Molecular Microbiology (2002) 44(2), 573–583
Nitrate and the GATA factor AreA are necessary for
in vivo binding of NirA, the pathway-specific
transcriptional activator of Aspergillus nidulans
Frank Narendja, Sabine P. Goller, Markus Wolschek†
and Joseph Strauss*
Zentrum für Angewandte Genetik, University of
Agricultural Sciences Vienna, Vienna, Austria.
Summary
In Aspergillus nidulans, the genes coding for nitrate
reductase (niaD) and nitrite reductase (niiA), are transcribed divergently from a common promoter region
of 1200 basepairs. We have previously characterized
the relevant cis-acting elements for the two synergistically acting transcriptional activators NirA and AreA.
We have further shown that AreA is constitutively
bound to a central cluster of four GATA sites, and is
involved in opening the chromatin structure over the
promoter region thus making additional cis-acting
binding sites accessible. Here we show that the
asymmetric mode of NirA–DNA interaction determined in vitro is also found in vivo. Binding of the
NirA transactivator is not constitutive as in other binuclear C6-Zn2+-cluster proteins but depends on nitrate
induction and, additionally, on the presence of a wildtype areA allele. Dissecting the role of AreA further,
we found that it is required for intracellular nitrate
accumulation and therefore could indirectly exert its
effect on NirA via inducer exclusion. We have tested
this possibility in a strain accumulating nitrate in the
absence of areA. We found that in such a strain the
intracellular presence of inducer is not sufficient to
promote either chromatin rearrangement or NirA
binding, implying that both processes are directly
dependent on AreA.
Introduction
In Aspergillus nidulans, most genes coding for activities
involved in the utilization of nitrogen sources are subject to
two forms of regulation, pathway-specific induction and
nitrogen metabolite repression. Nitrate and nitrite inducAccepted 28 January, 2002. *For correspondence. E-mail
[email protected]; Tel. (+43) 1 36006 6720; Fax (+43) 1
36006 6392. †Present address: Department of Internal Medicine IV,
University of Vienna, Währinger Gürtel 18–20, A-1190 Vienna,
Austria.
© 2002 Blackwell Science Ltd
tion require the pathway specific activator NirA, whereas
nitrogen metabolite repression is mediated by the positiveacting wide-domain regulatory protein AreA. Genetic
evidence shows that the two regulatory proteins NirA and
AreA act synergistically but with independently defined
roles to assure a co-ordinated regulation of nitrate utilization in A. nidulans (for a review, see Scazzocchio and Arst,
1989).
Nitrate assimilation involves nitrate uptake, and subsequent conversion via nitrite to ammonia. The genes which
code for these activities are clustered on chromosome
VIII; crnA, coding for a nitrate permease, niaD, encoding
nitrate reductase (NR) and niiA, encoding nitrite reductase (NiR). The two latter genes encoding these enzymes
are divergently transcribed and are separated by 1200
basepairs (bp). The expression of crnA, niiA and niaD
structural genes is strictly dependent on both functional
NirA and AreA (Brownlee and Arst, 1983;Scazzocchio and
Arst, 1989; Unkles et al., 1991). Genetic and biochemical
evidence suggests that at least one other nitrate transporter exists in A. nidulans as crnA– mutants are still able
to utilize nitrate as a sole nitrogen source. In plants, in
which nitrate serves as a prevalent nitrogen source, the
transport process is a key step in nitrate assimilation. On
the basis of chlorate resistance mutants and polymerase
chain reaction (PCR) approaches using crnA-based
degenerate primers, low- and high-affinity uptake systems
have been cloned (reviewed by Daniel-Vedele et al.,
1998). It appears that the high-affinity plant uptake
systems are regulated by available nitrate and reduced
nitrogen in a similar way to that described for fungi (Aslam
et al., 1992). So far, no regulatory genes have been identified in plants either for the transporters or for the structural genes encoding nitrate and nitrite reductase.
The corresponding regulatory genes of A. nidulans
have been cloned and characterized as DNA-binding
transcription factors (Kudla et al., 1990; Burger et al.,
1991a; b). NirA contains a binuclear Zn2+-cluster domain
of the GAL4p type but specific recognition does not
involve the typical inverted, everted or direct repeat
sequences as shown for many Zn2+-cluster proteins
(Schjerling and Holmberg, 1996). Instead, NirA binds as
a dimer to a non-repeated, asymmetrical 5¢-CTCCGHGG3¢ sequence. The NirA protein was the first to be described
in which a dimer binds such a sequence, presumably
574 F. Narendja et al.
each monomer making different contacts on each strand
(Strauss et al., 1998). AreA contains a C2-C2 Zn2+-finger
and belongs to the so called GATA factor family of transcription factors (recognizing a DNA motif with a 5¢WGATAR-3¢ core sequence (Omichinski et al., 1993a; b;
Starich et al., 1998). This class of eukaryotic transcriptional activators are known to be involved in the regulation of a variety of metabolic and developmental processes (Wilson and Arst, 1998; Scazzocchio, 2000
and references therein). It is proposed that promoter specific recognition and discrimination between the different
set of genes regulated by GATA factors is determined
by subtle differences in the hydrophobic protein-DNA
contacts (Ravagnani et al., 1997), by combinatorial interactions among GATA factors (Charron et al., 1999) or by
interaction with pathway-specific transcription factors
(Feng and Marzluf, 1998). In Saccharomyces cerevisiae,
regulated nitrogen catabolic gene transcription is also
mediated by GATA factors (reviewed by Coffman and
Cooper, 1997).
