Thermal proteome profiling reveals fructose-1,6-bisphosphate as a phosphate donor to activate phosphoglycerate mutase 1

ABSTRACT Deep understanding of sugar metabolite-protein interactions should provide implications on sugar metabolic reprogramming in human physiopathology. Although tremendous efforts have

been made for determining individual event, global profiling of such interactome remains challenging. Here we describe thermal proteome profiling of glycolytic metabolite

fructose-1,6-bisphosphate (FBP)-interacting proteins. Our results reveal a chemical signaling role of FBP which acts as a phosphate donor to activate phosphoglycerate mutase 1 (PGAM1) and

contribute an intrapathway feedback for glycolysis and cell proliferation. At molecular level, FBP donates either C1_-O_-phosphate or C6_-O_-phosphate to the catalytic histidine of PGAM1 to

form 3-phosphate histidine (3-pHis) modification. Importantly, structure-activity relationship studies facilitate the discovery of PGAM1 orthostatic inhibitors which can potentially restrain

cancer cell proliferation. Collectively we have profiled a spectrum of FBP interactome, and discovered a unique covalent signaling function of FBP that supports Warburg effect via histidine

phosphorylation which inspires the development of pharmacological tools targeting sugar metabolism. SIMILAR CONTENT BEING VIEWED BY OTHERS MULTI-OMICS REVEALS NEW LINKS BETWEEN

FRUCTOSAMINE-3-KINASE (FN3K) AND CORE METABOLIC PATHWAYS Article Open access 03 June 2024 THE MOLECULAR BASIS OF HUMAN FN3K MEDIATED PHOSPHORYLATION OF GLYCATED SUBSTRATES Article Open

access 22 January 2025 CROSSTALK BETWEEN O-GLCNACYLATION AND PHOSPHORYLATION IN METABOLISM: REGULATION AND MECHANISM Article 05 March 2025 INTRODUCTION Sugar metabolic reprogramming has been

regarded as a hallmark of various diseases1,2. The altered sugar metabolism, for instance, accelerated aerobic glycolysis in cancer cells3, is adapted to support biomolecule assembly,

energy production and cellular redox homeostasis4. Emerging evidences have shown that disregulated sugar metabolites from such reprogramming process act as signaling molecules to regulate

various cellular events through interacting with important proteins5,6,7,8,9,10,11,12,13. Dissecting the mechanisms of sugar metabolite signaling is therefore crucial for understanding the

impact of sugar metabolic rewiring on human physiology and diseases. As an example, the central glycolytic metabolite fructose-1,6-bisphosphate (FBP) has been revealed as an endogenous

regulator of physiopathological processes including AMP-activated protein kinase (AMPK) activation in response to glucose deprivation14, epidermal growth receptor (EGFR) binding and

activation in triple-negative breast cancer15, activation of oncogenic protein Ras through direct binding to son of sevenless homolog 1 (SOS1)16, inhibition of mitochondrial oxidative

phosphorylation17, as well as binding to high mobility group box 1 (HMGB1) to impair cancer cell viability18. Despite tremendous efforts have been made in identifying individual FBP binding

proteins, a full spectrum of its protein targets and the underlying molecular mechanisms of FBP-target interactions remain largely unexplored. Recently, derivatization-free quantitative

chemoproteomic strategies have become powerful tools to capture the interacting proteins of small molecules in their native forms and enable comprehensive profiling of metabolite-protein

interactome. Limited proteolysis-mass spectrometry (LiP-MS)19,20, the mass spectrometry integrated with equilibrium dialysis for the discovery of allostery systematically (MIDAS) platform21,

as well as target-responsive accessibility profiling (TRAP)22 have been developed for mapping metabolite-protein interactions in intact cells. As a complementary approach, MS-based thermal

proteome profiling (TPP) found broad applications in comprehensive and unbiased profiling of small molecule targets23,24 as well as protein post-translational modifications (PTMs)25,26,27.

It enables the proteome-wide assessment of drug–protein interactions or PTMs by combining quantitative mass spectrometry with cellular thermal shift assay (CETSA)28. In this work, we utilize

a TPP approach to profile the interactome of FBP in HepG2 cells and identify 66 unreported proteins with various functions and subcellular locations as creditable FBP targets. Through

functional validations of metabolic enzymes that involve in the catalytic process of phosphate transfer, we find that FBP is able to donate either its C1-_O_-phosphate or its

C6_-O_-phosphate to the catalytic histidine (His11) of glycolytic enzyme PGAM1 to form 3-pHis modification and activate its enzymatic activity. Such specific metabolite-metabolic enzyme

interaction act as a positive intrapathway feedback to support the Warburg effect and cancer cell proliferation. Molecular docking and molecular dynamics simulations, as well as structural

modifications of FBP allow for structure-activity relationship (SAR) studies of FBP-PGAM1 interaction and give rise to PGAM1 orthostatic inhibitors to reshape glucose metabolism and restrain

cancer cell proliferation, showing promises in strategies for drug discovery in cancer treatment by targeting sugar metabolism. RESULTS THERMAL PROTEOME PROFILING OF FBP-INTERACTING

PROTEINS IN HEPG2 CELLS In TPP strategy, targets can be identified through comparing proteins’ stability within proteome over a temperature range23. The protein abundance at each temperature

was quantified to draw a thermal shift curve, from which the corresponding melting temperatures (Tm) could be calculated. By comparing the difference of Tm before and after small molecule

treatment, one can report the thermal stability change (ΔTm) in an intuitive way. As a proof of concept, we first tested if FBP is able to modulate the thermal stability of whole proteome

and known FBP-interacting proteins in cell lysates heated from 37 °C to 62 °C with a 5 °C gradient (Supplementary Fig. 1). CESTA demonstrated that FBP reduced the thermal stability of its

known target pyruvate kinase isoform M2 (PKM2)29,30 with ΔTm of −2.7 °C, and greatly decreased the thermal stability of SOS1 with ΔTm of −6.2 °C. Unexpectedly, FBP greatly increased the

thermal stability of glyceraldehyde-3-phosphate dehydrogenase (GAPDH), which is not a known FBP target, with ΔTm of 5.9 °C (Supplementary Fig. 1b). These results indicated that the TPP

strategy is suitable for capturing FBP-interacting proteins that allows for global mapping of FBP interactome in the whole cell lysates. We then accessed the FBP-interactome by TPP coupled

with stable isotope dimethyl labeling for quantitative mass spectrometry in a human liver cancer cell line HepG2 (Fig. 1). HepG2 cell lysates were treated with FBP or H2O and subsequently

divided into two groups to be heated to either 37 °C or 57 °C, respectively (Fig. 1a). The “heavy” groups represent cell lysates heated to 37 °C, while the “light” groups represent cell

lysates heated to 57 °C. After evaluation and quantification by liquid chromatography-tandem mass spectrometry (LC-MS/MS), we got a 57/37 (light/heavy) ratio for each protein in either FBP

or H2O treated group respectively, and then we compared the light/heavy ratio of each protein in either group to yield a new ratio (FBP/H2O) for analysis. 2202 overlapping proteins were

identified out of three biological replicates (Fig. 1b and Supplementary Dataset 1), and we assigned the proteins with a mean ratio (FBP/H2O) over 1.5 as credible FBP targets with increased

thermal stability, and proteins with a mean ratio under 0.66 as credible FBP targets with reduced thermal stability (Fig. 1c). With high confidence, we identified 6 sugar metabolic enzymes

with increased thermal stabiliies by FBP treatment, including GAPDH, phosphomannomutase 2 (PMM2), PGAM1, phosphoglucomutase 1 (PGM1), ribose-phosphate pyrophosphokinase 1 (PRPS1) and a

protease dipeptidyl peptidase 1 (CTSC). On the other hand, 60 proteins were identified FBP targets with reduced thermal stability, such as GTP-binding nuclear protein Ran (RAN), glycine-tRNA

ligase (GARS), histone deacetylase 2 (HDAC2), serine/threonine-protein kinase 24 (STK24), and aminopeptidase B (RNPEP). For the 66 highly confident FBP targets (Supplementary Dataset 2), we

performed bioinformatic analysis to identify their molecular functions, subcellular distributions, as well as the biological pathways. Gene ontology (GO) enrichment analysis indicated that

these targets are functionally involved in transport (24.1%), metabolic process (21%), ribonucleoprotein (15%), transcription and translation (12%), chaperon (4.5%), etc. (Fig. 1d). They are

located in various cellular components including the cytoplasm (45.5%), nucleus (13.6%), cytoplasm & nucleus (25.8%) and so on. Kyoto Encyclopedia of Genes and Genomes (KEGG) functional

pathway analysis showed that they participate in various biochemical pathways such as translation, glycolytic process, ribosomal small subunit biogenesis and export from the nucleus (Fig. 

