaD (Chiou et al.,

2002). First, the percentage of GUS+ transformant is similar (approximately 24%). This

59

study utilized a plasmid pAPGUSNNP (Fig. 3.1, B) containing the nor-1::GUS fusion
(derived from pAPGUSN, Fig. 3.1, A) plus pyrG as a selectable marker; while
pAPGUSNNB (Fig. 3.1, C) and pBNG3.0 (Fig. 3.1, D) contained a similar nor-1::GUS
construct and niaD as a selectable marker. Transformation of pAPGUSNNB and
pBNG3.0 into A. parasiticus NR1 (niaD) resulted in 23% (25 out of 149 transformants
analyzed) and 25% (29 out of 117 transformants analyzed) GUS+ transformants,
respectively (data summarized in Table 3.2).

Second, these data confirmed the preliminary observation by Liang et al. (1997)
that chromosomal location plays a role in regulation of AF gene expression. Expression
of nor-1::GUS at the nor-1 locus was regulated similarly to the native nor-I gene, while
placement at two other positions (niaD and pyrG) outside of the aflatoxin gene cluster
resulted in no detectable GUS activity. The targeting of reporter constructs to specific
chromosomal locations for promoter analysis has been advised by other investigators
studying expression of fungal genes (Hammer, and Timberlake, 1987; Timberlake and
Marshall, 1989; 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 AF genes is reasonable, since removal of AF
reporter fusions from the AF gene cluster results in reduced GUS expression.

An alternative explanation for the lack of GUS activity at the niaD and pyrG loci
is that a transcriptionally inactive chromatin structure exists at these two sites. We think
that such inactivation is unlikely for two reasons. First, NR-l transformed with
pGAPN2B (B—tubulin::GUS) expressed GUS activity at similar levels at 24, 48, and 72 h
when integrated at the niaD locus, although the level of GUS activity in the pGAPN2B

transformants was approximately ten-fold lower than GUS activity in D8D3

60

Table 3.2. Summary of GUS activities and plasmid integration sites exhibited in A.

paraciticus transformants carrying nor: :GUS fusion in CS 1 0 transformants carrying

pAPGUSNP (Chiou, in this study). and NR-l harboring pAPGUSNNB (Wilson, 1998)

and pBNG3.0 (Miller, 2003).

 

 

 

 

 

 

9'38"“ pAPGUSNP“ pBNG3.0b pAPGUSNNBc

Target locus pyrG niaD niaD

Recipient strain CS10 NR-1 NR-1

it of Total transformants 178 117 149

# of GUS+ 42 29 35

% GUS + 24% 25% 23%

“WNW "WW“ “Chiou leller cwrison (1995)
(this study) (2003)

PCR on GUS+ transformants 14

Integration at 5' nor-1 11

Integration at 3' nor-1 3

Southern analysis on GUS+ 10 21 11

transformants

Integration at 5' nor-1 7 12 4

Integration at 3' nor-1 2 9 6

Multiple intgegration at 5' and 3' nor-1 1

Unverified 1

Southern analysis for 608- 11 22 6

transformants

Integration at 5' nor-1 0 0 0

Integration at 3' nor-1 0 0 0

Integration at pyrG 1 1

Integration at niaD 22 2

Single crossover at niaD 4

 

 

61

(pAPGUSNNB integrated at 3' nor-1; Wilson, 1998) at 48 and 72 h. In contrast,
transformant D8D4 (pAPGUSNNB integrated at niaD; Wilson, 1998), when grown under
the same conditions, produced no detectable GUS activity at any of these time points
(data not shown). Second, in transformants carrying pAPGUSNP, the nor-1 promoter
carried on this plasmid was not active at the pyrG locus even though growth on YES
medium required expression of the pyrG gene. In addition, several pAPGUSNP
transformants had no detectable GUS activity even when they carried multiple copies of
the plasmid (none integrated at nor-1). In contrast, other multiple integrrants with at least
one copy of pAPGUSNP integrated at nor-l were GUS": Based on these data and
preliminary data reported by Liang et al. (1997) I propose that aflatoxin gene expression
is influenced by an enhancer element.

The clustering 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 an enhancer element 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-1
region, and at least 1.8-kb downstream the transcription termination site in the 3' region
of nor-1 or within the nor-I coding region. Because similar position-dependent
expression was observed with the ver-1::GUS reporter construct (Liang et al., 1997), it
also is possible that the same cis-acting element is influencing the regulation of both nor-
1 and ver-I . Studies performed on the SpoCl gene cluster in Aspergillus 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; Timberlake and Barnard, 1981). The physical linkage
of aflatoxin biosynthesis genes also may have a regulatory role. Studies designed to

establish such a phenomenon and to determine the nature of the regulatory mechanism,

62

may eventually lead to a broader understanding of the expression of secondary

metabolism genes and the ability to manipulate their expression in microorganisms.

ACKNOWLEDGEMENTS
We thank Dr. Frances Trail, Department of Botany and Plant Pathology at
Michgan State University, for providing plasmid pAPGUSN.
Data in Chapter 3 is published in Applied and Environmental Microbiology in a
manuscript entitled "Chromosomal location plays a role in regulation of aflatoxin gene

expression in Aspergillus parasiticus" (Chiou et al., 2002).

63

CHAPTER 4

Immuno-localization of Nor-1, Ver-l, and OmtA proteins involved in aflatoxin
biosynthesis in Aspergillus parasiticus

INTRODUCTION

The biosynthesis of aflatoxin is a complex process that involves at least 18
enzyme activities (Minto and Towsend, 1997; Payne and Brown, 1998, Cary et al., 2000).
The mechanisms that coordinate these enzyme activities to synthesize AF within a cell
are still not clear. In addition, the distribution of these AF enzymes in a fungal colony
and their sub-cellular localization within a fungal cell have yet to be determined. The
objective of this study was to investigate the distribution and sub-cellular location of
representative enzymatic activities in the aflatoxin biosynthetic pathway. This study
focused attention on Nor-1 (Trail et al., 1994), Ver-l (Skory et al., 1992) and OmtA (Yu
et al., 1993) that catalyze early, middle and late enzymatic steps in the aflatoxin
biosynthetic pathway, respectively. Nor-1 is a NADPH-dependent keto-reductase
involved in the conversion of norsolorinic acid (NA) to averantin (AVN) (Zhou and Linz,
1999). Ver-l is another keto-reductase involved in conversion of versicolorin A (VERA)
to demethyl-sterigmatocystin (DMST)(Skory et al., 1992; Liang et al., 1996). OmtA is a
methyltransferase that specifically converts sterigmatocystin (ST) to O-
methylsterigrnatocystin (OMST) by transfer of a methyl group fi'om S-
adenosylmetlrionine (SAM) (Yu et al., 1993; Lee et al., 2002). Our laboratory produced
polyclonal antibodies (PAb) against Nor-l (Zhou, 1998; Lee et al., manuscript
submitted), Ver-l (Liang, et al., 1997), and OmtA (Lee et al., 2002) to localize these
enzymes in A. parasiticus SU-l and CSIO. An omtA disruptant strain, LW 1432 (th,
ver-I , omtA), was also generated to serve as a negative control (Lee et al., 2002).

A model for coordinate regulation of sterigmatocystin (ST) production (an AF

biosynthetic pathway intermediate in A. parasiticus) and asexual sporulation has been

64

described in A. nidulans (Hicks, et al., 1997; Adams et al., 1998). Based on this model, I
hypothesized that the enzymes involved in AF biosynthesis are localized within specific
developmental structures; this study focused on an asexual spore-bearing structure, the
conidiophore. At the sub-cellular level, according to the complexity of this AF
biosynthetic pathway, I hypothesized that enzymes (or some of the enzymes) involved in
AF biosynthesis are compartmentalized or closely associated within a cell. Physical
interaction of these enzymes may facilitate efficient and accurate catalysis.

Preliminary cellular localization of Nor-1 and Ver-l in the wild type AF
producing strain A. parasiticus SU—l was performed by Liang (1996) and Zhou (1998)
using a slide-culture cel] preparation procedure followed by indirect irnmuno-
fluorescence microscopy. However, artifacts were generated by this sample preparation
method for several reasons. First, growth conditions and colony morphology were
different from colonies generated by center-inoculation of conidiospores on solid media.
In the slide-culture method, conidiospores were inoculated on the surface of agar blocks
(approximately 8 m2) placed between 2 sterile coverslips. Mycelia as well as the
conidiophores attached onto one surface of the coverslips as growth continued.
Acquiring nutrients from the agar block likely became less efficient as the colony
extended outwards, unlike regular fungal growth on an agar plate in which the nutrients
were obtained from the agar surface directly underneath the mycelia. Slide-culture may
induce stress responses and cause alteration of fungal development. Second, the fungal
cell wall represents a barrier that obstructs cell permeablization and antibody
accessibility; these conditions are extremely critical to achieve success in subsequent
irnmuno-labeing. Thus, lysing the cell wall is an important step to immuno-localize
proteins in the fungal cells. However, digestion of A. parasitius cell wall utilizing
Novozyme 234, a partially-purified cell wall lysing enzyme isolated from T richoderma
harzianum, did not give a constant degree of cell wall digestion, thus causing artifacts in

immuno-labeling. Preliminary indirect fluorescence microscopy detected Nor-l in the

65

cytoplasm in the conidiophores and the substrate level mycelia (Zhou 1998). Ver-l was
detected mainly in the conidiophores and the signals were confined to particle-like
structures (Liang, 1996). However, these results were not consistently reproducible, and
no reliable negative control was available.

The goal of my study was to develop a growth model that would closely mimic
regulation of toxin synthesis in soil and on the host plant. An improved colony
fractionation scheme as well as cell preparation methods were developed. A 72 h-old
colony was fractionated based on 24-h intervals to correlate the time of growth with AF
protein distribution. Colony fiactions were then fixed and embedded in paraffin to
generate sections (4-5 pm) for immuno-labeling without the need to lyse cell walls. In
addition, instead of using a conventional epi-fluorescence microscope, fluorescence
microscopy was performed using a Confocal Laser Scanning Microscope (CLSM) which
provides better resolution of confocal images (optical sections) and broader image
analysis capabilities (described in Chapter 1).

The results of my study demonstrated that Nor-1, Ver-l and OmtA were evenly
distributed among different cell types and were not concentrated in conidiophores,
contrary to the hypothesis. In a 72 h-old colony, these proteins were most abundant in
colony fraction 2 where the cells had grown for 24 to 48 h. Within a cell, Nor-l and Ver-
1 were primarily localized to the cytoplasm, suggesting that they are cytosolic enzymes.
OmtA was also detected in the cytoplasm and the fluorescent signal was confined to
discrete areas (patches). Yet due to limitations in resolution of CLSM, it was not clear if
those patches represent specific sub-cellular organelles. The pattern of labeling using
anti-OmtA was not consistent with localization of OmtA only to nuclei, peroxisomes, or
Woronin bodies. However, the data do not rule out the possibility that OmtA was
localized to other cell locations as well as these specific organelles. Although Nor-1,
Ver-l and OmtA appeared to localize in the cytoplasm, compartrnentalization of these

proteins needs to be further explored.

