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MESH1 is a cytosolic NADPH phosphatase that regulates ferroptosis

Critical to the bacterial stringent response is the rapid relocation of resources from proliferation toward stress survival through the respective accumulation and degradation of (p)ppGpp by RelA and SpoT homologues. While mammalian genomes encode MESH1, a homologue of the bacterial (p)ppGpp hydrolase SpoT, neither (p)ppGpp nor its synthetase has been identified in mammalian cells. Here, we show that human MESH1 is an efficient cytosolic NADPH phosphatase that facilitates ferroptosis. Visualization of the MESH1-NADPH crystal structure revealed a bona fide affinity for the NADPH substrate. Ferroptosis-inducing erastin or cystine deprivation elevates MESH1, whose overexpression depletes NADPH and sensitizes cells to ferroptosis, whereas MESH1 depletion promotes ferroptosis survival by sustaining the levels of NADPH and GSH and by reducing lipid peroxidation. The ferroptotic protection by MESH1 depletion is ablated by suppression of the cytosolic NAD(H) kinase, NADK, but not its mitochondrial counterpart NADK2. Collectively, these data shed light on the importance of cytosolic NADPH levels and their regulation under ferroptosis-inducing conditions in mammalian cells.


Results
MESH1 is an efficient NADPH phosphatase. We reasoned that MESH1 may function through alternate metabolic substrate(s) from ppGpp in mammalian cells and examined metabolites with a similar molecular architecture to ppGpp. The metabolite NADPH shares many similarities with ppGpp, including a purine nucleoside, a 2′-phosphate group and a 5′-pyrophosphate group (Fig. 1a). Although NADPH differs from ppGpp in that it contains a 2′-phosphate instead of a 3′-pyrophosphate in ppGpp, the crystal structure of the bifunctional RelA/SpoT homologue from Streptococcus dysgalactiae subsp. equisimilis captured an unusual ppGpp derivative, GDP-2′,3′-cyclic monophosphate, in the active site of the hydrolase domain 6 , suggesting that the enzyme may accommodate a 2′-substituted phosphate group 6 . Based on the observation that SpoT catalyses the hydrolysis of the 3′-pyrophosphate group of ppGpp, we predicted that MESH1 would similarly hydrolyse the 2′-phosphate group of NADPH to yield NADH and an inorganic phosphate (Fig. 1a).
Indeed, treatment of NADPH with hMESH1 readily released inorganic phosphate, yielding a green solution in the malachite green assay 7 (Fig. 1b). Importantly, the phosphate accumulation was linear over time ( Fig. 1c), reflecting continuous enzymatic turnover of NADPH by hMESH1. Furthermore, mass spectrometry analysis revealed a peak at m/z 664.141 as [M-H] − , verifying the product as NADH (Fig. 1d). We then analysed the steady state kinetics parameters of hMESH1 toward NADPH (Fig. 1e). hMESH1 is an efficient NADPH phosphatase, with a catalytic efficiency (k cat /K M ) of 14.4 ± 1.1 s −1 mM −1 ; its Michaelis constant (K M ) value of 0.12 ± 0.01 mM is on par with that from other reported cellular enzymes utilizing NADPH (for example, K M value of 0.11 mM for the human phagocytic NADPH oxidase 8 ), supporting its role as a physiologically relevant NADPH phosphatase in cells. While hMESH1 also displays measurable activity towards NADP + in vitro, it is approximately tenfold less efficient (k cat /K M = 1.4 ± 0.1 s −1 mM −1 ), and its K M value of 0.43 ± 0.04 mM is higher than the estimated cytosolic concentration of NADP + , rendering hMESH1 an ineffective phosphatase for NADP + in cells (Fig. 1e).
Previous biochemical and structural analysis has revealed MESH1 as a Mn 2+ -dependent enzyme 4 . Accordingly, we found that the NADPH phosphatase activity of hMESH1 was reduced by >10 4fold when Mn 2+ was substituted with other metal ions, such as Zn 2+ (Fig. 1f). Likewise, mutations of hMESH1 residues near the Mn 2+ ion on the active site, including E65A and D66K, severely compromised the NADPH phosphatase activity by >10 4 -fold in enzymatic assays (Fig. 1f).