1e). Chromatographic peaks of representative peptides from FBP targets with 57/37 (light/heavy) ratio in the FBP treated group and control group (H2O) are shown in Fig. 1f and they provide

information of stabilization or destabilization effects of FBP on these proteins by isotope quantification. BIOCHEMICAL VALIDATION OF IDENTIFIED FBP TARGETS We next validated the

representative FBP targets by CETSA in HepG2 cell lysates (Fig. 2a). For the proteins with increased thermal stability by FBP treatment, the glycolytic enzyme PGAM1 demonstrated a ΔTm of 6.8

 °C, while the other sugar metabolic enzymes PGM1, PMM2 and PRPS1 exhibited ΔTm of 3.0 °C, 7.1 °C and 10.8 °C, respectively (Fig. 2b). We also validated proteins with reduced thermal

stability by FBP treatment including GARS (ΔTm = −8.9 °C), HDAC2 (ΔTm = −3.7 °C), DNA-directed RNA polymerase II subunit RPB1 (POLR2A, ΔTm = −5.5 °C), Pyridoxal-dependent decarboxylase

domain-containing protein 1 (PDXDC1, ΔTm = −3.6 °C), polyribonucleotide nucleotidyltransferase 1 (PNPT1, ΔTm = −2.7 °C), E3 ubiquitin/ISG15 ligase (TRIM25, ΔTm = −1.6 °C), RAN (ΔTm = −2.9 

°C) and serine/threonine-protein kinase OSR1 (OXSR1, ΔTm = −1.0 °C) (Fig. 2c and Supplementary Fig. 2a). These results revealed that FBP indeed induced stability changes of its interacting

proteins and suggested that our TPP strategy is an effective tool to profile the interactome of FBP. Our study delineated a global portrait of FBP-interacting proteins and provide a

comprehensive resource for the following functional investigations. To further confirm the direct regulation of protein thermal stabilities by FBP treatment at the molecular level, we

incubated the recombinant proteins with FBP or H2O. Consistent with the CESTA results in cell lysates, FBP increased the thermal stabilities of recombinant human GAPDH, PMM2, PRPS1, PGM1 and

PMM2 proteins, while it reduced the thermal stabilities of recombinant human RAN, HDAC2 and OXSR1 proteins (Fig. 2d). For the five FBP targets with increased thermal stability, we observed

concentration-dependent stabilizing effects of FBP on the recombinant proteins when they were heated from 37 °C to 62 °C (Supplementary Fig. 2b). And the proteins turned to be stabilized by

FBP immediately upon incubation. FUNCTIONAL REGULATION OF FBP TOWARDS SUGAR METABOLIC ENZYMES PGAM1, PGM1 AND PMM2 We then focused on the functional consequences of interactions between FBP

and its identified targets. Through analyzing the 66 credible FBP targets, we found that 25 of them (37.9%) are proteins involved in phosphate transfer or recognition, while 26 of them

(39.4%) are ATP/GTP or DNA/RNA binding proteins (Supplementary Fig. 3a). Further we found that 33% of FBP targets with reduced thermal stability and 83% of the FBP targets with increased

thermal stability are enzymes involved in the catalytic processes of phosphate transfer (Supplementary Fig. 3b, c). Interestingly, PGAM1, PGM1 and PMM2 share a common “ping-pong” mechanism

through phosphate transfer to catalyze the interconversion of sugar phosphates (Supplementary Fig. 3d)31,32,33. Such catalytic processes raised our strong interests to investigate how FBP,

as an endogenous metabolite, interacts with these phosphate-transfer related enzymes at the molecular level and how it regulates their functions. PGAM1 catalyzes the interconversion of the

lower glycolytic metabolite 3- phosphoglycerate (3-PG) and its downstream isomer 2-phosphoglycerate (2-PG), by using 2,3-bisphosphoglycerate (2,3-BPG) as both an intermediate and a cofactor

that phosphorylates the catalytic histidine 11 to activate the enzyme31. Besides 2,3-BPG, either lower glycolytic metabolite phosphoenolpyruvate (PEP)34 or 1,3-bisphosphoglycerate

(1,3-BPG)35 has been reported to act as a phosphate donor to activate PGAM1 through His11 phosphorylation using a similar mechanism. PGM1 catalyzes the interconversion of

d-glucose-6-phosphate (G6P) into α-d-glucose-1-phosphate (G1P) through α-d-glucose-1,6-bisphosphate (GBP) as an intermediate32 and the catalytic step branches glycolysis into glycogen

synthesis. PMM2 catalyzes the interconversion of d-mannose-6-phosphate (M6P) into α-d-mannose-6-phosphate (M1P) and requires a sugar bisphosphate, either α-d-mannose-1,6-bisphosphate (MBP)

or α-GBP for its activity33. PMM2 is very important in the biosynthesis of the GDP-mannose and dolichol-phosphate mannose and associated with the most common congenital disorder of

glycosylation (CDG)36. The three FBP targets share a common catalytic mechanism for the isomerization of sugar phosphates and we assumed that FBP as a sugar bisphosphate might have specific

interaction and regulation towards such kind of enzymes, to perform both intra- and inter-pathway regulations of glucose metabolism. From enzymatic activity assays and microscale

thermophoresis (MST) analysis we found that FBP is able to activate PGAM1 with an EC50 value of 0.48 mM and Kd value of 166 μM, while it inhibits PGM1 with an IC50 value of 2.79 mM and Kd

value of 235 μM, as well as PMM2 with an IC50 value of 7.91 mM and Kd value of 247 μM (Fig. 3a, b). Such weak interactions are consistent with previous observations on the interactions

between endogenous metabolites with their target proteins21, indicating FBP might regulate the activities of its targets and the downstream signaling events only when it is accumulated under

centain stressed or pathological conditons. We next compared FBP with the well known PGAM1 activator 2,3-BPG in regards of PGAM1 interaction and activation, and found that FBP is around

1000 times less reactive than 2,3-BPG. FBP has been reported to be highly accumulated in hepatocellular carcinoma and exhibit a cytosolic concentration up to 7–25 mM17,37. While in the

cultured mammalian cells, the absolute cellular concentration of FBP has been determined as 1.52 mM38. In order to investigate the biological or pathological relevance of such relatively

weak binding affinities between FBP and its targets, we performed LC-MS/MS based targeted metabolomics using 13C6-FBP as internal standard (IS) for quantification to determine the

intracellular concentrations of FBP in HepG2 cells (Supplementary Fig. 4a, b). The FBP concentrations in HepG2 cells were determined as 0.416 ± 0.056 mM (Supplementary Fig. 4c) with normal

glucose supplement, suggesting the binding affinities of FBP with PGAM1, PGM1 and PMM2, as well as the activation of PGAM1 by FBP are biologically relevant, while the inhibition of PGM1 and

PMM2 might be pathologically relevant. FBP ACTIVATES PGAM1 THROUGH PHOSPHORYLATION OF ITS CATALYTIC SITE HIS11 As a glycolytic metabolite, FBP is known to be cleaved into two three-carbon

metabolites glyceraldehyde-3-phosphate (G3P) and dihydroxyacetone phosphate (DHAP) by glycolytic enzyme ALDOA39. Meanwhile it is a well-studied allosteric activator of lower glycolytic

enzyme PKM229. Yet it’s unreported whether FBP could interfere or participate in the phosphate transfer processes of metabolic enzymes. We assume FBP functions as an activator of PGAM1

through phosphorylation of its catalytic histidine. Firstly, we validated this hypothesis using histidine phosphorylation antibodies and Phos-tag reagent. Recombinant human PGAM1 (PGAM1) was

incubated with FBP and subsequently subjected to immunoblotting or Phos-tag SDS-PAGE analysis (Fig. 3c). The N3-phosphohistidine (3-pHis) or 1-phosphohistidine (1-pHis) monoclonal

antibodies, developed by Prof. Tony Hunter and his colleagues40, were utilized to confirm that FBP is able to phosphorylate PGAM1 to form 3-pHis modification instead of 1-pHis species. It

was consistent with the Phos-tag SDS-PAGE41 results that the phosphorylated PGAM1 by FBP treatment exhibited a mass shift. Through comparison with the known 1-pHis modified protein NME2 by

