Non-enzymatic Lysine Lactoylation of Glycolytic Enzymes

Brief Communication

Non-enzymatic Lysine Lactoylation of Glycolytic Enzymes Graphical Abstract

Authors Dominique O. Gaffney, Erin Q. Jennings, Colin C. Anderson, ..., Eli Chapman, James R. Roede, James J. Galligan

Correspondence [email protected]

In Brief Gaffney et al. describe a lysine modification derived from a nonenzymatic acyl transfer from the secondary glycolytic intermediate, lactoylglutathione. This modification, lysine lactoylation, is enriched on primary glycolytic enzymes and regulates metabolic output.

Highlights d

Lysine lactoylation occurs via a non-enzymatic acyl transfer from lactoylglutathione

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Glycolytic enzymes are heavily modified by lactoylation

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Glyoxalase 2 is the critical regulator for lactoylglutathione and lysine lactoylation

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Glycolytic output is significantly repressed in glyoxalase 2 knockout cells

Gaffney et al., 2020, Cell Chemical Biology 27, 1–8 January 16, 2020 ª 2019 Elsevier Ltd. https://doi.org/10.1016/j.chembiol.2019.11.005

Please cite this article in press as: Gaffney et al., Non-enzymatic Lysine Lactoylation of Glycolytic Enzymes, Cell Chemical Biology (2019), https:// doi.org/10.1016/j.chembiol.2019.11.005

Cell Chemical Biology

Brief Communication Non-enzymatic Lysine Lactoylation of Glycolytic Enzymes Dominique O. Gaffney,1,5 Erin Q. Jennings,1,5 Colin C. Anderson,2 John O. Marentette,2 Taoda Shi,1 Anne-Mette Schou Oxvig,3 Matthew D. Streeter,4 Mogens Johannsen,3 David A. Spiegel,4 Eli Chapman,1 James R. Roede,2 and James J. Galligan1,6,* 1Department

of Pharmacology and Toxicology, College of Pharmacy, University of Arizona, Tucson, AZ 85721, USA of Pharmaceutical Sciences, Skaggs School of Pharmacy and Pharmaceutical Sciences, University of Colorado, Aurora, CO 80045, USA 3Department of Forensic Medicine, Aarhus University, 8200 Aarhus, Denmark 4Department of Chemistry, Yale University, New Haven, CT 06520, USA 5These authors contributed equally 6Lead Contact *Correspondence: [email protected] https://doi.org/10.1016/j.chembiol.2019.11.005 2Department

SUMMARY

Post-translational modifications (PTMs) regulate enzyme structure and function to expand the functional proteome. Many of these PTMs are derived from cellular metabolites and serve as feedback and feedforward mechanisms of regulation. We have identified a PTM that is derived from the glycolytic by-product, methylglyoxal. This reactive metabolite is rapidly conjugated to glutathione via glyoxalase 1, generating lactoylglutathione (LGSH). LGSH is hydrolyzed by glyoxalase 2 (GLO2), cycling glutathione and generating D-lactate. We have identified the non-enzymatic acyl transfer of the lactate moiety from LGSH to protein Lys residues, generating a ‘‘LactoylLys’’ modification on proteins. GLO2 knockout cells have elevated LGSH and a consequent marked increase in LactoylLys. Using an alkyne-tagged methylglyoxal analog, we show that these modifications are enriched on glycolytic enzymes and regulate glycolysis. Collectively, these data suggest a previously unexplored feedback mechanism that may serve to regulate glycolytic flux under hyperglycemic or Warburg-like conditions. INTRODUCTION Cell metabolism generates numerous intermediates that serve to regulate metabolic feedforward and feedback (Moellering and Cravatt, 2013). These mechanisms of regulation are achieved, in part, through the post-translational modification (PTM) of target enzymes. This is perhaps most notable with acetyl-coenzyme A (CoA), which serves as the primary source for Lys acetylation, regulating enzymatic activity and structure (Zhao et al., 2010). Lys acetylation, like many PTMs, is regulated enzymatically through the addition (Lys acetyltransferases) and removal (Lys deacetylases) of the acetyl group (Zhao et al., 2010). While

most PTMs are tightly regulated, there is growing interest in non-enzymatic PTMs and their role in metabolic regulation and cell homeostasis (Moellering and Cravatt, 2013; Zheng et al., 2019b). The reactive glycolytic by-product, methylglyoxal (MGO), plays a critical role in regulating transcription, chromatin structure, and activation of the Nrf2 pathway (Bollong et al., 2018; Galligan et al., 2018; Zheng et al., 2019a). Intracellular concentrations of this reactive molecule are kept in the low-micromolar range via the glyoxalase cycle, which is composed of two enzymes, glyoxalase 1 (GLO1) and GLO2 (Rabbani et al., 2014). GLO1 facilitates the isomerization of the labile glutathione (GSH)-MGO hemithioacetal, generating the stable product, S,D-lactoylglutathione (LGSH); GLO2 then hydrolyzes LGSH, recycling GSH and generating D-lactate (Rabbani et al., 2014). GLO1 has been investigated in the context of diabetes, cardiovascular disease, and numerous cancers (Rabbani et al., 2014). Additionally, decreased GLO1 expression is associated with elevations in MGO and MGO-derived PTMs, and has been identified as a biomarker for diabetic nephropathy (Giacco et al., 2014; Qi et al., 2017). Furthermore, MGO-modified histones are elevated in breast cancer (Zheng et al., 2019a). Despite these thorough investigations, unfortunately the role of GLO2 in controlling cellular homeostasis and MGO modifications remains far less understood. In a recent report by James et al. (2017), acetyl-CoA was found to be a major source of mitochondrial Lys acetylation through non-enzymatic, pH-driven acyl transfer; this mechanism was also demonstrated with S-acetylGSH. Interestingly, GLO2 is capable of regulating a broad range of S-acyl GSH conjugates, including S-acetylGSH (James et al., 2017; Talesa et al., 1989). Although the contribution of acetylGSH to the pool of acetylated Lys is thought to be minimal, this led us to hypothesize that the canonical GLO2 substrate, LGSH, may serve as an acyl donor for a new class of Lys modifications, LactoylLys. In this Brief Communication, we describe LactoylLys as a functionally relevant PTM in cells. Using a reactivity-based proteinprofiling approach, we identify 350 LactoylLys modified proteins, which are significantly enriched on glycolytic enzymes. The levels of these modifications are dependent on the intracellular concentrations of LGSH, which are significantly elevated in

