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This is to certify that the
dissertation entitled

Functional Analysis of OmtA and Subcellular Localization of
Aflatoxin Enzymes in Aspergillus parasiticus

presented by

Li-Wei Lee

has been accepted towards fulfillment
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Environmental Toxicology

 

 

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6/01 cJCIRC/DmeOmpes-sz

 

 

FUNCTIONAL ANALYSIS OF GMT A AND SUBCELLULAR LOCALIZATION OF
AF LATOXIN ENZYMES IN ASPERGILL US PARASIT IC US

By

Li-Wei Lee

A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
DOCTOR OF PIHLOSOPHY
Department of Food Science and Human Nutrition

2003

ABSTRACT

FUNCTIONAL ANALYSIS OF 0M T A AND SUBCELLULAR LOCALIZATION OF
AF LATOXIN ENZYMES IN ASPERGILL US PARASI TIC US

By

Li-Wei Lee

Biosynthesis of aflatoxin is a complex process that involves the activities of at
least 17 pathway enzymes. The distribution of these enzymes within fungal colonies and
location within fungal cells are not known. The objective of this study was to investigate
the distribution and sub-cellular location of Nor-1, Ver—l and OmtA that represent early,
middle and late enzymatic activities, respectively, in the aflatoxin biosynthetic pathway.
My initial goal was to determine if OmtA is necessary and sufficient to catalyze the
conversion of the aflatoxin pathway intermediate sterigmatocystin to O-
methylsterigmatocystin in vivo and if this reaction is necessary for aflatoxin synthesis. I
generated A. parasiticus omtA null mutant LW1432 and a maltose binding protein-OmtA
fusion protein expressed in Escherichia coli. Enzyme activity analysis of OmtA fusion
protein in vitro confirmed the reported catalytic function of OmtA. Feeding studies
conducted on LW1432 demonstrated a critical role for OmtA and the reaction catalyzed
by this enzyme in aflatoxin synthesis in viva. Because of a close regulatory link between
aflatoxin synthesis and asexual sporulation (conidiation), I hypothesized a spatial and
temporal association between OmtA expression and conidiospore development. I
developed a novel time-dependent colony fractionation protocol to analyze the
accumulation and distribution of OmtA in fungal colonies grown on a solid medium that

supports both toxin synthesis and conidiation. Using Western blot analysis, OmtA was

detected in all fractions of a 72h old colony including cells (0 to 24h) in which little
conidiophore development was observed. OmtA in older fractions of the colony (24 to
72h) was partly degraded. F luorescence-based immunohistochemical analysis
demonstrated that OmtA is evenly distributed among different cell types and is not
concentrated in conidiophores. These data suggest that OmtA is present in newly formed
fungal tissue and then is proteolytically cleaved as cells in that section of the colony age.
Western blot analysis and immuno-fluorescence microscopy demonstrated the
highest abundance of Nor-1, Ver-l and OmtA in colony fraction 2 (24 to 48h) of SUl.
Fungal tissues in this fraction were subjected to freeze-substitution, immuno-gold
labeling with highly specific polyclonal antibodies, and analysis by transmission electron
microscopy. Nor-1 and Ver-l were primarily localized to the cytoplasm suggesting that
they are cytosolic enzymes. OmtA was also detected in the cytoplasm. However, in cells
located near the basal (substrate) surface of colony fraction 2 of SUI, large quantities of
OmtA were detected in organelles identified as vacuoles. The role of this organelle in
toxin biosynthesis is unclear. Based on data from this and previous studies, it is
reasonable to assign two potential roles for the vacuole in aflatoxin synthesis: 1) as a
means to reduce or limit aflatoxin synthesis, OmtA is transported to and inactivated in
vacuoles via proteolytic cleavage; 2) as a means to shield the cell from the potential toxic
effects of aflatoxin in the mycelium, OmtA is transported to the vacuole together with the
late aflatoxin pathway intermediate ST. In the vacuole, OmtA is activated and ST is
converted to OMST and further converted to AF B] by OrdA localized in or on the same

vacuole. These hypotheses form the basis for my future studies on aflatoxin synthesis.

Copyright by
Li-Wei Lee
2003

ACKNOWLEDGMENTS

I would like to thank my major professor, Dr. John Linz. Without him, this
dissertation would not have been possible.

I also would like to thank my committee members, Dr. James Pestka, Dr. Karen
Klomparans, Dr. Gale Strasburg and Dr. Jay Schroeder.

Several people in the Linz laboratory provided a great deal of help including
Ching-Hsun Chiou, Dr. Michael Miller, Matt Rarick, Dr. Shun-Hsin Liang and Dr. Ren-

Qing Zhou. This help was greatly appreciated.

TABLE OF CONTENTS

LIST OF TABLES ................................................................................. ix
LIST OF FIGURES ................................................................................. x
CHAPTER 1
Literature Review ........................................................................... 1
Impact of aflatoxin contamination on health and the economy ....................... 1
Environmental factors affecting toxin production on crops - Temperature and
Humidity ............................................................................. 3
Aflatoxin biosynthesis and asexual sporulation ......................................... 4
Aflatoxin biosynthesis ..................................................................... 7
Aflatoxin gene cluster ............................................................. 7
Aflatoxin genes ..................................................................... 9
AflR ......................................................................... 9
A flJ .......................................................................... 9
fasl,fas2 andpks ........................................................ 10
nor-l ....................................................................... 10
avnA ....................................................................... 10
adhA and norA .......................................................... 11
ava ........................................................................ 11
estA ........................................................................ 1 1
vbs and verB .............................................................. 12
ver-l and moxY .......................................................... 12
omtB ....................................................................... 13
omtA ....................................................................... 13
ordA ....................................................................... 14
Organelles involved in secondary metabolism in filamentus fungi- the microbody
and vacuole ................................................................................. 15
Microbody .......................................................................... 16
Vacuole ............................................................................ 19
Compartmentalization of penicillin ..................................................... 25
REFERENCES ............................................................................ 28
CHAPTER 2
Genetic Disruption of omtA in Aspergillus parasiticus .............................. 36
INTRODUCTION ......................................................................... 36
MATERIALS AND METHODS ....................................................... 37
Fungal strains ..................................................................... 37
Construction of omtA disruption vector pLW14 ............................. 37
Transformation and screening for omtA gene-disruption strains ........... 38
Southern hybridization analysis ................................................ 40
Northern hybridization analysis ................................................. 40

vi

Feeding studies of omtA gene disruption strains .............................. 41

RESULTS .................................................................................. 41
Generation of omtA disruption mutants ....................................... 41
In vivo feeding experiments ..................................................... 46
DISCUSSION .............................................................................. 46
REFERENCES ............................................................................ 50
CHAPTER 3
Expression/Distribution of the OmtA Protein in Colonies of Aspergillus
parasiticus ................................................................................. 52
INTRODUCTION ......................................................................... 52
MATERIALS AND METHODS ........................................................ 53
Fungal strains ..................................................................... 53
Construction of OmtA expression vector pLW12 ........................... 53
Conversion of sterigmatocystin to O-methylsterigmatocystin by maltose
binding protein—OmtA .................................................. 54
OmtA antibody production and purification .................................. 55
Western blot analysis ............................................................ 56
Time-dependent fractionation of colonies grown on solid medium ....... 57
Immunolocalization of OmtA protein .......................................... 58
Preparation of paraffin-embedded fungal sections .................. 58
Immunolabeling ......................................................... 59
Confocal Laser Scanning Microscopy (CLSM) ..................... 59
RESULTS .................................................................................. 60
Western bolt analysis and protein localization using Pab against native
OmtA ...................................................................... 60
Purification of Maltose Binding Protein-OmtA .............................. 6O
Enzymatic conversion of sterigmatocystin to O-methylsterigmatocystin
by Maltose Binding Protein-OmtA .................................... 61
Production of PAb against OmtA fusion protein ............................. 61
Western blot analysis of OmtA in colonies grown on solid medium ......67
Confocal Laser Scanning Microscopy ......................................... 67
DISCUSSION .............................................................................. 74
ACKNOWLEDGEMENTS .............................................................. 80
REFERENCES ............................................................................ 81
CHAPTER 4
Sub-cellular Localization of Aflatoxin Biosynthetic Enzymes in Time-Dependent
Fractionated Colonies of Aspergillus parasiticus ..................................... 84
INTRODUCTION ......................................................................... 84
MATERIALS AND METHODS ........................................................ 86
Fungal strains and time-dependent fractionation of firngal colonies ...... 86
Western blot analysis of proteins isolated from colony fractions .......... 86
Immuno-fluorescence labeling and confocal laser scanning microscopy
(CLSM) ............................................................................ 87
Quantitative fluorescence intensity analysis .................................. 87

vii

Sample preparation for immuno-electron microscopy ...................... 88

CHAPTER 5

CHAPTER 6

Immuno-gold labeling and electron microscopy ............................. 88
RESULTS .................................................................................. 89
DISCUSSION ............................................................................ 105
ACKNOWLEDGEMENTS ............................................................. 109
REFERENCES ........................................................................... 110
Detection of Aflatoxin in Fungal Spores and Colonies ............................. 113
INTRODUCTION ....................................................................... 1 13
MATERIALS AND METHODS ...................................................... 116

Fungal strains .................................................................... 116

Aflatoxin distribution in time-dependent fractionated colonies of SUl

grown on solid medium ............................................... 116

Spores harvested by water (wet conditions) ................................. 116

Binding of AF B1, AF G1, OMST, and VA in water by spores ............. 120

Spores harvested by cotton-tipped applicators .............................. 120
RESULTS ................................................................................. 121

Aflatoxin distribution in time-dependent fi'actionated colonies of SUI

grown on solid medium ............................................... 121

Binding of AF BI and pathway intermediates in water by spores 124

AFB] concentrations in spores harvested under dry condition ............ 130
DISCUSSION ............................................................................ 130
REFERENCES ........................................................................... 134
Future Studies ............................................................................ 135

Model 1 ........................................................................... 135

Model 2 ........................................................................... 135

Model 3 ........................................................................... 136

viii

LIST OF TABLES

Table 4.1 . Quantative fluorescence intensity analysis of A. parasiticus SUI and AF SlO
irnmuno-fluorescence labeled with Nor-1 , Ver-l and OmtA antibodies ..................... 94

Table 5.1. Fractionation of fungal colonies at different ages ............................... 118

Table 5.2. Total aflatoxin detected in the colony, agar and membrane from different aged
colonies ............................................................................................. 122

Table 5.3. Dry weight (g) of fractionated mycelium from different aged colonies ...... 122
Table 5.4. Total aflatoxin (pg) in fractionated mycelium fiom different aged colonies.123

Table 5.5. Normalized aflatoxin levels (ug of aflatoxin/g dry weight mycelium) in
fractionated mycelium from different aged colonies ......................................... 123

Table 5.6. Aflatoxin detected in conidiospores isolated under wet and dry conditions from
A. parasiticus grown on YES and PDA agar ................................................... 125

ix

LIST OF FIGURES

Figure 1.1. A model for genetic control of Aspergillus growth, asexual sporulation and
sterigmatocystin biosynthesis ...................................................................... 6

Figure 1.2. Schematic presentation of the aflatoxin gene cluster ............................. 8

Figure 1.3. A schematic presentation of the perosisomal-targeting sequence-dependent
protein-import pathway for proteins destined for the peroxisomal matrix .................. 20

Figure 1.4. PSORT (Prediction of Protein Localization Sites version 6.4) prediction for
the protein targeting sequences (PTSl and PTSZ) in (A) Aflatoxin enzymes, AflJ, Nor-l ,
AvnA, VBS, Ver-l, OmtA and OrdA from A. parasiticus, and (B) 6-aminopenicillanic

acid acyltransferase from Penicillium chrysogenum and A. nidulans ....................... 21

Figure 2.1. Restriction endonuclease map of plasmid pLW14 .............................. 39
Figure 2.2. Southern hybridization analysis of omtA gene disruption strains .............. 42
Figure 2.3. Northern blot analysis of omtA gene disruption strains ......................... 44

Figure 2.4. Thin layer chromatography of extracts of CSIO and omtA-disruption strains
LW1418 and LW1432 ............................................................................. 47

Figure 3.1. Western blot analysis and fluorescence microscopy using polyclonal
antibodies raised against partially purified fungal OmtA proteins ............................ 62

Figure 3.2. Western blot analysis of fungal protein extracts using subtraction-purified
PAb raised against native OmtA. ................................................................. 64

Figure 3.3. Conversion of ST to OMST by affinity purified MBP-OmtA fusion protein
.......................................................................................................... 65

Figure 3.4. Western blot analysis of fungal protein extracts using affinity purified OmtA
PAb ................................................................................................... 68

Figure 3.5. OmtA protein localization in time-fractionated colonies of A. parasiticus
SUI, AF SlO (AfR knockout), C810 and LW1432 (omtA knockout) grown on YES agar
for 72 h. ............................................................................................... 71

Figure 3.6. Protein localization using OmtA PAb and anti-SKL, and detection of nuclei
using SYTOX Green. ............................................................................... 73

Figure 4.1. Western blot analysis of fungal protein extracts using Ver-I and Nor-1
polyclonal antibodies. .............................................................................. 90

Figure 4.2. Immuno-fluorescence confocal microscopy of Nor-1 and Ver-l in A.
parasiticus SUI and AF S10 (aflR knockout mutant) grown on YES agar for 72 h ....... 92

Figure 4.3. Immuno-gold labeling of A. parasiticus SUI and AF S10 ...................... 95
Figure 4.4. Immuno-gold labeling of OmtA in Aspergillus parasiticus AF S10 and SUI 98
Figure 4.5. Immuno-gold labeling of OmtA in Aspergillus parasiticus SUI ............ 100
Figure 4.6. Immuno-gold labeling using anti-SKL and anti-Ver-I in A. parasiticus
AFSIO and SUI. .................................................................................. 103
Figure 5.1. Mutants in the aflatoxin biosynthetic pathway ................................. 115
Figure 5.2. Diagram of colony fractionation .................................................. 117
Figure 5.3. Thin layer chromatography of aflatoxin and OMST in water used for spore
harvest ............................................................................................... 126
Figure 5.4. Thin layer chromatography of RHNl spore extracts ........................... 127

Figure 5.5. Thin layer chromatography of SUI and ATCC36537 spore extracts ....... 128

Figure 5.6. Thin layer chromatography of SUI and RI-INI spore extracts after binding
with water used to harvest ATCC36537 spores ............................................... 129

Figure 6.1. Proposed models for synthesis and export of aflatoxin from fungal cells 1 38

Figure 6.2. Western blot analysis of fungal protein extracts using affinity-purified OrdA
PAb ................................................................................................. 139

Figure 6.3. Production of MBP-AflJ fusion protein and AflJ antibody ................... 140

xi

CHAPTER 1

Literature Review

1.1. Impact of aflatoxin contamination on health and economy.

Ingestion of aflatoxins by humans and animals can cause acute and chronic
aflatoxicosis. The effects vary depending on the dose of aflatoxin ingested, time of
exposure and nutritional status of patients (CAST, 2003). Aflatoxins were identified as
the cause of thousands of turkey deaths in 1960 (known as Turkey X disease) (Asao et
al., 1963). Chronic toxicity can cause lowered levels of reproduction, feed efficiency,
weight gain, meat and egg production, and enhance liver cancer in animals.
Immunosupression resulting from chronic toxicity can lower animal resistance to
bacterial, fungal and parasitic infection. Aflatoxins can cause acute hepatitis in humans.
AFBl was detected in liver tissues of deceased patients (CAST, 2003) and patients
suffering two childhood diseases, Kwashiorkor and Reye's syndrome (Becrofi and
Webster, 1972).

Chronic aflatoxicosis in humans is usually associated with liver cancer.
Epidemiological studies showed a positive correlation between dietary aflatoxins and
liver cell cancer (LCC) in Asia and Africa (Eaton and Gallagher, 1994). Humans exposed
to both hepatitis B virus (HBV) and aflatoxin increased the possibility of developing liver
cancer. The synergism between the two factors was also observed in experimental studies
in HBV transgenic mice and woodchucks (Sell et al., 1991; Bannasch et al., 1995). The
mechanism of interaction between aflatoxin and HBV is still not clear. Two hypotheses

were suggested (Wild and Turner, 2002). One possibility is that HBV infection alters the

expression of aflatoxin metabolizing enzymes and consequently increases the binding of
aflatoxin to DNA. The second hypothesis is that aflatoxin exposure enhances viral
infection and replication. HBV may also inhibit DNA excision repair enzyme activity and
increase AF Bl adduct persistence.

The role of aflatoxin in liver cancer may be associated with mutations in the
tumor suppressor gene p53. p53 is known to regulate several cell activities such as cell
cycle, apoptosis, and DNA repair. Mutations in p53, therefore, may render a selective
growth advantage to the cell (Smela et al., 2001). In liver cells, aflatoxin B1 can be
activated to the 8,9-epoxy AF BI by Cytochrome P450 mono-oxygenase enzymes. This
epoxy AF B1 is particularly reactive with the guanine in the third position of codon 249
(AGQ) of p53 and forms a DNA adduct. It was proposed that the DNA composition
surrounding codon 249 of p53 favors this reaction. It has been shown that aflatoxins
prefer to bind a guanine base with a guanine or cytosine as an adjacent 5’ base (Smela et
al., 2001; Wild and Turner, 2002). After DNA repair, mutation with a G to T transversion
occurrs and results in an Arg (AGQ) to Ser (AG1) alteration in the p53 protein. This
mutation may disable the normal function of p53 protein (Smela et al., 2001). The 249s"r
mutant is insufficient to immortalize human liver cells but confers a grth advantage to
previously immortalized cells (Wild and Turner, 2002).

The economic impact of aflatoxin contamination is huge. The potential economic
cost resulting from the loss of crops and livestock contaminated with aflatoxins was
estimated and reported by the Council for Agricultural Science and Technology (CAST,
2003). The mean simulated potential value of crops lost because of aflatoxin
contamination was 5 47 million per year in food crops (corn and peanuts) and $ 225

million per year in feed corn. The potential value of livestock lost was $ 4 million per

year (CAST, 2003). As a result of human and economic issues, a standard to restrict
aflatoxin levels in food and feed for human and animal safety was set by the US. Food
and Drug Administration. The action levels of aflatoxin are 20 p.p.b. (0.5 p.p.b. for milk)

for food for human consumption and 20-300 p.p.b. for animal feeds (CAST, 2003).

1.2. Environmental factors affecting toxin production on crops- Temperature and
Humidity.

The colonization of crops and production of aflatoxins by A. flavus are known to
be affected by environmental factors (CAST, 2003). Among them, temperature and
humidity are two of the most important factors affecting aflatoxin contamination in the
field and during storage of crops. It has been observed that abnormally high temperatures
and below average moisture (drought) are highly associated with the incidence of
aflatoxin contamination in corn and peanuts (Payne, 1998). This can be explained by the
fact that, under high temperature and drought conditions, A. flavus can grow better than
other fungi on crops with low water activity. Conditions for aflatoxin production are
more restrictive than those for fungal growth. A. flavus can grow in the temperature range
from 12 to 48°C with the optimal temperature at 37°C. Asperigillus species grow at water
activity above 0.85 (Wilson and Payne, 1994). The optimum conditions for colonization
are a temperature between 25 to 35°C with a substrate water activity at 0.90. However,
the optimal conditions for the fungus to produce aflatoxin are 30°C with a water activity
of 0.996 (anleni et al., 1997). At temperatures of 20 and 37°C, A. parasiticus did not
produce aflatoxin (F eng and Leonard, 1998).

Another important factor that has a positive effect on aflatoxin production is

insect infestation. Insect damage in crops provides the fungus entry through the hard hull

of pistachios, cotton bolls and corn, and allows to successful infection of crops (CAST,
2003).

The interaction between environmental factors (temperature and humidity) and
crop species (which provides nutrient and water activity) are the most critical
determinants for successful colonization and aflatoxin production by A. flavus. In
addition, the nutrient composition of crops and their effects on aflatoxin production have
been reported (Luchese and Harrigan, 1993 and Calvo et al., 2002). Simple sugars such
as sucrose, glucose, fructose and sorbitol support aflatoxin production (Luchese and
harrigan, 1993). The seed oil, linoleic acid, also supports aflatoxin production (Calvo et

aL,2002)

1.3. Aflatoxin biosynthesis and asexual sporulation.

It has been shown that fungal cells after a period of vegetative grth (at least 18
h) become competent to receive developmental signals (conidiation). The signals include
carbon and nitrogen starvation. In A. nidulus, asexual sporulation and sterigmatocystin
(ST) production are coordinately up-regulated by FluG and Fle, and down- regulated by
a G-protein-mediated signaling pathway (FadA-dependent proliferation signaling) (Hicks
et al., 1997; Adams and Yu, 1998). F luG is involved in the production of an extracellular
factor(s) required for initiation of the developmental pathway (D’Souza et al., 2001). An
extracellular factor(s) can bind to receptors on the cell surface and transmit a positive
signal for asexual sporulation by activating transcription factor brlA (an early regulator of
development), and by activating transcription factor F le (fluffy low brlA). Activated
Fle then represses the F adA-dependent signal for proliferation by inactivating the or-

subunit (FadA, fluffy autolytic dominant) of a herterotrimeric G-protein. Both inactive

FadA and active Fle result in induction of ST production possibly through activating
transcription factor AflR (Adams and Yu, 1998 and Figure 1.1).

