Metabolic Enzymes in Saccharomyces cerevisiae: History
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Subjects: Biochemistry & Molecular Biology
Contributor:
Natsuko Miura

Condensate formation by a group of metabolic enzymes in the cell is an efficient way of regulating cell metabolism through the formation of “membrane-less organelles.” Because of the use of green fluorescent protein (GFP) for investigating protein localization, various enzymes were found to form condensates or filaments in living Saccharomyces cerevisiae, mammalian cells, and in other organisms, thereby regulating cell metabolism in the certain status of the cells. 

  • Saccharomyces cerevisiae
  • liquid–liquid phase separation
  • metabolic enzymes

1. Introduction

The intracellular distribution of proteins in cells is strongly related to their function. Protein localization has been extensively studied using budding yeast cells since the development of the GFP-fused protein library in budding yeast cells [1]. First reported in 2003, the library of yeast cells of which fused C-terminus of protein-coding sequences in the genome with GFP (G65T)-coding sequence [2] covers about 75% (4159) of genome-coded proteins in Saccharomyces cerevisiae cells (for more details, a haploid strain BY4741 (ATCC 201388), now commercially available). As the localization of most proteins in the library have been recorded and can now be searched in a database (Yeast GFP Fusion Localization Database; https://yeastgfp.yeastgenome.org/ accessed on 13 December 2021), research on protein localization entered a new phase; many proteins exist in locations that vary from those predicted from the amino acid sequence. This is not surprising because proteins, especially metabolic enzymes, exert different functions according to time and occasion, which are called “moonlighting proteins” [3][4]. However, the mechanisms for regulating the intracellular localization of moonlighting proteins are largely unknown. The differential protein localization in cells was sometimes observed depending on the cellular growth [5][6][7][8] or under environmental stress, including nutrient depletion [8][9] and hypoxia [10][11]. There is still a mystery about the biological meanings of spatial reorganization of metabolic enzymes in cells. Some reports have shown the effect of protein condensates to control cellular metabolism [6][10][11]. Here, the recent progress in the protein condensates formed by metabolic enzymes, biological roles of the condensate, and its regulation, mainly focusing on hypoxia-induced protein condensates, were overviewed.
It has now been suggested that metabolic enzymes produce multiple condensates in the cell, and some of these condensates associate with each other in a specific intracellular environment. The term “META body” was first coined in 2021 [12], and the acronym “META” stands for “Metabolic Enzymes Transiently Assembling”. As indicated by the term, the META body is a higher-order enzyme condensate in the cell, comprising condensates formed by glycolytic and purine biosynthesis enzymes that are formed under hypoxic conditions. 

