Effect of posttranslational modifications on enzyme function and assembly

JPROT-01368; No of Pages 30 JOURNAL OF P ROTEOM IC S XX ( 2013) X XX–X XX

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Review

Effect of posttranslational modifications on enzyme function and assembly☆ Helena Ryšlaváa,⁎, Veronika Doubnerováa , Daniel Kavana, b , Ondřej Vaněka a

Department of Biochemistry, Faculty of Science, Charles University in Prague, Hlavova 8, CZ-12840 Prague 2, Czech Republic Institute of Microbiology, Academy of Sciences of the Czech Republic, Vídeňská 1083, CZ-14220 Prague 4, Czech Republic

b

AR TIC LE I N FO

ABS TR ACT

Article history:

The detailed examination of enzyme molecules by mass spectrometry and other techniques

Received 29 December 2012

continues to identify hundreds of distinct PTMs. Recently, global analyses of enzymes using

Accepted 11 March 2013

methods of contemporary proteomics revealed widespread distribution of PTMs on many key enzymes distributed in all cellular compartments. Critically, patterns of multiple enzymatic

Keywords:

and nonenzymatic PTMs within a single enzyme are now functionally evaluated providing a

Enzyme

holistic picture of a macromolecule interacting with low molecular mass compounds, some of

Posttranslational modification

them being substrates, enzyme regulators, or activated precursors for enzymatic and

Structure

nonenzymatic PTMs. Multiple PTMs within a single enzyme molecule and their mutual

Catalytic activity

interplays are critical for the regulation of catalytic activity. Full understanding of this

Cellular localization

regulation will require detailed structural investigation of enzymes, their structural analogs,

Stability

and their complexes. Further, proteomics is now integrated with molecular genetics, transcriptomics, and other areas leading to systems biology strategies. These allow the functional interrogation of complex enzymatic networks in their natural environment. In the future, one might envisage the use of robust high throughput analytical techniques that will be able to detect multiple PTMs on a global scale of individual proteomes from a number of carefully selected cells and cellular compartments. This article is part of a Special Issue entitled: Protein Modifications. © 2013 Elsevier B.V. All rights reserved.

Abbreviations1 Standard IUPAC abbreviations are used for amino acids, nucleic acids, sugars, and coenzymes.: ABRF, Association for Biomolecular Resource Facilities; AGE, advanced glycosylation endproduct; ALE, advanced lipooxidation endproduct; AML, acute myeloid leukemia; APC/C, anaphase-promoting complex/cyclosome; CCT, chaperone containing TCP-1; CDK, cyclin-dependent kinase; CHO, Chinese hamster ovary; COS-1, CV-1 in Origin carrying SV40 genetic material (cell line); CSC, cell surface capture technology; EC, enzyme commission of IUPAC; ECD, electron capture dissociation; EGF, epidermal growth factor; ERAD, ER-associated protein degradation; GFP, green fluorescent protein; HECT, homologous to the E6-AP carboxyl terminus; HEK, human embryonic kidney; IP3, inositoltrisphosphate; MDM, murine double minute; MMP, matrix metalloproteinase; MRM, multiple reaction monitoring; RAGE, receptor for advanced glycosylation end products; RING, really interesting new gene; RNS, reactive nitrogen species; SAH, S-adenosyl-L-homocysteine; SAM, S-adenosyl-L-methionine; SIL, stable isotope labeling; TAS, tagging via substrate approach; TCP-1, tailless complex polypeptide-1 ☆ This article is part of a Special Issue entitled: Protein Modifications. ⁎ Corresponding author. Tel.: + 420 2 2195 1282; fax: + 420 2 2195 1283. E-mail address: [email protected] (H. Ryšlavá). 1

1874-3919/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.jprot.2013.03.025

Please cite this article as: Ryšlavá H, et al, Effect of posttranslational modifications on enzyme function and assembly, J Prot (2013), http://dx.doi.org/10.1016/j.jprot.2013.03.025

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Contents 1. 2.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Posttranslational modifications of enzymes followed by proteomic techniques . . . . . . . . 2.1. Most common posttranslational modifications of enzymes . . . . . . . . . . . . . . . 2.1.1. Glycosylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.2. Phosphorylation and O-GlcNAc modification . . . . . . . . . . . . . . . . . . . 2.1.3. Ubiquitylation and related modifications . . . . . . . . . . . . . . . . . . . . . 2.1.4. Alkylation, acylation, and prenylation . . . . . . . . . . . . . . . . . . . . . . . 2.1.5. Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.6. Glycation, lipooxidation, and modification by reactive carbonyl compounds . . 2.1.7. Other posttranslational modifications . . . . . . . . . . . . . . . . . . . . . . . 2.2. Proteomic methods for global detection of posttranslational modifications of enzymes 2.3. Compartmentalization of enzyme modifications and function . . . . . . . . . . . . . 3. Posttranslational modifications regulating catalytic activity of enzymes . . . . . . . . . . . . 3.1. Regulation by targeted proteolysis and destruction . . . . . . . . . . . . . . . . . . . . 3.2. Regulation by reversible phosphorylation and related PTMs . . . . . . . . . . . . . . . 3.3. Regulation by modifying cysteine residues under dynamic redox state . . . . . . . . . 3.4. Regulation of secreted and membrane-associated enzymes by glycosylation . . . . . 3.5. Regulation of enzyme activities by glycation and other nonenzymatic PTMs . . . . . . 3.6. Multiple posttranslational modifications regulating enzyme activities . . . . . . . . . 4. Practical aspects of enzymes posttranslational modifications . . . . . . . . . . . . . . . . . . 5. Conclusions and perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1.

Introduction

One of the strategic directions in the postgenomic era of today's biosciences is the monitoring of the complete life cycle of key cellular enzymes, the critical effector molecules and regulatory components of living cells. The main stages of this complex tracking process include genetic coding, synthesis of mRNA transcript, proteosynthesis, enzyme transport within the cell, its folding and assembly from individual protein subunits, and its numerous PTMs, both enzymatically catalyzed and nonenzymatic [1]. Enzymatic PTMs can be detected in ever increasing amounts and they appear to be critical for folding and assembly (e.g. glycosylation), function as key regulators of catalytic activity of enzymes (e.g. binding of prosthetic groups, phosphorylation), or mark enzyme molecules for targeted destruction (e.g. ubiquitylation and related modifications). In parallel with these processes there are nonenzymatic PTMs caused by reactive chemical species continuously acting on individual enzyme molecules as a consequence of aerobic metabolism of most forms of today's life. Nonenzymatic PTMs are contributing to molecular aging; however, these modifications may also be involved in regulation of enzymes' catalytic activity [2]. Nonenzymatic PTMs involve oxidation and nitration resulting from direct action of ROS and reactive nitrogen species (RNS) on enzyme molecules as well as glycation, “carbonyl stress” caused by reactive carbonyl compounds, carbamylation, and other subsequent chemical reactions such as isomerization, deamidation, and racemization [2]. The number of enzymes the complete life cycle of which has been followed remains so

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far limited. However, since such a tracking is essential to gain a deep insight into their structure and function, considerable efforts have been concentrated towards this direction. Abundant cytoskeletal proteins and chromatin-associated histones provide prototypic polypeptides displaying a rich mixture of PTMs. In the former example, the products of lysine acetylation, lysine methylation, glutamic acid methylation, arginylation, glutamylation, glycosylation, palmitoylation, phosphorylation, ubiquitylation, sumoylation, and glycation have all been detected in α- and β-tubulins, the building blocks of microtubules [3]. In the latter case, the complete posttranslational code of histone deacetylases (EC 3.5.1.98) has been investigated by Segré and Choica [4], and found to involve phosphorylation, acetylation, ubiquitylation, sumoylation, nitrosylation, and carbonylation. The dynamics of intracellular palmitoylation may be controlled through interplay with distinct PTMs such as phosphorylation and nitrosylation [5]. The continuous and intertwining processes of enzymatic and nonenzymatic PTMs of enzymes are expressed in organism-, tissue-, and organelle-specific manner, and are critical in cellular regulations during normal as well as pathological processes (such as diabetes, neurodegenerative, autoimmune, and many other diseases). Thus, the development of new large throughput proteomic methods for the global identification of individual posttranslational modifications remains critical. The aim of the present paper is to review this rapidly developing area of contemporary biomedical research with an emphasis on newly developed robust proteomic techniques capable of simultaneous monitoring of enzyme PTMs and their functional consequences on a large scale.

Please cite this article as: Ryšlavá H, et al, Effect of posttranslational modifications on enzyme function and assembly, J Prot (2013), http://dx.doi.org/10.1016/j.jprot.2013.03.025

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2. Posttranslational modifications of enzymes followed by proteomic techniques 2.1. Most common posttranslational modifications of enzymes Enzymatically catalyzed posttranslational modifications of enzymes can be divided into two main categories. The first group involves all proteolytic processes involving breakage of any peptide bond in enzyme's polypeptide, although peptide bonds following single or double basic amino acids appear to be the most common targets [6]. The classical examples of this category include proteolytic activation of inactive proenzymes resulting in fully active enzyme molecules [7]. The protective prosequences are usually rather short, although an example of a large propeptide regulating the activity of fungal hexosaminidases (EC 3.2.1.52) has been recently reported [8]. The cleaved propeptides are either metabolized or reassociated with the catalytic subunit of the parent enzymes (either covalently or noncovalently) to sustain their full activity [8]. The second category of enzymatically catalyzed PTMs of enzymes consists of processes that modify the side chains of various amino acid residues (or the free N-terminal amino group or the free C-terminal carboxyl) without any chemical changes of the polypeptide backbone. The chemical nature and function of these modifications is diverse (see Table 1 for the current survey of the most important enzymatic modifications) and occur in many cellular compartments (Fig. 1). Of primary importance for enzymes is the covalent attachment of a prosthetic group essential for their catalytical activity. The most common examples include the condensation of biotin (prosthetic group of carboxylases, EC 6.4.1.2, 3) or lipoate

(essential for acetyltransferases, EC 2.3.1.12) to ε-amino group of lysine, binding of 4′-phosphopantetheine (prosthetic group of acyltransferases, EC 2.3.1.85) to serine hydroxyl, binding of FAD (prosthetic group of oxidoreductases, EC 1.3.99.1) to the nitrogen of the imidazol ring of histidine, and binding of heme to methionine and other amino acids in hemoproteins [1]. All of these prosthetic groups remain covalently bound to the apoenzyme through the entire catalytic process with the exception of pyridoxal enzymes (e.g. aminotransferases EC 2.6.1.X) in which a bond between pyridoxal phosphate and the lysine amino group is converted into the bond between the coenzyme and the substrate amino acid during the catalytic cycle [1].

2.1.1.

Glycosylation

Glycosylation of secreted enzymes is one of the most frequent and abundant enzyme modifications contributing significantly to their solubility, stability, folding, and assembly into fully active complexes [9–11]. The protective role of glycosylation against toxic radicals generated by xenobiotics was recently described by Martinek et al. [12]. Glycosylation starts with N-glycosylation defined as the transfer of large oligosaccharide composed of 2 GlcNAc, 9 Man and 3 Glc residues from the dolichol diphosphate activated precursor onto the amide of Asn occurring in the sequon Asn-Xxx-Ser/Thr (where Xxx is any amino acid except Pro) catalyzed by the oligosaccharyltransferase enzyme complex (EC 2.4.1.119). This process occurs cotranslationally as soon as the unfolded enzyme polypeptide passes through the secretory protein channel into the oxidative environment of the lumen of ER (Fig. 1). After the attachment to enzyme polypeptide, the above oligosaccharide precursor is further modified by many enzymes in ER and Golgi making N-glycosylation one of the most complex PTM. Later in the Golgi complex, O-glycosylation may

Table 1 – Most common enzymatic posttranslational modifications of enzymes. Modification

Organism/tissue

Organelle

Target aa

MS increase

Ref.

Proteolytic processing Biotin or lipoate binding 4′-Phosphopantetheine FAD Heme B Pyridoxal phosphate N-Glycosylation O-Glycosylation Phosphorylation O-GlcNAc Ubiquitylation a Methylation Prenylation (farnesyl) Acetylation b Myristoylation Palmitoylation –SH group modification ADP ribosylation Hydroxylation Glu carboxylation Sulfation

Universal Universal Universal Universal Universal Universal Eukaryotes Eukaryotes Universal Eukaryotes Eukaryotes Eukaryotes Eukaryotes Universal Eukaryotes Eukaryotes Universal Eukaryotes Eukaryotes Eukaryotes Eukaryotes

Lysosome, cytoplasm Mitochondria Mitochondria, cytoplasm Mitochondria Mitochondria Cytoplasm ER, Golgi Golgi Cytoplasm Cytoplasm, nucleus Cytoplasm Nucleus Cytoplasm Nucleus Cytoplasm Cytoplasm Periplasm, ER Nucleus Nucleus Extracellular Cytoplasm

Many Lys Ser His Met Lys Asn Ser, Thr Tyr, Ser, Thr, His, Asp, Arg Ser, Thr Lys Lys, Arg Cys Lys Gly (N-term.) Cys Cys Arg, Asn, Cys Pro, Lys, Asn Glu Tyr

Many 188, 226 340 783 616 229 >1000 203, 365 80 203 >1000 14, 28, 42 204, 206 42 210 238 −2 541 16 44 80

[6–8] [1] [1] [1] [1] [1] [9,11] [9,11] [18,19,326] [22] [23,24] [33,34] [37,38] [35] [40] [41] [42] [62] [48] [61] [63]

a

A number of ubiquitin-related proteins may participate in this type of PTMs leading to sumoylation [27], neddylation [28], Apg12 conjugation, and other modifications. b Lysine propionylation and butyrylation has been also detected in histones [327].

