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Glycolytic flux control by drugging phosphoglycolate phosphatase

Targeting the intrinsic metabolism of immune or tumor cells is a therapeutic strategy in autoimmunity, chronic inflammation or cancer. Metabolite repair enzymes may represent an alternative target class for selective metabolic inhibition, but pharmacological tools to test this concept are needed. Here, we demonstrate that phosphoglycolate phosphatase (PGP), a prototypical metabolite repair enzyme in glycolysis, is a pharmacologically actionable target. Using a combination of small molecule screening, protein crystallography, molecular dynamics simulations and NMR metabolomics, we discover and analyze a compound (CP1) that inhibits PGP with high selectivity and submicromolar potency. CP1 locks the phosphatase in a catalytically inactive conformation, dampens glycolytic flux, and phenocopies effects of cellular PGP-deficiency. This study provides key insights into effective and precise PGP targeting, at the same time validating an allosteric approach to control glycolysis that could advance discoveries of innovative therapeutic candidates.

Inhibiting glycolysis is an important strategy to modulate immune responses and to treat cancer, but therapeutic progress has been limited so far 1-3 . It is an open question whether all metabolic targets of interest have been exhausted 3 . Recently, a previously unrecognized layer of control has emerged with the identification of metabolite repair enzymes that maintain proper flux in central carbon metabolism 4,5 , but pharmacological approaches to target these enzymes are still lacking.
Metabolic enzymes, including those involved in glycolysis, have long been regarded as perfect catalysts, but it is now clear that they can process non-cognate substrates, catalyze different types of reactions, or mediate side-reactions that produce accidental metabolites 6,7 . Such metabolic by-products can be wasteful or even toxic, either because they are inherently reactive, or because they resemble main substrates or effectors of other enzymes and cause their inhibition. Numerous enzymes have now been identified that specifically repair problematic metabolites and thus mitigate metabolite damage 4,5 . In glycolysis alone, 10 such metabolite repair proteins work in conjunction with 10 glycolytic core enzymes to maintain proper pathway flux, and inborn errors in some of these repair enzymes can cause severe human diseases 8,9 . Specific chemical tools to target repair enzymes in glucose metabolism are largely lacking, but could facilitate further studies on their roles in physiology and disease.
Phosphoglycolate phosphatase (PGP, also called glycerol 3-phosphate phosphatase 10 or AUM 11 ) is an evolutionarily conserved metabolite damage control enzyme that can dephosphorylate and thereby repair three adventitious glycolytic metabolites: 2-phospho-L-lactate, 4-phosphoerythronate 12 , and 2-phosphoglycolate [12][13][14] . An accumulation of 2-phospho-L-lactate or of 4-phosphoerythronate inhibits glycolysis or pentose phosphate pathway flux, respectively 12 . PGP is the only mammalian enzyme that has been described to act on 2-phosphoglycolate, 2-phospho-L-lactate and 4-phosphoerythronate, and there are no known functional overlaps with other metabolite repair processes in the context of glycolysis 8,12 (see also Supplementary Note 1). The whole-body genetic inactivation of Pgp in mice caused lethal growth defects in midgestation and blocked the proliferation of embryonic fibroblasts, whereas the targeted PGP inactivation in e.g. endothelial cells during embryogenesis and adulthood did not lead to an overt phenotype 13 . These observations are consistent with a fundamental role of PGP in balancing glycolytic and pentose phosphate pathway flux in rapidly proliferating cells, thus suggesting that PGP could be an interesting target to probe the physiological and pathological roles of a glycolysis-associated metabolite repair protein.
PGP belongs to the haloacid dehalogenase (HAD)-type superfamily of phosphatases, a large, widespread and evolutionarily ancient class of hydrolases with emerging roles in human physiology and disease [15][16][17][18] . PGP consists of a catalytic core harboring a Rossmanoid fold, and a large cap domain restricting access to the active site 11 . Commonly used phosphatase inhibitors do not inhibit HAD phosphatases, and natural inhibitors that may serve as lead compounds are unknown. Furthermore, chemical tools to specifically target mammalian HAD phosphatases are scarce [19][20][21] , and small molecules that selectively inhibit PGP have not yet been described.
Here, we report the identification and extensive biochemical, structural and functional evaluation of CP1, a specific small-molecule PGP inhibitor that blocks PGP-dependent glycolytic flux in mammalian cells. Our study thus validates a pharmacological target class and lays the groundwork for the development of innovative therapeutic approaches to target cellular metabolism.

A high-throughput screening campaign identifies PGP inhibitors
We screened the FMP small molecule repository containing 41,536 compounds for molecules able to modulate the phosphatase activity of recombinant, highly purified murine PGP (Supplementary Fig. 1a; see Methods for further details). Difluoro-4-methylumbelliferyl phosphate (DiFMUP) was used as a fluorogenic phosphatase substrate in a primary screen, and a counterscreen was performed with pyridoxal 5'phosphate phosphatase (PDXP), the closest PGP relative 11 . The PGP inhibitor hits that were inactive against PDXP were subsequently validated in a secondary assay, using 2-phosphoglycolate (PG) as a physiological PGP substrate. IC 50 values were determined for the 23 PGP inhibitor hits that blocked PG dephosphorylation by >50% (Supplementary Table 1), and three of the most potent PGP inhibitors (compounds 1-3/CP1-3) were selected for further characterization.
As shown in Fig. 1a, CP1-3 potently inhibited the activity of both human and murine PGP towards PG. CP1-3 also inhibited the activity of murine PGP towards the generic phosphatase substrate paranitrophenylphosphate (pNPP). Furthermore, CP1 effectively inhibited the dephosphorylation of 2-phospho-L-lactate (2PL), 4-phosphoerythronate (4PE) and glycerol 3-phosphate (G3P; see also Supplementary Fig. 1b). We confirmed that CP1-3 did not inhibit PDXP activity towards pyridoxal 5'-phosphate (PLP) 22 . CP1-3 were also able to block PG dephosphorylation catalyzed by an artificial protein consisting of the PDXP catalytic core and the PGP specificitydetermining cap (PDXP/PGP hybrid 11 ). However, for CP1,~37-fold higher concentrations were required for a half-maximal inhibition of the murine PDXP/PGP hybrid compared to full-length murine PGP. These data suggest that CP1-3 interact with the PGP cap domain, and that binding to both the cap and the core domain is essential for an optimal inhibition of PGP by CP1.
To exclude the possibility that the inhibitory effects of CP1-3 were caused by protein denaturation, we investigated the thermal stability of PGP in the absence or presence of increasing compound concentrations by differential scanning fluorimetry. Supplementary Fig. 1c shows that the inhibitors in fact stabilized PGP in a concentrationdependent manner, as reflected by an increase in the maximal PGP melting temperature from 66.4°C (control conditions) by 4.6°, 2.2°, or 7.1°for CP1, −2, or −3, respectively. Thus, the observed PGP inhibition was not due to protein destabilization.
We performed isothermal titration calorimetry (ITC) experiments to further characterize inhibitor binding to PGP. CP1 and CP3 showed a relatively high-affinity interaction with PGP characterized by K D -values of 0.78 and 0.43 μM, respectively, and bound PGP with monovalent binding stoichiometries (Fig. 1b). CP2 binding to PGP could not be analyzed by ITC due to its limited solubility.
Structure-activity relationships CP1 and CP2 are structurally related diphenylether and diphenylmethylene dibenzoic acid derivatives with similar inhibitory properties (Fig. 1a). Taking these hits as model compounds, we examined whether commercially available analogs might be more potent PGP inhibitors. We tested a pyrrolidinedione-derivative (1B), two isomers (1C and 1D) and two isomeric fragments (1E and 1F) of CP1, as well as four CP2 analogs ( Supplementary Fig. 2a) in the PG dephosphorylation assay. However, none of the tested CP1 analogs were anywhere as potent as CP1 ( Supplementary Fig. 2b). In contrast, the CP2 analogs with the exception of 2E still retained inhibitory potency, with 2B being even slightly more potent than the parental compound (Supplementary Fig. 2b).