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the absence of GLO2. Lastly, we demonstrate that ablation of GLO2 results in a significant reduction in primary glycolytic metabolites. RESULTS Lactoylation of Lys Residues Results from a Non-enzymatic Acyl Transfer The non-enzymatic transfer of the acetyl group on acetyl-CoA to the e-amine of Lys is a major source of mitochondrial Lys acetylation (James et al., 2017; Wagner and Payne, 2013). A similar acyl-transfer mechanism occurs with acetylGSH, generating acetylLys (James et al., 2017). Due to the chemical similarities between acetylGSH and LGSH, we hypothesized that a similar acyl transfer would take place, yielding a lactate-modified Lys (LactoylLys) (Figure 1A). MGO is also capable of modifying Lys residues directly, generating the isomeric compound Ne-(carboxyethyl)Lys (CEL) (Figure 1B). We have previously demonstrated that CEL, compared with MGO-Arg PTMs, is not abundant in biological settings (Galligan et al., 2018). To distinguish between these isomers, we used deuterated CEL (CEL-d4) as an internal standard and synthesized an authentic LactoylLys reference standard (Figure 1B and Supplemental Information). Using a previously reported method to quantify Lys and Arg PTMs in various biological settings, we are able to chromatographically separate LactoylLys and CEL (Figure 1B) (Galligan et al., 2017). To evaluate the direct transfer of the lactate group from LGSH to Lys residues, we incubated purified histone H4 with 1 mM LGSH or 1 mM MGO for 24 h (Figure 1C). H4 is devoid of Cys residues, thus removing the requirement for Cys migration as described by James et al. (2017). As shown in Figures 1D–1F, incubation of H4 with MGO results in robust generation of the Arg-derived PTMs, MGO hydroimidazolone (MG-H1) and carboxyethylArg (CEA), and modest generation of CEL, which is consistent with our previous findings (Galligan et al., 2018). Conversely, when incubated with LGSH, no Arg modifications were detected; however, a marked increase in LactoylLys was observed (Figure 1G) with representative tandem mass spectrometry (MS/MS) shown in Figure 1H (H4K31Lactoyl). These findings were also confirmed using phosphoglycerate kinase 1 (PGK1) (Figure S1), which contains four Lys residues in close proximity to nucleophilic Cys residues as described by James et al. (2017). The MS/MS of one of these sites, Lys94, is also shown in Figure S1. These findings thus justified the investigation of LactoylLys modifications in a moe physiologically relevant setting. GLO2 Regulates Cellular GSH and LGSH LGSH is enzymatically generated via GLO1, facilitating the isomerization of the spontaneously generated MGO-GSH hemithioacetal (Rabbani et al., 2014). This stable product is then hydrolyzed by GLO2, regenerating GSH and producing D-lactate (Figure 2A). GLO2 is the only reported enzyme to hydrolyze LGSH in cells (Rabbani et al., 2014). Therefore, to investigate LactoylLys modifications in cells, we performed CRISPR/Cas9 genome editing to generate HEK293 cells lacking GLO2 (GLO2 / ). To isolate the effects of LGSH and LactoylLys from those of MGO, we used a previously characterized GLO1 / cell line to generate cells deficient in any glyoxalase activity 2 Cell Chemical Biology 27, 1–8, January 16, 2020

(GLO1/2 / ) (Galligan et al., 2018) (Figure 2B). We have previously demonstrated deficient MGO metabolism in GLO1 / cells, with MGO-derived PTMs peaking following 6 h of treatment (Galligan et al., 2018). Consistent with our previous findings, GLO1 / cells displayed significantly elevated concentrations of MGO when challenged with 50 mM MGO for 6 h, while wild-type (WT) cells are able to sufficiently metabolize the bolus dose (Galligan et al., 2018). Conversely, GLO2 / cells are able to efficiently detoxify MGO with no significant differences compared with vehicle-treated controls (Figure 2C). Consistent with this metabolism, GLO2 / cells have elevated LGSH, which is exacerbated when challenged with MGO (Figure 2D); these effects are concomitant with a significant reduction in GSH (Figure 2E). Thus, the elevations in LGSH in GLO2 / cells provided us with the opportunity to study the biological abundance and effects of LactoylLys modifications in human cells.

LactoylLys Levels Are Dependent on LGSH and GLO2 To evaluate LactoylLys levels in cells, we applied a recently characterized chemical probe (Sibbersen et al., 2018). This alkynetagged MGO analog (alkMGO) is metabolized by the glyoxalase cycle at similar rates to that of MGO (Kold-Christensen et al., 2019; Sibbersen et al., 2018). By appending an alkyne tag, we are able to perform click chemistry using an N3-IR fluorophore to visualize protein modifications in cells (Figure 3A). As shown in Figure 3B, WT cells display minimal protein labeling when exposed to 50 mM alkMGO for 6 h, consistent with the efficient metabolism of this probe through the glyoxalase cycle. GLO1 / and GLO1/2 / cells, however, display marked increases in protein modification, consistent with the non-enzymatic generation of MG-H1 and CEA. Despite efficient conjugation of alkMGO to GSH (yielding alkLGSH) in GLO2 / cells, a marked increase in protein labeling was also observed when compared with WT controls (Figure 3B). To determine the composition of these modifications, we treated cells with either vehicle or 50 mM MGO for 6 h (consistent with Figures 2C–2E) and performed QuARKMod to quantify Lys and Arg PTMs. In support of our previous findings, GLO1 / cells display a marked increase in MG-H1, CEA, and CEL compared with WT controls (Figures 3C–3E) (Galligan et al., 2018); in contrast, GLO2 / cells display a significant elevation in LactoylLys (Figure 3F). These data also demonstrate that the generation of LactoylLys modifications in cells is dependent on the generation of LGSH, as no significant elevations were observed in GLO1/2 / cells compared with WT controls. We next evaluated the abundance of these PTMs under varying glycemic conditions by treating WT and GLO2 / cells with 0, 5, or 25 mM glucose for 24 h. As shown in Figure S2, a gradual rise in LactoylLys modifications was observed in WT cells, which was exacerbated in GLO2 / cells. These data further confirm glucose as the source of these PTMs. As cell culture conditions are notably high in glucose concentrations, we sought to evaluate the abundance of these PTMs in a more physiologically relevant system. Thus, we isolated tissues from WT C57BL/6J mice and quantified MGO-derived PTMs. As shown in Figures 3G–3J, LactoylLys levels are not only basally abundant but highest in tissues associated with a high glycolytic output (e.g., liver and muscle).

Please cite this article in press as: Gaffney et al., Non-enzymatic Lysine Lactoylation of Glycolytic Enzymes, Cell Chemical Biology (2019), https:// doi.org/10.1016/j.chembiol.2019.11.005

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Figure 1. LactoylLys Is Generated through a Non-enzymatic Acyl Transfer from LGSH to Lys (A) Mechanism of LactoylLys formation. (B) Synthetic standards for CEL-d4 and LactoylLys demonstrate chromatographic separation, allowing for quantitation via MRM-MS. (C) Recombinant histone H4 (2 mg) was treated with either MGO or LGSH (1 mM) for 24 h at 37 C. n = 3. +/ StDev. (D–G) QuARKMod was performed to quantify MG-H1 (D), CEA (E), CEL (F), and LactoylLys (G) modifications following incubation with either MGO or LGSH, as described in (C). (H) MS/MS of a LactoylLys modification detected on H4K31 (5.72 ppm mass error).

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Figure 2. GLO2 Regulates GSH and LGSH in Cells (A) The glyoxalase cycle. (B) Generation of GLO1 / , GLO2 / , and GLO1/2 / HEK293 cells. (C) MGO detoxification is achieved through GLO1. n = 3, +/ StDev; **p < 0.01, ***p < 0.001. (D and E) Significant increases in LGSH are observed in GLO2 / cells (D), resulting in a concomitant decrease in GSH (E). n = 3, +/ StDev; *p < 0.05, **p < 0.01, ***p < 0.001.

