NADPH-consuming PrxA is essential to cell survival under H2O2 stress conditions

We have previously shown that A. nidulans lacking PrxA displayed pronounced sensitivity to H2O2 (Xia et al. 2018), whereas, in many living beings, catalases act as the key H2O2-detoxifying enzyme (Rodriguez-Segade et al. 1985). Considering that catalases are abundant in A. nidulans (Kawasaki et al. 1997; Kawasaki et al. 2001), we directly compared the H2O2 protection functions exerted by PrxA and those of catalase B (a major catalase in A. nidulans) (Fig. 1A). Mutants carrying deletions in these genes (∆prxA or ∆catB) were viable with identical growth on agar plates to wild-type A. nidulans (WT) under normal growth conditions. Growth of ∆prxA was completely inhibited with 0.5 mM H2O2, whereas ∆catB exhibited little sensitivity to H2O2, clearly indicating that PrxA, rather than catalase B, is the indispensable enzyme that protects A. nidulans against H2O2 stress.

Fig. 1
figure1

A. nidulans PrxA in tandem with G6PD is essential to fungal ROS defense. A Effects of disrupting prxA and catB on protection against H2O2. Fresh conidia (1 × 105) of WT (WT_argB), ∆prxA, and ∆catB were inoculated on MM plates containing H2O2 at indicated concentrations and then incubated at 37 °C for 2 days. B Lack of PrxA resulted in intracellular accumulation of NADPH. After culture in liquid MM for 16 h, strains were treated with or without 1 mM H2O2 for 30 min before collection for determination of the NADPH/NADP+ ratio in cell lysates. NADPH/NADP+ is presented in relative quantitation; NADPH/NADP+ value of unstressed WT was set to 1 (mean ± SD; n = 3, *P < 0.05, **P < 0.01, one-way ANOVA). CE Phenotypes of WT (WT_pyrG) and nP.gsdA strains on MM plates under H2O2 conditions using ammonium tartrate (left), proline (middle), or nitrate (right) as the sole nitrogen source. F NADPH/NADP+ ratios in WT (WT_pyrG) and nP.gsdA strains. Strains were cultivated in liquid MM with ammonium tartrate (left), proline (middle), or nitrate (right) as the sole nitrogen source for 16 h, and then the NADPH/NADP+ ratios in cell lysates were quantified and compared; NADPH/NADP+ value of WT cultivated with each nitrogen source was set to 1 (mean ± SD; n = 3, *P < 0.05, **P < 0.01, t-test)

To confirm the process where PrxA employs NADPH to decompose H2O2 in vivo, we calculated the changes of intracellular NADPH/NADP+ ratio in WT and ∆prxA under oxidative stress conditions caused by H2O2. As expected, exposing WT to 1 mM H2O2 significantly decreased the NADPH/NADP+ ratio, which is considered to be the result of the NADPH being used for H2O2 decomposition (Fig. 1B). The ∆prxA strain was determined to have a slightly increased NADPH/NADP+ ratio under normal conditions in comparison with that of WT. In sharp contrast to the WT, H2O2 exposure further increased the NADPH/NADP+ ratio in ∆prxA (Fig. 1B), indicating that PrxA consumes NADPH to decompose H2O2. Taken together, we concluded that the NADPH-consuming PrxA plays an essential role in H2O2 detoxification.

Decreasing NADPH impairs antioxidant ability

One of the major NADPH-producing enzymes in A. nidulans is identified as glucose-6-phosphate dehydrogenase (G6PD, encoded by AN2981) (Wennekes et al. 1993). To reveal the direct link between intracellular NADPH and cellular defense against oxidative stress in A. nidulans, we attempted to construct and phenotypically characterize the G6PD deficiency strain (∆gsdA). However, only heterokaryon mutants were obtained (data not shown), suggesting that gsdA may be essential for cell development and growth in A. nidulans. To analyze the functions of the potentially essential gene on oxidative stress resistance, we used the conditional promoter replacement strategy (Marchegiani et al. 2015). This strategy uses the niaD promoter (niaD.P), a nitrogen-regulated promoter from A. nidulans, to replace the endogenous promoter of a target gene to enable strict regulation. Up- and down-regulated expression can be achieved in the presence of NO3 and NH4+ as the sole nitrogen source, respectively. Additionally, proline can be used as a neutral nitrogen source to partially derepress the activity of niaD.P from NH4+ suppression. Using this strategy, the conditional mutant nP.gsdA was successfully constructed (Additional file 1: Fig. S2).