Selectivity of PGP inhibitors
PGP is a member of the vast and ancient class of haloacid dehalogenase (HAD)-type phosphatases. The overall structure of HAD phosphatases and their catalytic residues are highly conserved. All HAD phosphatases consist of a catalytic core domain with a Rossmanoid-fold. This core is accessorized with so-called cap domains, which are mobile elements that can shield the entry to the catalytic cleft. The cap size (tiny C0 caps versus larger C1 or C2 type caps), their structure and insertion site determine the substrate binding pocket of the respective enzyme: C0-type HAD phosphatases can dephosphorylate macromolecules, whereas C1/C2-type HAD phosphatases are typically directed against small molecules that are able to enter the catalytic cleft. In addition, the caps contain substrate specificity loops that interact with the substrates. Hence, cap types are key differentiators of HAD phosphatases 15,16,23,24 .

Mode of PGP inhibition
To probe the mechanism of PGP inhibition, we assayed the steady state kinetics of PG dephosphorylation in the presence of increasing CP1-3 concentrations (Supplementary Table 2). Analysis of the derived kinetic constants suggested different modes of PGP inhibition. CP1 increased the K M up to~3-fold, and reduced v max values~3-fold, whereas CP2 and −3 increased the K M between~3-fold without affecting v max . Thus, CP1 shows a mixed mode of inhibition (apparently between competitive and non-competitive), and CP2 and −3 act as competitive inhibitors.

Observation of open and closed active site conformations in PGP
To understand mechanisms of PGP inhibition on a molecular level, we set out to obtain structural information of PGP in complex with an inhibitor. As a first step, we defined suitable crystallization conditions for full-length PGP, whose structure was previously unknown. We had shown earlier that PGP exists in a dynamic equilibrium between homodimers and -tetramers 11 , and that the mutation of Cys297 to Ser (PGP C297S ) abolished disulfide-mediated PGP tetramer formation 14 . We therefore hypothesized that PGP C297S might be a suitable PGP variant for crystallization. We verified that PGP C297S was enzymatically active and sensitive to compound 1 (PGP C297S IC 50~0 .25 ± 0.05 µM; Supplementary Fig. 3a, b), and that CP1 did not affect PGP C297S self-association (Supplementary Fig. 3c; see below). For subsequent crystallization attempts, we additionally exchanged the catalytically essential Asp34 to Asn, which abolishes PGP activity towards its substrates 11 . Full-length murine PGP D34N/C297S (here referred to as apo-PGP) crystallized in space group P1 with four dimers per asymmetric unit ( Supplementary Fig. 4), referred to as AB, CD, EF and GH dimers. The structure was solved by molecular replacement with the murine PDXP/PGP hybrid protein 11 (PDB code 4BKM). The structure was rebuilt and refined against data truncated anisotropically with Staraniso (http://staraniso.globalphasing.org/cgi-bin/staraniso.cgi) to 2.31/2.74 Å in the best/worst direction, resulting in an R work of 0.2068 and an R free of 0.2308 (PDB code 7PO7). Data collection and refinement statistics are summarized in Table 1. While monomers A-F and H are well defined in the electron density maps, large sections of the catalytic core of monomer G display very weak density. The electron density map of the active site of monomer E is shown in Supplementary Fig. 5. Figure 2a shows the structure of the EF-dimer of full-length apo-PGP. Like its paralog PDXP 22 and the PDXP core/PGP cap hybrid protein 11 , PGP homodimerizes via its cap domain. Aligning the core domains (Cα atoms of residues 11-99 & 238-320) of the B-F and H monomers (the G-monomer was omitted from the analysis due to its poor density for the core domain) on the A-monomer resulted in a close superposition with values for the root mean square deviation (RMSD) between 0.41 Å and 0.50 Å for the Cα atoms for monomers B, C, F and H and slightly higher deviations of 0.58 Å and 0.62 Å for monomers D and E. The disparity in the alignment for monomers D and E correlates with the presence of a distinct conformation of the cap domain in both monomers. An analysis with the DynDom server 25 revealed that in five (chains A-C and F-H) of the seven PGP monomers which were part of the analysis, the cap domain was rotated bỹ 11-15°relative to the conformation present in the other two monomers (chains D and E). Coupled to this rotation is a shift of the substrate specificity loop (amino acids 202-215) by up to~6 Å for corresponding Cα atoms towards the catalytic core. Conversely, the cap domains (Cα atoms of residues 100-237) can be superimposed, which then identifies a movement of the PGP 'flap' (amino acids 39-46 of the core domain) by >5 Å towards the catalytic core (Fig. 2b, c). The conformational changes do not appear to be induced by crystal packing forces since the cap domains of monomers D and E as well as that of monomer A, which together represent both conformations, are hardly involved in crystal packing contacts in contrast to the more extensively participating monomers B, C, F and H ( Supplementary Fig. 4).
Flap and cap movements are known to gate access to the catalytic site in C1-capped HAD phosphatases 26 . An inspection of the PGP catalytic cleft showed that the active site of PGP is organized as a channel, and that Asn34 (corresponding to the Asp34 nucleophile) is centrally buried within this cleft (Fig. 2d). Upon rotation of the substrate specificity loop and the flap, channel entrances become obstructed, and access to the active site is blocked. To our knowledge, experimental proof for the existence of open (chains D and E) and closed (chains A-C, F and H) conformations in a C2-capped mammalian phosphatase has not been previously provided.
Co-crystal structure of PGP bound to CP1 Because of its relatively high potency, selectivity profile, and highaffinity binding of CP1 to PGP (Fig. 1), we focused our structural analyses on this inhibitor. CP1-bound PGP crystallized in the tetragonal space group P4 1 with one dimer per asymmetric unit. The structure was solved by molecular replacement with the AB dimer of the apo-PGP structure, followed by refinement against anisotropically truncated data (again with Staraniso) to resolutions of 3.16/3.41 Å in the  amplitudes. f R free same as R work for 2.5% (apo) and 5% (inhibitor) of the data randomly omitted from refinement. g Ramachandran statistics indicate the fraction of residues in the favored, allowed, and disallowed regions of the Ramachandran diagram as defined by MolProbity 71 . best (a*b*-plane) and worst direction (along c*), resulting in an R work of 0.1984 and an R free of 0.2420 ( Fig. 3a and Table 1; PDB code 7POE). Already after molecular replacement both protomers showed clear density for CP1 (Fig. 3b). The electron density map of the active site of CP1-bound monomer A is shown in Supplementary Fig. 6. The temperature factors of the atoms in CP1 were similar in magnitude to those of the surrounding atoms, therefore suggesting that the inhibitor is bound with an estimated occupancy of~1. CP1 binds adjacent to, but not directly into, the PGP active site with one of the benzoic acid moieties (referred to in the following as proximal) at a distance of just below 6 Å from the catalytic Asp34 (Asn34). Inhibitor binding appears to be primarily stabilized by nonpolar interactions and a very small number of polar interactions (Fig. 3c). The proximal benzoic acid moiety of CP1 forms the only direct hydrogen bonds via its carboxylic acid to the nitrogen atom of the carboxamide function of Asn199 and to the side chain hydroxyl group of Thr215. At the limited resolution of this structure these contacts should be considered potential interactions. Overall, CP1 is embedded in a cavity that extends from the active site of monomer A towards the dimerization interface with monomer B. One side of this cavity is formed by the more polar residues 35-37, 68-71, 125-127 and 143-145, which originate entirely from monomer A. In contrast, the opposite side is formed by both monomers and is generally more hydrophobic in character. Key residues include stretches 172-175, 201-204/212-215 within monomer A, alongside residues 151-156 of monomer B. The primary driving force for binding seem to be van der Waals interactions involving the side chains of the non-polar residues Pro127, Phe172, Pro174, Leu204 and Trp156 of monomer A. In addition, the non-polar atoms of Asp36, Asn69, Ser71 and His175 engage in van der Waals interactions with CP1. All PGP residues found to engage in CP1 interactions (see also Fig. 4) are identical in murine and human PGP ( Supplementary Fig. 7). Towards the end of the refinement, residual density could be tentatively interpreted with a second CP1 molecule, which is binding to each monomer on the opposite side of PGP. The binding site is quite remote from the active site (closest distance of~15 Å) and is in close proximity to the non-crystallographic twofold axis of symmetry. While the ITC data indicate binding of a single copy of CP1 to each monomer ( Fig. 1), it cannot be ruled out that this second binding event does not lead to a change in enthalpy. The crystallographic evidence documents, however, that the second CP1 molecule is weakly bound and hence no biological significance was attributed to the second binding site.
To verify the putative CP1 -PGP interactions in the primary binding site, we introduced single or multiple mutations into the binding interface by exchanging PGP residues for the corresponding amino acids of the CP1-insensitive PGP paralog PDXP (Fig. 3d). Hence, Ser71 was changed to the large, positively charged Arg (PGP S71R ), Pro127 was altered to Glu (PGP P127E ), and the hydrophobic sequence 172 FDPH was swapped with the respective residues in PDXP (PGP 172FDPH>YDEQ ). In addition to the S71R variant, we also mutated Phe172 (facing Ser71 on the opposite side of the active cleft) to the less hydrophobic Trp (PGP F172W ). Figure 3d (left panel) shows that all mutations resulted in a strongly reduced potency of CP1, supporting the role of these residues in inhibitor binding. Exchanging both Ser71 and Phe172 (PGP S71R; F172W ) or ablating the hydrophobic pocket (PGP 172FDPH>YDEQ ) increased the CP1 IC 50 value~600-fold compared to PGP WT . These data are consistent with the binding mode of CP1: a replacement of Ser71 with Arg obstructs the binding site, a Glu at position 127 as well as an Asp at position 174 (both instead of a Pro) are electrostatically repulsive with respect to the distal carboxylate of CP1, and a Trp instead of Phe172 is sterically too demanding for CP1 binding. Figure 3d (right panel) shows that also the phosphatase activity of PGP S71R was strongly decreased. This indicates a key role of Ser71 for active site accessibility. The almost complete loss of phosphatase activity in PGP-Phe172 mutants (PGP S71R;