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Fig. 1 – Subcellular localization of individual PTMs. Only the most important PTMs are shown. AGE abbreviates advanced glycosylation endproducts interacting with the specific receptor (RAGE).

also be added in the form of shorter oligosaccharide sequences attached to Ser or Thr residues, especially in enzymes rich in these amino acids or displaying them in clusters [11]. Many enzymes thus acquire N- and O-glycosylation at several glycosylation sites, which may contribute to their complex molecular architecture and maintain their long-term stability [10,13–16].

2.1.2.

Phosphorylation and O-GlcNAc modification

Phosphorylation is the most frequent PTM; it is proposed that up to 30% of mammalian proteins could be phosphorylated. Reversible protein phosphorylation affects many physiological processes such as basic cellular metabolism and energy balance, growth, division, differentiation, and motility of cells, organelle trafficking and membrane transport, muscle contraction, signaling in nervous system, and response to altered

environment. Phosphorylation is catalyzed by enzyme protein kinases (EC 2.7.X.X); the reverse reaction by protein phosphatases (EC 3.1.3.16 and EC 3.1.3.48). Both reactions are separately controlled. The phosphate group can be bound to the protein in several ways. The most common are phosphomonoesters generated on –OH group of serine, threonine or tyrosine. Other possibilities are phosphoramides (histidine, arginine, lysine), acylphosphate (aspartate), and thiophosphate (cysteine). Phosphohistidine, phosphoarginine, and phospholysine in proteins are less stable than phosphomonoesters, so this modification could remain undiscovered in phosphoproteomic analyses performed under acidic conditions [17]. Protein tyrosine kinases (EC 2.7.10.X) and protein serine/threonine kinases (EC 2.7.11.X) are the most prominent effectors of protein phosphorylation able to transfer γ-phosphate from ATP onto a

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hydroxyl of the respective amino acids [18]. Histidine kinases (EC 2.7.13.X) appear to be universally distributed. These enzymes in plants and bacteria may function in a two step signal transduction system in which the inorganic phosphate is first attached to the nitrogen N1 of the imidazol ring of histidine in the enzyme itself and then transferred onto an aspartate residue in the target protein [19]. Protein phosphorylation may involve multiple sites in a single enzyme and rarely proceeds to completion. A well documented example of this modification includes eukaryotic RNA polymerase II (EC 2.7.7.6), the C-terminus of which contains many repeats (up to 52 in mammals, 26–27 in yeasts) of the heptapeptide sequence Tyr-Ser-Pro-Thr-Ser-Pro-Ser. Multiple phosphorylation of Ser and Thr residues of this repeats enhances the binding of transcription elongation factors and associated proteins thus converting the transcription preinitiation complex into a stable elongation complex [20]. In view of its regulatory role, the protein phosphorylation is a typical reversible protein modification and the enzymes of signaling cascades are eventually dephosphorylated through the action of protein phosphatases [21]. A competing modification of the target Ser/Thr hydroxyls occur through their modification with O-glycosidically linked GlcNAc bringing additional complexity to this regulatory system (Fig. 1) [22]. This modification occurs in cytoplasm and nucleus distinguishing it from O-glycosylation in ER or Golgi mentioned above (in Section 2.1.1 dealing with glycoprotein enzyme glycosylations).

2.1.3.

Ubiquitylation and related modifications

Another PTM of enzymes critical for many aspects of their fate in eukaryotic cells is acylation of lysine residues in proteins by activated C-terminal carboxyl group of glycine in ubiquitin (76 amino acid long peptide of approximately 8 kDa) and ubiquitin-related proteins termed ubiquitylation (or sometimes ubiquitination) [23,24]. The main purpose of this reaction is to

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mark the modified protein for degradation in the proteasome. Thus, this PTM is widespread in either damaged proteins or those for which a regulated destruction is necessary for their physiological role (e.g. targeted destruction of cyclins during precisely defined stages of the cell cycle of eukaryotic cell). Conjugation of the target protein with ubiquitin or related proteins is a three-stage process (Fig. 2). First, ubiquitinactivating enzyme E1 activates the C-terminal glycine using ATP forming ubiquityl-CO-AMP, and then ubiquityl-CO-S-E1. The second stage involves the transfer of activated ubiquitin onto a –SH group of cysteine in the ubiquitin-transporting protein E2. In the third stage the ubiquitin-protein ligase E3 (EC 6.3.2.19) catalyzes the transfer of ubiquitin onto protein substrate forming an amide bond between the activated C-terminal Gly76 of ubiquitin and lysine residue of the target protein [25]. Usually, conjugation is directed to a specific short amino acid sequence within the target protein named a degron. The conjugation process can be repeated several times: Lys of the first molecule of conjugated ubiquitin can be further modified by a second molecule of activated ubiquitin etc. thus leading to polyubiquitylation increasing the probability of the destruction of the target protein in the proteasome. In addition to constitutive ubiquitylation, a regulated variant of this PTM exists in which the degron peptide has to be phosphorylated, hydroxylated, glycosylated or N-terminally aminoacylated for efficient E3 ligase recognition. Several ubiquitin-like proteins have been identified including SUMO [26], Nedd8 [27], and others [23] sharing similar amino acid sequence and spatial structure. In case when an extended C-terminal sequence beyond the GlyGly* activation sequence occurs, peptidase cleaves the extra peptide and prepares ubiquitin-like proteins for ATP-mediated activation (Fig. 2). Ubiquitin-like proteins contribute to the diversification of the cellular systems of targeted proteolysis. The biological importance of ubiquitylation is emphasized by the fact that genes coding for its individual components amount a significant percent of the entire

Fig. 2 – Chemical reactions of ubiquitylation and deubiquitylation. Similar reaction schemes apply to both ubiquitin and ubiquitin-like protein marks. Modified from [333–336]. Please cite this article as: Ryšlavá H, et al, Effect of posttranslational modifications on enzyme function and assembly, J Prot (2013), http://dx.doi.org/10.1016/j.jprot.2013.03.025

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eukaryotic genomes [28]. Protein monoubiquitylation has been demonstrated to play many nondestructive functions as well mediating protein–protein interactions and participating in processes such as endocytosis, DNA repair, transcription, translation, and signal transduction [29–31]. During these functions protein may also be deubiquitylated by deubiquitylating enzymes that are themselves tightly regulated by substrateinduced conformational changes, binding to adaptor proteins, proteolytic cleavage, or even other PTMs [32].

2.1.4.

Alkylation, acylation, and prenylation

The charged groups of basic amino acids Lys and Arg can be modified by alkylation (most often methylation) or acylation (most often acetylation), both of which represent common and functionally very important PTMs of enzymes [33–35]. Protein methylation is maintained by dynamic equilibrium between the activities of methyltransferases (EC 2.1.1.X) using S-adenosyl methionine as substrate, and demethylases acting as FAD-dependent aminooxidases or enzymes requiring Fe2+ ions, 2-oxoglutarate and ascorbate. During methyltransferase catalyzed reactions lysine can be converted into mono-, dior trimethyllysine while arginine can form mono- or dimethylarginine [36]. Protein methylation has been most extensively studied on histone modifications where the participating enzymes are themselves subject to regulation by a large number of PTMs as already mentioned above [4]. Prenylation is modification of Cys residues by isoprene residues such as farnesyl or geranylgeranyl [37,38]. Modifications with these alkyl chains are used for membrane anchoring of molecular switches from Ras-, Rab-, and Rhofamilies. Prenylations are catalyzed by protein farnesyl transferases (EC 2.5.1.58), type I protein geranylgeranyl transferases (EC 2.5.1.59) targeting a sequence Cys-Aaa-Aaa, where Aaa is a small aliphatic amino acid, and type II protein geranylgeranyl transferases (EC 2.5.1.60) targeting a sequence Cys-Cys-XxxXxx or Xxx-Xxx-Cys-Cys or Xxx-Cys-Xxx-Cys, where Xxx may be any amino acid, respectively. The functional consequences of enzyme acylations are very similar to those of enzyme methylations: substitutions of lysine residues by short chain acyls such as acetyls neutralize their charges while the attachment of long alkyl acyls (palmitoylation, myristoylation, etc.) provides the target enzymes with hydrophobic lipid anchors allowing them to associate with a plasma membrane. Lysine acetylation has been again most extensively studied for histones; this process is catalyzed by histone acetyltransferases (EC 2.3.1.48) that transfer an acetyl group from acetyl coenzyme A to ε-amino groups of lysine residues. Interestingly, acetylation of lysine residues in the C-terminal domains of enzymes protects them from ubiquitylation, and increases significantly their lifespan and functional activities [39]. Myristoylation and palmitoylation are the two most common modifications by long fatty acid residues [40,41] transferred onto proteins from the respective acyl coenzyme A derivatives produced during oxidative cleavage of fatty acids. While the former modification involves the transfer of myristylate to the free N-terminal amino group of Gly (most often formed after the cleavage of the initiation Met) catalyzed by myristoylCoA: protein N-myristoyltransferase (EC 2.3.1.97) and is considered irreversible, the latter acylation involves the addition of palmitoyl

to cysteine residue forming a thioester bond and is subject to the palmitoylation/depalmitoylation dynamics resembling other reversible PTMs.

2.1.5.

Oxidation

Oxidation of –SH groups of cysteines in enzymes and the formation of disulfide bonds is connected with relatively minor mass shifts and proceeds through apparently simple mechanism [1] yet it represent one of the most important PTMs from the point of stabilization of enzyme structure [42]. Moreover, the proper redox status of these groups is also a critical determinant for many thiolate enzymes that use this group as part of their catalytic centers [43]. In addition to the oxidation of –SH group into disulfide, additional reactions such as the formation of sulfi- and sulfoacids as well as binding of glutathione are also possible [44]. Individual cysteines differ in their reactivity since the propensity to form thiolate anions depends on the surrounding electrostatic environment in the structure of a given protein. Reduction of the disulfide bonds in vivo is accomplished mostly by the glutathione tripeptide (γ-Glu-Cys-Gly, GSH) in combination with high levels of NADPH and enzymes glutathione reductase (EC 1.8.1.7) and thioredoxin reductase (EC 1.8.1.9). As a secreted enzyme moves into endoplasmic reticulum and down the secretory pathways of eukaryotic cells, the levels of GSH and NADPH decrease, and stable disulfide bonds are eventually formed catalyzed by protein disulfide isomerase (EC 5.3.4.1) in ER lumen [45]. Oxidative hydroxylation reactions can proceed at the nonnucleophilic –CH2– groups of Pro, Lys or Asn leading to the formation of 3-hydroxyproline, 4-hydroxyproline, 5-hydroxylysine, and 3-hydroxyasparagine catalyzed by nonheme iron (Fe2+), 2-oxoglutarate dependent dioxygenase family of enzymes (EC 1.14.11.X). 2-Oxoglutarate undergoes a decarboxylation reaction consuming remaining oxygen atom to form succinate and CO2. Important substrate of prolyl-4-hydroxylase (EC 1.14.11.2) is collagen, where hydroxyprolyl residues participate in the formation of compact helical structure [46,47]. The introduced hydroxyl can be further glycosylated and short oligosaccharide sequences are then used for cross-linking and stabilization of collagen and other fibrous proteins [48]. Hydroxylation of proline is a key reaction in hypoxiainducible factor (HIF), which mediates cellular response to low oxygen by expression of many genes including glucose metabolism, erythropoiesis, angiogenesis, and apoptosis. If oxygen is available, proline is hydroxylated and then the entire protein proteasomally degraded [49]. Peptides aspartyl/ asparaginyl beta-hydroxylase (EC 1.14.11.16), lysine hydroxylase (EC 1.14.11.4), and trimethyllysine hydroxylase (EC 1.14.11.8) also belong to the family of dioxygenases (EC 1.14.11.X) with similar mechanisms of reaction [46,50,51]. Aromatic amino acids could also be hydroxylated by members of the family of non-heme iron monooxygenases (EC 1.14.16.X) using tetrahydrobiopterin as the source of electrons. Phenylalanine hydroxylase (EC 1.14.16.1), tyrosine hydroxylase (EC 1.14.16.2), and tryptophane hydroxylase (EC 1.14.16.4) are involved in metabolic pathways important for normal functioning of nervous system. These enzymes have similar catalytic mechanisms but divergent regulatory properties [52,53]. Nonenzymatic posttranslational modifications (see the survey in Table 2) of enzymes result from exposure of their

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Table 2 – Most common nonenzymatic posttranslational modifications of enzymes. Modification Glycation Lipooxidation Oxidation, nitration Carbamylation S-nitrosylation Cyclopentenone PG Homocysteinylation Deamidation Transamination Cyclization a