66

MATERIALS AND METHODS

Fungal strains. A. parasiticus SU-l (NRRL5862, ATCC 56775) is a wild-type,
aflatoxin-producing strain. A non-aflatoxin producing aflR knockout strain, AF S10
(kindly provided by Dr. Jeff Cary, USDA, New Orleans, LA.), was used as a negative
control. AF $10 was generated fiom A. parasiticus NR-l (niaD) which was derived from
parental strain SU-l (Cary et al., 2002). Two additional strains, A. parasiticus CSIO (ver-
1 wh-I pyrG) (Skory et al., 1993) derived from A. parasiticus ATCC36537 (ver-I wh-I),
and an omtA disrupted mutant strain LW1432 (ver-I wh-I omtA) generated by disrupting
the omtA gene in CS10 (Lee et al., 2002) were also used to study localization of OmtA.

Time-dependent fractionation of colonies grown on solid medium. To
determine the accumulation and distribution of Nor-1 , Ver-l, and OmtA in fungal
colonies grown on solid culture media, conidiospores (2 x 105) of A. parasiticus SU-l
and AF S10, as well as CSIO and LW1432 (to specifically localize OmtA), were
inoculated onto the center of YES (2% yeast extract, 6% sucrose, pH 5.8) agar overlaid
with sterile cellophane membranes and incubated at 29° C in the dark. Three-day-old
colonies of SU-l (S) and AF S10 (R) were fractionated into three concentric rings based
on area covered at 24, 48, or 72 h of growth (fiaction 1, 48-72 h old; fraction 2, 24-48 h,
and fraction 3, 0-24 h) to generate fractions 81, S2, S3 and R], R2, R3 (Lee et al., 2002).
A. parasiticus CSIO, and LW1432 colonies were fractionated following the same scheme
to generate analogous fractions C1, C2, C3, and L1 , L2, L3, respectively (Figure 4.1).

Preparation of paraffin-embedded fungal sections. Samples from fungal
colony fiactions were embedded in paraplast (Sigma, St. Louis, MO) using a published
procedure (Ausubel et al., 2003) with the following modifications. Fungal tissues were

fixed with Streck tissue fixative (Streck Laboratory Inc., Omaha, NE) at 4° C overnight,

67

SU1:wild type AFS1O (afIR)

CS10 (wh 1. var-1.10m?) LW1432 (wh1,ver-1, omtA)

0% ca L2 L3

81, R1, 01, L1: 48-72 h
82, R2, C2, L2 : 24-48 11
SS, R3, CB, L3 : 0-24 h

Figure 4.1. Time-dependent fractionation of firngal colonies. A 72 h culture of SU-l
(wild type AF producing) grown on YES medium (2% yeast extract, 6% sucrose) was
divided into 3 fractions based on 24 h grth intervals (S1: 48- 72 h growth; S2: 24- 72 h
growth; S3: 0- 24 h growth). The control strain AF 810, an qflR knockout mutant, was
prepared similarly to generate R1, R2 and R3. For detection of OmtA, additional strains,
LW1432(ver-1 wh-I omtA) and its parent strain CSIO(ver-1 wh-I pyrG), were used to
generate L1, L2, L3 and C1, C2, C3, respectively.

68

and then dehydrated in a graded series of ethanol: 30% (30 min, room temperature); 50%
(30 min, room temperature); 70% (overnight, 4° C); 85% (30 min, room temperature, 2
times); 95% (30 nrin, room temperature, 2 times); 100% (30 min, 4° C, 2 times), followed
by incubation in 100% xylene (10 min, room temperature, 3 times). Fungal tissues were
then incubated in a paraffin/xylene mixture (1:1; v/v) for 15 min, 60 rrrin and overnight at
60°C and finally for 8 h in 100% paraffin at 60° C (three changes of paraffin during
incubation). The paraffm embedded sample blocks were hardened in a plastic mold
(V WR Scientific, Detroit, M1) at RT. Sample blocks were cut into 4 pm thick sections
using a tissue section microtome (AO Spencer 820 microtome, Fisher Scientific). The
sections were attached to poly-L-Lysine (Sigma) coated coverslips (22 mm square).

Immuno-fluorescence labeling of fungal cells. Coverslips with paraffin-
embedded ftmgal sections were placed in a coverslip holder (EMS, Fort Washington, PA)
for de-paraffinization and antigen retrieval. Sections were de-paraffinized twice in 100%
xylene for 10 min, then rehydrated with a decreasing concentration of ethanol: twice in

100% for 10 min, once in 95%, 70%, 50% for 5 min each, and finally in deioned distilled
H20. Antigen retrieval was performed by heating the sections in 10 mM citrate buffer
(pH 6.0) at 95° C for 5 min followed by cooling at room temperature for 20 min.
Coverslips were rinsed with TBS (Tris buffer saline, pH 7.5) and incubated in blocking
solution (1% BSA with 0.1% saponin in TBS) at 4°C overnight. The samples were
probed with primary antibodies against Nor-1 (1 to 500 dilution)(Zhou, 1998; Lee et al.,
submitted), Ver-l (20 pg/ml) (Liang et al., 1997), OmtA (20 pg/ml) (Lee et al., 2002), or
anti-SKL (1:500, Zyrned, So. San Francisco, CA) as a positive control, followed by
secondary antibody conjugated to a fluorescent probe (goat anti-rabbit IgG- Alexa 488
conjugate [5 pg/ml]; Abs 495 nm/ Em 519 nm) at RT for l h. Coverslips were washed
after each antibody treatment with TBS containing 1% BSA and 0.1% saponin followed

by two washes with TBS for 10 min. Fungal nuclei were detected using SYTOX Green

69

fluorescence dye (Molecular Probes). The samples were mounted onto microscopic
slides with Prolong anti-fade mounting media (Molecular Probes, Eugene, OR).

Confocal Laser Scanning Microscopy (CLSM). Fluorescence image detection
was performed on a Zeiss 210 Laser scanning microscope or Zeiss LSM5 Pascal Laser
Scanning Microscope with a 488 nm laser line. The 40 X oil objective lens (Zeiss Plan-
NeoFlura, NA= 1.3) was used to obtain most images. The Alexa 488 fluorescence probe
(Abs 495 nnr/Em 519 nm) was detected using LP 520 or BP 520-560 barrier filters.
Fluorescence image analysis of SU-l, AF SlO, CS10 and LW1432 was conducted under
the same instrument parameter settings. High resolution images were generated using a
Zeiss Plan-APOCHROMAT (63 X/ 1.4 Oil Dic, N.A.=1.4) objective along with higher
zoom levels in the Zeiss LSM5 Pascal CLSM specifically to demonstrate the localization
of Nor-1, Ver-l , and OmtA within a SU-l cell. Single optical sections or extended focus
images from CLSM Z- stacks were obtained.

Quantitative fluorescence intensity analysis. To quantitate fluorescence
intensity, a Meridian IN SIGHT® plus laser-scanning system (Meridian Instruments, Inc.,
Okemos, MI) connected to a Zeiss Axioskop microscope equipped with IQ Master
Program (V2.31) image analysis software was used. The Alexa 488 green fluorescence
was detected using a BP 530/30 barrier filter under the 488 nm laser line. All images
were captured using a 40X Zeiss Plan-NEOFLUAR oil objective lens (N .A.=1 .3) (Carl
Zeiss Inc., Germany) with a 1X cooled charged-couple detector (CCD) allowing analysis
of a large number of cells in one 512 x 480 pixel resolution image. Fluorescence image
analysis of strains SU-l and AF 810, as well as CSIO and LW1432 was conducted using
the same instrument parameter settings. For each colony fiaction, twenty images were

acquired for intensity analysis and the average pixel number was reported.

70

RESULTS

Immuno-Iocalization of Nor-1 and Ver—l. CLSM detected Nor-1 and Ver-l in
all colony fiactions of SU-l (Fig. 4.2a for Nor-1 and Fig. 4.2d for Ver-l). Under the
same instrument settings, very little signal was detected in AF 810 (Fig. 4.2b for Nor-l
and Fig. 4.2e for Ver-l). Therefore, bright field images of the AF SlO colony fi‘actions are
shown to illustrate the typical size and numbers of cells analyzed (Fig. 4.2e and 4.21).
Highest fluorescence intensity was detected in fraction 82 for both Nor-1 and Ver-l.
Both proteins were equally distributed in substrate level mycelia and conidiophores (Fig
4.3 for Nor-l; Fig. 4.5, C for Ver-l).

Within a cell, high resolution optical sections taken using Zeiss LSM5 PASCAL
microscope with a Zeiss Plan-APOCHROMAT (63 X Oil Dic, NA=1.4) objective
showed that both Nor-l and Ver-l were mainly localized in the cytoplasm. Nor-1 did not
associate with specific organelles in either hyphae (Fig 4.3, A to D) or conidiophores
(Fig. 4.3, E). Ver-l was mainly present in the cytoplasm (Fig. 4.4, A, and B), frequently
associated with the cell wall as well as spore coat, and occasionally detected in particle-
like structures (Fig. 4.4, C, and E). However, the presence of Ver-l in locations other
than cytoplasm appeared to be non-specific since these labeling patterns were also
observed in the control strain AF 810 (Fig. 4.5, A). Thus, both Nor-1 and Ver-l are
concluded to be cytosolic proteins. Although the Ver-l antibodies bound to AF S10 non-
specifically, the overall fluorescence intensities detected in AF S10 were significantly
lower than in SU-l (Fig. 4.2, B and Fig. 4.5).

Immuno—localization of OmtA. To compare the fluorescence labeling intensity
between SU-l and CS10 and their non-aflatoxin producing counterparts, AF $10 and
LW1432, respectively, the samples were viewed under the lowest zoom level using the
Zeiss 210 CLSM (zoom = 20) in order to acquire maximum cell number within a field.

Under the same contrast and brightness settings, the samples prepared from the control

71

Figure 4.2. Immune-fluorescence confocal microscopy of Nor-1 and Ver—l in A.
parasiticus SU-l and AF SIO (aflR knockout mutant) grown on YES agar for 72 h.
Colonies of SU-l and AF 810 were divided into 3 fiactions (see Material and Methods).
The paraffm embedded fungal sections were immuno—labeled with anti-Nor-l antiserum
(panel A) (1:500) or anti-Ver-l IgG (panel B) (20 pg/ml) followed by Alexa 488
conjugated goat anti-rabbit IgG. Fluorescence intensities of SU-l sections irnmuno-
labeled with anti-Nor-l and anti-Ver-l are shown in Rows a and d, respectively; and of
AF 810 sections in Rows b and e. The related bright field images of the AF SlO colony
fractions are shown in Rows c and f (legends labeled in black). Bars, 100 pm.

