Enzymatic oxidation of NADP+ to its 4-oxo derivative is a side-reaction displayed only by the adrenodoxin reductase type of ferredoxin-NADP+ reductases


A. Aliverti, Dipartimento di Scienze Biomolecolari e Biotecnologie, via Celoria 26, 20133 Milano, Italy
Fax: +39 02 50314895
Tel: +39 02 50314897
E-mail: alessandro.aliverti@unimi.it
Website: http://www.sbb.unimi.it/index.htm


We have previously shown that Mycobacterium tuberculosis FprA, an NADPH-ferredoxin reductase homologous to mammalian adrenodoxin reductase, promotes the oxidation of NADP+ to its 4-oxo derivative 3-carboxamide-4-pyridone adenine dinucleotide phosphate [Bossi RT, Aliverti A, Raimondi D, Fischer F, Zanetti G, Ferrari D, Tahallah N, Maier CS, Heck AJ, Rizzi M et al. (2002) Biochemistry41, 8807–8818]. Here, we provide a detailed study of this unusual enzyme reaction, showing that it occurs at a very slow rate (0.14 h−1), requires the participation of the enzyme-bound FAD, and is regiospecific in affecting only the C4 of the NADP nicotinamide ring. By protein engineering, we excluded the involvement in catalysis of residues Glu214 and His57, previously suggested to be implicated on the basis of their localization in the three-dimensional structure of the enzyme. Our results substantiate a catalytic mechanism for 3-carboxamide-4-pyridone adenine dinucleotide phosphate formation in which the initial and rate-determining step is the nucleophilic attack of the nicotinamide moiety by an active site water molecule. Whereas plant-type ferredoxin reductases were unable to oxidize NADP+, the mammalian adrenodoxin reductase also catalyzed this unusual reaction. Thus, the 3-carboxamide-4-pyridone adenine dinucleotide phosphate formation reaction seems to be a peculiar feature of the mitochondrial type of ferredoxin reductases, possibly reflecting conserved properties of their active sites. Furthermore, we showed that 3-carboxamide-4-pyridone adenine dinucleotide phosphate is good ligand and a competitive inhibitor of various dehydrogenases, making this nucleotide analog a useful tool for the characterization of the cosubstrate-binding site of NADPH-dependent enzymes.


adrenodoxin reductase

Amplex Red



atomic mass units


charge transfer complex between FprA and NADPH


ferredoxin-NADP+ reductase


iodo-nitro-tetrazolium chloride


3-carboxamide-4-pyridone adenine dinucleotide phosphate


3-carboxamide-4-pyridone mononucleotide



Ferredoxin-NADP+ reductases (FNRs, EC can be classified into two phylogenetically distinct subgroups: the mitochondrial-type and the plastid-type (or plant-type) enzymes [1]. The prototype of the former enzymes is the mammalian adrenodoxin reductase (AdR), whereas the latter subgroup is best exemplified by the photosynthetic FNR. Although both FNR types catalyze the same physiologic reaction, i.e. the transfer of a couple of electrons from NADPH to two successive ferredoxin molecules, they markedly differ in their structure and functional properties [2,3].

In the recent past, we have obtained the crystal structure of FprA, a Mycobacterium tuberculosis homolog of AdR (41% identity spread over the entire polypeptide length) at a very high (1.05 Å) resolution [4]. This accomplishment, together with the fact that the structures of AdR and FprA are very similar, made the bacterial protein an ideal representative of mitochondrial-type FNRs for structure–function relationship studies. The atomic resolution of the FprA structure allowed us to discover that this enzyme, in addition to the well-known NADPH-dependent ferredoxin reduction, catalyzed an unprecedented reaction, i.e. the oxidation of NADP+ to yield its 4-oxo derivative, which was named 3-carboxamide-4-pyridone adenine dinucleotide phosphate (NADPO). The first evidence that such a reaction was occurring was the observation that crystals of FprA grown in the presence of NADP+ contained NADPO bound to the active site instead of NADP+[4]. Comparison of the FprA–NADPO and FprA–NADPH structures clearly showed that in the latter complex an additional ordered water molecule (water 1) was sitting in the active site in a position very close to that occupied by the carbonyl oxygen atom of NADPO in the former complex [4]. This observation prompted us to propose the hypothetical mechanism for the FprA-catalyzed NADPO formation reaction depicted in Fig. 1. NADP+ oxidation is initiated by water 1 addition to the electron-poor C4 atom of the nicotinamide ring of NADP+. His57 and Glu214 have been previously proposed to increase the nucleophilicity of the water molecule by favoring its deprotonation [4]. Both residues, highly conserved in AdR-like enzymes, have been recently changed to nonionizable ones by site-directed mutagenesis [5]. Characterization of the resulting FprA variants allowed us to conclude that His57 but not Glu214 played a significant role in the physiologic, NADPH-dependent activity of FprA. However, crystals of FprA-H57Q grown in the presence of NADP+ again displayed NADPO as the bound nucleotide by X-ray diffractometry [5], showing that the H57Q mutation did not abolish the NADP+ oxidation activity of FprA.