To determine whether MESH1 is a significant contributor to the cellular NADPH phosphatase activity, we measured the enzymatic activity in cell lysates extracted from two human cell lines, RCC4 cells (a human clear cell renal carcinoma cell line) and HEK-293T cells (a human embryonic kidney cell line). We found that lysates from MESH1-silenced cells had a noticeable decrease of the NADPH phosphatase activity ( Fig. 1g and Extended Data Fig. 1a). Conversely, overexpression of wild-type (WT) hMESH1, but not the catalytically deficient E65A hMESH1 mutant, markedly enhanced the cellular NADPH phosphatase activity (Extended Data Fig. 1b). We also investigated the primary subcellular compartment where MESH1 functions. Cellular fractionation experiments 9 indicated that MESH1 was predominately enriched in the cytosolic, but not the mitochondrial or nuclear, pools (Fig. 1h). Taken together, these observations verify hMESH1 as a significant contributor to the cytosolic NADPH phosphatase activity in human cells. It is worth noting that the short interfering RNA (siRNA) of MESH1 did not suppress NADPH phosphatase activity completely, indicating the possible existence of another NADPH phosphatase or nonspecific phosphatase activity released during sample processing that also contributed to the NADPH phosphatase activity in mammalian cells. One possible candidate is the NADP(H) phosphatase activity previously observed in rat liver lysates 10 , though the reported enzymatic activity favours NADP + and the identity of the enzyme has remained unknown in animal cells 11 . Another contributor could be an unrelated mitochondrial NADPH phosphatase, Curl 12 , reported after the release of our discovery at bioRxiv 13 . Therefore, to the best of our knowledge, the cytosolic NADPH phosphatase activity of MESH1 is distinct from the previously observed activity and MESH1 represents a new instance of a cytosolic NADPH phosphatase in human cells.  Fig. 1 | MESH1 is a mammalian NADPH phosphatase. a, Chemical similarity between NADPh and ppGpp, and the proposed chemical reaction of MESh1. b, Detection of the hMESh1-catalysed phosphate release from NADPh by colour change (yellow to green) in the malachite green assay. c, Linear product accumulation of the NADPh dephosphorylation reaction catalysed by hMESh1 (n = 3 biologically independent samples; error bars indicate s.d.). d, Validation of the formation of NADh by mass spectrometry analysis. This experiment was repeated three independent times with similar results. e, Enzymatic characterization of hMESh1 toward NADPh and NADP + (mean ± s.d.; n = 4 independent experiments). f, Effects of Zn 2+ substitution and active site mutation on the specific activity of purified recombinant hMESh1 (mean ± s.d.; n = 3 independent experiments). g, hMESh1 is a significant contributor to the NADPh phosphatase activity in RCC4 lysates. Black dots represent siNT samples, and blue dots represent siMESh1 samples. n = 4 biologically independent samples for each condition; samples were analysed by western blot to confirm knockdown efficiency. h, Cell fractionation separating cytosol from nucleus and mitochondria, showing that MESh1 is specifically enriched in the cytosolic compartment. This representative western blot was repeated independently three times on RCC4 cells and hEK-293T cells with similar results. Error bars indicate s.d. One-way analysis of variance (ANOVA) with Tukey's honest significant difference (hSD) post-hoc test, *P < 0.05, **P < 0.01. Source data are provided in the Source Data File. Mit, mitochondria; NT, nontargeting; Nuc, nucleus.
cocrystal structure of the specific hMESH1-NADPH complex by comparing it with the catalytically compromised D66K mutant enzyme and by substituting the catalytic Mn 2+ ion with Zn 2+ . The structure was refined to 2.1-Å resolution (Supplementary Table 1). The crystallographic asymmetric unit consists of two protomers of hMESH1 (Extended Data Fig. 2a), each adopting a compact fold of ten α-helices and a short β-hairpin (Fig. 2a) previously observed in the structure of apo hMESH1 (ref. 4 ). Although the active sites of both protomers were occupied by the adenosine portion of the substrate (Extended Data Fig. 2b), the molecular recognition of the entire NADPH molecule, including that of the nicotinamide moiety, was only visible in one of the two protomers (Extended Data Fig. 2b), which is described.
The catalytic site of hMESH1 is formed at the centre of the helical architecture, surrounded by the short β-hairpin (β1 and β2), α2, the α3-α4 loop, α4, α7, α8, the α8-α9 loop and α9 (Fig. 2a). Anomalous scattering revealed the presence of a single Zn 2+ ion in the active site, which substitutes the catalytic Mn 2+ ion reported in the apo hMESH1 structure (Extended Data Fig. 2c). The Zn 2+ ion is hexa-coordinated in a distorted octahedral geometry by H35 of α2, H61 and D62-the signature HD motif of the α2-α3 loop that defines this family of enzymes, D122 of α8, the catalytic water molecule and the 2′-phosphate group of NADPH (Fig. 2b).