F172W
; PGP 172FDPH>YDEQ ) might likewise be explained by impeded substrate entry into the active site channel due to side chain interference and charge repulsion effects. Interestingly, the phosphatase activity of PGP P127E was comparable to PGP WT . Furthermore, ITC experiments demonstrated that the affinity of CP1 for PGP P127E was 15-fold lower than for PGP WT (K D 11.93 ± 0.41 µM; n = 1.15 ± 0.04; mean values ± S.E.M. of two independent experiments; see Fig. 1b for comparison). This result is in line with the higher IC 50 of CP1 for PGP-P127E compared to PGP-WT (Fig. 3d, left panel). Hence, PGP P127E represents a PGP variant that retains full phosphatase activity while being CP1-resistant.
The comparison of the CP1-bound PGP structure with apo-PGP conformers with the DynDom server 25 revealed that CP1 -PGP adopts a conformation that resembles the cap-closed state (Fig. 3e). Domain movements were recognized for both subunits of the inhibitor-bound structure only with respect to chains D and E of the apo-structure, thus indicating that both monomers of the CP1-bound structure and the majority of apo-PGP monomers adopt the same conformational state. Despite being in the same conformational state, there are differences between the closed state in the apo and inhibitor bound structures, e.g. the substrate binding loop, rather than moving solely towards the active site, is also translated outwards. This additional movement is needed to create the space required for incorporating CP1 (Fig. 3e).
Hence, CP1 appears to inhibit PGP activity by locking it in a cap-closed conformation.
Based on the inhibitor-bound structure and the competitive component of PGP inhibition by CP1 (increased K M , see Supplementary  Table 2), it seems likely that CP1 sterically hinders substrate access to the active site. In addition, the cap-closed conformation of CP1-bound PGP will block solvent entry into the enzymatic pocket, and thus prevent the hydrolysis of the phospho-aspartate intermediate during the final catalytic step. The resulting reduced rate of product formation is consistent with the observed non-competitive component of CP1-mediated PGP inhibition (reduction of v max , see Supplementary Table 2).

CP1 locks PGP in a closed-cap conformation during molecular dynamics simulations
To further investigate the PGP conformations as well as the interactions of CP1 at the active site, we carried out molecular dynamics (MD) simulations. For this purpose, PGP was restored to the wild-type active site with Asp34 and a Mg 2+ ion (see Methods). The crystal structures of the homodimeric apo-PGP (chains C and D) and CP1-bound PGP (chains A and B) were used as starting conformations to sample two 100 ns trajectories. The binding mode of CP1 to the wild-type active site was obtained by docking calculations, which confirmed the crystallographically obtained binding mode with an RMSD of~0.5 Å (see Methods and Supplementary Figs. [8][9][10][11]. Analyzing the protein RMSD profile along the trajectory based on different fittings using either the entire dimer or the individual monomers (Fig. 4b) revealed the overall stability of the monomers, but considerable fluctuations and conformational changes across the dimer interface, where the caps are located (see Supplementary Figs. 12-13 for further characterization of the dynamics by means of RMSF profiles and principal component analyses, and Supplementary Movies 1-4 related to Supplementary  Fig. 13). Within these cap domains, the substrate specificity loop (SSL; responsible for the substrate access gate) was predominantly in a closed state in the CP1-PGP simulation, whereas it moved and sampled the open form in the simulation of the apo structure (Fig. 4a). These results are consistent with the aforementioned findings of an open and closed PGP conformation, where the active site is locked by CP1. Although CP1 undergoes some configurational and conformational changes along the trajectory, as indicated by the 2D-RMSD plots of the ligand in each active site (Fig. 4d), the superimposed snapshots illustrate that CP1 remains at its binding site, occupying the same area as in the reference structure (Fig. 4a, lower right panel).
To clarify which interactions are formed and stably maintained by CP1 and how this inhibitor is supporting the cap-closed PGP state, we measured CP1-PGP interaction distances for putative hydrogen bonds and van der Waals interactions ( Supplementary Fig. 14). Interestingly, the crystallographically observed hydrogen bonds of the proximal carboxylic group with Asn199 and Thr215 were not stably maintained. Instead, hydrogen bonds with the side chain atoms of Ser70 and Ser71 were formed after initial fluctuations and moderate conformational changes (Fig. 4c) and observed for almost the entire second half of the trajectory in each monomer (Supplementary Fig. 12 and Supplementary Table 3). The diphenylether moiety engages in continuous interactions with Phe172, leading to distances around 5 Å between the centers of the aromatic rings. The distal benzoate moiety shows some early oscillations in the distances to His175, but rapidly stabilizes in between the side chains of Pro127 and His175, apparently supported by temporary hydrogen-bond formation of the adjacent amide with the side chain of Ser126, as observed in 64-81% of the trajectory.