Figure 1.

 Hypothetical mechanism of the reductive half-reaction of the catalytic cycle of FprA in the oxidation of NADP+ to yield NADPO. The reaction scheme was based on the crystal structures of the FprA–NADPO and FprA–NADPH complexes [4]. The depicted water molecule is referred to in the text as water 1. B1 and B2 represent hypothetical groups acting as base catalysts. B1 has been proposed to be the imidazyl of His57 [4].

The previous studies summarized above only gave a qualitative evidence of the ability of FprA to catalyze the production of NADPO. In the present article, we provide the first quantitative description of this reaction, along with a detailed analysis of the spectroscopic properties of the product of NADP+ oxidation.

Results and Discussion

NADPO isolation, quantitation, and spectral characterization

In order to study the kinetics of NADP+ oxidation to NADPO catalyzed by FprA, an NADPO assay method was required. After testing different analytical chromatographic procedures, we found the ion exchange method described by Orr & Blanchard to be particularly reliable and robust for our purposes [6]. Inclusion of AMP as an internal standard increased precision and accuracy, and replacement of the original mobile phase with a volatile ammonium formate buffer allowed the recovery of purified nucleotide as salt-free preparations after vacuum drying. Figure 2 shows the typical analysis of aliquots sampled at increasing incubation times from a reaction mixture where FprA was incubated with NADP+ in air. As can be seen, incubation over several hours resulted in a gradual decrease of the NADP+ peak while a single new peak appeared and progressively increased in intensity. By tandem MS, the latter peak was found to unambiguously contain an NADP derivative bearing an oxygen atom bound to the nicotinamide ring. Previously, the same NADP+ modification was observed by MS analysis of the whole reaction mixture [4]. However, even though only 4-oxo-NADP was observed by X-ray crystallography in complex with FprA, in principle it cannot be excluded that FprA could also introduce an oxygen atom at other positions of the nicotinamide ring of NADP+, leading to other nucleotide derivatives with a lower affinity for the enzyme active site. Thus, it was of interest to ascertain the identity of the isolated compound by a detailed spectroscopic characterization. Values of 17 100 and 15 600 m−1·cm−1 were found for the extinction coefficient of the NADP derivative at 260 and 254 nm, respectively, as calculated by determining the concentration of the nucleotide on the basis of the phosphate released by alkaline phosphatase treatment. The absorbance spectrum of the nucleotide is shown in Fig. 3A in comparison to those of NADP+ and NADPH. A peculiar feature of the modified NADP+ is a relatively high absorption in the 280–310 nm region. To study the effect of pH on its spectral properties, the adenylate portion of the modified nucleotide was removed by enzymatically splitting the pyrophosphate link in order to get rid of its large absorbance contribution. As shown in Fig. 3B, the spectrum of chromatographically purified NMN derivative undergoes a large spectral transition when the pH is decreased from 7.7 to 1.0. The two spectra are very similar to those described for N-methyl-4-pyridone-5-carboxamide, and completely different from those of N-methyl-2-pyridone-5-carboxamide, under similar pH conditions [7]. Furthermore, the isolated NADP derivative was found to lack any fluorescence when excited with light in the UV region. This observation is relevant for excluding the presence of species modified in position 6 of the nicotinamide ring, because, unlike the compounds with the oxo group in positions 2 and 4, N-methyl-6-pyridone-5-carboxamide has been shown to emit blue light by fluorescence [8]. We exclude the formation of the species modified in position 3 of the nicotinamide ring because of the poor reactivity of this position towards nucleophilic attack. Indeed, the positive charge of the pyridinium moiety favors attack by nucleophiles at positions 2 and 4 under mild conditions [9]. Furthermore, and more importantly, the modification at position 3 can be excluded because the resulting N-substituted 3-oxo-nicotinamide moiety would not be neutral, resulting in a dinucleotide with spectral and chromatographic properties expected to be markedly different from those of 4-oxo-NADP. These data allow us to conclude that FprA does not produce detectable amounts of NADP derivatives bearing a carbonylic oxygen at sites other than position 4 of the nicotinamide ring, indicating that the NADP+ oxidation reaction is highly regiospecific.