In addition to coordinating the active site metal ion, the 2′-phosphate group of NADPH forms additional polar interactions with Y32 of α2, K66 of α4 and N126 of α8 that are located one layer above the equatorial plane of the zinc ligands (Fig. 2c,d). K66, the mutated residue substituting D66 in the WT enzyme, not only significantly diminishes the catalytic activity, but also forms a direct salt bridge with the 2′-phosphate group of NADPH, a likely contributor to the successful capture of NADPH in the cocrystal structure. Another catalytically important residue, E65 of α4, the neighbouring residue of D66, is located over 4 Å away from the catalytic water molecule, and its indirect interactions with the catalytic water molecule and the 2′-phosphate of NADPH are bridged by a second water molecule in the active site (Fig. 2c,d).
The NADPH adenosine ribose group and its adjacent 5′-pyrophosphate group are extensively recognized (Fig. 2c,d). The 3′-hydroxyl group and the 4′-oxygen atom form hydrogen bonds with Y146 of α9 and K25 of β1, respectively. The 5′-diphosphate group adopts a tight turn, aided by salt bridges with K97 of α7, K25 of β1 and a sodium ion. While sodium is an unusual cation to coordinate the NADPH diphosphate, our purification procedure and crystallization condition lack divalent cations other than Zn 2+ but contain over 600 mM sodium chloride. Given that this density lacks anomalous Zn 2+ signals (Extended Data Fig. 2b), it is interpreted as a sodium ion, though Mg 2+ or Ca 2+ might be more suitable ions in vivo. The adenine moiety of NADPH is largely coordinated by π-stacking with R24 of β1 and a direct hydrogen bond between its N7 atom and the amide group of K25 of the β1 loop (Fig. 2c,d). As both of these interactions are also found in a guanine base, hMESH1 lacks the ability to differentiate the adenine nucleotide from the guanine nucleotide as was found in ppGpp. The ribose ring of the nicotinamide riboside is indirectly recognized by a water-mediated hydrogen bond of its ring oxygen atom with R142 of α9, whereas the nicotinamide moiety is supported by a π-stacking network with W138 and R142 that emanate from the α8-α9 loop and from α9 (Fig. 2c,e). The reduced electron density of the nicotinamide ring in NADP + would weaken such interaction and likely contribute to the reduced substrate affinity in comparison with NADPH. Finally, the terminal amide group is additionally buttressed by two hydrogen bonds with H141 and E145 of α9. These structural observations reinforce the notion that hMESH1 is a bona fide NADPH phosphatase.

MESH1 regulates cellular NADPH levels and ferroptosis.
After establishing hMESH1 as a significant contributor to the cellular NADPH phosphatase activity and elucidating its molecular recognition of NADPH, we investigated whether the NADPH phosphatase activity of MESH1 plays a role in ferroptosis, a regulated form of cell death driven by the lethal accumulation of lipid hydroperoxides 14 . Various genetic determinants of ferroptosis have been identified [15][16][17][18] . Importantly, ferroptosis can be triggered by erastin, an inhibitor of the cystine importer xCT 5 , or by cystine deprivation, and is alleviated by high concentrations of NADPH 19 . Accumulation of NADPH promotes the glutathione-reductase-catalysed regeneration of reduced glutathione (GSH), which serves as the reductive cofactor for glutathione peroxidase 4 (GPX4) to detoxify lipid hydroperoxidates, thus protecting cells from ferroptotic cell death (Fig. 3a). We found that MESH1 was significantly upregulated in RCC4 cells at both messenger RNA and protein levels by erastin treatment or cystine Erastin Erastin-siMESH1-3′ UTR 4%  Fig. 3a,b), implicating a functional role of MESH1 in these ferroptosis-inducing conditions. Overexpression of WT hMESH1 (MESH1-WT), but not the catalytically deficient mutant (MESH1-E65A), significantly lowered the intracellular NADP(H) level (Extended Data Fig. 3c) and sensitized cells to ferroptosis upon erastin treatment (Extended Data Fig. 3d). Taken together, these observations implicate a direct role of MESH1 in the execution of ferroptosis through degradation of NADPH.
To further assess the MESH1 function in ferroptosis, we investigated whether depletion of MESH1 and its associated NADPH phosphatase activity could mitigate lipid peroxidation and ferroptosis through the accumulation of NADPH and alteration of the glutathione redox potential. Erastin treatment dramatically reduced the level of NADP(H) (Fig. 3c) and increased degree of lipid peroxidation (Fig. 3d) in control (nontargeting siRNA, siNT) cells. Importantly, MESH1 silencing sustained a significantly higher level of NADP(H) under ferroptosis-inducing conditions ( Fig. 3c and Extended Data Fig. 3e), and markedly reduced the level of lipid peroxidation (Fig. 3d) under erastin. In addition, MESH1 silencing robustly protected cells from erastin-induced death over a broad range of erastin concentrations (Extended Data Fig. 3f) when monitored for over a 7-d period using CellTox-Green assay, which measures cell death by DNA staining (Fig. 3e). The ferroptosis protection of MESH1 silencing was further validated by multiple additional types of viability and death assays (Extended Data Fig. 4a-c). Such a ferroptosis-rescuing effect appeared to be general, as MESH1 depletion similarly promoted resistance to erastin-triggered ferroptosis in multiple cell lines, including HEK-293T, H1975, MDA-MB-231, PC3, HT1080, A673, PANC1 and 786-O (Extended Data Fig. 5a-h).