The proposed relationship between fungal growth, aflatoxin production and
sporulation in A. parasiticus is similar to that in A. nidulus but the genetic evidence is not
yet clear. The following observations, however, suggest that they are closely associated.
l) The chemicals, fluoroacetic acid and diaminobutanone, which inhibit sporulation also
reduce aflatoxin accumulation (Reiss, 1982). Using a concentration that had little effect
on growth, fluoroacetic acid could greatly reduce the accumulation of aflatoxin. When the
concentration was adjusted to allow sporulation, toxin accumulation returned to normal
levels (Reiss, 1982). Added after spore germination, diaminobutanone could completely
suppress sporulation and aflatoxin production (Guzman-de-Pena et al., 1997). 2) In
submerged shake culture, no sporulation was observed, and a much lower concentration
of aflatoxin was produced than that in solid culture (Guzman-de-Pena et al., 1997). 3)
Two mycotoxins, Aurasperone C (in A. niger) and Fumigaclavine C (in A. fumigatus),
were found mainly in spores but not in mycelia (Palmgren et al., 1986). Aflatoxin
concentrations in spores isolated from A. parasiticus and A. flavus were reported
(Wicklow and Shotwell, 1983; Palmgren et al., 1986). Wicklow (1983) reported that the
aflatoxin concentration in spores (harvested in water) ranged from 1,106 to 54,300 ppb,
and in sclerotia from 109 to 73,200 ppb in Aspergillus parasiticus. Using a vacuum
method, Palmgren et al. reported that 161,000 ppb of aflatoxin was detected in the spores
and 245,000 ppb in the mycelia in Aspergillus parasiticus (Palmgren et al., 1986). In the
same report, norsolorinic acid in a nor-1 mutant (mutation in the nor-1 gene) was mainly

detected in mycelia (730,000 ppb) and only 280 ppb in the spores (compared with

/? —’ flbE —*flbD '—'> flbB\

FluG

4L Asexual

\7‘ flbC ‘3' sporulation

, —Flb ~' 3.

 

 

 

 

FadA-GT? + SfaD (Y) ——> Colony growth

, Sterigmatocystin
T T Biosynthesis

 

 

 

 

ND

 

 

Figure 1.1. A model for genetic control of Aspergillus growth, asexual sporulation and

sterigmatocystin biosynthesis.

161,000 ppb of aflatoxin in wild-type spores) (Palmgren et al., 1986). This result implied
that aflatoxin is synthesized mainly in mycelia. The question of whether aflatoxin is

synthesized in spores or requires transport to spores remains unanswered.

1.4. Aflatoxin biosynthesis.

Aflatoxin gene cluster. Aflatoxins are secondary metabolites produced by
filamentous fungi, including A. flavus, A. parasiticus, A. nomius and A. pseudotamarii
(Ito et al., 2001). More than 17 enzymes are involved in aflatoxin biosynthesis and the
genes encoding these proteins are found clustered in a 75 kb genomic DNA fragment
(Figure 1.2; and Bhatangar et al., 2003). Recently, it was demonstrated that location
within the aflatoxin cluster plays an essential role in the regulation of aflatoxin gene
expression. When nor-1 was relocated to other chromosomal positions outside the cluster
(pyrG and niaD), the expression of this gene was highly reduced (Chiou et al., 2002). An
enhancer element is proposed to be located inside or surrounding the cluster to positively
regulate aflatoxin gene expression. In addition, clustering of secondary metabolite genes
may provide an advantage for horizontal gene transfer and, therefore, favor cluster
dispersal and survival as a unit (Walton, 2000). Dispersed pathway genes can be
transmitted vertically but do not favor horizontal transfer because a single pathway gene
has no advantage to the recipient host. The synthesis of secondary metabolites may be
subject to the natural selection to benefit the producing fungi. A dothistromin gene
cluster, homologous to the aflatoxin and sterigmatocystin clusters, was isolated from the
pine needle pathogen Dathistrama pim' (Bradshaw et al., 2002). Dothistromin is a

difuranoanthraquinone toxin with structure similar to versicolorin B. However,

norB
cypA

One pathway-specific I
regulator aflR

aflJ
adhA
estA

norA
ver- 1 A

verA
avnA

verB

ava
omtB

omtA
ordA

vbs

fl— ¢— —O —O-D—O IOQ—Q-I-C-CIQ—O d—I-Dfi—I-DQ—Q—I-b

cpr

moxY
0rdB

‘-

Figure 1.2. Schematic presentation of the aflatoxin gene cluster. Arrows indicate the

direction of gene transcription.

dothistromin can not be converted to aflatoxin by A. parasiticus, suggesting that A.
parasiticus has no enzyme to convert this toxin to an aflatoxin intermediate. One DNA
fragment containing four biosynthetic genes, datA, B, C and D was cloned from D. pini.
The amino acid sequences of dotA and dotC have 80.2 and 31.2% identity to ver-l and
aftT (toxin transporter), respectively. Because dothistromin is a phytotoxin targeted to
mature pine embryos and leaf callus, DotC was proposed to transport dothistromin out to
the target in host plant. This proposed general secretion mechanism may protect fungal
cells from this toxin.

Aflatoxin genes.

aflR. aflR encodes a pathway-specific regulator for many genes in the aflatoxin
gene cluster. AflR can bind to a consensus DNA sequence, 5’-TCG N5 CGA-3’
(consensus AF LR binding site) in the promoters of regulated genes. Disruption of aflR
eliminated accumulation of aflatoxin and pathway enzymes (Cary et al., 2002) except for
the transcript from aflJ (Chang and Yu, 2002). The function of AflR may be necessary
but not sufficient to activate certain pathway genes whose promoters do not contain
perfect AflR binding site, for example, ava. Other transcription factors may be also
involved in the regulation of aflatoxin genes and responsible for basal level of aflatoxin
gene expression.

:11”. aflJ is located adjacent to aflR in the aflatoxin gene cluster. A gene
disruption experiment showed aflJ is involved in Aflatoxin biosynthesis. No aflatoxin
was detected in aflJ mutant while other aflatoxin gene transcripts including pksA, nor-1,
ver—l and omtA were detected (Meyers et al., 1998). The actual function of this gene is
not clear. No aflJ homolog can be found in DNA/protein sequence databases to date.

Based on the Peroxisomal Targeting Sequence 2 (PTS2) found in this gene, one proposed

function of Afll is the transport of Aflatoxin proteins into microbodies (Meyers et al.,
1998). Recently, several reports suggested that AflJ may function to regulate early—
pathway gene transcription (Du, ASM Meeting 2000) possibly through interaction with
AflR as demonstrated in a yeast two hybrid system (Bhatangar, 2000; Chang, 2003).

fasl, fas2 and pks. The structural genes involved in aflatoxin biosynthesis
include pksA, fas-IA, fas-2A, nor-1, avnA, ava, ver—I, omtB, omtA, ordA and vbs
(Figure 1.2; Bhatangar et al., 2003). The 6-carbon compound hexanoyl-CoA is the starter
unit in aflatoxin biosynthesis and is generated by fatty acid synthases Fasl and FasZ
using one acetyl-CoA and two malonyl-CoA molecules (Mahanti et al., 1996; Watanabe
et al., 1996). Malonyl-CoA and acetyl-CoA are first converted to hexanoyl-CoA by these
two enzymes. The hexanoyl-CoA is further extended to noranthrone (NA) by polyketide
synthase (PKS) using 7 molecules of malonyl-CoA (Minton and Townsend, 1997).

nor-1. Norsolorinic acid (NOR) is the first stable intermediate in aflatoxin
biosynthesis. NOR is convert to averantin (AVN) by the enzyme Nor-1 (Zhou and Linz,
1999). Mutation of the nor-1 gene in A. parasiticus greatly reduced aflatoxin production
and resulted in accumulation of the red pigment, NOR (Trail et al., 1994). A low level of
aflatoxin production remained suggesting that an alternative pathway or enzyme(s) could
substitute Nor-I function but with lower enzymatic efficiency.

avnA. The reaction to convert AVN to 5’-hydroxyaverantin (HAVN) is catalyzed
by the enzyme AvnA. Disruption of avnA in A. parasiticus resulted in a non-aflatoxin
producing strain and accumulation of the aflatoxin precursor AVN (Yu et al., 1997).
AvnA is a cytochrome P-450 monoxygenase and contains 495 amino acids with a

calculated molecular mass of 56.3 kDa.

10

adhA and norA. HAVN can be converted to averufin (AVR) by AdhA and NorA
(Bhatangar et al., 2003). An adhA mutant was reported to produce trace amounts of
aflatoxins (Chang, ASM Meeting 2000). Based on the observation that adhA and nor-1
mutants still produced aflatoxins, it seems likely that enzymes involved in the early
stages of secondary metabolism share some activities with other primary metabolic
enzymes.

ava. AVR is converted to versiconal hemiacetal acetate (VHA) by Ava (Yu et
al., 2000) and may also involve Cpr (Bhatangar et al., 2003). ava is located
downstream of omtB and upstream of verB. Ava contains 285 amino acid with a
calculated molecular mass of 30.9 kDa (Yu et al., 2000). The function of Ava has not
been determined yet but it may function as a dehydrogenase based on its amino acid
sequence.

estA. The conversion of VHA to versiconal (VAL) is catalyzed by an esterase
(Anderson and Green, 1994). The gene thought to encode this esterase is estA, located
downstream of adhA and upstream of narA in the gene cluster (Yu, et al., 2002). Based
on the cDNA sequence, this enzyme has a putative molecular mass of 34 kDa (Yu, et al.,
2002). In the native form, however, this enzyme may form a dimer of 60 kDa as shown
by gel filtration (Kusumoto and Hsieh, 1996). The expression pattern of estA may differ
from other aflatoxin genes based on its ability to be expressed in non-aflatoxin inducing
medium (PMS) (Anderson and Green, 1994; Yu, et al., 2002). However, this conclusion
was based on RT-PCR; additional data generated from different methods such as Western
blot analysis or Northern blot analysis may be required to confirm this statement. The
second copy of this gene, estA2, located outside of this gene cluster, however, was not

expressed in either inducing or non-inducing media (Yu, et al., 2002). This provides

11

additional evidence for a positional effect on the aflatoxin gene cluster reported
previously (Chiou et al. 2002).

vbs and verB. The side-chain of Versiconal (VAL) is cyclized by Versicolorin B
synthase (VBS) to generate Versicolorin B (VERB) (Lin and Anderson, 1992; Silva,
1996 and 1997). The bisfuran ring structure of AFB, is formed at this step. VBS may
require glycosylation to obtain enzyme activity (Silva, 1996 and 1997). The native VBS
purified from A. parasiticus has a molecular mass of 78 kDa. Compared to the deduced
molecular mass of 70.3 kDa, a post—translational modification (glycosylation) of the
native protein was suggested. A double bond is introduced into the bifuran structure of
VERB by a desaturase called VerB (verB) to generate versicolorin A (VERA). The
bifuran ring is responsible for the mutagenic nature of aflatoxin AF 8,.

ver-l and moxY. The conversion of versicolorin A (VERA) to demethyl
sterigmatocystin (DMST) may require several steps (oxidation, keto-reduction, and
decarboxylation) and several enzymes (Keller et al., 1995). Ver-l, a keto-reductase, was
confirmed to be involved in this conversion (Liang, 1996 and 1997). Based on similarity
with a polyhydroxynaphalene reductase (66% identity and 82% similarity) which is
involved in melanin biosynthesis in Magnapartha grisea, the proposed function for Ver—l
is to catalyze a deoxygenation of VA to 6-deoxy VA. Disruption of the ver-I gene in
Aspergillus parasiticus NR-l (NiaD') resulted in a non-aflatoxin producing strain, VAD-
102 (Liang et al.,1996). The moxY gene, located in the aflatoxin gene cluster, also was
suggested to be involved in conversion of VERA to DMST (Bhatnagar et al., 2003).
However, no specific data were presented to support this idea. In A. nidulans, stcS, which
encodes a monooxygenase, was demonstrated to be involved in VERA to DMST

conversion (Keller et al., 1995).

12

omtB. The conversion of DMST to sterigmatocystin (ST) is carried out by OmtB
(Yaba et al., 1989 and Yu et al., 2000). The calculated molecular mass of OmtB is 43.1
kDa. OmtB as well as OmtA are the only two methyl-transferases utilized in the aflatoxin
biosynthetic pathway. omtB is located downstream of omtA and upstream of ava in the
aflatoxin gene cluster. The promoter of omtB contains an AflR binding site but the
downstream gene, ava, does not. The intergenic region between the omtB translation
stop codon and the start codon of ava is only 173 bp. Yu et al. (2000) suggested that
ava may be co-transcribed with omtB from the omtB promoter and subsequently the
transcript is processed into two mRNAs.

omtA. OmtA is a methyltransferase that catalyzes the conversion of ST to O-
methyl-sterigmatocystin (OMST) by transferring a methyl group from the cofactor S-
adenosyl-methionine (SAM) (Bhatnagar etal., 1987 and Cleveland et al., 1987).
Although the molecular structure of OMST is close to ST, several chemical
characteristics of OMST are more similar to AF 8,. Under UV illumination, the
fluorescence emitted from ST shows a weak brick-red color while emission is pale blue

from OMST and blue from AF B,. The migration rate (Rf) in TLC analysis showed that
the Rf of ST is 0.97, OMST is 0.44, and AFB, is 0.37 using ether-methanol-water

(96:3:1) as a development system. In a chloroform-acetone (95:5) development system,
the Rfof ST is 0.74, OMST is 0.24 and AFBI is 0.22 (Bhatnagar et al., 1987).

The first native OmtA purified from a cell-free extract of A. paraciticus showed
one band on a non-denaturing gel with a molecular mass of 168 kDa. This protein was
shown to contain two subunits, 110 and 58 kDa, by SDS-PAGE (Bhatnagar er al., 1988).
This large enzyme was extremely unstable and displayed a low reaction rate in vitro.

Therefore, Bhatnagar et al. suggested that a (hydrophobic) environment is required to

13

stabilize the hydrophobic ST (substrate), SAM (cofactor), OMST (product) and this
enzyme (Bhatnagar et al., 1988). However, using a similar method, a different protein
(OmtA) was purified with the same O-methyl-transferase activity. The second purified
protein had a smaller molecular mass of 40 kDa (Keller et al., 1993; Liu et al., 1993).
Antisera raised against the 40 kDa purified protein was used to screen a cDNA expression
library resulting in the cloning of omtA (Yu et al., 1993). The molecular mass derived
from the OmtA cDNA is 46 kDa. Comparing the N-terminal sequence of the purified
protein (40 kDa) with the deduced amino acid sequence (46 kDa), Yu et al. suggested
that OmtA might contain a leader sequence of 41 amino acids. This leader sequence may
function to translocate OmtA across a membrane structure (Yu et al., 1993). To explain
the relationship between the two purified enzymes, 40 kDa and 168— kDa, Yu et al.
suggested that OmtA subunits were posttranslationally modified and polymerized to
become the large protein (Yu et al., 1993). However, the antibody generated from the
large protein (168- kDa) has very little reactivity with the 40 kDa OmtA protein,
suggesting that these two enzymes are different proteins (Keller, 1993). No further study
was carried out on this large purified protein. In the current study, we conducted a
knockout of the omtA gene in A. paraciticus C810 and conducted an in viva feeding
experiment. Our results suggested that OmtA (46 kDa) plays a major role in conversion
of ST to OMST in viva (Lee et al., 2002).

ordA. OrdA is the final enzyme involved in the aflatoxin biosynthetic pathway.
The proposed reactions for the conversion of OMST to AFB, include parahydroxylation,
demethylation, reduction, ring cleavage, and conversion of a six-membered ring to a five-
membered ring (Bhatnagar et al., 1991). Therefore, more than one enzyme was proposed

to be involved in this conversion. In one early study, the partially purified oxidoreductase

14

showed four major bands on a non-denaturing gel with a molecular mass about 180 - 200
kDa and seven major proteins (165, 105, 88, 64, 43, 38, and 26 kDa) on SDS-PAGE
(Bhatnagar et al., 1991). However, none of the bands excised from the native gel showed
enzymatic activity. Recently, ordA encoding an enzyme activity able to convert OMST to
AF B1 was cloned from A. flavus (Prieto and Woloshuk, I997). The putative molecular
mass of OrdA is 60.2 kDa. Based on the amino acid sequence, OrdA is predicted to
function as a monooxygenase. Yeast transformed with a single gal l-ordA construct
(ordA fused with inducible gall promoter) were able to convert exogenous OMST to
AFB, (Yu et al., 1998). Because no other intermediates between OMST and AFB, were
reported, it is possible that this final reaction step requires only OrdA. However, the
possibility can not be ruled out that other enzymes may help OrdA in more general
reactions rather than the specific step in both filamentous fungi and yeast. The co-purified
proteins with oxidoreductase activity observed in the early study could represent these
helper proteins (Bhatnagar et al., 1991).

Two additional proteins were identified that are required for production of the G
group aflatoxins and these are detected in microsomal and cytosolic fractions (Yabe et

al., 1999).

1.5. Organelles involved in secondary metabolism in filamentous fungi- the
microbody and vacuole.

For most secondary metabolites including aflatoxin, the biological functions are
not clear. Specific secondary metabolites (including penicillin and preheliminthosporol)
are believed to enhance the survival of the producing organism by inhibition of other

micro-organisms in the same grth environment (Calvo etal., 2002; Akesson, et al.,

15

1996.). Characterization of the sub-cellular compartments of secondary metabolites can
aid in understanding their biological functions; storage and secretion mechanism, and
even efficient production via industrial fermentation.

Penicillin is perhaps the best-studied secondary metabolite (Van der Kamp et al.,
1999). The biosynthesis of penicillin in Penicillium chrysagenum involves three enzyme
activities that are distributed in at least in two sub-cellular compartments. The major
storage compartment for the substrates, or-aminoadipic acid, L-cysteine and L-valine, is
the vacuole (Van der Kamp et al., 1999). The first and second pathway enzymes were
localized in the cytosol (Van der Lende, 2002). The final enzyme was localized in the
microbody (Muller et al., 1991; Van der Lende, 2002). It was suggested that penicillin is
possibly synthesized and stored in this membrane-bound organelle (Muller et al., 1991).

To have full enzyme activity, it is important for biosynthetic enzymes to be
confined in the proper environment under conditions in which optimal pH, required
cofactors and a transport system for substrate are provided. Based on these observations, 1
hypothesized that microbodies and vacuoles represent two compartments for aflatoxin
synthesis. The biogenesis, protein import pathway and biological function of these two
organelles are briefly reviewed in the following sections.

Microbody. Micobodies can be divided into two subgroups based on the
enzymes they contain: peroxisomes (contain flavin-linked oxidase) and glyoxysomes
(contain enzymes of the glycoxylate cycle). The specific fiinction of microbodies also
varies in different cell types. In hepatocytes, peroxisomes can catalyze several metabolic
pathways including the biosynthesis of ether lipids, catabolism of long chain fatty acids

and production of cholesterol (Faumgart and Baumgart, 1993). Unlike animals where B-

16

oxidation occurs primarily in the mitochondria, B-oxidation in plants and fungi is
exclusively a peroxisomal function (Waterham and Cregg, 1997).

The biogenesis of peroxisomes can occur either by fission or budding from an
existing peroxisome or proliferation independent of a pre-existing peroxisome (Markham
et al., 1994; Rapp et al., 1996; Valenciano et al., 1998). The proliferation of peroxisomes
involves several maturation steps- from low-density organellar structures (pre-
peroxisome) to mature peroxisomes (Hettema et al., 1998). The low-density organellar
structures fine-peroxisome) contain several peroxisomal proteins including Pex10p and
ScPexl 1p that are responsible for independent peroxisome proliferation in yeast
(Waterham et al., 1997). PexlOp (34-48 kDa) is a peroxisomal integral-membrane
protein. ScPexl 1p (27 kDa) is a peroxisomal membrane-associated protein. Induction of
both genes leads to an over-proliferation of peroxisomes.

Proteins targeted to the peroxisome are mainly dependent on the Peroxisomal
Targeting Sequence (PTS) on the protein. Three kinds of PTS have been identified: PTS-
1, 2 and internal PTS (Subramani, 1998). PTS-1 and PTS-2 are commonly found in
peroxisomal proteins whereas the internal PTS is not yet clearly defined. PTS-l is a
tripeptide sequence, Ser-Lys-Leu (SKL) or conserved variants (S/A(/C))(K/R/H)L,
located at the carboxyl-terminus (Waterham et al., 1997). PTS-l is not cleaved after
import into the peroxisome. PTS-2 is an amino-terminal motif with a consensus sequence
Arg-Leu-Xs-His/Gln-Leu (RLX5H/QL). PTS-2 may be cleaved after import to the
peroxisome (Waterham et al., 1997). Internal PTS motifs may be contained in some
peroxisomal matrix proteins with or without PTS-1 and/or PTS-2. For example, the yeast
catalase A with PTS-l deleted is still able to be imported into the peroxisome (Purdue

and Lazarow, 1994). The potential position of internal PTS for this protein is located

17

between residues 104 and 126. Since the receptor for internal PTS has not been identified,
a proposed function for intemal PTS motifs is to associate and co-translocate with other
PTS-l-containing proteins into the peroxisome (Subramani, 1998).