The contribution of the four NirA and 10 AreA binding
sites in the niaD–niiA intergenic region (IGR) to nitrateresponsive expression of NR and NiR was analysed by
deletion studies in vivo (Punt et al., 1995). The central
NirA site 2 acts bidirectionally and contributes to more
than 80% of niiA and niaD transcription in an areA+ background. It is located in the proximity of a cluster of four
AreA sites. These GATA sites are responsible for about
80% of the transcriptional activity in a nirA+ background
(Muro-Pastor et al., 1999). These authors showed in vivo
that a GATA factor is directly involved in the remodelling of chromatin. NirA site 2 is located at the border of
positioned nucleosome-1 and the clustered GATA sites
5–8 are situated in a nucleosome-free region. AreAdependent remodelling occurs upon nitrate induction in
the absence of ammonia. Mutation of all four GATA sites
could not prevent this nucleosomal rearrangement, which
suggests either a mechanism acting at distance from the
remaining six AreA binding sites in the IGR, or a mechanism which is independent of DNA binding. The repositioning of four nucleosomes out of five monitored in this
region is independent from the NirA-specific transcription
factor and hence from the onset of transcription. In vivo
footprinting, which was adapted to filamentous fungi by
Wolschek and colleagues (Wolschek et al., 1998) showed
that AreA binds to site 5 under derepressed conditions
(regardless of whether non-induced or induced conditions). Upon repression, occupancy of AreA site 5
decreases but protection of this site is still clearly visible
even under conditions of full repression. Only in areA
loss-of-function mutants, such as areA600 or areA18, is
protection completely lost.
Here we report the in vivo binding characteristics of the
pathway-specific NirA transcription factor. NirA recognizes
in vivo the same asymmetrical, non-repeated sequence
as it does in vitro. We show that in contrast to the yeast
GAL4p activator which is bound under non-induced conditions (Kodadek, 1993), binding of NirA is dependent on
intracellular nitrate and on a functional AreA protein.
Results
The kinetics of nitrate induction
We have previously shown that the genes coding for
nitrate reductase (niaD) and nitrite reductase (niiA) are
co-regulated in response to the availability of nitrate (Punt
et al., 1995). To monitor the appearance and disappearance of the niaD transcript in detail, we have set up an
experimental procedure in which mycelium was precultured under non-inducing conditions (on urea as sole
nitrogen source, NI). These cultures were aliquoted and
transferred to medium containing only nitrate (induced, I),
simultaneously nitrate plus ammonia (induced/repressed,
I/R) or ammonia following 20 min of nitrate induction
(I20 + R). Aliquots were incubated for different times
(5–90 min) and the cultures were harvested for Northern
analysis, reporter enzyme assays, in vivo footprinting and
chromatin analysis.
The results of the Northern experiments are shown in
Fig. 1 for the wild-type (Fig. 1A) and for nirA and areA
mutant strains (Fig. 1B). In the wild type, under noninduced conditions, niaD transcript can hardly be
detected. Nitrate induction results in a rapid accumulation
of niaD transcripts already after 10 min. The simultaneous
Fig. 1. Northern blot analysis of niaD transcription in Aspergillus
nidulans wild-type strain (A) after incubation on different nitrogen
sources and in A. nidulans wild-type, nirA and areA loss-of-function
mutants (B) after 20 min of nitrate induction. The strains were
pre-grown and processed as detailed in Experimental procedures.
Different nitrogen sources (NI, non-induced; I, induction by nitrate;
I/R, simultaneous induction by nitrate and repression by ammonia;
I20 + R, induction by nitrate for 20 min and subsequent addition of
ammonia) were then added and the cultures were incubated further
for the time indicated (10, 20, 45 and 90 min). Finally, the samples
were harvested by filtration and total RNA was isolated. Per
sample, 20 mg of total RNA was loaded onto the gels.
Rehybridization with a probe of the A. nidulans actin gene was
used as a loading control. Numbers below the panel indicate the
normalized relative niaD signal intensity for each condition in
relation to the loading control.
© 2002 Blackwell Science Ltd, Molecular Microbiology, 44, 573–583
NirA binding is dependent on nitrate and AreA 575
addition of nitrate plus ammonia prevents the induction
process (I/R10 to I/R90). Induction for 20 min and subsequent addition of ammonia to the culture (I20 + R10 to
I20 + R90) medium results in a rapid decrease in niaD
mRNA abundance, which is in accordance with an areAmediated repression mechanism (Kudla et al., 1990). As
expected, the nirA 637 and areA600 loss-of-function
mutant strains, which are unable to utilize nitrate as a sole
nitrogen source, did not accumulate niaD transcript over
the investigated time-period (Fig. 1B).