In the absence of H2O2, nP.gsdA exhibited drastically attenuated growth under NH4+ repression conditions (Fig. 1C). The addition of proline partially relieved growth inhibition from NH4+ repression, whereas the addition of NO3 almost recovered the growth rate compared with that of WT under unstressed conditions (Fig. 1D–E). These diverse phenotypes of the conditional mutant responding to the three nitrogen sources further supported the deduction that G6PD is important for fungal development. To provide insights into how intracellular NADPH levels affect the cell growth rate, we measured the intracellular NADPH/NADP+ ratios of nP.gsdA and found profound fluctuation of the NADPH/NADP+ ratio in response to different nitrogen sources (Fig. 1F). NO3 induced a sixfold higher NADPH/NADP+ ratio in nP.gsdA than that in WT, whereas proline and NH4+ decreased the ratio to 2/3 and 1/5 of that of WT, respectively. Obviously, depressing of G6PD decreased intracellular NADPH, which should be responsible for fungal growth retardation.

Next, we investigated how NADPH decrease affects resistance ability of the fungus to oxidative stress. Although nP.gsdA remained alive under NH4+ conditions, the poor cellular growth should make it difficult to estimate the severity of the H2O2 damage under these conditions (Fig. 1C); therefore, we compared conidial viabilities in response to H2O2 treatment between WT and nP.gsdA strains by counting the colonies formed. The survival of nP.gsdA was poorer than that of WT under the oxidative stress conditions induced by 1 mM H2O2 (Additional file 1: Fig. S6A). Consistent with this result, the activity of G6PD and the corresponding NADPH/NADP+ ratio were significantly repressed by NH4+ (Additional file 1: Fig. S6B–C). Together with the fact that the slight derepression of gsdA by proline partially alleviated the H2O2 resistance defect of the mutant (Fig. 1D), we may conclude that the artificial down-regulation of NADPH levels impairs fungal H2O2 resistance ability. This is in agreement with the above-mentioned finding that the indispensable antioxidant PrxA employs NADPH for ROS elimination.

Increasing NADPH also impairs cell antioxidant ability

Given that the intracellular NADPH level is crucial for fungal antioxidant ability, increasing intracellular NADPH levels may be beneficial for fungal oxidative defense, as in Drosophila melanogaster and some other animal cells (Salvemini et al. 1999; Leopold et al. 2003; Legan et al. 2008; Zhang et al. 2012; Xiao et al. 2018, 2020). In our study, we found that NO3 significantly induced gsdA expression (Fig. 2A) and accelerated G6PD activity (Fig. 2B), which resulted in at least a fivefold higher NADPH/NADP+ ratio in nP.gsdA than that in WT under either unstressed or stressed conditions (Fig. 2C). However, unexpectedly, nP.gsdA showed higher H2O2 sensitivity than that of WT (Fig. 1E), which led us to consider that increasing NADPH did not promote and, on the contrary, impaired cell antioxidant ability.

Fig. 2
figure2

NO3 induction promoted transcription (A), activity (B), NADPH production (C) of G6PD in nP.gsdA strains. AC Fresh conidia (1 × 108) of WT (WT_pyrG) and nP.gsdA strains were cultivated in MM medium using NO3 as the nitrogen source for 16 h and then exposed to the indicated concentrations of H2O2 for 30 min for the following analysis (mean ± SD; n = 3, *P < 0.05, **P < 0.01, one-way ANOVA)

For further verification of this hypothesis, we constructed two other NADPH-high producing strains, gP.gsdA and gP.gndA. A constant and high-yield of NADPH was expected to be realized by replacing the native promoters of gsdA and gndA (6PGD encoding gene) with gpdA promoters, which is a strong constitutive promoter derived from the A. nidulans gpdA gene that encodes glyceraldehyde-3-phosphate dehydrogenase (Umemura et al. 2020). In gP.gsdA, the gpdA promoter produced approximately 100-fold more gsdA mRNA than that produced by the native gsdA promoter, but only half of that was produced by the niaD promoter (Additional file 1: Fig. S7A). The intracellular NADPH levels ranged from high to low across the nP.gsdA, gP.gsdA, and WT strains (Additional file 1: Fig. S7B), which was contrary to the orders of fungal H2O2 resistance (Additional file 1: Fig. S7C). In gP.gndA, both gndA mRNA and intracellular NADPH levels were significantly elevated by the gpdA promoter, which also lowered its antioxidant ability (Additional file 1: Fig. S8A–C). These results strengthened the fact that artificial increasing NADPH levels has adverse effects on fungal H2O2 resistance.