ATP, it was then confirmed by immunoblotting analysis that there’s no 1-phosphohistidine formation on PGAM1 by FBP treatment (Supplementary Fig. 5). We observed a clear concentration

dependent phosphorylation of PGAM1 by FBP, in a similar way as known PGAM1 activator 2,3-BPG (Fig. 3d). 2,3-BPG is able to phosphorylate PGAM1 at a much lower concentration of 0.01 mM than

that of FBP (1 mM), which is consistent with the EC50 and Kd value of these two molecules towards PGAM1 (Fig. 3a, b). The weak phosphorylation of PGAM1 by PEP was consistent with the report

that PEP might need an extra enzyme or cofactor to phosphorylate and activate PGAM134. FBP was found to immediately phosphorylate the histidine of PGAM1 upon incubation and the 3-pHis

modification decayed after 4 h (Fig. 3e). These results suggested that FBP phosphorylates and activates PGAM1 directly with no cofactor and no time required. In order to identify the PGAM1

phosphorylation sites induced by FBP, we incubated PGAM1 with FBP, then digested the protein with trypsin and subjected it to LC-MS/MS analysis. It was shown that FBP could directly

phosphorylate PGAM1 at its catalytic site His11 (Fig. 3f and Supplementary Fig. 6a). We then generated a mutant of PGAM1-H11A through site-directed mutagenesis and confirmed by

immunoblotting analysis that the catalytic His11 was the only histidine phosphorylated by FBP to generate 3-pHis modification (Fig. 3g). Meanwhile, FBP lost its activation effect towards

PGAM1-H11A mutant, suggesting the phosphorylation of catalytic His11 is the key for PGAM1 activation (Fig. 3a). His11 was found to be correlated with the thermal stability of PGAM1, and H11A

mutant was slightly less stable than PGAM1 WT (Supplementary Fig. 6b–d). However, loss of His11 did not influence the stabilizing effect of FBP towards PGAM1 (Supplementary Fig. 6c, d),

indicating FBP might increase the thermal stability of PGAM1 through other mechanism but not pHis11 modification. To clarify the fate of FBP after its phosphate donation to PGAM1, we

performed LC-MS/MS analysis of fructose monophosphate products using a derivatization reagent 2-(diazo-methyl)-_N_-methyl-_N_-phenyl-benzamide (2-DMBA, Supplementary Fig. 4d) for

distinguishing F6P and F1P by retention time and fragment ions (Supplementary Fig. 4e)42. Upon treatment of PGAM1 with FBP in vitro, both F6P and F1P were generated immediately and

accumulated following reaction time (Fig. 4a), indicating that both C1-_O_-phosphate and C6-_O_-phosphate of FBP could be transferred to PGAM1 (Fig. 4b). MOLECULAR DOCKING AND MOLECULAR

DYNAMICS SIMULATIONS OF FBP-PGAM1 INTERACTION To access the molecular details of the interaction between FBP and PGAM1, we performed molecular docking to examine how FBP donates its

phosphate to His11 of PGAM1, and to confirm that the two phosphates attached to the sugar backbone behave similarly. Of special note, when FBP exists as an α-anomeric cyclic form or a linear

form, the calculated distances out of thirty confirmations between the atom P1 and atom N3 at PAGM1-His11 were all much shorter than those between P6 and N3 (Supplementary Dataset 3). In

sharp contrast, when FBP exists as a β-anomeric cyclic form that is favored (80%) in solution43, the calculated distances between P1 and N3 at PAGM1-His11 showed no significant difference

with those between P6 and N3, indicating that FBP donates either its C1-_O_-phosphate or its C6-_O_-phosphate to the N3 position of PGAM1 catalytic His11 (Supplementary Fig. 7a). It’s

consistent with the formation of both F6P and F1P products upon LC-MS/MS analysis of FBP-PGAM1 interaction (Fig. 4a, b). We also observed hydrogen bonds between C1-_O_-phosphate, C2-OH,

C3-OH, C6-_O_-phosphate of β-FBP and a few crucial amino acid residues within the catalytic pockets of PGAM1 (Fig. 4c). In detail, when donating C1-_O_-phosphate, the phosphate group at C1

position forms hydrogen bonds with the known substrate binding sites Arg10, Gly187 and Arg62, the transition state stabilizer His18644,45. And the anomeric C2-OH of FBP forms hydrogen bonds

with proton donating active site Glu89 and substrate binding site Asn188, while the C3-OH of FBP forms hydrogen bonds with substrate binding site Tyr92. On the other hand, when donating

C6-_O_-phosphate, the phosphate group at C6 position forms hydrogen bonds with the same three amino acids Arg10, Arg62 and His186, while the C2-OH and C3-OH form hydrogen bonds with Asn17

and C1-_O_-phosphate forms hydrogen bonds with Arg10. Overall the overmentioned hydrogen bonding network enabled the position of either C1-_O_-phosphate or C6-_O_-phosphate of FBP placed

exactly opposite to the N3 of His11, thereby facilitating the phosphate transfer from FBP to PGAM1. As a comparision, molecular docking of 2,3-BPG with PGAM1 showed that the C2-_O_-phosphate

is orientated to the His11 of PGAM1 through hydrogen bonding network between C2-_O_-phosphate and Arg10, Gly187, Arg62, His186, those between C1 carboxylic acid and Arg62, Gly24, as well as

those between C3-_O_-phospahte and Arg10, Ser14 (Supplementary Fig. 7c). To validate the molecular docking simulation of the binding mode between FBP and PGAM1, we generated six PGAM1

single site mutants including PGAM1-R10A, R62A, E89A, Y92A, H186A, N188A through site-directed mutagenesis of the key residues participating in the FBP binding with PGAM1, as well as

PGAM1-3A with three sites mutated (R10A, E89A and H186A). Through immunoblotting and MST analysis of the mutated proteins, we concluded that mutation of the substrate binding sites Arg10, or

the proton donating active site Glu89 or the transition state stabilizer His186 drastically compromised both the binding and histidine phosphorylation of PGAM1 by FBP (Fig. 4d and

Supplementary Fig. 6e). To further confirm the FBP-PGAM1 interaction with molecular details, molecular dynamics (MD) simulations were performed to investigate the dynamic process of the

binding and phosphorylation of PGAM1 by FBP (Fig. 4e and Supplementary Fig. 8). Steady state of PGAM1-FBP complex during the process of phosphate transfer from either C1-_O_-phosphate (Fig. 