Simultaneously, we investigated niiA and niaD expression using the GUS and lacZ double reporter vector
pTRAN3–1 A (Punt et al., 1995), which monitors expression of niaD-GUS and niiA-lacZ from the intergenic
region. We found that the expression profiles detected in
the Northern experiments were mirrored by the enzyme
assays of the reporter (data not shown).
NirA binding in vivo to its cognate sequence is
nitrate-dependent
We have previously shown by in vitro binding and a
number of protection and interference in vitro footprinting
studies that NirA, a member of the Zn6-cluster family of
transcription factors, binds as a dimer to a non-repeated,
asymmetrical sequence with the consensus 5¢CTCCGHGG-3¢ (Strauss et al., 1998). We have further
established the physiological role of the four NirA binding
sites in the niiA-niaD intergenic region by deletion and
mutagenesis studies (Punt et al., 1995). These studies
revealed a major bi-directional contribution of the central
NirA site 2 of about 80% of the total induction potential.
A cluster of four AreA-binding GATA sites is neighbouring
this central NirA site 2 and, in a recent mutagenesis study
(Muro-Pastor et al., 1999), we have allocated the main
physiological role to these four sites acting synergistically
with NirA site 2 to achieve maximal induction levels.
We were now interested to see whether the unusual
recognition mode of NirA in vitro binding is also mirrored
in vivo and whether binding of the activator is inducerdependent or not. Numbering of the bases within the NirA
site was carried out as in the work by Strauss and
colleagues (Strauss et al., 1998), i.e. 1–8 on the strand
showing the 5¢3¢-consensus sequence (5¢-C1TCCGC
GG8) and 1* to 8* in the 5¢3¢-orientation of the complementary strand (5¢C1*CGCGGAG8*). The in vivo methylation protection pattern shown in Fig. 2 reveals three
conserved guanines within the complementary strand of
the consensus sequence C1*CGCGGAG8* as strongly
protected, i.e. G3*, G5* and G6*. Footprints carried out on
the opposite strand of NirA site 2 (Fig. 3) showed a protection pattern consistent with the in vitro NirA-DNA
contacts, i.e. guanines G5, G7 and G8 are protected from
methylation under inducing conditions. Additionally,
© 2002 Blackwell Science Ltd, Molecular Microbiology, 44, 573–583
Fig. 2. In vivo methylation protection of NirA site 2 in the niiA-niaD
intergenic region of wild-type (left) nirA and areA mutant strains
(right). A relevant section of the sequence of the niaD coding strand
is shown at the right of the figure. The sequence highlighted in bold
letters corresponds to the complementary sequence of the
asymmetric NirA binding site (consensus 5¢-CTCCGCGG-3¢). Filled
squares () indicate protected guanines within the NirA binding
site. ‘Vitro’ indicates protein-free control DNA methylated after
isolation of total DNA from mycelium pre-grown under the standard
conditions (12 h, 30∞C on 1.25 mM ammonia). Protection of G3*, G5*
and G6* (see text for nomenclature of base numbering in the NirA
binding site) is clearly visible already after 5 min of induction and
pertains throughout the induction period of 90 min. Neither wild-type
under non-induced conditions nor the nirA and areA mutant strains
show relevant protection of the guanines within the NirA binding
site 2. Growth conditions were exactly as described for the
Northern experiment in Fig. 1. Left, wild-type cultures induced for
different times: I5, I10, I20, I45 and I90, induced for 5, 10, 20, 45
and 90 min respectively. Right, comparison of wild-type noninduced (NI WT), with 20 min induced cultures of areA600
(I areA600)and nirA637 (I nirA637) loss-of-function strains.
adenine A10, which is hypersensitive in in vitro methylation assays (Strauss et al., 1998), also becomes hypersensitive in the in vivo assay (Fig. 3; A10 is marked with
arrows). From these results, it becomes apparent that
the unusual DNA–protein interaction profile obtained in
in vitro studies with a truncated NirA protein (representing the bi-nuclear Zn-cluster and the dimerization domain)
is also seen with the native, full-length transcription factor
in vivo.
Our data show that binding of NirA is strictly dependent
on the presence of inducer (Figs 2 and 3, lanes I10 to
I90). Binding of NirA to its DNA target is very fast and can
be monitored even 5 min after induction (Fig. 2, I5). The
nirA637 mutant, which contains an in-frame deletion of
the DNA-binding domain (Muro-Pastor et al., 1999), as
576 F. Narendja et al.
Fig. 3. In vivo methylation protection of NirA site 2 of the niiA coding strand (opposite strand compared with Fig. 2) in the niiA-niaD intergenic
region of wild-type, nirA and areA mutant strains under conditions of non-induction (NI), induction (I), induction for 20 min and subsequent
repression (I20 + R) and induction and simultaneous repression (I/R). The nirA637 and areA600 mutant strains were induced for 20 min.
Numbers below the panel indicate the relative intensity of bands within the NirA binding site for each condition (see Experimental procedures).