Organism Universal Universal Universal Universal Universal Animals Animals Universal Bacterial GFP

Organelle

Target aa

Extracellular, cytoplasm Cytoplasm Mitochondria, cytoplasm Mitochondria, cytoplasm Mitochondria, ER Cytoplasm Cytoplasm Cytoplasm Cytoplasm Extracellular

Lys, Arg Lys, Arg Many Lys Cys Cys (His) Lys Asn, Gln Gln + Lys Ser-Tyr-Gly

MS increase a

162 16 Variable 43 40 335 117 1 Cross-link −18

Ref. [328] [102] [54] [329] [58] [60] [330] [331] [1] [1]

Many additional end products formed due to subsequent rearrangements and cross-linking reactions.

molecules to reactive chemicals occurring inside the cell due to aerobic oxygen metabolism in today's forms of life. The most important group of these compounds includes ROS, RNS, reactive aldehydes originating from carbohydrate, or lipid metabolism (reducing sugars, oxidized lipid intermediates, glyoxal, malondialdehyde, and others). These chemicals tend to react nonspecifically attacking a wide range of enzyme's amino acids irreversibly, thus retaining their molecular marks on individual enzyme molecules. Unlike the enzymatic PTMs described above, these modifications most often impair the catalytic activities of modified enzymes functioning as the reaction centers for cross-linking, aggregation, precipitation, and other detrimental processes related to enzyme aging. More recently, however, we began to appreciate the physiological role of these modifications in signaling and other normal processes of homeostasis. Altogether, nonenzymatic PTMs of enzymes are situated between their early covalent enzymatically catalyzed modifications and noncovalent interactions of enzymes with their substrates, allosteric regulators, coenzymes, metal ions, physiological buffer components, and other components with which enzymes react in a fully reversible fashion. Because of their participation in many molecular dysfunctions accompanying age related diseases such as diabetes, renal insufficiencies, atherosclerosis, or neurodegenerative diseases the end products of nonenzymatic PTMs have a great potential to serve as early biomarkers of these diseases [2]. Enzymes can be also directly altered by ROS to generate advanced oxidation products [54]. The most abundant products involve methionine sulfoxide formed by Met oxidation, kynurenin and N-formylkynurenin obtained by Trp oxidation, 3-nitrotyrosine formed by Tyr nitration, and 3-chlorotyrosine coming from chlorination of this amino acid. The recent review by Møller et al. [54] lists a total of 64 documented amino acid modifications including several on amino acids considered to be rather “nonreactive” (i.e. one change in Ala and two changes in Val, Leu, and Ile). Thus, ROS can modify side chains of amino acids in proteins, especially of sulfur containing amino acids. Methionine, as a rather hydrophobic amino acid, is often localized inside the protein structure but methionine residues exposed to the surface are oxidized to methionine sulfoxide. This modification is reversible; methionine sulfoxide could be returned to methionine by the catalysis of methionine S-oxide reductase (EC 1.8.4.11, 12) using thioredoxin disulfide as a reducing power

[55,56]. Methionine sulfoxide seems to have physiological functions in the cell. Ca2+/calmodulin dependent protein kinase II is a key enzyme in Ca2+ signaling pathway. Oxidation of two methionine residues to methionine sulfoxide activates this protein kinase in Ca2+/calmodulin independent manner similar to phosphorylation of specific threonine residues. Methionine oxidation also affects the secondary structure of proteins. The transition of normal cellular prion protein to infectious one means an α-helix to β-sheet shift and methionine oxidation plays an important role in this process [56]. Also other amino acids are susceptible to oxidation: tryptophane oxidation leads to kynurenin and N-formylkynurenin, and histidine forms 2-oxohistidine. Tyrosine could form 3,4dihydroxyphenylalanine, dityrosine or chlorotyrosine and phenylalanine ortho- or meta-tyrosine [57]. NO and other RNS can also modify amino acids in proteins, e.g. tyrosine forms 2-nitrotyrosine [57].

2.1.6. Glycation, lipooxidation, and modification by reactive carbonyl compounds Glycation represent a widespread PTM for both intracellular and secreted proteins (Table 2). It refers to binding of Glc and other reducing sugars to proteins. The initial reactions of carbonyl (aldehyde) group of Glc with ε-amino group of Lys leads to the formation of Schiff base (N-glycosylimine) that undergoes Amadori rearrangement into a stable ketoamine (1-amino-1-deoxyfructose, Amadori product). Amadori products are further processed by dehydration, cyclization, fragmentation, and oxidation leading to advanced glycation end products (AGEs). The levels of glycated proteins in blood (glycated hemoglobin and albumin) reflect long term (integral) values of blood Glc levels, and thus became invaluable in monitoring diabetic patients. The second common nonenzymatic posttranslational modification of enzymes results from their reaction with several reactive carbonyl compounds such as glyoxal, methylglyoxal, and malondialdehyde. These compounds are generated from glucose and other reducing sugars by their direct oxidation, from Amadori products by their further rearrangement, and from peroxidation of lipids [2]. Increased cellular concentration of these carbonyl compounds occurring under pathological conditions contribute to “carbonyl stress” and subsequent protein carbonylation leading to both AGEs and ALEs [2]. Obviously, all modifications caused by the above reagents are

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not the result of a simple linear processes but form a very complex network of reactions in which the three above listed pathways (glycation, lipooxidation, and carbonylation) are often mutually intertwined. Another nonenzymatic PTM of enzymes is their carbamylation due to a reaction of isocyanic acid primarily with ε-amino group of Lys leading to the formation of homocitrulline, the most characteristic carbamylation end product. Isocyanic acid is, in turn, formed by a spontaneous dissociation of urea (abundant in uremic patients) or by transformation of thiocyanate by myeloperoxidase in the presence of hydrogen peroxide. S-nitrosylation is a recently described nonenzymatic PTM that appears to be reversible and apparently selective [58]. This PTM can regulate enzyme activity, localization, and stability in a wide variety of tissues and cellular systems [58]. S-nitrosylation of enzymes occurs in vivo through several mechanisms including oxidative S-nitrosylation by higher oxides of NO, transnitrosylation by small molecular weight NO carriers such as S-nitrosoglutathione. It can be catalyzed by metalloproteins, or proceed as protein-assisted transnitrosylation documented for S-nitrosylation of caspase-3 (EC 3.4.22.56) by S-nitrosothioredoxin [59]. Similarly, cyclopentenone prostaglandin enzyme adducts have been described only recently as key regulators of proinflammatory transcription factors, signaling kinases, and proteins involved in control of the redox environment [60].

2.1.7.

Other posttranslational modifications

There are virtually hundreds of additional PTMs that may occur in enzymes and the reader is thus referred to the corresponding databases that are regularly updated (see Table 3). Several blood-clotting factors contain γ-carboxyglutamic acid formed through the action of vitamin K dependent γ-glutamylcarboxylase (EC 4.1.1.90) [61]. Among the newly emerging enzyme PTMs, we would like to briefly mention ADP-ribosylation. ADP-ribosylation was shown to be involved in the regulation of enzymes participating in many key cellular processes such as DNA repair, apoptosis, and function of mitotic spindle during cellular division [62]. NAD serves as a donor of ADP-ribosyl residue. The nicotinamide bond in this coenzyme is cleaved by ADP-ribosyltransferase (EC 2.4.2.31) forming a ribooxocarbene cation, which then interacts with various nucleophilic groups in proteins (Arg, Asn, Cys). In response to DNA breaks, poly-ADPribosylation occurs in which the additional ADP-ribose residues are attached to 2′-OH groups in adenylate in a manner similar to polyubiquitylation. Related to phosphorylation and resulting in identical mass increment (Table 1) is protein sulfation

occurring in some cell surface adhesion molecules such as selectins [63]. This modification is catalyzed by protein-tyrosine sulfotransferase (EC 2.8.2.20) that uses phosphoadenosyl phosphosulfate as substrate, and it is cleaved by arylsulfatases (EC 3.1.6.1).

2.2. Proteomic methods for global detection of posttranslational modifications of enzymes The modification of enzyme's molecules by enzymatic or chemical processes described in the preceding chapter contributes significantly to the fact that the proteome of a cell or an organism consists of many more components than the absolute number of genes coding for individual polypeptides. However, the exact nature of these modifications makes their faithful dissection a formidable analytical challenge. There are many factors that contribute to difficulties in monitoring enzymes PTMs during their entire lifespan. In particular, the modifications are numerous, dynamic, often mutually interlinked (as has been briefly discussed above), and rarely proceed to completion. Traditionally, the detection of individual PTMs has been relying on spectroscopic or chemical derivatization techniques [64], but these are most often not enough specific and rarely allow a simultaneous monitoring of several PTMs in one sample at the same time. Today's proteomics can employ modern high-resolution mass spectrometry techniques and take snapshots of the complete spectrum of PTMs imposed upon individual molecules [65–69]. However, since defining a particular PTM by its mass increment only is difficult and can lead to misinterpretations, the combination of a specific chemical probing of a particular PTM (using techniques such as stable isotope metabolic labeling, specific fluorescent or otherwise labeled tags, or affinity capture and concentration of the target modified peptides) with high resolution MS and MSn techniques as analytical tools has provided the most promising strategies for complete proteomic dissections of individual PTMs at the global scale [70]. In fact, successful and robust protocols along these lines of research have already been reported for nearly all of the most critical PTMs (Table 4). Proteomic identification of free N-terminal ends in proteins represents a powerful approach for monitoring numerous physiologically relevant proteolytic processes, protease networks, and network-sculpted proteomes [71]. This approach may also be used in a more targeted way for the substrate degradomes of specific proteases. For instance, Prudova et al. developed iTRAQ labeling method that simultaneously modified and blocked all primary amines including

Table 3 – Databases for the record of posttranslational modifications. Database

Modification

Entries

Ref.

UniProtKB/Swiss-Prot Human Proteinpedia Unimod Delta Mass (ABRF) Resid Phosida GlycosuiteDB

All All All All All Phosphorylation Glycosylation

75,291 15,231 948 350 589 80,082 9436

http://www.expasy.org http://www.humanproteinpedia.org http://www.unimod.org http://www.abrf.org http://www.ebi.ac.uk http://www.phosida.com http://www.expasy.org

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Table 4 – Proteomic identification of enzyme's posttranslational modifications performed at global scale. Modification

Model

Principle of analysis

# identified proteins/modifications

Ref.

Enzymatic Proteolysis Glycosylation Glycosylation Phosphorylation O-GlcNAc Ubiquitylation Lys methylation Arg methylation Lys acetylation Farnesylation Palmitoylation –SH group modification ADP ribosylation

Fibroblasts, MMP D. melanogaster Human AML cells EGF stimulated HeLa cells Human HEK293 cells Human HEK293T cells Human histones Human HeLa cells Human AML cells COS-1 cells Human Jurkat T cells Yeast mitochondria CHO cells

iTRAQ Affinity purification Affinity purification SIL, TiO2 Chemical modification Immunopurification LC–MS-MRM Immunopurification Immunopurification Substrate tagging Substrate tagging Affinity purification Affinity purification

1054/3152 202 505 2244/6600 374 4273/11,054 2/20 200 1750/3600 18 125 139 12

[72] [75] [76] [81] [87] [89] [93] [34] [94] [37] [41] [99] [62]

Nonenzymatic Glycation Lipooxidation Carbonylation S-nitrosylation Nitroxylation

Human plasma proteins Rat skeletal muscle Rat skeletal muscle Mouse liver Human platelet

Radiolabeling LC–MS/MS 18 O labeling Affinity purification MS (ECD)