4e up) or C6-_O_-phosphate (Fig. 4e down) of FBP were captured after 750 ns simulation. We observed the hydrogen binding of FBP with key residues of PGAM1 including Arg10, Asn17, Arg62,

His186, Gly187, Asn188 that align well with the docking model, as well as the position of FBP phosphate group towards His11 of PGAM1. Based on the non-bond monitor of MD simulation of

PGAM1–FBP complex (Supplementary Fig. 8c, d), we calculated the probability of hydrogen bonding interactions between FBP and the PGAM1 active site residues. Within the interaction network,

we found that Arg10 exhibited the highest probability during the process of either C1-_O_-phosphate transfer (Fig. 4e up) or C6-_O_-phosphate transfer (Fig. 4e down). Taken together, these

data showed that the active site of PGAM1 is suitable to locate a six-carbon sugar phosphate, although its natural substrates 3-PG, 2-PG and cofactor 2,3-BPG are all three-carbon sugar

phosphates. Three-carbon units on FBP that holds C1-_O_-phosphate, 2-OH and 3-OH, makes the molecule easy to be installed into PGAM1 catalytic pocket through binding of key residues Arg10,

Glu89 and His186, which facilitates His11 phosphorylation of PGAM1 by FBP. STRUCTURE-ACTIVITY RELATIONSHIP (SAR) OF FBP-PGAM1 INTERACTION As mentioned above, we found that FBP can activate

PGAM1 by covalent signaling mediated by phosphate transfer, but little is known about its specificity for such process. There are different kinds of metabolites with sugar phosphate

structure in cells, it’s important to know whether they could activate PGAM1 in a similar way. We next collected glucose-1,6-bisphosphate (GBP), 2,3-BPG, F1P, F6P, G1P, G6P, M1P and M6P and

test whether they were able to phosphorylate PGAM1 (Fig. 5a). It turned out that only FBP or 2,3-BPG act as a phosphate donor to form 3-pHis modification on PGAM1 by immunoblotting and

Phos-tag SDS-PAGE analysis (Fig. 5b). Interesting, GBP can phosphorylate PGAM1 even though it could not form 3-pHis. From the PGAM1 enzymatic assay, bisphosphates containing metabolites FBP

and 2,3-BPG could activate the enzyme while GBP inhibited its activity (Fig. 5c). Sugar monophosphates F6P, F1P, G6P, G1P, M6P and M1P did not regulate the enzymatic activity of PGAM1. In

the meantime, the bisphosphate-containing structure has been found to be essential for PGAM1 activation. F1P holds the same three-carbon units containing C1-_O_-phosphate, C2-OH and C3-OH,

but it could not position its C1-_O_-phosphate to the opposite position of His11 from molecular docking (Supplementary Fig. 7d left). Likewise, F6P could not position its C6-_O_-phosphate to

PGAM1 His11 (Supplementary Fig. 7d right). This explains why neither F1P nor F6P could activate PGAM1 through His11 phosphorylation. GBP bearing two phosphates is able to inhibit the PGAM1

activity through occupying the active site but not donating its phosphate to His11 (Supplementary Fig. 7b). Compared to FBP, neither of the two phosphates in GBP, either in the α-cyclic form

or the β-cyclic form, could be positioned opposite to the catalytic His11 on PGAM1. Taken together, we could draw the conclusion that sugar bisphosphates (FBP, GBP and 2,3-BPG) show

influences on the phosphorylation and activation of PGAM1, but sugar monophosphates do not have such effect. To gain deeper molecular insights, we next designed and synthesized a series of

FBP analogues for SAR studies (Figs. 5d and 6). As the C1-_O_-phosphate, C6-_O_-phosphate, C2-OH and C3-OH are involved in the hydrogen-bonding network within the PGAM1 active site (Fig. 

4c), we generated α- and β-methoxide on the anomeric C2 of the furanose ring to give α-MeFBP and β-MeFBP, to block the hydrogen-bonding and fix the sugar phosphate into its cyclic form with

absolute anomeric configuration (Fig. 6a). We modified the C1-_O_-phosphate of FBP with dimethyl phosphates to form 1-DMeFBP, with the purpose of inhibiting the phosphate donation process

(Fig. 6e). Likewise, we synthesized 6-DMeFBP that masks the C6-_O_-phosphate and 1,6-DMeFBP that masks both phosphates with methyl groups (Fig. 6c, d). 4-DeoxyFBP was also synthesized to act

as a mimic of FBP to retain the bisphosphates for PGAM1 phosphorylation (Fig. 6b). From Phos-tag SDS-PAGE and immunoblotting analysis, all of the six FBP analogues were found to be unable

to phosphorylate PGAM1 (Fig. 5e). From the PGAM1 enzymatic assay, α-MeFBP, β-MeFBP, 4-deoxyFBP, 1-DMeFBP, 6-DMeFBP and 1,6-DMeFBP all demonstrated inhibitory properties against PGAM1 (Fig. 

5f). Among them, we measured the IC50 of the methyl phosphates 1-DMeFBP, 6-DMeFBP, 1,6-DMeFBP and gave 1.08 mM, 1.23 mM and 1.52 mM respectively (Fig. 5g). Molecular docking of PGAM1 with

six analogues showed that they are able to enter into the catalytic pocket of the enzyme but the C1-_O_-phosphate of such analogues cannot be positioned towards His11 of PGAM1 (Supplementary

Fig. 9a). As an example, we looked into the details of the interaction between 6-DMeFBP and PGAM1, and found that even the C1-_O_-phosphate, 2-OH and 3-OH of 6-DMeFBP form similar hydrogen

bonding network with the PGAM1 active site, the phosphate cannot be positioned at the opposite position of His11 (Supplementary Fig. 9b, c). These results suggested that the entire part of

FBP including bisphosphates and fructose backbone are necessary for PGAM1 phosphorylation. FBP ACTIVATES PGAM1 BY HIS PHOSPHORYLATION IN LIVING CELLS TO FEEDFORWARD GLYCOLYSIS AND SUPPORT

CANCER CELL PROLIFERATION We next investigated the downstream consequences of PGAM1 activation by FBP in living cells (Fig. 7). Dose-dependent histidine phosphorylation of PGAM1 by FBP

treatment was confirmed by immunoblotting analysis in living cells with overexpressed Flag-tagged PGAM1 (Fig. 7a). Through LC-MS/MS quantification we observed upregulation of cellular FBP

level after exogenous treatment of HepG2 cells with 10 mM FBP, ranging from 0.416 ± 0.056 mM to 1.435 ± 0.062 mM, indicating the successful uptake of FBP (Fig. 7b left). Accumulation of FBP

at such level may not reach the disease conditions that were reported as 7–25 mM.17,37 PGAM1 catalyzed the intervention of 3-PG to 2-PG and branched the glycolytic flux into the serine

synthesis pathway (SSP). As a downstream consequence of PGAM1 activation, the 2-PG/3-PG ratio was significant increased by 10 mM exogenoues FBP treatment (Fig. 7b right and Supplementary

Fig. 4f, g). Similar as FBP activation of PKM2, we regard PGAM1 activation by FBP phosphorylation as an intrapathway feedforward mechanism to support glycolysis in cancer cells. We observed

significant increases of both lactate production and glucose consumption by FBP treatment (Fig. 7c), suggesting upregulation of glycolysis by FBP treatment in living cells. PGAM1 has been

regarded as a therapeutic target in hepatocellular carcinoma46 and reported to play a significant role in coordinating glycolysis and biosynthesis to promote tumor growth47 and cancer cell

migration48. PGAM1 activation by FBP resulted in significantly upregulated cell proliferation and the effect of FBP was abolished in HepG2 cells with PGAM1-knockout (KO) by a specific sgRNA

(Fig. 7d and Supplementary Fig. 10), suggesting a positive intrapathway feedback role of central glycolytic metabolite FBP in promoting cancer cell proliferation by activating its downstream

metabolic enzyme PGAM1. The supportive role of FBP on accelerating glycolysis and cancer cell proliferation was confirmed by the opposite effects of its synthetic analogue 1-DMeFBP that

inhibits PGAM1 activity. Synthetic PGAM1 inhibitors 1-DMeFBP, 6-DMeFBP and 1,6-DMeFBP were all found to be able to inhibit cell preliferation (Fig. 7e). Among the three FBP analogues,

1-DMeFBP showed strongest inhibition against HepG2 cells proliferation, therefore we continued detailed studies using this analogue. We observed dose-dependant decrease of histidine

phosphorylation level of Flag-tagged PGAM1 in living HepG2 cells by 1-DMeFBP treatment (Fig. 7f). 1-DMeFBP significantly downregulated lactate production and glucose consumption, indicating

its inhibitory role on glycolysis by targeting PGAM1 in HepG2 cells (Fig. 7g). Consistently, cell proliferation was significantly inhibited by 1-DMeFBP treatment and and it was abolished in