After confirming the general role of MESH1 in ferroptosis regulation, we investigated at which signalling steps MESH1 functions during ferroptosis. We found that MESH1 depletion similarly conferred resistance to ferroptosis induced by cystine deprivation and sulfasalazine, another xCT inhibitor (Extended Data Fig. 3f,g), which trigger ferroptosis by depleting intracellular GSH. In comparison, inhibition of GPX4 downstream of NADPH and GSH also triggers ferroptosis (Fig. 3a), but MESH1 depletion could not rescue ferroptosis induced by different GPX4 inhibitors, such as ML-162 or RSL3 (Fig. 3h,i). These observations indicate that MESH1 depletion functions upstream of GPX4 and rescues ferroptosis by NADPH-driven repletion of GSH (Fig. 3a). Consistent with this analysis, although no statistically significant effects on the levels of intracellular cysteine or oxidized glutathione (GSSG) were observed in MESH1-depletion cells, (Extended Data Fig. 6a,b), we observed a significant increase of the GSH level and the GSH/GSSG ratio (Extended Data Fig. 6c,d).
NADPH phosphatase activity of MESH1 contributes to ferroptosis. Next, we investigated the specific roles of the enzymatic activities of MESH1 in the ferroptosis-related phenotypes. We found that the resistance to erastin-induced ferroptosis is directly correlated with the loss of the catalytic activity of MESH1, as the ferroptotic survival phenotype in MESH1-depletion cells was abolished when WT MESH1, but not the catalytically deficient E65A mutant of MESH1, was expressed (Fig. 4a).
Besides NADPH, the bacterial ppGpp is also a previously identified in vitro substrate of MESH1 (ref. 4 ), which has been suggested as a signalling molecule despite the lack of evidence of its presence in mammalian cells. To verify that the enhanced survival under ferroptosis-inducing erastin treatment is dependent on a higher sustained level of NADPH, but not due to accumulation of any other in vitro substrate(s) of hMESH1, such as ppGpp, we tested whether the survival advantage of MESH1 depletion is reversed by simultaneous depletion of NAD(H) kinases that convert NAD(H) to NADP(H). Human cells have two NAD kinases, NADK and NADK2, that are predominantly located in the cytosol and mitochondria 20,21 , respectively (Fig. 3a). As the cytosolic and mitochondrial pools of NADP(H) and NAD(H) are compartmentalized in mammalian cells due to the impermeability of these molecules across the mitochondrial membranes 22 , we reasoned that removal of the NAD kinase either from the cytosol (NADK) or mitochondria (NADK2) would reduce the distinct pools of NADP(H) and only the cytosolic NADK depletion would compromise the ferroptosis survival phenotypes of MESH1-silenced cells. Indeed, when the gene encoding the cytosolic enzyme NADK was silenced, the survival benefit of silencing MESH1 was largely eliminated (Fig. 4b and Extended Data Fig. 7). In contrast, silencing the gene encoding the mitochondrial enzyme NADK2 did not affect the MESH1-depletion-mediated ferroptosis survival (Fig. 4b and Extended Data Fig. 7). Consistent with this notion, we found that the cellular NADP(H) was significantly diminished when NADK was silenced, whereas silencing of NADK2 had little effect on cellular NADP(H) (Fig. 4c,d), suggesting a limited role of NADK2 in this process. Importantly, the mRNA expression level of MESH1, as measured by the normalized reads in the RNA sequencing, was comparable (within 0.5-4 fold) to that of other genes in the NADPH-glutathione pathway (Fig. 3a), including NADK, NADK2, NNT (NAD(P) transhydrogenase, mitochondrial), G6PD (glucose-6-phosphate dehydrogenase), PGD (phosphogluconate dehydrogenase), GSR (glutathione reductase) and NOX1 (NADPH oxidase 1) (Extended Data Fig. 8), supporting a functional role of MESH1 in mediating the essential cytosolic metabolite NADPH. Taken together, our results establish MESH1 as a cytosolic NADPH phosphatase that is induced by ferroptosis-inducing conditions and is involved in the execution of ferroptosis by degrading NADPH. Conversely, depletion of MESH1 and its NADPH phosphatase activity under ferroptosis-inducing conditions promotes stress survival by preserving NADPH, elevating GSH and mitigating lipid peroxidation.

