Structure–function analysis of NADPH:nitrate reductase from Aspergillus nidulans: analysis of altered pyridine nucleotide specificity in vivo

Neil Halla,1 and A. Brian Tomsett1

School of Biological Sciences, The University of Liverpool, Donnan Laboratories, Liverpool L69 7ZD, UK1

Author for correspondence: A. Brian Tomsett. Tel: +44 151 794 3616. Fax: +44 151 794 3655. e-mail: tomsett{at}liverpool.ac.uk


   ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
REFERENCES
 
Nitrate reductase (NaR) catalyses the reduction of nitrate to nitrite via a two-electron transfer. In fungi, the electron donor for NaR is NADPH whereas plants can have two enzymes, NADH:NaR and a bispecific NAD(P)H:NaR. PCR mutagenesis was employed to introduce mutations into the niaD gene of Aspergillus nidulans in order to identify residues involved in co-enzyme specificity. The niaD3000 mutation (NiaD T813D, K814Q) altered co-enzyme specificity: the new enzyme had high levels of NADH:NaR activity in vitro, whilst all NADPH-associated activity was lost. However, strains carrying this mutation did not grow on nitrate. Enzyme assays suggested that this was not due to inhibition of the mutant enzyme by NADPH. All revertants of the niaD3000 mutants had restored NADPH activity and lost NADH activity. Sequence analysis of these revertants showed that they all contained a single amino acid change at Asp-813, suggesting that this position is crucial to co-enzyme specificity. Further studies have shown that the mutant enzyme was not protected from deactivation by either co-factor in cell-free extracts (unlike the wild-type), and that induction of the glucose-6-phosphate dehydrogenase occurred independently of NADPH levels. These data highlight the importance of functional tests in vivo under physiological conditions.

Keywords: Aspergillus nidulans, nitrate reductase, site-directed mutagenesis, NADPH specificity, pentose phosphate pathway

Abbreviations: CbR, cytochrome b reductase; G6PD, glucose-6-phosphate dehydrogenase; NaR, nitrate reductase; PPP, pentose phosphate pathway

a Present address: IACR Long Ashton, University of Bristol, Long Ashton, Bristol BS41 9AF, UK.


   INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
REFERENCES
 
In eukaryotes, nitrate reductase (NaR) catalyses the first step in nitrate assimilation, the conversion of nitrate to nitrite. NaR is a multicentre redox protein that has three distinct regions (Kinghorn & Campbell, 1989 ): an N-terminal catalytic region, which binds molybdopterin and is similar to human sulphite oxidase; a central region, which is a cytochrome b557 sequence that binds haem; and a C-terminal region, which binds FAD and a reduced pyridine nucleotide and is a cytochrome b reductase (CbR).

Eukaryotic assimilatory NaRs form three classes based upon which reduced pyridine nucleotide they oxidize (Kinghorn & Campbell, 1989 ). Aspergillus nidulans, in common with other fungi, has an NADPH-specific NaR (NADPH:nitrate oxidoreductase, EC 1 . 6 . 6 . 3). Higher plants usually contain an NADH-specific NaR (EC 1 . 6 . 6 . 1), which has also been found in some algae. A third type, NAD(P)H-bispecific NaR (EC 1 . 6 . 6 . 2), has been found in some higher plants and algae, but is rarely the sole NaR in these organisms (Friemann et al., 1991 ).

To examine co-enzyme specificity, Friemann et al. (1991) aligned birch bispecific NaR with NADH- and NADPH-specific NaRs. They concluded that in NADH-specific enzymes, three successive proline residues are conserved whilst only one is conserved in the birch enzyme; a different proline residue is conserved in NADPH-specific enzymes. The importance of this sequence was tested in birch NaR by site-directed mutagenesis, which demonstrated that the NADPH-dependent activity of the bispecific enzyme could be reduced (to 7%) by replacing the proline residues which are missing in the wild-type birch enzyme; however, NADH-dependent activity was also reduced by 50% (Shondorf & Hachtel, 1995 ). A similar study in A. nidulans, however, did not alter the specificity or activity of NaR at all (Garde et al., 1995 ), suggesting that other residues must be involved.