PGAM1-KO cells (Fig. 7h). These studies suggested that FBP activates PGAM1 in living cells by histidine phosphorylation to support glycolysis and cancer cell proliferation, while its

analogues exhibit a contrary effect as a type of orthostatic inhibitor of PGAM1 (Fig. 7i). Our study hence provides a unique perspective on developing pharmacological molecules to regulate

and study PGAM1 based on its specific interaction with endogenous sugar metabolite FBP. DISCUSSION Highly abundant intracellular metabolites actively engage with various metabolic enzymes

from both intra- and interpathway in the forms of allosteric or orthosteric modulation, which fabricates a complex and dynamic network to regulate the cell behaviors and fates in response to

nutrient availability and environmental cues19,21,49. In terms of the interactions between sugar metabolites and enzymes, emerging evidences show that sugar metabolites not only regulate

specific upstream or downstream metabolic enzymes for intrapathway feedforward and feedback regulations, but also crosstalk with inter-pathway metabolic enzymes to maintain cellular redox

homeostasis47,50. Comprehensive identification of the interaction network between sugar metabolites and their target proteins especially those intra- and interpathway metabolic enzymes will

facilitate deeper understandings of the underlying regulatory mechanisms of sugar metabolism reprogramming and provide directions for disease diagnosis and treatment through targeting

aberrant sugar metabolism. Derivatization-free chemical proteomic strategies enable the profiling of the native forms of sugar metabolite interactomes which serve as powerful tools to

interpret the precise mechanism of actions of sugar metabolite signaling in living systems19,20,21,22. From our unbiased global profiling of the interacting proteins of the central

glycolytic metabolite FBP based on a TPP strategy in HepG2 cells, a series of unreported FBP-interacting proteins have been discovered containing a considerable population of enzymes

involved in the phosphate transfer process. Following with such a unique feature of sugar metabolite with bisphosphate, we validated the functional regulation role of FBP towards sugar

metabolic enzymes PGAM1, PGM1 and PMM2 that share a communal “ping-pong” catalytic mechanism for the interconversion of sugar phosphate substrates. PGAM1 is overexpressed in many kinds of

cancers, PGM1 and PMM2 are two important biomarkers closely related to glycosylation disorder diseases. Studies on endogenous regulations of these enzymes should facilitate better

understanding of their roles in human diseases and the development of intervention strategy. Here we disclosed the unreported activating property of FBP towards its intrapathway downstream

metabolic enzyme PGAM1, as well as its inhibitory activity against inter-pathway sugar metabolic enzymes PGM1 and PMM2. These findings may provide fundamental references for further

exploration of their psychopathological relevance. Comprehensive studies have been conducted to determine the molecular details of PGAM1 activation by FBP. As an interesting discovery, we

found that FBP acts as a phosphorylation donor to activate the catalytic His11 by generating 3-pHis modification on PGAM1. And it was confirmed by LC-MS/MS, immunoblotting and Phos-tag

SDS-PAGE analysis. Such 3-pHis modification was generated immediately upon FBP-PGAM1 interaction, with no cofactor or enzyme required, and we defined the phosphorylation process as an

autocatalytic reaction. Molecular docking and MD simulations demonstrated that the FBP interacts with PGAM1 in a very specific manner mediated by the hydrogen bonding network between its

C1_-O_-phosphate, C6_-O_-phosphate, C2-OH, C3-OH and the binding sites within the catalytic pocket of PGAM1. FBP is very likely to be fitted into the pocket and donate either its

C1-_O_-phosphate or its C6-_O_-phosphate to the N3 of catalytic His11 and then activates the enzyme immediately. It shows that the flexibility of PGAM1 catalytic pocket which allows a

six-carbon sugar metabolite besides its three-carbon substrates 2-PG, 3-PG and cofactors 2,3-BPG, PEP and 1,3-BPG34,35. Such metabolite-metabolic enzyme interaction has been proved to be

specific since other endogenous sugar phosphates with a six-carbon sugar backbone such as F1P, F6P, G1P, G6P, M1P and M6P cannot activate PGAM1. FBP analogues, including α-MeFBP, β-MeFBP,

4-deoxyFBP, 1-DMeFBP, 6-DMeFBP and 1,6-DMeFBP that block the hydrogen bonding interaction between FBP and PGAM1 were designed and obtained by chemical synthesis. Compared with FBP, they all

show inhibiting effects against PGAM1 enzymatic activity and therefore offered a type of PGAM1 inhibitors based on the endogenous sugar metabolite scaffold. FBP activates PGAM1 in living

cancer cells to support the Warburg effect and cell proliferation. Given the significant role of PGAM1 in tumor growth and cancer cell migration, such specific interaction between the

endogenous glycolytic metabolite FBP with the glycolytic enzyme PGAM1 might work as a unique feedforward mechanism, mediated by covalent signaling through histidine phosphorylation, to

support energy and biomass demands in proliferating cancer cells. Representative PGAM1 inhibitor 1-DMeFBP was found to be able to reduce its histidine phosphorylation level in living cells

and afterwards reduce glycolysis and inhibit cell proliferation. 1-DMeFBP served as a valuable tool compound to confirm the signaling role of FBP in intrapathway PGAM1 phosphorylation,

glycolysis acceleration as well as the supportive effect for cancer cell proliferation. Importantly, 1-DMeFBP is an orthostatic inhibitor of PGAM1 that derivatized from an identified

endogenous PGAM1 activator. Our study offers an approach of manipulating cell fates through chemical regulation of the interactions between sugar metabolite and metabolic enzymes. METHODS

CELL CULTURE HepG2 and HEK293T cells were obtained from China Center for Type Culture Collection and maintained in DMEM supplemented with 10% (vol/vol) FBS, 100 U/mL penicillin, and 100 

mg/mL streptomycin in a humidified atmosphere at 37 °C with 5% CO2. The cells were harvested by scraping and the pellets were washed with phosphate buffered saline (PBS). After

centrifugation, the cells were collected, frozen and stored at −80 °C. SGRNA AND PLASMIDS TRANSFECTION LentiCRISPR v2 system was employed to knock out PGAM1 in HepG2 cell51, and the sgRNA

sequences were designed as following: sgPGAM1-F1 CACCGACAGCAACATCAGTAAGGTA sgPGAM1-R1 AAACTACCTTACTGATGTTGCTGTC HEK293T cells were grown in DMEM supplemented with 10% FBS to 60% confluence.

The transfection complex was prepared as following according to the manufacturer’s instructions. The pLKO.1-PGAM1 sgRNA vector was added to Lipofectamine3000 (Invitrogen) in 500 µL OPTI-MEM

serum-free medium along with psPAX2 packaging vector and pMD2.G envelop vector (4:3:1). About 6 h after transfection, the media was replaced by fresh DMEM containing 10% FBS. After

additional 48 h incubation at 37 °C, the supernatant was collected, filtered by 0.45 µm microfiltration membrane and infected HepG2 cells in the presence of 10 µg/mL polybrene (Millipore)

for 24 h. Cells stably expressing PGAM1 sgRNA were selected in the presence of puromycin (1 mg/mL) before plating for experiments. WESTERN BLOT ANALYSIS Cells were collected by RIPA lysate

(Beyotime) with cocktail protease and phosphatase inhibitors (Beyotime) and protein concentrations were determined using a BCA assay (Thermo Scientific), normalized to 2 mg/mL in 25 µL

lysate. After adding 6.25 µL loading buffer, proteins were subjected to SDS-PAGE separation and transferred to PVDF membranes (Immobilon-P, Millipore). Membranes were blocked for 1 h at room

temperature with 5% milk in Tris-buffered saline with 0.1% Tween-20 (TBST) and blotted with primary antibodies at 4 °C more than 12 h. The following antibodies were used: anti-PGAM1,

anti-PGM1, anti-1pHis, anti-3pHis, anti-PMM2, anti-GAPDH, anti-PRPS1, anti-SOS1, anti-ALDOA, anti-GARS, anti-HDAC2 (Abcam). Anti-POLR2A, anti-PDXDC1, anti-PNPT1 and anti-TRIM25, anti-RAN,

anti-OXSR1, anti-PKLR (Proteintech). After washing with TBST three times for 30 min, membranes were incubated with horseradish peroxidase-conjugated anti-rabbit IgG (1:5000) or anti-mouse

IgG (1:5000) antibodies at room temperature for 1 h and the immunoblots were scanned with ECL (Thermo Fisher Scientific). MOLECULAR DOCKING The crystal structure of human B type

phosphoglycerate mutase H11 phosphorylated form (PDB: 4gpz) is uploaded from the RCSB database at a resolution of 1.65 Å. A molecular docking study was carried out to determine the

interactions between candidate molecules and the PGAM1 active site. Referring to the template of His11 phosphorylated PGAM1 by cofactor 2,3-BPG with ligand 2-(_N_-morpholino) ethanesulfonic

acid (MES)52, amino acids within 10 Å around His11 were selected for molecular docking of sugar phosphates. Waters were deleted except the one form water bridge with the small molecule and

residues of the protein. After the addition of missing hydrogen atoms by standard protein preparation protocol. The structure of FBP was minimized using Tripos force field. The docking of

FBP was carried out using Flexx with standard precision protocol in Tripos software. In the end, the default 30 binding conformations were given. MOLECULAR DYNAMICS (MD) SIMULATION The

atomic coordinates of the PGAM1 complex were extracted from the docking model. These PGAM1 complex structures were constructed by Flexx with standard precision protocol in Tripos software.