Discussion
MESH1, the metazoan homologue of the bacterial ppGpp hydrolase SpoT, has been implicated in the cellular response to metabolic starvation in Drosophila, though the functional role and pertinent mammalian cell substrate of MESH1 have not been elucidated 4 . The discovery of MESH1 as a cytosolic NADPH phosphatase that contributes to the execution of ferroptosis through degradation of the central metabolite NADPH directly addresses this mystery and implicates a physiological role of MESH1 in disposing of cells damaged by metabolic and redox disorders.
Ferroptosis is a distinct form of iron-dependent cell death 5 that is triggered by oxidative stresses and characterized by the accumulation of lipid peroxidation products 14 . Even with the recent discovery of many genetic determinants [15][16][17][18]23,24 of ferroptosis by regulating lipid metabolism, iron availability, reactive oxygen species (ROS) and anti-oxidant capacity, much still remains unknown about the molecular mechanisms regulating ferroptosis. Our discovery of MESH1 as an NADPH phosphatase reveals an important role of the MESH1-mediated NADPH depletion and impairment of the glutathione regeneration in the execution of ferroptosis. Ferroptosis also has emerging roles in other pathophysiological processes and human diseases, such as neurotoxicity 25 , acute renal failure 26 , hepatic 27 and cardiac 28 injuries, and ischaemia-reperfusion injury 29 . Therefore, modulating ferroptosis may have therapeutic potentials, and the newly discovered MESH1-NADPH phosphatase activity and our structural elucidation of the hMESH1-NADPH may be harnessed for developing potential therapeutics for the treatment of these devastating diseases involving dysregulated ferroptosis.

Methods
Purification of recombinant hMESH1. The gene encoding hMESH1 was codon optimized for E. coli expression, synthesized and cloned into a modified pET28a vector as a C-terminal fusion to the His 6 -tagged SUMO protein.
Cultures of transformed E. coli strain BL21(DE3)* were grown to an optical density at 600 nm between 0.4 and 0.5 and induced with 1 mM isopropyl β-d-1thiogalactopyranoside at 37 °C for 2 h. Following cell lysis, the target protein was purified using Ni 2+ -nitrilotriacetic acid affinity chromatography following standard protocols (Qiagen). The SUMO tag was cleaved using the SENP1 protease, and both the tag and protease were removed by a second round of Ni 2+ -nitrilotriacetic acid chromatography. The target protein was further purified using size-exclusion chromatography (Superdex 75, GE Life Sciences) in a buffer containing 50 mM Tris, pH 8.0, 200 mM NaCl and 0.1% 2-mercaptoethanol. Mutants of hMESH1 were generated using the QuikChange site-directed mutagenesis kit (Agilent) and prepared using the same procedure.
Mass spectrometry. Reversed-phase liquid chromatography-electrospray ionisation tandem mass spectrometry for NADH and NADPH was performed using a Shimadzu liquid chromatography system (comprising a solvent degasser, two LC-10A pumps and an SCL-10A system controller) coupled to a highresolution TripleTOF5600 mass spectrometer (Sciex). Liquid chromatography was operated at a flow rate of 200 μl min −1 with a linear gradient as follows: 100% mobile phase A was held isocratically for 2 min and then linearly increased to 100% mobile phase B over 5 min, and held at 100% B for 2 min. Mobile phase A was a mixture of water/acetonitrile (98:2, v/v) containing 0.1% acetic acid. Mobile phase B was a mixture of water/acetonitrile (10:90, v/v) containing 0.1% acetic acid. A Zorbax SB-C8 reversed-phase column (5 μm, 2.1 × 50 mm 2 ) was obtained from Agilent. The liquid chromatography eluent was introduced into the electrospray ionization (ESI) source of the mass spectrometer. Instrument settings for negative ion electrospray ionization/mass spectrometry and tandem mass spectrometry analysis of lipid species were as follows: ion spray voltage = −4,500 V; curtain gas = 20 pounds per square inch (psi); ion source gas 1 = 20 psi; de-clustering potential = −55 V; focusing potential = −150 V. Data acquisition and analysis were performed using the Analyst TF1.5 software (Sciex).