To investigate whether excess NADPH increased the levels of oxidants, we used fluorescent probes to measure and compare superoxide and H2O2 accumulated in WT and nP.gsdA strains. Although excess NADPH theoretically can be utilized by NOXs to produce superoxide (Leopold et al. 2003; Gupte et al. 2007; Lee et al. 2011), overexpression of A. nidulans gsdA did not lead to an increase in intracellular superoxide under both stressed and unstressed conditions (Fig. 3A). However, a high level of NADPH appeared to directly contribute to the production of H2O2 because a slight but significant increase of H2O2 accumulation was detected in NO3-induced nP.gsdA than that in WT under normal conditions (Fig. 3B). H2O2 exposure has further promoted intracellular H2O2 accumulation in both strains and enlarged the difference in H2O2 level between WT and nP.gsdA (Fig. 3B). Moreover, the elevation of H2O2 accumulation was prevented by the H2O2 scavenger N-acetyl-l-cysteine (NAC, 10 mM) (Fig. 3B), which also eliminated the H2O2-sensitivity difference between both strains (Additional file 1: Fig. S9). Therefore, it can be concluded that excess NADPH directly contributes to toxic level of H2O2 accumulation in fungal cells under oxidative stress conditions.

Fig. 3
figure3

prxA suppression is responsible for the impairment of fungal resistance to H2O2 in G6PD-overexpression strains. All cultivations used NO3 as the sole nitrogen source. A, B Quantification analysis of intracellular O2·− and H2O2 in WT (WT_pyrG) and nP.gsdA. After precultivation, both strains were exposed to 0 or 1 mM H2O2 for 30 min followed by addition of ROS fluorescent probes. The ROS scavenger NAC (10 mM) was applied 1 h before the probe incubation. Fluorescence intensities of BES-So-AM and BES-H2O2-Ac were used to measure the level of intracellular superoxide and H2O2, respectively. All values were normalized by that in the unstressed WT (set to 100) (mean ± SD; n = 3, *P < 0.05, **P < 0.001, one-way ANOVA). C Relative expression levels of prxA in WT (WT_pyrG) and nP.gsdA. Strains were precultivated for 16 h, and then exposed to 1 mM H2O2 for 30 min. The level of prxA in WT without H2O2-treatment was set to 1 (mean ± SD; n = 3, *P < 0.05, one-way ANOVA). D Relative Prx-GFP levels in the WT (P_Gfp) and nP.gsdA (nP.gsdA/P_Gfp) strains. Inset, fluorescence spectra of Prx-GFPs from the corresponding cell lysates. After preculture, both strains were exposed to 0 and 1 mM H2O2 for 2 h. Cell lysates (1 mg/ml) were used for fluorescence analysis. E Effects of constitutive expression of prxA on fungal oxidative resistance. Conidia (1 × 105) of the strains were spotted and cultivated for 2 days on NO3-MM plates with or without 2 mM H2O2. Newly constructed strains are as follows: gP.prxA (replacing prxA promoter with gpdA promoter) and nP.gsdA/gP.prxA (replacing gsdA promoter with niaD promoter in gP.prxA)