Figure 2.

 Anion exchange chromatographic pattern of NADP+ oxidation reaction mixtures. NADP+ at 500 µm was reacted with O2 at its air equilibrium concentration (c. 250 µm) in the presence of 100 µm FprA at 25 °C. Forty-microliter aliquots were analyzed by high-performance chromatography as described in Experimental procedures. Reaction times were 0 h (dotted trace), 3 h (dashed trace), 6 h (dot-dashed trace), and 24 h (continuous trace).

Figure 3.

 Extinction coefficients and absorption spectra of NADPO and NMNO. (A) Dinucleotides in 20 mm Tris/HCl (pH 7.7). (B) NMNO in 20 mm Tris/HCl (pH 7.7), and after bringing the pH to 1 by the addition of HCl.

Kinetics of NADPO formation as catalyzed by FprA and AdR

Figure 4A shows the time courses of NADP+ oxidation to NADPO catalyzed by FprA or AdR in the presence of air oxygen. Clearly, both enzymes are able to catalyze this reaction. Studies at various NADP+ concentrations were performed, showing that the substrate concentration of 500 µm was fully saturating. The initial rate of NADPO formation promoted by AdR was slightly lower (75%) than that of FprA under the same conditions. A peculiar feature of the NADP+ oxidation kinetics is the progressive decrease in the reaction rate. When FprA was the catalyst, this behavior was particularly marked: NADPO formation sharply decreased after the first enzyme turnover. We excluded the possibility that this was a consequence of enzyme denaturation by assaying the enzyme during incubation. A reasonable explanation is that NADPO was acting as a competitive inhibitor of the enzyme with respect to NADP+ (see below). In the case of FprA, the reaction was studied at different enzyme concentrations. As shown in Fig. 4B, the initial rate of the reaction was proportional to FprA concentration, showing that the NADP+ oxidation, although very slow, was strictly enzyme-dependent. The NADP+ oxidation reaction catalyzed by FprA was much slower (0.14 h−1) than that of its NADPH-dependent, physiologic reaction (336 min−1 when Mycobacterium smegmatis FdxA was used as electron acceptor [10]).

Figure 4.

  (A) Time-courses of NADPO production catalyzed by FprA and AdR in the presence of O2. The reaction was carried out at 25 °C in the presence of 100 µm FprA (filled circles) or bovine AdR (open circles) and 500 µm NADP+. A control experiment was performed, omitting the enzyme (squares). (B) Initial rate of NADPO production calculated over the first 6 h of the reaction plotted as a function of the enzyme concentration.

Plant-type FNRs do not catalyze NADPO formation

To verify whether NADP+ oxidation to yield NADPO was a common feature of the members of the FNR class of enzymes, we assayed Toxoplasma gondii FNR [11], Plasmodium falciparum FNR [12] and spinach (Spinacia oleracea) leaf FNR [13] (which all are plant-type FNRs) for their ability to catalyze this reaction, and found them to be completely inactive. Although FNRs from other sources should be assayed before drawing general conclusions, it is suggested that NADP+ oxidation to NADPO most probably represents a unique feature of AdR-type FNRs and reflects very specific organization and reactivity of their active sites.