The sample preparation process for quantification of cysteine, GSH and GSSG was carried out using an automated MicroLab STAR system (Hamilton). After removal of the protein fraction, the extract was divided into two fractions: one for analysis by liquid chromatography (GSH, GSSG) and one for analysis by gas chromatography (cysteine). The liquid chromatography/mass spectrometry portion of the platform is based on a Waters ACQUITY UPLC and a Thermo-Finnigan LTQ mass spectrometer, which consists of an ESI source and linear ion-trap mass analyser. The sample extract was analysed using acidic positive ion optimized conditions. Extracts reconstituted in acidic conditions were gradient eluted using water and methanol both containing 0.1% formic acid, while the basic extracts, which also used water/methanol, contained 6.5 mM ammonium bicarbonate. The mass spectrometry analysis alternated between mass spectrometry and datadependent tandem mass spectrometry scans using dynamic exclusion. The samples destined for gas chromatography/mass spectrometry analysis were re-dried under vacuum desiccation for a minimum of 24 h before being derivatized under nitrogen using bistrimethyl-silyl-triflouroacetamide. The gas chromatography column was 5% phenyl and the temperature ramp was from 40 °C to 300 °C in a 16-min period. Samples were analysed on a Thermo-Finnigan Trace DSQ fast-scanning singlequadrupole mass spectrometer using electron impact ionization. The information output from the raw data files was automatically extracted. For ions with counts greater than 2 million, an accurate mass measurement could be performed. Accurate mass measurements could be made on the parent ion as well as fragments. The typical mass error was less than 5 p.p.m. Identification of known chemical entities was based on comparison with metabolomic library entries of purified standards. The combination of chromatographic properties and mass spectra gave an indication of a match to the specific compound or an isobaric entity.
Enzymology. Enzymatic assays for hMESH1 were performed in a buffer containing 50 mM Tris, pH 8, 200 mM NaCl and 1 mM MnCl 2 or ZnCl 2 . Michaelis-Menten kinetics assays for NADPH and NADP + were carried out using 50 nM or 500 nM enzyme, respectively, and serial dilutions of substrate concentration starting at 1 mM. Reactions were stopped by the addition of formic acid (3 M final concentration) at 2-min, 4-min, 6-min, 8-min and 10-min time points. Specific activities of purified hMESH1 mutants were determined using 93 μM Zn-replaced, 33 μM E65K, 100 μM D66K or 50 nM WT enzyme with 1 mM NADPH, with the reaction stopped at 30 min for each mutant and 10 min for the WT. Specific activities of the knockdown cell lysates were performed using diluted cell lysate (1:40,000 in Tris-NaCl with 1 mM MnCl 2 ) and reactions were stopped at 15 min, 30 min, 45 min, 60 min, 75 min and 90 min. The amount of released phosphate was assessed using the malachite green reagent 7 by mixing equal parts of reaction mixture and malachite green reagent and measuring the absorbance at 630 nm. Values for K M and maximal enzyme rate, V max , were calculated from the Michaelis-Menten equation.
X-ray crystallography. Crystals of hMESH1 bound to NADPH were formed using the hanging drop vapour diffusion method. Immediately before crystallization, hMESH1 was treated with 10 mM EDTA for 30 min and exchanged into a buffer containing no metal before addition of 1 mM ZnCl 2 . Diffracting-quality crystals were obtained by mixing equal volumes of the protein solution (9 mg ml −1 hMESH1, 100 mM NADPH, 50 mM Tris, pH 8.0, 200 mM NaCl and 0.1% 2-mercaptoethanol) with the mother liquor solution (200 mM ammonium acetate, 100 mM sodium citrate and 25% PEG 4000) by streak-seeding with apo hMesh1 crystals. To improve NADPH occupancy, crystals were additionally soaked in the mother liquor solution containing 750 mM NADPH for 1 h before cryoprotection in a solution containing 525 mM NADPH, 10 mM ZnCl 2 , 20% ethylene glycol and 70% precipitant solution and flash freezing in liquid nitrogen.
X-ray data were collected at the Southeast Regional Collaborative Access Team 22-BM beamline at the Advanced Photon Source, Argonne National Laboratory and processed using the XDS package 30 . The structure was determined by molecular replacement with the program Phaser 31 and by using the previously reported structure of apo hMESH1 (3NR1) as the search model. Iterative model building and refinement were carried out using COOT 32 and PHENIX 33 .

MESH1 overexpression.
Stable RCC4 cell lines with MESH1-WT (RefSeq: NM_001286451.1) or MESH1-E65A expression were generated using lentiviral vector pLX302 (ref. 34 ) (a gift from David Root, Addgene plasmid no. 25896) without expression of the V5 tag. Briefly, virus was generated by transfecting HEK-293T cells with a 0.1:1:1 ratio of pMD2.G/psPAX2/pLX302 with TransIT-LT1 Transfection Reagent (Mirus, MIR2305). pMD2.G and psPAX2 were gifts from Didier Trono (Addgene plasmid no. 12259 and no. 12260, respectively). Virus was collected 48 h after transfection. Stable cell lines were generated by adding 200 μl of virus to a 60-mm dish of RCC4 cells with 8 μg ml −1 polybrene. Then, 1 μg ml −1 puromycin was used for selection. Complete death in blank infection dishes was used to determine the success of infection and the length of puromycin selection. The efficiency of overexpression was determined by western blotting.