Glycolytic Enzymes Are Major Targets for LactoylLys Modification We next sought to identify and quantify proteins modified by LactoylLys using a SILAC (stable isotope labeling of amino acids in cell culture) model (Figure 4A) (Beavers et al., 2017). Both WT (‘‘Light’’) and GLO2 / (‘‘Heavy’’) were treated with 50 mM alkMGO for 6 h (consistent with Figures 2 and 3). Click chemistry was performed using a UV-cleavable N3-biotin and modified proteins were enriched with streptavidin beads (Figure 4A) (Kim et al., 2009). Using this approach, we identified 350 modified proteins; GLO1 and glyoxalase domain-containing protein 4 (GLOD4) were among the most heavily modified proteins, which is consistent with their utilization of MGO as a substrate (Figure S3 and Table S1). To gain insight into the biological role of these PTMs, we performed bioinformatics clustering with DAVID (Huang da et al., 2009) and KEGG analysis. These results reveal enrichment for proteins associated with carbon metabolism and glycolysis (Figure 4B and Table S2). We thus mapped identified proteins to these metabolic pathways, which revealed a striking enrichment for glycolysis (Figure 4C: identified proteins are displayed in red). These proteomic results were then validated via immunoblotting for key glycolytic enzymes in both input and eluate fractions from WT and GLO2 / cells exposed to 50 mM alkMGO for 6 h, showing no alterations in the basal levels of these proteins (Figure S4). Due to this enrichment for glycolytic and carbon metabolism enzymes, we performed a targeted metabolomics approach in WT and GLO2 / cells exposed to 50 mM MGO for 6 h. As shown 4 Cell Chemical Biology 27, 1–8, January 16, 2020

in Figure 4D, GLO2 / cells display a relative decrease in glycolytic metabolites downstream of DHAP/GA3P, which is exacerbated in the presence of MGO. These effects are not due to significant alterations in glucose uptake nor related to the committed step in glycolysis (PFK), as no significant differences were observed upstream of bisphosphoglycerate. We also investigated tricarboxylic acid (TCA) cycle metabolites, as well as co-factors used in these biochemical pathways. No significant alterations in any of these metabolites or co-factors were observed, supporting our proteomics data and indicating a possible regulatory feedback for glycolysis.

DISCUSSION In this article, we describe a Lys acylation derived from the non-enzymatic reaction with LGSH. This modification, LactoylLys, is basally present, abundant in highly glycolytic tissues, and enriched on glycolytic enzymes, resulting in decreased glycolytic output. This proposed feedback mechanism is similar to that of 3-phosphoglyceryl-Lys, which is derived from the primary glycolytic metabolite, 1,3-bisphosphoglycerate (Moellering and Cravatt, 2013). This non-enzymatic modification is enriched on glycolytic enzymes, inhibits enzymatic activity, and decreases glycolytic metabolites (Moellering and Cravatt, 2013). Our results demonstrate a similar non-enzymatic mechanism of metabolic regulation, resulting

Please cite this article in press as: Gaffney et al., Non-enzymatic Lysine Lactoylation of Glycolytic Enzymes, Cell Chemical Biology (2019), https:// doi.org/10.1016/j.chembiol.2019.11.005

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Figure 3. Reactivity-Based Protein Profiling Demonstrates Marked Elevations in Modified Proteins in GLO2–/– Cells (A) Overview of the reactivity-based protein-profiling approach performed. (B) Modified proteins are observed in each knockout cell line treated with 50 mM alkMGO for 6 h. (C–F) QuARKMod was performed to determine the composition of MGO-derived PTMs in each cell line in vehicle and 50 mM MGO-treated cells. The absence of GLO1 results in a significant increase in Arg-derived MGO modifications, while ablation of GLO2 leads to a robust increase in LactoylLys modifications. n = 6, +/ StDev; *p < 0.05, **p < 0.01, ***p < 0.001. (G–J) Basal levels of MGO-derived PTMs were quantified in tissues collected from WT C57Bl6/J mice (n = 3, +/ StDev).

in a global decrease in glycolytic output that is dependent on LGSH levels. GLO2 is the rate-limiting enzyme in the glyoxalase pathway and is the only enzyme known to hydrolyze LGSH in mammalian cells (Rabbani et al., 2014); not surprisingly, ablation of GLO2 results in the accumulation of LGSH (Figure 2D). These data demonstrate the importance of GLO2 in regulating cellular GSH concentrations and suggest that a sizable pool of GSH may be dedicated to dealing with MGO. In healthy individuals, it is estimated that 0.1% of the glucotriose flux results in MGO, yielding micromolar concentrations (Rabbani et al., 2016b). Although the glyoxalase cycle is highly efficient, the sustained generation of MGO results in quantifiable protein adducts, and disruptions in GLO1 amplify these PTMs (Galligan et al., 2018;

Rabbani et al., 2016a). As a result, GLO1 has been studied extensively in the context of diabetes and cancer (Rabbani et al., 2014); the role of GLO2, however, is largely unknown. In a report by Antognelli et al. (2017), GLO2 was found to be elevated in prostate cancer. These studies demonstrate that silencing of GLO2 through small interfering RNA knockdown is an effective therapeutic strategy to slow prostate cancer cell migration and invasion (Antognelli et al., 2017). In addition to the results presented here, these findings suggest that perhaps the balance between GLO1 and GLO2 is the critical component in evaluating glyoxalase function in disease. Amplification of GLO1, without compensatory induction of GLO2, may tip the balance in favor of LactoylLys generation, and thus hinder glycolytic flux and cell growth. Cell Chemical Biology 27, 1–8, January 16, 2020 5

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Figure 4. LactoylLys Modifications Are Enriched on Glycolytic Proteins (A) Enrichment strategy to identify and quantify protein targets for LactoylLys modifications in cells. (B) A total of 350 proteins were identified using this approach; DAVID analysis was performed on identified proteins to reveal carbon metabolism and glycolysis as heavily enriched pathways in this dataset. (C) Proteins identified (red) are mapped to glycolysis, TCA cycle, and pentose phosphate pathway demonstrating clear enrichment for glycolysis. (D) Targeted metabolomics reveals a global reduction in glycolytic metabolites in GLO2 / cells, which is exacerbated in the presence of MGO (50 mM, 6 h). Data are presented as a log2 fold change (n = 6).