The three-dimensional structure of the C-terminal domain of corn NaR has now been deduced by X-ray crystallography (Lu et al., 1994 , 1995 ; Campbell, 1996 ), demonstrating that this CbR fragment is a member of the FNR family of flavoenzymes, named from ferredoxin NADP+ reductase (Karplus et al., 1991 ). Whilst members of this family have little overall amino acid similarity, their three-dimensional structures in the NADH/NADPH domains are very similar, that is a variation of the Rossman or dinucleotide fold found in many dehydrogenases and reductases (see Shiraishi et al., 1998 ). The fine structure of the binding pocket for the ribose 2' hydroxyl of the adenine of NADH and 2' phosphate of NADPH has been defined for some FNR family members (Campbell, 1996 ; and see Shiraishi et al., 1998 ). The presence of a negatively charged side-chain, such as aspartic acid, at the end of the third ß-strand of this pocket is characteristic of NAD+/NADH-specific enzymes (residue 205 in Lu et al., 1994 ), whereas NADP+/NADPH-specific enzymes, including A. nidulans NaR, have serine or threonine at this position (see Shiraishi et al., 1998 ). Equivalent residues were identified as being involved in cofactor specificity in glutathione reductase (Scrutton et al., 1990 ).

It is only recently that a mutation at this position (the S920D mutation of Neurospora crassa) has been induced (Shiraishi et al., 1998 ), but this was studied only as the CbR fragment of this enzyme expressed in Escherichia coli. In the wild-type, NADPH-dependent activity was 113-fold greater than the NADH-dependent activity. Substitution of this serine by aspartic acid reduces electron donation by NADPH to 2·4% of the wild-type. Concomitantly, NADH-dependent activity was increased by 15-fold relative to the NADH-dependent activity in the wild-type; however, this only represents 13·5% of the wild-type NADPH-dependent activity, and is only 5·6 times the NADPH-dependent activity of this mutant. Whilst the NADPH/NADH specificity had essentially been reversed, the levels of activity clearly show that other residues are involved.

Such studies of NaR fragments in heterologous systems are ideal for kinetic analyses of the enzyme. However, they do not allow investigation of the activity of the holoenzyme, of the impact of pyridine nucleotide alterations on the physiology of the organism, nor of any pleiotropic effects on gene regulation. For the ideal functional genomics approach, site-directed mutations of holoenzyme must be integrated back into the native locus in a host genome lacking any residual gene sequences: this prevents spurious genomic effects from duplicate sequences (either mutant or wild-type) and heterologous dimerization between wild-type and mutant proteins. A. nidulans offers such a system (Garde et al., 1995 ). In this fungus, the structural genes for NaR (niaD) and nitrite reductase (niiA) are tightly linked and are divergently transcribed (Tomsett & Cove, 1979 ; Johnstone et al., 1990 ); transformation of a deletion strain allows selection for niiA+ arising from homologous targeted integration (Garde et al., 1995 ).

In A. nidulans, NaR expression is regulated both by substrate induction via the nirA gene product and by nitrogen metabolite repression mediated by ammonium/glutamine and the areA gene product (see Crawford & Arst, 1993 ). Furthermore, the interaction between NaR and NAD(P)H has been implicated in the regulation of the pentose phosphate pathway (PPP) and in the post-transcriptional regulation of NaR activity (Dunn-Coleman & Pateman, 1977 ; and see Cove, 1979 ). Hankinson & Cove (1974) have demonstrated that the enzymes of the PPP are induced in the presence of nitrate; this has also been observed in plants (Redinbaugh & Campbell, 1998 ). This induction could occur via the nirA pathway or via another pathway which detects the NADPH/NADP+ ratio in the cell, causing induction when NaR is active and NADPH is oxidized. Most evidence points to the former of these two models, as constitutive niaD mutants are constitutive for NADPH synthesis. However, the latter theory cannot be ruled out as these constitutive mutants may retain diaphorase activities, which could induce the PPP by oxidizing NADPH.