A relevant section of the sequence of the niiA coding strand is shown at the right of the figure. The sequence highlighted in bold letters
corresponds to the sequence of the asymmetric NirA binding site (consensus 5¢-CTCCGCGG-3¢). Filled squares () indicate protected
guanines within the NirA binding site. The arrows indicate an adenine, which becomes hypersensitive to methylation after binding of NirA (see
text). ‘Vitro’ indicates protein-free control DNA methylated after isolation of total DNA from mycelium pre-grown under the standard conditions.
Growth conditions and strain descriptions are as described for the Northern experiment in Fig. 1.
expected, does not show protected bases in the binding
site.
NirA dissociates under conditions of induction
plus repression despite sufficient intracellular
nitrate concentrations
When we assayed binding of NirA under conditions of
induction and repression (Fig. 3, I/R10 to I/R90 and
I20 + R10 to I20 + R90), we found that NirA is bound to its
cognate site for 20 min even in the presence of ammonia.
At 45 and 90 min of repression, occupancy of NirA site 2
is lost. As described above, there are neither niaD transcript nor reporter enzyme activities accumulating under
conditions of induction plus repression, and the lack of
transcriptional activation can be attributed to the inactivation of AreA by ammonium. As binding of NirA is
dependent on inducer, a possible explanation for NirA
dissociation from its target after 20 min would be low internal nitrate concentrations after this prolonged repression
time. We therefore analysed intracellular nitrate concentrations of cells incubated with the relevant nitrogen
sources.
Figure 4 shows intracellular nitrate concentrations of
cells subjected to non-inducing, inducing and inducingrepressing conditions. The nitrate uptake systems are
rapidly induced and this results in an intracellular nitrate
level of more than 200 ng mg–1 dry weight (DW) after
10 min (Fig. 4A). Under conditions of prolonged induction
(90 min) this level drops to roughly 30 ng mg–1 DW. This
level is sufficient for NirA to remain bound to site 2 when
nitrate alone serves as nitrogen source, and is also sufficient to maintain transcription. Under conditions of induction and repression (Fig. 4B), nitrate accumulates to a
level of 250 ng mg–1 DW, which is to be expected as nitrate
is taken up without being metabolized. Despite the fact
that almost no NR activity is present, the internal nitrate
level drops to roughly 100 ng mg–1 DW after 20–45 min
(Fig. 4B and C). This finding suggests that a nitrate secretion mechanism is operating in A. nidulans.
However, this result indicates that low internal nitrate
concentrations do not account for the loss of NirA binding
after 45 min under I/R and I20 + R conditions. The intracellular nitrate concentrations measured under these
conditions (70–150 ng mg–1 DW) are well above the concentrations of I90 (30 ng mg–1 DW) in which NirA remains
bound.
Repression does not lead to NirA degradation
We also considered the possibility that NirA might be
subjected to ammonia-triggered protein degradation.
Owing to the extremely low NirA level in cells that do not
allow detection of the protein by Western blotting (M.
Muro-Pastor and C. Scazzocchio, personal communication), we were not able to monitor a putative nitrogen
source-dependent degradation process. We therefore
tested this process indirectly. We pre-grew the wild-type
mycelium under repressing conditions, and transferred
aliquots to induction in the presence and absence of the
protein synthesis inhibitor cyclohexamide (Strauss et al.,
1999). We reasoned that cyclohexamide should prevent
© 2002 Blackwell Science Ltd, Molecular Microbiology, 44, 573–583
NirA binding is dependent on nitrate and AreA 577
Binding of NirA to its DNA target is dependent on a
functional areA allele
Fig. 4. Nitrate uptake in a wild-type strain under conditions of
induction (A), induction and repression simultaneously (B) and
subsequent repression following 20 min of induction (C). Growth
conditions and intracellular nitrate measurements are as described
in Experimental procedures. At time-points indicated in the figure,
aliquots of mycelia were taken and the amount of intracellular
nitrate was determined using the enzymatic assay. Error bars
indicate standard errors from at least three independent uptake
experiments. Abbreviations of induction and repression conditions
are as in Figs 1, 2 and 3.
de novo synthesis of NirA in ammonia-grown cultures,
which are then transferred to nitrate. In in vivo footprinting
assays we did not see any difference between the control
mycelium and the cyclohexamide-treated mycelium. In
both cases, NirA binding site 2 was occupied already after
5 min of induction (data not shown). Thus, it seems very
unlikely that NirA is subject to ammonia-mediated degradation and the results are consistent with the view that loss
of NirA site 2 occupancy under inducing/repressing conditions is not due to increased turnover of the activator.
© 2002 Blackwell Science Ltd, Molecular Microbiology, 44, 573–583
In Figs 2 and 3, we also show that NirA binding does not
respond to nitrate in the absence of a functional AreA
protein (Figs 2 and 3, lanes I areA600). A possible explanation is that in an areA mutant background, complete
inducer exclusion prevents nitrate-dependent association
of NirA with binding site 2.