James et al. (2017) have recently described a thioester exchange between acetyl-CoA (and acetyl-GSH) and protein Cys residues; a subsequent intramolecular SN-transfer then gives rise to acetylLys. It is estimated that this acyl-transfer mechanism is responsible for the bulk of mitochondrial Lys acetylation (James et al., 2017). To prevent accumulation of acetylated proteins in the mitochondria, sirtuins selectively remove the acetyl group, restoring homeostatic enzyme activity (Houtkooper et al., 2012). Our proteomics inventory of LactoylLys modified proteins reveals enrichment for glycolytic enzymes, suggesting that these PTMs may be regulated at the ‘‘eraser’’ level. Although the existence of a LactoylLys ‘‘writer’’ cannot be dispelled, our data suggest that LGSH levels are the 6 Cell Chemical Biology 27, 1–8, January 16, 2020

key driver to the generation of these PTMs in cells. Future studies will be focused on the identification and site-specific regulation of LactoylLys modifications by enzymatic ‘‘eraser’’ proteins. Agreeing with the principal findings presented here, during the review process Zhang et al. (2019) have demonstrated the presence of histone lactylation derived from a lactyl-CoA intermediate. These data provide further evidence for the existence and critical role of this PTM in mediating cellular homeostasis. SIGNIFICANCE Post-translational modifications regulate enzymatic activity, serving as sensors for metabolic feedback. In this

Please cite this article in press as: Gaffney et al., Non-enzymatic Lysine Lactoylation of Glycolytic Enzymes, Cell Chemical Biology (2019), https:// doi.org/10.1016/j.chembiol.2019.11.005

work, we identify a Lys modification, LactoylLys, that is derived from a non-enzymatic acyl-transfer from the glycolytic metabolite, lactoylglutathione. LactoylLys levels are dependent on the levels of lactoylglutathione, which is hydrolyzed in cells by glyoxalase 2. Cells lacking glyoxalase 2 display marked elevations in lactoylglutathione and LactoylLys modifications. Reactivity-based protein profiling reveals glycolytic enzymes as primary targets for LactoylLys modification, resulting in a global decrease of glycolytic output. Collectively, these data reveal a functional Lys modification that serves as a feedback mechanism to regulate glycolysis.

DECLARATION OF INTERESTS

STAR+METHODS

Antognelli, C., Ferri, I., Bellezza, G., Siccu, P., Love, H.D., Talesa, V.N., and Sidoni, A. (2017). Glyoxalase 2 drives tumorigenesis in human prostate cells in a mechanism involving androgen receptor and p53-p21 axis. Mol. Carcinog. 56, 2112–2126.

Detailed methods are provided in the online version of this paper and include the following: d d d

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KEY RESOURCES TABLE LEAD CONTACT AND MATERIALS AVAILABILITY EXPERIMENTAL MODEL AND SUBJECT DETAILS B Mouse Tissues B Cell Culture METHOD DETAILS B Reagents B In Vitro Treatment of Recombinant Proteins with MGO or LGSH B CRISPR-Cas9-Mediated GLO1, GLO2, and GLO1/2 Knockout (-/-) HEK293 Cells B SDS-PAGE and Immunoblotting B Reactivity-Based Protein Profiling B Quantification of Cellular MGO B Quantification of GSH and LGSH B Synthesis of LactoylLys B QuARKMod Analysis for PTM Quantitation B Targeted Metabolomics B Identification of MGO Modified Proteins QUANTIFICATION AND STATISTICAL ANALYSIS DATA AND CODE AVAILABILITY

SUPPLEMENTAL INFORMATION Supplemental Information can be found online at https://doi.org/10.1016/j. chembiol.2019.11.005. ACKNOWLEDGMENTS Financial support was provided by National Institutes of Health grants (R01 ES027593 to J.R.R.), the SENS foundation (D.A.S.), the American Diabetes Association Pathway to Stop Diabetes grant 1-17-VSN-04 (D.A.S.), and the Velux Foundations (VELUX34148 to A.-M.S.O.). Mass spectrometry and proteomics data were acquired by the University of Arizona Analytical and Biological Mass Spectrometry Facility supported by NIH/NCI grant CA023074 to the University of Arizona Cancer Center, UA Research Development and Innovation Office, and the BIO5 Institute of the University of Arizona.

The authors declare no competing interests. Received: August 19, 2019 Revised: October 14, 2019 Accepted: November 8, 2019 Published: November 22, 2019 REFERENCES Andon, N.L., Hollingworth, S., Koller, A., Greenland, A.J., Yates, J.R., 3rd, and Haynes, P.A. (2002). Proteomic characterization of wheat amyloplasts using identification of proteins by tandem mass spectrometry. Proteomics 2, 1156–1168.

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AUTHOR CONTRIBUTIONS

Moellering, R.E., and Cravatt, B.F. (2013). Functional lysine modification by an intrinsically reactive primary glycolytic metabolite. Science 341, 549–553.

Conceptualization and methodology, J.J.G. and J.R.R.; Investigation, D.O.G., E.Q.J., C.C.A., and J.O.M.; Writing, J.J.G.; Resources, E.C., T.S., A.-M.S.O., M.J., M.D.S., and D.A.S.; Funding Acquisition, J.J.G., J.R.R., E.C., and D.A.S.; Supervision, J.J.G., J.R.R., E.C., and D.A.S.

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Wagner, G.R., and Payne, R.M. (2013). Widespread and enzyme-independent Nepsilon-acetylation and Nepsilon-succinylation of proteins in the chemical conditions of the mitochondrial matrix. J. Biol. Chem. 288, 29036–29045.

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Zhang, D., Tang, Z., Huang, H., Zhou, G., Cui, C., Weng, Y., Liu, W., Kim, S., Lee, S., Perez-Neut, M., et al. (2019). Metabolic regulation of gene expression by histone lactylation. Nature 574, 575–580.

Rabbani, N., Xue, M., and Thornalley, P.J. (2016b). Methylglyoxal-induced dicarbonyl stress in aging and disease: first steps towards glyoxalase 1-based treatments. Clin. Sci. (Lond.) 130, 1677–1696. Sibbersen, C., Palmfeldt, J., Hansen, J., Gregersen, N., Jorgensen, K.A., and Johannsen, M. (2013). Development of a chemical probe for identifying protein targets of alpha-oxoaldehydes. Chem. Commun. (Camb.) 49, 4012–4014. Sibbersen, C., Schou Oxvig, A.M., Bisgaard Olesen, S., Nielsen, C.B., Galligan, J.J., Jorgensen, K.A., Palmfeldt, J., and Johannsen, M. (2018). Profiling of methylglyoxal blood metabolism and advanced glycation endproduct proteome using a chemical probe. ACS Chem. Biol. 13, 3294–3305.

8 Cell Chemical Biology 27, 1–8, January 16, 2020

Zhao, S., Xu, W., Jiang, W., Yu, W., Lin, Y., Zhang, T., Yao, J., Zhou, L., Zeng, Y., Li, H., et al. (2010). Regulation of cellular metabolism by protein lysine acetylation. Science 327, 1000–1004. Zheng, Q., Omans, N.D., Leicher, R., Osunsade, A., Agustinus, A.S., FinkinGroner, E., D’Ambrosio, H., Liu, B., Chandarlapaty, S., Liu, S., et al. (2019a). Reversible histone glycation is associated with disease-related changes in chromatin architecture. Nat. Commun. 10, 1289. Zheng, Q., Prescott, N.A., Maksimovic, I., and David, Y. (2019b). (De)Toxifying the epigenetic code. Chem. Res. Toxicol. 32, 796–807.