CP1 impairs glycolytic flux
To assess the impact of CP1 on central carbon metabolism, we profiled cellular metabolites by untargeted nuclear magnetic resonance (NMR) spectroscopy. As a cellular model system, we used human fibrosarcoma HT1080 cells, a line whose malignant phenotype is driven by an activated N-ras oncogene 27 . HT1080 cells are known to form highly aggressive tumors in murine xenograft models 28 , suggesting potential translational relevance 29 . We compared the effects of CP1 with PGP wildtype (PGP-WT, clone D5) or PGP knockout (PGP-KO, clone C2) HT1080 cells generated with the CRISPR/Cas9 system. Strikingly, the majority of the 45 identified metabolites was significantly different in CP1-versus mock-inhibited cells, and in PGP-WT versus PGP-KO cells. Sparse partial least squares discriminant analyses (sPLS-DA; Supplementary Fig. 15) of the metabolomes clearly separated both groups in the two trials, and the ten metabolites that contributed to the decisive principal component 1 were also detected as significantly different in the trial groups by biomarker analysis. Pathway enrichment analyses and topology scoring indicated significant effects on 'glycolysis and gluconeogenesis', among other pathways (Supplementary Tables 4-7; Supplementary Fig. 16).
Because of the conserved role of PGP for glycolytic flux 12 , we next compared the effects of CP1 inhibition and PGP deletion on glycolysis in detail. PGP is known to dephosphorylate the pyruvate kinase side product 2-phospho-L-lactate (2PL), thereby preventing the 2PLmediated inhibition of phosphofructokinase-2 (PFK-2). Through this mechanism, PGP safeguards the PFK-2-catalyzed formation of fructose-2,6-bisphosphate, an allosteric activator of the key glycolytic regulator phosphofructokinase-1 (PFK-1), thus contributing to the maintenance of glycolytic flux 12 (Fig. 5a). Hence, the loss of PGP activity or -expression is expected to reduce the formation of the glycolytic end products pyruvate and lactate. While 2-PL was not resolved in our NMR spectra, we found significantly reduced pyruvate and lactate levels in both CP1-inhibited and PGP-deficient cells, consistent with a reduced glycolytic flux upon PGP-targeting (Fig. 5b). CP1 treatment (3 h, 33 µM CP1; see Supplementary Note 2 and Supplementary Discussion for an explanation of the required CP1 concentrations) reduced pyruvate levels (−10%), lactate levels (−45%) and acetate levels (−42%; potentially reflecting pyruvate decarboxylation 30 ) (Fig. 5b). Since alanine levels were also significantly lower in CP1-inhibited cells (−52% after 3 h; Supplementary Fig. 17), pyruvate levels may be buffered by the gluconeogenic formation of pyruvate from alanine under these conditions. Finally, CP1 treatment or PGP-KO markedly decreased cellular glucose concentrations (−30%, −61%, −74% in cells treated for 3 h with 33 µM CP1, for 24 h with 100 µM CP1, or in PGP-KO cells, respectively; Fig. 5b and Supplementary Fig. 17), in line with the previously reported decrease in the consumption of extracellular glucose observed in PGP-KO HCT116 cells 12 . In sum, these data support the conclusion that CP1 diminishes glycolytic flux, as expected for a PGP inhibitor.

CP1 impairs PGP-dependent cell proliferation
Glucose consumption and -catabolism, including flux through the pentose phosphate pathway, are intimately connected with proliferative metabolism 31 . Indeed, PGP phosphatase-inactivated mice (Pgp D34N/D34N ) die in utero with severe growth defects, and murine embryonic fibroblasts obtained by dissociating viable E8.5 Pgp D34N/D34N embryos do not proliferate under standard in vitro culture conditions 13 , indicating that PGP activity can influence cell proliferation. To characterize the suitability of CP1 as a tool to modulate cellular PGP functions, and to address the on-target activity of this compound, we next investigated cell proliferation.
The comparison of PGP-WT and PGP-KO HT1080 cells (see Supplementary Fig. 18a for protein blots) showed that PGP deletion reduced the proliferation of single cell clones derived from two different single guide RNAs (gRNAs), or from five pooled gRNAs in a comparable manner (Supplementary Fig. 18b, left and middle panel). CP1 was not cytotoxic at the employed concentration of 100 µM ( Supplementary Fig. 18c), but decreased the proliferation of PGP-WT cells to the level of PGP-KO cells, which were insensitive to CP1 treatment ( Supplementary Fig. 18b, middle panel). Despite this initial proliferation defect, PGP-KO C2 cells eventually proliferated like their PGP-WT counterparts ( Supplementary Fig. 18b, right panel), suggesting that cells can adapt to PGP deficiency (see also Supplementary Note 3). The adaptation of PGP-KO C2 cells reproducibly happened around passage 15, and occurred each time after the PGP-KO C2 cells were put into culture. Nevertheless, long-term PGPdeficient cells remained insensitive to CP1, and CP1 inhibited the proliferation of PGP-WT cells overexpressing wildtype PGP, but not of PGP-WT cells expressing the CP1-insensitive PGP-P127E variant ( Supplementary Fig. 18b, right panel).
To better evaluate the on-target activity of CP1, we additionally characterized a CRISPR/Cas9 PGP-knockdown clone (PGP-KD clone B1-10;~25% remaining PGP expression; Fig. 5c). Similar to PGP-KO cells, PGP-KD cells proliferated less than their WT counterparts (Fig. 5c). CP1-treatment decreased the proliferation of PGP-WT, but not of PGP-KD cells. The re-expression of PGP-WT in PGP-KD cells normalized their proliferation to the level of PGP-WT cells, and CP1 treatment of these cells again inhibited their proliferation. Importantly, the expression of the enzymatically active, but CP1-resistant PGP variant PGP-P127E (Fig. 3d) in PGP-KD cells reverted their proliferation back to the level of PGP-WT cells, but did not restore responsiveness to CP1 (Fig. 5c). These findings strongly support the conclusion that CP1 mitigates cell proliferation in a PGP-dependent manner.

CP1 induces PGP-dependent lipid droplet accumulation
Given the PGP-dependent block in glycolytic flux, we finally investigated the impact of CP1 and PGP-KO on a cellular nutrient stress response. Upon energy stress, cells remodel their metabolism and shift from a reliance on glycolysis to fatty acid breakdown for survival 32 . Fatty acids are stored as triacylglycerides (TAGs) in intracellular lipid droplets (LDs) 33,34 . During a cellular energy crisis, the abundance of LDs increases in an adaptive response to buffer lipotoxicity and ER stress, and to increase cellular viability 35,36 . Indeed, we have previously observed that genetic Pgp-inactivation or PGP knockdown leads to intracellular TAG and LD build-up in mice, rats, and murine cell lines 10,13,37 .
Supplementary Figure 18d shows that CP1 triggered a time-and concentration-dependent accumulation of LDs in HT1080 cells. To test whether CP1-induced LD formation was dependent on PGP inhibition, we re-examined the drug response in PGP-deficient cells. Figure 5d shows that CP1-treatment effectively increased LDs in PGP-WT but not in PGP-KO cells, whose mean basal LD content resembled CP1-treated PGP-WT cells. The re-expression of PGP-WT in PGP-KO cells decreased their LD content to control cell levels, whereas CP1 treatment of these cells again triggered a strong LD accumulation. In contrast, the expression of the CP1-insensitive mutant PGP-P127E in PGP-KO cells reverted the elevated LDs of PGP-KO cells back to WT levels, but did not restore their responsiveness to CP1 (Fig. 5d). In sum, these results strongly argue that the observed LD accumulation was caused by CP1induced PGP inhibition, and validate CP1 as a pharmacological inhibitor of PGP.