Role of O2 in enzyme-catalyzed NADP+ oxidation

When the O2-dependent NADPO production reaction catalyzed by FprA was allowed to proceed in the presence of excess peroxidase and its fluorogenic substrate 10-acetyl-3,7-dihydroxyphenoxazine (Amplex Red), a progressive increase in fluorescence emission at 585 nm was observed (not shown), indicating accumulation of the fluorescent product resorufin. No fluorescence built up in the absence of peroxidase, allowing us to attribute resorufin formation entirely to the 1 : 1 reaction between H2O2 and Amplex Red catalyzed by the peroxidase. This conclusion was confirmed by the effect of the presence of catalase in the reaction mixture, which completely abolished resorufin formation. On the basis of the quantum yield experimentally determined for resorufin, a rate of 0.08 mol H2O2 (mol FprA)−1 was calculated, a value comparable to the rate of NADPO formation. In addition to H2O2, a small amount of superoxide was apparently produced in the reaction, as judged from the slight increase in the rate of fluorescence appearance after superoxide dismutase addition. However, it could not be excluded that the peroxide/superoxide ratio was substantially lower than that found, as the low rate of the reaction would allow enough time for spontaneous disproportionation of O2 to H2O2 and O2. In any case, the production of H2O2 and/or O2 strongly supports the FAD prosthetic group as the direct oxidant of the nicotinamide ring, as these species are the usual products of FADH2 reoxidation by O2 in most flavoproteins.

To verify whether the direct reaction between NADP+ and O2 was not required in the enzymatic production of NADPO, the latter reaction was studied in the presence of electron acceptors different from O2. NADP+ was incubated with FprA under anaerobic conditions in the presence or in the absence of K3Fe(CN)6, an artificial electron acceptor of the enzyme. As shown in Fig. 5A, as NADP+ was added to the reaction mixture, a progressive bleaching of the absorbance contributed by ferricyanide was observed, indicating its reduction to ferrocyanide. The spectrum of enzyme-bound FAD remained unaltered, until all the ferricyanide was consumed. Starting from this point, the FAD spectrum undergoes a progressive perturbation (not shown), similar to that observed by incubating FprA with NADP+ in the absence of ferricyanide (Fig. 5B). The ability of FprA to carry on NADP+ oxidation using either O2 or ferricyanide supports our previous proposal, made on the basis of structural data [4], that the mechanism of NADPO formation can be split in two half-reactions: the first leads to NADPO formation coupled to FAD reduction; and the second consists of the reoxidation of FADH2 by O2 or other oxidants (Fig. 1). With the aim of trapping possible intermediates produced in the first half-reaction, the reaction between NADP+ and FprA was studied in the absence of any oxidant, thus preventing the second, oxidative, half-reaction of the catalytic cycle. During incubation, the A340 of the mixture progressively increased, and the visible absorbance spectrum of the enzyme-bound FAD underwent a progressive partial bleaching, leading to a final stable spectrum strongly reminiscent of that of the charge transfer complex between NADPH and oxidized FprA [10]. The time course of the FAD spectral change approximately followed a single exponential decay equation (inset of Fig. 5B) with a first-order rate constant of 0.095 ± 0.009 h−1, a value similar to the rate of NADPO formation measured under aerobic conditions. The enzyme present in the endpoint reaction mixture was denatured and precipitated before admitting air into the anaerobic cuvette, and the enzyme-free solution was analyzed by anion exchange chromatography as previously described. The sample was found to contain both NADPO and NADPH. Thus, in the absence of oxidants, FprA catalyzes the disproportionation of NADP+ according to the following equation:

Figure 5.

 Spectral changes resulting from the incubation of FprA with NADP+ in the presence and absence of potassium ferricyanide under anaerobic conditions. (A) FprA at c. 20 µm was incubated with 200 µm NADP+ and 140 µm K3Fe(CN)6 at 25 °C in 20 mm Hepes/NaOH (pH 7.0), containing 100 mm NaCl and 10% glycerol, within a gas-tight cuvette made anaerobic by several vacuum–N2 flushing cycles. Reaction times were 0 min to 330 min. (B) Same conditions as in (A), with the omission of K3Fe(CN)6. Reaction times were 0 h to 19 h. Inset: absorbance at 580 nm as a function of incubation time. The curve represents the best fit according to a single exponential decay equation (k = 0.095 h−1).