Please cite this article in press as: Gaffney et al., Non-enzymatic Lysine Lactoylation of Glycolytic Enzymes, Cell Chemical Biology (2019), https:// doi.org/10.1016/j.chembiol.2019.11.005

STAR+METHODS KEY RESOURCES TABLE

REAGENT or RESOURCE

SOURCE

IDENTIFIER

Antibodies Mouse monoclonal anti-Lactoylglutathione lyase

MilliporeSigma

Cat# 05-1925; RRID: AB_10563505

Rabbit polyclonal HAGH

ThermoFisher

Cat# PA5-28292; RRID: AB_2545768

Rabbit monoclonal HK1

Cell Signaling Technologies

Cat# 2024; RRID: AB_2116996

Rabbit monoclonal LDHA

Cell Signaling Technologies

Cat# 3582; RRID: AB_2066887

Rabbit monoclonal PKM1/2

Cell Signaling Technologies

Cat# 3190; RRID: AB_2163695

Rabbit monoclonal ALDOA

Cell Signaling Technologies

Cat# 8060; RRID: AB_2797635

Chemicals, Peptides, and Recombinant Proteins Human recombinant PGK1

BioVision

Cat# 7819

alkMethylglyoxal

Sibbersen et al., 2013

N/A

Methylglyoxal

MP Biomedicals

Cat# 02155558-CF; CAS: 78-98-8

Histone H4

New England Biolabs

Cat# M2504S

GSH-(glycine-13C2,15N, internal standard)

MilliporeSigma

Cat# 683620

(1-Carboxyethyl)-L-lysine-d4

Toronto Research Chemicals

Cat# C178072

ftp://ftp.thegpm.org/fasta/cRAP

N/A

Galligan et al., 2018

N/A

Vanderbilt University

Cat# 5656552, RRID:MGI:5656552

GLO2 gRNA targeting sequence: TACGGGGGTGACGACCGTAT

This paper

N/A

Primer: GLO2 Forward: CCACTGCACCAGGACAAGAAATCCACC

This paper

N/A

Primer: GLO2 Reverse: GGCTGAAGACACCCTCGCAGGG

This paper

N/A

Cong and Zhang, 2015

N/A

Mendeley Data

https://doi.org/10.17632/9ccms3gnf6.2

Xcalibur v 4.0.27.19

Andon et al., 2002

https://www.thermofisher.com/order/ catalog/product/OPTON-30965

Scaffold Q+S v 4.8.7

Proteome Software Inc., Portland OR

http://www.proteomesoftware.com/ products/scaffold/

Deposited Data homo sapiens protein database, SwissProt (Nov 5 2018, 42252 sequences) Experimental Models: Cell Lines HEK293 GLO1-/Experimental Models: Organisms/Strains Mouse: C57BL/6J Oligonucleotides

Recombinant DNA Plasmid: pSpCas9(BB)-2A-Puro Deposited Data Proteomics data set of lactoylated proteins Software and Algorithms

Other Kintetix C8 column

Phenomenex

50 x 2.1mm, 3mm particle diameter Ascentis C18 column

Supelco

150 x 2.1mm, 3.5 mm particle diameter Eclipse XDB-C8 column

Agilent, Santa Clara, CA

Cat# 930990-906

2.1 mm x 100 mm, 3.5 mm particle diameter XBridge Amide column

Waters, Milford, MA

Cat# 186004860

Acclaim Pepmap 100 trap column, 75 micron ID x 25 cm

ThermoScientific

Cat# 164569

Cat# SU581300-U

LEAD CONTACT AND MATERIALS AVAILABILITY Further information and requests for resources and reagents should be direct to and will be fulfilled by the Lead Contact, James Galligan ([email protected]). All unique reagents generated in this study are available from the Lead Contact with a completed Materials Transfer Agreement. Cell Chemical Biology 27, 1–8.e1–e6, January 16, 2020 e1

Please cite this article in press as: Gaffney et al., Non-enzymatic Lysine Lactoylation of Glycolytic Enzymes, Cell Chemical Biology (2019), https:// doi.org/10.1016/j.chembiol.2019.11.005

EXPERIMENTAL MODEL AND SUBJECT DETAILS Mouse Tissues Male C57BL/6 mice (8 weeks) were housed in temperature and humidity-controlled specific pathogen-free conditions with 14-hour light/10-hour dark cycle. Mice were fed a standard chow diet (NIH-31, Envigo) consisting of 6.2% fat-derived calories, 45% carbohydrate-derived calories and 18% crude protein. Mice were fed ad libitum for 12 weeks had free access to water. Non-fasted mice were anesthetized via isoflurane and sacrificed via exsanguination and cervical dislocation. Tissues were excised and immediately frozen in liquid nitrogen. All animal use was approved by the Animal Care and Use Committee of the University of Arizona and adhered to the NIH Guide for the Care and Use of Laboratory Animals. Cell Culture HEK293 cells were cultured in low-glucose DMEM supplemented with 10% FBS. Cells were incubated at 37 C under 5% CO2. Following treatments, cells washed and scraped into ice-cold PBS. Cells were pelleted via centrifugation at 1000 x g for analysis as described. METHOD DETAILS Reagents All reagents were purchased from ThermoFisher Scientific (Waltham, MA) unless otherwise stated. Fetal bovine serum (FBS) was purchased from Atlas Biologicals (Ft. Collins, CO). Purified histones were purchased from New England Biolabs (Ipswich, MA). Recombinant PGK1 was purchased from BioVision (Milpitas, CA). Methylglyoxal was purchased from MP Biomedicals (Solon, OH). alkMGO was synthesized as described (Sibbersen et al., 2013). In Vitro Treatment of Recombinant Proteins with MGO or LGSH Recombinant histone H4 (2 mg) or PGK1 (2 mg) was incubated with either 1 mM LGSH or 1 mM MGO for 24 h at 37 C in a final volume of 20 mL PBS (pH 7.4). Following treatments, protein was precipitated with 300 mL ACN and placed at -20 C for 1 h. Samples were centrifuged at 14,000 x g for 10 min and the supernatant was removed. Precipitated proteins then digested for QuARKMod analysis (see below) or resolved via SDS-PAGE and stained with coomassie for proteomics analysis (see below). CRISPR-Cas9-Mediated GLO1, GLO2, and GLO1/2 Knockout (-/-) HEK293 Cells gRNA oligonucleotides were designed to target restriction enzyme recognition sites in the initial exons of the GLO1 or GLO2 locus and ligated into the pSpCas9(BB)-2A-Puro plasmid according to Cong et al. (Cong and Zhang, 2015). GLO1-/- cells were generated as described (Galligan et al., 2018); TACGGGGGTGACGACCGTAT was inserted into the plasmid to target GLO2. To generate GLO2-/-, 2 x 105 HEK293 cells were plated in 2 mL DMEM supplemented with 10% FBS in 6-well plates. The following day, 5 mg of each construct was combined with 10 mL lipofectamine 2000 (Life Technologies, Carlsbad, CA) reagent in 1 mL Opti-MEM and incubated at room temperature for 30 min. The DMEM was replaced with the plasmid-lipofectamine solution, and the cells were incubated at 37 C for 24 h. The medium was then replaced, and cells were allowed to recover for 24 h at 37 C. The medium was then replaced with serum-containing DMEM with 0.75 mg/mL puromycin, and the cells were incubated at 37 C for 48 h. The medium was then replaced with puromycin-free medium, and cells were incubated for 24 h before the medium was replaced to remove any traces of puromycin. Cultures were then pelleted and resuspended in sorting buffer (PBS + 4% FBS), and strained. Solutions were sorted by flow cytometry using a BD FACSAria III cell sorter to isolate single cell cultures in 3 x 96-well plates for each cell line, yielding approximately 100 viable clones. To generate GLO1/2-/- cells, the outlined transfection procedure was performed using the GLO2 gRNA in GLO1-/- cells, rather than WT cells. Flow cytometry experiments were performed in the Vanderbilt University Medical Center Flow Cytometry Shared Resource. Clones were maintained to 80% confluency and passaged until enough cells could be harvested for indel analysis. Restriction-fragment length polymorphism assay was employed to assess for indel mutations; target genes were PCR-amplified and subjected to digestion by restriction enzymes specific to the WT sequence at the gRNA target site. The absence of restriction enzyme activity was indicative of mutations in both alleles at the gRNA target site and clones with homozygous mutations of GLO1 or GLO2 were validated as genetic knockouts. PCR products from RFLP assays were purified using the Nucleospin PCR Clean-Up Kit and sequenced with the forward (5’-CCACTGCACCAGGACAAGAAATCCACC-3’) and reverse (3’-GGCTGAAGACACCCTCGCAGGG-5’) primers. Sequencing samples were analyzed by GenHunter Corporation. SDS-PAGE and Immunoblotting Samples were denatured in SDS loading buffer and heated at 95 C for 5 minutes. Proteins were then resolved via SDS-PAGE and transferred to PVDF membranes (GE Healthcare, Piscataway, NJ). Membranes were blocked with Odyssey Blotting Buffer (Li-Cor Biosciences, Lincoln, NE) for 45 minutes at room temperature. Primary antibodies were incubated with membranes overnight at 4 C as described: GLO1 (1:2000, MilliporeSigma, 05-1925), GLO2 (1:1000, ThermoFisher, PA5-28292), HK-1 (1:2000, Cell Signaling Technologies, #2024), LDHA (1:2000, Cell Signaling Technologies, #3582), PKM1/2 (1:2000, Cell Signaling Technologies, #3190),