In fact, measuring intracellular nitrate levels under
inducing conditions (Fig. 5), we found that areA600 does
not follow wild-type kinetics but only accumulates up to
15 ng mg–1 DW intracellular nitrate over the whole incubation time of 90 min. It is interesting to note that in the
non-induced control experiments (NI), carried out with
nitrate and nitrite-free urea as sole nitrogen source, a
considerable intracellular nitrate level accumulates in the
mutant strains. We have repeatedly observed this effect
also in other mutants lacking nitrate reductase activity,
such as niaD26 (unpublished). For the moment, it is not
clear how nitrate can be generated in the fungal cell as
the known pathway via nitric oxide synthase activity
(Stuehr, 1997) has not been described in ascomycetes.
However, strongly reduced intracellular nitrate levels in
the induced areA mutant strain suggest that inducer
exclusion could be at least in part responsible for our
observation that NirA does not bind in an areA loss-offunction strain. To circumvent the inducer exclusion effect,
we constructed a strain expressing the crnA nitrate
transporter gene (Unkles et al., 1991) under the AreAindependent gpdA promoter (Punt et al., 1990) in an
areA600 background. The gpd-crnA construct was tested
for functionality and was found to complement the crnA1
mutation. We transformed this construct into an
areA600/argB2 strain as a single copy integrated at the
argB locus. Figure 6A compares the expression pattern of
crnA and niaD of the recipient strain areA600/argB2 with
Fig. 5. Nitrate uptake of wild-type, areA600 mutant strain (defective
in nitrate uptake) and of gpd-crnA/areA600, which takes up nitrate
independently of AreA. Growth conditions and intracellular nitrate
measurements are as described in Experimental procedures. At
time-points indicated in the figure, aliquots of mycelia were taken
and the amount of intracellular nitrate was determined using the
enzymatic assay. Error bars indicate standard errors from at least
three independent experiments.
578 F. Narendja et al.
Fig. 6. Northern blot (A) and chromatin structure analysis (B) of wild-type, areA600 mutant strain (defective in nitrate uptake) and of gpdcrnA/areA600, which takes up nitrate independently of AreA.
A. The cultures were grown under standard conditions and transferred to non-inducing conditions (NI), inducing conditions for 90 min (I90) and
simultaneous inducing and repressing conditions (I/R 90) for 90 min. The membranes were probed to monitor the expression of the nitrate
reductase gene (niaD), the nitrate transporter (crnA) and reprobed again with the actin gene as a loading control.
B. Chromatin organization of the niiA-niaD intergenic region in a wild-type, areA600 and a gpd-crnA/areA600 strain. Growth conditions are as
described in Experimental procedures. Crude nuclei were treated for 5 min with 125 or 250 U MNase per gram of mycelia (wet weight) at
30∞C. Increasing quantities of enzyme are symbolized on top of the lanes. DNA was digested with HindIII and subjected to indirect endlabelling using a HindIII–EcoRV fragment of the niiA gene as a probe. The positions of nucleosomes are pictured at the right as ellipses and
are numbered divergently from the nucleosome free region (nfr) as referred to by Muro-Pastor and colleagues (Muro-Pastor et al., 1999).
Under induced conditions in the wild type (I90 wild type), in which the open chromatin structure prevails, nucleosomes are not positioned,
whereas under the same conditions in gpd-crnA/areA600 (I90 gpd-crnA/areA600) and areA600 (I90 areA600) strains positioned nucleosomes
indicate a closed chromatin structure.
these of the gpd-crnA/areA600 transformant together with
a wild-type control. crnA is not expressed to a detectable
level in either condition in an areA600 background but is
clearly expressed under all conditions in the gpd-crnA/
areA600 transformant. On the other hand, the constitutive
expression of the transporter does not lead to induction
of niaD in the areA600 strain. Thus, reversal of inducer
exclusion does not bypass an areA loss-of-function mutation. These data confirm our earlier observation that gpdcrnA/areA600 transformants are not able to utilize nitrate
as sole nitrogen source.
When we tested the gpd-crnA/areA600 strain for nitrate
incorporation under inducing conditions (Fig. 5), we
observed during a 90 min incubation time a slowly increasing intracellular nitrate level of up to 35 ng mg–1
DW. Interestingly, this strain did not reconstitute wild-type
uptake kinetics. This effect might be due to areA dependent function of the second nitrate transporter ntrB
(Unkles et al., 2001) and/or another factor necessary for
efficient nitrate uptake. However, this intracellular nitrate
level is sufficient to promote NirA DNA binding in the wild
type. We therefore used these conditions to test in vivo if
NirA binding is directly dependent on AreA. In the gpdcrnA/areA600 strain, site 2 is not occupied by NirA under
these conditions (data not shown) which provides evidence that, independently of sufficient intracellular nitrate,
functional AreA is necessary for in vivo NirA–DNA interaction at least for site 2.
It was previously shown also that the rearrangement of
chromatin structures in the niiA-niaD IGR is inducer- and
AreA-dependent. Our nitrate accumulation studies in an
areA– background prompted us to question whether AreA
dependence could be indirect because of inducer exclusion. We therefore analysed the gpd-crnA/areA600 strain
under inducing conditions for chromatin remodelling in the
nitrate-responsive niiA-niaD IGR (Fig. 6B). We observed
that in this strain the nucleosomal structure in the
bidirectional promoter is not disrupted despite sufficient
intracellular nitrate levels. This result extends our earlier
observation in respect to AreA-dependent chromatin
remodelling: AreA seems to be directly involved in this
process and not only indirectly via inducer exclusion.