As FprA has a single binding site for NADPH [4,5], the reaction must proceed through a ping-pong mechanism, with the transferred electron couple being transiently stored on the enzyme prosthetic group. Thus, experimental evidence points to the involvement of FAD in the mechanism of NADPO formation. The observed spectral changes can be interpreted as resulting from the following reaction mechanism, where E(FAD) and E(FADH) represent the oxidized and fully reduced form of FprA, respectively:


In the reaction shown in Eqn (4), the FprA–NADPH complex represents a charge transfer species (CT1) [10]. The reactions shown in Eqn (2) and Eqn (4) are expected to be much faster than that shown in Eqn (3), as dinucleotide binding and release to and from FprA have never been found to be limiting steps in reactions of the enzyme with NADPH or NADH, which occurred at rates much higher than that of NADPO formation [5]. Thus, the only process accessible to experimental observation was the conversion of the E(FAD)–NADP+ complex to CT1, precluding any further dissection of the reaction mechanism by time-resolved spectral analysis. The fact that CT1 formation follows single-phase kinetics without the formation of any observable transient intermediate suggests that water molecule addition to the nicotinamide moiety (Fig. 1) might be the limiting step of the whole reaction.

Investigating the role of Glu214 and His57 of FprA in the catalysis of NADPO production

As it was not possible to obtain detailed information on the catalytic mechanism of the NADP+ oxidation reaction by kinetic studies, we attempted to gain insights into the role of specific residues of FprA by site-directed mutagenesis. On the basis of structural data, Glu214 and His57 of FprA have been proposed to be involved in promoting nucleophilic attack by a water molecule on the C4 of the nicotinamide moiety of NADP+[4]. The production and purification of the enzyme variants, where these residues were replaced with Ala or Gln, have been described elsewhere [5]. The physiologic NADPH-dependent activity of the mutant FprA forms was characterized in detail. Unlike Glu214, which had essentially no effect on this activity, His57 turned out to decrease by 4–5-fold the hydride transfer rate from NADPH or NADH to the enzyme-bound FAD [5]. Here, we have assayed FprA-E214A and FprA-H57Q for their ability to catalyze NADP+ oxidation to NADPO. As shown in Fig. 6, both mutant forms supported the production of NADPO, with time-courses similar to that of the wild-type enzyme. It is noteworthy that both single mutations slightly increased the initial rate of NADPO synthesis. At 500 µm NADP+, the reaction rates were 0.21 h−1 and 0.19 h−1 for FprA-E214A and FprA-H57Q, respectively. If the rate-determining step of NADPO production is water addition, the proposed activating role of Glu214 and His57 should be dismissed. The crystal structure of the complex between FprA-H57Q and NADPO has been obtained at 1.8 Å resolution [5]. Slight alterations in the positioning of the nicotinamide ring in the active site have been observed by comparing the mutant with the wild-type enzyme. The described 0.7 Å shift of the nicotinamide moiety could represent the structural basis of the observed small increase in NADP+ oxidation rate observed with this mutant.

Figure 6.

 Time-courses of NADPO production catalyzed by wild-type and mutant forms of FprA. The reaction was carried out at 25 °C in the presence of 100 µm wild-type FprA (filled circles), FprA-E214A (open circles) or FprA-H57Q (squares) and 500 µm NADP+ in air-equilibrated buffer.

NADPO as an inhibitor of FNRs

To the best of our knowledge, this is the first time that enzymatic oxidation of NADP+ has been reported and the spectral properties of the resulting NADP derivative have been described. It was thus of interest to provide a first characterization of NADPO as a possible probe for studying the NADPH-binding site of dehydrogenases. By inhibition studies under steady-state conditions, we have found that NADPO acts as a competitive inhibitor with respect to NADPH on various FNRs (both photosynthetic and nonphotosynthetic), with Ki values ranging between 1 and 30 µm (Table 1). FprA is the enzyme most strongly inhibited by this nucleotide. To better characterize the interaction of NADPO with FprA, enzyme–ligand binding was studied by difference spectrophotometry using various nucleotides: NADPO, NADP+, thio-NADP+ and 2′-phospho-AMP (2P-AMP). The Kd determined for the complex between the enzyme and NADPO was slightly but significantly lower than that measured for NADP+ (Table 2). As 2P-AMP, which lacks the NMN portion of NADP+ but still strongly binds to FprA, yielded very weak perturbations of the absorption spectrum of the enzyme, it is clear that the intense difference spectra of the FprA–dinucleotide complexes are due to the alteration of the isoalloxazine microenvironment induced by the binding of the NMN group. The small structural differences in the nicotinamide rings of NADP+, NADPO and thio-NADP+ resulted in small but significant differences in the corresponding difference spectra (Table 2). In plant FNRs, we found a correlation between the intensity of the difference spectrum induced by binding and the degree of occupancy of the nicotinamide ring of the bound ligand in the active site [14]. In this view, it is significant that the perturbations induced by NADPO binding to FprA were more intense than those induced by the other dinucleotides tested. This suggests that the 4-pyridone-5-carboximide ring of NADPO is particularly well fitted to stack over the isoalloxazine ring, resulting in a higher occupancy than with dinucleotides carrying other pyridine derivatives. These observations support the hypothesis that accumulation of NADPO would substantially inhibit its own production by FprA, due to competition with NADP+ for binding to the enzyme active site.