A. nidulans and N. crassa NaRs have also been shown to be protected from deactivation in the presence of NADPH, and deactivation can be reversed by subsequent incubation with NADPH; NADH cannot protect NaR in these organisms (Subramanian & Sorger, 1972 ; Dunn-Coleman & Pateman, 1977 ). The mechanism for this deactivation and subsequent reactivation is not understood.

Here, the effect of mutations altering pyridine nucleotide specificity, created by site-directed mutagenesis and reintroduced back into a niaD deletion strain using a homologous transformation system, have allowed analysis of their effect not only on NaR catalytic activity, but also on the growth and metabolism of A. nidulans, in particular the regulation of the PPP.


   METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
REFERENCES
 
A. nidulans strains and phenotypic studies of mutants.
The strains, media, growth conditions and phenotypic tests used in this study were as described previously (Cove, 1966 ; Tomsett & Cove, 1979 ; Garde et al., 1995 ). The niaD mutations created by Garde et al. (1995) are referred to here as follows: niaD3004 (niaDE850P), niaD3005 (niaDEA850/1PP), niaD3010 (niaDH654A). The mutations created in this study (niaD3000, 3001, 3002, 3006, 3007, 3008, 3009) are defined in Fig. 1.



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Fig. 1. Comparison of A. nidulans wild-type (GenBank accession no. M58291), mutant and revertant niaD gene sequences and the resulting NaR amino acid sequence with the equivalent sequences from corn (AF153448) and N. crassa (X61303.1). Sequences are shown only in the region of interest. The amino acid sequence is shown in upper-case letters and the DNA sequence in lower case. The DNA and amino acid sequence which is altered from the wild-type is shown in bold, and the sequence which is altered in the revertants with respect to the original mutation is underlined. The gap between the two mutation sites is denoted by two slashes.

 
Site-directed mutagenesis and vector construction.
PCR was carried out on a cloned fragment of the niaD gene using a mutagenic primer (i.e. containing mismatch bases) to introduce two mutations: T813D, K814Q. This PCR product was eluted from an agarose gel and combined with a second PCR product by recombinant PCR (Vallette et al., 1989 ) to create a 472 bp mutant fragment. This fragment was cloned into three vectors (pUC/niaD/KK, pUC/niaD/KK/E850P and pUC/niaD/KK/EA850/1PP), each containing a KpnI fragment from the niaD gene, one wild-type and two containing mutations created by Garde et al. (1995) . The PCR fragments were cloned in at NcoI and SacII sites. The KpnI fragments were subcloned into the transformation vector pUC/5'niiA/niaD (Garde et al., 1995 ). All cloning was carried out using the methods of Sambrook et al. (1989) . To ensure that no PCR errors were introduced into the gene and to check for the presence of the desired mutations, the area of interest was sequenced using an ABI 373 automated sequencer (Applied Biosystems) using the manufacturer’s instructions.

Protoplast preparation and transformation.
Protoplasts of A. nidulans strain niiA niaD{Delta}503 (pabaA1) (Tomsett & Cove, 1979 ) were made by the method of Johnstone et al. (1990) , including modifications of Garde et al. (1995) . DNA (5 µg) was added to 100 µl protoplasts in a 0·4 mm electroporation cuvette and incubated on ice for 1 min. The protoplasts were then transformed by electroporation (Garde et al., 1995 ). The protoplast suspension was plated onto supplemented minimal medium (Cove, 1966 ) containing 1·2 M sorbitol and 10 mM potassium nitrite to select for niiA+ transformants. Transformant homokaryons were purified twice by streaking single colonies onto selective medium. DNA was isolated using the method of Tilburn et al. (1983) . The genomic DNA was cut with SalI, and Southern blotting (Southern, 1975 ) was performed onto ZetaProbe (Bio-Rad) membrane following the manufacturer’s protocol to ensure that only single-copy transformants were studied.