All initial structures were first minimized in Discovery studio 2023 to eliminate any possible overlaps or clashes. CHARMm was used to perform efficient simulations with periodic boundary

conditions. Hydrogen atoms were added using the Discovery studio 2023. Counterions were used to maintain system neutrality. All systems were solvated in a truncated octahedron box of waters

with a buffer of 12 Å. The pairwise interactions (van der Waals and direct Coulomb) were computed with a cutoff distance of 8 Å. All MD simulations were accelerated with the CUDA version of

PMEMD in GPU cores of NVIDIA Tesla K20. In order to impress the importance of different residues during the MD simulation, we found the non-bond monitor of MD simulation of PGAM1–FBP complex

(FBP donates its C1-_O_-phosphate to the catalytic His11 of PGAM1) and PGAM1–FBP complex (FBP donates its C6-_O_-phosphate to the His11 of PGAM1), then we calculated the probability of

hydrogen bonding between FBP and different PGAM1 active site residues. PGAM1 ENZYME ACTIVITY ASSAY The PGAM1 enzyme activity assay was operated as previously reported53. Briefly, after

incubation of FBP dissolved in H2O of indicated concentration with 49 µL of 50 nM recombinant PGAM1, 50 µL enzyme mix containing 100 mM Tris-HCl, 100 mM KCl, 5 mM MgCl2, 1 mM ADP, 0.2 mM

NADH, 4 mM 3-PG, 3 units/mL enolase (Sigma–Aldrich), 3 units/mL recombinant pyruvate kinase M2 (Sigma–Aldrich), and 0.6 units/mL recombinant LDH (Sigma–Aldrich) was added. The decrease in OD

at 340 nm was measured as PGAM1 activity. PGM1 AND PMM2 ENZYME ACTIVITY ASSAY The enzyme activity of PGM1 was monitored using Phosphoglucomutase Activity Assay kit (ab155896). The PMM2

enzyme activity assay was operated as previously reported54. The assay was carried out for 3–4 h, with absorbance readings every 30 min at 340 nm. All of the reagents needed to convert

mannose 1-phosphate to gluconate 6-phosphate and NADPH were added to the samples except for the enzyme measured in the assay. The reaction solution consisted of 50 mM HEPES (pH 7.1), 5 mM

MgCl2, 2 µg/mL phosphoglucose isomerase (Roche), 1.7 µg/mL phosphomannose isomerase (Sigma-Aldrich), 10 µg/mL glucose 6-phosphate dehydrogenase (Roche), 1 µM glucose 1,6-bisphosphate, 0.25 

mM NADP (Roche), and 0.34 mM mannose 1-phosphate (Sigma-Aldrich). Assay buffer without the substrate mannose 1-phosphate was used as the control. THERMAL PROTEOME PROFILING Cellular thermal

shift assay experiments were performed as previously described23. Briefly, HepG2 cells were resuspended in ice-cold PBS (containing EDTA-free cocktails and 0.1% Triton) and lysed by

sonication in ice and the cell lysates were collected by centrifugation (20,000 g, 30 min) at 4 °C to remove the debris. The protein concentration was determined by using the BCA protein

assay kit (Pierce). 1 M FBP stock or H2O was added to the cell extract to HepG2 cell lysates (2 mg/mL) with 5 mM final FBP concentration. The samples were then incubated for 30 min at 25 °C,

divided into 6 aliquots and transferred into 0.2 mL polymerase chain reaction (PCR) tubes. Each sample was heated in parallel for 3 min to the respective temperature (range: 37–62 °C).

Subsequently, the samples were centrifuged at 20,000 g for 20 min at 4 °C and supernatant was collected. The supernatant was divided into two parts. One part was subjected to

gel-electrophoresis for western blotting analysis, the other part of 57 °C group and 37 °C group were for in-solution digestion, dimethyl labeling and LC-MS/MS analysis (For the proteome

profiling, we determined the temperature points 57 °C according to the previous TPP studies where Tm of the majority of the proteins fall at 57 °C). IN-SOLUTION DIGESTION AND DIMETHYL

LABELING Each sample was transferred into a 10-K filter and changed with 6 M urea/100 mM TEAB buffer for three times to remove FBP. 10 mM DTT was added and incubated at 37 °C for 30 min

followed by the addition of 20 mM iodoacetamide (IAA) for 30 min at 37 °C in the dark. The samples were changed with 100 mM TEAB buffer for four times to remove excess urea, DTT and IAA. 2.5

 µg trypsin resuspended in 100 mM TEAB was added into per 100 µg of proteome samples and digested overnight at 37  °C. Then peptide samples were labeled with dimethyl reagents. The 57 °C

heated group were reacted with 4 μL of 4% “light” formaldehyde (CH2O) (Sigma) every 100 μL peptides, 37 °C heated group were reacted with “heavy” formaldehyde (13CD2O) (Sigma), respectively.

The resulting solution were treated with 4 μL of 0.6 M sodium cyanoborohydride (Sigma) and incubated at room temperature for 1 h. The reaction was quenched by adding 16 μL of 1% ammonia and

8 μL of 5% formic acid. The “light” and “heavy” samples were combined and subjected for fraction. The peptides were separated into 20 fractions by a high-pH reverse phase C18 column (Agela

Technologies) using Agilent HPLC system. Mobile phase A: 2% MeCN–98% H2O (adjusted to pH 10 with NH3·H2O); B: 98% MeCN–2% H2O (adjusted to pH 10 with NH3·H2O). Samples were separated using a

20 min gradient of buffer B at a flow rate of 1 mL/min, as follows: 0 min 0% B; 0.1 min 5% B; 1 min 8% B; 7 min 14% B; 12 min 24% B; 16 min 40% B; 18 min 95% B; 20 min 15% B. The columns

were operated at 45 °C and the temperature was controlled by a built-in column heater. The 20 fractions were combined into 12 fractions, dried in a SpeedVac and subjected to LC-MS/MS

analysis. PROTEIN EXPRESSION AND PURIFICATION DNA encoding PGAM1 (NP_002620.1), PKM2 (NP_001193725.1), PMM2 (NP_000294.1), PGM1 (NP_001166289.1), PKLR (000289.1), EHHADH (NP_001957.2), HDAC2

(NP_001518.3), OXSR1 (NP_005100.1), PDXDC1 (NP_055842.2), RAN (001287725.1), PGI (NP_000166.2), G6PD (NP_000393.4), PRPS1 (NP_001191331.1), ENO1(NP_001188412.1), LDHA (NP_001128711.1), NEM2

(NP_002503.1) and GAPDH (NP_001243728.1), were amplified from HepG2 cell line cDNA libraries as the template and individually cloned into a modified pET-32M vector with Trx tag removed. All

mutants were created using the standard two-step PCR methods. The primers used for constructing the mutants are shown in Supplementary Dataset 4. All plasmids were transferred into _E.