Excess NADPH suppresses prxA transcription by downregulating NapA

Logically, intracellular H2O2 accumulation can be attributed to the inefficiencies of the key H2O2-decomposing enzymes. To explore whether excess NADPH impaired the antioxidant function of A. nidulans PrxA, we compared the transcriptional levels of prxA in WT and NO3-induced nP.gsdA. As expected, external H2O2 greatly increased PrxA transcriptional levels in WT (Fig. 3C), which was consistent with previous findings (Thon et al. 2010; Xia et al. 2018). H2O2-induced prxA expression was also observed in NO3-induced nP.gsdA strains; however, the induction strength was approximately 50% lower than that of WT (Fig. 3C). To investigate whether the transcriptional induction of prxA results of the corresponding changes of PrxA at protein level, we constructed GFP-tagged PrxA expression strains P_Gfp and nP.gsdA/P_Gfp, facilitating the quantification estimation of intracellular PrxA by fluorescence intensity measurements. The P_Gfp strain restored the oxidative resistance caused by prxA deletion (Additional file 1: Fig. S3), indicating the full function of Gfp-tagged PrxA. The same change tendency between gene transcription and protein expression was observed: the induction strength of PrxA in nP.gsdA was lower than in WT, which was indicated by the fluorescence intensity of PrxA-GFP in P_Gfp and nP.gsdA/P_Gfp under H2O2 treatment conditions (Fig. 3D). We hypothesized that the adverse induction of prxA expression caused by excess NADPH may account for the H2O2 accumulation and subsequent H2O2 defense defect in NO3-induced nP.gsdA. To verify this, we constructed two prxA-constitutively expressing strains (gP.prxA and nP.gsdA/gP.prxA) using WT and nP.gsdA as parent strains, respectively (Additional file 1: Fig. S2), and analyzed their antioxidant abilities. In both strains, constitutive expression of prxA was realized by replacing the prxA promoter with gpdA promoter. As expected, constitutive expression of prxA abrogated the distinct of H2O2-resistance between gP.prxA and nP.gsdA/gP.prxA, which was in sharp contrast to WT and nP.gsdA (Fig. 3E). Collectively, these data further illustrate that NADPH may determine the antioxidant ability of fungi via regulating the gene transcription of PrxA, the frontline defender against H2O2.

Repression of prxA transcription by accelerating intracellular NADPH production led us to infer that the function of NapA, the common transcriptional activator of fungal antioxidant genes, including prxA, is impaired under these conditions since NADPH should be the electron donor for NapA reduction and result in consequent deactivation of NapA (Thon et al. 2010). To validate this prediction, we first examined whether NapA can correctly localize in response to H2O2 exposure in the presence of excess intracellular NADPH. A GFP-tagged NapA was introduced to replace the original NapA in WT to construct N_Gfp (NapA-GFP) (Additional file 1: Fig. S3). The H2O2 resistance of N_Gfp was similar to that of WT (Additional file 1: Fig. S3), indicative of the functionality of this NapA::GFP fusion. The strain N_Gfp was further transformed with the pyrGniaD.P-gsdA cassette to construct a new strain nP.gsdA/N_Gfp which can realize the NO3-inducible overexpression of gsdA in the fluorescent strain. Then, we characterized NapA::GFP localization and found that H2O2 exposure quickly resulted in NapA::GFP nuclear accumulation in both strains, indicating that activation of NapA was not interfered by excess intracellular NADPH (Fig. 4). Surprisingly, we found that nP.gsdA/N_Gfp showed significantly reduced fluorescence intensity compared with that of N_Gfp regardless with or without the presence of H2O2 (Fig. 4), indicating that excess intracellular NADPH impaired NapA production, which occurred prior to H2O2 exposure. Thus, we demonstrated that excess intracellular NADPH modulates fungal antioxidant activity by downregulating the amount of NapA rather than by affecting the redox state of NapA.

Fig. 4
figure4

Overexpressing G6PD downregulated expression of NapA but did not interfere with its nuclear localization. Fresh conidia (1 × 105) of N_Gfp (NapA::GFP) and nP.gsdA/N_Gfp (replacing gsdA promoter with niaD promoter in N_Gfp) was incubated in NO3-MM for 10 h and was then exposed to 0 or 2 mM H2O2 for 20 min; nuclei were stained with Hoechst 33258 for 15 min. Scale bar, 20 μm. Images were captured using a laser confocal microscope