Enzyme assays.
Cultures (200 ml) were inoculated with 2x106 fresh conidia and grown by shaking for 17 h at 30 °C in liquid minimal medium with supplements and 10 mM urea as the nitrogen source. Induced cultures also contained 20 mM potassium nitrate. The mycelium was strained, washed through muslin and cell-free extracts produced by the method of MacDonald & Cove. (1974) . The mycelium was extracted in fresh 0·1 M sodium phosphate buffer pH 7·2, 5 mM EDTA, 5 mM cysteine, 10% (v/v) glycerol for the nitrite reductase assay, and 0·1 M sodium phosphate buffer pH 6·8, 1 mM ß-mercaptoethanol, 0·5 mM EDTA, 1% (w/v) NaCl for the NaR assay and its partial activity assays. Nitrite reductase activity was measured as the oxidation of NADPH by a decrease in absorbance at 340 nm over 4 min as described by Pateman et al. (1967) . NaR activity was measured as the production of by an increase in absorbance at 340 nm according to Garrett & Cove (1976) . Reduced methyl viologen and bromophenol blue partial activities of NaR were measured as described by Garrett & Nason (1969) and Campbell (1986) , respectively. NaR was assayed in triplicate and nitrite reductase in duplicate; all assays were carried out over two or more reaction times that gave a linear response. The glucose-6-phosphate dehydrogenase (G6PD) assay was adapted slightly from Hankinson & Cove (1974) . Protein was extracted as previously described in 50 mM Tris/HCl pH 7·4. The assay mixture, made up in 1 ml quartz cuvettes, contained 50 µl glucose 6-phosphate (disodium salt), 50 µl 12 mM NADP+ (disodium salt) and 50 or 20 µl of cell-free extract, and was made up to 1 ml with Tris/HCl pH 8·0. In each case it was checked that there was negligible activity before glucose 6-phosphate was added, then the initial rate at 340 nm was determined. The activity was measured as the rate of reduction of NADP+ to NADPH. Protein concentrations were measured using the Bradford (1976) method.

UV-induced reversion of A. nidulans conidia.
Conidial suspensions [108 conidia ml-1 in a 1% (v/v) Tween 80 solution] were exposed to 1200 J m-2 UV radiation (to achieve 5% survival) whilst shaking under a UV lamp, and then plated onto appropriate nitrate-selective media allowing only revertant colonies to grow. DNA was extracted from revertants (as above) and a region of niaD was amplified by PCR and sequenced using an ABI 370 automated sequencer (Applied Biosystems) using the manufacturer’s recommended protocol.


   RESULTS AND DISCUSSION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
REFERENCES
 
Production and characterization of mutants with altered pyridine nucleotide specificities
Alignment of the sequence of A. nidulans NaR with other nitrate reductases indicated that T813 and K814 are in equivalent positions to those residues (D205/Q206 in corn leaf; S920/R921 in N. crassa) implicated in conferring NADH/NADPH specificity (Lu et al., 1994 ; Shiraishi et al., 1998 ). Three mutant genes were created by site-directed mutagenesis and subcloning (Fig. 1). The first, niaD3000, incorporated the putative NADH-specific ‘DQ’ sequence from corn. The other two, niaD3001 and niaD3002, combined these mutations with changes to the prolines described earlier (Friemann et al., 1991 ; Garde et al., 1995 ; Shondorf & Hachtel, 1995 ). The plasmids were transformed into A. nidulans and the genomic DNA of a single transformant from each transformation was extracted and Southern-blotted to ensure each contained a single homologous integration of the pUC5'niiA/niaD plasmid.