72

A. Nor-l "

 

B. Ver—1 5'

(I

Figure 4.2

73

Figure 4.3. CLSM irnmuno-fluorescence detection of Nor-1 in A. parasiticus SU-l.
Fungal cells derived fi'om colony fraction 82 were irnmuno-labeled with the Nor-1
antibody followed by goat-anti-rabbit IgG conjugated with Alexa 488 probe.
Fluorescence images were acquired in a Zeiss LSM5 Pascal CLSM using a Plan-
APOCHROMAT (63 X Oil Dic, NA=1.4) objective and zoom level 3. Immuno-
fluorescence of Nor-1 was detected in the cytoplasm of SU-l hyphae shown in single
optical sections (panels A, B, C, and D). Immuno-fluorescence of Nor-1 was also
detected in conidiophores (panel E). The DIG image related to panel B is shown in panel

F. Bars=2pm.

74

 

Figure 4.3

75

Figure 4.4. CLSM immuno-fluorescence detection of Ver-l in A. parasiticus SU-l .
Fungal cells derived from colony fiaction 82 were immuno-labeled with the Ver-l
antibody followed by goat-anti-rabbit IgG conjugated with Alexa 488 probe.
Fluorescence images were acquired in a Zeiss LSM5 Pascal CLSM using a Plan-
APOCHROMAT (63 X Oil Dic, NA=1.4) objective and zoom level 3. Immuno-
fluorescence of Ver-l was detected in the cytoplasm of SU-l hyphae shown in single
optical sections (panels A and B). Immuno-fluorescence of Ver—l was also detected in
conidiophores (panels C and E). The DIG images related to C and E are shown in D and
F, receptively. Bars = 2 pm. The non-specific signals of Ver-l associated with cell wall,

spore coat, or particle structures are indicated with arrows.

76

 

‘W

1.33,.

‘3'
r
U.
A

 

Figure 4.4

77

 

Figure 4.5. Immuno-fluorescence labeling by Ver—l antibodies in the control snain A.
parasiticus AF S 10 and the wild-type strain SU-l. Fungal cells derived from colony
fraction R2 and S2 (see Material and Methods) were immuno-labeled with the Ver-l
antibody followed by goat-anti-rabbit IgG conjugated with an Alexa 488 probe.
Fluorescence images were acquired in a Zeiss 210 CLSM using a Plan-NeoFluar (40 X
Oil Dic, NA=1.3) objective. (A) Fluorescence signals detected in the control strain

AF 810. (C) Fluorescence signals detected in the wild-type strain SU-l. The bright-field
images related to A and C are shown in B and D, respectively. Representative non-

specific signals are indicated with arrows. Bar, 50 pm.

78

strain AFSIO (fractions R1, R2, and R3) and LW1432 (fractions L1, L2 and L3) did not
show significant fluorescent signals compared to samples prepared from the same colony
fractions from wild type SU-l (fractions S1, 82, and S3) or CSIO (fractions C1, C2 and
C3) (Fig. 4.6). These data are consistent with Western blot analysis of protein extracts
isolated fi'om the same colony fiactions (Lee et al., 2002). OmtA was detected in the
substrate level mycelium (Fig. 4.7, A; Fig 4.8, B) as well as conidiospore-bearing
structures (Fig. 4.7, B; Fig. 4.8, A) located in fiactions S1 and $2 at nearly equal intensity
as in substrate level mycelium. The fluorescent signal was confined to discrete areas
(patches) within cells (Fig. 4.7, A). However, due to limitations in resolution of CLSM, it
was not clear if these areas are associated with particular organelles. Similar samples
were also probed with anti-SKL antibodies that are expected to detect proteins targeted to
peroxisomes and Woronin bodies (Fig. 4.7, C) (Keller et al., 1991; Valenciano et al.,
1998). The observed fluorescent pattern was consistent with localization to these
organelles. In addition, we used SYTOX Green to stain nuclei in fungal fractions (Fig.
4.7, D); again the fluorescent pattern was consistent with the expected results. Under the
same magnification, immuno-fluorescence labeling pattern of OmtA (Fig. 4.7, A and B)
was different from SKL or SYTOX staining (Fig. 4.7, C and D). Neither anti-SKL not
SYTOX generated the same “patchy” pattern as anti-OmtA.

In summary, the data suggest that, like Nor-l and Ver-l, OmtA is evenly
distributed in all cell types in a fungal colony and does not accumulate to highest levels in
conidiophores. The fluorescent signal is not consistent with localization of OmtA to only
peroxisomes, Woronin bodies, or nuclei. However, the data do not rule out the
possibility that OmtA is localized to other cell locations as well as these specific
organelles.

Quantitative fluorescence intensity analysis. The CLSM fluorescence

intensities represent the abundance of Nor-l , Ver-l , and OmtA shown in three colony

79

Figure 4.6. OmtA protein localization in time-fiactionated colonies of A. parasiticus
SU-l, AF SlO (afR knockout), CSIO and LW1432 (omtA knockout) grown on YES agar
for 72 h. Paraffin-embedded fimgal sections were irnmuno-labeled with affinity purified
OmtA PAb (20 pg/ml) followed by Alexa 488 conjugated goat anti-rabbit IgG. (A)
Fluorescence images of SU-l (81, S2, and S3) and AF S10 (R1, R2, and R3). (B)
Fluorescence images of CSIO (C1, C2 and C3) and LW1432 (L1, L2 and L3). All colony

fiactions were analyzed under the same instrument settings. Bars, 50 pm.

80

 

Figure 4.6

81

Figure 4.7. Protein localization using OmtA PAb and anti-SKL, and detection of nuclei
using SYTOX Green. (A) Z-series overlay image of SU-l (fiaction $2) immuno-labeled
with OmtA PAb. The image consists of an overlay of ten consecutive optical sections
taken at Z—interval of 800 nm (step size). (B) Fluorescence images of vesicles of
conidiophores of SU-l (fraction S1) immuno-labeled with OmtA PAb. (C) Irnmuno-
labeling of SU-l (fiaction S2) with anti-SKL. The magnification for the left panel is
400X and for right pane] is 2,000 X. (D) Fluorescence image of nuclei in SU-l (fraction
82) stained with SYTOX. The magnification for the left panel is 400X and for the right
panel is 2,000 X. The bar in panels A, B, C, and D represents 10 pm.

82

 

Figure 4.7

 

Figure 4.8. CLSM immuno-fluorescence detection of OmtA in A. parasiticus SU-l.
Fungal cells derived from colony fraction S2 were immuno-labeled with the OmtA
antibody followed by goat-anti-rabbit IgG conjugated with an Alexa 488 probe. Single
optical sections were acquired using a Zeiss LSM5 Pascal CLSM with a Plan-
APOCHROMAT objective (63 X Oil Dic, NA=1.4). (A) Immuno-fluorescence of OmtA
was detected in conidiophores. (B) Immuno-fluorescence of OmtA was detected in

hyphae. Bars = 2 pm.

84

fiactions of SU-l. Fluorescence intensities were quantified by measuring the pixel
numbers on CLSM micrographs using the IQ Master Program (V2.31) image analysis
software. The quantitative pixel analysis to analyze distribution of AF enzymes in colony
fractions was consistent with qualitative detection by Western blot analysis and
fluorescence microscopy (Lee et al., 2002; Lee et al. submitted). The control strain

AF S10 did not produce significant levels of Nor-l, Ver-l and, OmtA compared to SU-l.
These data confirm that the highest quantity of these three aflatoxin enzymes occurs in
fiaction 2 (Table 4.1).

DISCUSSION

Because a close regulatory association between aflatoxin synthesis and
conidiation has been demonstrated in several studies (Guzman-de-Pena and Ruiz-Herrera,
1997; Hicks et al., 1997, Adams et al., 1998), a close temporal and spatial association
between AF protein expression and conidiospore development was hypothesized. To test
this hypothesis, antibodies against several AF proteins were generated to immuno-
localize these proteins. Previous studies (Liang, 1996; Zhou, 1998) utilized a fungal cell
preparation technique for immuno-labeling that required digestion of the cell wall using
Novozyme (Harris et al., 1994). However, in these early localization studies, variation in
cell wall digestion resulted in significant artifacts. The paraffm-embedment and
sectioning procedures developed and described in this study successfully preserved fungal
mycelium, developmental structures, organelle structure, and protein antigenicity; they
also eliminated the need for cell wall digestion, in turn generating consistent and
reproducible immuno-labeling results. This technique provides a useful tool to localize
proteins and possibly other compounds in fungal cells and colonies.

Despite the close regulatory relationship between sporulation and aflatoxin
production, my data suggest that Nor-1, Ver-l, and OmtA are not produced exclusively in

conidiophores as hypothesized. On the contrary, CLSM micrographs showed that these

85

Table 4.1. Quantative fluorescence intensity analysis of A. parasiticus SU-l and AF S 1 0

immuno-fluorescence labeled with Nor-l , Ver-l and OmtA antibodies.

 

 

 

 

 

(Fungal colony) (SU-l) (AF S10)

Protein / Colony inaction 81 S2 S3 R1 R2 R3
Nor-l 229.5 858.4 260.9 15.2 18.7 62.3
Ver-l 144.4 297.9 166.5 68.7 89.2 42.3
OmtA 120.0 940.1 146.7 4.3 3.9 21.6

 

* The data represent the average pixel number in 20 images from each colony fraction.

86

proteins are distributed throughout the colony and in both conidiophores and vegetative
hyphae. In fraction S2, the highest intensity of AF proteins was detected by CLSM and
the signal was observed at similar levels in vegetative hyphae and conidiophores.

CLSM detected Nor-1 and Ver-l mainly in the cytoplasm. These data are
consistent with data from previous subcellular fiactionation studies along with Western
blot analysis. Nor-1 was mainly detected in the cytosolic fiaction (Zhou, 1998), while
Ver-l was present in the cytosol (80%) and the rrricrobody fraction (14%) (Liang, 1996).
Based on CLSM data generated in the current study, detection of Ver-l in particle-like
structures by Liang ( 1996) (presumably microbodies), was an artifact. Cytosolic
immuno-fluorescence labeling of Ver—l was specific in SU-l and not observed in the
negative control strain AF S10. However, immuno-fluorescence of Ver—l on the cell wall,
spore coat, as well as particle-like structures was observed in both SU-l and AF S10.
Ver-l displays 56% protein sequence similarity to scytalone reductase (V idal-cros et al.,
1994), and 66% identity and 82% similarity to polyhydroxynaphthalene reductase; both
enzymes are involved in melanin biosynthesis. Melanins are dark pigments that
accumulate in distinct layers of cell walls and spores of many fungi (Wheeler and Bell.,
1988). Thus, the PAb against Ver-l used in this study possibly exilribited cross-reactivity
to the cell wall and the spores. In addition, immuno-gold labeling of Ver-l observed by
transmission electron microscopy (TEM) revealed that the labeling pattern of Ver-l in
particle-like structures in SU-l and AF S10 was not specific to Ver-l. The organelle
irnmuno-labeled by Ver-l antibody in both SU-l and AF S10 was identified to be the
Woronin bodies. The appearance of this organelle was recently demonstrated in A.
parasiticus using PAb against a C-terminal tri-peptide Peroxisomal Targeting Sequence 1,
SKL (Ser-Lys-Leu) (Lee, 2003). The fourteen percent of the Ver-l associated with the
nricrobody fraction detected by Liang (1996) was possibly due to this non-specific

interaction.