Discussion
The discovery of metabolite repair proteins has revealed additional layers of control in central carbon metabolism 4,8 , but target molecules that can be pharmacologically addressed to modulate pathway flux remain to be defined. Here, we demonstrate the druggability of PGP, a haloacid dehalogenase-type phosphatase and prototypical metabolite repair enzyme that is required to prevent a block in the pentose phosphate pathway and glycolysis 12 . We identify and characterize CP1 as a selective small-molecule PGP inhibitor. Detailed structural analyses reveal that CP1 locks PGP in a catalytically incompetent conformation by targeting a hydrophobic pocket located between the enzyme's core and cap domains. This intriguing mechanism may be broadly applicable to capped HAD phosphatases, and facilitate the future design of highly selective HAD phosphatase inhibitors.
Although phosphatases are considered notoriously difficult to target, the successful development of allosteric protein phosphatase inhibitors is now finally changing this view 38,39 . While only a few mammalian HAD phosphatases have been studied in depth, substantial progress has been made in targeting e.g. EYA2 tyrosine phosphatase in cancer 19,40 . Importantly, HAD phosphatases can show remarkable intrinsic substrate specificity, and do not require an association with regulatory or targeting subunits to impart substrate selectivity 11,26,41 . Inhibiting the activity of a particular HAD phosphatase therefore offers an approach to precisely modulate a specific substrate or small set of  substrates. As such, HAD phosphatases conceptually represent a rich source of pharmacologically largely untapped targets. CP1 treatment of HT1080 human fibrosarcoma cells recapitulates three previously described effects of PGP loss: a reduction of glycolytic flux 12 , a decrease in cell proliferation 12,13 , and an increase in lipid droplet formation 13,37 . We therefore anticipate that CP1 will be a useful tool compound to probe the functional consequences of acute PGP inhibition. The broader implication of this work lies in the interest in innovative approaches to target cellular metabolism. Future studies in e.g. a range of cancer cells and suitable disease models will need to evaluate whether the approach of blocking a metabolite repair protein such as PGP to control flux through the pentose phosphate pathway and glycolysis is effective with an adequate therapeutic index. If this turns out to be the case, perhaps more potent PGP inhibitors could be exploited therapeutically to modulate e.g. immune responses in inflammation, autoimmunity or cancer 2 .

Expression and purification of recombinant phosphatases
All constructs for bacterial protein expression and purification contained an N-terminal His 6 -tag and were in pETM11 (EMBL). Constructs were transformed into Escherichia coli BL21 (DE3) (Stratagene); to increase solubility, proteins were co-expressed with the chaperones groES-groEL-tig from the pG-Tf2 plasmid (Takara Bio Inc.) according to the manufacturer's instructions. All purification steps were carried out at 4°C. PGP was expressed for 20 h at 28°C after induction with 0.5 mM isopropyl β-D-1-thiogalactopyranoside (IPTG). Cells were harvested at 8000 × g for 10 min and resuspended in TNM-1 (50 mM triethanolamine, 200 mM NaCl, 5 mM MgCl 2 ; pH 7.5) supplemented with 10 mM imidazole and protease inhibitors (EDTA-free protease inhibitor tablets; Roche Applied Science). Cells were lysed in the presence of 150 units/mL DNase I (Applichem) using a cell disruptor (Constant Systems), and cell debris was removed by centrifugation (10,000 × g, 30 min, 4°C). Cleared supernatants were loaded on a HisTrap HP nickel sepharose column (Cytiva) operated on an ÄKTA liquid chromatography system equipped with Unicorn 5.11 software (GE Healthcare) in TNM-1, and His 6 -tagged proteins were eluted using a linear 10-400 mM imidazole gradient in TNM-1. PGP peak fractions were pooled, and the His 6 -tag was cleaved with TEV protease for 2-3 days at 4°C. Subsequently, cleaved PGP was separated from uncleaved PGP and the His-tagged TEV protease on a HisTrap HP nickel sepharose column. Fractions containing untagged PGP were pooled, concentrated (10 kDa MWCO; Amicon Ultra-15, Millipore), further purified on a HiLoad 16/60 Superdex 200 pg size exclusion chromatography column (GE Healthcare) and eluted in TNM-1.
Murine PGP C297S (ref. 14) was obtained by gene synthesis (GeneArt/ Thermo FisherScientific). The D34N mutation was subsequently introduced into murine PGP C297S by nested PCR, using the Pgp D34N primers listed above under Molecular Cloning. PGP D34N; C297S was purified and the His 6 -tag was removed as described above for PDXP protein with the following modifications: cultures were incubated for 20 h at 28°C, protein overexpression was induced by addition of IPTG to a final concentration of 1 mM, and metal affinity chromatography was performed using a HisTrap excel nickel sepharose column (Cytiva).

High-Throughput Screen for PGP Modulators
Chemical library: The FMP small molecule repository used for primary screening consisted of 41,536 compounds in total, divided into subsets with compounds that represent a structural diversity set (33,088 compounds comprising a World Drug Index-derived collection of small molecules structurally related to about 500 different core scaffolds of drugs, according to the concept of maximumcommon-substructures 42 ; compounds with known pharmacological activity (LOPAC), FDA-approved drugs, and drug candidates which failed in clinical trials (3168 compounds). In addition, the library contains compounds obtained from academic donators world-wide (5280 compounds).
High-throughput screening protocol: Compounds were screened at a final concentration of 10 µM in a total reaction volume of 30 µL in black, non-transparent 384-well assay plates (Corning, #3573). Measurements were conducted in the following assay buffer: Triethanolamine (30 mM), NaCl (30 mM), MgCl 2 (5 mM), DTT (1 mM), Triton X-100 (0.01% v/v), pH 7.5. DTT was included as a reducing agent to suppress the redox-dependent formation of PGP oligomers and to ensure maximal PGP activity 14 . For measurements using PGP, final concentrations of 10 µM DiFMUP and 4 ng/µL (0.12 µM) PGP enzyme were used (the K M value of DiFMUP could not be determined as a saturation could not be observed with DiFMUP concentrations up to 400 µM). For measurements using PDXP, 5 µM DiFMUP and 10 ng/µL (0.3 µM) PDXP were used (K M~5 .1 µM). For both target proteins, screening was therefore conducted at a substrate concentration corresponding to K M or lower. DiFMUP and target protein solutions were prepared as 3-fold concentrated solutions in assay buffer. Using a microplate dispenser (MicroFlo Select Microplate Dispenser, Biotek), 10 µL of assay buffer were dispensed into columns 1-24 of a 384-well microtiter plate. 0.3 µL of compound solution were transferred from 384-well compound stock plates (containing compounds dissolved to 1 mM in DMSO in columns 1-22 and DMSO in columns 23 and 24) to the assay plates using a liquid handling workstation equipped with a 384channel pipetting head and disposable tips (Biomek FxP, Beckman Coulter). 10 µL of enzyme solution was dispensed into columns 1-23 of the assay plate, and 10 µL of assay buffer was added to column 24 using a manual multi-pipet. This pipetting scheme created control samples in column 23 with enzyme and DMSO (100% enzyme activity) and control samples in column 24 without enzyme (0% enzyme activity). The plates were then incubated at room temperature for 15 min to allow binding of the compounds to the enzyme. Finally, 10 µL of DiFMUP substrate solution were dispensed to columns 1-24 of the assay plate, the plate shaken and centrifuged with a quick spin to remove air bubbles, and immediately read in a microplate reader (Safire II, Tecan). Fluorescence was measured from the top with an excitation wavelength of 360 nm and emission wavelength at 460 nm (20 nm bandpass each). Five time points were measured over a duration of 9 min. The PerkinElmer LabChip-3000 software was used to generate the .csv flat text files containing the substrate/product ratio.
Concentration-dependent assays: 5 µL of concentrated stock solutions (10 mM in DMSO) of compounds active in primary screening were picked onto a new compound mother plate by the FMP compound management facility, and serial 2-fold inter-plate dilutions were created as follows: using a liquid handler equipped with a 384-channel pipetting head, 5 µL of DMSO were added to the compound mother plate, the samples mixed using the pipetting head, and 5 µL of the diluted compound samples were transferred onto a new compound mother plate. This process was repeated 8 times, thereby creating a set of 9 compound mother-plates with decreasing concentrations. These serially diluted compound plates were then assayed in 2 technical replicates as described above, giving concentration-dependent activity data in the concentration range from 0.2 µM to 50 µM. Because the highest compound concentration used was 50 µM, we defined this value as an IC 50 cutoff to ensure that the IC 50 estimate is an interpolation of generated data, according to the recommendations of the NIH Assay Guidance Manual (https://www.ncbi.nlm.nih.gov/books/ NBK91994/#assayops.Determination_of_EC50IC50).
Data analysis: Using in-house developed software, the slope (as indicator of the enzyme activity) and intercept (as indicator of compound autofluorescence) were derived from the kinetic trace measured in each well. The slopes from the 100%-activity and 0%-activity control samples of each plate were used to calculate the Z'-factors of the plates as a quality control parameter for the assay. Data of the compound samples were normalized separately for each plate, by calculating Z-scores using all sample wells of a plate, and calculating activities relative to the 100% and 0% control samples of a plate using statistically robust estimators 43 . Final reports and IC 50 /EC 50 curve fits were then created using the Pipeline Pilot software 9.5 (Biovia).