2. Filaments Formed by a Single Metabolic Enzyme in S. cerevisiae

Before GFP clones were developed, condensate or filament formation by metabolic enzymes in cells has been forecasted by electron microscopy using purified enzymes. The filament formation by some metabolic enzymes was later confirmed in S. cerevisiae or other organisms such as Drosophila and mammalian cells [13]. For example, acetyl coenzyme A carboxylase (ACC), an enzyme involved in the fatty acid synthesis, purified from animal tissues, formed filaments in electron microscopy [14]. The filament formation of ACC (yeast ortholog, Acc1p) was later confirmed in S. cerevisiae cells under prolonged starvation [7][8]. Regulatory mechanisms of ACC filament have been extensively studied on human ACC. The filament formation by ACC is regulated through phosphorylation at serine residue, in addition to adenosine monophosphate (AMP)-activated protein kinase (AMPK) and cAMP-dependent protein kinase (PKA) [15]. Kleinshmidt reported that dissociation of the enzyme filament inactivates the enzyme, and reassembly restored catalytic activity [14]. In vitro studies have indicated that 5 mM citrate concentration induces filament formation of ACC [16]. Although the concentration seemed too high with the plasma concentration of citrate being ~150 μM [17], the intracellular citrate levels of Escherichia coli [18] and human breast cancer cells [19] may fall within the range, indicating that the filament formation can be regulated by intracellular metabolites.
Another enzyme, cytidine triphosphate (CTP) synthase, which plays an important role in polynucleotide and lipid synthesis, formed filaments in E. coli [20], S. cerevisiae [6], Drosophila [21], and human-derived HeLa cells [22]. In S. cerevisiae, CTP synthase forms filaments under starvation and depends on intracellular pH below 7.0 [23]. The filament formation inactivates CTP synthase [24][25][26] while prolonging the half-life of the enzyme [27]. CTP induces filament formation; in growing bacterial cells, excess CTP possibly induces inactive CTP synthase filaments and control CTP levels in cells [28]. In contrast to bacterial CTP synthase, the polymerization of human CTP synthase increases the enzymatic activity [29]. Furthermore, in Drosophila eggs, the polymerization of CTP synthase improves the egg production [30]. These opposite effects of polymerization of CTP synthases among biological species are considered to depend on the three-dimensional conformation of enzymes [30].
In S. cerevisiae, an increasing number of metabolic enzymes have been reported to form filaments. Glucokinase (Glk1p) forms filaments in the presence of glucose during the stationary phase, and the polymerization inhibits the enzymatic activity [31]. Glk1p is one of the three enzymes in the hexokinase family. Other members of the hexokinase family, hexokinase 1 (Hxk1p) and 2 (Hxk2p), did not form any filament [31]. Pyruvate kinase (Cdc19p), an ATP-producing enzyme in the glycolytic pathway, also forms molecular condensates or filaments in S. cerevisiae under starvation [9][32][33][34], depending on its low-complexity region [35].
Recently, phosphofructokinase (PFK), one of the ATP-consuming enzymes in the glycolytic pathway, formed filaments in breast cancer cells [36] or nerve synapses [37], with unknown mechanisms. These findings imply that the additional enzymes present in S. cerevisiae produce filaments in certain environments. The filament formation by a single metabolic enzyme is considered to be a faster technique for regulating the enzymatic activity than protein degradation or de novo protein synthesis, hence conserving cellular energy [38]. While the number of metabolic enzymes that form filaments in cells is increasing, it is not known whether filament formation by a single metabolic enzyme can trigger condensate formation by a group of enzymes that govern the metabolic pathway.

3. Condensate Formation by a Group of Enzymes in the Purine Synthesis Pathway

While some enzymes formed filaments or condensates in cells under certain conditions, it was unknown until 2008 whether a group of enzymes that govern some metabolic pathway form condensates to regulate cell metabolism. In 2008, the formation of an enzyme condensate “Purinosome” was reported in human-derived HeLa cells [39]. Purinosome is composed of multiple enzymes in the purine synthesis pathway and is formed in association with microtubules during purine depletion and the G1 phase of the cell cycle [39][40][41][42]. In yeast cells, some enzymes in the purine synthetic pathway form condensates. However, the number of enzymes observed to form condensates varies from mammalian cells, suggesting that there are various mechanisms of purinosome formation between mammalian cells and S. cerevisiae cells [5]. Among the purine synthesis pathway enzymes, those located at the branch of the metabolic pathway were found to form protein condensates in S. cerevisiae after five days of growth [5]. The effect of the condensate or filament formation by enzymes located at the branch of metabolic pathways on cell metabolism requires further investigation. Some enzymes in the purine biosynthetic pathway, including Ade4p, Ade12p, Ade17p, and Ade5,7p, were shown to form punctate foci in S. cerevisiae under nutrient-depleted conditions [9]; the optimization of cell growth conditions might enable the accumulation of all enzymes in the purine biosynthetic pathway in S. cerevisiae as well as in mammalian cells.