87

Cells immuno-labeled with OmtA PAb and analyzed by CLSM showed patches of
fluorescence within fungal cells suggesting that OmtA is confined to sub-cellular
compartments. An alternative explanation is that OmtA is present in the cytoplasm and is
thus excluded from cell organelles. To gain more information about sub-cellular
localization, we labeled paraffm embedded sections to identify specific organelles; for
example anti-SKL antibodies (Keller et al., 1991) were used to label peroxisomes and
SYTOX Green was used to detect nuclei. Both probes generated small, regularly shaped
signals consistent with the expected organelles, indicating that the double-membrane
bound nuclei as well as single-membrane bound microbodies were well-preserved. These
observations strongly suggest that we have minimized artifacts due to poor sample
preparation. Images using both probes were different than with OmtA PAb.

Quantitative fluorescence intensity analysis performed in this study supported
qualitative CLSM data for the distribution of AF enzymes in colony fractions. CLSM
also generated important imformation for subsequent determination of sub-cellular
localization of AF proteins using high resolution TEM. The specimen size required for
TEM cell preparation is extremely small (approximately 1 mm2 dimension to be
sectioned into 100 nm thickness). Therefore, the finding that the S2 fi'action exhibited
the highest density of AF proteins allowed sample acquisition efficiently (from fraction

S2) in order to detect these proteins at the sub-cellular level (Lee, 2003).

88

ACKNOWLEDGEMENTS

Special thanks to Dr. Shirley Owens (Center for Advanced Microscopy at
Michigan State University) for help in quantification of fluorescence intensity and Dr.
Jeff Cary (USDA, New Orleans, LA.) for providing the aflR knockout strain, AF S10.

Immuno-localization of OmtA is published in Applied and Environmental
Microbiology (Lee, L.W., C.H. Chiou, and J .E. Linz. 2002. Function of native OmtA in
vivo and expression and distribution of this protein in colonies ofAspergillus parasiticus.
Appl. Environ. Microbiol. 68:5718-5727).

Immuno-fluorescence analysis of Nor-1 and Ver-l is submitted for publication
(Lee, L. W., C. H. Chiou, K. L. Klomparens, and J. E. Linz. 2003. Sub-cellular
localization of aflatoxin biosynthetic enzymes in time-dependent fiactionated colonies of

Aspergillus parasiticus. Manuscript submitted to Arch. Microbiol.).

89

 

CHAPTER 5

Distribution and sub-cellular localization of the aflatoxin enzyme versicolorin B
synthase (VBS) in time-fractionated colonies of Aspergillus parasiticus

ABSTRACT

Aflatoxins (AF) are highly toxic and carcinogenic fungal secondary metabolites.
Aflatoxin biosynthesis in the filamentous fungi Aspergillus parasiticus and A. flavus
requires a complex pathway consisting of at least eighteen enzymatic activities; one of
these enzymes, versicolorin B synthase (V BS), catalyzes closure of the bisfuran ring in
the racemic versiconal hemiacetal substrate to form versicolorin B. Bisfuran ring
formation is important for AF toxicity as the structure is required for the subsequent
creation of AFB1-8,9 epoxide, a potent alkylating agent. I analyzed the localization of
VBS in the AF producing strain A. parasiticus SU-l grown on solid media using a colony
fiactionation technique developed previously. A highly specific polyclonal antibody,
raised against a Maltose Binding Protein-VBS firsion protein synthesized in Escherichia
coli, was utilized to detect VBS in SU-l grown on a rich solid medium via irnmuno-
fluorescence confocal laser scanning microscopy (CLSM) and irnmuno-gold transmission
electron microscopy (TEM). VBS was found not only in vegetative hyphae but also in
asexual developmental structures, called conidiophores. Western blot and CLSM analyses
demonstrated the highest abundance of VBS in colony fraction S2 where cells had grown
for 24 to 48h; this fraction also contained the highest levels of newly developed
conidiophores. The highest abundance of AF Bl also was detected in fiaction S2,
consistent with VBS abundance. At the sub-cellular level, CLSM and TEM detected
VBS at similar levels in the cytoplasm and in ring-like structures surrounding nuclei; it is

uncertain if enzymatically active VBS is present in either or both locations.

90

INTRODUCTION

Aflatoxins (AF), fungal secondary metabolites produced by filamentous fungi
including Aspergillus parasiticus, A. flavus, A. nomius, and A. tamarii, are highly toxic
and carcinogenic compounds (Cary et al., 2000; Eaton & Groopman, 1994; Cotty and
Cardwell,l999). AF contamination in commercial crops including com, peanuts, tree
nuts, and cottonseed has generated significant economic and human health concerns
throughout the world (CAST, 2003). Among the members of the AF family, AFB] is
considered the most dangerous as it is efficiently bioactivated by cytochrome P450
enzymes (particularly in the liver) to form the AF 31- 8,9 epoxide, a potent alkylating
agent. The epoxide interacts with the N—7 position of guanine residues in double stranded
DNA to generate AFB1-N7-Gua DNA adducts (Bailey, et al., 1996; Mace, et al., 1997;
Aguilar et al., 1993; Eaton and Gallagher,1994). The tumor suppressor gene p53 appears
to be a frequent target of AF B] since mutation of p5 3 is often seen in hepatic-cancer

patients with high levels of AF B] exposure (Yang, et al., 1997). Our goal is to eliminate

AF contamination in agricultural commodities and in turn to ensure product safety and
consumer health.

AF biosynthesis utilizes a complex pathway requiring at leastl 8 enzymatic
activities (Minto & Townsend, 1997; Payne, 1998; Bhatnagar et al., 2003). The structural
- genes that encode these aflatoxin enzymes, including nor-1 (Trail et al., 1994), ver-l
(Skory et al., 1992), omtA (Yu, et al., 1993), vbs (Silva et al., 1996), as well as one
regulatory gene aflR (Chang et al., 1993; Woloshuk et al., 1994), are clustered in a 75 kb
chromosomal region (Trail et al., 1995; Minto & Townsend, 1997; Payne and Brown,
1998; Cary et al., 2000; Bhatnagar et al., 2003). Although most AF biosynthetic genes
have been cloned and characterized, the mechanisms that regulate AF biosynthesis are not
completely understood. The positive-acting GAL4-type transcription factor AflR is
required for AF biosynthesis (Chang et al., 1993; Ehrlich et al., 1999a; Ehrlich, et al.,

2003). A recent report also described cluster-dependent positional effects on AF

91

biosynthesis. The nor-1 promoter is positively regulated within the AF gene cluster
whereas outside the cluster (niaD or pyrG), nor-1 promoter activity decreases
dramatically (Chiou et al., 2002).

To date, little is known about how or where aflatoxin enzymes interact at the
cellular level to synthesize AFB]. To address this issue, our laboratory developed a
colony fiactionation protocol and irnmuno-fluorescence and immuno-gold based
microscopic methods to localize proteins involved in AF biosynthesis. Previously, we
analyzed the distribution of OmtA, an enzyme involved in the later stages of AF
biosynthesis (Lee et a]. 2002). The highest abundance of OmtA was detected in firngal
cells grown for 24 to 48 h (fiaction S2) in A. parasiticus strain SU-l grown on a solid
medium (YES). Immuno-fluorescence confocal laser scanning microscopy (CLSM)
detected OmtA in the cytoplasm and the protein frequently was confined to discrete areas
in cells in this colony fiaction. Transmission electron microscopy (TEM) together with
immuno-gold labeling suggested that OmtA and the aflatoxin enzymes Nor-1 and Ver-l
are found in the cytoplasm; of particular interest, OmtA was also detected at high levels
in vacuole-like organelles in cells residing near the substrate surface of the colony (Lee et
al., manuscript submitted). These observations caused us to propose that early aflatoxin
enzymes function in the cytoplasm while later pathway enzymes are localized to
organelles either to protect cells from the toxic effects of late pathway intermediates or to
reduce the level of aflatoxin synthesis via protein turnover. In this report, we investigate
the localization of another key enzyme involved in AF biosynthesis, versicolorin B
synthase (V BS), in A. parasiticus to determine if the data support or refute the proposed
model.

VBS catalyzes the cyclization of the racemic versiconal hemiacetal substrate and
closure of the bisfuran ring to form versicolorin B (Lin and Anderson, 1992; Anderson

and Chung, 1994; Silva et al., 1996). The bisfuran ring structure is required for the

subsequent creation of the AF B1-8,9 epoxide and toxicity. vbs encodes a protein of 634

92

11111110 301
116. 33 1.
11111116]
and does
The pied
VBS 210]
by $08
atVI
mass '1‘.
VBS 1
lltltll
main'

trans

IECO

loca

in r
leni

811g

amino acids (V BS) that contains 3 putative N—glycosylation sites at amino acid residues
116, 221, and 507 (Silva et al., 1996). VBS displays sequence similarity to several
flavin—dependent oxidases and dehydrogenases but it does not catalyze a redox reaction
and does not bind FAD or F MN to its truncated FAD binding domain (Silva et al., 1996).
The predicted molecular mass of VBS based on the cDNA sequence is 70 kDa, yet native
VBS appears as a homo-dimer with a monomeric molecular mass of 78 kDa (estimated
by SDS-polyacrylamide gel electrophoresis). N-glycosidase F treatment of native VBS
and VBS heterologously expressed in yeast revealed that the discrepancy in molecular
mass is attributed to N-glycosylation (Silva et al., 1997). The N-tenninus of the native
VBS was also reported to be post-translationally modified (Silva et al., 1996). VBS
heterologously expressed in E. coli was not active, whereas VBS expressed in yeast
maintained its activity, suggesting that N—glycosylation and perhaps other post-
translational modifications are essential for VBS activity (Silva et al., 1997).

In the current study, a highly specific PAb was produced in rabbits using a
recombinant maltose binding protein (MBP)-VBS fusion expressed in E. coli. VBS
localization was conducted by fluorescence-based CLSM and irnmuno-gold TEM. VBS
was observed throughout a 72 h-old colony and was detected not only in vegetative
hyphae but also at similar levels in conidiophores. The highest levels of VBS, AF B], and
newly developed conidiophores were present in colony fiaction S2 where the cells had
grown for 24 to 48 h. These observations suggest a strong spatial association between
secondary metabolism and asexual sporulation; a regulatory and temporal association
between these processes was reported previously (Adams et al., 1998; Guzrnan-de Pann
and Ruiz-Herrera, 1997; and Calvo et al, 2002).

At the sub-cellular level, VBS was detected in the cytoplasm and at similar levels
in ring-like structures surrounding nuclei; based on morphology, these structures were
tentatively identified as endoplasmic reticulum (ER). Computer-based sequence analysis

suggested that VBS contains a putative signal peptide at the N-terminus. Because VBS is

93

 

 

N-link glycosylated, we propose that VBS is associated with the ER/golgi secretary
pathway for transient co- and post-translational modification and localizes to the
cytoplasm where it participates in aflatoxin synthesis. However, based on the data, it is
currently uncertain if the ER, the cytoplasm, or both locations represent the final

destination for VBS activity.