To characterize these mutants, the activities of a series of enzymes were measured: NADPH:NaR and NADH:NaR for activity of the holoenzyme; partial activities for mutant NaR using the artificial electron donors bromophenol blue and methyl viologen, which transfer electrons to the cytochrome and molybdopterin regions of the protein, respectively (see Solomonson & Barber, 1990 ); and NADPH nitrite reductase, the second enzyme of the pathway which is co-regulated with NaR. As shown in Table 1, a very low background NADPH:NaR activity was detected in all single or double mutant strains (niaD3000, 3001, 3002) containing the T813D, K814Q substitutions. Bromophenol blue:NaR activity levels in these mutants were normal, indicating that the mutation affected only the CbR region of the enzyme. In contrast, the niaD3004 (E850P) and niaD3005 (E850P, A851P) mutants displayed wild-type levels of all enzyme activities, as described previously (Garde et al., 1995 ). All mutants were inducible for NADPH:NiR activity.


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Table 1. Enzyme activities of wild-type, mutant and revertant strains

 
NADH:NaR was also measured to detect whether the co-enzyme specificity had altered. Like the wild-type, the niaD3004 and niaD3005 mutants had essentially no NADH:NaR activity. In all strains carrying the T813D, K814Q mutation, the level of activity had increased 40-fold relative to the wild-type NADH-dependent activity and was approximately 33% of induced NADPH:NaR activity in the wild-type. There was no significant difference between any of the single or double T813D, K814Q substitution strains, indicating that this mutation alone determines the observed alteration in co-enzyme specificity. This was consistent with the data of Shiraishi et al. (1998) , indicating that an acidic side-chain at this position is crucial for NADH specificity of NaR.

The phenotype of each mutant was tested by growth on a range of nitrogen source and chlorate media (data not shown). The niaD3004 and the niaD3005 mutants displayed wild-type phenotypes on all media as expected (Garde et al., 1995 ). However, all single or double mutant strains (niaD3000, 3001, 3002) containing the T813D, K814Q mutation were unable to utilize nitrate as sole N source and displayed wild-type sensitivity to chlorate. This inability to utilize nitrate cannot be due to lack of expression or reduced synthesis of the NiaD protein because significant NADH:NaR activity was detected in vitro. The specific activity of the NADH:NaR in these strains [39–45 nmol NO2 min-1 (mg protein)-1] was more than sufficient to support growth. Garde et al. (1995) demonstrated that the niaD3010 (H654A) mutant grew normally on nitrate as sole nitrogen source, despite having an NADPH:NaR with a specific activity of only 28 nmol NO2 min-1 (mg protein)-1. For this reason, these strains were not tested by Northern or Western analysis. This suggests that the lack of growth on nitrate occurs for other reasons: a metabolic effect in vivo, for example NADPH/NADH competition or insufficient NADH pools to drive nitrate assimilation in addition to other cellular reactions; a conformational effect in vivo which reduces catalytic efficiency, for example the pyridine nucleotide pocket for NADH is not optimal, but in vitro this is overcome by high concentrations of NADH; a regulatory effect, for example post-transcriptional inactivation/phosphorylation; or a toxicity effect arising from increased levels of NADPH in the mutant background.

Is lack of growth on nitrate due to NADPH competition in vivo?
If the mutant enzyme was binding but not oxidizing NADPH in vivo, NADH may be excluded; whereas in standard in vitro assays, NADPH concentrations would be too low. This hypothesis was tested in a competition assay. The mutant extracts were tested for activity in comparison with the wild-type, both with a varying amount of NADPH against a fixed amount of NADH and vice versa. In both strains, activity decreased slightly as the total NADPH/NADH concentration increased, but there was no evidence for competition (data not shown).

Dunn-Coleman & Pateman (1977) have shown that whilst NADPH protects wild-type NaR from deactivation in cell-free extracts, NADH does not. Using similar protection assays (Table 2), the half-life of the wild-type enzyme was >42 min when incubated with NADPH, but was around 8 min in the absence of any reduced cofactor or in the presence of NADH. However, all of the mutant enzymes had half-lives of 3–5 min in the presence and absence of NADPH and NADH. This indicates that NADPH is specific for protection of NaR against deactivation and that NADH cannot substitute in the mutant NADH-specific NaR. The mechanism of this protection is unknown. If catalytic, it is possible that a conformational change results from binding of the phosphate group of NADPH to the amino acids T813 and K814. However, a regulatory response triggered by NADPH oxidation is also possible.