Unlike mitochondria and ER proteins that require an unfolded conformation for
translocation, peroxisomal proteins can be imported in either extended monomeric
conformation or large preassembled structures (Waterham and Cregg, 1997). The
monomer without a PTS-2 sequence can be dimerized with a PTS-Z-containing monomer
and translocated into peroxisomes (Rachubinski and Subramani, 1995). Formation of
large structures via PTS-1 receptor-PTS-2 receptor by protein-protein interaction has
been demonstrated in a yeast system (Rachubinski and Subramani, 1995; Tabak et al.,
1999). The yeast (Saccharomyces cerevisiae) PTS-1 receptor (PTS-1R5), Pex5p, contains
a loosely conserved 34 amino acid repeat called the tetratricopeptide (TPR) domain. The
yeast PTS-2 receptor (PTSZR), Pex7p, contains six copies of a motif of 40 amino acid
residues (WD40 motif). WD is a Trp (VD-Asp (D) pair at the carboxyl terminus of the
motif. The results from two-hybrid experiments indicate that the binding between Pex7p
and Pex5p is mediated by interaction between the TPR domain and the WD motif
(Rachubinski and Subramani, 1995). Other proteins such as Pexlp and Pex6p with AAA
domain (ATPases associated with diverse cellular activities) are also involved in protein
import (Tabak et al., 1999). These proteins are associated with vesicles and interact in an
ATP-dependent manner in Pichia pastoris (Tabak et al., 1999).

A simple model for operation of a PTS—containing protein-import pathway is
demonstrated in Figure 1.3 (modified from Tabak et al., 1999). First, a PTS-containing
protein forms a complex with its receptor in the cytosol. Second, this complex binds to

docking proteins (translocon) in and on the peroxisomal membrane. Third, the PTS

18

protein is released into the peroxisomal matrix and the receptor is recycled. The docking
proteins consist of three proteins, Pex13p (an integral peroxisomal membrane protein),
Pexl4p (a peroxisomal membrane-associated protein) and Pex17p (membrane-associated
protein). Two additional helper proteins, hsp70—class cytoplasmic chaperones (Hsp70)
and J-domain—containing proteins (Djplp), are required in the recognition of PTS-
containing proteins by PTS receptors and/or the formation of PTS-1 and PTS-2 receptors
complex during the translocation process (Hettema et al., 1998). Based on a PSORT
search, the 6-aminopenicillanic acid acyltransferase from Penicillium chrysogenum and
Aspergillus nidulans contain a carboxyl-tenninal tripeptide ARL (PTS-1). The
experiments also confirmed that 6-aminopenicillanic acid acyltransferase is localized in
microbodies. A PSORT search suggested that several aflatoxin enzymes contain PTS-l
sequences but the locations of PTS-1 sequences in those enzymes are different from the
suggested location (carboxyl-terminal). Among them, AflJ even contains both PTS—1 and
2, located at the appropriate sites in the protein. The PSORT report for aflatoxin enzymes
is shown in Figure 1.4. So far, no published data have been obtained to confirm the
functional significance of these targeting signals.

Vacuole. The origin of vacuoles is not clear. It was suggested that the
endoplasmic reticulum may be involved directly in vacuole biogenesis and the Golgi
complex involved in increasing vacuole size (Klionsky, 1990). In vacuole biogenesis,
more than 40 proteins are required. Mutations in vacuole biosynthetic genes usually result
in fragmentation of the vacuole into small vacuoles, or vesicles, or unusual membrane
inclusions (Ohsumi et al., 2002). Proteins such as newly synthesized hydrolytic enzymes

destined for the vacuole are transported through the endoplasmic reticulum and Golgi

19

PTS protein

62

Helpers

A Hel ers
\/ \:/ P
. Receptors

 
 

 

 

   

    

 

 

Peroxisomal l Translocon
membrane L
Peroxisomal matrix ©

Figure 1.3. A schematic presentation of the peroxisomal-targeting sequence-dependent

protein-import pathway for proteins destined for the peroxisomal matrix.

20

&

AflJ

l. SKL motif (signal for peroxisomal protein): ARL (a.a. 354; n.t. 438)

2. SKL2: 2nd peroxisomal targeting signal: found [KLASIPLHL at a.a. 286]
Nor-1

SKL motif (signal for peroxisomal protein): ARL (a.a. 225; n.t. 271)
AvnA

1. SKL motif (signal for peroxisomal protein): SHL (a.a. 211; n.t. 396)

2. Peroxisomal proteins: Status: positive

VBS

SKL motif (signal for peroxisomal protein): No.

Ver-l

SKL motif (signal for peroxisomal protein): AKL (a.a. 75; n.t. 262)
OmtA

SKL motif (signal for peroxisomal protein): SHL (a.a. 81; n.t. 418)

OrdA

1. GvH: Examining signal sequence (von Heijne) Possible cleavage site: 23
2. Seems to have a cleavable N-term signal sequence.

3. SKL motif (signal for peroxisomal protein): AHL (a.a. 84; n.t. 528)

B_._

Acyl coenzyme A: 6-aminopenicillanic acid acyltransferase (from Penicillium

chrysogenum and Aspergillus nidulans)

21

1. SKL motif (signal for peroxisomal protein): SKL motif found at the C-terminus

2. Peroxisomal proteins: positive.

Figure 1.4. PSORT (Prediction of Protein Localization Sites version 6.4) prediction for
the protein targeting sequences (PTS-I and PTS-2) in (A) Aflatoxin enzymes, AflJ, Nor-
1, AvnA, VBS, Ver-l, OmtA and OrdA from A. parasiticus, and (B) 6-aminopenicillanic

acid acyltransferase from Penicillium chrysogenum and A. nidulans.

22

secretion pathway (Klionsky, 1990 and 1997). These proteins are carried by vesicles and
finally released into the vacuole. The process is completed‘by fusion between vesicles
and the vacuole with the help of small GTPases. The small GTPases recycle between
donor (vesicle) and receptor (vacuole) membranes (Ohsumi et al., 2002). GTP-bound
GTPase directs the fusion of vesicle to the vacuole. Upon fusion, GTPase hydrolyzes
GTP to GDP and is released from the membrane with the assistance of GTPase—activating
protein. The GDP-bound GTPase then returns to the donor membrane and GDP is
replaced by GTP with the assistance of guanine exchange factor.

The vacuole is a single-membrane bound organelle with highly dynamic structure
(Markham, 1994). Morphological changes in the vacuole occur rapidly and may
correspond to the grth stage (Markham, 1994; Paul, et al., 1994). Hyphal vacuolation
and fragmentation was only observed after a rapid growth phase during fungal
fermentation (Paul, at al., 1994). Recently, it was discovered that the vacuole is not just
an empty space as its electron microscopic appearance suggests but an organelle with
multiple possible functions (Markham, 1994). Therefore, vacuoles may consist of a series
of functionally distinct subclasses of organelles to compartmentalize a variety of
biological activities (Markham, 1994). The detailed functions of vacuoles in filamentous
fungi are not completely understood but may be similar to those in yeast (Klionsky,
1990). Here, vacuoles are large storage sites for metabolites, ions and amino acids. Ions
(Cafi, Mg”, Zn”, Fe”), including potential toxic ions (Srii, Co“, Pb”), and inorganic
phosphorus (in the form of polyphosphate) (Markham, 1994). S-adenosyl-methionine
(AdoMet) is an important methyl donor involved in a great variety of transmethylation
reactions. In S. cerevisiae, S-adenosyl-methionine accumulates to high levels (about 60%

of intracellular S-adenosyl-methionine) in vacuoles by a specific transport system

23

(Farooqui et al., 1983; Schwencke et al., 1976; Svihla et al., 1969). S-adenosyl-
methionine in vacuoles can be directly detected by microscopy under utraviolet light (at a
wavelength of 260 or 265 nm) (Farooqui et al., 1983; Schwencke et al., 1976; Svihla et
al., 1969). Results from subcellular fractionation revealed that two metabolically distinct
S-adenosyl-methionine pools exist in yeast. The labile pool exists in the cytosol while the
stable pool is in the vacuole (Farooqui et al., 1983). The vacuole is also an important
nitrogen reserve. Basic amino acids like arginine are concentrated in vacuole. The amino
acids a-aminoadipic acid, L-cysteine and L-valine used in penicillin synthesis are also
stored in vacuoles. Recently, the vacuole was found to be an important compartment for
lipid degradation and glycerol production at a specific stage during appressoria formation
in Magnaparthe grisea , suggesting that partial or complete enzymatic pathways may be
compartmentalized in vacuoles (Weber et al., 2001). Vacuoles control the intracellular
pH homeostasis (Thumm, 2000). The pH in vacuoles is maintained at pH6 in Neurospora
crassa. The membrane ATPase can pump H+ from the cytosol into the vacuole,
generating a membrane potential of about 25-40 mV in N. crassa. This electron-chemical
potential is the driving force to transport ions and amino acids across the vacuolar
membrane. Vacuolar hydrolases (e.g. aminopeptidase I) mature in this organelle after
cleavage of a signal peptide. Fungal vacuoles are an equivalent of mammalian lysosomes
in that they can be involved in degradation and recycling of unneeded proteins and even
whole organelles (Amor et al., 2000). In Saccharomyces, cytoplasmic proteins can be
transported into the vacuole by macroautophagy or the cytoplasm to vacuole targeting
(cvt) pathway without involvement of receptors. These two pathways are regulated by
nutrients (e.g. glucose or ethanol) and nutritional conditions (nutrition limitation)

(Klionsky, 1997 and Thumm, 2000).

24

1.6. Compartmentalization of penicillin in Penicillium chrysogenum.

The biosynthesis of penicillin in Penicillium chrysagenum involves three enzyme
activities and three amino acids (or-aminoadipic acid, L-cysteine and L-valine) (Van der
Kamp et al., 1999). In the first reaction, or-aminoadipic acid, L-cysteine and L-valine are
condensed to a 6-(L-a-aminoadipyl)-L-cysteinyl-D-valine tripepetide (ACV) catalyzed
by the enzyme 8—(L-or-aminoadipyl)-L-cysteinyl-D-valine synthetase (ACV synthetase).
The substrate amino acids (a-aminoadipic acid, L-cysteine and L-valine) are stored in
vacuoles. In early studies, ACV synthetase was localized in two subcellular
compartments, a Golgi-like organelle (Kurylowicz et al., 1987) and vacuoles (Lenderfeld
et al., 1993). Recently, ACV synthetase was also suggested to localize in the cytosol
(Van der Lende et al., 2002). The second reaCtion is conversion of ACV to isopenicillin
N (IPN) by the enzyme, isopenicilin N-synthetase. This enzyme has been localized to
Golgi-like organelles (Kurylowicz et al., 1987) and the cytosol (Van der Lende et al.,
2002). Most evidence suggests that the cytosol is the true compartment for this enzyme
(Muller et al., 1991 and Van der Lende et al., 2002). The final reaction entails the
conversion of IPN to penicilin G by Acyl-coenzyme A: isopenicillin N acyltransferase
(IAT). IAT was localized in peroxisomes (microbodies) (Mfiller et al., 1991 and Van der
Lende et al., 2002). In this step, the aminoadipyl side-chain of IPN is substituted by
phenylacetyl (PA) or phenoxylacetyl (POA) to yield penicilin G or penicilin V,
respectively (van de Kamp et al., 1999). Prior to this reaction, the PA and POA have to
be activated to their thioesters (PA-CoA and POA-CoA) by acetyl-CoA synthetase (ACS)

or PA-CoA ligase (PCL). ACS is a cytosolic enzyme. PCL contains a PTS-1 sequence,

25

suggesting that this enzyme may target to microbodies where it could function together
with IAT (van de Kamp et al., 1999). Based on localization results, Miiller et al.
suggested that penicillin is stored in the microbody together with IAT (Mfiller et al.,
1991). To summarize these studies, Van der Lende et al. suggested that the penicillin
biosynthetic activities are likely confined to two major compartments, the cytosol and
microbodies (Van der Lende et al., 2002). Other compartments observed in earlier studies
were proposed to be artifacts generated from the experimental procedures. A detailed
discussion is presented in two publications (Muller et al., 1991 and Van der Lende etal.,
2002). For the localization of ACV synthetase in vacuoles, van der Lende et al. suggested
that this artifact occurred during the isolation of protoplasts and organelles (Van der
Lende et al., 2002). Because the procedure is time-consuming (one day) and is conducted
under nutrition starvation conditions (cell wall digestion), this enzyme potentially could
relocate from the cytosol into the vacuole for protein degradation. For the localization of
ACV synthetase and isopenicilin N-synthetase in Golgi-like organelles, Muller et al.
suggested that organelles would be disrupted during grinding of mycelia and subsequent
fractionation procedure (Muller et al., 1991). The broken organelles could be reformed
into new organelles with morphology similar to other organelles.

To avoid potential artifacts introduced by vigorous cell disruption and time-
consuming organelle isolation procedures, another strategy for in situ localization of
protein is immuno-electron microscopy. In immuno-electron microscopic analysis, the
sample is processed first by fixation to prevent protein relocation and organelle
disruption. Therefore, the fixation method plays a crucial role in this analysis. In general,
the sample can be fixed by chemical crosslinking (chemical fixation) or low temperature

freezing (cryofixation). It is widely accepted that it is very difficult to obtain

26

simultaneously good preservation of both cellular ultrastructure and protein antigenicity.
For example, chemical fixation usually requires a longer incubation time because
crosslinking will inhibit chemical penetration and slow down the fixation process.
Prolonged crosslinking with chemicals like glutaradehyde may provide a good
preservation of ultrastructure but result in loss of antigenicity. The cryofixation method is
commonly considered superior to chemical fixation in preservation of ultrastructure and
protein antigenicity. This method physically fixes cellular components in a very short
time (ms) without any modification of protein antigenicity (Akesson et al., 1996 and
Muller et al., 1991). However, membrane components are easily extracted during
substitution of water with organic solvent. Although a low concentration of chemical
fixative can be added in the substitution solvent to maintain the membrane structure, this

may cause the loss of antigenicity.

27

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35

CHAPTER 2

Genetic Disruption of omtA in Aspergillus parasiticus C810

INTRODUCTION

Aflatoxins, highly toxic and carcinogenic secondary metabolites produced by the
filamentous fungi Aspergillus parasiticus, A. flavus, A. nomius, and A. tamarii (6,11),
frequently contaminate food and feed crops such as corn, cotton, peanuts and tree nuts
resulting in health risks to animals and humans (6). Most genes involved in aflatoxin
biosynthesis are clustered in a 75-kb genomic DNA region that carries a positive
regulator, AflR, and structural genes including omtA (17). Although much has been
learned about the molecular biology and biochemistry of aflatoxin biosynthesis, little is
known about the location of aflatoxin enzymes within a fungal colony or within a fungal
cell.

Early studies identified two enzymes, a 168 kDa protein and a 40 kDa protein
(OmtA), in Aspergillus parasiticus capable of converting sterigmatocystin to O-
methylsterigmatocystin (3,5) in vitro; about 60% of the enzyme activity in cell extracts
was associated with the 168 kDa protein and 40% of the activity was associated with

OmtA (4). Both enzymes required the co-factor S-adenosylmethionine. The 168-kDa
protein was purified to near homogeneity (3) and consisted of two subunits (58 and 110
kDa). OmtA (40 kDa) was also purified to homogeneity (8,9). The gene encoding OmtA
(omtA) was cloned from a cDNA expression library using OmtA-specific polyclonal

antibodies raised to OmtA protein expressed in E. coli (9, 16); omtA was later shown to

36

reside in the aflatoxin gene cluster (17) supporting its proposed role in aflatoxin
synthesis. Because sterigmatocystin and O-methlysterigmatocystin could be converted to
aflatoxin B, in vivo (2), they were generally accepted as aflatoxin pathway intermediates.
Two important issues remained unresolved. 1) Although the 168 kDa protein and OmtA
converted sterigmatocystin to O-methylsterigmatocystin in vitra, it was not clear which
of these activities was necessary and sufficient to catalyze this reaction in vivo (7, 10, 11,
14). 2) Because the methyl group on O-methlysterigmatocystin is removed during the
subsequent oxidoreduction reaction and because structurally related molecules are
incorporated into AFB, in feeding experiments with similar or higher efficiency than
sterigmatocystin (7), it was not clear if this reaction was necessary for aflatoxin synthesis.
To address these issues, I generated an omtA gene disruption mutant (LW1432). Feeding
studies conducted on LW1432 demonstrated a critical role for OmtA and the reaction

catalyzed by this enzyme in aflatoxin synthesis in viva.

MATERIALS AND METHODS

Fungal strains. A. parasiticus SU1(NRRL5862, ATCC 56775) is a wild—type,
aflatoxin-producing strain. A. parasiticus CSIO (ver-I wh-I pyrG) was derived from A.
parasiticus ATCC36537 (ver-I wh-I) (l3). LW1418 and LW1432 (ver-I wh-I omtA)
were generated in this study by disrupting the omtA gene in CSIO. AF 810 is a non-
aflatoxin producing aflR knockout strain derived from A. parasiticus NR1 (niaD) derived
fi'om SUI (provided by Dr. Jeff Cary, USDA, New Orleans, LA.).

Construction of omtA disruption vector pLW14. Plasmid pLW14 was

constructed by inserting pyrG, which encodes orotidine monophosphate decarboxylase

37

(13), into the coding region of omtA at the SphI site (Figure 2.1). A plasmid carrying
omtA genomic DNA was kindly provided by Dr. Fun Sun Chu (University of Wisconsin-
Madison, WI). The 2.5kb pyrG fragment was generated by polymerase chain reaction
(PCR) using plasmid pPG3J (12) as template. The primers used to amplify pyrG carried
an SphI (GCATGC) restriction site to facilitate cloning (5'-
GTAGAAGTTCAGQATGCTGATGG-3' and 5'-GAGTATCACAGTCAG_GCAT§C
ACGTC-3'). The reaction conditions for thermal cycling were: 95° C for 5 min followed
by 35 cycles at 95° C for 1 min, 62° C for 1 min, and 72° C for 3 min. The reaction was
completed by incubation at 72° C for 10 min.

Transformation and screening for omtA gene-disrupted strains. Circular or
linear plasmid (8 ug) digested with HindIII was used in transformation experiments using
A. parasiticus CSIO as the recipient strain (13). Protoplasts were generated by digestion
of mycelium (harvested 17 h after initiation of germination) with Novozyme 234 and
transformation was conducted by a PEG method as described previously (13). The

selection of omtA-disrupted strains was achieved in four steps. 1) pyrG-positive clones

(pyrG+; uridine prototrophs) were selected by grth on CZ agar (Czapek Dox Agar;
DIFCO Laboratories, Detroit, MI), a minimal growth medium (13). 2) Spores from 3 to 4
day old transfonnant colonies grown on CZ plates were transferred by sterile toothpick to
coconut agar medium (1) supplemented with sterigmatocystin at a concentration of 20
11ng and also to CZ agar medium. CSIO protoplasts regenerated and tolerated
sterigmatocystin supplementation of the agar during active growth and then utilized
sterigmatocystin to synthesize aflatoxin BI as active growth slowed or ceased. 3)

Colonies that did not fluoresce blue under long-wavelength UV light (254 nm) after a

38

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Figure 2.1. Restriction endonuclease map of plasmid pLW14.

39

3-day incubation at 29° C in the dark on coconut agar medium were selected and
secondary metabolites extracted and analyzed by thin-layer chromatography by a
previously published procedure (15). 4) The genotype of suspected omtA-disrupted
strains was confirmed by Southern hybridization analysis. OmtA null mutants were
further analyzed for ability to convert sterigmatocystin or O-methylsterigmatocystin to

aflatoxin B1 by a feeding experiment, as described below.

Southern hybridization analysis. Conidiospores (5 x 106) isolated from
transforrnants were inoculated into 100 ml of YES and incubated with shaking at 150 rpm
for 48 to 72 h at 29° C in the dark. One gram of freshly harvested mycelium was used for
isolation of genomic DNA using a published procedure (13). Southern hybridization
analysis was performed using a Non-Radioactive Hybridization Kit (Roche Molecular
Biochemicals, Indianapolis, IN) and a procedure provided by the manufacturer. To
identify the presence of pyrG sequences integrated within the chromosomal omtA gene,
genomic DNA was digested with HindIII and EcoRI, separated by electrophoresis on a
1% agarose gel, transferred to Nytran membrane, and probed with digoxigenin-labeled

2.9 Kb HindIII fiagment contained in the omtA gene.

Northern hybridization analysis. Conidiospores (5 x 106) isolated from
LW1418, LW1432, CSIO and SUI were inoculated into 100 ml of YES and incubated
with shaking at 150 rpm for 48 h at 29° C in the dark. Total RNA was purified from
freshly ground mycelium used TRIzol reagent (GibcoBRL, Rockville, MD). Northern
hybridization analysis was performed using a Non-Radioactive Hybridization Kit (Roche
Molecular Biochemicals, Indianapolis, IN) and a procedure provided by the
manufacturer. The total RNA (35 u g) was separated by electrophoresis (60 V, 4 h) on a

1% formaldehyde-agarose gel, transferred to Nytran membrane, and probed with a

40

digoxigenin-labeled 2.9 Kb HindIII fragment contained in the omtA gene. The gel was
stained with ethdium bromide (in 0.5 M ammonium acetate, at a final concentration of
0.5 pg per ml) for 40 min before the transferring procedure.

Feeding studies of omtA gene disruption strains. One gram of mycelia from
the same culture as used in Southern hybridization analysis was inoculated into 10 ml of
YES media supplemented with either sterigmatocystin or O-methylsterigrnatocystin at a
concentration of 20 pg per ml and incubated at 29° C for 24 h in the dark with shaking at
150 RPM. The fungal mycelium and culture medium were extracted with 10 ml of

chloroform at 4° C for 16 h. The chloroform fraction was evaporated under N2 gas and

resuspended in 200 pl of acetone. Ten pl samples were analyzed by TLC using

ether/methanol/water (96:32]) as the solvent system.