MATERIALS AND METHODS

Fungal strains and growth conditions. The wild type aflatoxin producing strain
A. parasiticus SU-l (NRRL5862, ATCC 56775) and a non-aflatoxin producing strain
AF S10 (aflR knockout mutant, kindly provided by Dr. Jeffrey Cary, USDA, New
Orleans, LA) (Cary et al., 2002) were used to study VBS and other AF enzymes. For
liquid culture, a conidiospore suspension (2 x 107 conidiospores) was inoculated into a
250 ml Erlenmeyer flask containing 100 ml YES media (2% yeast extract, 6% sucrose,
pH5.5) and incubated at 29°C with constant shaking (150 rpm) for 24 h to 72 h. For solid
culture, conidiospores (2 x 105/pl) were center inoculated onto YES agar (1.5%) overlaid
with sterile cellophane membranes and cultured in the dark at 29°C for 24 h, 48 h, or 72
h.

Production of antibodies against native VBS. Native VBS purified fi'om A.
parasiticus SU-l was kindly provided by Dr. Craig Townsend, Johns Hopkins University
(McGuire et al., 1996). Approximately 30 pg of protein was mixed in a l: 1 (V N ) ratio
with TiterMax adjuvant (Cthx, Norcross, GA) and used to immunize two New Zealand
White rabbits. After 30 d, a booster injection with the same dose of VBS/TiterMax
emulsion was utilized. Twenty-one (I later, the anti-serum was collected and the serum
IgG was precipitated in 33% ammonium sulfate according to standard protocols (Ausubel
et al., 2003).

94

 

Generation of MBP (Maltose Binding Protein)-VBS fusion protein. Three
possible reading frames of the vbs exon H fragment were amplified by PCR based on the
published sequence (Silva et al., 1996) using A. parasiticus SU-l genomic DNA as a
template. To generate VBS clones with each of the three possible reading fiames, PCR
amplification was conducted using three forward primers containing 23 nucleotides
including an EcoRI restriction endonuclease site (underlined) [(1) 5' agaGAATTC-
agAGGCTCGAAAGG-3', (2) 5' tagGAATTCcag AGGCTCGAAAG-3', and (3) 5'-
ctaGAATTC-tcagAGGCTCGAAA—3'] and one reverse primer (5'-ATCAAGCTT-
GGTCTACTGCCCAG-3') containing a HindIII restriction endonuclease site
(underlined). Each 1.4 kb vbs fiagment was individually cloned into the pMAL-c2
expression vector (New England Biolabs, Beverly, Mass.) to generate expression
plasmids, pMAL-c2-vbsl , pMAL-c2-vbs2, and pMAL-c2-vbs3. These plasmids were
transformed into E. coil to express the 94 kDa MBP-VBS fusion protein. The fusion
protein was induced in large scale culture by addition of 0.5 mM IPTG at optimum cell
density (O.D.600 approximately 0.6). Cells were disrupted by a brief freeze and thaw
cycle followed by sonication on ice (Sonifier cell disruptor W-350; Fisher Scientific,
Pittsburgh, PA). The induced soluble MBP-VBS was purified with amylose resin
according to the manufacturer's instructions (New England Biolabs, Beverly, Mass).

Production and purification of PAb against MBP-VBS. Purified recombinant
MBP-VBS was injected into two New Zealand White rabbits. Each rabbit was
immunized subcutaneously with approximate 50 pg purified MBP-VBS mixed at a l: 1
(V N ) ratio with TiterMax adjuvant (Cthx, Norcross, Ga). After 32 d, a booster
injection with the same amount of protein/TiterMax emulsion was utilized. Anti-sera
collection was performed after an additional 30-d and approximately 60 ml of anti-sera
were obtained from each rabbit. Serum IgG was precipitated in 33% ammonium sulfate
as described previously (Ausubel et al., 2003). Because immuno-gold TEM studies
require low background signals, anti-VBS IgG was further purified by affinity

95

 

chromatography. The anti-VBS IgG was passed through a column containing proteins
extracted from AF S10 (non-AF producing, does not synthesize VBS) grown in liquid
YES medium for 48 h as described by Lee et al., (2002); the extracted proteins were
conjugated to a resin rrrixture of Affi-Gel 10 and Affr-Gel 15 (BioRad, Hercules, Calif).
The flow-through fraction (containing the cleaned-up IgG) was dialyzed against
phosphate buffered saline (PBS)(Ausubel et al., 2003) and used for immunogold-labeling
on ultra-thin fungal sections (described below).

Time-dependent fractionation of fungal colonies. A 72 h-old colony grown on
YES agar was fiactionated according to Lee et al. (2002). Briefly, at 24 h intervals during
growth, the culture plate was marked on the bottom to indicate the location of the outer
margin of the colony. After 72 h of culture, the colony was dissected into 3 fractions
(concentric rings) using these marks. Fractionation of 72 h-old SU-l colonies (AF
producing strain) generated fractions S1 (48 to 72 h), S2 (24 to 48 h), and S3 (0 to 24 h).
Fractionation of AF S 1 0 (aflR knockout mutant) resulted in analogous fractions R1, R2,
and R3 (using the same scheme).

Fungal protein extraction. Mycelia from liquid culture were collected by
filtration through miracloth (Calbiochem, La Jolla, Calif.) by aspiration. Mycelia fi'om
solid culture fractions were generated as described above and peeled directly from the
cellophane membrane. The fungal mycelia were then pulverized under liquid nitrogen
with a mortar and pestle and fungal proteins were extracted using TSA buffer (2 mM
Tris-Cl, pH 8.0; 40 mM NaCl; and 0.025% sodium azide) containing complete protease
inhibitors (2 tablets/10 ml buffer, Roche Molecular Biochemicals, Indianapolis, Ind.).
The resulting extracts were centrifuged at 12,000 g for 20 min at 4°C to remove mycelia.
The supernatant (fungal protein extract) was transferred to microcentrifuge tubes in small
aliquots and stored at -80°C. Protein concentration was determined using Bio-Rad
protein assay reagent (Bio-Rad, Hercules, Calif.) and bovine serum albumin (BSA,

Sigma-Aldrich, St. Louis, Mo.) as a standard.

96

SDS—Polyacrylamide Gel Electrophoresis (SDS-PAGE) and Western blot
analysis. SDS-PAGE was conducted using a standard procedure (Ausubel et al., 2003) to
analyze expression of recombinant MBP-VBS and other fungal proteins. The molecular
mass was determined by comparison to Low-Range Prestained SDS-PAGE standards or
SDS-PAGE Molecular Mass Standards-low range (Bio-Rad, Hercules, Calif.). For
immunoblotting, 30 to 60 pg of fungal proteins were separated by SDS-PAGE
(electrophoresis on 10% or 12% polyacrylamide gels) and transferred onto PolyScreen
PVDF membrane (Dupont Biotechnology Systems, NEN Research Products, Boston,
Mass). The membranes were blocked with 1% casein in TBST (10 mM Tris-Cl, pH 8.0;
150 mM NaCl; and 0.05% Tween 20) at RT for 1 h. The blots were probed with anti-
VBS IgG (2 pg/ml) followed by goat anti-rabbit IgG alkaline phosphate conjugate
(Sigma-Aldrich, St. Louis, Mo.) as the secondary Ab probe (1: 5,000 dilution) at RT for
30 min. Reactive proteins were visualized using the BCIP/NBT colorimetric detection
system (Roche Molecular Biochemicals, Indianapolis, Ind.).

Detection of AFB] in colony fractions. (1) Thin Layer Chromatagraphy

(TLC). Fungal colony fiactions (from replica plates) were placed into glass scintillation
vials for measurement of dry weight, followed by extraction of AF B] using 5 ml
chloroform. The chloroform extract (5 pl) was dotted onto silica plates (LHPKD Silica
Ge] 60 A, 10 cm x 10 cm; Whatrnan International Ltd., Maidstone, England) for TLC

with a solvent system consisting of 5% acetone/ 95% chloroform. AF B] was detected

under long-wave UV light. (ii) Enzyme-Linked-Immunosorbent—Assay (ELISA).
AFB1 in each colony fraction was quantified by a competitive-sandwich ELISA method

described previously (Pestka et al., 1980). One ml of AF extract (in chloroform) was
evaporated under N2 and re-suspended in 0.5 ml methanol. The assay plates were coated
with anti-AFB] antibodies (Sigma-Aldrich), incubated at 42°C over night and blocked
with 1% BSA-PBS at 37°C for 30 to 60 min. Serially diluted samples as well as AF B]
standard (Sigma-Aldrich) were mixed with the AF B1-horse radish peroxidase (HRP)

97

conjugates (kindly provided by Dr. Pestka, Michigan State University) as a competitor,
followed by incubation with the substrate ABTS (2,2'-Azino-bis(3)-ethylbenzothiazoline-

6-sulfonic acid) to generate a detectable signal at an absorption wavelength at 405 nm.

The concentration of AFB] was determined according to the standard curve. The levels

of AFB1 measured in a colony fiaction were normalized to dry weight (AF B1 [pg]/

fraction weight [g]).

Preparation of paraffin-embedded fungal sections and immuno-fluorescence
labeling. Fungal samples were derived from a whole 48 h colony or colony fractions
generated from a 72 h-old colony. The detailed procedure to prepare the paraffin-
embedded fungal sections for immuno-flluorescence microscopy was described
previously (Lee, et al., 2002). Specimens (5 mm) were cut from each colony fraction,
fixed with Streck tissue fixative (Streck Laboratory Inc., Omaha, Nebr.) at 4° C overnight,
followed by dehydration in a graded series of ethanol, and finally embedded in Paraplast
(Sigma-Aldrich). Fungal sections (45 pm thickness) were generated using a tissue
section nricrotome (AO Spencer 820 nricrotome, Fisher Scientific) and attached to poly-
L-Lysine (Sigma-Aldrich) coated coverslips (22 mm square) (Corning Glass Works,
Corning, NY). Prior to immuno-labeling, sections required removal of paraffin by 100%
xylenes, and re-hydration with decreasing concentrations of ethanol, followed by an
antigen retrieval procedure (heating the sections in 10 mM citrate buffer, pH 6.0 at 95° C
for 5 min and cooling at room temperature for 20 min). After blocking the samples with
TBS containing 1% BSA and 0.1% saponin at 4°C overnight, the samples were immuno-
labeled with one of the following primary antibodies: 20 pg/ml of anti-VBS IgG
(generated in this paper); 20 pg/ml purified anti-OmtA IgG (Lee et al., 2002): 20 pg/ml
anti-Ver—l IgG (Liang et al., 1997); or a 1:500 dilution of Nor-l antiserum (Zhou, 1997;
Lee et al., manuscript submitted) at room temperature for 1.5 h, followed by incubation

with the secondary antibody conjugated with a fluorescent probe (Goat anti-rabbit IgG-

98

 

Alexa 488 conjugate [5 pg/ml]; Molecular Probes; Eugene, OR), at RT for l h. Selected
coverslips were subsequently stained with SYTOX orange dye (Molecular Probes;
Eugene, OR) to localize the nucleus.