50/161 3 60/210 192/328 10

[104] [232] [332] [58] [106]

protein N-termini and lysine side chains. Using this approach these authors could follow 3152 N-terminal peptides corresponding to 1054 unique proteins of which 201 were cleaved by matrix metalloproteinase-2 (gelatinase A) (EC 3.4.24.24) and only 19 by matrix metalloproteinase-9 (gelatinase B) (EC 3.4.24.35) [72]. The global view on glycosylation of cell surface glycoproteins has proved essential for our better understanding of many roles of glycoproteins and has found many practical applications in experimental oncology, pharmacoproteomics, etc. The two major technologies (referred to as glycoproteomics and glycomics) are based on affinity capture and subsequent analysis of glycosylated peptides after their controlled periodate oxidation and subsequent conjugation to biotin hydrazide, or labeling, release, and analysis of all surface oligosaccharide sequences, respectively [73,74]. The former approach turned useful for monitoring the changes in cell surface glycoproteome upon prolonged insulin stimulation [75], cell surface phenotyping of myeloid leukemia [76], and quantitative proteomic identification of proteins regulating cell migration [77]. Glycomic technologies, on the other hand, have found use for characterization of human stem cells [78], evaluation of human induced pluripotent stem cells [79], or evaluation of surface glycome in glycosylation-defective CHO cell variants [80]. Using SIL, strong cation exchange chromatography, and TiO2 phosphopeptide enrichment, it was possible to monitor dynamic changes in global phosphoproteome including 6600 phosphorylation sites on 2244 proteins upon stimulation of human HeLa cells with epidermal growth factor in vivo [81]. In addition to kinases, the major targets were ubiquitin ligases, guanine nucleotide exchange factors and transcriptional regulators. The analysis of more limited sub-phosphoproteomes has become even more popular, e.g. phosphotyrosine interactome of ErbB receptor or studies on protein kinase C (EC 2.7.11.13) in the heart [82,83]. Development of mild β-elimination followed by Michael addition of dithiothreitol provided a suitable

method for proteomic analysis of the dynamic relationship between nucleocytoplasmic O-GlcNAc modification and phosphorylation [84,85]. Recently, even more sensitive methodology based on metabolic incorporation of azido derivatives of GalNAc via the GalNAc salvage pathway has been applied by Bertozzi and coworkers [86,87]. Immunoenrichment of modified peptides has been combined with high resolution MS for proteome-wide mapping of ubiquitylation sites or modification by ubiquitin-related proteins [88–90]. The global survey revealed 11,054 endogenous sites of ubiquitylation on 4273 human proteins including 800 already known and 10,254 novel modifications on enzymes of cellular signaling cascades, enzymes important for DNA replication, DNA damage repair, and cell cycle progression [89]. Christianson et al. investigated functional organization of the human ER-associated degradation network integrating proteomics, functional genomics, and the transcriptional response to ER stress [91]. Using an advanced systems-level strategy, a complete functional model describing individual protein interactions could be constructed and functionally evaluated. Yet again, mutual influence of enzymes' PTMs was documented by the fact that nonglycosylated proteins are preferably subjected to ER-associated degradation [92]. Global proteomic techniques for analysis of lysine and arginine methylation and acetylation based on immunoaffinity enrichment in combination with high resolution MS emerged quite recently [34,93–95]. The major impact of these techniques has been in the detailed characterization of the “histone PTM code”, in particular the catalytic and inhibitory loops describing the cross-talk between H2B ubiquitylation and H3 Lys79 methylation [93]. The global analysis of Lys acetylation by high resolution MS was able to monitor 3600 lysine acetylation sites on 1750 proteins and reveal how this PTM preferentially targets large macromolecular complexes involved in chromatin remodeling, cell cycle, splicing, nuclear transport, and actin nucleation [94]. Once again, a cross-talk of individual enzyme

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PTMs was apparent from the way in which acetylation impaired phosphorylation-dependent interactions of 14-3-3 protein and regulated the cyclin-dependent kinase Cdc28 (EC 2.7.11.22) [94]. Tagging through metabolic incorporation of substrate analog 8-anilinogeranyl and subsequent immunocapture using antibodies against the anilinogeranyl moiety proved useful for large scale profiling of protein prenylation and palmitoylation [37,41,96,97]. Using these techniques more than a hundred of palmitoylated proteins could be detected in mammalian cells including a family of hitherto uncharacterized hydrolases localized at the plasma membrane due to the occurrence of this PTM [41]. Monitoring of the global thiol proteome is critical for both the subsequent evaluation of thiol: disulfide equilibrium and for documentation of several nonenzymatic PTMs. Towards this goal, Hill and colleagues have standardized the usage of biotinylated fluorophore-labeled alkylating reagents such as iodoacetamide or N-ethylmaleimide at the global proteomic scale [98]. Experimental strategies of proteomics and transcriptomics have been combined to analyze redoxins involved in maintenance of thiol redox homeostasis [99]. Pompach et al. optimized electrophoretic and digestion conditions allowing a simple MS evaluation of disulfide bonding from a single protein spot opening this technique for the large-scale proteome analyses [100]. The technical development in the area of global proteomic monitoring of nonenzymatic PTMs of enzymes (Table 4, lower part) has been somewhat lagging behind that of enzymatic PTMs, many protocols still relying on chemical or spectral techniques for identification of these modifications and their end products [64,101–103]. However, modern techniques based on quantitative proteomics and MS are emerging fast and have already proven valuable for the evaluation and monitoring of glycation [104], oxidation and carbonylation [54,105], S-nitrosylation, nitroxylation, and others [106]. The rapid development of optimized protocols will hopefully soon enable to monitor the global changes in most individual PTMs of enzymes making it possible to start looking at multiple modifications in a single sample which is supposed to fasten our progress towards full understanding of their functional importance. In this respect, the development of those techniques suitable for high-throughput robust screening of nonenzymatic PTMs should be emphasized. Considering the number and complexity of enzyme PTMs, bioinformatics institutes and companies developed specialized databases reserved for the deposition of increasing amount of data, especially data coming from automated MS and (MS)n experiments. For PTMs concerning human enzymes, human proteinpedia has been established and is available at http://www.humanproteinpedia.org [107]. The list and brief description of other useful databases is provided in Table 3.

2.3. Compartmentalization of enzyme modifications and function Most enzymatic PTMs of enzymes are fixed to certain cellular locations defined by the corresponding milieu of enzymes executing them (Fig. 1). Thus, their presence on enzymes can per se give evidence for their cellular fate and individual history of the particular enzyme molecule. For instance, in

normal cell under physiological conditions the occurrence of glycosylation or oxidized disulfide bonds in enzyme's molecule can be taken as a trace of its movement through ER and the subsequent organelles [70]. Success of the global proteomic techniques in mapping of cell surfaces in tissues [108], in studies of unique membrane transport systems in bacteria and eukaryotes [109], or in profiling mitochondrial enzyme changes related to physiological processes or aging [110] proved already to have a critical impact on cellular biology. Modern proteomic studies are now changing our somewhat simplistic view on cellular localization of many key enzymes [111] even allowing the construction of their cellular interaction networks such as those involved in regulation of ER-associated degradation [72]. Finally, in plants, in which their subcompartmentation is particularly intriguing (in addition to chloroplasts they possess many other specific compartments not found elsewhere), many cellular compartments have been for the first time functionally clarified using proteomic techniques [112,113]. Nonenzymatic PTMs of enzymes are again connected with specific cellular compartments that create the chemical environment enabling their occurrence. The main compartment for ROS production in mammalian cells and in nonphotosynthesizing parts of plants is the mitochondrion where superoxide is produced mainly in complex I and complex III of the respiratory chain [54]. Modern tracing and proteomic studies revealed that superoxide is released mainly into mitochondrial matrix where it can be converted to H2O2 by Mn-dependent superoxide dismutase (EC 1.15.1.1). H2O2 can diffuse across the inner mitochondrial membrane through aquaporins and across the outer membrane through porin into the cytosol [114]. There are specific aquaporins facilitating the diffusion of hydrogen peroxide through membranes of individual compartments [104]. The most common mechanism of protein carbonylation in living cells is the metal catalyzed oxidation occurring when reduced metal ions like Fe2+ and Cu+ react with H2O2 producing extremely reactive hydroxyl radicals in Fenton reaction: Fe2+ + H2O2 → Fe3+ + OH− + OHU. The hydroxyl radicals oxidize amino acid side chains or cause protein backbone cleavage both resulting in the introduction of carbonyl groups [54]. Although the total metal ion concentration in mitochondria is very high, the free metal ion concentration is strictly regulated by metal binding proteins such as ferritin and frataxin. The critical importance of these proteins is illustrated both in humans (where frataxin deficiency causes cardio- and neurodegeneration designated as Friedreich's ataxia [115]) and in the yeast experimental model (frataxin deficiency leads to accumulation of carbonylated proteins while its overexpression increases oxidative stress tolerance [116]). It has been experimentally found that many mitochondrial matrix enzymes are carbonylated indicating their proximity to metal ion release sites [117]. The major source of iron release in mitochondrial matrix appears to be aconitase (EC 4.2.1.3) which comes into contact with other Krebs cycle enzymes during the formation of temporary multienzyme complex [118]. The redox environment is critical for both enzymatic and nonenzymatic PTMs including cysteine oxidation (a critical step during protein folding), cysteine S-nitrosylation, and other cysteine modifications. Mitochondria associated membranes of ER have been pinpointed as a specific ER subcompartment responsible for

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protein folding [119]. This region is enriched in ER folding chaperones and oxidoreductases. It appears to be functionally linked to the folding of newly synthesized proteins bridging the control of oxidative protein folding in ER with calcium homeostasis. Three critical proteins regulating these reactions have been identified: mitofusin-2, mortalin, and PACS-2. Mitofusin-2 is an outer membrane GTPase (EC 3.6.5.X) regulating contact and fusion between mitochondria as well as ER morphology, calcium homeostasis, and tethering of ER to mitochondria [120]. Mortalin is a homologue of hsp70 cytosolic chaperone that bridges the mitochondrial voltage dependent anion channel to the large N-terminal IP3 binding regulatory domain of the IP3R. PACS-2 localizes the calcium release channel transient receptor potential protein 2 and the chaperone calnexin into the mitochondria associated membranes and stabilizes its formation [121]. Calcium concentration in ER is dependent on the ubiquitylation of the ER stress protein Herp by E3 ubiquitin-ligase named Plenty Of SH3 (POSH) [122]. The ER maintains an optimal chemical environment to produce secreted proteins including millimolar concentration of free calcium, sufficient ATP supply, as well as an appropriate redox buffer in addition to specific ER chaperones and oxidoreductases. ER chaperones (in particular calreticulin) buffer calcium concentration within ER and together with oxidoreductases modulate calcium exchange with mitochondria. Many ER chaperones, such as glucose-regulated protein BiP and calnexin, depend on a constant supply of ATP provided both by glycolysis and mitochondrial oxidative phosphorylation. Finally, the ER enzyme critical for the regulation of redox environment identified initially in the yeast is Ero1p (EC 1.8.4.X), a glycosylated membrane associated flavoenzyme localized to the lumenal site. Ero1p uses molecular oxygen to oxidize protein disulfide isomerase, which can then catalyze the formation of disulfide bonds in the newly synthesized polypeptides [123]. Formation of disulfide bonds by Ero1-Lα and Ero1-Lβ leads to the production of H2O2 in ER making it (together with mitochondria) an organelle with the highest hydrogen peroxide concentration. High levels of this ROS occur inside ER under stress leading to inactivation of SERCA (sarco/endoplasmic reticulum Ca2+ ATPase; EC 3.6.3.8) by S-glutathionylation and the activation of IP3R by oxidation increasing the levels of calcium on the cytosolic side of ER. Thus, ROS production by Ero1 proteins may provide an additional mechanism for ER to attach to mitochondria under conditions of ER stress illustrating the dynamic nature of interactions between cellular compartments under normal and pathological conditions [81,111]. Certain PTMs may serve not only as markers of cellular fate and history of individual enzymes, but themselves enable and define some cellular location. A typical example includes the assembly of G proteins, critical transmembrane signaling molecules for the biggest family of membrane receptors [124] requiring a cooperation of four cellular compartments (Fig. 3). The process begins with the folding of Gβ subunit assisted by cytoplasmic CCT chaperone and association with the Gγ subunit. Gγ is farnesylated or geranylgeranylated by cytoplasmic transferases and acylated. Gβ/Gγ dimer thereafter associates with ER enabling its modification by specific ER enzymes. ER localized protease cleaves the three C-terminal amino acids of Gγ subunit and thus created isoprenylcysteine is carboxymethylated by isoprenylcysteine carboxymethyl

11

transferase (EC 2.1.1.100). Thereafter, Gα subunit is folded cytoplasmatically, associates with the Gβ/Gγ heterodimer, and is palmitoylated by ER/Golgi localized palmitoyl acyl transferase (EC 2.3.1.B1) before the final transport of the complex from ER through Golgi to plasma membrane (Fig. 3) [124]. Indeed, the recent review by Salaun et al. [5] emphasizes the Golgi as a possible “super-reaction center” for the palmitoylation of peripheral membrane proteins, whereas palmitoylation reactions in post-Golgi compartments contribute to the regulation of membrane association of specific substrates as a result of balance between palmitoylation and depalmitoylation [5]. There are about 23 putative S-palmitoyl transferases (EC 2.3.1.B1) in mammals characterized by the presence of the AspHisHisCys motif within a 50 amino acid cysteine-rich domain. Protein modified with initial prenylation or N-myristoylation acquires a weak membrane affinity allowing its transient membrane interaction. Palmitoylation by membrane-bound AspHisHisCys proteins promotes stable membrane association by kinetic trapping [125]. Also worth mentioning is the control of intracellular palmitoylation dynamics through interplay with other PTMs, notably phosphorylation and nitrosylation [5]. Another example includes the regulated degradation of excessive or damaged membrane proteins or oligomeric complexes that fail to fold correctly in ER by ubiquitylation [91,126].

3. Posttranslational modifications regulating catalytic activity of enzymes There are four groups of enzyme functions that require PTMs of amino acid residue side chains. Many PTMs directly or indirectly influence the spatial structure and subunit assembly of enzyme molecule. Second, the functional activity of a wide number of enzymes requires the presence of certain prosthetic groups covalently bound to apoproteins. The third important group of PTMs includes reversible protein modifications that switch the enzymatic activity on and off representing principal components of the cellular regulatory circuits. Finally, some PTMs in enzymes do not influence their structure or activity but serve as tags or addresses transporting enzyme molecules into individual cellular compartments or functional sites. In the following paragraphs we are going to discuss the details of these functional categories.