RESULTS

Generation of omtA disruption mutants. TLC analysis of cell extracts from pyrG+
transforrnants tentatively identified 5 omtA gene disruption isolates. Two isolates
(LW1418 and LW1432) were identified from among 41 pyrG+ colonies transformed with
circular plasmid LW14. Southern hybridization analysis of genomic DNAs isolated from
LW1418 and LW1432 detected a 2.5 kb pyrG DNA fragment within the omtA gene
(Figure 2.2.). Northern hybridization analysis of total RNAs isolated from LW1418 and
LW1432 did not detect any bands but a 1.6 kb band was detected in CS 10 and SUI,

confirming the gene disruption event in these isolates (Figure 2.3.).

41

Figure 2.2. Southern hybridization analysis of omtA gene disruption strains. A.
Restriction endonuclease map of omtA locus in omtA disruption strains LW1418 and
LW1432. B. Southern hybridization analysis. Genomic DNAs isolated from LW1418
and LW1432, CSIO (parent strain) and SUI (wild-type) were digested with restriction
enzymes HindIII and EcoRI. DNAs were resolved and hybridized to a digoxigenin-
labeled 2.9 kb HindIII omtA DNA fragment using standard methods. Strains with omtA-
disruption were predicted to contain a 5.4 kb HindIII fragment consisting of the 2.5 kb
pyrG selectable marker inserted into the omtA locus. A 5.2 kb fragment in the omtA-
disrupted strains was also predicted to replace the wild-type EcoRI fragment (2.7 kb).
Numbers to the right of the blot represent the approximate size of the detected fragments

in Kilobase pairs.

42

 

 

 

 

 

 

 

 

 

 

 

omtA
E 1H 1; IS H E
E: EcoRI, a: Hindu], s: Sphl l ‘—— ] ”'6
Probe: 2.9 Kb HindIII fragment
Digestion: Hind III EcoRl
Wild type: 2.9 Kb 8 Kb + 2.7 Kb
Mutant: 5.4 Kb 8 Kb + 5.2 Kb
mum Ecol“
F I l 1
1°. a 9.". 2
~ 2 1!. 1'. '7 9. E. 2'.
#9: 8 5 a. a 8 g 3 a,
__ 5.4 Kb
Figure 2.2.

43

Figure 2.3. Northern blot analysis of omtA gene disruption strains. (A) omtA transcript
accumulation assessed by Northern hybridization analysis. Total RNAs isolated from
LW1418 and LW1432, CSIO (parent strain) and SU-l (wild-type) were resolved and
hybridized to a digoxigenin-labeled 2.9 kb HindIII omtA DNA fragment using standard
methods. Strains with omtA-disruption were predicted to contain no omtA transcript and
a 1.6 kb transcript was predicted to be detected in both CSIO and SU-l. Equal loading of
RNA is demonstrated by Ethidium bromide staining of RNA as shown in panel (B).
Numbers to the left of the gel represent the approximate size of standard molecular

fragments in Kilobase.

44

9.49 l
7.46

4.40
2.37
1.35

0.24

Figure 2.3.

Northern Blot Analysis

no N
E E a
a E E a
_- .. - <— OmtA
EtBr Stain

‘— 1.6 kb

 

45

In vivo feeding experiments. No O-methylsterigmatocystin or aflatoxin B1

could be detected in LW1418 or LW1432 supplied with (fed) exogenous
sterigmatocystin; in contrast, each isolate converted exogenously supplied O-

methylsterigrnatocystin to aflatoxin B1 (Figure 2.43). The parental strain CS10
converted either sterigmatocystin or O-methylsterigmatocystin to aflatoxin B1 (Figure

2.4.A). These data suggest that LW 1418 and LW1432 carry mutant copies of omtA, but
the genes encoding later pathway enzymes and the regulatory genes involved in aflatoxin

biosynthesis are still functional.

DISCUSSION

The initial goal of this study was to determine if OmtA is necessary and sufficient
to convert sterigmatocystin to O-methylsterigmatocystin in vivo and if this reaction is
necessary for aflatoxin synthesis. When the same concentration of either sterigmatocystin
or O-methylsterigmatocystin was fed to the parent strain CSIO (wild type OmtA and
OrdA activities) nearly equal concentrations of O-methylsterigmatocystin and aflatoxin
B, were generated. This observation could result for one of at least three potential reasons.
I) The intermediates were fed in excess of the capacity of the pathway enzyme to carry
out complete conversion in the 24 hour period allotted. 2) The entry of the intermediates
into the cell or into the enzyme active sites was rate-limiting. 3) The intermediates
stimulate a feedback repression system downregulating activities of one or both enzymes.
Nevertheless, it was clear that CSIO could convert either sterigmatocystin or O-

methylsterigmatocystin to aflatoxin B, while LW1432 and LW1418 could convert

46

Figure 2.4. Thin layer chromatography of extracts of CS 1 0 and omtA-disruption strains
LW1418 and LW1432.
(A) Thin layer chromatography of extracts of CS10 supplied with (fed) exogenous

sterigmatocystin (ST) or O-methylsterigmatocystin (OMST). Lane 1 and 7, AF 81, AF 82,
AF G1 and AFGz standard mixture. Lane 2 and 6, OMST standard. Lane 3, CSIO fed no

pathway intermediate. Lane 4, CSIO fed sterigmatocystin. Lane 5, CSIO fed 0-
methylsterigmatocystin. Ten pl samples were analyzed using chloroform/acetone (95: 5)
as the solvent system.

(B) Thin layer chromatography of extracts of omtA -disrupted strains LW1418 and
LW1432 supplied with (fed) exogenous sterigmatocystin (ST) or 0-
methylsterigmatocystin (OMST). Metabolic scheme for aflatoxin biosynthesis in strain
LW1418 and LW1432 is shown at the top of the thin layer chromatograph. The ver-I and
omtA genes involved in aflatoxin biosynthesis are non-functional in LW1418 and 1432.

Lanes 1 and 8, OMST standard. Lanes 2 and 7, AF B1, AF Bz, AF G1 and AFG2 standard

mixture. Lane 3, LW1418 fed OMST. Lane 4, LW1418 fed ST. Lane 5, LW1432 fed
OMST. Lane 6, LW1432 fed ST. Abbreviations: VA, versicolorin A; DMST,
demthylsterigmatocystin. Ten pl samples were analyzed using ether/methanol/water (96:

3:1) as the solvent system.

47

OMST
AFB,
AFG,

 

B

ver-I dmrA omtA ordA
VA ’X" omsri>sr WX+ OMST ans,
1 2 3 4 5 6 7 8

OMST

AFB,

AFG,

 

Figure 2.4.

48

O-methylsterigmatocystin to aflatoxin B, but were unable to convert sterigmatocystin to
either O-methylsterigmatocystin or aflatoxin B,. These data clearly demonstrate that
OmtA is necessary for efficient conversion of sterigmatocystin to O-
methylsterigmatocystin in viva in A. parasiticus CSIO under the specified growth and
assay conditions. The data also demonstrate that this reaction is necessary for aflatoxin
biosynthesis. It is not yet clear if OmtA alone is sufficient to carry out this reaction in
viva. We cannot rule out the possibility that the 168 kDa protein contributes to this
reaction, but its contribution appears negligible compared to OmtA. In support of this
idea, LW1432 (omtA disruption) could not convert exogenously supplied
sterigmatocystin to detectable levels of either O-methylsterigmatocystin or aflatoxin B,

but could convert O-methylsterigmatocystin to aflatoxin B1 in viva. In contrast, CSIO

(parent of LW1432) converted either sterigmatocystin or O-methylsterigmatocystin to
aflatoxin B,. These data demonstrate that the omtA disruption is gene-specific and does

not affect the activity of late pathway enzymes or regulators of aflatoxin synthesis.

49

REFERENCES

l. Arseculeratne, S. N., L. M. De Silva, S. Wijesundera, and C. H. S. R. Bandunatha.
1969. Coconut as a medium for the experimental production of aflatoxin. Appl.
Microbiol. 18:88-94.

2. Bhatnagar, D., S. P. McCormick, L. S. Lee , R. A. Hill. 1987. Identification of O-
methylsterigmatocystin as an aflatoxin B1 and GI precursor in Aspergillus parasiticus.
Appl. Environ. Microbiol. 53:1028-33.

3. Bhatnagar, D., A. H. Ullah, and T. E. Cleveland. 1988. Purification and
characterization of a methyltransferase from Aspergillus parasiticus SRRC 163 involved
in aflatoxin biosynthetic pathway. Prep. Biochem. 18:321-49.

4. Bhatnagar, D., T. E. Cleveland, and E. B. Lillehoj. 1989. Enzymes in aflatoxin B1
biosnythesis: Strategies for identifying pertinent genes. Mycopathologia. 107:75-83.

5. Cleveland, T. E., A. R. Lax, L. S. Lee, and D. Bhatnagar. 1987. Appearance of
enzyme activities catalyzing conversion of sterigmatocystin to aflatoxin BI in late-
growth-phase Aspergillus parasiticus cultures. Appl. Environ. Microbiol. 53:1711-3.

6. Eaton, D. L. and J. D. Groopman (eds.). 1994. The toxicology of aflatoxins: human
health, veterinary, and agricultural significance. San Diego, CA: Academic.

7. Gengan R. M., A. A. Chuturgoon, D. A. Mulholland and M. F. Dutton. 1999.
Synthesis of sterigmatocystin derivatives and their biotransformation to aflatoxins by a
blocked mutant of Aspergillus parasiticus. Mycopathologia 144:115-122.

8. Keller, N. P., H. C. Dischinger, JR., D. Bhatnagar, T. E. Cleveland and A. H. J.
Ullah. 1993. Purification of a 40-kDa methyltransferase activity in the aflatoxin
biosynthetic pathway. Appl. Environ. Microbiol. 59:479-484.

9. Liu, B. H., N. P. Keller, D. Bhatnagar, T. E. Cleveland and F. S. Chu. 1993.
Production and characterization of antibodies against sterigmatocystin O-
methyltransferase. Food Agric. Immunol. 5:155-164.

10. Mito, R. E. and C. A. Townsend. 1997. Enzymology and molecular biology of
aflatoxin biosynthesis. Chem. Rev. 97:2537-2555.

11. Payne, G. A. and M. P. Brown. 1998. Genetics and physiology of aflatoxin
biosynthesis. Annu. Rev. Phytopathol. 36:329-362.

50

12. Reichard, U., G. T. Cole, T. W. Hill, R. Ruchel and M. Monod. 2000. Molecular
characterization and influence on fungal development of ALP2, a novel serine proteinase
from Aspergillusfumigatus. Int. J. Med. Microbiol. 290:549—558.

13. Skory, C. D., J. S. Horng, J. J. Pestka, and J. E. Linz. 1990. Transformation of
Aspergillus parasiticus with the homologous gene (pyrG) involved in pyrimidine
biosynthesis. Appl. Environ. Microbiol. 56:3315-3320.

14. Sweeney, M. J. and A. D. W. Dobson. 1999. Molecular biology of mycotoxin
biosynthesis. FEMS Micobiology Letters. 175:149-163.

15. Trail, F., P-K. Chang, J. Cary and J. E. Linz. 1994. Structural and functional
analysis of the nor-1 gene involved in the biosynthesis of aflatoxin by Aspergillus
parasiticus. Appl. Environ. Microbiol. 60:3315-3320.

16. Yu, J., J. W. Cary, D. Bhatnagar, T. E. Cleveland, N. P. Keller, F. S. Chu. 1993.
Cloning and characterization of a cDNA from Aspergillus parasiticus encoding an O-
methyltransferase involved in aflatoxin biosynthesis. Appl. Environ. Microbiol. 59:3564-
71.

17. Yu, J., P-K. Chang, J. W. Cary, M. Wright, D. Bhatnagar, T. E. Cleveland, G. A.

Payne, and J. E. Linz. 1995. Comparative mapping of aflatoxin pathway gene clusters in
Aspergillus parasiticus and Aspergillusflavus. Appl. Environ. Microbiol. 61 :2365-2371.

51

CHAPTER 3

Expression/Distribution of the OmtA Protein in Colonies of Aspergillus parasiticus

INTRODUCTION

Previous studies on accumulation of aflatoxin enzymes and expression of
aflatoxin genes including omtA were conducted primarily in liquid medium using
submerged shake culture (batch fermentation) (6,22,25); these conditions induce aflatoxin
synthesis but do not support asexual sporulation (conidiation). Because a close regulatory
association has been demonstrated between aflatoxin synthesis and conidiation (5,7,9) we
hypothesized a spatial and temporal association between omtA expression and
conidiospore development. Our goal was to develop a grth model that would closely
mimic regulation of toxin synthesis in soil and on the host plant. We developed a novel
time-dependent colony fractionation protocol to study OmtA accumulation in fungal
colonies grown on solid medium; these conditions support toxin synthesis and
conidiation. This protocol also allowed analysis of OmtA distribution to different cell
types in fungal colonies. OmtA-specific polyclonal antibodies were generated against
OmtA fusion protein (MBP-OmtA) and purified by affinity chromatography using a
LW1432 protein extract. OmtA was not detected in 24 h old colonies but was detected in
48 h old colonies using Western blot analysis; the protein accumulated in all regions of a
72 h old colony including cells (0 to 24 h old near the colony margin) in which little
conidiophore development was observed. OmtA in older parts of the colony (24 to 72 h)

was partly degraded. F luorescence-based immuno-histochemical analysis conducted on

52

thin sections of paraffin-embedded fungal cells from time-fractionated fungal colonies
demonstrated that OmtA is evenly distributed among different cell types and is not
concentrated in conidiophores. These data suggest that OmtA accumulates in newly
formed fungal tissue and then is proteolytically cleaved as cells in that section of the
colony age. The data also suggest that OmtA is localized to specific areas within a fungal
cell but it is not yet clear if these areas correspond to 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.

MATERIALS AND METHODS

Fungal strains. A. parasiticus SU1(NRRL5862, ATCC 56775) is a wild-type,
aflatoxin-producing strain. A. parasiticus CSIO (var-1 wh-I pyrG) was derived from A.
parasiticus ATCC36537 (ver-I wh-I) (21). LW1418 and LW1432 (ver-I wh-I omtA)
were generated in this study by disrupting the omtA gene in CSIO. AF $10 is a non-
aflatoxin producing aflR knockout strain derived from A. parasiticus NR-I (niaD) which

in turn was derived from SUI.

Construction of OmtA expression vector pLW12. An omtA cDNA was
generated by reverse transcriptase- PCR (RT-PCR). Template RNA was isolated from A.
parasiticus strain SUI cultured in YES media for 48 to 72 h using Trizol reagent and a
procedure supplied by the manufacturer (GibcoBRL, Rockville, MD). For first strand
cDNA synthesis, 48 pg of total RNA were incubated in the RT-PCR mix at 37° C for 2 h.

All chemicals used in the RT-PCR were purchased from GibcoBRL (Rockville, MD).

53

The 20 pl reaction mixture contained: 4 pl of 5 X first strand buffer, 2 pl of 0.1 M DTT,
1 pl of 10 mM dNTP, 2 pl of M-MLV Reverse Transcriptase (200 U per pl), and 1 pl of
Oligo (dT) primer (0.5 pg per pl). One primer for omtA amplification contained a HindIII
(AAGCTT) restriction site and the other primer an XbaI (TCTAGA) restriction site to
facilitate cloning (5'-CCCTCTAGAATG GCACTACCGAGCAAAG-3' and 5'-
TGCAAQCTTCTACTTGCGCAAACGCAGT-3'). One pl of the resulting cDNA
mixture was used as template for PCR. The cycling conditions were: 95° C for 5 min
followed by 35 cycles of 95° C for 1 min; 55° C for 1 min, and 72° C for 2 min. The
mixture was incubated at 72° C for 10 min to complete the reaction. The omtA PCR
fragment (1260 bp) was digested with restriction enzymes XbaI and HindIII and cloned
into plasmid pMAL-c2 (New England Biolabs, Beverly, MA) digested with the same
enzymes. The resulting plasmid construct, pLW12, was transformed into E. cali DHSa.
The proper construction of pLW12 in clones expressing MBP-OmtA was confirmed by
restriction enzyme analysis of purified plasmid DNA isolated by the Qiagen mini prep
plasmid kit (Qiagen, Valencia, CA). The size of the fusion protein was determined by
small-scale expression studies. E cali DHS or carrying pLW12 was incubated in 5 m1 of
LB broth containing Ampicillin (100 pg per ml) for 16 h. One ml of bacterial culture was
saved as non-induced control. The remaining 4 ml of culture was induced to express

fusion protein by addition of 0.3 mM of IPTG for 3 h.

Conversion of sterigmatocystin to O-methylsterigmatocystin by maltose
binding protein -OmtA. The pMAL Protein Fusion and Purification System from New
England Biolabs (Beverly, MA) was utilized to express and purify Maltose Binding

Protein-OmtA. Large-scale preparation of Maltose Binding Protein-OmtA was conducted

54

for the study of enzymatic activity and antigen purification. E cali DH5 or carrying
pLW12 was grown in 500 ml of rich media (10 g tryptone, 5 g yeast extract, 5 g NaCl,
2% glucose) containing ampicillin (100 pg per ml). Fusion protein synthesis was induced
by addition of IPTG (0.3 mM), cells were sonicated (Sonifier cell disrupter W-350;
Fisher Scientific, Pittsburgh, PA) and the fusion protein was purified by amylose affinity
column chromatography according to a protocol supplied by the manufacturer (New
England Biolabs, Beverly, MA).

Amylose resin-purified Maltose Binding Protein and Maltose Binding Protein -
OmtA were used for analysis of enzyme activity. In a time-course experiment, three l-ml
reactions were prepared. Each reaction mixture contained 100 pg of Maltose Binding
Protein-OmtA, 20 pg of sterigmatocystin, and 600 pg of S-adenosylmethionine.
Sterigmatocystin, O-methylsterigmatocystin and S-adenosylmethionine were purchased
from Sigma (St. Louis, MO). The reaction mixtures were incubated at room temperature
for appropriate times (20-min time points for 60 min; one tube per time point). Three
control reactions were also incubated for 60 min. Each contained the same reagents as
above except the Maltose Binding Protein-OmtA or sterigmatocystin or S-
adenosylmethionine were omitted. Reactions were stopped by extraction with 4 volumes
of chloroform. Chloroforrn extracts were dried under nitrogen gas and resuspended in 100
pl of acetone. Five pl of extract was analyzed by TLC using chloroform lacetone (95:5)

as a development system (23).

OmtA antibody production and purification. OmtA polyclonal antibodies
were generated in rabbits using purified MBP-OmtA as antigen. Each of 2 rabbits (New

Zealand White) was injected subcutaneously with 300 pg of protein in TitreMax adj uvant

55

(Cthx, Norcross, GA) at 1: 1 (V/V) ratio. After 35 days, the animals received one
booster injection with 200 pg of protein in TitreMax adjuvant. Serum was obtained 4
weeks after the boost. The IgG fraction was purified from rabbit serum by precipitation
with ammonium sulfate using standard procedures (2).

The IgG fraction of OmtA antibodies was firrther purified by affinity
chromatography using a protein extract prepared from LW1432 grown in YES media for
48 h. This column was prepared by conjugating 11 mg of total protein to 2 ml of
Aminolink coupling gel (coupling efficiency approximately 80%) using a procedure
supplied by the manufacturer (Pierce Chemical Company, Rockford, IL). For antibody
purification, one ml of IgG (8 mg) was loaded onto the gel bed in this column and an
additional 2 ml of PBS was loaded to cover the gel bed. The column with antibodies was
incubated at room temperature for 1.5 h at room temperature. After incubation, PBS was
allowed to flow-through and antibody-containing fractions were identified by absorbance
at 280nm. The protein concentration was determined by a BIO-RAD protein assay
(Hercules, CA) using Dye Reagent and BSA (Fraction V, Sigma, St. Louis, MO) as
standard. This highly purified antibody preparation (3 mg per ml) was used in Western
blot analysis and immuno-fluorescence microscopy.

Western blot analysis. To determine specificity of anti-OmtA antibodies, fungal
proteins were extracted from mycelia for Western blot analysis. Fungal strains including
SUI, CSIO, LW1418, LW1432, LW1468 and LW1470 were grown in the dark at 29°C
(shaking at 150RPM) in 100 ml of YES liquid media for 24, 48 and 72 h or only 48 h.
The mycelia were harvested, pulverized under liquid nitrogen and the fungal proteins
were extracted in TSA buffer (2 mM Tris-Cl, pH8.0; 40 mM NaCl and 0.025 % sodium

azide) containing complete protease inhibitors (Roche Molecular Biochemicals,

56

Indianapolis, IN). Proteins were resolved by SDS polyacrylamide gel electrophoresis
using standard methods (2). For Western blot analysis, each lane on the 12% SDS-PAGE
contained 30pg protein. The primary Ab was column-purified anti-OmtA IgG (1 pg per
ml) and the secondary Ab consisted of a 10,000-fold dilution of goat anti-rabbit IgG
alkaline phosphate conjugate (Schleicher & Schuell, Keene, NH). A BCIP/NBT
colorimetric detection system was utilized (Roche Molecular Biochemicals, Indianapolis,
IN).