4. Coalesced Metabolic Enzymes under Hypoxia

In 2013, another group of metabolic enzymes governing the glycolytic pathway were found to form large-scale molecular condensates in S. cerevisiae under hypoxia [11]. The existence of the condensate in cells under hypoxia was reconfirmed in 2017, and the condensate was named “Glycolytic body” or “G-body” [10]. Moreover, similar condensate was also observed in mammalian cells, and the condensate in mammalian cells was named “G-body” [10] or “Glucosome” [50]. The number of G-body-associated proteins identified through pull-down assay followed by liquid chromatography and mass spectrometry (LC-MS) was more than 100, and microscopic observations using GFP clones showed that most glycolytic enzymes coalesce under hypoxia in a similar localization in the cell [10][11]. While the machinery of G-body formation is still unclear, it was suggested that enzymes that form molecular condensate under hypoxia coalesce in order [12], suggesting the existence of coordinated regulatory mechanisms of the condensate formation. Not only glycolytic enzymes, but also Ade57p—a member of purinosome—were specifically immunoprecipitated with condensate-forming enolase [11], suggesting that enzymes of other metabolic pathways, or other types of condensates, are also involved in the condensate formed by glycolytic enzymes. Indeed, it was later reported that enzymes of the purine synthetic pathway that forms purinosome also form molecular condensate in cells under hypoxia [51]. However, the effect of synthetic purine enzymes assembled under hypoxia is unspecified. The large-scale protein assembly in cells formed under hypoxia, “META body” [12], is now presenting novel cellular machinery that potentially newly regulates cell metabolism. 

This entry is adapted from the peer-reviewed paper 10.3390/microorganisms10020232