e2 Cell Chemical Biology 27, 1–8.e1–e6, January 16, 2020

Please cite this article in press as: Gaffney et al., Non-enzymatic Lysine Lactoylation of Glycolytic Enzymes, Cell Chemical Biology (2019), https:// doi.org/10.1016/j.chembiol.2019.11.005

ALDOA (1:2000, Cell Signaling Technologies, #8060). Following 3x washes with TBS +0.1% Tween-20, infrared secondary antibodies (Li-COR) were added in blocking buffer (1:5000) for 45 minutes. Blots were developed following 3 additional washes with TBST using a c600 Azure Imaging System (Azure Biosystems, Dublin, CA). Reactivity-Based Protein Profiling 3 x 106 cells were plated in 100 mm plates and allowed to seed overnight. Cells were treated with either 50 mM alkMGO of vehicle control for 6 h (Sibbersen et al., 2018). Following treatments, media was removed and cells were washed once with ice-cold PBS, scraped, and pelleted at 250 x g. Cell pellets were sonicated in PBS containing an EDTA-free protease inhibitor tablet (Pierce, San Jose, CA). Insoluble debris was removed via centrifugation at 16,000 x g for 10 min at 4 C. Soluble protein (25 mg) was diluted to a volume of 27 mL in PBS and click chemistry was performed via the addition of 85 mM TBTA (final concentration, TCI America, Portland, OR), 1 mM CuSO4, 1 mM TCEP, and 10 mM N3-IR (Li-COR Biosciences, Lincoln, NE). Samples were incubated in the dark at room temperature for 90 min prior to the addition of 4x loading buffer. Proteins were separated on a 12% SDS-PAGE gel and visualized on an Azure c600 Imaging System. Equal loading was monitored via Coomassie blue staining (Thermo Scientific). Quantification of Cellular MGO MGO was quantified as previously described (Galligan et al., 2018). Briefly, 0.4 x 106 cells were plated in 6-well plates and treated as described. Following treatments, media was removed and the cells were washed once with ice-cold PBS and scraped into 1 mL cold PBS. Cells were pelleted at 1,000 x g for 5 min at 4 C. Cell pellets were immediately extracted using 100 mL of methanol (-80 C) containing 50 pmol 13C3-MGO and placed at -80 C for 1 h. Protein was pelleted via centrifugation at 14,000 x g for 10 min at 4 C and the supernatant was derivatized with 1 mM o-phenylenediamine (final) in PBS (150 mL final volume) for 2 h in the dark with end-over-end rotation. Samples were centrifuged at 14,000 x g for 10 min and the supernatant (20 mL) was chromatographed using a Shimadzu LC system equipped with a 50 x 2.1mm, 3mm particle diameter Atlantis C18 column (Waters, Milford, MA) at a flow rate of 0.350 mL/min. Buffer A (0.1% formic acid in water) was held at 99% for 0.5 min then a linear gradient to 98% solvent B (0.1% formic acid in acetonitrile) was applied over the next 4 min. The column was held at 98% B for 2 min and then washed at 1% A for 2 min and equilibrated to 99% A for 2 min. Multiple reaction monitoring (MRM) was conducted in positive mode using an AB SCIEX 4500 QTRAP with the following transitions: m/z 145.1/77.1 (analyte); m/z 148.1/77.1 (13C3-MGO, internal standard). Quantification of GSH and LGSH 3 x 105 cells were plated in 6-well plates. Following treatments, cells were washed once with ice-cold PBS and scraped into 1.0 mL. Cells were pelleted via centrifugation at 250 x g and pellets were sonicated in 80 mL of a solution containing 20 mM ethyliodoacetate and 2.5 nmol of GSH-(glycine-13C2,15N, internal standard) in 50 mM PBS, pH 7.4. Samples were derivatized for 45 min at room temperature protected from light. Protein was precipitated via addition of 10 mL of 20% (w/v) 5-sulfosalicylic acid (2% final) and removed via centrifugation at 10,000 x g for 5 min at room temperature. Supernatants were removed 10 mL of 200 mM heptafluorobutyric acid (HFBA) was added to each sample. Clarified supernatant (20 mL) was chromatographed using a Shimadzu LC system equipped with a 50 x 2.1mm, 3mm particle diameter Atlantis C18 column (Waters, Milford, MA) at a flow rate of 0.35 mL/min. Solvent A (10 mM HFBA in H2O) was held at 96% for 0.5 min, then a linear gradient to 95% B (10 mM ACN) was applied over the next 4.5 min. The column was held at 95% B for 0.5 min and then equilibrated to 96% A for 2 min. The needle was washed prior to each injection with a buffer consisting of 25 mM NH4OAc in MeOH. Multiple reaction monitoring was performed in positive ion mode using an AB SCIEX 4500 QTRAP with the following transitions: m/z 394.2 / 265.2 for GSH; m/z 380.1 / 233.1 for LGSH; 397.2 / 268.2 for GSH-(glycine-13C2,15N). GSH and LGSH were quantified using GSH-(glycine-13C2,15N). Synthesis of LactoylLys