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Table 2. Half-lives (min) of wild-type and mutant enzymes when incubated in the presence and absence of NADPH and NADH

 
Since NaR and NAD(P)H have been implicated in the regulation of the PPP (see Cove, 1979 ), these mutants were tested for G6PD activity (Fig. 2). All strains showed induction. The increase in activity of G6PD was about threefold for each strain. This would appear to rule out an effect of the NADPH/NADP+ ratio in the regulation of the PPP since the T813D, K814Q (niaD3000, 3001, 3002) NaR enzyme cannot oxidize NADPH, and since the mutant does not grow on nitrate, there is insufficient NADPH oxidation from NADPH:NiR. Rather, it suggests that nitrate induces the PPP via the nirA gene product, in much the same way as the enzymes of nitrate assimilation are induced.



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Fig. 2. Nitrate-induced ({blacksquare}) and uninduced ({square}) activities of G6PD in wild-type and T813D, K814Q substitution mutants. Activity is expressed in nM NADP+ reduced min-1 (mg total protein)-1. Data are means and standard deviations of three separate experiments.

 
Reversion analysis of niaD3001
As the T813D, K814Q mutants did not grow on nitrate, it was possible to select UV-induced revertants by growth on nitrate as the sole nitrogen source. These could arise from mutations within the niaD gene which optimize the interaction of NADH with the enzyme or restore NADPH:NaR activity. Alternatively, mutations could be selected in other genes which, for example, increased the available pool of NADH in vivo or disengaged a regulatory inactivation of the enzyme. Fourteen revertants of the niaD3001 mutant were isolated and their niaD gene was sequenced in the region spanning the primary mutations. In growth tests, all were indistinguishable from the wild-type (data not shown).

All revertants contained nucleotide alterations (mostly single base changes) which substituted the single amino acid residue D813; an acidic residue at this position is thus responsible for elimination of growth on nitrate. No alterations occurred at the Q814 site or at P850. Four classes of revertant were identified (Fig. 1); all amino acid substitutions replaced the negatively charged aspartic acid residue with an uncharged amino acid (I, V, G, N). This further supports the view that it is principally the presence of an acidic residue at this position that provides specificity to NADH:NaR (Shiraishi et al., 1998 ). Conclusive proof will require the reciprocal experiment on an NADH-specific NaR to show that replacement of the acidic residue by a small neutral residue also destroys NADH specificity (Shiraishi et al., 1998 ).

In enzyme assays, none of the revertants had NADH activity, one from each class was analysed in detail (Table 1). Substitution of the aspartic acid residue restored NADPH:NaR activity to levels capable of supporting normal growth: asparagine 65% of wild-type; glycine 42%; isoleucine 70%; valine 47%.

Reversion of the niaD3000 and niaD3002 mutants produced no new classes (data not shown). The reversion frequencies for each strain were almost identical (data not shown), suggesting that the number of possible reversions were the same for each mutation; i.e. the alterations at E850 and A851 did not affect the reversion frequency and hence were not involved in preventing enzyme activity or restoring it.

The NAD(P)H specificity of NaR
Site-specific mutagenesis demonstrated that the T813D, K814Q mutation alters co-enzyme specificity and that the E850P and A851P substitutions had no effect, even in a niaD3000 background. This would seem to contradict the evidence from birch (Shondorf & Hachtel, 1995 ). The removal of the acidic side-chain at position 813 was sufficient to eliminate NADH as electron donor and restore NADPH-dependent activity. Since a series of isolates all had this change, this suggests that either a more complex change is required to mutate niaD to increase the efficiency of NADH utilization, or that lack of growth on nitrate is a metabolic or regulatory control (see below). The fact that the mutant enzymes had significant NADH:NaR in vitro suggests that although subsequent alterations to the CbR region of this enzyme may be able to further optimize the NaR/NADH interaction, perhaps through modification to the size/shape of the binding pocket, these changes may not be detected in the reversion analysis of niaD3001 as they would not restore growth. The lack of growth is probably not due to a lack of NADH:NaR activity but is most likely caused by other factors (see below).