Discussion
Almost all members of the fungal binuclear Zn-cluster
proteins have been shown to recognize DNA sequences
of diad symmetry and as homodimers (reviewed in
Schjerling and Holmberg, 1996). Exceptions to this rule
are the A. nidulans NirA and AlcR proteins and the
Saccharomyces cerevisiae CYP1p (HAP1p) activator. In
A. nidulans, AlcR binds as a monomer to multiple
repeated sites in the ethanol regulon (Cerdan et al., 1997;
Panozzo et al., 1997). CYP1p was shown to recognize
tandemly repeated half sites in a highly asymmetric
manner as a homodimer (King et al., 1999a; b). In CYP1p,
asymmetric dimerization and a head-to-tail orientation of
the two monomers align the protein to the half sites of
different polarities. It is not known if the mode of AlcR or
CYP1p binding determined in vitro is also mirrored in vivo.
© 2002 Blackwell Science Ltd, Molecular Microbiology, 44, 573–583
NirA binding is dependent on nitrate and AreA 579
NirA is the only known protein of this family of transcription factors which recognizes a non-repeated, fully asymmetric sequence as a dimer (Strauss et al., 1998). In vivo
footprinting of both strands of the crucial NirA site 2 in the
niiA-niaD IGR has provided evidence that the base
contacts determined in vitro are also protected from
methylation in vivo. Additional features such as hypersensitive adenines outside the consensus sequence
(A10) are also revealed in the in vivo footprint. From the
in vitro work, it was proposed that NirA contacts bases
within the recognition sequence in such a way that amino
acid side chains in the binding domain of each monomer
are able to make specific base and phosphate contacts
to the entirely separate half sites. To our knowledge, NirA
is the only case showing such a binding mode, not only
among Zn-cluster proteins but also among all DNA
binding proteins.
In our in vitro binding assays, we could not detect any
effect of nitrate on the affinity of the truncated NirA protein
carrying about one third of the native form (J. Strauss and
C. Scazzocchio, unpublished). In contrast to this finding,
we now observe inducer-dependent DNA binding in vivo.
NirA is the first member of the binuclear Zn-cluster family
to be shown to bind in vivo only in response to the induction signal. So far, all regulators of this family are known to
interact without inducer with their cognate sites. In S. cerevisiae GAL4p is bound to UASGAL in the presence of noninducing glycerol as well as inducing galactose (Giniger
et al., 1985; Lohr and Hopper, 1985; Selleck and Majors,
1987). PUT3p, the transcriptional activator of the proline
utilization pathway (Axelrod et al., 1991) and LEU3p, a
transcriptional activator that regulates leucine biosynthesis (Kirkpatrick and Schimmel, 1995) are both bound constitutively to their targets in vivo. To perform their activation
function, however, these factors need the presence of an
inducing compound, which can be either an extracellular
inducer or intracellular metabolites. Recent data indicate
that at variance with PUT3 also the PrnA activator, regulating proline utilization in A. nidulans, needs the presence
of proline to bind to its cognate sites in vivo (Gomez et al.,
2002). Inducer-dependent DNA binding mechanisms are
only known for other DNA binding protein families such
as the hormone receptor family of transcription factors
(reviewed in Dittmar and Pratt, 1997).
Regulatory processes, traditionally assayed after
several hours of induction and/or repression in A. nidulans and other fungal experimental systems, were shown
here to result in transcriptional responses within a few
minutes. This suggests that the specific molecular
machinery for the nitrate induction process must pre-exist
and only needs to be activated. areA transcripts exist
under all physiological conditions (Platt et al., 1996a; b)
and AreA is bound permanently but with different affinity
to GATA site 5 in the niiA-niaD IGR under the different
© 2002 Blackwell Science Ltd, Molecular Microbiology, 44, 573–583
physiological conditions. The nirA gene has been shown
to be constitutively transcribed at a low level (Burger
et al., 1991a; b) and, in this work, we provided evidence
that occupancy of NirA site 2 in the IGR is independent of de novo NirA synthesis, but is strictly dependent
on a functional AreA. NirA is never bound in an areA–
strain.