Table 1.   Inhibition constants of NADPO for different FNRs. 2,6-Dichloroindophenol reductase activity was measured using either 62 nm FprA or 2.5 nmT. gondii FNR in 0.2 m Tris/HCl (pH 8.2) at 25 °C at a fixed concentration of 66 µm 2,6-dichloroindophenol; the NADPH concentration was varied between 0.1 and 10 µm in the case of FprA, and between 2 and 27 µm in the case of T. gondii FNR. The concentration of NADPO was varied between 0 and 20 µm. INT reductase activity was measured using 2.5 nm spinach leaf FNR in 0.2 m Tris/HCl (pH 9.0), 70 mm NaCl and 0.1% Triton X-100 at 25 °C at a fixed concentration of 100 µm INT; the NADPH concentration was varied between 2 and 27 µm; the NADPO concentration was varied between 0 and 20 µm. All assay mixtures included an NADPH-regenerating system comprising glucose 6-phosphate and glucose-6-phosphate dehydrogenase.
EnzymeDiaphorase reactioninline image
inline image
  • a

     Values reported in the table should be considered as ‘apparent’ kinetic parameters, as they were determined at a nonsaturating fixed concentration of the electron acceptor.

FprANADPH→2,6-dichloroindophenol0.2 ± 0.014.1 ± 0.051.2 ± 0.2
Spinach leaf FNRNADPH→INT5.0 ± 0.454 ± 125 ± 5
T. gondii FNRNADPH→2,6-dichloroindophenol3.5 ± 0.264 ± 1.530 ± 5
Table 2.   Affinities of various nucleotides for FprA and extent of the spectral perturbations induced by their binding to the enzyme.
LigandKdm)Δεa (mm−1·cm−1)
  • a

     Difference extinction coefficients at the wavelength indicated in parentheses were calculated by subtracting the absorbance of free FprA from that extrapolated at infinite ligand concentration.

2P-AMP3.6 ± 20.22 (489 nm)
NADP+6.3 ± 0.71.2 (499 nm)
Thio-NADP+2.0 ± 0.20.83 (496 nm)
NADPO3.0 ± 11.8 (494 nm)


In this article, we provide clear evidence that M. tuberculosis FprA and bovine AdR, but not plant-like FNRs, catalyze the FAD-dependent oxidation of NADP+ to NADPO. This enzyme activity, possibly shared by all AdR-like enzymes, is highly regiospecific, in that it targets only the 4-position of the pyridine ring of the substrate. The very low reaction rate tends to exclude a physiologic role for NADPO, although at least one example exists of an NADP derivative, i.e. nicotinic acid adenine dinucleotide phosphate, with documented signaling functions [15]. Rather, we feel that the NADP+ oxidation activity of AdR-like enzymes reflects a specific and conserved reactivity of their active sites, where a water molecule exerts considerable strain and possibly a polarizing effect on the C4 atom of the nicotinamide moiety of the bound substrate. The strict interaction between a zinc-bound water molecule and the nicotinamide ring has been suggested to have a role in activating NADH for efficient hydride transfer in the catalytic cycle of horse liver alcohol dehydrogenase [16]. It can be speculated that water 1 plays a similar role in the active site of AdR-type FNRs, i.e. it favors the hydride transfer between NADPH and FAD in the physiologic reaction catalyzed by these enzymes. When NADP+ is the enzyme ligand, the electrophilicity of the nicotinamide C4 promotes water 1 addition to the nicotinamide and subsequent FAD-dependent oxidation to give NADPO. It is interesting to note that no ordered water molecules are present in the proximity of the nicotinamide ring in the crystal structure of the complexes between plant FNR and NADP+ or NADPH [14]. This observation could provide a rationale for the lack of NADP+ oxidation activity in plant-type FNRs. Further work will be required to fully elucidate the actual role of active site water molecules in modulating the reactivity of NADP+ and NADPH bound to FprA.