Site-directed mutagenesis of the equivalent sequence has been undertaken on N. crassa NADPH:NaR to examine pyridine nucleotide specificity. Gonzalez et al. (1995) showed that a serine to alanine (S920A) substitution reduced NADPH:NaR activity by 40% when transformed back into N. crassa, whilst a S920T substitution increased activity by 75%. The latter observation is not surprising since A. nidulans has threonine at this position (T813), except that it indicates that N. crassa may have evolved a suboptimal NaR activity.

Is lack of growth on nitrate due to NADH limitation or altered post-transcriptional regulation?
The strains carrying the NADH:NaR enzyme do not grow on nitrate, which would not be expected from the enzyme assay results. It has been shown that mutants with 20% wild-type NADPH-dependent activity still grow well on nitrate (Garde et al., 1995 ). It could be that NADH is, for some reason, not available to NaR in vivo; however, this seems unlikely as it has been demonstrated in A. niger that NADH is abundant in cells and that levels are maintained even when it is in demand (Fuhrer et al., 1980 ). It would therefore seem likely that whilst the mutant enzyme is able to bind and oxidize NADH in cell-free extracts, in vivo conditions prevent this.

NADH and NADPH have also been implicated in the post-translational regulation of NaR activity in a number of different organisms (Minagawa & Yoshimoto, 1982 ; Dunn-Coleman & Pateman, 1977 ; Vega et al., 1975 ; Solomonson et al., 1973 ). The mechanism by which NADPH stabilizes NaR is not known but it would seem likely that the binding of the co-enzyme induces a conformational change that protects the enzyme from cellular factors that deactivate it. Dunn-Coleman & Pateman (1977) showed that the deactivation is reversible, so it would seem likely that the deactivation occurs via a phosphorylation event rather than by proteolysis. Lillo & Ruoff (1992) , working on Cucurbita maxima, suggested that this deactivation may occur at a second binding site, as the protection could occur with NADH or NADPH, whilst only NADH could bind to the catalytic binding site of the enzyme. This work presents evidence that protection occurs via binding of NADPH to the catalytic binding site as disruption of this site prevents NADPH protection. Also, the fact that the mutant enzymes have a shorter half-life than the wild-type in the absence of reducing cofactor would suggest that the mutation must affect the inherent stability of the enzymes.

Could increased levels of NADPH in the mutant background be toxic?
The results shown in Fig. 2 demonstrated that the PPP was induced in the mutant strains just as in the wild-type; however NADPH, its product, would not have been oxidized by the altered NaR enzyme. It is thus possible that increased levels of NADPH in these strains could be toxic and hence could be responsible for the inability of the mutants to grow on nitrate. However, the mutants had a nitrogen-starved morphology when grown on nitrate as the sole nitrogen source, perhaps suggesting that NADPH toxicity was not the cause of this phenotype. In an attempt to test for NADPH toxicity, a niaD3001 mutant was crossed into a strain carrying the pppA1 mutation in the G6PD enzyme of the PPP (Hankinson & Cove, 1974 ). Preliminary data suggested that the double mutant showed slightly increased growth on nitrate compared to the niaD3001 mutant alone (data not shown). This suggested that whilst NADPH may be slightly toxic, it is not the principal factor causing the inability of the niaD3001 mutant to grow on nitrate.

This study has demonstrated the importance of functional genomic analysis in vivo; investigations of an enzyme function must analyse changes to the physiology of the organism within which it has evolved.


   ACKNOWLEDGEMENTS
 
This work was supported by a BBSRC studentship to N.H. We wish to thank Bill Campbell and Mark Caddick for many useful discussions throughout the course of this work.


   REFERENCES
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ABSTRACT
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METHODS
RESULTS AND DISCUSSION
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Received 5 November 1999; revised 23 February 2000; accepted 9 March 2000.



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