3.1.

Regulation by targeted proteolysis and destruction

Our view on the targeted proteolysis of enzymes and its functional importance has changed dramatically during the recent period due to discovery of new forms of extralysosomal intracellular proteolysis, namely ubiquitin-targeted proteasome degradation [127]. Regulation of polyubiquitin-dependent proteolysis occurs at several levels. First, by combining different E2 and E3 enzymes, which can be regulated by phosphorylation and dephosphorylation reactions, exquisite temporal and spatial control of ubiquitylation can be achieved [128]. Second, a large family of deubiquitylating enzymes functions to remove ubiquitin chain from substrates and to recycle ubiquitin [129]. Monoubiquitylation has been shown to be involved in nondestructive functions such as regulation of other PTMs, membrane

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Fig. 3 – Interplay of PTMs and cellular compartments during G protein assembly and plasma membrane translocation. Individual stages include Gβ and Gγ folding and CCT/phosducin-like protein chaperone dissociation (1–3), GβGγ farnesylation or geranylgeranylation (shown in red) (4), cleavage of C-terminal tripeptide in Gγ (5), isoprenyl cysteine methylation (6), Gα folding (7), Gα palmitoylation (8), and transfer to plasma membrane (9). aaX denotes a tripeptide composed of two aliphatic amino acids followed by any amino acid. According to [124].

protein trafficking, and others [130]. The recent discovery of new ubiquitin-like proteins (Nedd8, Sentrin/SUMO, Apg12, and others) has further broadened the horizon of this type of post-translational enzyme modification [27]. A functional catalog of human ubiquitylation has been recently provided by the hUbiquitome web resource listing 1 E1 enzyme, 12 E2 enzymes, 138 E3 ligases or complexes, 279 substrate proteins, and 17 deubiquitylating enzymes [131]. Analyses revealed that proteins ubiquitylated by RING E3 ligases are related to regulation of p53 tumor suppressor, regulation of receptors and transporters, protein quality control, angiogenesis, and apoptotic death, whereas those modified by other E3 ligases such as HECT are important for the regulation of gene expression [132,133].

Eukaryotic cell cycle transitions are driven by E3 ubiquitin ligases that catalyze the ubiquitylation and destruction of specific protein targets. For example, the anaphase promoting complex/cyclosome (APC/C) promotes the exit from mitosis via destruction of securin and mitotic cyclins, whereas E3 ligase CRL1Skp2 allows entry into S phase by targeting the destruction of the cyclin-dependent kinase inhibitor p27. Recently, an E3 ubiquitin ligase called CRL4Cdt2 has been characterized, which couples proteolysis to DNA synthesis via display of substrate degrons in the DNA polymerase (EC 2.7.7.7) processivity clamp. It thus emerges as a master regulator that prevents re-replication in S phase that would compromise genome integrity [134]. Cdt1, which is required

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for the recruitment of the MCM2-7 helicase (EC 3.6.4.12) to origins of DNA replication in G1 (the “licensing” reaction), but is destroyed in S phase, ensures that each origin of DNA replication undergoes only one initiation event per cell cycle. The data suggest a model wherein Cdt1 first uses its degron to dock onto the sliding clamp that is engaged in replication or repair synthesis. Subsequently, CRL4Cdt2 is recruited, whereupon Cdt1 is polyubiquitylated on chromatin and destroyed. The Mdm2/p53 pathway is considered the most important guardian of the genome whose activity is critical for maintaining the genome integrity. In response to stresses such as DNA damage, activation of the ATM/ATR/CHK kinase (EC 2.7.11.1) pathway results in p53 phosphorylation at Ser15 and Ser20, which serve to disrupt the interaction between Mdm2 and p53. Phosphorylated p53 can escape Mdm2-mediated proteolysis [135]. The coordination of individual reactions of posttranscriptional RNA processing within the nucleus and nucleocytoplasmic export of mRNA is also regulated by ubiquitylation [136]. Messenger RNA export proceeds via transient, consecutive interactions between the soluble protein transport receptors named nuclear export factors and the PheGly repeats of certain nucleoporins, proteins forming the nuclear pore complex. This huge protein complex spans the nuclear envelope and possesses the double basket molecular architecture working as a tissue-specific regulator of nucleocytoplasmic transport [137]. Ubiquitin-associated domain of nuclear export factor Mex67 synchronizes recruitment of the mRNA export machinery with transcription [138]. Collectively, these factors form a complex which has been named a transcription export complex [139] necessary for the “gene gating” [140,141]. Since transcription and mRNA processing events are regulated by PTMs of histones, transcription factors, and mRNA processing factors, nuclear export of mRNA is indirectly controlled by several PTMs (mainly ubiquitylation and phosphorylation). Moreover, several recent studies have implicated ubiquitylation in direct regulation of mRNA export making this process even more complex [142,143]. Recent studies have suggested functional links of protein ubiquitylation with the regulation of cellular growth and intracellular signaling. Cullin7 E3 ubiquitin ligase (CUL7) has been identified as a novel player in growth control acting probably through insulin receptor substrate 1. Genetic ablation of CUL7 in mice resulted in intrauterine growth retardation and perinatal lethality underscoring its importance for growth regulation [144]. Similarly, ubiquitylation dependent dynamics in EGF signaling has been characterized using quantitative proteomics [145] revealing possible ubiquitylation cross-talk between the EGF signaling pathway and centrosomaldependent rearrangement of the microtubules. Ubiquitylation of GTPase regulators also makes this PTM a key feature within the most widespread category of GTPase-dependent signaling pathways [146]. In plants the ubiquitin–proteasome system is utilized to modulate nearly every aspect of growth and development [147]. A considerable interest has been raised in the role of ubiquitylation in the immune recognition. In particular, many details have been revealed about the role of interferonstimulated gene 15, one of the most upregulated genes upon type I interferon treatment or pathogen infection. The 17 kDa product of this gene was the first ubiquitin-like modifier

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identified [148] which has a role in the innate immunity against viruses and bacterial infections. Pharmacologists have been attracted by ERAD dependent degradation of cytochromes P450 [149], and their potential use for the modulation of drug metabolism. Worth mentioning are complex interactions between individual forms of ubiquitin-like modifications and between ubiquitylation and certain nonenzymatic PTMs. Ubiquitin and SUMO are small single domain proteins sharing the same fold. However, their surface residues and charge distributions are significantly different pointing to their distinct, even antagonistic roles [150]. Apparent competition between ubiquitylation and sumoylation may be, however, regulated by phosphorylation [28]. Many of these interactions can be assigned to the action of a new class of SUMO-targeted ubiquitin ligases described recently [28]. Ubiquitin ligase has proved critical for mammalian oxygen sensing. Molecular oxygen in the presence of Fe2+ ions causes hydroxylation of proline residue in hypoxia-inducible factor triggering its destruction by E3 ubiquitin ligase, thus regulating the adaptation to changes in oxygen availability on transcriptional level [151–154]. Nondestructive monoubiquitylation has been found in plant metabolic enzyme phosphoenolpyruvate carboxylase (EC 4.1.1.31) at absolutely conserved Lys 628 located proximal to PEP binding site. This ubiquitylation has caused changes in kinetic properties, increase of Km value for PEP and sensitivity to allosteric activators and inhibitors [31]. Our ultimate understanding of the molecular mechanisms of regulated proteolysis is based on structural studies with highly purified enzymes. Recent structural studies revealed how E3 enzymes carry out the final step in target protein ubiquitylation [155]. This elusive mechanism was difficult to capture because of a labile thioester bond linking the E2 conjugating enzyme and ubiquitin. Two structural works were published recently linking ubiquitin through more stable peptide [156] or oxyester [157] bond types, respectively. Further studies are required, however, to address how E2–E3 ubiquitin complexes interact with their protein substrates [158].

3.2. PTMs

Regulation by reversible phosphorylation and related

Modification of a protein molecule by phosphate group alters its charge and therefore potentially also its structure. These changes can concern conformation of active site of an enzyme, binding sites for enzyme effectors, or alter the enzyme's interaction with other proteins. Phosphorylation catalyzed by protein kinases (and partly dephosphorylation catalyzed by protein phosphatases) often means activation (switch on) or inactivation (switch off) of a metabolic pathway. Therefore, these reactions are highly regulated. The activity of a protein kinase could be controlled by signals from outside the cell, by a hormone or growth factor; sometimes a cascade of phosphorylated enzymes alters the activity of a target enzyme. The total set of genome encoded kinases (kinome) can be classified into families and subfamilies according to primary structure of their catalytic domains, sequence similarity, domain structure on amino- and carboxy-terminal part of their molecules, and known biological functions. In human,

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518 kinases were found forming 1.6% of the entire genome pointing to the biological importance of these enzymes. In addition to catalytic domain, 258 of these kinases contain other domains that regulate kinase activity, link them to other signaling modules, or localize the protein to specific subcellular compartment. On the other hand, 260 kinases contain only the catalytic domain, but may be controlled by additional regulatory subunits. Moreover, human kinome contain approximately 10% of kinases without enzyme activity. They may act as kinase substrates and scaffolds for assembly of signaling complexes. These kinases lack at least one of the conserved catalytic residues, but maintain a typical kinase domain fold [159]. Many kinases are associated with serious diseases including cancer, so various types of kinase inhibitors belong to important drugs [160,161]. The plant kinome seems to be very important as well. About 1000 genes for protein kinases were detected in Arabidopsis thaliana representing 4% of its entire genome. The genes encoding proteins similar to those present in animal signaling pathways such as Hedgehog, JAK/STAT, Notch, and receptor tyrosine kinase-Ras pathways are absent in Arabidopsis genome, while protein Ser/Thr receptor kinases, receptor-like kinases, and two component histidine kinases are abundant. The reason why plants have more genes for protein kinases when compared with Ceanorhabditis elegans, Drosophila melanogaster, or Homo sapiens could be a higher number of environmental and intracellular signals to be integrated and used to adapt their metabolism, physiology, and morphology. Plants have evolved their own signaling molecules and their own signal transduction pathways by combining original ligands with receptors and universal kinase cascades [162]. The kinases are divided to groups according to the catalyzed reactions. The kinases transferring phosphoryl group to tyrosine belong to EC 2.7.10.X (receptor kinase EC 2.7.10.1, unspecific EC 2.7.10.2), to serine/threonine EC 2.7.11.X (unspecific EC 2.7.11.1, cAMP dependent protein kinase A EC 2.7.11.11, protein kinase C EC 2.7.11.13, Ca2+ calmodulin dependent protein kinase EC 2.7.11.17, cyclin dependent kinase EC 2.7.11.22, mitogen-activated protein kinase (MAPK) EC 2.7.11.24). Kinases with dual specificity fall into group EC 2.7.12.X (mitogen-activated protein kinase kinase (MAPKK) EC 2.7.12.2), histidine kinases into EC 2.7.13.X. Further details may be found at the web server www.expasy.org. The metabolism of saccharides is closely related to energetic balance of the cell and therefore the regulation of many reactions of saccharide metabolism is of key importance for healthy cell and could be altered under pathological conditions. Namely, the degradation of glycogen is controlled very carefully; the inactive glycogen phosphorylase (EC 2.4.1.1) is activated by phosphorylation catalyzed via phosphorylase kinase (EC 2.7.1.38). This enzyme is an example of protein kinase with very complex structure. It has one catalytic (γ) and three different regulatory (α, β, δ) subunits. The catalytic γ subunit comprises an N-terminal catalytic domain with typical protein kinase fold and C-terminal regulatory domain that binds both δ (calmodulin) subunit and regulatory α subunit. The activity of γ subunit is regulated by the presence and phosphorylation status of regulatory subunits α, β, and δ. The whole molecule forms a hexadecameric complex composed of two lobes, each composed of two αβγδ protomers packed head-to-head with the

lobes associated through interconnecting bridges [163]. Moreover, the activation of phosphorylase kinase is regulated on hormonal and neuronal level. Detailed explanation of these processes is, however, beyond the scope of this review, and the reader is thus referred to the previously published reviews, e.g. to reference [164]. In case of pyruvate kinase (EC 2.7.1.40) the phosphorylation means inactivation of the enzyme, even though the regulation is controlled hormonally by glucagon. Low concentration of blood glucose causes glucagon release, which activates protein kinase A that phosphorylates liver isoenzyme of pyruvate kinase; this mechanism is used for reduction of glycolysis in liver but not in the muscle. The M1 isoenzyme of pyruvate kinase could not be phosphorylated and after adrenalin secretion the glycolysis is activated as energy source for “fight or flight”. Differently spliced isoenzyme called M2 is expressed in highly proliferating tissues and is also regulated by phosphorylation. The overexpression of M2 was found in cancerous cells and seems thus to be a marker for malignant transformation [165]. The dephosphorylation of proteins is catalyzed by protein phosphatases. An interesting example of bifunctional protein, which fulfills both kinase and phosphatase function, is 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase (EC 2.7.1.105/EC 3.1.3.46). The phosphorylation of Ser residue near the N-terminus catalyzed by protein kinase A inhibits the kinase activity and activates the phosphatase activity. This enzyme catalyzes the formation or degradation of fructose-2,6-bisphosphate, an important regulator of glycolysis [166–168]. The phosphorylation cannot only change activity of an enzyme significantly, but also modulate enzyme activity in a fine manner, or modulate the sensitivity to enzyme effectors. In plants the effect of phosphorylation on phosphoenolpyruvate carboxylase is well documented. The phosphorylation of a conserved Ser residue near the N-terminus is catalyzed by the smallest known protein kinase: PEPC kinase, which consists only of a core kinase domain without any regulatory domain or subunit. The activity of PEPC kinase is controlled by changes in rates of PEPC kinase synthesis and degradation. The phosphorylated form of PEPC often differs from the dephosphorylated form in kinetic properties such as maximal reaction rate, affinity to PEP, and sensitivity to allosteric effectors that relieve its inhibition by malate and enhance its activation by glucose-6-phosphate [31,169,170]. Phosphoryl group in phosphohistidine may be linked to N1 or N3 nitrogen of imidazole ring; low stability of phosphoramines causes dephosphorylation in acidic microenvironment in the cell without phosphatase treatment (i.e. in nonenzymatic manner) [17]. Phosphorylated histidine is localized in active site of some enzymes involved in transfer of high-energy phosphoryl groups to other compounds via a phosphohistidine intermediate. An obvious example is phosphoglycerate mutase (EC 5.4.2.1); 3-phosphoglycerate is attached to the phosphorylated enzyme (containing phosphohistidine) and thus formed 2,3-bisphosphoglycerate intermediate is subsequently hydrolyzed to 2-phosphoglycerate and phosphorylated enzyme [166]. Pyruvate, orthophosphate dikinase (EC 2.7.9.1), a key enzyme in C4 photosynthesis (also found in C3 plants and some