Confocal Laser Scanning Microscopy (CLSM). To determine the distribution
of VBS in colony fiactions, fluorescence image acquisition and quantitative intensity
analysis of one colony of SU-l and AF S10 was conducted using a Zeiss Axioskop
microscope equipped with a Meridian Insight confocal laser scanning system (Meridian
Inc. Okemos, Mich.) as described by Lee, et al. (manuscript submitted). Due to damage
to the detector in this microscope, the remaining samples were viewed using a Zeiss
LSM5 Pascal Laser Scanning Microscope with the same 40X objective lens (Plan
NEOFLUAR 40 X/ 1.3 Oil Dic, NA=1.3). High resolution single confocal images,
extended focus image made from Z-stacks, or DIC (Differential Interference Contrast)
images were taken using a Zeiss Plan-APOCHROMAT (63 X/ 1.4 Oil Dic, N.A.=l .4)
objective. Detection of the Alexa 488 fluorescence probe (A bs/Em 495/519 nm) was
conducted using a BP 505-530 emission filter set under excitation with the 488 nm argon-
ion laser line and detection of SYTOX orange (Abs/Em 547/570 nm) was performed
using the BP 560-615 nm emission filter set under the 543 nm helium-neon laser.

Quantitative fluorescence intensity analysis. Samples derived fiom three SU-l
replicate colonies were immuno-labeled with anti-VBS or OmtA and subjected to
fluorescence intensity analysis by quantifying the pixel number in 20 randomly selected
images. Pixels were analyzed using IQ Master Program (V2.31) image analysis software
that accompanied the Meridian Insight laser-scanning microscope (Meridian Instrument,
Inc., Okemos, Mich.) or on a Macintosh using a NIH image program (developed at the
US. National Institutes of Health and available on the web: httpz/lrsb. info. nih.gov/nih-
irnagel). For the samples derived from the same immuno-labeling experiment (both SU-l

and the negative control AF S 1 0), the CLSM images were acquired under identical

99

 

 

instrument settings. For each colony fraction, average pixel number in the 20 images
was reported.

Statistical analysis. Fluorescence intensity data were statistically analyzed by a
complete random design with a split-plot model using SAS software (SAS Institute Inc.
Cary, NC) or two-way ANOVA using SigrnaStat (SPSS Inc. Chicago, Ill.). The amount
of AF B1 in three colony fractions was analyzed by one-way AN OVA using SigrnaStat.
Statistical significance among samples was defined by a P-value not greater than 0.05.

Preparation of ultra-thin fungal sections, immuno-gold labeling, and TEM
analysis. Ultra-thin sections were generated from colony fiactions 82 and R2. Sample
preparation and immuno-gold labeling followed the procedures described by Lee et al.
(manuscript submitted). Briefly, samples were prepared by a freeze-substitution method
(F legler et al., 1995) and embedded in LR White resin (Ted Pella Inc., Redding, CA).
Ultra-thin sections (90-100 nm) were generated using an MTX ultra-microtome (RMC,
Tucson, AZ) and mounted onto formvar-coated nickel grids. Sections were irnmuno-
labeled with column-purified anti-VBS IgG (50 pg/ml), followed by goat anti-rabbit IgG
conjugated with 10 nm gold (Ted Pella. Inc., Redding, CA); sections were subsequently
post-stained with 3% uranyl acetate for 20 nrin. Samples were observed using a JEOL

IOOCX II transmission electron microscope (Tokyo, Japan) at 100 kV acceleration

voltage .

RESULTS
Generation of PAb against native fungal VBS. An anti-VBS PAb preparation
was generated using native VBS purified from A. parasiticus SU-l as an imrnunogen.
Only 60 pg of VBS protein was available for immunization split between a primary and
booster injection into two rabbits. The resulting PAb recognized VBS heterologously
expressed in yeast but also appeared to cross-react with fungal proteins other than VBS.

To remove cross-reactive antibodies, we pre-absorbed the IgG fraction with proteins in a

100

 

fungal extract isolated from SU-l grown for 24 h in liquid YES culture (conditions which
generate little detectable VBS expression). The resulting IgG recognized a single sharp
band in heterologous VBS expressed in yeast (78 kDa) but several proteins in SU-l.
Among these SU-l proteins, two closely associated protein bands (one band equal to 78
kDa and a smaller band of approximately 72 kDa) appeared to be VBS based on pattern
of accumulation; they were only detected at 48 h and 72 h but not at 24 h in the SU-l
extracts (Fig. 5.1). Although this clean-up procedure reduced the cross-reactivity, the
remaining background signals could generate considerable artifacts during an immuno-
cytochenristry study. Thus, anti-VBS antibodies with higher specificity were produced
using an alternate procedure.

Expression of MBP-VBS fusion protein and production of PAb against VBS.
To assure production of full-length protein, we amplified (PCR) three reading frames of
VBS exon II based on the published sequence (Silva et al., 1996) and cloned these PCR
fiagrnents into the pMAL-c2 expression vector to generate the MBP-VBS fusion proteins.
E. coli harboring pMAL-c2-vbs1 expressed the expected 94 kDa MBP-VBS firsion
protein (Fig. 5.2) whereas E. coli harboring pMAL-c2-vbs2 and pMAL-c2-vbs3 generated
pre-maturely terminated protein products (molecular masses of 45 kDa and 55 kDa,
respectively). Control E. coli carrying the backbone vector, pMAL-c2 (malE::lacZ)
produced a 52 kDa MBP-fl-galactosidase fusion protein (Fig. 5.2).

The MBP-VBS fusion protein was purified and injected into two rabbits to
generate PAb. We initially intended to isolate VBS by removing MBP from the fusion
protein using factor Xa (IEER, Ile-Glu-Glu-Arg recognition). However, treatment
fiagrnented VBS as well (Fig. 5.3) even though there is no factor Xa cleavage site found
in the VBS exon 11 sequence. Thus cleavage within VBS appeared to be non-specific.
We consequently used the entire MBP-VBS fusion protein to generate PAb. ’

Based on Western blot analysis, PAb raised against MBP-VBS displayed a higher
specificity (Fig. 5.4) than PAb raised against the native VBS (Fig. 5.1). The serum

101

 

SU1

 

l 1 Yeast
24h 48h 72h vas

 

m

4.4.

75 kDa—p «—--- 2:1,;

 

 

 

Figure 5.1. Western blot analysis of VBS detected by PAb raised against native fungal
VBS. Fungal proteins were extracted from A. parasiticus SU-l grown in liquid YES
medium for 24 h, 48 h, and 72 h. Proteins were separated by SDS-PAGE (12%
acrylamide gel), transferred onto PVDF membrane and probed with rabbit serum IgG
raised against native VBS (2 pg/ml) that had been preabsorbed with protein isolated fi'om
SU-l cultured in liquid YES medium for 24 h. Heterologous VBS expressed in yeast is

shown as a control. The apparent molecular mass for the target protein is 78 kDa.

102

 

pMAL-cz pMAL-c2 pMAL-c2 pMAL-c2
-vbs3

 

    
 
  
 
 

      

MW1 -vbs1 -vbs2 uwz
IPTG: '- +II. +11- +31- +1
“may; 135,1 , ~ Wm”; .. --.97
MBP-VBS ~ .

“108
~..—v I 31

21‘ .023

.414

Figure 5.2. SDS-PAGE of recombinant MBP-VBS fusion proteins. Three reading
flames of vbs exon 11 sequences were amplified from genomic DNA of SU-l and PCR
fiagments cloned into pMAL-cZ expression vector generating three flame-shift plasmids,
pMAL-c2-vbsl, pMAL-cZ-vsz, and pMAL-cZ-vbs3 (see Materials and Methods) that
were subsequently transformed into E. coli DH501. The in-fiame MBP-VBS had the
predicted molecular mass of approximately 94 kDa (a**). As a control, E. coli carrying
the backbone vector, pMAL-cZ containing malE::lacZ, expressed only the MBP-fl-
galactosidase fusion protein (52 kDa). The induced fusion proteins (IPTG+) are indicated

with symbols * or **.

103

Figure 5.3. MBP-VBS treated with Factor Xa to generate MBP (42 kDa) and VBS (52
kDa). The purified MBP-VBS (94 kDa) produced in E. coli was incubated with Factor
Xa at RT for 0 h, 2 h, 4 h, 8 h, and 24 h. As a control, an equal amount of MBP-VBS
without incubation with Factor Xa was set at RT for 8 h and 24 h. (A) SDS-PAGE
analysis. Samples were separated by electrophoresis on a 10% polyacrylamide gel and
stained with coomassie blue. Molecular mass markers are shown at both sides of the gel.
(B) Western blot analysis. Samples separated by SDS-PAGE were transferred onto
PVDF membrane. The blot was incubated with anti-VBS IgG (2 pg/ml, raised against
native VBS) and Goat anti- rabbit IgG conjugated with alkaline phosphatase, followed by
colorimetric detection by BCIP/NBT.

104

Xa factor treated no Xe

I l I I
MW 0h 2h 4h 8h 24h 24h 8h MW

 

Him—:25“ ~—-- --~-- --=-.1 :2: 4— MBP-VBS(94kDa)

1".E‘fi ' 1 -... .. r '1':- ""1

43...- - *r 7 “-7- 7-“- --~—- ,, e—*VBS(521toa)
"' "" ”*5 "— MBP(42 kDa)
34-1. .. .
25. ..
20-... s
on 211 an 511 24hcontrol
2:13-23— —-_-3 2'994—uep-vas
”‘— .'- _ u. 1.. . “#VBS
- 34
- 25
- 20
Figure 5.3

105

 

 

recognized VBS expressed in yeast and native VBS present in a 72 h culture of SU-l
grown in YES (Fig. 5.4, A). SU-l cultured in YEP medium (contains peptone instead of
sucrose and does not support AF biosynthesis) did not produce detectable VBS (Fig. 5.4,
B). VBS was not observed in a 24 h SU—l in YES liquid-shake culture (Fig. 5.4, A and
C) while VBS was abundant at 48 h (Fig. 5.4, C).

Expression of VBS in A. parasiticus grown on solid medium. In liquid-shake
culture, fungal cells are maintained as vegetative hyphae and also produce secondary

metabolites at highest levels during a transition between active growth and stationary

 

phase. However, normal developmental processes such as asexual sporulation are not
observed under such growth conditions. Thus, I cultured A. parasiticus on a solid
medium that supports native morphology and asexual development including the genesis
of conidiophores and asexual conidiospores.

Western blot analysis using the highly specific VBS PAb (MBP-VBS) was
conducted on fungal total proteins isolated from SU-l and AF SlO colonies grown on
solid YES medium for 24 h, 48 h, and 72 h (Fig. 5.5). No VBS was detected in the
control strain AF 810 (Fig. 5.5, A and B). A 24 h-old SU-l colony did not contain
detectable VBS whereas colonies cultured for 48 h and 72 h did contain detectable levels
of VBS (Fig. 5.5, A), a result similar to liquid-shake culture (Fig. 5.4, C). We then
analyzed time-dependent expression of VBS; a 72 h—old SU—l colony grown on solid
YES was fractionated into S] (48 to 72 h), S2 (24 to 48 h), and S3 (0 to 24 h); AF S10
was similarly fractionated to generate R1, R2, and R3. Western blot analysis detected
VBS in all three SU-l fractions but not in any AF S1 0 fraction (Fig. 5.5, B). Fraction 82
appeared to contain the highest abundance of VBS. Similar results for OmtA (Lee et al.,
2002), Ver-l , and Nor-1 were reported previously (Lee et al., manuscript submitted). The
intact form of VBS (78 kDa) appeared in all SU-l samples and little degradation was

apparent.