Excess NADPH obligatorily activates AnCF to repress napA expression

To elucidate the mechanism whereby NapA expression was downregulated by excess NADPH, we focused on the dynamics of the levels of AnCF, which is a key transcriptional repressor of NapA (Thon et al. 2010; Hortschansky et al. 2017). The HapB, HapC, and HapE subunits of AnCF are all necessary for DNA binding (Thon et al. 2010; Hortschansky et al. 2017). AnCF senses the redox status of the cell via oxidative modification of thiol groups within HapC; oxidized HapC is then unable to participate in AnCF assembly, but can be reduced by the thioredoxin system (TrxA and TrxR) for recycling in the AnCF assembly. Thus, we questioned if excess NADPH can over-reduce HapC, leading to the ROS-resistant defect of nP.gsdA. If this is the case, deletion of hapC should relieve fungal H2O2 sensitivity caused by excess NADPH. Thus, we have constructed a hapC deletion strain (∆hapC) (Additional file 1: Fig. S1) and overexpressed G6PD in this mutant (nP.gsdA/∆hapC) to understand the relationship among HapC, excess NADPH, and cell H2O2 resistance. Deletion of hapC resulted in a great growth defect under normal conditions (Additional file 1: Fig. S4), which is consistent with previous reports (Papagiannopoulos et al. 1996). Overexpression of gsdA using niaD.P under NO3 conditions also realized excess NADPH accumulation in nP.gsdA/∆hapC (Fig. 5A). Next, we compared conidia survival rates of WT, nP.gsdA, ∆hapC, and nP.gsdA/∆hapC strains in response to oxidative stress induced by H2O2 under NO3 induction (Fig. 5B). The ∆hapC strain showed significantly decreased survival rate in all H2O2 stress conditions compared with that of the WT strain, indicating that AnCF is indispensable to A. nidulans oxidative stress resistance. Notably, overexpressing gsdA in ∆hapC did not impair fungal oxidative stress resistance, and, in contrast, substantially rescued the survival rate of nP.gsdA/∆hapC strain (Fig. 5B), clearly indicating that, in the absence of AnCF, extra NADPH supply is advantageous for fungal ROS defense. That is to say, impairment of the antioxidant ability caused by excess NADPH in WT is mediated by the AnCF complex.

Fig. 5
figure5

Intracellular NADPH levels determine the oxidative stress resistance via regulating AnCF complex assembly. A Overexpression of gsdA resulted NADPH accumulation in ΔhapC. WT (WT_pyrG), nP.gsdA, ∆hapC, and nP.gsdA/∆hapC were cultivated in NO3-MM liquid media for 16 h, then the lysates were used for the relative NADPH/NADP+ ratio calculation. NADPH/NADP+ value of WT was set to 1 (mean ± SD; n = 3, **P < 0.01, one-way ANOVA). B AnCF is involved in impairment of H2O2 resistance caused by excess NADPH. Fresh conidia (1 × 108) of four strains were spread on NO3-MM plates containing the indicated concentrations of H2O2. Colonies were counted after a 48-h incubation, and survival rates were expressed as percentages of the CFU for strains incubated without H2O2 (mean ± SD; n = 3). C Immunoblot quantification of AnCF complex levels (top) and HapC levels (bottom box) in WT and nP.gsdA during H2O2-treatment; H_Flag (WT expressing Flag-tagged HapC) and nP.gsdA/H_Flag (nP.gsdA expressing Flag-tagged HapC). Cell-free lysates (100 µg) from each sample were loaded to native- (top) and SDS-PAGE (bottom box). AnCF and HapC were detected using anti-Flag antibody; actin was used as a control and was detected using an anti-Actin antibody. D Quantitated graph for intracellular AnCF level normalized to actin (mean ± SD; n = 3, *P < 0.05, one-way ANOVA). E Time course analysis of NADPH/NADP+ level in H_Flag and nP.gsdA/H_Flag during 1 mM H2O2 treatment. Strains were precultivated in NO3-MM liquid media for 16 h and then exposed to 1 mM H2O2 for 30, 60, and 90 min (mean ± SD; n = 3, t-test). F Relative napA mRNA levels in these strains with or without treatment of H2O2. Strains were cultivated in NO3-MM liquid media for 16 h, and then treated by H2O2 for 30 min and 60 min (mean ± SD; n = 3, *P < 0.05, **P < 0.01, n.s., not significant, two-way ANOVA)