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bacteria), is regulated in very interesting manner. A histidine residue located in active site is involved in transfer of phosphoryl group from ATP to pyruvate with phosphohistidine intermediate and phosphoenolpyruvate is formed. Furthermore this enzyme is subjected to diurnal dark–light regulation via reversible phosphorylation of a Thr residue. Both phosphorylation (inactivation) and dephosphorylation (activation) is catalyzed by one bifunctional regulatory protein which uses unusual mechanism of reaction: the phosphoryl group comes from ADP instead from ATP, while the dephosphorylation is catalyzed by orthophosphate-dependent (and pyrophosphate generating) phosphotransferase activity of regulatory protein instead of hydrolysis by phosphatase [171]. In order to adapt to their fluctuating environment bacteria, yeast, and plants have evolved a two-component system consisting of a sensor histidine kinase and a response regulator. Sensor histidine kinases consist of an N-terminal domain that detects various stimuli such as pH, temperature, chemoattractants, or osmolarity, a histidine containing phosphoacceptor domain, and an ATP-binding kinase domain. Most of the response regulators consist of an N-terminal receiver domain and an effector domain. The conserved aspartate residue of the response domain receives the phosphoryl group from the histidine residue of the sensor histidine kinase and this phosphorylation regulates the activity of the C-terminal effector domain. Most response regulators have a DNA-binding effector domain and function as transcription factors, and a few have effector domains with enzymatic activity. The two-component system known as His-Asp phosphorelay can be arranged as multistep phosphorelay system, which additionally requires both an intermediate response domain and a histidine-containing phosphotransfer domain (His-Asp-His-Asp), although a special two step system (His-Asp-His) is also known [172,173]. 14-3-3 proteins mediate the effects of certain protein kinases through their ability to bind well-defined phosphoserine or phosphothreonine containing peptide motifs [174], which finally results in modification of the activity, stability, subcellular localization, or interaction capability of the client protein [175]. The fact that 14-3-3s form clamp-like dimers with each monomer capable of binding a phosphopeptide within an amphipathic groove immediately suggests that 14-3-3s function as an intermolecular bridge linking two different phosphoproteins [176]. More commonly, a 14-3-3 dimer seems to engage with two tandem phosphorylated sites in the same protein [174]. Research in recent years has revealed an impressive list of putative 14-3-3 targets in both animals and plants as reviewed in ref. [177]. Several recent high-throughput proteomic studies have suggested 14-3-3s acting also as a key regulatory components of signaling cascades, in particular phytohormone mediated processes. 14-3-3 protein also regulates activity of a key enzyme in nitrogen assimilation: nitrate reductase (EC 1.6.6.1). Nitrate reductase is composed of three domains each containing a cofactor and thus transferring electron from NAD(P)H to nitrate. The activity of nitrate reductase is regulated by phosphorylation of Ser residue localized in the linker between the heme and molybdenum cofactor domains. 14-3-3 protein can bind to this phosphorylation site, cause conformational changes and thus reversibly inhibit the enzyme. Mg2+ ions participate in this regulation process [178,179].

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O-GlcNAcylation is the addition of β-N-acetyl-D-glucosamine to serine or threonine residues of nuclear and cytoplasmic proteins. O-linked N-acetylglucosamine (O-GlcNAc) was not discovered until the early 1980s and still remains difficult to detect and quantify [22]. Nonetheless, O-GlcNAc is highly abundant and cycles on proteins with a timescale similar to protein phosphorylation. O-GlcNAc occurs in organisms ranging from some bacteria to protozoans and metazoans, including plants and nematodes, up the evolutionary tree to man. O-GlcNAcylation is mostly observed on nuclear proteins, but it occurs in all intracellular compartments, including mitochondria. Recent glycomic analysis has shown that O-GlcNAcylation has surprisingly extensive cross-talk with phosphorylation where it serves as a nutrient/stress sensor to modulate signaling, transcription, and cytoskeletal functions [180].

3.3. Regulation by modifying cysteine residues under dynamic redox state Specific redox proteome and redox environment is actively maintained using different strategies within individual cellular compartments. It defines the formation of cysteine thiolates and disulfide bonds which can serve structural, catalytic, or signaling functions [181]. The cytoplasmic redox potential in most animal cells is controlled mainly by glutathione found at concentration around 5 mM, less than 10% of which is present in its oxidized form. The redox potential set by the glutathione system and other redox buffers defines the formation of disulfide bonds — the most common PTM of cysteine during which the pair of cysteines is oxidized and bonded. Analysis of the human genome predicts that more than half of the proteins secreted into ER and transported further by secretion machinery acquire disulfide bonds [182]. Disulfide bonds play important roles in protein structure, catalysis, and signaling. Disulfide bonds can be observed in the extracellular parts of proteins more often than in intracellular ones enhancing the rigidity and stability of receptors or enzymes occurring in the fairly damaging extracellular environment. In this respect, disulfide bonds just lower the entropy by decreasing the number of possible conformations and thus improve the durability of the protein. However, disulfide bonding can also be essential for proper protein folding during which the mispairing of cysteines can lead to misfolding with the loss of protein functionality, or even to protein aggregation. Moreover, disulfide bonds can participate in metal ion coordination or cofactor binding [183]. The most important catalytical role of disulfide bonds is arguably played in oxidoreductases that are involved in creation or isomerization of other disulfide bonds. The original Anfinsen experiment led the scientific community to an erroneous conclusion that disulfide bonds are formed spontaneously in vitro as well as within the living cell [184] requiring only electron acceptor such as oxygen. This dogma was questioned in 1991 with the discovery of the first enzyme in Escherichia coli periplasm required for disulfide bond formation [185]. Disulfide bonds can be reduced, oxidized, or isomerized during protein folding. An oxidoreductase catalyzing the electron transfer is necessary for all these processes. These oxidoreductases typically comprise Cys-Xxx-Xxx-Cys sequence in the active site. However, various factors determine whether the enzyme is oxidase or reductase in vivo, namely the relative stability of

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oxidized/reduced state of the enzyme, the chemical environment of the compartment where the enzyme is localized, and the amino acids occurring in the vicinity of and within the catalytical motif [181]. The pKa of cysteine thiol is normally around 8.5. It means that cysteine thiols are considerably stable at the physiological pH and it is necessary to stabilize the more reactive thiolate form by ambient basic amino acids, usually arginines or lysines. The latter can lower the pKa of protein incorporated cysteine thiols and influence their ability to form disulfides. While oxidases are more abundant in compartments possessing oxidative environment such as ER influenced by dehydroascorbic acid, reductases are more common in the cytoplasm. Thioredoxin A can serve as an example as it behaves as a reductase when natively expressed into the cytoplasm with catalytic cysteines kept reduced while it can behave partially as an oxidase when expressed into the oxidative periplasm [186]. Another important role of cysteines is in oxidative or nitrosative stress signaling since the oxidation or reduction of protein thiols is the main way how the reactive oxygen/nitrogen species initiate signaling pathways [187]. These signals can be transmitted by oxidation/reduction of the reporter disulfide bond as in the case of OxyR initiating the transcription of oxidative stress response proteins after formation of disulfide bond [188]. Alternatively, signaling may involve other important cysteine PTMs such as S-nitrosylation or S-glutathionylation [189]. S-glutathionylation can be considered as another type of disulfide Cys-Cys bridge with protein cysteine on one side and low molecular mass partner (i.e. glutathione) on the other. The exact mechanism for this modification is not well understood. Nevertheless, several potential pathways were suggested and discussed [190]. Protein tyrosine phosphatases (EC 3.1.3.48) may serve as best examples of regulatory oxidation, where the key cysteine residues are initially oxidized to sulfenic acid. In case of PTP1B this step is then followed by glutathionylation [191]. Formation of sulfenic acid is generally the first step of cysteine thiol modification. This oxidation is mediated by H2O2, various organic peroxides, or hypohalous acids and may be either stabilized or followed by formation of other reversible (disulfides, S-nitrosyles, sulfenamides) or irreversible (sulfinic and sulfonic) oxidation products [192]. The maintenance of proper redox state in the cell is very important especially under conditions of oxidative stress. Thioredoxin system is present in all types of organisms from bacteria to plants and animals. Thioredoxins are small proteins with characteristic fold and conserved sequence TrpCysGlyProCys in active site, which catalyze protein disulfide reduction while turning into oxidized state. The reduction of oxidized thioredoxins is catalyzed by thioredoxin reductases using NADPH as reducing power. In plants there is a large number of thioredoxins localized in chloroplasts. Chloroplasts' thioredoxins are reduced by catalysis of ferredoxin dependent thioredoxin reductase (EC 1.8.7.2) using reduced ferredoxin as reducing power formed during photosynthetic electron transport. This ferredoxin–thioredoxin system is important for activation of target enzymes, which are often enzymes of Calvin cycle [193,194]. Recently, a novel type of NADP thioredoxin reductase localized also in chloroplasts was discovered. It acts as a bifunctional enzyme with N-terminal thioredoxin reductase domain and C-terminal thioredoxin domain. Due to a high affinity of NADP thioredoxin reductase

to NADPH, this system might participate as an alternative electron donor for the chloroplast detoxification system especially during darkness when the source of NADPH is oxidative pentose phosphate pathway [194–196].

3.4. Regulation of secreted and membrane-associated enzymes by glycosylation The role of glycosylation in maintenance of enzyme solubility, stability, folding, and assembly into fully active complexes has already been mentioned in the introductory Section 2.1.1 [11,15,197]. Novel modes of glycosylation are being continuously discovered e.g. O-mannosylation conserved from bacteria to humans [198] necessary for normal embryonic development in zebrafish [199] and compromised in patients with Walker–Warburg syndrome [200]. Sensitive modern techniques allow to analyze individual glycopeptides of a human glycoprotein produced in different tissues providing novel insights into tissue-specific and site-specific glycosylation in normal human tissues [201]. Details of specific roles of enzyme glycosylations in many production organisms are emerging fast. The presence of multiple glycosylations was necessary for proper folding and transport of carbonic anhydrase (EC 4.2.1.1) and ENDO2 endonuclease in A. thaliana (EC 3.1.30.1) [202,203], catalytic and biochemical properties of β-D-glucuronidase (EC 3.2.1.31) from Penicillium purpurogenum [204], and human prostatic acid phosphatase (EC 3.1.3.2) [205]. Moreover, glycosylation is also required for full enzyme activity (although not oligomerization) of membrane acetylcholinesterase (EC 3.1.1.7) in HEK293T cells [206], human α1,3-fucosyltransferase IX (EC 2.4.1.214) [207], and for the autocatalytic cleavage of human γ-glutamyl transpeptidase (EC 2.3.2.2) [208]. Finally, glycosylation modulates the action and signaling of membrane receptors as it has proved necessary for sustaining the function of membrane Mid2p sensor in yeast Saccharomyces cerevisiae [209], for modulation of FGF and TGF-β induced cell signaling [210,211], for stability and function of collagen adhesin of Aggregatibacter actinomycetemcomitans [212], and for binding of H1N1 virus to target cells through its lectin-type adhesin [213]. Protein glycosylation depends on the concerted action of glycosyltransferases (EC 2.4.1.X) and glycosidases (EC 3.2.1.X) [11] and is exquisitely sensitive to the physiological conditions of the producing cell. Therefore, glycosylation marks left on the secreted glycoproteins can provide noninvasive and sensitive information about the whole range of normal as well as pathological processes. Vanhooren and colleagues reported on three specific changes in N-glycome related to aging that affects undergalactosylated biantennary, biantennary, and core α-1,6-fucosylated biantennary glycans [214]. Their findings have been further expanded using (kl/kl) and (dw/dw) mice having shortened or prolonged lifespan, respectively. Increase in the activity of fucosyltransferase VIII (EC 2.4.1.68) responsible for core fucosylation could be linked to altered IGF-1 signaling providing an interesting connection between cell surface and intracellular determinants of aging [215]. Extensive efforts have been made to use changes in dynamic glycosylation for diagnostic purposes, and, in particular, for specific diagnosis of tumors and prognosis in tumor patients [216]. In this respect, there is a significant hope that