106

 

 

 

 

A B C
Yeast SU1 Yeast SU1 SU1
VBS YES VBS l—‘—'I YES
l—'l YEP YES r—-. _
I 24h 72h 72h 72h 24h 48h 72h 103
as . as
If 28
a1
22' d ”j

 

 

 

 

 

 

 

Figure 5.4. Western blot analysis of VBS produced in liquid shake culture using highly
specific PAb raised against MBP-VBS. (A) Analysis of protein isolated from A.
parasiticus SU-l grown in liquid-shake YES (yeast extract sucrose) culture for 24 h or 72
h. (B) Analysis of proteins isolated from SU-l grown in liquid-shake YES or YEP (yeast
extract peptone) culture for 72 h. In panels A and B, samples were electrophoresed on
10% polyacrylamide gels and subsequently probed with a 1:5000 dilution of anti-serum.
VBS expressed in yeast was used as a control. (C) Time-course expression of VBS in
SU-l grown in liquid-shake YES culture for 24 h, 48 h, and 72 h. Proteins were
separated by electrophoresis on a 12% polyacrylamide gel and subsequently probed with
2 ug/ml of serum IgG. The primary signal is shown with the molecular mass of 78 kDa.

107

 

A su1 AFS10 3 5m AFS10

 

 

 

 

24h 48h 72h 48h 72h MW ;1 S2 :3 IF" R2 R2 MW
73., '-*:2::7skoa WW7
.22--—.—.-m _66 —>-—-—-m “liaise
- 45 “3:13? 45

-30 £Q31

'20

 

 

 

 

 

Figure 5.5. Western blot analysis of VBS expressed in A. parasiticus SU-l colonies
(solid culture) and colony fiactions. (A) VBS expressed in A. parasiticus colonies of
different ages. Proteins were isolated from colonies of SU-l and AF SlO (aflR knockout
strain) grown on solid YES medium for 24 h, 48 h and 72 h. (B) Distribution of VBS in
time-dependent colony fractions. The 72 h-old colonies of SU1 and AFSIO grown on
YES solid media overlaid with a cellophane membrane were divided into 3 fractions
based on time of growth (81: 48-72 h; S2: 24-48 h; S3: 0-24 h) according to Materials
and Methods. Approximately 60 pg of fimgal proteins were loaded in each lane of a 12%
polyacrylamide gel. Both blots were probed with specific anti-VBS IgG (2 ug/ml). The
molecular mass markers (MW) are shown on the right of each blot. Native VBS has a

mass of 78 kDa.

108

 

Confocal Laser Scanning Microscopy (CLSM) of time-dependent expression
of VBS in a 72 h-old fungal colony. Fungal tissues from three colony fractions (72 h-old
SU-l or AF SlO colonies grown on YES solid medium) were fixed and embedded in
parafi'm to generate sections for immuno-labeling. CLSM detected significant
fluorescence in all SU-l fi'actions (Fig. 5.6, A), while little fluorescence was observed in
AF S10 (F ig. 5.6, B). Thus, the corresponding AFSIO bright field images are shown to
indicate that similar numbers of the AF S10 cells were analyzed (Fig. 5.6, B, insets).
Fluorescence intensity analysis via CLSM (Fig. 5.6, A) appeared to parallel Western blot
analysis (Fig. 5.5, B) in that highest intensities were observed in fraction S2; here, VBS
was present in both substrate level mycelia (Fig. 5.8, A, B, and C) and in conidiophores
(Fig. 5.8, D). To confirm this observation, we immuno-localized VBS and OmtA in 2
additional colonies and analyzed the fluorescence intensity quantitatively.

Fluorescence intensity quantification and statistical analysis. Pixel analysis
was performed on images generated from three independent colonies (designated A, B,
and C)(Table 1). Pairwise comparison confirmed that the levels of OmtA in fraction S2
(cultured for 24 to 48 h) were significantly higher than the other two fractions of SU-l
(S2 vs. S1, P= 0.0003; S2 vs. S3, P < 0.0001). The levels of VBS detected by fluorescent
antibody were consistently higher in fraction S2 than in fi'actions SI and S3 although the

difference was not statistically significant (82 vs. 81, P= 0.3529; 82 vs. SB, P= 0.1230).
Production of AFB] in colony fractions. TLC analysis did not detect AFB1 in

the control strain AF 810 while significant quantities of AF B1 were detected in all 3 SU-l
fiactions (Fig. 5.7). The quantity of AF B] extracted from fraction S2 was higher than

from fractions 81 and S3 based on TLC (Fig. 5.7) and ELISA (Table 5.2), yet the
difference was not statistically significant (P: 0.0589).

Sub-cellular localization of VBS. Using CLSM and anti-VBS probes,
fluorescent signals appeared in the cytoplasm of hyphae of SU-l in fraction S2 and these

109

 

A
SU-1

 

 

Figure 5.6. CLSM of VBS protein distribution in time-fractionated colonies of A.

parasiticus SU-l (SI, 82, S3) and AFSIO (R1, R2, and R3) grown on YES agar for 72 h.
Paraffin-embedded fimgal sections derived from various colony fractions (see Material
and Methods) were immuno-labeled with anti-VBS IgG (20 ug/ml) followed by Alexa
488 conjugated goat-anti rabbit IgG. The associated bright field images (insets in B)
demonstrate the numbers of AF 810 cells being analyzed. Bars, 10 pm.

110

 

AFS10 SU1
Std R1 R2 R3 Std S1 82 S3 Std

AFB1 —>
AFG1 —’

 

Figure 5.7. Thin Layer Chromatagraphy analysis of AFB1 accumulation in colony
fractions. Colony fractions of A. parasiticus SU-l (Sl, S2, and S3) and AF 810 (R1, R2,
and R3) were extracted in 5 ml chloroform and 5 ul were resolved on silica plates.
Aflatoxin standard (Std) is shown on both sides and in the middle lane. Fluorescence of

AF was detected under long-wave UV light (2F 360 mn).

111

 

 

Figure 5.8. CLSM immuno-fluorescence of VBS of A. parasiticus SU-l grown in solid
culture. Immune-fluorescence of VBS was detected in the hyphae of SU-l shown as

extended focus images of CLSM Z-stacks (A and B) and a single optical section (C). (D)

Presence of VBS in a conidiophore shown in an extended focus image. Bars, 5 um.

 

Table 5.1. Quantitative fluorescence intensity analysis“ of A. parasiticus SU-l and

AF SlO labeled with polyclonal antibodies to VBS and OmtA.

 

 

 

 

 

 

 

(Fungal colony) (SU-l) (AF S10)
Protein / Colony frafction 81 S2 S3 R1 R2 R3
VBS
Colony .49 18.46 42.37 15.02 9.48 9.77 8.06
ab 1 1.60 29.00 6.22 5.75 5.73 7.25
Cb 6.48 1 1.92 5.05 3.00 3.26 3.89
OmtA
Colony .19 7.53 58.79 9.20 0.29 0.28 1.38
ab 15.22 37.08 2.55 1.04 1.65 2.85
Cb 9.27 28.28 2.29 0.86 1.84 1.69

 

* The data represent the average pixel number of 20 images fiom each colony fraction.

a. Images of colony A were generated by the Meridian Insight confocal laser scanning

system and pixel analysis by NIH image program.

b- Images of colony B and C were generated using the Zeiss LSM5 Pascal Laser Scanning

Microscope and pixel analysis by NIH image program.

113

 

Table 5.2. Quantitative detection (ELISA) of AF B1 in colony fractions of SU-l cultured

 

on YES solid media for 72 h.
(Fungal strain) (SU-l)
Colonv fraction 81 S2 S3

 

 

 

ColonyA *AFBI(118/g) 295.59 490.41 245.83
ColonyB 204.82 451.24 252.82

 

* The levels of AF B1 measured in the colony fraction were normalized to AF B1 (ug)/

fraction weight (g).

114

 

 

 

Figure 5.9. CLSM dual detection for localization of VBS and nuclei in A. parasiticus

SU-l. Fungal sections generated from colony fraction 82 were immuno-labeled with
anti-VBS IgG (20 ugml) followed by Alexa 488 conjugated goat-anti rabbit IgG
(represented in green) and further stained with SYTOX orange dye to contrast the
localization of nuclei (represented in red). Irnmuno-fluorescence of VBS (single green
channel) is shown in panels A, C, E, and G. The ring-like fluorescence signals are
indicated with arrows. The merged dual channel images indicating localization of VBS
(represented in green) and nuclei (represented in red) are shown at the right of each of the
image pairs (panels B, D, F, and H). Immuno-fluorescence pattern of OmtA (panel I,
immuno-labeled with 20 ug/ml anti-OmtA IgG) and nuclear staining are shown in the

lower right corner. Bars, 2 pm. Images in this dissertation are presented in color.

115

were fiequently associated with ring-like structures (Fig. 5.9, A, C, E, and G). We
stained nuclei using SYTOX-orange fluorescent dye (Abs/Em 543/570) and found that the
fluorescent ring-like structures were associated with the outside surface of the nuclei (Fig.
5.9, B, D, F, and H). The cytoplasmic labeling pattern of VBS was clearly different from
OmtA (Fig. 5.9, 1, lower right panel). Ultra-thin sections were immuno-labeled with
column purified anti-VBS IgG followed by immuno-gold labeling; TEM analysis
revealed that VBS was distributed within the cytoplasm and also concentrated in ring-like
structures surrounding the nuclei in SU-l (Fig. 5.10, C, D, E, and F), while no labeling
was observed in those structures in AF S10 (Fig. 5.10, A and B). Within a single cell,
several ring-like structures immuno-labeled with VBS antibodies were observed in both
CLSM (Fig. 5.9, A, C, E, and G) and TEM (Fig. 5.10, panel B). It is not clear if the
signals were associated with a membrane structure. The data suggested that VBS is
localized in the cytoplasm and also likely in the ER (see below).

DISCUSSION

I initially used native VBS to generate PAb but encountered significant cross
reactivity. This could possibly be due to the limited amount of purified VBS available
(60 pg for two rabbits) or that purified VBS still contained contaminating fimgal proteins.
Similar cross-reactivity was encountered using the glycosylated form of A. niger glucose
oxidase to generate PAb to the native enzyme (Witteveen et al., 1992). Native VBS has
been confirmed to be N-link glycosylated (Silva et al., 1997). If the glycosylated form of
the antigen affects antibody specificity, the native VBS should have been treated with
deglycosylating enzyme to remove the N-glycan moiety prior to immunization.
Nevertheless, I chose an alternative way to generate highly specific antibodies using
methods reported previously, e.g., anti-Ver-l (Liang et al., 1997), anti-Nor—l (Zhou and
Linz, 1999), and anti-OmtA (Lee et al., 2002), and this approach proved fruitful.