To a 50 mL round bottom flask with a magnetic stir bar, was added 1 (1 mmol, 118 mg), 2 (0.2 mmol,50mg) and 10 mL ACN, the solution was heated up to 80 C and reacted until the 2 disappeared. 10 mL NH4Cl (aq) was added to quench the reaction and the resulting mixture was extracted with EtOAc for 3 times (15 mL). The combined EtOAc phase was washed with brine (30 mL), and then dried with Na2SO4. An hour later, the Na2SO4 was removed by filtration. The resulting solution was concentrated to obtain crude 4 as colorless oil. 4 was used directly in the deprotection without further purification. To a 20 mL vial with a magnetic stir bar, 4 and TFA at 0 C were added. The reaction material was allowed to stir overnight, and the reaction was quenched with NaHCO3 (aq) at 0 C. The mixture was then extracted with EtOAC for 3 times (9 mL). The combined EtOAc phase was washed with brine (9 mL), and then

Cell Chemical Biology 27, 1–8.e1–e6, January 16, 2020 e3

Please cite this article in press as: Gaffney et al., Non-enzymatic Lysine Lactoylation of Glycolytic Enzymes, Cell Chemical Biology (2019), https:// doi.org/10.1016/j.chembiol.2019.11.005

dried with Na2SO4. An hour later, the Na2SO4 was removed by filtration. The resulting solution was concentrated to obtain crude 5 as colorless oil. The resulting MS/MS is shown.

QuARKMod Analysis for PTM Quantitation Lys and Arg PTMs were quantified from 100 mg of cell lysate as described (Galligan et al., 2017). 0.4 x 106 cells were plated in 6-well plates and treated as described. Following treatments, media was removed and the cells were washed once with ice-cold PBS and scraped into 1 mL cold PBS. Cells were pelleted at 1,000 x g for 5 min at 4 C. Cell pellets were lysed in 100 mL of a buffer containing 150 mM NaCl, 50 mM HEPES, and 1% IGEPAL, pH 7.4. Samples were sonicated and insoluble debris was removed via centrifugation at 14,000 x g for 10 min at 4 C. Soluble protein concentration was determined via BCA and 100 mg was precipitated using ice-cold ACN at -20 C for 1 h. Proteins were pelleted via centrifugation at 14,000 x g for 10 min at 4 C. 70 mL of 50 mM ammonium bicarbonate, pH 8.0 was added to each pellet followed by the addition of an internal standard mix (10 mL, see table below), and 1 mg of sequencing-grade trypsin (Promega, 10 mL); proteins were digested for 16 h at 37 C. Trypsin was denatured via heating to 95 C for 10 min and samples were allowed to cool to room temperature prior to the addition of 15 mg of aminopeptidase M (Millipore) for 16 h at 37 C. Following digestion, 5 mL of heptofluorobutyric acid was added to each sample and insoluble debris was removed via centrifugation at 14,000 x g for 10 min. 10 mL of the supernatant was then chromatographed using a Shimadzu LC system equipped with a 150 x 2.1mm, 3.5 mm particle diameter Eclipse XDB-C8 column (Agilent, Santa Clara, CA) at a flow rate of 0.425 mL/min. Mobile phase A: 0.1% HFBA in water; mobile phase B: 0.1% HFBA in ACN. The following gradient was used: 0.5 min, 5% B; 8 min, 50% B; 8.5 min, 80% B; 9 min 80% B; 9.5 min, 5% B. The column was equilibrated for 2 min at 5% B. Scheduled MRM was conducted in positive mode using an AB SCIEX 4500 QTRAP. The MRM detection window was 50 sec with a target scan time of 0.75 sec. The following parameters were used for detection:

Species

Q1 (m/z)

Q3 (m/z)

Time (min)

CE (V)

Lys

147.1

84.1

4.7

29

13

155.1

90.1

4.7

29

Arg

175.1

70.1

5.0

47

13

185.1

75.1

5.0

47

acLys

189.2

84.1

3.5

39

acLys-d8

197.2

91.1

3.5

39

me3Lys

189.2

84.1

5.0

39

13

197.2

90.1

5.0

31

Leu

132.1

86.1

5.4

17

13

139.1

93.1

5.4

17

meLys

161.1

84.1

4.9

31

me2Lys

175.1

84.1

5

31

ADMA

203.1

70.1

5.4

C615N2 Lys C615N4 Arg

C615N2 me3Lys C615N Leu

51 (Continued on next page)

e4 Cell Chemical Biology 27, 1–8.e1–e6, January 16, 2020

Please cite this article in press as: Gaffney et al., Non-enzymatic Lysine Lactoylation of Glycolytic Enzymes, Cell Chemical Biology (2019), https:// doi.org/10.1016/j.chembiol.2019.11.005

Continued Species

Q1 (m/z)

Q3 (m/z)

Time (min)

CE (V)

ADMA-d7

210.1

77.1

5.4

51

MG-H1

229.2

70.1

5.5

53

13

230.2

70.1

5.5

53

CEA

247.2

70.1

5.4

55

13

C-CEA

248.2

70.1

5.4

55

LactoylLys

219.2

84.1

3.3

41

CEL

219.2

84.1

4.7

41

CEL-d4

223.2

88.1

4.7

41

C-MG-H1

Targeted Metabolomics 1 x 106 cells were plated in 100 mm dishes and allowed to seed for 24 h. Cells were treated with either vehicle (ddH2O) or 50 mM MGO for 6 h. Culture media was removed and the cells were washed twice with warm PBS. Cells were extracted on dry ice with 3 mL of 80:20 MeOH:ddH2O chilled to -80 C. Plates were placed at -80 C for 15 min and then scraped into a 15 mL conical tube containing 100 pmol of IS (CEL-d4). The plates were washed once with an additional 3 mL chilled extraction buffer and pooled with the previous extraction. Insoluble metabolites were removed via centrifugation at 14,000 x g for 10 min. Pellets were washed once with extraction buffer (1 mL) and pooled with the previous extraction. Extracted metabolites were dried under N2 and resuspended in 200 mL of 50:50 ACN:20 mM NH4OAc in ddH2O. Metabolites were chromatographed using a Shimadzu LC system equipped with a 2.1 mm x 100 mm, 3.5 mm particle diameter XBridge Amide column (Waters, Milford, MA) at a flow rate of 400 mL/min. Buffer A (20 mM NH4OAc in ddH2O) and Buffer B (ACN) were chromatographed as follows: 0.5 min, 95% B; 12 min, 30% B; 13 min, 30% B; 14 min, 95% B; 15 min, 95% B. The column was equilibrated at 95% B for 5 min between runs. Multiple reaction monitoring was performed in negative ion mode using an AB SCIEX 4500 QTRAP with the parameters defined in the table below. All compounds were quantified against the IS (CEL-d4) and normalized to total protein. Data are presented as a Log2 fold change compared to WT vehicle control.