Transient overexpression of MESH1 in HEK-293T cells for the NADP(H) content assay or enzymology assay was achieved by using the pCMV6-neo vector (OriGene, SC334209). Briefly, 2 × 10 5 HEK-293T cells were seeded in one well of a six-well plate for 24 h, and then 1 μg of the plasmid (pCMV6-neo empty vector or with MESH1-WT or MESH1-E65A) was transfected with TransIT-LT1 Transfection Reagent (Mirus) for an additional 48 h before collection.
Cell culture. The RCC4 cell line was provided by Denise Chan (University of California, San Francisco, San Francisco, CA) and was authenticated by DNA Diagnostics Center Medical using the short tandem repeat method in November of 2015. HEK-293T, H1975, MDA-MB-231, 786-O, PC3, HT1080, A673 and PANC1 were obtained from the Duke Cell Culture Facility whose origins were ATCC. All cell lines were tested negative for mycoplasma before use. All cells were cultured in DMEM with 4.5 g l −1 glucose and 4 mM glutamine (11995-DMEM, ThermoFisher Scientific) and 10% heat-inactivated fetal bovine serum (Hyclone no. SH30070.03HI) in a humidified incubator, at 37 °C with 5% CO 2 .

NADP(H) measurement.
The Amplite colorimetric NADPH assay kit (no. 15272, ATT Bioquest) was used to measure NADP(H) (both NADP + and NADPH) in this study. Briefly, cells were seeded into a six-well plate with siRNA or plasmid as described in the MESH1 silencing using RNA interference section. NADP(H) was collected using 100 μl of the provided lysis buffer. The NADP(H) content was measured and normalized by protein content of the lysate, and quantified by the bicinchoninic acid assay. In cells treated with erastin or cystine deprivation, the NADP(H) change is normalized with NADP(H) measured in dimethylsulfoxidetreated cells or cells in regular full media.

NADPH sensor experiments.
For the NADPH biosensor assay, HEK-293T cells (6,000 cells per well in a 96-well plate) were counted and reverse transfected with control or indicated MESH1 siRNA using Lipofectamine RNAiMAX (Invitrogen). After 24 h, the cells were transfected with iNAP1 biosensor 38 using TransIT-LT1 (Mirus Bio). After overnight incubation, the medium was replaced with 20 µM erastin. After 16 h, dual-excitation ratios were acquired with excitation filters 420 nm and 485 nm, and emission filter 528 nm for both. Background subtraction was performed by subtracting the intensity of the sample without transfecting iNAP1.
Viability and death measurement. Multiple viability and cell death assays were used to probe cell viability and death. The cell viability was measured by the CellTiter-Glo assay (Promega, no. G7570), which quantifies intracellular ATP by luminescence as an indicator of viability, or by crystal violet staining of the attached live cells, which is quantified by dissolving in 10% acetic acid and measured by the absorbance at 570 nm. Cell death under erastin with multiple time points was evaluated by the CellTox-Green Cytotoxicity assay (Promega, no. G8741), which stains DNA in dead cells. The cell death was further confirmed with the CytoTox-Fluor assay (Promega, no. G9260), which quantified protease release from broken cells in the cell culture media. The fluorescence intensity, absorbance and luminescence intensity were measured by using a plate reader (FLUOstar Optima, BMG Labtech).
Lipid peroxidation measurement. The C11-BODIPY dye (D3861, ThermoFisher Scientific) was used to determine lipid ROS levels according to the manufacturer's instructions. Cells were reverse transfected with siRNAs as indicated in six-well plates for 2 d, and then treated with 1 µM erastin overnight. The next day, the cells were incubated with 10 µM C11-BODIPY-containing medium for 1 h. Later, the cells were collected by 0.25% trypsin-EDTA treatment and washed three times with ice-cold PBS followed by re-suspension in PBS containing 1% BSA. The amount of ROS within cells was determined by flow cytometry analysis (FACSCanto II, BD Biosciences).