Compound solubility and solvent controls
Compound solubilities in 100% DMSO were as follows. CP1: 100 mM; CP2: 15 mM; CP3: 50 mM. 2B: 10 mM, 2D: 50 mM, 2E: 100 mM, 2F: 50 mM. DMSO solvent control concentrations in the different assays varied depending on the requirements of the respective experiment (see below for further details). However, the DMSO concentration was kept constant across each assay. For SAR experiments, all compound stocks were prepared at 10 mM in 100% DMSO. A constant final DMSO concentration of 0.4% was maintained under all conditions, and solvent control samples contained 0.4% DMSO without compounds.

IC 50 determinations, enzyme kinetics, and compound selectivity
Experiments were generally conducted at room temperature (RT) in TMN30 (30 mM triethanolamine, 5 mM MgCl 2 , 30 mM NaCl; pH 7.5), supplemented with 5 mM DTT and 0.01% (v/v) Triton X-100, with the following exceptions: PHOSPHO1/2 activity was measured in a buffer containing 50 mM imidazole (pH 7.0), 5 mM MgCl 2 , 0.01% (v/v) Triton X-100 and 5% (v/v) glycerol. The buffer for PHOSPHO1 was further supplemented with 5 mM DTT, and its activity was measured at 37°C. Calcineurin activity against the PKA regulatory subunit type II (phosphopeptide DLDVPIPGRFDRRVpSVAAE) was assayed in calcineurin assay kit buffer (Millipore #207005); PTP-1B activity against the insulin receptor β-subunit phosphopeptide (amino acids 1142-1153, TRDI-pYETDYYRK) was measured in 150 mM NaCl, 50 mM 2-(N-morpholino) ethanesulfonic acid, 1 mM DTT, 1 mM EDTA, 0.05% (v/v) NP-40; pH 7.2. Calf intestinal alkaline phosphatase activity was assayed in 100 mM triethanolamine (pH 9.4), 1 mM MgCl 2 , 20 µM ZnCl 2 . The 3'-P-oligonucleotide 5'-ATTACGAATGCCCACTCCTC-[PO 4 3− ]−3' was used as a PNKP substrate. Purified phosphatases were pre-incubated for 5 min (for compound selectivity assays) or 15 min (for IC 50 determinations and enzyme kinetics) at RT with serial dilutions of CP1-3. A fixed concentration of 0.1% DMSO was employed for all compounds. Solvent control samples contained 0.1% DMSO without compound. Dephosphorylation reactions were started by the addition of the indicated substrate; buffer without enzyme served as a background control. Prior to compound testing, time courses of inorganic phosphate release from the respective phosphatase substrates were conducted to ensure assay linearity. 2-Phospho-L-lactate was a kind gift of Dr. Guido Bommer; 4-phosphoerythronate was from BLD Pharmatech GmbH. Inorganic phosphate release was detected with malachite green solution (Biomol Green; Enzo Life Sciences); the absorbance at 620 nm (A 620 ) was measured on an Envision 2104 multilabel reader, using the Wallac Envision Manager software 1.2 (Perkin Elmer Life Sciences). Glycerol 3-phosphate (G3P) dephosphorylation was measured with the EnzChek Phosphate Assay Kit (Thermo Fisher Scientific), and analyzed at 330/360 nm on a Clariostar multiplate reader and Clariostar software 5.21 R2 (BMG Labtech). Para-nitrophenyl phosphate (pNPP) dephosphorylation to the colored product para-nitrophenol was monitored colorimetrically on an Envision 2104 multilabel reader at A 405 . Released phosphate was determined by converting the values to nmol P i with a phosphate standard curve. Data were analyzed with GraphPad Prism 7.04. For IC 50 determinations, log inhibitor versus response was calculated for a Hill slope of −1. To derive K M and k cat values, data were fitted by nonlinear regression to the Michaelis-Menten equation.
For the analysis of phosphatase activities in cell lysates, LIPIN-1, NT5C and NT5M (myc-DDK tagged) were expressed in HEK-AD293 cells. Cells were cultured as described below, seeded at a density of 0.5 × 10 6 cells/3 cm dish overnight, and transfected with Lpin1, Nt5c, Nt5m or empty vector control (1.5 µg plasmid each) using TransIT-LT1 (Mirus Bio). The next day, cells were rinsed twice with 1 mL 0.9% NaCl at RT, and scraped in 200 µL ice-cold lysis buffer (TNM30 without DTT, supplemented with 0.2% (v/v) Triton X-100, 5 µg/mL aprotinin, 1 µg/mL pepstatin, 5 µg/mL leupeptin and 1 mM Pefabloc). Cells were lysed by passing through a 25 G × 5/8 needle on ice. Unbroken material and cellular debris were removed by centrifugation (21,000 × g, 15 min, 4°C), and the protein concentration of the clarified lysates was determined using the Micro BCA assay (Pierce). Time courses of P i release were performed to find linear assay conditions. Phosphatase activities in the absence or presence of the indicated compounds were determined in 5 µL of the respective supernatants (~5 µg cellular proteins), as described above for purified phosphatases. Lysates of vector control transfected cells were used for background correction. All data are mean values of three independent experiments.

Analysis of CP1 and CP3 combinations on PGP activity
Enzyme inhibition assays were performed as described above, except that murine PGP was incubated in the simultaneous presence of serial CP1 and CP3 dilutions. All wells contained 0.1% DMSO as a solvent control; control wells contained DMSO only. Matrix combination data were processed using the Loewe synergy model and Combenefit software 2.021 (http://sourceforge.net/projects/combenefit/) 44 . Briefly, the software extracts the effects of CP1 and CP3 as single agents from the combination data, which are then fitted with a concentration-response curve. Based on the two single agent concentration-response curves, a combination concentration-response surface is derived. This surface provides a reference for a non-synergistic (additive/independent) combination. The experimental combination concentration-response surface is then compared to the model-generated one, resulting in a synergy distribution in concentration space 44 .

Isothermal titration calorimetry (ITC)
Purified murine PGP-WT or PGP-P127E was dialyzed overnight against ITC buffer (30 mM HEPES, 30 mM NaCl, 5 mM MgCl 2 , 1 mM tris(2-carboxyethyl)phosphine/TCEP; pH 7.5). All calorimetric measurements were performed at 25°C on an ITC200 microcalorimeter (GE Healthcare). Experiments were conducted with a fixed concentration of 2% DMSO. The heat signals released by 15 consecutive 2.4 μL injections of 1 mM CP1 or 0.5 mM CP3 solutions that were titrated into 100 μM or 50 μM PGP, respectively, were measured under continuous mixing at 750 rpm. Both CP1 and CP3 were diluted from a 50 mM stock solution in 100% DMSO, and PGP solutions likewise contained 2% DMSO. To reduce the effects of ligand leakage from the syringe during baseline equilibration, the heat release of a single 1.2 μL injection was recorded prior to the actual titration experiment, and discarded during data analysis with Nitpic 2.0.7 45 . To determine background heats of dilution, control experiments of ligand into buffer, buffer into protein and buffer into buffer titrations were performed.