References

  1. Huh, W.K.; Falvo, J.V.; Gerke, L.C.; Carroll, A.S.; Howson, R.W.; Weissman, J.S.; O’Shea, E.K. Global analysis of protein localization in budding yeast. Nature 2003, 425, 686–691.
  2. Tsien, R.Y. The green fluorescent protein. Annu. Rev. Biochem. 1998, 67, 509–544.
  3. Jeffery, C.J. Moonlighting proteins. Trends Biochem. Sci. 1999, 24, 8–11.
  4. Curtis, N.J.; Jeffery, C.J. The expanding world of metabolic enzymes moonlighting as rna binding proteins. Biochem. Soc. Trans. 2021, 49, 1099–1108.
  5. Noree, C.; Begovich, K.; Samilo, D.; Broyer, R.; Monfort, E.; Wilhelm, J.E. A quantitative screen for metabolic enzyme structures reveals patterns of assembly across the yeast metabolic network. Mol. Biol. Cell 2019, 30, 2721–2736.
  6. Noree, C.; Sato, B.K.; Broyer, R.M.; Wilhelm, J.E. Identification of novel filament-forming proteins in Saccharomyces cerevisiae and Drosophila melanogaster. J. Cell Biol. 2010, 190, 541–551.
  7. Shen, Q.J.; Kassim, H.; Huang, Y.; Li, H.; Zhang, J.; Li, G.; Wang, P.Y.; Yan, J.; Ye, F.; Liu, J.L. Filamentation of metabolic enzymes in Saccharomyces cerevisiae. J. Genet. Genom. 2016, 43, 393–404.
  8. Suresh, H.G.; da Silveira Dos Santos, A.X.; Kukulski, W.; Tyedmers, J.; Riezman, H.; Bukau, B.; Mogk, A. Prolonged starvation drives reversible sequestration of lipid biosynthetic enzymes and organelle reorganization in Saccharomyces cerevisiae. Mol. Biol. Cell 2015, 26, 1601–1615.
  9. Narayanaswamy, R.; Levy, M.; Tsechansky, M.; Stovall, G.M.; O’Connell, J.D.; Mirrielees, J.; Ellington, A.D.; Marcotte, E.M. Widespread reorganization of metabolic enzymes into reversible assemblies upon nutrient starvation. Proc. Natl. Acad. Sci. USA 2009, 106, 10147–10152.
  10. Jin, M.; Fuller, G.G.; Han, T.; Yao, Y.; Alessi, A.F.; Freeberg, M.A.; Roach, N.P.; Moresco, J.J.; Karnovsky, A.; Baba, M.; et al. Glycolytic enzymes coalesce in g bodies under hypoxic stress. Cell Rep. 2017, 20, 895–908.
  11. Miura, N.; Shinohara, M.; Tatsukami, Y.; Sato, Y.; Morisaka, H.; Kuroda, K.; Ueda, M. Spatial reorganization of Saccharomyces cerevisiae enolase to alter carbon metabolism under hypoxia. Eukaryot. Cell 2013, 12, 1106–1119.
  12. Yoshimura, Y.; Hirayama, R.; Miura, N.; Utsumi, R.; Kuroda, K.; Ueda, M.; Kataoka, M. Small-scale hypoxic cultures for monitoring the spatial reorganization of glycolytic enzymes in Saccharomyces cerevisiae. Cell Biol. Int. 2021, 45, 1776–1783.
  13. Park, C.K.; Horton, N.C. Structures, functions, and mechanisms of filament forming enzymes: A renaissance of enzyme filamentation. Biophys. Rev. 2019, 11, 927–994.
  14. Kleinschmidt, A.K.; Moss, J.; Lane, D.M. Acetyl coenzyme a carboxylase: Filamentous nature of the animal enzymes. Science 1969, 166, 1276–1278.
  15. Hunkeler, M.; Hagmann, A.; Stuttfeld, E.; Chami, M.; Guri, Y.; Stahlberg, H.; Maier, T. Structural basis for regulation of human acetyl-coa carboxylase. Nature 2018, 558, 470–474.
  16. Meredith, M.J.; Lane, M.D. Acetyl-coa carboxylase. Evidence for polymeric filament to protomer transition in the intact avian liver cell. J. Biol. Chem. 1978, 253, 3381–3383.
  17. Costello, L.C.; Franklin, R.B. A review of the important central role of altered citrate metabolism during the process of stem cell differentiation. J. Regen Med. Tissue Eng. 2013, 2, 1.
  18. Bennett, B.D.; Kimball, E.H.; Gao, M.; Osterhout, R.; Van Dien, S.J.; Rabinowitz, J.D. Absolute metabolite concentrations and implied enzyme active site occupancy in Escherichia coli. Nat. Chem. Biol. 2009, 5, 593–599.
  19. Peng, M.; Yang, D.; Hou, Y.; Liu, S.; Zhao, M.; Qin, Y.; Chen, R.; Teng, Y.; Liu, M. Intracellular citrate accumulation by oxidized atm-mediated metabolism reprogramming via pfkp and cs enhances hypoxic breast cancer cell invasion and metastasis. Cell Death Dis. 2019, 10, 228.
  20. Ingerson-Mahar, M.; Briegel, A.; Werner, J.N.; Jensen, G.J.; Gitai, Z. The metabolic enzyme CTP synthase forms cytoskeletal filaments. Nat. Cell Biol. 2010, 12, 739–746.
  21. Liu, J.L. Intracellular compartmentation of CTP synthase in Drosophila. J. Genet. Genom. 2010, 37, 281–296.
  22. Carcamo, W.C.; Satoh, M.; Kasahara, H.; Terada, N.; Hamazaki, T.; Chan, J.Y.; Yao, B.; Tamayo, S.; Covini, G.; von Muhlen, C.A.; et al. Induction of cytoplasmic rods and rings structures by inhibition of the CTP and GTP synthetic pathway in mammalian cells. PLoS ONE 2011, 6, e29690.