Experimental procedures

Enzymes and chemicals

Calf intestine alkaline phosphatase was obtained from GE Healthcare (Milano, Italy). Crotalus durissus phosphodiesterase, beef liver catalase, superoxide dismutase from bovine erythrocytes and yeast glucose-6-phosphate dehydrogenase were all from Roche Diagnostics (Monza, Italy). Horseradish peroxidase was bought from Invitrogen (San Giuliano Milanese, Milano, Italy). Recombinant M. tuberculosis wild-type FprA was produced and isolated in two different molecular forms: without extra residues [10], and with an N-terminal poly-His extension [5]. The two enzyme forms were found to be indistinguishable in their functional properties. The site-directed mutants FprA-H57Q and FprA-E214A were obtained in poly-histidinylated form only [5]. Recombinant S. oleracea leaf FNR, T. gondii FNR and P. falciparum FNR were purified as described elsewhere [11,17,18]. Purified recombinant bovine AdR was a generous gift of R Bernhardt (Universität des Saarlandes, Saarbrücken, Germany). NADP+, NADPH, NAD+ and AMP were purchased from Sigma-Aldrich (Milano, Italy). Amplex Red was from Invitrogen. All other chemicals were of the highest possible grade.

Chromatographic separation of nucleotides

Variable volumes of enzyme reaction mixtures (see below) were treated with equal volumes of acetonitrile to denaturate and precipitate the protein. After centrifugation at 12 000 g for 10 min, the supernatants were dried, resuspended in 50 mm ammonium formate, and chromatographed by a modification of the high-performance ion exchange procedure described in Orr & Blanchard [6]. Using an ÄKTA FPLC apparatus (GE Healthcare), samples were loaded on a MonoQ HR 5/5 column (1 mL; GE Healthcare), equilibrated in the above volatile buffer. Nucleotides were separated at room temperature using a 50–600 mm ammonium formate gradient in 25 column volumes at a flow rate of 1 mL·min−1. The eluate was monitored continuously by measuring its absorbance at 254 nm. Fractions containing NADPO or 3-carboxamide-4-pyridone mononucleotide (NMNO) were dried under vacuum and stored at − 20 °C.


MS and MS/MS data were obtained using an LCQ ADV MAX ion trap mass spectrometer equipped with an ESI ion source and controlled by xcalibur software v.1.3 (Thermo-Finnigan, San Jose, CA, USA). ESI experiments were carried out in positive ion mode under the following constant instrumental conditions: source voltage 5.0 kV, capillary voltage 10 V, capillary temperature 250 °C, and tube lens voltage 55 V. MS/MS spectra obtained by collision-induced dissociation were performed with an isolation width of 2 Th (m/z), and the activation amplitude was around 35% of the ejection RF amplitude of the instrument, which corresponds to 1.58 V. ESI-MS of NADPO yielded ions at m/z 760.18 [MH]+ and 782.06, amu [MNa]+ ESI-MS/MS of the former ion yielded fragment ions at m/z 741.74 [MH–H2O]+, 624.71 [MH–adenine]+, 603.66 [MH−4-oxo-nicotinamide–H2O]+, 489.71 [MH−4-oxo-nicotinamide-ribose–H2O]+, and 329.86 [MH−4-oxo-nicotinamide-ribose-diphosphate–H2O]+, amu.

Determination of the absorption and fluorescence properties of NADPO and NMNO

All absorption and fluorescence emission spectra were recorded on a UV–visible 8453 diode array spectrophotometer (Agilent, Cernusco sul Naviglio, Milano, Italy) and a Cary Eclipse spectrofluorimeter (Varian, Leini, Torino, Italy), respectively. In order to determine its extinction coefficient, NADPO was quantified on the basis of the amount of the phosphate released by phosphatase treatment. NADPO at 10–20 nmol was incubated for 1 h with 0.25 units of alkaline phosphatase at 25 °C in 0.5 m Tris/HCl (pH 9.0), containing 10 mm MgCl2, to hydrolyze the 2′-phosphate group. Free phosphate content was determined by the method of Chen et al. [19]. Known amounts of NADP+ were used as controls, verifying the accuracy of the procedure. To isolate NMNO, c. 10 nmol of NADPO was treated with 2 µg of phosphodiesterase for 20 min at 25 °C in 20 mm Tris/HCl (pH 7.7). NMNO was then purified chromatographically as described above. The absorbance spectrum of NMNO was recorded both in 20 mm Tris/HCl (pH 7.7) and after adjusting the pH to c. 1 by the addition of HCl.