Species

Q1 (m/z)

CE (V)

RT

Glucose

179

Q3 (m/z) 89

-21

7.0

Glucose/Fructose 6-phosphate

259

79

-19

8.0

Fructose 1,6-Bisphosphate

339

79

-73

8.6

Glyceraldehyde 3-phosphate

169

79

-26

7.9

Dihydroxyacetone phosphate

169

79

-26

8.6

Bisphosphoglycerate

265

97

-38

8.2

2/3-bisphosphoglycerate

185

79

-38

7.8

Phosphoenolpyruvate

167

79

-25

7.8

Lactate

89

43

-16

5.5

AMP

346

134

-30

7.1

ADP

426

134

-30

7.7

ATP

506

159

-32

8.1

NAD+

662

540

-27

7.5

NADP+

742

620

-27

8.4

Citrate/Isocitrate

191

111

-16

7.9

cis-Aconitate

173

129

-16

7.9

a-Ketoglutarate

145

101

-16

6.7

Succinate

117

73

-16

7.9 7.1

Fumarate

115

71

-10

Malate

133

115

-16

7.1

Oxaloacetate

131

87

-16

8.5

GTP

522

79

-32

8.8

GDP

442

79

-32

8.1

CEL-d4

221

149

-26

7.8

Cell Chemical Biology 27, 1–8.e1–e6, January 16, 2020 e5

Please cite this article in press as: Gaffney et al., Non-enzymatic Lysine Lactoylation of Glycolytic Enzymes, Cell Chemical Biology (2019), https:// doi.org/10.1016/j.chembiol.2019.11.005

Identification of MGO Modified Proteins WT or GLO2-/- cells were cultured in DMEM Flex medium for Stable Isotope Labeling of Amino Acids in Cell Culture (SILAC) (ThermoFisher Scientific) supplemented with 1 g/L sterile filtered glucose, 2 mM glutamine, 1 mM pyruvate, and 10% dialyzed FBS (Gibco). ‘Heavy’ cells were 0.1 g/L 13C615N2 Lys and 0.1 g/L 13C615N4 Arg for ‘heavy’ cells or natural abundance isotopes for ‘light’ cells. Cells (5 x 106) were plated on 150 mm dishes and allowed to seed for 24 h. The following day, each plate was treated with 50 mM alkMGO for 6 h. Plates were washed once with ice-cold PBS and scraped into PBS; to obtain sufficient input protein, two plates were pooled for each replicated. Pellets were harvested via centrifugation and lysed via sonication in PBS containing EDTA-free protease inhibitors. Insoluble protein was removed via centrifugation at 14,000 x g for 20 min at 4 C. Protein concentrations were determined via BCA assay and 3 mg of WT ‘light’ was mixed with 3 mg of GLO2-/- ‘heavy’ and diluted to 2 mg/mL in PBS. Click chemistry was carried out with 0.2 mM N3-UV-biotin (final concentration, Kerafast, Boston, MA), 1 mM TCEP, 1 mM CuSO4, and 0.1 mM TBTA for 2 h in the dark. Protein was precipitated with ice-cold ACN for 2 h at -20 C protected from light to remove excess biotin. Protein was then resolubilized via sonication in PBS containing 1% SDS. SDS was diluted 10-fold in PBS and biotin-conjugated protein was enriched with 200 mL of a 50% streptavidin bead slurry (GE Lifesciences, Pittsburgh, PA) overnight at 4 C. Enriched protein was isolated through centrifugation and the beads were washed as follows: 1% SDS in PBS (2x), 2M urea in PBS (2x), 1M NaCl in PBS (2x), PBS (2x), and ddH2O (2x). Beads were resuspended in 300 mL of ddH2O and proteins were released from the beads through UV-irradiation (365 nm, 2 h). Eluted proteins were concentrated to dryness under vacuum and briefly resolved (1 cm) on an SDS-PAGE gel. Gel bands were excised and disulfide bonds were reduced and alkylated prior to digestion with trypsin/lysine C (20 ng/mL, Promega Corporation, Madison, WI) overnight at 37 C using ProteaseMax Surfactant trypsin enhancer (Promega Corporation, Madison, WI). LC-MS/MS analysis was carried out using a QExactive Plus mass spectrometer (Thermo Fisher Scientific, San Jose, CA) equipped with a nanoESI source. Peptides were eluted from an Acclaim Pepmap 100 trap column (75 micron ID x 25 cm, ThermoScientific) onto an Acclaim PepMap RSLC analytical column (75 micron ID 3 25 cm, ThermoScientific) using a 5-20% gradient of solvent B (acetonitrile, 0.1% formic acid) over 50 minutes, 20-50% solvent B over 5 minutes, 50-95% of solvent B over 3 minutes, 95% hold of solvent B for 5 minutes, and finally a return to 5% solvent B in 1 minute and another 10 minute hold of 5% solvent B. Solvent A consisted of water and 0.1% formic acid. All flow rates were 300 nL/min using a Dionex Ultimate 3000 RSLC nano System (Thermo Scientific). Data dependent scanning was performed by the Xcalibur v 4.0.27.19 software (Andon et al., 2002) using a survey scan at 70,000 resolution scanning mass/charge (m/z) 353-1550 at an automatic gain control (AGC) target of 1e5 and a maximum injection time (IT) of 65 msec, followed by higher-energy collisional dissociation (HCD) tandem mass spectrometry (MS/MS) at 27nce (normalized collision energy), of the 10 most intense ions at a resolution of 17,500, an isolation width of 1.5 m/z, an AGC of 1e5 and a maximum IT of 65 msec. Dynamic exclusion was set to place any selected m/z on an exclusion list for 20 seconds after a single MS/MS. Ions of charge state +1, 7, 8, >8 and unassigned were excluded from MS/MS, as were isotopes. Tandem mass spectra were searched against homo sapiens protein database from SwissProt (Nov 5 2018, 42252 sequences) to which additional common contaminant proteins (e.g. trypsin, keratins; obtained at ftp://ftp.thegpm.org/fasta/cRAP) were appended. All MS/MS spectra were searched using Thermo Proteome Discoverer v 2.2.0388 (ThermoFisher Scientific) considering fully tryptic peptides with up to 2 missed cleavage sites. Variable modifications considered during the search included methionine oxidation (15.995 Da), cysteine carbamidomethylation (57.021 Da), heavy Lys (+8.014) and heavy Arg (+10.008). Proteins were identified at 99% confidence with XCorr score cut-offs as determined by a reversed database search(Qian et al., 2005). The protein and peptide identification results were also visualized with Scaffold Q+S v 4.8.7 (Proteome Software Inc., Portland OR), a program that relies on various search engine results (i.e.: Sequest, X! Tandem, MASCOT) and which uses Bayesian statistics to reliably identify more spectra (Keller et al., 2002). Protein identifications were accepted that passed a minimum of two peptides identified at 0.1% peptide False Discovery Rate and 90-99.9% protein confidence by the Protein Profit algorithm within Scaffold. SILAC quantification was performed using Proteome Discoverer (ThermoFisher) and only proteins with reported H:L ratios in both replicates were considered positive identifications. QUANTIFICATION AND STATISTICAL ANALYSIS Data were quantified using a two-way ANOVA using a Tukey post-hoc test. Differences were considered to be significant when P < 0.05 with the N indicated in each figure legend. All analyses were carried out using Prism 8 for Macintosh, GraphPad Software. DATA AND CODE AVAILABILITY The proteomics inventory generated during this study is available on Mendeley, https://doi.org/10.17632/9ccms3gnf6.2.

e6 Cell Chemical Biology 27, 1–8.e1–e6, January 16, 2020