coli_ BL21 (DE3) cells. The cells were cultured in LB media (containing 5 g yeast extract, 10 g NaCl and 10 g tryptone per 1 L) at 37 °C until OD at 600 nm reached between 0.6 and 0.8. The

protein overexpression was induced by the addition of IPTG to a final concentration of 0.1 mM. After IPTG induction at 16 °C for 16 h, bacterial cells were collected by centrifuging at 4000 

rpm for 20 min, and frozen at −20 °C until further use. The collected bacteria were resuspended in 50 mM Tris, pH 8.0, 500 mM NaCl, 5 mM imidazole. The bacteria cells were lysed by

sonication. The supernatant was collected after centrifugation at 15,000 g for 30 min and loaded on Ni-chelating Ni2+ Sepharose 6 FF beads (Cytiva). After washed by 50 mM Tris, pH 8.0, 500 

mM NaCl, 30 mM imidazole, the target proteins were eluted by buffer containing 50 mM Tris, pH 8.0, 500 mM NaCl, 30 mM imidazole. The elution was purified by size-exclusion chromatography

(HiLoad 26/60 Superdex 200 pg/ HILOAD 26/600 SUPERDEX 75 PG, Cytiva) in 50 mM Tris, PH 7.5, 100 mM NaCl, 1 mM DTT. The recombinant PGAM1 were subjected to a further purification step by

ion-exchange chromatography (HiPrep Q HP 16/10, Cytiva) followed by desalting (PD-10, Cytiva) to a buffer containing 50 mM Tris, pH 7.5, 100 mM NaCl, 5 mM MgCl2. THERMAL SHIFT ASSAY WITH

PURIFIED RECOMBINANT PROTEINS Each purified protein (0.2 μg/μL) was treated with FBP or H2O at 25 °C for 30 min. The mixture was divided into six aliquots and transferred into 0.2 mL PCR

tubes. All of the FBP or H2O-treated samples were heated in parallel for 3 min to their specified temperatures, respectively. The samples were subsequently centrifuged at 20,000 g for 20 min

at 4 °C and the supernatants were collected for SDS-PAGE analysis. The gel was stained by Coomassie blue. CELL VIABILITY ASSAY HepG2 cells (5000 cells/well) were seeded in 96-well plates

allowed to adhere overnight and treated with a diluted series of FBP with no glucose medium or 5 mM glucose for 24 h, or treated with a diluted series of FBP derivatives with 25 mM glucose

medium for 24 h or 48 h. Cell viability was determined using the Cell Counting Kit-8 (CCK8) colorimetric assay. The results were analyzed and displayed using GraphPad Prism 9.0. LC-MS/MS

QUANTIFICATION OF INTRACELLULAR FBP CONCENTRATIONS HepG2 cells were seeded in 6-well plates (2 × 106 cells/well) in triplicates allowed to adhere overnight. Treatment groups were stimulated

by 10 mM FBP for 0.5 h in medium with 5 mM or 25 mM glucose supplement. Control groups were treated with H2O. All cells samples were washed with PBS three times and extracted with 500 µL of

MeOH: H2O mixture (8:2, v/v, −80 °C) with 0.1 µg/mL 13C6-FBP in it as internal standard for quantification. Samples were centrifuged for 15 min at 20,000 g and 4 °C to settle any particulate

matter. Supernatants were dried in a SpeedVac and submitted to LC-MS/MS analysis on Agilent 6490 Triple Quadrupole LC-MS system. Chromatographic separations were carried out on an ACQUITY

UPLC BEH Amide (2.1 × 100 mm, 3.5 mm) at an oven temperature of 40 °C. The mobile phase of LC-MS/MS was composed of 10 mM NH4HCO3 in Milli-Q Water (A) and Acetonitrile (B) in negative ion

mode. Maintained at a flow rate of 0.3 mL/min for 2 min in the 5% phase B, the B ratio rose to 23% in 2–3 min, then returned to the 5% B phase to in 3–5 min and remained until 8 min.

DERIVATIZATION BASED LC-MS/MS METHOD FOR DETECTION OF F6P AND F1P The purified PGAM1 (10 mg/mL in enzyme buffer) was diluted to 1.5 mg/mL by H2O, then incubated with 10 mM FBP (sample) or

H2O (control) as the reaction system at 25 °C. At the pointed time, fixed volume of the reaction system was diluted to ten-time volume and then PGAM1 was removed by filter in 5 minutes at 4 

°C. The remaining buffer with metabolites was collected. The derivatization reagent, 2-(diazo-methyl)-_N_-methyl-_N_-phenyl-benzamide (2-DMBA) was used to obtain better separation and

identification of F1P and F6P. Briefly, 80 μL of borate buffer (50 mM, pH 7.0) and 10 μL of 2-DMBA were mixed with 10 μL sample, and the reaction solution was vortexed and then incubated at

30 °C for 1 h. LC-MS/MS analysis was performed on Agilent 6490 Triple Quadrupole LC-MS system, and chromatographic separations were carried out on an ACQUITY UPLC HSS T3 C18 Amide (2.1 × 150

 mm, 1.8 mm) at an oven temperature of 50 °C. The mobile phase was composed of 0.1% formic acid in Water (A) and Acetonitrile (B) in negative ion mode. The modified gradients (v/v) were as

following: 0–2 min, 5% (B); 2–40 min, 5–35% (B); 40–40.5 min, 35–90% (B); 40.5–45 min, 90–5% (B), followed by 7 min of re-equilibration with 5% B. MS parameters for the MRM transitions were

optimized by direct infusion. LC-MS/MS DETECTION OF 3-PG AND 2-PG HepG2 cells were seeded in 6-well plates (2 × 106 cells/well) in triplicates allowed to adhere overnight. Treatment groups

were stimulated by 10 mM FBP for 0.5 h in 5 mM glucose or 1 mM 1-DMeFBP for 24 h in 25 mM glucose. Control groups were treated with H2O. All cell samples were extracted with 500 µL of MeOH:

H2O mixture (8:2, v/v, −80 °C). Samples were centrifuged for 15 min at 20,000 g and 4 °C to settle any particulate matter. Supernatants were dried in a SpeedVac and taken to LC-MS/MS

analysis. Chromatographic separations were carried out on an ACQUITY UPLC BEH Amide (2.1 × 100 mm, 3.5 mm) at an oven temperature of 4 °C. The mobile phase of LC-MS/MS was composed of 10 mM

NH4HCO3 in Milli-Q Water (A) and MeCN (B) in negative ion mode. Maintained at a flow rate of 0.3 mL/min for 2 min in the 5% phase B, the B ratio rose to 23% in 2–3 min, then returned to the

5% B phase to in 3–5 min and remained until 8 min. 3-PG and 2-PG both generate fragments with m/z 79 and 97, but at different ratios. The standard mixtures containing 3-PG and 2-PG (in the

ratio of 100:0, 80:20, 60:40, 40:60, 20:80, and 0:100) were used to generate a standard curve of 97/(79 + 97). The 97/(79 + 97) value from experimental samples was calculated and converted

into the 3-PG/total mono-PG ratio using the standard curve. The absolute concentration of mono-PG in WT cells was measured as described. The ion counts of 3-PG and 2-PG were converted to

absolute concentrations by normalizing to the WT total mono-PG concentration. GLUCOSE CONSUMPTION AND LACTATE PRODUCTION HepG2 cells were cultured in 96 wells overnight. The cells were

treated with H2O or 10 mM FBP for 0.5 h, or treated with H2O or 1 mM 1-DMeFBP for 24 h. Glucose consumption in the culture medium were measured using the Glucose (GO) Assay Kit (Abnova)

whereas lactate levels in the culture medium were determined using a Lactate Assay Kit (Cayman). MICROSCALE THERMOPHORESIS (MST) ANALYSIS Purified PGAM1, PGM1 and PMM2 was labeled on lysine

residues using the Protein Labeling Kit RED-NHS 2nd Generation (NanoTemper Technologies). Measurements were carried out in MST-buffer (50 mM Tris-HCl, pH 7.5, 100 mM NaCl, 0.05% Tween-20).