116

 

Figure 5.10. Immuno-gold labeling of VBS in A. parasiticus AF S10 and SU-l analyzed
by TEM. Ultra-thin fungal sections derived from colony fractions R2 (AF S10 shown in
Panels A and B) and S2 (SU-l shown in Panels C, D, E and F) were immuno-labeled
with column-purified anti-VBS IgG (50 ug/ml) followed by goat-anti rabbit IgG
conjugated with gold particles (10 nm diameter). The large organelles shown in panels B,
D, and F were generated at higher magnifications from the cognate hyphae shown in
panels A, C, and B, respectively. Abbreviations: CW, cell wall; M, mitochondria; N,
nuclei; V, vacuole. Bars represent 1 pm in panels A and E, 0.5 um in panels B and D, and

250 nm in panels D and F. (This figure was generated by Lee, 2003).

117

 

 

 

   

Figure 5.10

118

The E. coli- derived recombinant MBP-VBS fusion protein was generated and used as an
irnmunogen. Compared to other MBP fusion proteins previously generated in our
laboratory (Liang et al., 1996) (Zhou, 1997) (Lee, et al., 2001), the yield of MBP-VBS
fusion was relatively low. Silva et al. (1997) reported that E. coli expresses VBS in
inclusion bodies with low solubility. Although part of my recombinant MBP-VBS was
associated with cell particulates, part of the fusion protein was present in the soluble
fraction. This may be due to the nature (solubility) of VBS or just simply the large size of
the fusion protein (94 kDa).

A temporal and regulatory relationship between secondary metabolism and
microbial development has been postulated in various organisms (Bennett, 1983;
Horinouchi et al., 1994, Guzman-De-Pena and Ruiz-Herrera, 1997; Adams et al., 1998;
Calvo et al., 2002). In A. nidulans for example (contains the AF gene cluster but lacks
the final enzyme, OrdA), a regulatory association between asexual sporulation and
sterigmatocystin (ST) biosynthesis was demonstrated (Adams, et al., 1998). The balance
between vegetative growth, sporulation, and ST production is regulated by two
antagonistic factors, F le (Fluffy, I_.ow erA; brlA is a conidial vesicle specific gene) and
F adA (Fluffy Autolytic Dominant) involved in a G-protein signaling pathway (Yu et al.,
1996; Hicks et al, 1997). A dominant activation mutation in fadA completely blocked ST
production and sporulation (Hicks et al., 1997).

Based on these data, I hypothesized that AF biosynthesis is spatially associated
with fimgal development at the cellular level and that AF enzymes are localized
specifically in conidiophores (Lee et al., 2002?). However, recent data now clearly
demonstrate that this hypothesis is not fully correct (Chapter 4); similar levels of VBS
(this study), Nor-1, Ver-l , and OmtA (Chapter 4; Lee et al., submitted) were detected in
vegetative hyphae and conidiophores. However, immuno-localization of VBS (this

study) and OmtA, Nor-1, and Ver-l (Chapter 4) detected the highest levels of these
proteins and AFB1 in fraction 82; interestingly, fraction 82 also contained the highest

119

 

 

numbers of newly developed conidiophores. According to Adams et al. (1998),
conidiospores are generated approximately 24 h after germination of spores in the original
inoculum. Consistent with these findings, conidiophores and conidiospores were
observed in fractions S1 (48 to 72 h) and S2 (24 to 48 h) but not in fi'action S3 (0 to 24 h).
In addition, most conidiophores in fraction SZ were immuno-labeled with antibodies to
VBS and OmtA but significantly fewer were labeled in fraction 81 suggesting that these
proteins were present in newly developed conidiophores. Visual microscopic inspection

of the conidiophores in fi'action 2 confirms that many of these are immature and exist in

 

various stages of development. In conclusion, our data support a spatial and temporal
association between AF protein expression/AF production and sporulation at the colony
level. I cannot rule out the possibility that protein stability or proteolytic degradation may
contribute to our observations. The factors that directly participate in mediating this close
association in A. parasiticus require further investigation.

In terms of sub-cellular localization of VBS, both CLSM and TEM indicated the
presence of VBS in the cytoplasm as well as in ring-like structures adjacent to the
nucleus. I attempted to identify a subcellular targeting signal in VBS using protein
localization prediction programs found on the ExPASy Molecular Biology Server
(http://us.expasy.org). Since no database is available for filamentous fungi, in most cases
we used the sequence analysis database for yeast; occasionally data from animal or plant
cells were used. These analyses predicted that VBS has a signal peptide (analyzed by
iPOSRT, PSORT version 6.4, Signal IP V2.0, and Target P V1.0), suggesting VBS may
be associated with the ER/Golgi secretary pathway. Using PSORTH, the possible
subcellular localization for VBS was predicted as follows: Mitochondria: 43.5%;
Cytoplasm: 21.7%; Golgi: 17.4%; Endoplastic reticulum: 13.0%; and Vacuole: 4.3%.

The observed perinuclear localization of VBS is likely due to its association with
the ER and possibly the Golgi. First, the sequence prediction analysis suggests that the

close association of VBS with ring like structures is consistent with passage through an

120

ER/Golgi secretory pathway. VBS is a N-link glycosylated protein and the ER is the site
for core-glycan (Glc3-Man9-GlcNAcz) addition to the Asn residue.

Second, the perinuclear immuno-fluorescence labeling pattern of VBS is
reminiscent of several proteins known to target the ER including calreticulin (Roderick et
al., 1997; Dyer and Mullen, 2001; Wedlich-Soldner et al., 2002), BR mannosidase (Zuber
et al., 2000), nephrin (Yan et al., 2002), and the vacuolar system-associated protein
VASAP-60 (Brule et al., 2003).

Third, N-glycosylation of VBS is consistent with passage through an ER/Golgi

 

pathway. Native VBS appeared as a doublet (78 kDa and 72 kDa) upon SDS-PAGE.
Similar doublet bands have been detected by immuno-blotting of nephrin (Yan et al.,
2002) and VASAP-60 (Brule et al., 2003). These proteins localize to ER and are thought
to be post-translationally modified. In addition, it is well known that several co-
translational and post-translational modifications take place specifically in the ER but not
in the cytosol, including cleavage of the signal-peptide, N-link glycosylation, formation of
disulfide bonds, and glyco-phosphatidylinositol (GPI)-anchoring (Ellgaard and Helenius,
2003 review). These observations allow us to predict that VBS is present in the ER for
N-glycosylation and signal peptide cleavage; other VBS modifications might occur at that
location as well.

It is not clear if VBS is associated with the Golgi for complex glycan

 

modification. In filamentous fungi, the typical appearance of the Golgi apparatus with the
classic dictyosome organization of stacked disc shape cistemae was not observed
previously (Markham, 1995); however, a recent study detected the morphology of Golgi
bodies in A. niger (Khalaj et al. 2001). Our CLSM and TEM micrographs did not show
clear evidence that the Golgi bodies were being immuno-labeled. Also, the enzyme
PNGase F, which was used to confirm that VBS is N-Glycosylated (Silva et al., 1997),
cleaves intact oligosaccharide chains from the protein; therefore this method cannot

determine if VBS is modified with complex glycans (Ausubel et al., 2003). In order to

121

determine the degree of complexity of the glycan attached onto VBS, other types of
endoglycosidase and glycoamidase enzymes should be used (Ausubel et al., 2003).

N-glycosylation of VBS appeared to be significant for VBS activity and may play
a role in stability. In general, protein N—glycosylation along the secretary pathway plays
an important role in control of proper folding to the native tertiary and quaternary
structure, and for subsequent translocation to the final cellular destination (Ellgaard and
Helenius, 2003; Roth, 2002; Helenius and Aebi, 2001; Parodi, 2000). The N-
glycosylation and ER protein control system provide proof reading mechanisms to ensure
that the native conformation of newly synthesized proteins is maintained; in this regard,
they are important for the fidelity of cellular fimctions (Ellgaard and Helenius, 2003
review). My data support the idea that N-glycosylation contributes to protein stability;
VBS was significantly more stable than OmtA analyzed using similar methods (colony
fiactionation/Westem blot data). A high proportion of intact VBS (78 kDa) remained in
colony fraction S1 (48 to 72 h) compared to OmtA in this same fraction (Fig. 5.5 in this
study compared to Fig. 5, Lee et al., 2002).

It is still uncertain if enzymatically active VBS is present in the ER, cytoplasm, or
at both locations. VBS does not contain a typical C-terminal KDEL ER-retention signal
(Teasdale and Jackson, 1996; Harter and Wieland, 1996), so presumably it is not an ER
resident protein but is only transiently present for modification. However, in most cells
studied to date, glycoproteins are transported along the secretary pathway and localized in
the extracellular space, the plasma membrane, or in compartments that are involved in
secretion and endocytosis (Ellgaard and Helenius, 2003). Therefore, an important issue
to resolve is why VBS appears in the cytosol.

One possible explanation is that incorrectly folded, immature proteins can be
retro-translocated from the ER into the cytosol for degradation through an ER-associated
degradation (BRAD) process (Kostova and Wolf, 2003). However, the relative
proportion of VBS found in the cytosol and the ER (approximately 50% in each location)

122

 

 

strongly suggests that the cytosolic localization of VBS is not likely due to this type of
mechanism.

Another, more likely explanation is that, since VBS is involved in biosysthesis of
a secondary metabolite (i.e. AF ), targeting of this protein to its final cellular destination
may be different from typical glycoprotein targeting in primary metabolism in higher
eukaryotes. Perhaps the regular glycoproein transport machinery is altered during
secondary metabolism. In addition, we cannot rule out the possibility that although VBS
appeared to be a cytosolic potein, it may be associated with a particular sub-cellular
compartment yet the membrane structure was not preserved during cell preparation.
Alternatively, VBS may be anchored in membrane structures while the bulk of the protein
remains in the cytoplasm. In support of this, VBS is predicted to contain a putitive
transmembrane domain (PSORT version 6.4). Furthermore, mature VBS is a good
candidate for a cytosolic protein because addition of a polar glycan generally increases
solubility (Helenius and Aebi, 2001).

If VBS is functional in the cytosol, it will be interesting to determine if this
enzyme interacts with other AF synthesizing enzymes that appeared in the cytoplasm (e. g.
Nor-l , Ver-l, and OmtA; Lee et al., manuscript submitted). Based on our preliminary co-
immuno-gold labeling of VBS and OmtA using TEM, these two proteins are in close
proximity in the cytosol. Future experiments involved in co-localization of various AF
synthesizing proteins either by GFP-linked or co-immuno-labeling techniques, subcellular
fractionation to identify the organelles associated with AF synthesizing enzymes, and
immuno-precipitation studies to determine interactions between AF proteins may provide

insights about the coordination of these protein involved in AF biosynthesis.

123

ACKNOWLEDGEMENTS
Special thanks to Dr. Xudong Fan (Center for Advanced Microscopy at Michigan
State University) for help in Operating the jet-freezer.
Data in Chapter 5 were submitted for publication (Chiou, C. H., L. W. Lee, S. A.
Owens, J. Whallon, K.L. Klomparens, C. A. Townsend, and J. E. Linz. Distribution and
sub-cellular localization of the aflatoxin enzyme versicolorin B synthase (V BS) in time

fiactionated colonies of Aspergillus parasiticus.Fungal Genet. Biol.).

124

 

 

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