  23. Hansen, J.M.; Horowitz, A.; Lynch, E.M.; Farrell, D.P.; Quispe, J.; DiMaio, F.; Kollman, J.M. Cryo-em structures of CTP synthase filaments reveal mechanism of ph-sensitive assembly during budding yeast starvation. Elife 2021, 10I, e73368.
  24. Aughey, G.N.; Grice, S.J.; Shen, Q.J.; Xu, Y.; Chang, C.C.; Azzam, G.; Wang, P.Y.; Freeman-Mills, L.; Pai, L.M.; Sung, L.Y.; et al. Nucleotide synthesis is regulated by cytoophidium formation during neurodevelopment and adaptive metabolism. Biol. Open 2014, 3, 1045–1056.
  25. Barry, R.M.; Bitbol, A.F.; Lorestani, A.; Charles, E.J.; Habrian, C.H.; Hansen, J.M.; Li, H.J.; Baldwin, E.P.; Wingreen, N.S.; Kollman, J.M.; et al. Large-scale filament formation inhibits the activity of CTP synthetase. Elife 2014, 3, e03638.
  26. Noree, C.; Monfort, E.; Shiau, A.K.; Wilhelm, J.E. Common regulatory control of CTP synthase enzyme activity and filament formation. Mol. Biol. Cell 2014, 25, 2282–2290.
  27. Sun, Z.; Liu, J.L. Forming cytoophidia prolongs the half-life of CTP synthase. Cell Discov. 2019, 5, 32.
  28. Aughey, G.N.; Liu, J.L. Metabolic regulation via enzyme filamentation. Crit. Rev. Biochem. Mol. Biol. 2015, 51, 282–293.
  29. Lynch, E.M.; Hicks, D.R.; Shepherd, M.; Endrizzi, J.A.; Maker, A.; Hansen, J.M.; Barry, R.M.; Gitai, Z.; Baldwin, E.P.; Kollman, J.M. Human CTP synthase filament structure reveals the active enzyme conformation. Nat. Struct. Mol. Biol. 2017, 24, 507–514.
  30. Simonet, J.C.; Foster, M.J.; Lynch, E.M.; Kollman, J.M.; Nicholas, E.; O’Reilly, A.M.; Peterson, J.R. CTP synthase polymerization in germline cells of the developing Drosophila egg supports egg production. Biol. Open 2020, 9, bio050328.
  31. Stoddard, P.R.; Lynch, E.M.; Farrell, D.P.; Dosey, A.M.; Di Maio, F.; Williams, T.A.; Kollman, J.M.; Murray, A.W.; Garner, E.C. Polymerization in the actin atpase clan regulates hexokinase activity in yeast. Science 2020, 367, 1039–1042.
  32. Cereghetti, G.; Saad, S.; Dechant, R.; Peter, M. Reversible, functional amyloids: Towards an understanding of their regulation in yeast and humans. Cell Cycle 2018, 17, 1545–1558.
  33. Cereghetti, G.; Wilson-Zbinden, C.; Kissling, V.M.; Diether, M.; Arm, A.; Yoo, H.; Piazza, I.; Saad, S.; Picotti, P.; Drummond, D.A.; et al. Reversible amyloids of pyruvate kinase couple cell metabolism and stress granule disassembly. Nat. Cell Biol. 2021, 23, 1085–1094.
  34. Saad, S.; Cereghetti, G.; Feng, Y.; Picotti, P.; Peter, M.; Dechant, R. Reversible protein aggregation is a protective mechanism to ensure cell cycle restart after stress. Nat. Cell Biol. 2017, 19, 1202–1213.
  35. Grignaschi, E.; Cereghetti, G.; Grigolato, F.; Kopp, M.R.G.; Caimi, S.; Faltova, L.; Saad, S.; Peter, M.; Arosio, P. A hydrophobic low-complexity region regulates aggregation of the yeast pyruvate kinase Cdc19 into amyloid-like aggregates In vitro. J. Biol. Chem. 2018, 293, 11424–11432.
  36. Webb, B.A.; Dosey, A.M.; Wittmann, T.; Kollman, J.M.; Barber, D.L. The glycolytic enzyme phosphofructokinase-1 assembles into filaments. J. Cell Biol. 2017, 216, 2305–2313.
  37. Jang, S.; Nelson, J.C.; Bend, E.G.; Rodriguez-Laureano, L.; Tueros, F.G.; Cartagenova, L.; Underwood, K.; Jorgensen, E.M.; Colon-Ramos, D.A. Glycolytic enzymes localize to synapses under energy stress to support synaptic function. Neuron 2016, 90, 278–291.
  38. Gomes, E.; Shorter, J. The molecular language of membraneless organelles. J. Biol. Chem. 2019, 294, 7115–7127.
  39. An, S.; Kumar, R.; Sheets, E.D.; Benkovic, S.J. Reversible compartmentalization of de novo purine biosynthetic complexes in living cells. Science 2008, 320, 103–106.
  40. An, S.; Deng, Y.; Tomsho, J.W.; Kyoung, M.; Benkovic, S.J. Microtubule-assisted mechanism for functional metabolic macromolecular complex formation. Proc. Natl. Acad. Sci. USA 2010, 107, 12872–12876.
  41. Chan, C.Y.; Zhao, H.; Pugh, R.J.; Pedley, A.M.; French, J.; Jones, S.A.; Zhuang, X.; Jinnah, H.; Huang, T.J.; Benkovic, S.J. Purinosome formation as a function of the cell cycle. Proc. Natl. Acad. Sci. USA 2015, 112, 1368–1373.
  42. Pedley, A.M.; Benkovic, S.J. A new view into the regulation of purine metabolism: The purinosome. Trends Biochem. Sci. 2017, 42, 141–154.
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