PGP Crystallization and data collection
For the crystallization of full-length PGP, PGP D34N; C297S (30 mg/mL in TMN50, containing 50 mM triethanolamine, 5 mM MgCl 2 , 250 mM NaCl; pH 7.5) was supplemented with 5 mM unbuffered TCEP. Plateshaped apo-PGP D34N; C297S crystals were grown at 20°C in 0.1 M Tris-HCl pH 8.5, 0.2 M (NH 4 ) 2 SO 4 , 12% (w/v) PEG 8000 using the sitting-drop vapor diffusion method. For co-crystallization, PGP D34N;C297S (25 mg/mL in TMN50) was supplemented with 5 mM unbuffered TCEP and 1 mM CP1. Prism-shaped plates of CP1-bound PGP D34N;C297S were grown at 20°C in 0.49 M NaH 2 PO 4 and 0.91 M K 2 HPO 4 using the sitting-drop vapor diffusion method. Crystals were cryoprotected for flash-cooling in liquid nitrogen by soaking in mother liquor containing 25% (v/v) glycerol for apo-PGP D34N;C297S (apo-PGP) or 25% (v/v) PEG 400 for CP1bound PGP D34N;C297S (CP1-PGP complex). Diffraction data for apo-PGP were collected on the ESRF beamline ID23-1 and for the CP1-PGP complex on the EMBL beamline P13 at the PETRA III synchrotron (DESY, Hamburg). Diffraction data were processed using XDS 46 and further analyzed with Aimless 47 of the CCP4 suite 48 . As Aimless detected significant anisotropy for apo-PGP and the PGP-CP1 complex, data were reprocessed with Staraniso (http://staraniso.globalphasing.org/ cgi-bin/staraniso.cgi). The structure of apo-PGP D34N; C297S was solved by molecular replacement with the program Phaser 49 with the structure of the dimeric murine PDXP/PGP hybrid (PDB entry 4BKM) as search model. The structure of the inhibitor complex was solved with the ABdimer of the apo-PGP structure as search model. Initial real space refinement and restrained refinements were performed in Refmac 50 . Restrained refinement with the addition of TLS refinement, in the case of the apo-PGP structure, was continued in Phenix 51 and Buster (https://www.globalphasing.com/buster/). A polder electron density map 52 was calculated using Phenix.

Molecular dynamics simulations
MD simulations were carried out for apo-PGP and the CP1-PGP complex. Starting structures of PGP with wild-type active site were built from the crystal structures of the full-length PGP D34N; C297S (C-D chains, PDB: 7PO7) and CP1-bound PGP D34N; C297S (A-B chains, PDB: 7POE). The Molecular Operating Environment (MOE) 2022.02 (Chemical Computing Group ULC, 1010 Sherbooke St. West, Suite #910, Montreal, QC, Canada, H3A 2R7, 2022) was used for the initial preparation of the structures. The wild-type active site was modelled based on a structural alignment of PGP with crystal structures of PDXP available in the PDB. As PDXP structure 5AES showed the lowest RMSD (1.06 Å) after alignment with the core residues of CP1-PGP D34N; C297S , it was used to extract the catalytic Mg 2+ ion along with two coordinated water molecules and place it into the active site of PGP. A third water molecule was placed at the position where a phosphate oxygen of the PDXP inhibitor in PDB entry 5AES coordinates to the Mg 2+ ion. Finally, the mutated Asn34 was reverted to the wild-type Asp34, and all residues within 5 Å of the Mg 2+ ion were subjected to energy minimization with the AMBER14:EHT force field in tether mode to an RMS-gradient of 0.001 kcal/(mol·Å).
CP1 was docked to this structure (as described below) to obtain an energetically favourable starting configuration. The docking results obtained in chain A and chain B differed from the crystallographic binding mode in the CP1-PGP D34N; C297S by only 0.48 Å and 0.73 Å, respectively.
Except for the CP1 inhibitor bound to the active site of each monomer, all other non-protein molecules (i.e., glycerol, potassium ions and water, as well as the CP1 molecules localized outside the active site in the crystal structure) were removed from the system to prepare it for the MD simulation. The protein N-and C-termini were capped with ACE (acetyl, i.e. -COCH 3 ) and NME (N-methyl, i.e. -NH-CH 3 ) groups, respectively. The system was protonated at pH 7.5. Histidine residues were defined as HIE tautomers, i.e., with the hydrogen at the epsilon nitrogen. Cys104 and Cys243 were kept in the reduced state (CYS). CP1 was parametrized with AmberTools18 from Amber 2018 53 and the General AMBER Force Field (GAFF) 54 using antechamber and parmchk. The atomic charges were derived from a Restrained Electrostatic Potential (RESP) fit based on electrostatic potentials calculated with Gaussian09 55 at the Hartree-Fock level of theory with the 6-31 G* basis set. PGP parameterization was performed using the AMBER force field ff14SB 56 . After an initial energy minimization of the built structures with a maximum of 2000 cycles through pmemd, the systems were solvated in a TIP3P 57 cuboid water box by means of tleap with a minimum distance of 14.0 Å between solute and periodic box edge. Charge neutrality of the systems was ensured with sodium ions.
All MD simulations were carried out with pmemd of Amber18 under periodic boundary conditions. The systems were thermalized and equilibrated as follows: The solvent was first heated from 100 K to 300 K in the NVT ensemble, using a harmonic potential restraint of 100 kcal/(mol·Å 2 ) on the solute (protein and ligand) during 0.5 ns. The temperature was controlled via Langevin dynamics 58 with a collision frequency of 1 ps −1 . Then, the system was cooled to 100 K with a reduced positional restraint of 10 kcal/(mol·Å 2 ) for 0.25 ns. This restraint was further lowered to 1 kcal/(mol·Å 2 ) during the next 0.5 ns, while the system was reheated to 300 K. The relaxation at 300 K was extended for 0.25 ns maintaining the applied restraint of 1 kcal/(mol·Å 2 ). To reach the density of interest, the system was switched to an NPT ensemble, employing a Berendsen barostat 59 at 1 bar and a relaxation pressure time of 5 ps. Under NPT conditions, the system was further relaxed, conserving the restraint during the first 0.5 ns, followed by 1.0 ns of unrestrained simulation.
After this preparatory phase, a production run of 100 ns was carried out for each system. Electrostatic interactions were handled with the Particle Mesh Ewald method 60 , a cut-off of 10 Å was applied for the van der Waals interactions (8 Å during the preparatory phase). Bonds involving hydrogens were constrained using the SHAKE algorithm, allowing a time step of 2 fs in the simulations. The Langevin dynamics were applied with a collision frequency of 3.0 ps −1 . The relaxation pressure time was switched to 2 ps at 1 bar. Coordinates of the systems were saved every picosecond. The resulting trajectories were imaged and analyzed using cpptraj from AmberTools18 53 . Structural illustrations from the saved coordinates were prepared with PyMOL 2.4.1 61 .
Principal component analysis of the MD simulations was carried out with utilities from Bio3D 42 . The underlying PDB format trajectory for each principal component was obtained from Bio3D and postprocessed with PyMOL 2.4.1 to generate movies from two different views. The individual files were then assembled and labelled with Shotcut (www.shotcut.org, version 22.06.03, Meltytech, LLC). The same colour scheme as in the corresponding PCA figures in Supplementary Fig. 13 was applied.