This situation conflicts with our observation that under
induced/repressed conditions, in which AreA is nonfunctional, NirA is able to bind for a certain time (at least
20 min). Repression-mediated nuclear export of AreA
and/or NirA would be a possibility to explain this timedelayed response. In the yeast S. cerevisiae Gln3p, a
GATA factor similar to AreA involved in the activation of
nitrogen catabolite repressible genes is retained in the
cytoplasm under repressing conditions (Beck and Hall,
1999). In Aspergillus, this mechanism is not very likely to
play an important role in AreA regulation, as we have
shown in a previous report (Muro-Pastor et al., 1999) that
the physiologically essential GATA site 5 in the niiA-niaD
IGR remains at least partially protected even under fully
repressed conditions. However, future experiments using
tagged versions of these two transcription factors will
show if the nitrogen source in fact stimulates nuclear
translocation of AreA and NirA. Additionally, physical interaction between NirA and AreA might be another prerequisite for NirA binding. As a matter of fact, direct
protein–protein interactions have been shown for the NirA
and AreA homologues of Neurospora crassa, Nit4 and
Nit2 respectively. A Nit2 mutant, which possesses wildtype DNA binding activity in vitro but is greatly impaired
in its interaction with Nit4, showed significantly reduced
expression of the target gene, nitrate reductase (Feng
and Marzluf, 1998). Other members of the GATA-factor
family such as hGATA1 and hGATA2 also require cooperative interactions with specific transcription factors
for transcriptional activation (Kawana et al., 1995;
Gregory et al., 1996).
However, on the transcriptional level the repression
mechanism inactivating AreA seems to be effective
immediately after the addition of ammonia as under
induced/repressed conditions no niiA or niaD transcripts
are accumulating. The inactivation of AreA or its homologue Nit2 of N. crassa involves the interaction with a
negatively acting protein, NMR (Xiao et al., 1995;
Andrianopoulos et al., 1998) and hence it is likely that
NMR prevents the transcriptional activation function of
AreA but not its function necessary for NirA binding.
Experimental procedures
Strains, plasmids and genetic techniques
Aspergillus nidulans strains used throughout this work are
listed in Table 1. Escherichia coli strain JM109 [F¢ traD36
580 F. Narendja et al.
Table 1. Aspergillus nidulans strains.
Strain name
Genotype
Reference
Wild type:
nirA 637
nirA 87
areA600
areA600/argB
gpd-crnA/areA600
crnA1/argB2
pabaA1
pabaA1, nirA637
biA1, fwA1, nirA87
pantoB100, biA1, areA600
pabaA1, argB2, areA600
pabaA1, [pNTC3], areA600
biA1, crnA1, argB2
Pontecorvo et al. (1953)
Muro-Pastor et al. (1999)
Tollervey and Arst (1981)
Al Taho et al. (1984)
This work
This work
This work
lacIqD(lacZ)M15 proA+B+/e14– (McrA–) D(lac-proAB) thi
gyrA96 (Nalr) endA1 hsdR17 (rK–mK+) relA1 supE44 recA1]
was used for routine plasmid propagation.
Plasmid pNTC3 expressing the A. nidulans crnA nitrate
transporter gene under the A. nidulans gpd promoter was
constructed as follows: the argB gene of pMS12 (obtained
from Fungal Genetics Stock Center) was isolated as
BamHI fragment, the restriction site was filled in and cloned
into the unique XbaI site (filled in) of pAN52-1 (Punt et al.,
1987) resulting in plasmid pNTC1. The crnA gene was
amplified with polymerase chain reaction (PCR) using primer
crnA-for (5¢-GTTGAAGAGATAGATCTCGCCAAGCTGCTG3¢) and crnA-rev (5¢-GGAAATAAGATCTAGAATCCAGAT
GCCGCG-3¢), which overlap the translational start codon
and the stop codon of crnA (Unkles et al., 1991). Both primers
contain a BglII site not present in the original crnA sequence.
The resulting PCR fragment was cloned as BglII fragment
into the BamHI site of pNTC1 resulting in plasmid pNTC3.
The gpd-crnA construct was tested for functionality by
transformation into a crnA1/argB2 strain and found to
complement the crnA1 mutation for hypersensitivity to
75 mM caesium chloride in the presence of nitrate. The transformants of the areA600/argB2 recipient strain were analysed
using Southern hybridization, and the strain used in this
study showed integration of the plasmid into the argB locus
in a single copy. Sexual crosses to obtain the recipient
strain for transformation were carried out according to
standard procedures. Transformation protocols followed the
procedures published by Tilburn and colleagues (Tilburn
et al., 1983).
Culture conditions
Strains were grown for 12 h at 30∞C in minimal medium
(Pontecorvo et al., 1953) with appropriate supplements plus
5 mM urea and 1.25 mM ammonium D-(+)-tartrate as nitrogen
source. This medium allows the growth of all strains, including areA600. The mycelia were then harvested by filtration,
washed with sterile water, and transferred to the same
medium without any nitrogen source. Incubation was continued for 20 min in this medium. To this end, the cultures were
divided into aliquots of 20 ml when the following nitrogen
sources were added: 5 mM urea (non-induced conditions,
NI); 10 mM NaNO3 (induced conditions, I); 10 mM NaNO3 and
5 mM ammonium D-(+)-tartrate (simultaneous inducedrepressed conditions, I/R); or 10 mM NaNO3 as sole nitrogen
source for 20 min followed by the addition of 5 mM ammonium D-(+)-tartrate for the remaining incubation time (sequential induced-repressed conditions, I20 + R). The aliquots were
incubated for different times (10, 20, 45 and 90 min); the
mycelium was harvested by filtration and frozen in liquid nitrogen for further processing.