The labeled protein was mixed with FBP and incubated for 30 min at room temperature in the dark before being loaded into Monolith NT.115 Capillaries (Nano Temper Technologies). Samples were

then transferred into the Monolith NT.115 device and MST experiments were carried out at Nano-red. MST was measured using a Monolith NT.115 instrument (NanoTemper Technologies) at an ambient

temperature of 23.5 °C. Instrument parameters were 20% excitation power and Medium MST power. Error bars = SD from two independent experiments. MS IDENTIFICATION OF PHOSPHORYLATION SITES

INDUCED BY FBP Purified PGAM1 (0.2 μg/μL) was incubated with 10 mM FBP or H2O at 25 °C for 30 min. Then the sample was transferred into a 10-K filter and changed with 6 M urea/100 mM TEAB

buffer for three times to remove FBP. 10 mM 1,4-dithiothreitol (DTT) was added and incubated at 37 °C for 30 min followed by the addition of 20 mM iodoacetamide (IAA) for 30 min at 37 °C in

the dark. The samples were changed with 100 mM TEAB buffer for four times to remove excess urea, DTT and IAA. 1 µg trypsin resuspended in 100 mM TEAB was added into per 50 µg of proteome

samples and digested overnight at 37 °C. Then peptide samples were desalted by a C18 column tip, and elution was dried in a SpeedVac and subjected to LC-MS/MS analysis. LC-MS/MS BASED

PROTEOMIC ANALYSIS LC-MS/MS analysis was performed on an Orbitrap Fusion Lumos Tribrid mass spectrometer (Thermo Fisher Scientific) coupled with an Ultimate 3000 LC system. The peptides were

chromatographically separated by a 90 min gradient from 5% to 40% solvent B (A = 0.1% formic acid in water, B = 0.1% formic acid in 80% MeCN) at a flow rate of 300 nL/min. Under the

positive-ion mode, full-scan mass spectra were acquired over the m/z range from 350 to 1800 using the Orbitrap mass analyzer with mass resolution setting of 70,000. MS/MS fragmentation was

performed in a data-dependent mode, of which the 20 most intense ions were selected from each full-scan mass spectrum for high-energy collision induced dissociation (HCD) and MS/MS analysis.

MS/MS spectra were acquired with a resolution setting of 35,000 using the Orbitrap analyzer. Other parameters in the centroid format: isolation window, 1.2 m/z units; default charge, 2+;

normalized collision energy, 32%; maximum IT, 100 ms; dynamic exclusion, 20.0 s. DATA ANALYSIS For the dimethyl labeling-based experiments, LC-MS/MS data was first analyzed by ProLuCID55

with static modification of cysteine (+57.0215 Da, carbamidomethyl). The isotopic modifications (28.0313, and 34.0757 Da for light and heavy labeling respectively) are set as variable

modifications on the N-terminal of a peptide and lysines. The ratios of reductive dimethylation were quantified by the CIMAGE software56. CHEMICAL COMPOUNDS FBP, F1P, G6P, G1P, M6P and M1P

were purchased from Sigma-Aldrich. FBP analogues (α-MeFBP, β-MeFBP, 4-deoxyFBP, 1,6-DMeFBP, 6-DMeFBP and 1-DMeFBP) were synthesized as described in the Supporting Information. STATISTICS All

statistical analyses were performed using GraphPad Prism 9.0. The respective figure legends provide information on the specific statistical test used for each assay. REPORTING SUMMARY

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article. DATA AVAILABILITY The mass spectrometry proteomics data have been

deposited to the ProteomeXchange Consortium (https://proteomecentral.proteomexchange.org) via the iProX partner repository57,58 with the dataset identifier PXD048243

[http://proteomecentral.proteomexchange.org/cgi/GetDataset?ID=PXD016640] (2D-TPP experiments in Jurkat cells). Molecular dynamics (MD) simulation data for the process of phosphate transfer

from FBP to PGAM1 are available via https://doi.org/10.5281/zenodo.13836214. All NMR data are available in the Supplementary Information. All remaining data are available in the Article,

Supplementary Information, and Source Data files. Source data are provided with this paper. The data that support the findings of this study are available from the corresponding authors upon

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big data. _Nucleic Acids Res._ 50, D1522–D1527 (2021). Article  PubMed Central  Google Scholar  Download references ACKNOWLEDGEMENTS We thank Prof. Xing Chen (PKU) for helpful discussions,

Prof. Chu Wang (PKU) for support of chemical proteomics, Dr. Xueshi Mao and Dr. Yue Zhou (IMM) for support of MD simulation, Prof. Jinlan Zhang and Dr. Zhe Wang (IMM) for targeted

metabolomics, Prof. Qin Li and Dr. Qian Wang (SKLNBD) for NMR and MST analysis. This work was funded by National Key Research and Development Plan (Grant No. 2021YFC2102900), National

Natural Science Foundation of China (Grants No. 22377148 and 22207126), CAMS Innovation Fund for Medical Sciences (Grant No. 2021-I2M-1-026), R & D Program of Beijing Municipal Education

Commission (Grant No. KZ202210028043), Non-profit Central Research Institute Fund of Chinese Academy of Medical Sciences (Grant No. 2021-RC350-002), and the Special Research Fund for

Central Universities, Peking Union Medical College (Grant No. 3332022042). Y.G. and H.L. are supported by “1000 Talents Plan” Young Investigator Program respectively. AUTHOR INFORMATION

Author notes * These authors contributed equally: Yanling Zhang, Yafei Cao. AUTHORS AND AFFILIATIONS * State Key Laboratory of Bioactive Substance and Function of Natural Medicines, Beijing

Key Laboratory of Active Substance Discovery of Active Substances Discovery and Druggability Evaluation, Institute of Materia Medica, Chinese Academy of Medical Sciences and Peking Union

Medical College, Beijing, China Yanling Zhang, Yafei Cao, Zhenghui Chen, Bowen Chen, Anhui Wang, Yanshen Guo, Ruolan Xue, Tian Li, Ruiqin Cheng, Ning Zhou, Huixin Luo & Yiqun Geng *

State Key Laboratory of Natural and Biomimetic Drugs, Chemical Biology Center, School of Pharmaceutical Sciences, Peking University, Beijing, China Xia Wu, Wei Chen & Houhua Li *

Synthetic and Functional Biomolecules Center, Beijing National Laboratory for Molecular Sciences, College of Chemistry and Molecular Engineering, Peking University, Beijing, China Zihua Liu,

 Yuanpei Li & Yuan Liu * Beijing Key Laboratory of DNA Damage Response and College of Life Science, Capital Normal University, Beijing, China Jing Li * Jinhua Institute and College of

Pharmaceutical Sciences, Zhejiang University, Hangzhou, China Xiaohui Zhao * Department of Cardiology and Institute of Vascular Medicine, Peking University Third Hospital, Peking University,

Beijing, China Ming Xu Authors * Yanling Zhang View author publications You can also search for this author inPubMed Google Scholar * Yafei Cao View author publications You can also search

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inPubMed Google Scholar CONTRIBUTIONS Y.Z., H.L. and Y.G. conceived the project. Y.Z. conducted most of the experiments unless otherwise specified. Y.C. and R.X. carried out the synthesis of

six FBP analogues, X.W. and N.Z. conducted the expression and purification of target proteins. Z.C. helped with the biochemical validation of FBP targets. Y.C., B.C., and Y.-S.G. carried

out the molecular docking and MD simulations. Y.C., A.W., W.C. and T.L. performed targeted metabolomics experiments. R.C. helped with the MST experiments. Z.L. helped with the construction

of KO cell line, Y.-P.L. synthesized the Phos-tag reagent, Y.L. helped with MS/MS and MD data analysis. M.X., H.-X.L., X.Z. and J.L. helped with data analysis. Y.G. wrote the manuscript with

input from all authors. CORRESPONDING AUTHORS Correspondence to Yanling Zhang, Houhua Li or Yiqun Geng. ETHICS DECLARATIONS COMPETING INTERESTS The authors declare no competing interests.

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Wu, X. _et al._ Thermal proteome profiling reveals fructose-1,6-bisphosphate as a phosphate donor to activate phosphoglycerate mutase 1. _Nat Commun_ 15, 8936 (2024).

https://doi.org/10.1038/s41467-024-53238-w Download citation * Received: 25 January 2024 * Accepted: 08 October 2024 * Published: 16 October 2024 * DOI:

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