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powerful modern glycoproteomic and glycomic technologies will be able to suggest specific tumor markers with a high informative value with respect to tumor malignancy and progression based on blood samples without a need to resort to more invasive methods [217–220]. The recurring theme here has been in the integration and mutual interconnections between cell surface and intracellular processes. This has been recently emphasized by a link between N-glycosylation and phosphorylation of anaplastic lymphoma kinase (EC 2.7.10.1) [221], DNA methylation and glycosylation [222], and the possibility to resuscitate wild-type p53 expression upon disruption of ceramide glycosylation [223]. Finally, our increasing abilities to understand the complexity of protein glycosylation as well as the availability of technologies allowing to produce proteins bearing defined glycosylation resulted in considerable practical achievements in the area of glycoprotein therapeutics and carbohydrate-based vaccines [11]. In particular, defined glycosylation has proved critical for immune properties of therapeutical monoclonal antibodies [224] which are now produced using contemporary glycoengineering technologies [11,225]. Glycoproteins as well as other glycoconjugates are becoming important antitumor vaccines exploring the potential of many chemical modifications that might be introduced into the carbohydrate moieties [226,227]. Using today's technologies of protein engineering and glycoengineering, carbohydrate components can be added to proteins and enzymes that do not bear them naturally with the aim to improve their natural therapeutical or catalytical properties. Important examples of this approach include engineered insulin preparations bearing sialyl-α-2,6-lactosyl sequences that increase their serum half-life [228], or N-glycosylated porcine pepsin having unique enzymatic properties [229].

3.5. Regulation of enzyme activities by glycation and other nonenzymatic PTMs Nonenzymatic glycation begins with Schiff base formation between reactive glucose and ε-amino group of lysine followed by Amadori rearrangement and formation of other dicarbonyl compounds [230] that participate in a number of follow-up reactions leading to AGE (Fig. 1). One such pathway proceeds through Amadori dion (1-amino-4-deoxy-2,3-dion) and Amadori en-dion (1-amino-4-en-2,3-dion), the double bond of which can be attacked by other Lys ε-amino group resulting in protein cross-linking. Other AGEs can be formed by α-dicarbonyl compounds generated from free sugars or protein-bound carbohydrates, of which glyoxal, methylglyoxal, and 3-deoxyglucose are the most prominent examples (“reactive carbonyls”). Thus, glyoxal leads to the formation of Nε-carboxymethyllysine, methylglyoxal-derived Lys dimer, and Arg-derived imidazolone. Similarly, methylglyoxal gives rise to Nε-carboxyethyllysine, methylglyoxal-derived Lys dimer, arginine-derived methylimidazolone and argpyrimidine, while 3-deoxyglucosone initiates the formation of even more complex end products such as 4-deoxyerythritol modified glyoxal Lys dimer, 4-deoxyerythritol derivative of Arg imidazolone, and pyrraline (5-hydroxymethyl-1-alkylpyrrole-2-carbaldehyde). Additional identified products involve pentosidine ((2S)-2-amino-6-[2-[[(4S)-4-amino5-hydroxy-5-oxopentyl]amino]-4-imidazo [4,5-b]pyridinyl] hexanoic acid, crossline ((3R,4S)-3,4-dihydroxy-5-[(1S or 1R,2S,3R)-

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1,2,3,4-tetrahydroxybutyl] -1,7-bis[6- (N-acetyl-L-norleucyl)]-1, 2,3,4-tetrahydro-1,7-naphthyridinium), and vesperlysine (6-hydroxy-1,4-di{6-(L-norleucyl)}-1H-pyrrolo[3,2-b]pyridinium) [231]. Another source of reactive carbonyl compounds is lipooxidation [102], providing reactive lipid species (lipid electrophiles) in the form of unsaturated alkenals. The most prominent examples of these compounds are unsaturated lipids derived malondialdehyde and 4-hydroxy-2-nonenal, as well as arachidonate derived A2 isoprostane, 12-nitroarachidonate, leukotriene A4 and 15-deoxy-Δ12,14-prostaglandin J2 [92]. These electrophiles react with nucleophilic reaction centers in enzymes (most reactive of which is Cys followed by His and Lys) eventually leading to advanced lipooxidation end products (Michael adducts) such as 4-hydroxy-2-nonenal derivatives of Cys, His, and Lys [2]. These adducts formed with abundant small peptides GSH, carnosine, and anserine serve as early markers of oxidative stress in excitable tissues [232]. Extensive cross-linking through AGEs leads to vascular stiffness [231], one of the most critical factor of physiological aging exacerbated in diabetes and leading to its major complications such as diabetic retinopathy [233], diabetic nephropathy [234], and preterm birth [235]. Vascular receptor for AGEs belonging to immunoglobulin superfamily has been recently identified to represent a fundamental mechanism of signaling danger to the vulnerable vasculature [236]; soluble form of this receptor is now becoming an important prognostic marker in chronic inflammatory diseases [237]. Decoration of the polypeptide chain by inert carbohydrate extensions occurs essentially at the moment of their entry into the oxidative environment in ER pointing to the general protective role of carbohydrates against enzyme aging caused by chemical radicals, ROS, and other nonenzymatic PTMs [12]. Considering the universal occurrence of D-glucose (both intracellular and extracellular) as a major source of energy in wide range of organisms, the effect of glycation on a wide range of enzymes would merit more extensive investigations. Glycation-induced conformational changes and their effect on the activity of human paraoxonase 1 (EC 3.1.1.2), an important enzyme with a protective effect against LDL oxidation thus reducing the blood level of cholesterol with antiatherogenic effects, were studied by modeling, docking, and molecular dynamic simulations [238]. Mitochondrial catalase (EC 1.11.1.6) has been identified as the only antioxidant enzyme significantly glycated in old rats; treatment of catalase with the glycating agent fructose led to significant time-dependent inactivation of the enzyme, while methylglyoxal had no noticeable effect [239]. Plasmatic β2-glycoprotein I was shown to be effectively glycated by blood glucose leading to AGE formation and activation of dendritic cells that upregulate AGE receptor, which becomes eventually one of the major processes leading to chronic endothelial dysfunctions [240]. Since enzyme glycation and the formation of AGEs plays such a profound role in pathology of several principal human diseases, attention has been paid to minimization of AGE food intake [241] and to efficient inhibitors of AGE formation as therapeutics [230].

3.6. Multiple posttranslational modifications regulating enzyme activities Proteomic identification of many PTMs during the recent years including multiple modifications residing on a single

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protein or even single amino acid residue led to questions concerning the mutual interplay between individual PTMs and their effects on the final occurrence, activity, and cellular localization of the target enzymes. In this chapter we describe different modes of positive and negative feedback relations between individual PTMs. Enzymes and proteins regulating and maintaining chromatin consists of histones, transcription factors, scaffold proteins, and transcription enzymes. In order to coordinate their activities, multiple PTMs are used. The spectrum of the revealed histone modifications is shown in Fig. 4. These include lysine acetylation, lysine and arginine methylation, serine and threonine phosphorylation, and lysine ubiquitylation and sumoylation. These modifications occur primarily within the N-terminal tails protruding from the surface of the nucleosome [242]. Histone modifications are proposed to affect chromosome function through two distinct mechanisms: (a) alteration in the electrostatic charge of the histone resulting in structural change and differences in binding to DNA and (b) introduction of modification marks forming binding sites for recognition modules of various accessory proteins. The existence of these recognition mechanisms forms a basis of the “histone code” hypothesis [243]. Initially, individual histone modifications have been identified by mass spectrometry using middledown strategies [244]. Recently, histone modifications and variants have also been analyzed at the whole genome level by the ENCODE consortium using antibody-based ChIP-on-chip analyses developed by Life Technologies [245]. The individual analyzed marks included those for active regulatory elements associated with promoters/transcription starts (H3Lys4Me3, H3Lys9Ac, H3Lys27Ac), marks associated with enhancers and other distal elements (H3Lys4Me1), marks with preference for the 5′end of genes (H3Lys9Me1, H4Lys20Me1), marks associated with transcribed portions of genes (H3Lys36Me), repressive marks associated with constitutive heterochromatin (H3Lys9Me3), and repressive marks established by polycomb complex (H3Lys27Me3). In combination with other analyses of

chromatin, these experiments revolutionized our understanding of the human genome, and allowed to assign biochemical functions to as much as 80% of the DNA sequence (not just to its tiny portion representing protein coding genes) [245]. Histone acetylation and phosphorylation are transient events associated with the initiation and repression of transcription [246]. High resolution X-ray crystallography revealed the structure of nucleosome including flexible amino terminal tails (the sequences of which are shown in Fig. 4) and the globular histone octamer (H2AH2BH3H4)2 forming the nucleosome core [247]. Although it was known that histone acetylation marked actively transcribed genes, the mechanistic nature of the role of this PTM was unclear. Chemically, acetylation at lysine residues at amino terminal tails of histones (Fig. 4) neutralizes their positive charge decreasing their ability to bind DNA and increasing their accessibility to transcription factors and coactivators to chromatin. Functionally, an explanation came from the discovery that histone acetyltransferases are transcriptional coactivators [248], while histone deacetylases behaved as transcriptional repressors [249,250]. Thus, these studies not only confirmed histone acetylation as the key regulator of transcription, but also revealed the mechanism of recruitment of histone acetylases/deacetylases to the promoters of individual genes during transcriptional activation. Well documented examples of these activities include histone H3 and H4 acetyltransferase complexes, both recruited by certain acidic activators [251]. On the other hand, the same repressor can recruit different histone deacetylase complexes [252]. Moreover, acetylation of nonhistone proteins regulates cellular proliferation and survival, intracellular trafficking, and transcription [253]. In particular, acetylation of multiple substrates within the nuclear receptor signaling pathways give rise to an acetylation signaling network from the cell surface to the nucleus [254]. The recent finding that NAD+-dependent histone deacetylases, the sirtuins, can deacetylate nuclear receptors reveal a new complexity in regulation of these receptors in which local concentrations of nicotinamide coenzymes can

Fig. 4 – Schematic representation of PTMs in N-terminal tails of histones. According to [17,337]. Please cite this article as: Ryšlavá H, et al, Effect of posttranslational modifications on enzyme function and assembly, J Prot (2013), http://dx.doi.org/10.1016/j.jprot.2013.03.025

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regulate nuclear receptor physiology [254]. Furthermore, recent global proteomic analyses revealed large number of acetylated proteins in the cytoplasm and mitochondria including key metabolic enzymes being regulated through enzyme activation or inhibition, enzyme stability, and enzyme complex formation [255], thus pointing to a more widespread importance of this PTM beyond chromatin regulation and signaling in the nucleus. After the discovery of the first histone methyltransferase (EC 2.1.1.43) [256], an extended family of these enzymes has been characterized [257] featuring a SET (Su(var)3–9, Enhancer of Zeste, Trithorax) domain. The enzymes placing the lysine histone marks along the chromatin are well defined, however, not so much is known about protein arginine N-methyltransferases (EC 2.1.1.125) that are (similar to acetyltransferases) targeting a broad range of cellular proteins [258]. In contrast to yeast, mammals have a greatly expanded histone methylation profiles [259]. Mouse knockout models demonstrated important roles of methyltransferases in regulation of transcription, maintenance of genome integrity, and development [260] accelerating an effort to identify specific inhibitors of these enzymes as pharmacological targets in cancer and other diseases [261]. Histone methylations act as nucleation sites for binding of effector proteins. The crystal structures of effector proteins containing ankyrin repeats, chromodomain, MBT repeats, plant homeodomain finger, and double tudor domains responsible for detection of histone marks have been solved [262]. The proximity of lysine and arginine residues within the histon tail suggests a cross-talk between the adjacent modifications of these amino acids [263]. Histone lysine methylation can be dynamically regulated by histone demethylases (EC 1.14.11.X). The first class of histone demethylases are mammalian amine oxidases (EC 1.14.11.X) which use FAD as a cofactor and remove Me1 and Me2 modification states [264], complemented by a second class containing Jumonji-C domain in their catalytic core. The latter enzymes are iron and 2-oxoglutarate dependent oxygenases targeting all three histone lysine methylation states [265]. Recently, a member of this family was shown to reverse histone arginine methylation [266]. Interestingly, histone lysine demethylation reaction itself produces reactive byproducts including formaldehyde and H2O2 [267]. Surprisingly, DNA damage caused by H2O2 appears to be functionally important for efficient transcription of estrogen receptor regulated genes because single strand DNA breaks induced during DNA repair help to facilitate DNA bending permitting more efficient RNA polymerase III (EC 2.7.7.6) loading [268]. On the other hand, formaldehyde produced during lysine demethylation appears to be scavenged by NAD- and glutathione-dependent formaldehyde dehydrogenase (EC 1.1.1.284) [269]. Poly-ADP-ribosylation is another recently studied PTM important for chromatin structure and DNA metabolism, cell division, proliferation, differentiation, or death, viral infection, spermatogenesis, and other processes [270]. Its role in DNA damage response is particularly well documented [271]. An interesting example of complex interplay between protein sumoylation and protein acetylation machinery is provided by histone deacetylases which remove the acetyl groups from target proteins and are also reported to function as SUMO E3 ligases for some substrates. Furthermore, some histone deacetylases can be sumoylated themselves as well,