Monitoring of the time-course of NADPO formation catalyzed by various enzymes

The enzymatic conversion of NADP+ to NADPO was studied in both aerobic and anaerobic conditions. The aerobic reactions were carried out at 25 °C by mixing 50–150 µm enzyme with variable concentrations of NADP+, ranging from 150 µm to 10 mm, in 20 mm Hepes/NaOH (pH 7.0), containing 100 mm NaCl and 10% glycerol. At different incubation times, 40 µL aliquots were withdrawn, AMP was added as internal standard, and samples were analyzed by ion exchange chromatography as described above. The amount of NADPO was determined on the basis of peak integration data provided by unicorn 5 software (GE Healthcare), and the experimentally estimated ε254 value of 15.6 mm−1·cm−1 for NADPO. To monitor the reaction in the absence of molecular oxygen as the oxidant, c. 20 µm FprA in the same buffer as above, either in the absence or in the presence of 140 µm K3Fe(CN)6, was placed in an anaerobic cuvette, containing a concentrated NADP+ solution in the side arm to yield a final concentration of 200 µm. After anaerobiosis was established by successive cycles of N2-flushing and evacuation, reactants were mixed. Spectral changes were recorded over a period of several hours at 20 °C using a UV–visible 8453 diode array spectrophotometer (Agilent).

Identification of the reactive oxygen species produced in the reaction between NADP+ and O2 catalyzed by FprA

Air-equilibrated mixtures of 10 µm FprA and 0.5 mm NADP+ were incubated as described in the previous paragraph in the presence of 0.1 unit·mL−1 horseradish peroxidase and 100 µm Amplex Red. Peroxidase-catalyzed Amplex Red conversion to resorufin was monitored by measuring the fluorescence emission at 585 nm of the solution upon excitation at 571 nm. When superoxide dismutase and catalase were present, the concentrations were 0.5 µg·mL−1 and 1 µg·mL−1, respectively.

Enzyme activity assays

Assays of the NADPH-dependent catalytic activities of FprA, spinach leaf (S. oleracea) FNR and T. gondii FNR were performed under steady-state conditions with different artificial electron acceptors [iodo-nitro-tetrazolium chloride (INT) or 2,6-dichloroindophenol] by continuously monitoring the reactions using either an Agilent 8453 diode array or a Varian Cary 100 double-beam spectrophotometer. Reaction conditions have been described elsewhere [10]. To evaluate the inhibitory effect of NADPO, the concentration of NADPO was varied between 0 and 20 µm, whereas that of NADPH was independently varied between 0.1 and 10 µm and between 2 and 27 µm, in the case of FprA and FNRs, respectively. Ki values of NADPO were determined by fitting the experimental data points to the theoretical equation for the competitive inhibition mechanism, using the nonlinear fitting feature of grafit 5 (Erithacus Software Ltd, Horley, Surrey, UK).

Ligand-binding studies

Spectrophotometric titrations of FprA (c. 15 µm) with either NADPO, NADP+, 2P-AMP or thio-NADP+ were performed at 15 °C in 20 mm Hepes/NaOH (pH 7.0), containing 50 mm NaCl, using a Cary 100 double-beam spectrophotometer (Varian). The spectra were recorded before and after successive additions of equal amounts of the nucleotide to the sample and reference cells. Difference spectra were computed by subtracting the initial spectrum, corrected for dilution, from those recorded after each ligand addition. Kd values were computed by fitting the data points to the theoretical equation for 1 : 1 binding [20], using the nonlinear fitting feature of grafit 5 (Erithacus Software Ltd).


We are grateful to Rita Bernhardt for providing a sample of purified recombinant bovine AdR. We also thank Federico Fischer for his contribution to the initial part of this research. This work was supported by grants from Ministero dell'Università e della Ricerca of Italy (PRIN 2005) and Fondazione Cariplo, Milano, Italy.