Next, we obtained insight into the relevance between the level of intracellular NADPH and AnCF complex assembly. The Flag-tagged HapC was introduced into both WT and nP.gsdA strains to replace native HapC and construct H_Flag and nP.gsdA/H_Flag strains, respectively (Additional file 1: Fig. S4), enabling the measurement of the cellular level of AnCF in both strains by western blotting. We first confirmed that HapC-Flag protein in both H_Flag and nP.gsdA/H_Flag strains complemented the growth delay caused by hapC deletions (Additional file 1: Fig. S4), indicating that the fusion protein was functional. No bands were detected in cells expressing untagged HapC in WT (data not shown). Considering that the assembly and dissociation of AnCF may affect the dynamics of the intracellular AnCF content, we measured the content of AnCF with time. The HapC bands on reducing SDS-PAGE showed that the total amount of HapC in WT cells was relatively stable across the 120-min observation period under H2O2-treatment conditions (Fig. 5C, bottom box, and Additional file 1: Fig. S10), while levels of AnCF complex in non-reducing native PAGE fluctuated with time (Fig. 5C, top and Fig. 5D). During the earlier 30 min of H2O2 exposure, the intracellular AnCF level declined to 2/3 the level of the pretreatment sample in H_Flag strain. Extending the H2O2 exposure time to 90 and 120 min has gradually recovered and stabilized the AnCF formation to the original level. Interestingly, we found that changes in the level of intracellular NADPH in H_Flag strain kept pace with the fluctuation of AnCF: a sudden drop in the first 30 min, which then returned to the original level within the next 60 min (Fig. 5E). Thus, we deduced that the NADPH intracellular contents may determine the level of the AnCF complex. This supposition was further supported by investigating the AnCF and intracellular NADPH profiles in NO3-induced nP.gsdA/H_Flag, which was proved to be very similar to those present in H_Flag (Fig. 5C–E); moreover, the NADPH level in nP.gsdA/H_Flag was found to be well above that in H_Flag at any time (Fig. 5E). In response to the elevated NADPH, AnCF content also keeps higher in nP.gsdA/H_Flag than that in H_Flag (Fig. 5D). These data, taken together, showed that the initial “down then up” fluctuations of NADPH levels and AnCF contents are the first response of the fungal cells to the H2O2 stimulus.

We further deduced that the “down then up” fluctuation of AnCF content would result in a reverse fluctuation of NapA levels and corresponding up- and downregulated expression of A. nidulans prxA. This was verified by the following transcriptional changes of napA in strains upon H2O2 treatment (Fig. 5F). Exposure to H2O2 for the first 30 min induced napA expression in WT and nP.gsdA, as opposed to the downregulation of intracellular AnCF level in both strains (Fig. 5F). Notably, deletion of hapC drastically elevated napA induction amplitude compared with that of WT during the first 30 min of H2O2 exposure, confirming the transcriptional repression effect of AnCF on napA. Extending H2O2 exposure from 30 to 60 min decreased napA transcription in WT and nP.gsdA (Fig. 5F), which contrasted with the changes in the levels of AnCF (Fig. 5C, top). Conversely, in ∆hapC and nP.gsdA/∆hapC, extending H2O2 exposure from 30 to 60 min did not lower napA transcription levels (Fig. 5F), further confirming the involvement of AnCF in the negative regulation of napA. Since NapA is the transcription activator of prxA, the “down then up” content fluctuation of NADPH should be ultimately used to trigger and subsequently break the induction of prxA to provide the on-demand cellular level of PrxA for oxidative stress defense in A. nidulans.

Reversible inhibition of G6PD may account for the NADPH fluctuation

Under oxidative stress conditions, a sudden “down” of intracellular NADPH level at the initial stage should be the result of NADPH consuming by PrxA for fungal antioxidant machinery. The following “up” of NADPH content suggests a quick activity acceleration of the NADPH-producing enzyme. A. nidulans G6PD may act as the key enzyme, because G6PDs from other sources have been reported to be reversibly inhibited by NADPH, which can be broken by rapid withdrawal of NADPH (Ramos-Martinez 2017). To verify that, we prepared recombinant G6PD of A. nidulans to test the NADPH-dependent inhibition and disinhibition of fungal G6PD in vitro. As shown in Additional file 1: Fig. S5, premixing G6PD with NADPH effectively inhibited fungal G6PD activity, which is indicative of self-braking of G6PD by its product NADPH. Next, we have investigated the disinhibition of G6PD by employing A. oryzae flavohemoglobin (a NADPH-dependent nitric oxide dioxygenase) (Zhou et al. 2011) as an NADPH scavenger. A rapid G6PD activation was achieved by the addition of flavohemoglobin and nitric oxide release reagent to the reaction buffer (Additional file 1: Fig. S5), indicating that regulation of A. nidulans G6PD activity is dependent on disinhibition. Taken together, these results supported our view that the most possible mechanism of NADPH fluctuation may be the result of the rapid NADPH consumption by PrxA upon oxidative exposure coupling the subsequent regeneration of NADPH via the disinhibition of G6PD. Moreover, the fungus may take advantage of the fluctuation of intracellular NADPH to regulate AnCF assembly in response to oxidative stress.

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