The antibody against native OmtA was kindly provided by Dr. Fun Sun Chu
(University of Wisconsin-Madison, WI). This antibody was also subtraction-purified with
a LW1432 strain (omtA gene disrupted) protein extract before use in Western blot
analysis (Figure 3.2.). 1.4 pl of antibody preparation was incubated with 100 pl (800 pg)
of LW1432 protein extract for I h at 4° C. After incubation, the preparation was
centrifuged at 12,000 x g for 15 min at 4° C to pellet cross-reactive antibodies. The

supernatant was mixed with 14 ml of blocking reagent and used in Western blot analysis.

Time-dependent fractionation of colonies grown on solid medium. To

determine the accumulation and distribution of OmtA in fungal colonies grown on solid

culture media, conidiospores (2 x 105) ofA. parasiticus SUI, AFSIO, c310, and
LW1432 were inoculated onto the center of YES agar or PDA agar (for SUI only)
overlaid with sterile cellophane membranes and incubated at 29° C in the dark. Some
colonies were analyzed after 24 or 48 h of growth. 72 h-old colonies were fractionated
into three concentric rings based on area covered at three time points (72, 48 and 24 h).
For example (Fig. 5D), a SUI colony with a diameter of 4.2 cm was fractionated to SI

which contained mycelia from the colony center out to a distance of 0.8 cm (ages 48 to 72

57

h), 82 which contained mycelia from 0.8 to 2.5 cm (24 to 48 h) and S3 which contained
mycelia from 2.5 to 4.2 cm (0 to 24 h). The harvested mycelia from appropriate sections
of the colony were pulverized under liquid nitrogen and the fungal proteins were
extracted in TSA buffer containing complete protease inhibitors. Western blot analysis

was performed as described above.

Immuno-localization of OmtA protein. Immuno-localization of OmtA protein
was conducted on SUI colony fractions. In addition, AF S10, CSIO, and LW1432
colonies were fractionated following the same scheme to generate analogous fractions

R1, R2, R3, C1, C2, C3, and L1, L2, L3, respectively.

Preparation of paraffin-embedded fungal sections. Samples from fungal
colony fractions were embedded in paraplast (Sigma, St. Louis, MO) using a published
procedure (2) with the following modifications. Fungal tissues were fixed with Streck
tissue fixative (Streck Laboratory Inc., Omaha, NE) at 4° C overnight, 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 min, 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 paraffin/xylene mixture (1:1; v/v) for 15 min, 60 min and overnight at 600C
and finally for 8 h in 100% paraffin at 60° C (three changes of paraffin during
incubation). The paraffin embedded sample blocks were hardened in a plastic mold

(V WR Scientific, Detroit, MI) at RT. Sample blocks were cut into 4 pm thick sections

58

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-labeling. Coverslips with paraffin-embedded fungal 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 immuno-labeled with primary
antibody (purified anti-OmtA IgG; 20 pg/ml) or anti-SKL (1:500, Zymed, So. San
Francisco, CA) at room temperature for 1.5 h, followed by secondary antibody
conjugated with fluorescent probe (goat anti-rabbit IgG-Alexa 488 conjugate [5 pg/ml];
Molecular Probes; Eugene, OR), at room temperature 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
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 with 488 nm laser line. The 40

X oil objective lens (Zeiss Plan-NeoFlura, NA: 1.3) was used to acquire all images. The

59

Alexa 488 fluorescence probe (Abs 495 nm/ Em 519 nm) was detected using LP 520 or
BP 520-560 barrier filters. Fluorescence image analysis of SUI, AF S10, CSIO and

LW1432 was conducted under the same instrument parameter settings.

RESULTS

Western blot analysis and protein localization using PAb against native
OmtA. The antibodies against purified native OmtA were kindly provided by Dr. Fun
Sun Chu (University of Wisconsin-Madison, WI). The specificity of this PAb was tested
on SUI (wild type) cultured in YES liquid medium for 24, 48 or 72 h. One primary band
(larger than 45 kDa) was detected in Western blot analysis of protein extracts prepared
from 24 h culture and three primary bands were detected in 48 or 72 h cultures (Figure
3.1.A). Protein localization using a slide-culture method (12) showed the primary signal
was detected in conidiophores and conidiospores (Figure 3.1 .B). This antibody was
further purified by subtraction with a protein extract from LW1432 (omtA gene disruption
strain) and the specificity was tested by Western blot analysis. Two primary bands were
detected in SUI and CSIO and one intense band was detected in all omtA knock-out
strains, suggesting that this purified antibody still cross-reacted with one protein (Figure

3.2.).

Purification of Maltose Binding Protein-OmtA. Maltose Binding Protein-
OmtA in DH50L cell extracts (Figure 3.3.A) was purified by amylose column
chromatography. Most of the purified Maltose Binding Protein-OmtA fusion protein was

approximately 88 kDa in mass (Figure 3.3.8); this is consistent with a mass calculated

60

from the fusion of 42 kDa MBP and 45 kDa OmtA predicted using nucleotide sequence
data. Very little proteolytic cleavage of the protein was detected by this analysis. The
fusion protein remained soluble in the bacterial cytoplasm allowing purification of 39 mg

of MBP-OmtA per liter of bacterial culture.

Enzymatic conversion of sterigmatocystin to O-methylsterigmatocystin by
Maltose Binding Protein-OmtA. Purified OmtA fusion protein efficiently converted
sterigmatocystin to O- methylsterigrnatocystin in the presence of S-adenosylrnethionine
(Figure 3.3.C) within 60 min. The cofactor, S-adenosylmethionine, was required for this
conversion. Without exogenous S-adenosylmethionine, no O-methylsterigmatocystin
could be detected in the presence of the Maltose Binding Protein-OmtA. As expected, no
O-methylsterigmatocystin was detected in the reaction that contained S-
adenosylrnethionine without substrate (sterigmatocystin), in a reaction without added
Maltose Binding Protein-OmtA, or in a reaction with Maltose Binding Protein but

without the Maltose Binding Protein-OmtA (data not shown).

Production of PAb against OmtA fusion protein. To obtain highly specific
anti-OmtA polyclonal antibodies (PAb), amylose-purified MBP-OmtA was used as
antigen to produce polyclonal antibodies in two rabbits. The IgG fraction of the rabbit
serum was further purified by affinity chromatography using a protein extract from
LW1432 (omtA gene disruption strain). The specificity of anti-OmtA PAb was tested on
SUI (wild type) cultured in YES liquid medium for 24, 48 or 72 h. No OmtA could be
detected in Western blot analysis of protein extracts prepared from SUI grown on YES

liquid medium for 24 h while one primary band (approximately 45 kDa) could be

61

Figure 3.1. Western blot analysis and fluorescence microscopy using polyclonal
antibodies raised against partially purified fungal OmtA proteins. The native OmtA
antibody was kindly provided by Dr. Fun Sun Chu (University of Wisconsin-Madison,
W1).

(A) Analysis of SUI crude protein extracts isolated at three time points (24, 48 and 72 h)
from fungal cultures grown in YES liquid medium. The approximate mass of the primary
signal (45 kDa) is shown to the right of the blots.

(B) OmtA protein localization in a colony of A. parasiticus SUI grown on YES agar for
48 h. Fungal tissue from slide-culture was digested with Novezyme and probed with
OmtA PAbs (20 pg/ml) followed by FITC-conjugated goat anti-rabbit IgG (1:50). The

protein localization method used was reported previously (12).

62

Western Blot Analysis: anti-OmtA

24H 48H 72H
lfi F—I l_‘l

MW.
45 _ ————-—-—-...,...‘-m--—_-
--w-
31 _ — -— —- -—
B.
Bright Field OmtAFITC

IYNK
1111111111,, ,
willfl ‘11“ :3,“ ““3,“ \11 ll

 

. ‘
110““ “m“?
1 mill lsllllll

Figure 3.1.

63

 

ab 6'! co N no a
v-1 I") F! ("I ‘9 l‘
g c r. z z z z z a
t— "‘ 5 3 3 3 B 3 3
m a o .4 ...i ...r .1 .1 .2 Em"
- U 66kDa
OmtA» 19191:; - g __' ” """'"" """"‘" .2-.. ““'“"' _ 45 kDa
. J 31 kDa

Figure 3.2. Western blot analysis of firngal protein extracts using subtraction-purified
PAb raised against native OmtA. Crude protein extracts were isolated from wild-type
aflatoxin-producing strain SUI, parent strain CSIO, and omtA knock-out strains,
LW1418-1, LW1432-1, LW1418, LW1432, LW1468 and LW1470, grown in YES liquid
medium for 48 h. LW1418-l and LW1432-1 are single spore isolates of LW141 8 and
LW1432, respectively. The native OmtA antibodies used were subtraction-purified with
LW1418 protein extract two times. The possible OmtA reactive band (45 kDa) is
indicated by an arrow at the left of the blots. Lane STD contains molecular mass

standards; mass of standards is shown at the right of the blot.

64

Figure 3.3. Conversion of ST to OMST by affinity purified MBP-OmtA fusion protein.
(A) SDS-PAGE of IPTG- induced bacterial crude extracts containing MBP-OmtA or
MBP. (B) SDS-PAGE of affinity purified MBP-OmtA. Lanes MWI and MW2 represent
molecular mass standards; molecular mass is indicated to the left and right of the gel. (C)
Thin layer chromatography to catalyze the conversion of ST to OMST by affinity purified
MBP-OmtA and controls. Five pl of reaction mixture was analyzed after appropriate
time periods by TLC using chloroform /acetone (95:5) as the developing system. Control
reactions included all reagents except ST, SAM, or MBP-OmtA, respectively. OMST

standards are located in lanes on both ends of the TLC (STD: OMST).

65

E"
I”

 

Z
a:
'F
o z
i 96 MWI
.3’ - .-.. kDa Purified
MBP-OmtA
1"“ kDa MWI l—fi MWI MW2 RD:
201
97 116
- - - - - a so
66 - " -
31 - - g 32
-_ ..._ ,__. .fi
3
1% i
C a? 3
' S a 3
+ 3 +
w MBP-OmtA % + 3.” m
g +ST+SAM z ‘11 2 g
3 20’ 4o: 60’ 60’ 60’ 60’ E
U) m
-I -1

Figure 3.3.

 

66

detected in SUI grown for 48 or 72 h in the same medium (Figure 3.4.A).

Western blot analysis of OmtA in colonies grown on solid medium. The
accumulation and distribution of OmtA was analyzed in time-fractionated colonies grown
on solid YES or PDA agar medium for 72 h (Figure 3.4.B, C and D). OmtA protein was
detected in fractions SI, S2, and S3 of a 72 h old SUI colony but not in corresponding
colony fractions isolated from AF S10 (aflR knockout) (Figure 3.4.D) or LW1432 (omtA
knockout) (Figure 3.4.C). OmtA in colony fractions S1 (48 to 72 h) and 82 (24 to 48 h)
showed increased levels of smaller peptides and correspondingly less full-length protein.
In contrast, in fraction S3 (0 to 24 h) of a 72 h old colony, full length OmtA was found at
higher levels. A similar result was observed using SUI grown on PDA medium and CSIO
(parent strain of LW1432) grown on YES agar medium; very little OmtA protein could
be detected in fraction SI and C1 (48 to 72 h) while more full-length protein was detected
in the youngest colony fraction (S3 and C3; 0 to 24 h) (Figure 3.4.C). No OmtA protein

could be detected in any colony fraction of either LW1432 or AF SlO.

Confocal Laser Scanning Microscopy. Immuno-fluorescence microscopy was
conducted on samples prepared from fractionated colonies grown on YES solid media.
To compare the fluorescence labeling intensity between SUI and CSIO and their non-
aflatoxin producing counterparts, AF S10 and LW1432, respectively, the samples were
viewed under the lowest zoom level during the Confocal Laser Scanning Microscopy

(zoom = 20) in order to acquire maximum cell number within a field. Under the same

67

Figure 3.4. Western blot analysis of fungal protein extracts using affinity purified OmtA
PAb.

(A) Analysis of SUI crude protein extracts isolated at three time points (24, 48 and 72 h)
from fungal cultures grown in YES liquid medium. The approximate mass of the primary
signal (45 kDa) is shown to the right of the blots in panels A and B, and to the left of
blots in panels C and D. (B) Analysis of protein extracts from time—fractionated colonies
of SUI grown on PDA agar medium. 72 h-old colonies were fractionated into three
concentric rings based on area covered at three time points. SI, 48 to 72 h; S2, 24 to 48 h;
S3, 0 to 24 h. (C) Analysis of protein extracts from time-fractionated colonies of CSIO
and LW1432 grown on YES agar medium. (D) Analysis of protein extracts from time-
fractionated colonies of SUI and AF SIO grown on YES agar medium. Lane STD

contains molecular mass standards; mass of standards is shown at the right of panel C.

68

A

Medium Liquid YES

 

 

 

 

 

 

Strain SUI
24h 48h 72h
-—---—1-45kDa
YES Agar
CSIO LW1432
0.8 1.6 2.7 0.8 2.1 3.5
C1 C2 C3 STD L1 L2 L3
kDa
250
I48
60
! 42
30
22

 

 

 

Figure 3.4.

69

Medium
Strain

PDA Agar
SUI

Diameter (cm) 1.0 2.2 3.3

Fractions

818283

 

‘-- + 45 kDa

 

 

 

YES Agar
SUI AFSIO

l j l N

0.8 2.5 4.2 0.8 2.5 4.0
SI 82 S3 R1 R2 R3

 

 

45 kDa ->

 

Mb

 

 

contrast and brightness settings, the samples prepared from the control strain AF SIO
(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 SUI (fractions S1, 82, and S3) or CSIO (fractions C1, C2 and
C3) (Figure 3.5.). These data are consistent with Western blot analysis of protein extracts
isolated from the same colony fractions (Figure 3.4.C and D; i.e., OmtA was only
detected in SUI and CSIO but not in AF S10 or LW1432). OmtA was observed in the
substrate level mycelium throughout a 72 h old colony grown on YES media; the protein
was also detected in the conidiospore-bearing structures (particularly in vesicles; Figure
3.6.B) located in fractions S1 and S2 at nearly equal intensity as in substrate level
mycelium. The fluorescent signal was confined to discreet areas (patches) within cells
(Figure 3.6.A). However, due to limitations in resolution of confocal laser scanning
microscopy, 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 (Figure 3.6.C) (10,23). The
observed fluorescent pattern was consistent with localization to these organelles. In
addition, we used SYTOX Green to stain nuclei in fungal fractions (Figure 3.6.D); again
the fluorescent pattern was consistent with the expected results. Neither anti-SKL nor
SYTOX generated the same “patchy” pattern as anti-OmtA. In summary, the data
suggest that OmtA protein 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.

70

Figure 3.5. OmtA protein localization in time-fractionated colonies of A. parasiticus
SUI, AF S10 (AfR knockout), CSIO and LW1432 (omtA knockout) grown on YES agar
for 72 h. Paraffin-embedded fungal sections were immuno-labeled with affinity purified
OmtA PAb (20 pg/ml) followed by Alexa 488 conjugated goat anti-rabbit IgG.

(A) Fluorescence images of SU-I (SI, S2, and S3) and AF 510 (R1, R2, and R3).

(B) Fluorescence images of CSIO (C1, C2 and C3) and LW1432 (L1, L2 and L3). All

colony fractions were analyzed under the same instrument settings. Bar = 50 pm.

71

 

Figure 3.5.

72

 

Figure 3.6. Protein localization using OmtA PAb and anti-SKL, and detection of nuclei
using SYTOX Green. (A) Z-series overlay image of SUI (fraction S2) 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 81) immunolabled with OmtA PAb. (C) Immuno-
labeling of SUI (fraction 82) with anti-SKL. The magnification for the left panel is 400X
and for right panel is 2,000 X. (D) Fluorescence image of nuclei in SUI (fraction S2)
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.

73

DISCUSSION

Because a close regulatory association between aflatoxin synthesis and
conidiation has been demonstrated in several studies (7,9), we hypothesized a close
temporal and spatial association between OmtA expression and conidiospore
development. This study was designed to address this hypothesis. First, it was necessary
to develop highly specific OmtA antibodies and a method for analysis of protein
accumulation and distribution in cells and fungal colonies grown on solid growth
medium.

We initially experienced specificity problems with PAb (15) raised to native
OmtA that resulted in severe cross-reactivity in Western blot analysis and artifacts in
protein localization (Figure 3.1 .). Because aflatoxin enzymes are present at low
concentration in the fungus (I I), purification of OmtA from the fungus likely resulted in
co-purification of at least trace amounts of other proteins that, although undetectable by
SDS-PAGE, still possessed strong antigenicity.

To increase specificity, we generated PAb against affinity-purified MBP-OmtA
and further purified them by affinity chromatography with a column carrying fungal
proteins isolated from LW1432 (omtA knockout). This procedure helped eliminate cross-
reactive antibodies. Specificity was demonstrated via Western blot analysis. OmtA could
not be detected in either AF $10 or LW1432, strains that do not synthesize this protein.
OmtA PAbs did not detect OmtA after 24 h in YES liquid or solid media but did detect a
protein of appropriate size (45kDa) at 48 and 72 h. This pattern of accumulation is

similar to that observed for other aflatoxin proteins including Nor-1 (26), Ver-l (13,14)

74

and VBS (unpublished data) suggesting that accumulation of these proteins is
coordinately regulated. In addition, analysis of fungal extracts from SU-l, CSIO, AF 810
and LW1432 did not show significant cross-reaction of OmtA PAb to other cellular
proteins including DmtA; sequence identity between OmtA and DmtA was reported
previously (18).

Time-dependent colony fractionation provided a practical method for monitoring
protein accumulation and distribution in firngal tissues of different ages; similar
information could not be obtained in liquid shake culture (batch fermentation).
Knowledge of OmtA distribution was essential to identify fungal tissues that were rich in
the target protein and allowed successful immuno-histochemistry in this study and
immuno-electron microscopy in a study now underway. Since these protocols require
very small quantities of fungal tissue (1mm3 for immunoelectron microscopy for
example) one could easily miss the relevant protein if it was not uniformly distributed in
a colony.

Using this colony fractionation protocol, OmtA PAb, and Western blot analysis,
we observed OmtA in all fractions of a 72 h old colony (fractions S1, S2, and S3) grown
on YES agar. The youngest colony fraction (S3; 0 to 24 h) contained mostly full-length
OmtA, while fraction S1 (48 to 72 h) contained less full-length OmtA and more OmtA-
derived peptides. In fraction C1 (CSIO, 48 to 72 h) and the oldest fraction of SU-I (48 to
72 h) grown on PDA medium, OmtA was almost undetectable. Similarly, OmtA was not
observed in the oldest fraction of a 90 h old colony grown on YES medium. No OmtA
protein was detected in 24 h old colonies grown on YES; OmtA was detected in 48 h old
colonies on the same medium. Together the data suggest that OmtA biosynthesis in A.

parasiticus SUI grown on solid YES media initiates after 24 h. OmtA then accumulates

75

to relatively high levels in the youngest fraction, for example, in C3 and S3 (0 to 24 h) of
a 72 h old colony. As cells in that fraction age (at 48 or 72 h), OmtA synthesis appears to
decline, the protein is proteolytically cleaved, and disappears completely by 90 h. The
rate of OmtA proteolysis appears to be age- and grth medium-dependent. OmtA
proteolysis was not observed to the same extent in cells grown in liquid culture.

OmtA proteolysis may occur after “accidental” contact with fungal proteases
released during mechanical disruption of fungal tissues. However, because the tissue is
quickly frozen before disruption and because protease inhibitors are added to the
extraction buffer, we hypothesize that OmtA proteolysis occurs as part of a natural
process during fungal grth and development. To generate developmental structures in
solid culture, fungi may require proteases to digest unneeded structures or metabolic
proteins (20). For example, a mutation in one protease, subtilisin-related serine proteinase
(ALP2), resulted in smaller conidiophore vesicles (50% reduction) and a lower number of
conidia (80% reduction) in A. nidulans (20). We observed that SU-l fungal colonies on
PDA, in which colonies grew more slowly than on YES (smaller diameter), also had
more severe OmtA proteolysis. Conidiophore structures also could be found in fraction
C3 on YES whereas they were absent in S3 on YES.

Septal pores form “channels” between adjacent cells in the mycelium and appear
to play an important role in cell-to-cell communication in filamentous fungi (3).
Cytoplasm, mitochondria and nuclei migrate throughout the mycelium via septal pores
(3,16). Because we do not see accumulation of OmtA in 24 h old colonies on YES but do
see accumulation of OmtA in cells in the youngest fraction S3 (0 to 24 h) of a 72-h old
fungal colony, we hypothesize that migration of OmtA together with cytoplasm and

organelles occurs from older cells (fraction S2, 24 to 48 h) to younger cells (fraction S3, 0

76

to 24 h) on solid media. Alternatively, it is also possible that specific regulatory factor(s)
involved in aflatoxin biosynthesis (eg. AflR) move fiom the aflatoxin producing cells in
fraction S2 to fraction S3 inducing omtA expression in these younger tissues.

OmtA-specific PAb were also used in immuno-fluorescence microscopy. The
data provided strong evidence that our purification strategy resulted in PAb that were
sensitive and specific. Although the paraffin-embeddment procedure and sectioning
technique have been used for microscopic analysis of a variety of organisms (4), this is
the first reported use in the study of a fungus grown on solid media and the first
application in a study of Aspergillus parasiticus. We previously utilized a fungal cell
preparation technique for immuno-labeling that required digestion of the cell wall using
Novozyme (8). However, in these early localization studies, variation in cell wall
digestion resulted in significant artifacts. The paraffin-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, our data suggest that OmtA is not produced exclusively in conidiophores as
hypothesized. On the contrary, CLSM micrographs showed that OmtA is distributed
throughout the colony and in both conidiophores and vegetative hyphae. Western blot
analysis indicated that abundant full-length OmtA was observed in fraction S3, even
though conidiophores were nearly absent in this area. In fraction 82, the highest intensity

of OmtA was detected under CLSM, and OmtA accumulated to similar levels in

77

vegetative hyphae and conidiophores.