Quantitative real-time PCR. RNA was extracted using the RNeasy kit (Qiagen, 74104) following the manufacturer's instructions. Total RNA (500 ng) with or without reverse transcriptase was prepared using a SensiFast cDNA Synthesis Kit (Bioline, BIO-65054) for real-time PCR comparison with the Power SYBRGreen Mix (ThermoFisher Scientific, 4367659). Primers were designed across exons Extended Data Fig. 1 | MESH1 contributes to the NADPH phosphatase activity in cells. NADPh phosphatase activity of hEK-293T was estimated by incubating NADPh with cell lysates with different MESh1 status; phosphate release over time was measured by the malachite green assay and normalized by the total lysate protein (μmol/min/mg total lysate protein). a, Significantly decreased cellular NADPh phosphatase activity by silencing

NATuRE METAbolISM
Extended Data Fig. 3 | up-regulation of MESH1 protein and its regulation of NADP(H) and cell survival during ferroptosis. a, b, The levels of MESh1 protein were determined by Western blots of MESh1 and β-tubulin protein in RCC4 cells treated with (a) DMSO vs. erastin (3 µM; 12 h) or (b) 200 or 0 µM for 20 h. All the representative western blots were repeated for 3 biologically independent times with similar results. c, Overexpression of WT MESh1, but not the MESh1-E65A mutant, significantly reduced the intracellular NADP(h) content in RCC4 cells. d, Overexpression of MESh1-WT, but not MESh1-E65A, sensitized the cell death to erastin treatment in RCC4 cells. Black dots represent empty vector samples, red dots represent MESh1-WT samples, and grey dots represent MESh1-E65A samples. e, MESH1 silencing restored the intracellular NAPDh level decreased by erastin treatment. hEK-293T cells transfected with the NADPh fluorescent biosensor, iNAP1, and siNT or two independent siMESh1 RNAs were treated with 20 µM of erastin. The ratio of fluorescence excited at 420 nm and 485 nm indicated the cellular NADPh level. The fluorescence ratios of individual treatments were normalized to that of the control condition. (n=6 biologically independent samples). f, The viability of RCC4 cells transfected with siNT (black) or two independent siMESh1 (blue) and treated with indicated concentrations of erastin for one day. Each dot represents one sample; there are three biologically independent samples (n=3) in each group unless otherwise specified. Error bars indicate s.d. Statistical analysis: one-way ANOVA with Tukey's hSD posthoc test, *P<0.05, **P<0.01. Source data are provided in the Source Data File.

Statistics
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The exact sample size (n) for each experimental group/condition, given as a discrete number and unit of measurement A statement on whether measurements were taken from distinct samples or whether the same sample was measured repeatedly The statistical test(s) used AND whether they are one-or two-sided Only common tests should be described solely by name; describe more complex techniques in the Methods section.
A description of all covariates tested A description of any assumptions or corrections, such as tests of normality and adjustment for multiple comparisons A full description of the statistical parameters including central tendency (e.g. means) or other basic estimates (e.g. regression coefficient) AND variation (e.g. standard deviation) or associated estimates of uncertainty (e.g. confidence intervals) For null hypothesis testing, the test statistic (e.g. F, t, r) with confidence intervals, effect sizes, degrees of freedom and P value noted Give P values as exact values whenever suitable.

For Bayesian analysis, information on the choice of priors and Markov chain Monte Carlo settings
For hierarchical and complex designs, identification of the appropriate level for tests and full reporting of outcomes Estimates of effect sizes (e.g. Cohen's d, Pearson's r), indicating how they were calculated Our web collection on statistics for biologists contains articles on many of the points above.

Software and code
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Data collection
The X-ray diffraction data were collected at SER-CAT on the 22-BM line. The ROS measurement data by flow cytometry were collected by the BD FACSDiva v8.0.1 (BD FACSCanto II).

Data analysis
Raw X-ray diffraction data were processed with the XDS package (BUILT=20170215). The hMESH1-NADPH complex structure was determined by molecular replacement using the Phaser module in the Phenix package (version: 1.10.1_2155) and using the previously reported structure of apo hMESH1 (3NR1) as the search model. Iterative model building and refinement was carried out using COOT (version 0.8.9.1) and PHENIX (version: 1.10.1_2155). The structure images were made in PyMOL (version 1.8.0.4). Raw flow cytometry data were analyzed by the FlowJo v10.1.
For manuscripts utilizing custom algorithms or software that are central to the research but not yet described in published literature, software must be made available to editors/reviewers. We strongly encourage code deposition in a community repository (e.g. GitHub). See the Nature Research guidelines for submitting code & software for further information.

Data
Policy information about availability of data All manuscripts must include a data availability statement. This statement should provide the following information, where applicable: -Accession codes, unique identifiers, or web links for publicly available datasets -A list of figures that have associated raw data -A description of any restrictions on data availability The coordinate of the hMESH1 D66K-NADPH complex has been deposited at the PDB databank, with the accession number of 5VXA.