Docking
To investigate the putative binding mode of the inhibitors CP1-3 and the substrate 2-phosphoglycolate (PG) to the wild-type active-site of PGP, computational docking studies were carried out with Genetic Optimisation for Ligand Docking (GOLD) 2022.1.0 62 . Protein and ligand structures were prepared using MOE 2022.02. The wild-type active-site modelled from the CP1-PGP crystal structure as described above under Molecular Dynamics Simulations was used for docking studies at both active sites (i.e, AS-A of monomer A and AS-B of monomer B), using ChemPLP and ASP as scoring functions within GOLD. The search radius for the binding site was set to 10 Å around CP1 as reference structure; for PG, the search area was centred at the Mg 2+ ion with a radius of 10 Å. Water molecules coordinated to the Mg 2+ ion were considered in three different modes: off, toggle, and on, with a maximal translation distance of 1 Å. Parameters of the genetic algorithm were set as follows: population size 750; number of operations 500,000; crossover 90; mutation 95; and migration 25. For each scoring function and water setting 100 docking runs were carried out, resulting in 200 docking poses per water mode and a total of 600 docking poses per binding site. All poses were rescored with DrugScoreX (DSX) CSD 0.9 63 and structurally clustered with Fconv 1.24 64 using an RMSD cut-off of 2 Å. PyMOL 2.4.1 was used for visual inspection.

NMR spectroscopy, data normalization and statistical analysis
Parental HT1080 cells or HT1080 CRISPR/Cas9_PGP-WT (clone D5) or CRISPR/Cas9_PGP-KO (clone C2) cells were cultured as described above. For NMR experiments, 2.7 million cells were seeded in 10 cm dishes for all samples. The next day, PGP inhibition was performed using either 33 µM CP1 over 3 h, or 100 µM CP1 over 24 h; the same amount of pure DMSO solvent was added in parallel to the respective control cells. To ensure comparable conditions between CP1-treated and CRISPR/Cas9 HT1080 lines, the latter cells were also grown for a total of 48 h and treated with 0.1% DMSO for 24 h. For each condition, six samples were prepared; one of which was used to count the number of cells at the end of the sample preparation. Free metabolites were obtained by methanol extraction: After washing the samples with ice-cold PBS, they were placed in liquid nitrogen for 2 min, followed by mixing the samples cooled on ice with 5 mL methanol. The samples were mixed for 15 min at 4°C, and centrifuged at 2.5 × g for 30 min at 4°C. The solvent was removed by evaporation with a N 2 gas flow over 5 h on ice, followed by freeze-drying under vacuum. The remaining pellet was dissolved in 700 µL D 2 O, and transferred to a 5 mm borosilicate NMR tube. NMR experiments were performed with a BRUKER AVANCE NEO 600 spectrometer operating at 600.13 MHz for 1 H detection equipped with a cryogenic NMR probe offering enhanced sensitivity. The 1 H NMR spectra were acquired using a double watergate pulse sequence with excitation sculpting for water suppression. A relaxation delay of 4 s was sufficient. The π and π/2 pulses performed with a radio frequency power of 62.5 kHz were combined with 2 ms selective rectangular shape pulses optimized for efficient suppression of the H 2 O 1 H resonance, which was chosen as the transmitter frequency. The two blocks of gradient pulses were conducted over 1 ms per pulse using gradients of 0.155 T/m and 0.055 T/m with a smoothed rectangular shape profile (SMSQ10.100). Signal averaging was achieved over 512 scans using an acquisition time of 1.38 s. Acquisition and pre-processing of NMR spectra was performed under the control of a workstation with TopSpin 4.0.7. Data processing included Fourier transformation, zero order phase correction, 1 Hz line broadening using a Lorentzian window function and automatic baseline correction with a polynomial of fifth degree. A 9 mmol/L aqueous sucrose solution was used as a calibration compound to calibrate an ERETIC2 reference for absolute concentration determination 66,67 . Based on the sucrose reference, an electronic signal was placed in each 1 H spectrum to calibrate the deconvolution software Chenomx NMR suite. A standard protocol for quantitative NMR was applied: The signal corresponding to the calibration compound was integrated and normalized according to the number of protons of this compound. The unknown concentration of compound x in the presence of the calibration compound was calculated according to: With I, N, and C being the integral area, number of nuclei, or concentration of the calibration (cal) or the targeted compound (x), respectively. Due to spectral superposition, glycerol-3-phosphate could not be quantified in 1 H NMR spectra. Therefore, we conducted 31 P -1 H HMBC 2D experiments and quantified glycerol-3-phosphate using the phosphorylcholine peak and the respective concentration known from 1 H NMR spectra. Even though signal intensity in such experiments also depends on 31 P-1 H J-couplings, we assume that the error caused hereby is insignificant, especially as we aimed to investigate relative concentration differences caused by cell treatments. 1 H and 31 P pulses were performed at radio frequency powers of 62.5 kHz and 22.2 kHz, respectively. Three gradient pulses were used (sequence hmbcgpndqf) over 1 ms each with gradient strengths of 0.06 T/m, 0.06 T/m and 0.05 T/m. The relaxation delay was set to 2 s, the acquisition time to 0.328 s. 128 experiments were acquired for each 2D experiment using a time increment of 204 µs and averaging over 16 scans. Data processing was achieved as described above except for the Qsine window function in both dimensions. All measurements were performed at 7°C. Raw datasets are provided on Figshare under the accession code [https://doi.org/10.6084/m9.figshare.21316533] 68 .
Data were normalized by dividing the concentration of each metabolite by the cell number and by the absolute metabolite content of the respective sample; this value was multiplied with the group average of the absolute metabolite content. This approach removes effects caused by cell number differences, while maintaining those generated by the trial conditions. Multivariant data analysis was performed using custom-made R scripts employing the mixOmics package 69 . For sPLS-DA, the data were additionally centered. Classic univariate receiver operating characteristic (ROC) curve analysis used to calculated AUC values and pathway enrichment analysis was performed using MetaboAnalyst 70 using the web server MetaboAnalyst 5.0 (https://www.metaboanalyst.ca/MetaboAnalyst/ModuleView.xhtml). The pathway analysis procedure employed the options Global Test and relative betweenness centrality in combination with the KEGG Homo sapiens database.

Cell viability assays
HT1080 cells were seeded onto 96-well plates (5000 cells per well), and treated for 24 h with the indicated CP1 concentrations or with the appropriate DMSO solvent control. Cell viability was assessed using AquaBluer according to the manufacturer's recommendations (Multi-Target Pharmaceuticals, LLC). Fluorescence (excitation: 540 nm; emission: 590 nm) was analyzed on a Clariostar microplater reader.

Cell proliferation assays
Parental HT1080 cells or the indicated CRISPR/Cas9-engineered control (PGP-WT) or PGP-KO cells were grown in complete DMEM. For the expression of PGP-WT or PGP-P127E, cells were transfected with either the pcDNA3 empty vector control or the indicated constructs in pcDNA3 using X-tremeGENE (Roche/Merck), according to the manufacturer's instructions. The next day, cells were seeded at a density of 2000 cells per well of a 96-well microtiter plate, and incubated with 100 µM CP1 or 0.1% DMSO for 16 h. To assess DNA synthesis, cells were labeled for 2.5 h with 10 μM 5-ethynyl-2'-deoxyuridine (EdU), fixed and permeabilized according to the manufacturer's recommendations (Click-iT EdU Alexa Fluor 488 imaging kit, Invitrogen/Thermo Fisher). Cells were counterstained with 4′,6-diamidine-2-phenylindole (DAPI), embedded with Fluoromount-G (Thermo Fisher), and imaged on a