Northern blots
Strains were grown under conditions described above. Total
RNA isolation and Northern hybridization was carried out as
described in Strauss et al., 1999). The probe for detecting the
niaD transcript was obtained by PCR amplification with
primers niaD-F and niaD-R (Muro-Pastor et al., 1999) from
chromosomal DNA. The crnA transcript was detected with a
PCR-generated probe using primers crnA-F (5¢-GTTGAA
GAGATAGATCTCGCCAAGCTGCTG-3¢) and crnA-R (5¢GGAAATAAGATCTAGAATCCAGATGCCGCG-3¢). The actin
probe was derived from plasmid pSF5 (Fidel et al., 1988).
Probes were labelled by random priming. Relative intensities
of signals were calculated with the IMAGEQUANT software
(Molecular Dynamics) from densitometric analysis of autoradiographs. Normalized niaD signal intensity was obtained
by dividing the intensity of the niaD signal by the intensity of
the actin signal.
Nitrate measurement and reporter enzyme assay
For reporter enzyme assays and nitrate measurement,
mycelia were harvested and extensively washed with deionized water after incubation with the different nitrogen sources
for exactly the same time as indicated for Northern and in
vivo footprinting analysis. Filtered mycelia were immediately
frozen in liquid nitrogen. The GUS and b-GAL assays were
carried out as described by Punt and colleagues (Punt et al.,
1995).
For nitrate measurement, mycelia were ground under liquid
nitrogen and resuspended in 1 ml of 70% ethanol. Cell debris
was removed by centrifugation at 13 000 g for 15 min. The
supernatant was transferred into a new tube and ethanol was
removed by drying the samples in a speed-vac desiccator.
The remaining material was dissolved in 1 ml of distilled water
and 100 ml was used for nitrate assay. Nitrate concentrations
in samples were measured with the colorimetric nitrate/nitrite
assay from Roche according to the manufacturer’s instructions, except that the assay was carried out in microtitre
plates and the volume of reagents was downscaled to one
fifth of the original amounts. All measurements including the
determination of mycelial dry weight were carried out in three
independent repetitions.
In vivo footprint studies
For in vivo footprint studies, all strains were grown under
© 2002 Blackwell Science Ltd, Molecular Microbiology, 44, 573–583
NirA binding is dependent on nitrate and AreA 581
exactly the same conditions as for the Northern Blot analysis. Methylation conditions, DNA extraction and detection
of in vivo protein–DNA interactions were carried out as
described by Wolschek and colleagues (Wolschek et al.,
1998). Briefly, 2 min before harvesting the mycelia, dimethylsulphate was added to the cultures and further incubated.
Mycelia were harvested by filtration and methylated DNA
was isolated. Following cleavage of methylated guanines
by piperidine, DNA–protein interactions were visualized by
radioactive ligation-mediated PCR (LM-PCR).
The primers detecting NirA–site 2 interaction on the niaDcoding strand were: NIR II/1, 5¢-tggctagagccgcttgacgataatg3¢, NIR II/2, 5¢-gagccggcgataagcatgatg-ttggc-3¢; and NIR II/3,
5¢-ccggcgataagcatgatgttggcgctgtc-3¢.
The primers detecting NirA–site 2 interaction on the niiAcoding strand were: NIR R II/1, 5¢-tggctagagccgcttgacgata
atg-3¢; NIR R II/2, 5¢-gagccggcgataagcatgatgttggc-3¢; and
NIR R II/3: 5¢-ccggcgataagcatgatgttggcgctgtc-3¢.
Relative intensities of signals were calculated with the
IMAGEQUANT software (Molecular Dynamics) from densitometric analysis of autoradiographs. The percentage of signal
intensity within the binding site was obtained by dividing the
mean intensity of bands within the binding site by the mean
intensity of four bands located outside the binding site. The
ratio obtained for the bands inside and outside the binding
sequence in the ‘vitro’ lane was set to 100% and the ratios
obtained for the other lanes are relative to the ‘vitro’ lane.
Chromatin structure analysis
Strains were grown under exactly the same conditions as
described for Northern Blot analysis and in vivo footprinting.
For the analysis of the chromatin structure in the niiA-niaD
intergenic region, DNase I and MNase sensitivity was tested
using the method described by Gonzalez and Scazzocchio
(1997). Culture conditions, transfer of mycelium to different
media and further incubation under different nitrogen regimes
were as described for Northern and in vivo footprinting analysis. Details on incubation times are given in the legends to
the figures. The HindIII–EcoRV fragment of the niiA-niaD
intergenic region (IGR) was used for indirect end labelling
and was isolated from plasmid pAN302 (Strauss, 1993).
Acknowledgements
The authors thank Claudio Scazzocchio (IGM, Université
Paris Sud) and Michael J. Hynes (University of Melbourne,
Australia) for sharing unpublished results and critical reading
of the manuscript. We are grateful to Herb Arst Jr. (Department of Infectious Diseases, Imperial College, London) for
providing the crnA1 mutant strain. This work was supported
by the START Program Nr. Y114-MOB of the Austrian
Science Fund (FWF) to J.S.
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