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 altemative 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 paraffin embedded sections to identify specific organelles; for
example anti-SKL antibodies (10) 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. The anti-SKL
labeled organelles were recently demonstrated to be Woronin bodies by immuno-electron
microscopy (data not shown).

There are some indirect data available that suggest that at least some aflatoxin
proteins are localized in organelles. For example, in A. parasiticus, the aflatoxin enzyme
OrdA was found to be membrane associated during protein purification (6). Based on
amino acid sequence, another aflatoxin protein, AflJ, was predicted to contain a C-
terminal microbody targeting signal, NRY, and three membrane-spanning regions ( 1 7).
By comparing the protein sequence of purified OmtA with the predicted amino acid
sequence derived from the OmtA cDNA, this protein was proposed to contain a leader
sequence that apparently is processed to generate a 42-kDa protein (25). This leader
sequence may be required for this enzyme to interact with membranes (25). Based on

computer-assisted analysis (PSORT; prediction of protein localization sites Version 6.4:

78

http://psort.nibb.ac.jp/psort), we also speculate that putative peroxisomal targeting signals
are present in aflatoxin proteins including Nor-1, AvnA, OmtA and OrdA (unpublished
data). In contrast, OmtA (6) and Nor-1 (26) were observed in the post-microsomal
cytoplasmic fraction in cell fractionation studies. Therefore, the distribution of aflatoxin
enzymes within a fungal cell still is not clearly understood.

Localization to specific organelles has been demonstrated for enzymes involved in
secondary metabolism. Penicillin is a secondary metabolite produced by several
filamentous fungi including A. nidulans. The enzyme, 6-aminopenicillanic acid
acyltransferase, which catalyzes the final step of penicillin biosynthesis in Penicillium
was localized to a membrane bound organelle, the microbody (19); the authors suggested
that penicillin is synthesized in this organelle. In a separate study, prehelminthosporol, a
phytotoxin produced by Bipalaris sarokinianawas, was localized in the Woronin body (1)
which is thought to be derived from the peroxisome (24). Prehelminthosporol has been
shown to disrupt plant plasma membranes and inhibit growth of gram-positive bacteria.
Detailed sub-cellular localization of OmtA and several other aflatoxin enzymes using
immunoelectron microscopy should help clarify the cellular site of aflatoxin synthesis in

A. parasiticus.

79

ACKNOWLEDGEMENTS

I. This work was supported by the Michigan Agricultural Experiment Station (MSU), the
National Food Safety and Toxicology Center (MSU), the Center for Environmental
Toxicology (MSU), and the National Institutes of Health (ROI CA52003-11).

2. Data in chapter 2 and 3 were published in Applied and Environmental Microbiology
entitled “Function of native OmtA in viva and expression/distribution of this protein
in colonies of Aspergillus Parasiticus”.

3. The LSM imagines in Figure 5 and 6 were generated by Ching-Hsun Chiou.

80

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83

CHAPTER 4

Sub-cellular localization of aflatoxin biosynthetic enzymes in time-dependent

fractionated colonies of Aspergillus parasiticus

INTRODUCTION

Aflatoxins are highly toxic and carcinogenic secondary metabolites produced by
several filamentous fungi including Aspergillus parasiticus, A. flavus, A. namius, and A.
tamarii (12,23). Aflatoxins pose significant health and economic problems in the US and
many other locations throughout the world because they frequently contaminate food and
feed crops including com, peanuts, tree nuts and cottonseed (9,12).

The biosynthesis of aflatoxin is a complex process that involves at least 17
enzyme activities (23). The distribution of these enzymes within fungal colonies and
location within fungal cells remain unknown. The objective of this study was to
investigate the distribution and sub-cellular location of representative enzymatic activities
in the aflatoxin biosynthetic pathway. Toward this end, I focused attention on Nor-1
(29), Ver-l (19) and OmtA (33) that catalyze early, middle and late enzymatic steps in the
aflatoxin biosynthetic pathway, respectively. Nor-1 and Ver-I are NADPH-dependent
keto-reductases involved in the conversion of norsolorinic acid (NA) to averantin (AVN)
and versicolorin A (VERA) to demethylsterigmatocystin (DMST), respectively; OmtA is
a methyltransferase that converts sterigmatocystin (ST) to O-methylsterigmatocystin

(OMST).

84

Specific secondary metabolites such as penicillin and preheliminthosporol are
known to enhance the survival of the producing organism in the growth environment
(5,1). However, the biological function of most secondary metabolites, including
aflatoxin, is not clear. Characterization of the sub-cellular location of aflatoxin and/or its
biosynthetic enzymes should provide important clues about the biological function and
mechanisms that mediate intracellular transport, storage, and secretion of this toxic
secondary metabolite by the producing firngus.

As a potential mechanism to protect fungal cells from the potential deleterious
effects of aflatoxin accumulation in the mycelium, I hypothesized that aflatoxin enzymes
are compartmentalized in a sub-cellular organelle(s). Further, I hypothesized that the
end-product, aflatoxin, is synthesized within this organelle and then secreted directly into
the extracellular environment. To address these hypotheses, I analyzed enzyme
distribution in fungal colonies by Western blot analysis and immuno-fluorescence
microscopy, and canied out in situ localization of aflatoxin enzymes within fungal cells
using immuno-microscopic methods following cryofixation. For immuno-transmission
electron microscopy (TEM), it had been shown previously that a sample preparation
procedure that combines rapid freezing and freeze substitution is superior to chemical
fixation in terms of ultrastructure and antigenicity (1). Therefore, I used a modification of
published methods (32) to localize Nor-1, Ver-l and OmtA in time-fractionated colonies.
Nor-1 and Ver—l enzymes were primarily localized to the cytoplasm in cells 24 to 48 h
old (fraction 2) suggesting that they are cytosolic enzymes. OmtA was also detected in
the cytoplasm. However, in cells located near the basal (substrate) surface of the colony,

large quantities of OmtA were detected in organelles identified as vacuoles. Based on

85

current data, we hypothesize that the relative distribution of OmtA to cytoplasm or

vacuoles depends on the age and/or physiological condition of the fungal cells.

MATERIALS AND METHODS

Fungal strains and time-dependent fractionation of fungal colonies. A.
parasiticus SUI (NRRLS 862, ATCC 56775) is a wild-type, aflatoxin-producing strain.
AF S10 (aflR via gene disruption) is a non-aflatoxin producing strain derived from A.
parasiticus NR-l (niaD) that in turn was derived from SUI (7). Asexual conidiospores (5
x105) of A. parasiticus SUI and AF S10 were inoculated onto the center of YES agar
medium (2% yeast extract, 6% sucrose, pH 5.8) overlaid with sterile cellophane
membranes and incubated at 29° C in the dark. 72 h-old colonies of SUI (S) and AF S10
(R) were fractionated into three concentric rings based on area covered at 24, 48, or 72 h
of growth (fraction 1, 48-72 h old; fraction 2, 24-48 h, and fraction 3, 0-24 h) to generate
fractions S1, 82, S3 and R1, R2, R3 respectively as described by Lee et al. (17). The
harvested mycelia from appropriate sections of the colony were used in sample

preparation for Western blot analysis and for fluorescence and electron microscopy.

Western blot analysis of proteins isolated from colony fractions. Sample
preparation and Western blot analysis of A. parasiticus proteins isolated from colony
fractions were conducted using methods described in Lee et al. (1 7). Each lane on the
12% SDS-PAGE gel contained 60pg protein. Nor-1, Ver-I and OmtA polyclonal

antibodies were raised against maltose binding protein fusions generated for each

86

aflatoxin protein (35,18,19,l7). Anti-Nor-l serum (rabbit No. 126) was generated in this

study and used at a 1 to 5000 dilution. Anti-Ver-l IgG was used at 2 pg per ml.

Immuno-fluorescence labeling and confocal laser scanning microscopy
(CLSM). Preparation of paraffin-embedded fungal sections, immuno-labeling, and
microscopic analysis via CLSM utilized methods described by Lee et al. (17). The
samples were probed with primary antibodies against Nor-1 (1 to 500 dilution), Ver-l (20
pg/ml) (19) or OmtA (20 pg/ml) (17), 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). Fluorescence image detection was performed on a Meridian INSIGHT plus
laser-scanning microscope (Meridian Instruments, Inc., Okemos, MI) with a 488 nm laser
line. The Alexa 488 green fluorescence was detected using a BP 530/30 banier filter. All
images were captured using a 40X Zeiss Plan-NEOFLUAR oil objective lens (N .A.=1 .3)
(Carl Zeiss Inc., Germany) with a 1X CCD cool-charged detector allowing analysis of a
large number of cells in one 512 x 480 image. Fluorescence image analysis of strains SUI
and AF S10 was conducted using the same instrument parameter settings. For all AF 810
samples, bright field images were also generated to demonstrate the size and number of

cells analyzed.

Quantitative fluorescence intensity analysis. Alexa 488 fluorescence was
quantified using IQ Master Program (V2.31) image analysis software that accompanied
the Meridian Insight laser-scanning microscope (Meridian Instruments, Inc., Okemos,

MI). The CLSM images were acquired under the identical instrument settings for samples

87

analyzed in the same immuno-labeling experiments. For each colony fraction, twenty

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

Sample preparation for immuno—electron microscopy. Three to five mm
diameter circles of firngal tissue were obtained from fraction 2 of colonies of SUI and
AF S10 grown on YES agar plates. These circles were cut from the colony using sterile
200 pl pipette tips with the end removed with a sterile razor blade to generate a cutting
edge with the correct diameter. Samples were immediately cryofixed at ——190° C in a
commercial Jet-Freezer (RMC MF 7200) and then transferred to a tube containing acetone
that had been frozen by immersion in liquid nitrogen. The following substitution
procedure was modified from a published method (32). Samples in frozen acetone were
then stored at -80° C for 6 (1. Samples were washed twice with fresh, ice-cold acetone (-
20° C) and then immersed in flesh, ice-cold acetone (-20° C) containing 0.2%
glutaradehyde and stored at -20° C for 24 h. After washing with fresh,“ ice-cold acetone (-
20° C) three times, acetone in samples was replaced with 100 % ethanol by a graded
series of ethanol/acetone (35%, 50%, 75%, 90% [30 min each] and 100 % [two times, 30
min each]). Samples were then infiltrated with a graded series of LR White resin (Ted
Pella, Inc., Redding, CA) /ethanol: 5, 10, 20, 40, 60% (3 h each), 90% for 24h and 100%
(3 changes) for 2 d at -20° C. Polymerization was carried out under UV light (366 nm) at

4° C for 48 h.

Immuno-gold labeling and electron microscopy. Fungal sections for immuno-
gold labeling were cut to 90 ~ 100 nm thickness using an MTX ultrarnicrotome (RMC,

Tucson, Arizona). Sections were collected onto formvar-coated grids. Each grid

88

contained approximately 10-20 thin sections. For each labeling experiment, two grids
each from SUI and AF SIO tissues were used. For each antibody, at least three
independent labeling experiments were performed. First, grids with sections were
incubated with blocking solution (1% BSA plus 0.1% saponin in TBS) at 4° C overnight.
Primary antibody treatment (purified OmtA IgG, 50 pg per ml; anti-Verl IgG, 140 pg per
ml; anti-serum: anti-SKL serum, 1: 200; anti-Nor-I, 1: 500) was performed at room
temperature for 4 h. Secondary antibody treatment (goat anti-rabbit conjugated with 10
nm gold; 1 to 30-fold dilution) (Ted Pella, Inc., Redding, CA) was performed at room
temperature for 2 h. After each antibody incubation, grids were washed once with Tris-
buffered saline (pH 7 .5) containing 1% BSA and 0.1% saponin for 5 min followed by

five washes with TBS for 25 min total. Finally, grids were washed with dd H20 for 30 s

and post stained with 3% uranyl acetate for 20 min. Sections were observed using a JEOL

100CX II transmission electron microscope (Tokyo, Japan) at 100 kV.

RESULTS

Western blot analysis of Nor-land Ver-l isolated from colony fractions 1, 2, and
3 of A. parasiticus SUI and AF SlO demonstrated the specificity of the antibody
preparations used in microscopic analysis (Figure 4.1). Similar analysis was conducted
previously on OmtA only, in which increasing quantities of proteolytically cleaved
enzyme were detected in fractions 2 and 1 (17); the highest proportion of full-length
protein was observed in fraction 2. In contrast, Nor-1 and Ver-I in the current study
appeared predominantly in full-length form in all colony fractions. The proteolytically

cleaved forms of these two enzymes were not observed in cells grown on solid YES

89

SI 52 83 R1 R2 R3 MWIkDa)

1‘. mg

8 $3388

V'cr-I
zs-o -

I
l

B

81 82 SJ R1 R2 R3 MW(kDa)

E120

100

80

W 60

fl 5"

Nor-l ‘ n 40

31-0“ “mu—.m- .- a 30

- ’ 20

Figure 4.1. Western blot analysis of fungal protein extracts using Ver-I and Nor-1
polyclonal antibodies. Proteins were extracted from time-fractionated colonies of SUI
and AF S10 grown on YES agar medium and were subjected to Western blot analysis
with Ver-l (Panel A) and Nor-1 (Panel B) polyclonal antibodies. 72 h-old colonies were
fractionated into three concentric rings based on area covered at three time points: for S1
and R1, 48 to 72 h; 82 and R2, 24 to 48 h; S3 and R3, 0 to 24 h. The lanes to the left and
right contain molecular mass standards. The molecular mass of standards are marked at
the right of Panels A and B. The molecular masses of Ver-l and Nor-1 (indicated by

arrows at the left) are 28 kDa and 31 kDa, respectively.

90

medium. However, like OmtA, the highest quantity of Nor-l and Ver-l was observed in
fraction 2 of SUI.

Immuno-fluorescence microscopy via CLSM confirmed the specificity of the
Nor-1 and Ver-I polyclonal antibodies for microscopic analysis. The specificity of the
OmtA polyclonal antibody was confirmed in a previous study (17). Nor-1 and Ver-I were
detected in all colony fractions of SUI (Figure 4.23 for Nor-I and Figure 4.2d for Ver-l).
Under the same instrument settings, very little signal was detected in AF 810 (Figure 4.2b
for Nor-1 and Figure 4.2e for Ver-l). Therefore, bright field images of the AF S10 colony
fractions are shown to illustrate the typical size and numbers of cells analyzed (Figure
4.2e and 20. Quantitative analysis of the fluorescence intensity in the CLSM digital
images using IQ Master Program (V2.31) image analysis software confirmed that the
highest quantity of these three aflatoxin enzymes occurred in fraction 2 (Table 4.1).
Therefore, we focused our immuno-TEM analysis on the proteins in this colony fraction.

Using our sample preparation, labeling, and imaging procedures, very few gold
particles were observed on control sections of wild-type strain SUI treated with
secondary antibodies only (Figure 4.3A and B). Therefore, we interpreted an intense
signal (black dots representing 10 nm gold particles) on sections treated with both
primary and secondary antibodies to result from the specific interaction between primary
antibodies and their target proteins immobilized within the cell. However, we
occasionally detected labeling of the cell wall, especially for anti-Ver-l, in certain
sections of both SUI and AF S10 (no aflatoxin proteins produced), leading us to conclude
that this was non- specific labeling (data not shown). It was reported previously that non-
specific labeling may result either from non-specific binding of primary antibodies to the

cell wall or from cell wall-reactive antibodies that arise in rabbits exposed to or

91

Figure 4.2. Immuno-fluorescence confocal microscopy of Nor-l and Ver-l in A.
parasiticus SUI and AF SIO (aflR knockout mutant) grown on YES agar for 72 h.
Colonies of SUI and AF S10 were divided into 3 fractions (see Material and Methods).
The paraffin embedded fungal sections were immuno-labeled with anti-Nor-I 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 SUI sections immuno-
labeled with anti-Nor-l and anti-Ver-l are shown in Rows a and d, respectively; and of
AF SIO sections in Rows b and e. The related bright field images of the AF SIO colony

fractions are shown in Rows c and f (legends labeled in black). Bar, 100 pm.

92

 

A. Nor-l

B. Ver-l 5'

d

 

Figure 4.2.

93

Table 4.1. Quantative fluorescence intensity analysis of A. parasiticus SUI and AF S10

immuno-fluorescence labeled with Nor-1, Ver-I and OmtA antibodies.

 

(Fungal colony) (SU— 1) (AF S I 0)

Protein / Colony fraction S1 82 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.

94

Figure 4.3. Immuno-gold labeling of A. parasiticus SUI and AF 810. In Panels A and B,
ultra-thin sections of fungal tissues of the aflatoxin producing strain SUI were prepared
for TEM and labeled with secondary antibodies only. Ultra-thin sections of fungal tissues
of AF SIO (Panel C and E) and SUI (Panel D and F) were labeled with primary antibodies
against Nor-1 (Panel C and D) and Ver-l (Panels E and F), followed by 10 nm gold beads
conjugated to goat anti-rabbit IgG secondary antibodies (1 to 30 dilution) as described in
Methods. Bars represent 500 nm in A and 250 nm in B, C, D, E and F. Abbreviations:

cw, cell wall; mb, microbodies; M, mitochondria; N, nuclei; V, vacuole.

95

.111, ‘L'l'il‘l‘ip‘

“'iifl" . 4"”ijin . l =~' .. . -.

-i muwmlhulm‘m .3- mm lllillilnmi.‘ I!
' .1. , ll‘ ., , , . .

1 WW I
. . “if

j 1‘,” .151 “3‘1““ .‘1 .11 _'i‘
11,1 ,

 

Figure 4.3.

infected by yeast or molds before or during antibody production (3, 4).

Widely scattered gold particles were also found occasionally associated with the
nucleus and mitochondria in both AF S10 and SUI labeled with Nor-I and Ver-l
antibodies, suggesting that this was non-specific labeling or resulted from cross-reactive
proteins located in these organelles. This background “noise” could be largely eliminated
by a further step of antibody purification using affinity subtraction as described in Lee et
al. (17). As shown in this study, very little labeling was noted in the cell wall,
mitochondria, and nucleus using affinity-purified antibodies to OmtA (Figure 4.4 and
Table. 4.1).

When highly specific antibodies to Nor-1, Ver-l, and OmtA were used to label
sections from fraction 2 of SUI , high signal intensity was observed in the cytoplasm in
many but not all cells (Figure 4.3D, F and 4.4C, D). The gold particles frequently
aggregated to form clusters (Figure 4.3D and 4.4D); similar clusters were not seen in the
cytoplasm of control sections obtained from AF $10 or when only gold-labeled secondary
antibodies were used. The immuno-gold labeling results are in good agreement with
immuno-fluorescence data (Figure 4.2) in this and our previous study and may help
explain the “patchy” appearance of the fluorescent signal observed (17). Of particular
interest, OmtA antibodies specifically and intensely labeled small and large organelles
primarily in cells located near the basal surface (substrate surface) of the colony in
fraction 2 of SUI (Figure 4.4E and 4.5). Based on ultrastructure /morphology, these
organelles have been identified as vacuoles (2). The labeling intensity suggested a high
concentration of OmtA in these organelles. In contrast, antibodies against Nor-l or Ver-l

did not label these organelles in sections from the same location (basal portion of fraction

97

Figure 4.4. Immuno-gold labeling of OmtA in Aspergillus parasiticus AF 810 and SUI.
Ultra thin sections of fungal tissues of AF SIO (Panels A and B) and SUI (Panels C and
D) were labeled with primary antibodies against OmtA followed by gold-labeled
secondary antibodies as described in Methods. Panel B is a schematic representation of
the colony fractionation procedure and shows the cell distribution in a typical thin
section. The specific location of OmtA in cells near the basal surface of the colony
(shaded gray) is also shown in Panel E. Bars present 500 nm in panels A, C and E, and
250 nm in panels B and D. Abbreviations: cw, cell wall; M, mitochondria; N, nuclei; V,

vacuole.

98

 

Figure 4.4.

99

Figure 4.5. Immuno-gold labeling of OmtA in Aspergillus parasiticus SUI.

Sections cut from the basal surface of the colony were labeled with primary antibodies
against OmtA. The gold-labeled vacuoles inside of cells were numbered (1 to 8) and
higher magnification images of these organelles are shown in Panels B, C, D and E. Bars

present 2 pm in panels A and 1 pm in B, C, D and E. Abbreviations: cw, cell wall; M,

mitochondria; N, nuclei; V, vacuole.

100

\ .
04' rt
1, mini.
h '1

‘il

F'I

Its

‘,,,li11 . . " ,
.11. 31111141 ' .

 

Figure 4.5.

101

2) (data not shown). Cells located above the basal region contained smaller organelles
which were sporadically labeled by OmtA and not at all by antibodies to Nor-l and Ver-
1. Similar organelles in AF S10 did not label with polyclonal antibodies to any aflatoxin
enzymes.

The gross ultrastructure of cells observed in thin sections suggested that our
fixation, sectioning and labeling procedures successfully preserved the integrity of
organelles bounded by double (nuclei and mitochondria) and single (vacuoles,
microbodies, Woronin bodies) membranes. In order to confirm that fragile organelles
bounded by single membranes could retain both ultrastructure and the antigenicity of
internal proteins, we labeled some sections from fraction 2 of SUI and AF S10 with
antibodies against peroxisomal targeting signal 1 (PTS-l; anti SKL). Anti-SKL can label
proteins exported to peroxisomes (microbodies; mb) and Woronin bodies (derived from
microbodies; 30). Microbodies and Woronin bodies were specifically and intensely
labeled with anti-SKL (Figure 4.6A, B and C). However, not all microbodies within one
cell were labeled to the same signal intensity possibly because the specific protein
concentration or protein composition varies between microbodies in the same cell. It has
been demonstrated previously that certain proteins lacking PTS-1 can be localized in
peroxisomes (21,27); these proteins should not be recognized by the anti-SKL
preparation. In our studies, labeling of microbodies by Nor-land OmtA antibodies was
rarely observed. However, we did observe that Woronin bodies but not other microbodies
were labeled with anti-Ver-l (Figure 4.6D and E). Woronin bodies in AF S10 were also
labeled to a similar intensity, suggesting that this is non-specific labeling possibly due to
a high antibody concentration used or that “contaminating” antibodies bound to cross-

reactive proteins located in this organelle (Figure 4.6F and G).

102

Figure 4.6. Immuno-gold labeling using anti-SKL and anti-Ver-I in A. parasiticus
AF S10 and SUI. Ultra-thin sections of firngal tissues obtained from fraction 2 were
labeled with primary antibodies to PTS-1 (anti-SKL) (Panels A, B, and C) or primary
antibodies to Ver-l (Panels, D, E, F and G) followed by gold-labeled secondary
antibodies as described in Methods. Panels A, B, D, F, and G, represent ultra-thin
sections prepared from fungal tissues of AF 810 and Panels C and E represent ultra-thin
sections prepared from fungal tissues of SUI. Bars present 250 nm. Abbreviations: cw,

cell wall; mb, microbodies; wb, woronin body; st, septum; M, mitochondria; V, vacuole.

103

' i ll
1“ Illlllill
. .nll '.

\I 1‘ ‘
.ul'lil
.1,,IIIII

 

I III. “if.“ A
II'IIIIIIIII 1"
l "I

1" I‘Maimlr‘riliriilmnllllmi

‘i‘r'rllllilli l'."!'"‘"“

   

Figure 4.6.

104

DISCUSSION

My goal was to analyze the distribution and sub-cellular localization of the
aflatoxin enzymes Nor-1, Ver-I, and OmtA in Aspergillus parasiticus grown on YES
agar growth medium. Western blot analysis of protein extracts from time-fractionated
colonies demonstrated that fraction 2 (cells 24 to 48 h old) contained the highest

concentration of each target protein. In addition, proteolytic cleavage of OmtA occurred

 

to a greater extent in “older” fungal tissues (fractions 1 and 2) but this was not observed
for Nor-1 and Ver-l under these culture conditions. The specificity and timing of
expression of the proteolytic enzymes and the potential role of cleavage in activation/
inactivation of aflatoxin enzymes is clearly of interest for follow up studies.

Immuno-fluorescence microscopy confirmed that Nor-I, Ver-l and OmtA are
present at highest concentration in fraction 2; the “patchy” appearance of the fluorescent
signal suggested that the proteins aggregate in the cytoplasm or are compartmentalized
consistent with our previous studies on Nor-1, Ver-l, and OmtA (17, 18, 35).

At even higher resolution, TEM analyses confirmed that Nor-1, Ver-I and OmtA
are present in the cytoplasm of many, but not all cells in colony fraction 2. In addition,
most cells located near the basal surface of fraction 2 were closely packed and contained
one to several large organelles heavily labeled with gold-labeled antibodies to OmtA. In
some cells, the large and small organelles appeared to fuse together consistent with a
model for vacuolar development in Aspergillus reported previously (22). To our
knowledge, this is the first report of aflatoxin enzymes localizing to a specific cellular

organelle.

105

The morphology and apparent developmental origin of the labeled organelles in
fraction 2 prompted us to identify them as vacuoles. Vacuoles have been associated with
several biological functions in fungi including degradation or recycling of proteins and
whole organelles (2), storage of metabolites, ions and amino acids, enzyme maturation
(e.g. aminopeptidase I) and pH homeostasis (15,28). Vacuoles are also the site for lipid
degradation and glycerol production at specific stages of appressoria formation in
Magnaparthe grisea (31). In yeast, the biological sulfonium compound, S-adenosyl-
methionine (AdoMet), accumulates to high levels in vacuoles and cytoplasm (25,26,13).
S-adenosyl-methionine is an important cofactor that provides the methyl group in the
reaction catalyzed by OmtA. Based on the available data, it is difficult to determine if the
vacuolar localization of OmtA occurs for protein recycling, for enzyme activation, or for
some other purpose in A. parasiticus.

Although the data appeared quite convincing, it was possible that disruption of
organelles occurred during sample preparation and was in part or fully responsible for the
observed cytoplasmic location of Nor-1 , Ver-l , and OmtA. To minimize this possibility,
we demonstrated that the ultra-structure of Woronin bodies and microbodies (bounded by
single membranes) and the antigenicity of their content proteins were preserved using our
methods for sample preparation and microscopic analysis. These structures were
intensely labeled by anti-SKL but not by antibodies to Nor-1 or OmtA. Some labeling of
Woronin bodies did occur with anti-Ver-l, however, the observed labeling occurred in
both AF 810 (control) and SUI strains allowing us to interpret this as non-specific
labeling. This interpretation appears to be likely because disruption of aflR in A.

parasiticus (such as in AF S-IO) results in loss of nor-1, ver-I and omtA gene transcripts

106

 

(6,7) and proteins (this study). Therefore, it is unlikely that Ver-l exists at any location in
AF S I 0.

Similarly, the ultrastructure of double-membrane bound organelles (nuclei and
mitochondria) as well as other single-membrane bound organelles (vacuoles) were also
maintained in this study. With the exception of vacuoles in the basal region of fraction 2
of SUI , these organelles did not label with antibodies against the aflatoxin enzymes.
These results suggest that Nor-1, Ver-I, and OmtA do not localize to nuclei,
mitochondria, Woronin bodies, or microbodies. Because the absence of signal in these
organelles does not appear to be an artifact arising from organelle breakage, these data
strongly support the specificity of labeling of the vacuoles observed in the basal region of
fraction 2.

Two observations from this study are particularly noteworthy. First, intensely
labeled, weakly labeled and unlabeled cells could be found adjacent to each other in the
same thin section analyzed by TEM. It is known that cell components located under the
surface of a section are unable to be labeled. However, the cytoplasm proteins such as
Ver-l and Nor-1 in these unlabeled cells were remainly unlabeled even in serial sections.
These data suggest that the extent and timing of aflatoxin gene expression varies from
cell to cell, even in an area of the colony that is presumably the same age. We propose
that this variation may be related to the local concentration of available nutrients or the
relative age of fungal cells (5,10,20). Second, OmtA appeared in the cytoplasm in certain
cells and in the vacuole in other cells in the same colony fraction. In Saccharomyces,
cytoplasmic proteins can be transported into vacuoles by at least two alternative
pathways: 1) macroautophagy; and 2) the cytoplasm to vacuole targeting (cvt) pathway.

The cvt pathway is regulated by the availability or limitation of specific nutrients (e. g.

107

 

glucose or ethanol) (15,16,28); to date, no pathway similar to the yeast cvt has been
reported in filamentous fungi. However, we propose that transport of OmtA is similar to
the cvt pathway in that a specific set of enviromnental changes (cell age, physiological
conditions or nutrient availability) may trigger the transport of OmtA into the vacuole.
Based on data from this and previous studies, it is reasonable to assign two
potential alternative roles for the vacuole in aflatoxin synthesis: 1) as a means to reduce
or limit aflatoxin synthesis, OmtA is transported to and inactivated in vacuoles via
proteolytic cleavage; 2) as a means to shield the cell from the potential toxic effects of
aflatoxin in the mycelium, OmtA is transported to the vacuole together with the late )
aflatoxin pathway intermediate ST. In the vacuole, OmtA is activated and ST is
converted to OMST and further converted to AP B, by OrdA (24,34) (catalyzes the final
step of aflatoxin biosynthesis) localized in or on the same vacuole. These hypotheses

form the basis for our future studies on aflatoxin synthesis.

108

ACKNOWLEDGEMENTS

Chapter 4 was submitted for publication in Applied and Environmental
Microbiology (2003). Figure 4.2 and Table 4.1 were generated by Ching-Hsun Chiou. We
also thank Dr. Shirley Owens for help in quantification of fluorescence intensity and Dr.
Xudong Fan in operating the jet-freezer in Center of Advanced Microscopy at Michigan

State University.

109

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19. Liang, S.-H., T.-S. Wu, R. Lee, F. S. Chu and J. E. Linz. 1997. Analysis of
mechanism regulating expression of the ver-I gene, involved in aflatoxin biosynthesis.
Appl. Environ. Microbiol. 63:1058-1065.

20. Luchese, R.H., and W. F. Harrigan. I993. Biosynthesis of aflatoxin- the role of
nutritional factors. J. Applied Bacteriology. 74: 5-14.

21. Maeting, 1., G. Schmidt, H. Sahm, J. L. Revuelta, Y.-D. Stierhof, and K.-P.
Stahmann. 1999. Isocitrate lyase of Ashbya gossypii- transscriptional regulation and
peroxisomal localization. FEBS letters. 444: 15-21.

22. 0hsumi, K., M. Arioka, H. Nakajima, and K. Kitamoto. 2002. Cloning and
characterization of a gene (avaA) from Aspergillus nidulans encoding a small GTPase
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23. Payne, G. A. and M. P. Brown. 1998. Genetics and physiology of aflatoxin
biosynthesis. Annu. Rev. Phytopathol. 36:329-362.

24. Prieto, R., and C. P. Woloshuk. 1997. ardl, an oxidoreductase gene responsible for

the conversion of O-methylsterigmatocystin to aflatoxin in Aspergillusflavus. Appl.
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25. Schwencke, J., and H. De Robichon-Szulmajster. 1976. The transport of S-
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targeting of isocitrate lyases in Saccharamyces cerevisiae. Biochem. J. 319:255-262.

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29. Trail, F., P-K. Chang, J. Cary and J. E. Linz. 1994. Structural and functional
analysis of the nar-l gene involved in the biosynthesis of aflatoxin by Aspergillus
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30. Valenciano, S., J.R. De Lucas, 1. Van der Klei, M. Veenhuis, and F. Laborda.
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32. Xu, H and K. Mendgen. 1994. Endocytosis of 1,3-B-glucans by broad bean cells at
the penetration site of the cowpea rust fungus (haploid stage). Planta. 195:282—290.

33. Yu, J., J. W. Cary, D. Bhatnagar, T. E. Cleveland, N. P. Keller, F. S. Chu. 1993.
Cloning and characterization of a cDNA from Aspergillus parasiticus encoding an O-
methyltransferase involved in aflatoxin biosynthesis. Appl. Environ. Microbiol. 59:3564-
71.

34. Yu, J., P.-K. Chang, K. C. Ehrlich, J. W. Cary, D. Bhatnagar, and T. E.
Cleveland. 1998. Characterization of the critical amino acids of Aspergillus parasiticus
cytochrome P-450 monooxygenease encoded by ordA that is involved in the biosynthesis
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35. Zhou, R. 1997. The function, accumulation, and localization of the Nor-1 protein
involved in aflatoxin biosynthesis; the function of the fluP gene associated with
sporulation in Aspergillus parasiticus. PhD Dissertation, Michigan State University, East
Lansing.

112

CHAPTER 5

Detection of aflatoxin B, in fungal spores and colonies

INTRODUCTION

The biological function of aflatoxin in the producing fungus is still not clear.
Since aflatoxins are secondary metabolites and synthesized primarily during stationary
phase, they are not necessary for primary growth of the cell. It has been shown previously
that the production of sterigmatocystein (ST) in A. nidulus and aflatoxin in A. parasiticus
is closely associated with asexual sporulation (2), and positively regulated by F le (1).
Although the details of the connection between production of aflatoxin and fungal
sporulation are entirely not clear, the presence of aflatoxins in asexual conidiospores has
been reported (4,6). Since spores are important structures involved in dispersal and
colonization, one of the proposed function of aflatoxins is to provide the fungus with a
survival advantage by overcoming environmental threats (e.g. resistance to UV light).
The aflatoxin concentrations in spores harvested from different media were estimated.
Wicklow and Shotwell (1982) reported that conidiospores harvested from Czapek agar
cultures (grown in the dark at 28° C for 21 days) contained 1,106 to 54,300 ppb (ng/g) of
AFB, in four A. parasiticus strains and 36 to 97,400 ppb in five A. flavus strains.
Palmgren and Lee (1986) reported that conidiospores harvested from A. parasiticus
(SRRC-2004) cultured on autoclaved rice at 25°C for 7 days contained 166,000 ppb of

AFB,. In contrast to aflatoxin distribution in wild-type strains, the distribution of the

113

 

aflatoxin intermediate, Nosolorinic acid (NA), in a nor-l mutant is mainly in vegetative
mycelia.

In the present study, I wanted to compare: 1) the distribution of aflatoxin between
colony and substrate (agar); and 2) the aflatoxin concentration in spores harvested under
wet and dry conditions. Although the aflatoxin gene cluster contains an aflT gene, which
may function as a toxin transporter, no data in the literature have demonstrated that a
specific secretion system including AflT is used to transport aflatoxin into the
environmental media. In this study, I measured the aflatoxin concentration in time
fractionated colonies grown for different total times and compared the aflatoxin
distribution between fungal colonies and the substrate media. My results showed that, in
YES cultures ranging from 2 days to 5 days of age, the ratio of aflatoxin in the fungal
mycelium and total aflatoxin produced decreased with culture time. However, for an
unknown reason, the ratio increased at days 6 and 7. In addition, culture media affected
the distribution of aflatoxin in the mycelium. Compared with 60% in YES culture, only
20% of the total aflatoxin was detected in fungal mycelium cultured on PDA agar. Based
on the current data, the nature of the specific transport system for aflatoxin can not be
determined yet. My data also showed that there is a 10-fold difference in aflatoxin
concentration between conidiospores isolated under wet and dry conditions. The higher
concentration obtained from spores harvested under wet conditions is due to absorption of

aflatoxin from water used for harvest onto the surface of the spores.

114

Mutants or

 

 

 

 

 

 

Treatments
Norsolorinic acid ATCC 24690
Averantin ATCC 56774
Averufanin
Averufin ATCC 24551
1?
Hydroxyversicolorine HVN-I

 

Versiconal Hemiacetal Acetate Dichlorvos

2%:

versicolorin A ATCC 36537

 

 

 

 

 

AFBl «H.— O-methylsterigmatocystin 4—_ Sterigmatocystin
SRRC 2043

 

 

 

 

Figure 5.1. Mutants in the aflatoxin biosynthetic pathway.

115

 

MATERIAL AND METHODS

Fungal strains. A. parasiticus SU1(NRRL5862, ATCC 56775) is a wild-type,
aflatoxin-producing strain. A. parasiticus RHNI (SRRC2043; ordA) accumulates the

aflatoxin intermediate OMST and ATCC36537 (var-I) accumulates VA (Figure 5.1).

Aflatoxin distribution in time-dependent fractionated colonies of SUI grown

on solid medium. To determine the accumulation and distribution of aflatoxin in fungal

colonies grown on solid culture media, conidiospores (2 x 105) of A. parasiticus SUI
were inoculated onto the center of YES or PDA agar overlaid with steriled cellophane
membranes and incubated at 29° C in the dark. Colonies were analyzed after 2, 3, 4, 5, 6
and 7 d of growth. The 2 d-old colonies were harvested directly without fractionation.
Colonies older than 2 d were marked at the edge every 24-h as an indicator for future
cutting. For example, the 3 d-old colony was fractionated into two concentric rings (ring-
1 and 2) based on area covered at the two time points: Ring-l (48 h) contained mycelia
from the colony center out to a distance of 2.5 cm (24 to 72 h) and Ring-2 (0 t024 h)
contained mycelia from 2.5 to 4.2 cm (0 to 24 h) (Figure 5.2 and Table 5.1). The
harvested mycelia from appropriate fractions were weighed and extracted with
chloroform. The chloroform extracts were evaporated and resuspended in methanol. In
the same culture, the substrate (agar) and membrane were also analyzed for aflatoxin

levels. The aflatoxin concentrations were determined by ELISA (5)

Spores harvested by water (wet conditions). 1) In early experiments, the

fungus was grown on a nylon membrane on PDA agar plates at 29° C in the dark for 7 d.

116

 

   

Figure 5.2. Diagram of colony fractionation.

117

Table 5.1. Fractionation of fungal colonies at different ages.

 

 

 

Age\Diamter (cm) 2.5 2.5-4.2 4.2-5.0 5.0-6.5 6.5-8.0
2d Ring-1
3d Ring-I Ring-2
4d Ring-1 Ring-2 Ring-3
5d Ring-I Ring-2 Ring-3 Ring-4
6d Ring-l Ring-2 Ring-3 Ring-4 Ring-5

 

118

 

Spores were released from conidiophores into water (harvest water) by mechanical force.
Colonies with membranes were immersed into the water and the surface of the colony
was scraped with a spatula. The water containing spores was passed through glass wool
to remove any hyphal fragments. Spores were pelleted from the water by centrifugation at
3,000 x RPM for 20 min. Both water and spores were measured for aflatoxin
concentrations. 2) In later experiments, firngal strains, including SUI, RHNI
(SRRC2043) and ATCC36537, were grown on cellophane membranes laid onto PDA
agar plates and incubated at 29°C in the dark for 7 d. For spore isolation, the whole
colony was removed from the cellophane into a 50-ml conical tube and 35 ml of water
was dispensed into the tube. Spores were released into the harvest water by rocking the
tube back and forth for 1 min. The spore-containing water was then filtered through glass
wool. Spores were pelleted by centrifuged at 20,000 x g for 15 min and resuspend in 1 ml
of H20. The resulting spore pellet and the harvest water were used in binding assays as
described in the following section to test their ability to absorb aflatoxin (AFB, and G,)
and aflatoxin intermediates (OMST and VA) contained in the harvest water. The AFB,
and G, or OMST-containing water used in the binding experiment were firrther filtered
through 0.22 pm or 0.45 pm filters to eliminate any contaminating spores. In addition, to
make sure that the aflatoxin and pathway intermediates would not block the filter
membrane, the filtered water and only centrifuge-treated water (without passing through
filter) were examined under a microscope and analyzed by TLC analysis in parallel. No
spores were observed in the filtered water, but a small quantity of spores were observed
in the centrifuge-treated water. In both treatments, AF 8,, AF G, and OMST were all

detected in harvest water by TLC analysis (Figure 5.3).

119

Binding of AFB,, AFG,, OMST, and VA in harvest water by spores. Spores
isolated from the non-aflatoxin producing strain RHNI (accumulates only OMST) were
used to study the binding of AFB, and AFG, in harvest water. Spores isolated from
aflatoxin-producing strain SU-l (accumulates only AFB, and G,) were used to study the
binding of OMST and VA in harvest water. The same numbers of spores (2 x 107) were
used in the binding assay and for extraction of aflatoxin and OMST for comparison.

The RHNI spores were mixed with harvest water (1 ml) containing AFB, and G,
(saved from a SU-l spore preparation) for 10 min and then span down at 20,000 x g for
15 min. The spores were washed once with 1 ml of pure water and span down again
(same conditions). Spores were finally resuspend in 1 ml of water and extracted with 0.5
m1 of chloroform. 10 pl of spore extracts was analyzed in parallel with other extracts,
including the harvest water, spores before binding, and toxin extracted from a whole
colony, by TLC. The same procedure was applied to the binding of OMST and VA from

harvest water by SU-I spores.

Spores harvested by cotton-tipped applicators. A method to harvest spores
under dry conditions was developed. The colony surface was gently swabbed with cotton-
tipped applicators and the spores attached to the applicator were then released into water
by stirring the applicator in the water. A new applicator was used each time to touch the
colony surface. In this dry-harvested method, water was never directly in contact with the
colony during spore harvest. The spore-containing water (total 8 ml) was then filtered
through glass wool. Spores were pelleted by centrifugation as described above. The water
was saved for future aflatoxin measurement. The spore pellet was dried under vacuum

with heat (60 °C) for 4 h. The AFB, concentrations were determined by ELISA.

120

RESULTS

Aflatoxin distribution in time-dependent fractionated colonies of SUI grown on
solid medium. The total aflatoxin produced by a colony increased with time during the
first 4 days, declined at d 5, and then increased at d 6 and 7 (Table 5.2). The total
aflatoxin in the mycelium also reached a peak at d 4 and then decreased at d 5 (Table
5.2). A similar pattern was observed in the agar (Table 5.2). These results suggest that,
for unknown reasons, the detectable aflatoxin decreased between d 4 and 5. The ratio of
aflatoxin in the mycelium to total AF B1 tended to decline with time until (1 5 (Table 5.2)
and then increased at d 6 (40 and 46 %) and d 7 (61%). By monitoring dry weight in the
same fraction isolated from different age of colonies, the results suggest that the firngus is
not only growing outward (two-dimensions) but also growing upward (three dimensions)
with increased time (Table 5.3). As shown by dry weights measured in Ring-2, the
regular rate of growth is 0.12 g per (1 (Table 5.3). Approximately 10 pg of aflatoxin were
detected in the Ring 2 (0-24 h-old) of a 72 h-old colony. Over the next 24 h (24-48 h-old),
aflatoxin in Ring 2 rapidly accumulated (6-fold increase in concentration) and the
accumulation then declined (Table 5.4). The aflatoxin concentrations in each fraction
isolated from two 6-d colonies varied even they were cultured at the same time (Table 5.4
and 5.5). This variation may be explained by the observation that a part of the edge of
colony-No. 2 made contact with the Petri dish. That contact may affect a normal fungal
growth and the normal physiology of the fungal cells. Similarly for 7-d old colony, the
colony margin reached the edge of plate, which appeared to result in an increase in

aflatoxin content.

121

 

Table 5.2. Total aflatoxin detected in a colony, agar and membrane from different

aged colonies.

 

 

 

 

 

 

 

 

Day 0‘ sample Membrane Agar Mycelium Total Mycelium
‘"‘° (Its) (Iug) (118) (118) / Agar /T0ta|
Colony No. C/o)
YES
2 d-l 0.1 8 8 16 100 50
2 0.1 7 10 17 140 59
3 1 10 16 27 160 59
3 d-l 2 63 94 159 150 59
2 6 98 87 191 90 46
4 d-l 2 155 143 300 90 48
2 9 191 133 333 70 40
5 d-l 2 138 77 217 60 35
2 7 153 80 240 60 33
6 d 11 286 218 515 80 42
7 d 14 257 420 691 160 61
PDA
7 d-l 19 301 86 406 30 21
2 25 460 67 552 10 12

 

 

Table 5.3. Dry weight (g) of fractionated mycelium from different aged colonies.

 

 

 

 

 

 

 

 

 

 

Age/Fraction Ring 1 Ring 2 Ring 3 Ring 4 Ring 5 Total
L.__E__8__fl__fi___£_,
YES
2d ,,,,, 0.128 __ 0.128
3 d , 0.200 0.135 0.335
4 d 0.238 0.250 0.216 0.704 W
, ~6de 0.407 0.502 0.438 9548 03272222
-2w0306 0.369 0.420 0.55 1 l .273 2.919
7 d 3.107
PDA
7 d-l 1.263
-2 1.328

 

122

 

 

Table 5.4. Total aflatoxin (pg) in fractionated mycelium from different aged

 

 

 

 

 

 

 

colonies.
Age/Fraction Ring 1 Ring 2 Ring 3 Ring 4 Ring 5
118 pg pg $8 118
2 d 16
‘_ 3 d 1 75 12
5 d . 35 29 13 3
6 d-l 47 60 37 19 4
-2 54 87 57 59 12

 

 

Table 5.5. Normalized aflatoxin levels (pg of aflatoxin/g dry weight mycelium)

in fractionated mycelium from different aged colonies.

 

 

 

 

 

 

 

 

 

 

 

 

Age/Fraction Ring 1 Ring 2 Ring 3 Ring 4 Ring 5 Total/w.t
Its/g rig/g rig/g Its/g Its/g JIB/g
YES
2 d 125 125
3 d 375 .89.. __.____ - 260
4 d 193 296 60 189
5 d 147- 78 ._ 43 14 ..._...70..__.
6d-1 115 120 84 35 12 75
-2 176 236 136 107 .9..- .. 92 _
7 days 135
PDA
7 d-l 68
-2 50

 

 

 

123

Binding of AFB, and pathway intermediates in water by spores. Spores
harvested under wet conditions (by water) contained 10-fold higher AFB, concentration
(1350 rig/mg) than under dry conditions (by cotton-tipped applicator) (average 114
rig/mg) (Table 5.6). Based on TLC analysis, the explanation for the higher concentration
in wet-harvested spores is that they absorb aflatoxin from water used in spore harvest-
this presumably arises from the mycelium. This is supported by the binding assay. In this
experiment, SUI spores were demonstrated to bind OMST from water used to harvest
RHN-1 spores and RHN-I spores bound AFB, and AF G, from water used to harvest SUI
spores. Before the binding assay, TLC analysis showed that only AFB, and G, were
detected in extracts of SUI spores and water used to harvest SU-l spores, and only
OMST was detected in extracts of RHNI spores and water used to harvest RHN-1 spores
(Figure 5.3 and Fig. 5 .4, lane 3 and 4). After binding, AF B,, G,, and OMST were
detected in an extract of RHNI spores (Figure 5.4, lane 5 and 6); and were also detected
in extracts of SUI spores (Figure 5.6, lane 4). The AFB, and AF G, concentrations
detected on RHNI spores were very similar to the amount detected on SU-l spores
suggesting efficient binding (Figure 5 .4). VA was not detected in extracts of ATCC36537
spores and water used to harvest spores but was detected in extracts of the whole colony
(Figure 5.5). Because VA was not detected in the water used to harvest spore, it is not
surprising that it was also not detected in extracts of SU-l and RHN 1 spores after the
binding assay (Figure 5.5 and Figure 5.6, lane 2, 3, 6 and 7). Possible explanations for the
lack of detectable VA in water used to harvest spores are the solubility of VA in water is

very low or the quantity of VA secreted on the colony surface is very low.

124

 

Table 5.6. Aflatoxin detected in conidiospores isolated under wet and dry

conditions from A. parasiticus grown on YES and PDA agar.

 

 

 

 

 

 

 

Spore Water
Wt". (mg) Total (ng) Conc. ° Vol. ° (ml) Total (ng) Conc.
(ng/ mg) (118/ m1)
Dry'
YES 3 .0 l 56 52
1 .3 226 l 74 8 194 24
PDA 1 .0 1 3 5 1 3 5 8 72 9
Wet
PDA 10.0 16215 1622
1 0.0 1 4 l 77 141 8
9.9 11621 1174 50 63000 1260
8.5 10140 1193 50 42400 848

 

 

a: Water used in dry spore-harvest did not directly contact the colony. b: Dry weight. c:
Concentration. d: Volume.

125

1234567

U "I "

8 9

       

. 4
AFB] + OMST

AFG1 +

Figure 5.3. Thin layer chromatography of aflatoxin and OMST in water used for spore
harvest.

Lane 1. AFB,, B2, G1, G2 standards. Lane 2. Water used to harvest SUI spores
(centrifuged to remove spores). Lane 3. Water used to harvest SUI spores (0.22 um
filtrate). Lane 4. Water used to harvest SUI spores (0.45 um filtrate). Lane 5. Water used
to harvest RHNI spores (centrifuged to remove spores). Lane 6. Water used to harvest
RHNl spores (0.22 um filtrate). Lane 7. Water used to harvest RHNI spores (0.45 um
filtrate). Lane 8. Fluid (brown color) collected from the surface of RHNI colony. Lane 9.

OMST standard.

126

 

123456789

a OMST
AFB] ,

AFG1 +

 

Figure 5.4. Thin layer chromatography of RHNI spore extracts.

Lane 1. SUI whole colony extract. Lane 2. AFB1,B2,GI and G2 standards. Lane 3. SUI
spore extract. Lane 4. RHN 1 spore extract. Lane 5. RHN] spores mixed with water used
to harvest SUI spores (0.22 urn filtrate). Lane 6. RHN 1 spores mixed with water used to
harvest SUI spores (0.45 um filtrate). Lane 7. Fluid (brown color) collected fi'om the

surface of RHNI colony. Lane 8. OMST standard. Lane 9. RHN] whole colony extract.

127

STD 1 2 3 4 5

VA

AFB, ->
AFG, ->

 

Figure 5.5. Thin layer chromatography of SUI and ATCC36537 spore extracts.
Lane 1. SUI spore extract. Lane 2. ATCC 36537 spore extract. Lane 3. ATCC 36537

spore extract. Lane 4. ATCC 36537 whole colony extract. Lane 5. VA standard.

128

 

123456789

7

    

VA

OMST
AFB,

AFG1-P "

Figure 5.6. Thin layer chromatography of SUI and RHN] spore extracts after binding
with water used to harvest ATCC36537 spores.

Lane 1. AFB1, B2, G1 and G2 standards. Lane 2. Extract of SUI spores mixed with water
used to harvest ATCC36537 spores (0.22 pm filtrate). Lane 3. Extract of ATCC36537
spores. Lane 4. Extract of SUI spores mixed with water used to harvest RHNI spores
(0.22 pm filtrate). Lane 5. OMST standard. Lane 6. Extract of RHNI spores mixed with
water used to harvest ATCC36537 spores (0.22 pm filtrate). Lane 7. Water used to
harvest ATCC36537 spores (0. 22 pm filtrate). Lane 8. VA standard. Lane 9. SUI whole

colony extract.

129

 

AFB, concentrations in spores harvested under dry condition. The AFB,
concentrations in spores were also measured in spores harvested under dry conditions
(Table 5.6). Spores harvested from two cultures (PDA and YES) contained
concentrations from 50 to 174 ng of aflatoxin per mg of spores, which is about 10 fold
lower than the results using a wet harvest method. Only 73 to 194 ng of aflatoxin was

detected in water used to release spores from cotton tips of applicators.

DISCUSSION

In preliminary experiments, Aspergillus was cultured on nylon membranes on
which the mycelia was firmly attached; the bottom portion of the mycelia could not be
completely recovered for analysis and, therefore, this caused difficulty in estimating the
aflatoxin concentration in the colony. This problem was solved by replacing the nylon
membrane with a cellophane membrane; the mycelia could be fractionated and
completely removed from the cellophane membrane

The colony fractionation results showed that, from day 2 to day 5, the ratio of
aflatoxin in the fungal mycelium to total aflatoxin produced by the colony decreased.
However, the ratio then increased at day 6 and 7. One possible explanation is that the
colony edge made contact with the Petri dish at day 6. This possibly could be a signal to
the fungus that the space and nutrients are limited, resulting in a change in the normal
physiological conditions to promote aflatoxin production. Another possible explanation is
a pH change in the medium. As reported by Mellon et al., the pH in the medium dropped
to be its lowest value at day 5 (under their culture conditions), possibly stimulating higher

levels of aflatoxin synthesis in the mycelium (3).

130

 

The media used was observed to affect the distribution of aflatoxin within and
outside of the fiingal colony. Whereas 80% of the aflatoxin was released into the medium
when cultured on PDA agar for 7 days, more than 50% of the aflatoxin was found in the
mycelium cultured on YES agar. The reason for this is not clear but may be related to the
level of sporulation. I can observe two major differences in colony morphology on PDA:
more green spores and less mycelial mass. Although it has been previously hypothesized
that aflatoxin synthesis and asexual sporulation are tightly associated (2,4,6), no
estimation was conducted on the relative distribution of aflatoxin in the fungal mass and
the culture medium. Researchers previously measured aflatoxin in the medium only or in
whole culture extracts (mycelium and medium). I hypothesize that, since spores are
important structures for dispersal and colonization, if aflatoxin is harmful (carcinogen) to
the fungus, then the ability to reduce aflatoxin concentration inside of fimgal mycelium
prior to and during spore production could be necessary to prevent mistakes in DNA
replication caused by DNA adduct formation. Therefore, a related hypothesis is that the
substrate mycelium is still highly active in aflatoxin production but the toxin is secreted
outside of the mycelium to prevent toxin accumulation. This mechanism could enhance
the precise production of spores by conidiophores.

It is obvious that spores harvested under dry conditions have a much lower
aflatoxin concentration than those collected under wet conditions (with water). The water
used to harvest spores contains high aflatoxin concentrations and is responsible for the
higher aflatoxin detected in spores as compared to mycelium. Under dry harvest
conditions, the water used to release the spores from the cotton-tipped applicator
contained very low but detectable amounts of aflatoxin. The traces of aflatoxin in this

water may come fi'om the fluid on the colony surface, absorbed by applicator and released

131

 

into water. In support of this notion the brown-colored liquid collected from the colony
surface of RHNI contained significant levels of OMST (Figure 4, lane 7). We believe this
fluid on the colony surface was secreted by fungal cells.

It has been demonstrated previously that the variation in spore aflatoxin
concentration between strains was high and was affected by the culture conditions and
spore-harvest methods (4,6). Wicklow and Shotwell (1982) reported that condiospores
harvested from Czapek agar cultures (in the dark at 28° C for 21 days) using water
contained a range of 1,106 to 54,300 ppb (ng per g) of AFB, in four A. parasiticus strains
and a range of 36 to 97,400 ppb in five A. flavus strains. Palmgren and Lee (1986)
reported that the condia harvested from A. parasiticus (SRRC-2004) cultured on
autoclaved rice and incubated at 25°C for 7 days contained 161,000 ppb (161 ng per mg)
of AF B,. In this report, the spores were harvested by vacuum (dry conditions). Based on
our data, we suspect that the real aflatoxin concentration in spores in Wicklow (1982)
was likely 10-fold lower than that reported because Czapek agar media is not a good
aflatoxin-inducing media. In addition, the method used to harvest the spores (similar to
the wet condition reported in our study), caused an artificially high concentration of
aflatoxin detected in spores; the aflatoxin extracted from the colony bound to the spore
surface. The results reported by Palmgren (4) were similar to ours (proximately 126 ng
per mg); we believe these data generated using a dry-harvest method more closely
represent the actual concentration in spore.

My current data show that both fungal hyphae and spores isolated from aflatoxin-
producing strains contain aflatoxin. However, for a nar-I mutant, spores (harvested by
vacuum) contained very low amount of NA (280 ng per g of spores) as compared with

mycelia (760 ug per g) (4). This observation prompts us to propose that aflatoxin is

132

synthesized in the mycelium and then needs to be transported to spores. In this regard, it
will be interesting to determine if all aflatoxin intermediates or only certain pathway-

intermediates can be accumulated in spores.

133

 

REFERENCES

1. Adams T. H. and J-H. Yu. 1998. Coordinate control of secondary metabolite
production and asexual sporulation in Aspergillus nidulans. Curr Opinion Microbiol. 1:
674-677.

2. Guzman-de-Pena D and J. Ruiz-Herrera. 1997. Relationship between aflatoxin
biosynthesis and sporulation in Aspergillus parasiticus. Fungal Genet Biol. 21:198-205.

3. Mellon, J. E., P. J. Cotty, and M. K. Dowd. 2000. Influence of lipids with and
without other cottonseed reserve materials on aflatoxin B, production by Aspergillus
flavus. J. Agric. Food Chem. 48:3611-3615.

4. Palmgren MS. and LS. Lee. 1986. Separation of mycotoxin-containg sources in
grain dust and determination of threir mycotoxin potential. 66:105-108.

5. Pestka, J. J., P. K. Gaur, and F. S. Chu. 1980. Quantitation of aflatoxin BI and
aflatoxin Bl antibody by an enzyme-linked immunosorbent microassay. Appl. Environ.
Microbiol. 40: 1027-1031 .

6. Wicklow, D. T., and 0. L. Shotwell. I983. Intrafungal distribution of aflatoxins

among conidia and sclerotia of Aspergillus flavus and Aspergillus parasiticus. Can. J.
Microbiol. 29:1-5.

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CHAPTER 6

Future Studies

Based on the current data, it is difficult to determine if the vacuolar localization of
OmtA occurs for protein recycling, for enzyme activation, or for some other purpose in A.
parasiticus. The compartment for storage of aflatoxin B, in the fungus was also not
determined in this study. We propose three alternative models for synthesis and export of
aflatoxin from fungal cells that are consistent with data from this and previous studies.
We propose these models as a primary means to design future research directions (Figure
6.1).

Model I. Aflatoxin enzymes aggregate in large complexes in the cytoplasm to
carry out sequential reactions efficiently. Aflatoxins are synthesized in the cytoplasm and
are transported to the extra-cellular environment via AflT (with identity to a transproter
protein), via another transporter, or via direct release into the extra-cellular environment
when cells die. To fine tune the level of (reduce or limit) aflatoxin synthesis, aflatoxin
enzymes (OmtA in particular) are transported to and inactivated in vacuoles by
proteolytic cleavage.

Model 2. Aflatoxin enzymes form complexes in the cytoplasm to generate the
late pathway intermediate sterigmatocystin (ST); the intermediates in the pathway
upstream from ST are passed directly from one active site to the next in the enzyme
complex limiting the quantity of free intermediates in the cytoplasm. OmtA is
synthesized and participates as one enzyme in the complex but it is present in inactive

form. As ST bound to OmtA accumulates in the complexes, OmtA carries ST into

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vacuoles in cells near the basal surface of the colony. Proteolytic cleavage activates
OmtA that then converts ST to OMST. OrdA, also localized to the vacuole, converts
OMST to AFB, that accumulates in the vacuole. As it fills, the vacuole fuses with the cell
membrane and releases AFB, into the external environment at the basal surface (substrate
surface) of the fungal colony. Purified native OmtA has a molecular weight about 40
kDa. Based on the nucleotide sequence of the cDNA, OmtA has a predicted molecular
mass of 45 kDa suggesting that a 5 kDa peptide is cleaved from its N-terminus. This
cleavage could inactivate OmtA as proposed in Model 1 or activate OmtA as proposed in
Model 2. In support of this idea, Western blot analysis demonstrated that OmtA is
cleaved in fungal cells and the first cleavage releases a peptide of the predicted size.

Model 3. This model is similar to Model 2 except OmtA is firnctional in the
cytoplasm. OmtA and its product, OMST, are transported together to the vacuole where
OMST is converted to AFB, by OrdA.

Three experiments will be conducted in the future to allow discrimination
between these models: 1) co-immunoprecipitation, 2) subcellular fractionation, 3) in situ
localization of OrdA by immunoelectron microscopy (IEM). Results from Experiment 1
will answer the question whether aflatoxin proteins can form protein aggregates to carry
out several reactions together. Results from Experiment 2 and 3 will answer the question
regarding where OrdA (60 kDa) and aflatoxins are compartmentalized. The purified
organelles will be used to test their ability to convert OMST to AFB, in feeding
experiments; they will also be freeze-substituted and immunolabeled to confirm protein
localization. The protein content extracted from isolated organelles will be assayed to
determine their aflatoxin protein profile using Western blot analysis. To assist in these

experiments, the required OrdA antibody was generated using similar procedure as

136

described in Chapter 3. The specificity of this antibody was tested by Western blot
analysis, and the results are shown in Figure 6.2. In addition, a MBP-AflJ fusion protein
and AflJ antibody were also produced (Figure 6.3). These reagents will be used in
Experiment 1 to test the possibility that the protein-protein interaction between AM (47
kDa) and AflR (and other transcription factors) or proteins involved in microbody
targeting pathway.

The primers used in PCR to clone ordA cDNA were: 5’- ATG ATT TCTAGA

ATA ATT ATT TGT GCG G-3’ and 5’- GTA AAQQTT TCA AAT CAT CTG ATT-3’.

The primers used to clone aflJ cDNA were: 5’- TCC TCTAGA ATG ACC TTG ACT
GAC CTA G-3’ and 5’- CAT AAGQTT TTA ATA TCG GTT GTC ATC G -3’. (XbaI

restriction sequence: TCT AGA; HindIII: AAG CTT).

137

 

 

 

 

Model 1 Model 2

 

 

 

 

 

 

Cytoplasm fl Medium Vacuole ’ Medium

   

Figure 6.1. Proposed models for synthesis and export of aflatoxin from firngal cells.

138

123456 7 89STDkDa
120
100
so
60

50
40

30

OrdA ——> “*‘ "" ‘ “’

20

 

Figure 6.2. Western blot analysis of fungal protein extracts using affinity-purified OrdA
PAb. Proteins were extracted from time-fractionated colonies of SU-l and AF SIO grown
on YES agar medium. 60 pg of protein was loaded in each lane on SDS-PAGE. OrdA
antibody (#137) was purified by a column packed with AF S10 (aflR knockout) protein.
OrdA antibody was used at a concentration of 6 pg per ml in the experiment. Lane 1, SI
fraction; 2, SZ fraction; 3, S3 fraction; 4, R1 fraction; 5, R2 fraction; 6, R3 fraction; 7,

32cm fraction; 8, S fraction; 9, S6,m fraction. Lane STD contains molecular mass

4cm
standards; mass of standards is shown at the right side of blot. SW, SW, and S6cm are
fractions isolated from SU-l colonies grown for 90-h and were fractionated according to

their diameters. S covers from center to 2cm; Sm“, 2 to 4 cm; SW, 4 to 6 cm.

2cm

139

 

 

Figure 6.3. Production of MBP-AflJ fusion protein and AflJ antibody.

(A) SDS-PAGE of IPTG- induced bacterial crude extracts containing MBP-AflJ. Lane I,
Non-induced bacterial crude extract; 2, IPTG- induced bacterial crude extract; 3,
Molecular mass standards; molecular mass is indicated to the right of the gel.

(B) SDS-PAGE of affinity purified MBP-AflJ. Lanes 1, Affinity-purified MBP-AflJ; 2,
Molecular mass standards; molecular mass is indicated to the right of the gel.

(C) Western blot analysis of fungal protein extracts using AflJ PAb (#124). Proteins were
extracted from time-fractionated colonies of SU-l and AF SIO grown on YES agar
medium. 60 pg of protein was loaded in each lane on SDS-PAGE. AflJ antibody was
used at l to 1000 dilution in the experiment. Lane 1, Molecular mass standards; mass of
standards is shown at the left side of blot; 2, SI fraction; 3, 82 fraction; 4, S3 fraction; 5,

R1 fraction; 6, R2 fraction; 7, R3 fraction.

140

 

kDal

\llllll

\

Figure 6.2

1 2 kDa
+8." 97
MBP-ADJ H 66
.1 45
d 31
5 6 7
3......“ <— AflJ

141