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and can noncovalently bind other sumoylated proteins, pointing to complex interrelationships between these pathways [272]. However, the spectrum of PTMs identified in highly homologous histone deacetylases 1 and 2 is even much broader. This variety in PTMs is necessary for our understanding of both redundant and highly specific roles that these enzymes have in different cell types in response to different signaling pathways. PTMs identified so far in these enzymes involve both enzymatically catalyzed and nonenzymatic modifications such as phosphorylations, acetylations, ubiquitylations, sumoylations, nitrosylations, and carbonylations all pointing to complicated cross-competition creating a rational code for context dependent regulation of these enzymatic regulators [4]. A complex interplay between histone PTMs (Fig. 4) and PTMs of RNA polymerase II, critical for the regulation of transcription of all protein coding genes, have been described by Fuchs et al. [273]. In particular, these interactions concern those occurring between histone H3Lys4 methylations, histone H2B ubiquitylations, and phosphorylations of the C-terminal domain of RNA polymerase II, as well as those that go on between methyltransferase Kmt3 (EC 2.1.1.43), responsible for methylation of Lys36 in histone H3, and phosphorylation of the above mentioned C-terminal domain of RNA polymerase II. Obviously, our complete description of these numerous interactions should eventually lead to understanding of the molecular mechanisms of chromatin remodeling and to a unified theory of gene expression as suggested by Orphanides and Reinberg [274]. Deubiquitylating enzymes are also tightly controlled and regulated by multiple PTMs including monoubiquitylation, sumoylation, acetylation, and phosphorylation as a part of multiple feedback loops operating in living cells [32]. PTMs play a crucial role in the coordination of cellular response to DNA damage. Maintenance of the integrity of the genome is considered to be, together with the ability to produce metabolic energy, the most vital characteristics of every living cell. Recent evidence again suggests a complex interplay between numerous PTMs in this area that include (but are not limited to) phosphorylation, ubiquitylation, acetylation, and sumoylation that combine to link DNA damage signal to cell cycle arrest, DNA repair, or apoptosis. The p53 tumor suppressor protein plays a key role in coordination of these processes acting predominantly as a transcription factor regulating more than a hundred of target genes [275]. The p53 protein is composed of 6 domains bearing multiple phosphorylations on Ser/Thr residues and ubiquitylations, neddylations, and methylations on Lys residues [276]. Moreover, many enzymes delivering these multiple PTM combinations have been detected, including many kinases, ubiquitin E3 ligases, and methyltransferases activated by UV light, γ-irradiation, DNA damage, or ER stress. Functional importance of individual PTMs and their combinations have been extensively investigated using biochemical assays and cellular tests giving rise to more than forty thousand publications on this subject. However, these in vitro tests were not fully supported by in vivo results obtained using mice expressing numerous mutant p53 forms. Nevertheless, a more definitive picture is now coming out from mouse genetics studies indicating the critical role of p53 phosphorylation in the stabilization of p53 in response to DNA

Please cite this article as: Ryšlavá H, et al, Effect of posttranslational modifications on enzyme function and assembly, J Prot (2013), http://dx.doi.org/10.1016/j.jprot.2013.03.025

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damage, most likely by interfering with MDM2 binding and subsequent proteasomal degradation [276]. Moreover, additional p53 functions are related to its translocation to outer mitochondrial membrane where it interacts with the proand antiapoptotic members of Bcl-2 protein family [275]. Cyclin-dependent kinases that restrict DNA replication origin firing to once per cell cycle by preventing the assembly of pre-replication complex outside G1 phase cooperate with APC/C-dependent proteolysis [277]. The cyclin T/cyclindependent kinase 9 complex (EC 2.7.11.22 and 2.7.11.23) phosphorylates the C-terminal domain of the large fragment of RNA polymerase II which is a hallmark of transition from initiation to elongation of transcription. However, this kinase complex is itself modified by phosphorylation, ubiquitylation, and acetylation [278]. Serine/threonine protein kinase Akt orchestrates many biological functions such as cell proliferation, survival, metabolism, cell migration, and metastasis. The activity of this critical kinase is regulated through the phosphorylation of Thr308 and Ser473, but also by ubiquitylation as recently proved for cancer-associated Akt mutant shown to display enhanced phosphorylation [279]. Phosphorylation of Skp2 by Akt triggers its association with other polypeptides and formation of SCF E3 ligase activity [280] promoting its cytoplasmic location and impairing its own proteolytic destruction [281]. Moreover, molecular events involved in circadian rhythms signaling are dependent not only on phosphorylations but also destructive as well as nondestructive ubiquitylations [282]. N-glycosylation has been proved critical for protein quality control in ER, namely for ubiquitylation of unfolded glycoproteins by SCFFbx2 ubiquitin E3 ligase complex for subsequent translocation to cytosol end degradation by ERAD system [283]. The structural basis for this quality control pathway has been explained from the solved three dimensional structure of the saccharide-binding domain of the ubiquitin E3 ligase complex recognizing Man3GlcNAc2 oligosaccharide [284]. The SCFFbx2 ubiquitin E3 ligase complex is physiologically very important for the degradation of β-secretase (EC 3.4.23.46) which has a central role in β-amyloidogenesis initiating neurodegenerative disorders such as Alzheimer's and Parkinson's diseases [285]. Similarly, there exists a significant cross-talk between protein phosphorylation and protein sumoylation as well as between protein sumoylation and protein acetylation both in terms of coregulation of substrate proteins and in terms of cross-regulation of both PTMs [28]. For instance, sumoylation can regulate phosphorylation dynamics through modification of several kinases such as focal adhesion kinase (EC 2.7.10.2) [286], glycogen synthase kinase 3b (EC 2.7.11.1), homeodomain-interacting protein kinase 2 (EC 2.7.11.1) [287], and extracellular signal-regulated kinase 5 (EC 2.7.11.1) [288]. Modern techniques of contemporary proteomics have been detecting multiple PTMs on the key cellular enzymes and multienzyme complexes, although the functional relevance of these modifications has not been fully evaluated so far. Several PTMs such as phosphorylations, lysine methylations, lysine acetylations, oxidations, and nitrations have been detected in the ATP synthase complex (EC 3.6.3.14) [289] some of which may serve as sensitive markers of the functional state of the heart, e.g. S-nitrosylation and Tyr

nitration following ischemia/reperfusion [290]. In plants, PTMs of the key enzyme of photosynthesis ribulose-1,5bisphosphate carboxylase (EC 4.1.1.39) have been analyzed. These PTMs include N-terminal deformylation, proteolytical cleavage of N-terminal Met and Ser, acetylation of the newly formed Pro3 N-termini, and Lys14 trimethylation (all in the large subunit of this enzyme), together with signal peptide cleavage and N-terminal methylation in its small subunit [291]. The functional significance of these individual PTMs remains unclear despite extensive identification of enzymes responsible for these modifications. For functional photosynthesis ribulose-1,5-bisphosphate carboxylase requires activation by carbamylation of the ε-amino group of Lys localized in the active site by a CO2 molecule distinct from the substrate CO2. The carbamylated Lys is stabilized by the binding of the Mg2+ ion to the carbamate [292,293]. Glutamate-cysteine ligase (EC 6.3.2.2) is an enzyme catalyzing the rate-limiting step in the synthesis of glutathione, a key peptide regulating the cellular redox environment. While PTMs occurring in the catalytic subunit of this enzyme were shown to have only minor effects on its catalytic activity, subtoxic concentrations of H2O2 and other oxidants increase enzyme's activity without stimulation of de novo proteosynthesis [294]. Similar plant enzyme is regulated by a unique posttranslational redox switch mechanism based on modification of redox-sensitive intramolecular bridges [295]. The widespread occurrence of acetylation in key metabolic enzymes and their complexes mentioned above also belong to the growing list of PTMs pertinent to key cellular enzymes [244]. Finally, a completely different set of PTMs is pertinent to the function of the second most important fibrous cellular structures after chromatin, the cytoskeleton. Microtubules possess unique PTMs in the form of polyglutamylation and polyglycylation acting in a reciprocal way (i.e. polyglycylation acting as a suppressor of polyglutamylation) [296,297]. Polyglutamylation is involved in the functional adaptation of microtubules since it occurs in functionally important carboxy terminal tails [296]. Extensive proteomic characterization of this PTM and enzymes responsible for its synthesis and degradation is ongoing [298]. Additional PTMs characteristic for microtubules are discussed in Section 3.6. However, worth mentioning here is the acetylation of lysine regulating all three major types of filaments, namely microfilaments (actin filaments), intermediate filaments, and microtubules [299]. As the most abundant cellular proteins, cytoskeletal proteins are highly prone to nonenzymatic modifications (glycations, carbonylations, tyrosine nitrations) that in turn bear a significant influence on their functional properties [300–302]. Thus, the recent identification of numerous PTMs in tubulins, together with the characterization of tubulin modifying and tubulin demodifying enzymes, revealed key roles of tubulin PTMs in the regulation of organization and dynamics of microtubules and in the regulation of motor proteins associating with tubulin [3,303,304]. Similarly, some pathological mutations in huntingtin, the elongated polyglutamine tract of which is responsible for the neurodegenerative disorder Huntington disease, could be related to sites of numerous PTMs in this protein consisting of phosphorylation, sumoylation, ubiquitylation, acetylation, palmitoylation, and proteolytic cleavage [305].

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4. Practical aspects of enzymes posttranslational modifications As discussed in previous chapters, both enzymatic and nonenzymatic PTMs can have a significant impact on enzymes' structure or function. While current techniques of MS analysis are beginning to be able to resolve the vast and highly heterogeneous posttranslationally modified proteome, they have still limited capabilities when an individual structure– function relationship of a given protein is under question on a molecular level. Our understanding of this relationship (as already pinpointed in Section 3.1) is based on structural studies with highly purified enzymes, in most cases enzymes prepared by various techniques of recombinant protein expression. In this respect, PTMs might represent a serious hurdle along the process of enzyme structure determination and are therefore considered from a completely opposite point of view. Since for most techniques of structural biology homogeneity of the studied material (i.e. of the purified enzyme) is a necessary key to eventual success, heterogeneous and inherently incomplete PTMs may represent a problem. However, this inevitably leads to a paradox — solved structures of artificially prepared homogeneous proteins might not correspond in all details to their native counterparts from living organism. At least partial solution of this contradiction might be granted by molecular modeling coupled to molecular dynamics simulations with protein structures in silico decorated with PTMs identified by MS studies from native raw PTM-proteome. The two most widely used techniques of structural biology that are able to provide structural details at atomic resolution, namely NMR and X-ray protein crystallography, require both a considerable amount of pure protein sample. While isolation from natural source might be convenient in certain cases, the high demand for protein sample is solved most commonly by protein overproduction in a suitable expression host. Also, the possibility to introduce changes to the studied enzyme (be it a mutation or an affinity tag) together with at least some chance to control PTMs all represent significant advantages of this approach. Since the protein molecule needs to be labeled with 15 N/13C isotopes for NMR measurements, the choice of expression host is for practical reasons limited almost exclusively to E. coli. Given the rather limited range of PTMs found in bacteria, this expression system is well suited for protein crystallography, too. On the other hand, the same limitations of prokaryotic protein translation machinery might be detrimental to the yield of expressed protein–disulfide bonds might not be formed, enzymes' cofactors might not be bound properly, protein might form insoluble denatured aggregates, etc. For eukaryotic proteins and especially for human ones, more advanced expression systems with complete eukaryotic protein translation machinery are often necessary. While nonenzymatic PTMs are basically out of our control during recombinant protein expression, some of the enzymatic PTMs could be controlled and others might be even beneficial to structural research. From the point of view of structural biology, the most frequent PTM found in eukaryotically expressed proteins, and arguably also the most heterogeneous and bulky, is protein N-glycosylation. Luckily, this PTM is also the one (and more or less the only one) for which nowadays exist well-paved roads

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enabling its modification relatively easily [11,306,307]. The choice of the expression host represents the first level of N-glycosylation control. While insect cell lines and certain protozoa (e.g. Leishmania tarentolae — LEXSY [308]) produce N-glycans that are sufficiently small and homogeneous, wild-type oligosaccharides synthesized by yeasts and mammalian cell lines are in general too complex for protein crystallization. As yeast N-glycans are of high-mannose type, these oligosaccharide chains can be cleaved off leaving single GlcNAc residue using endoglycosidase H (Endo H), but fucosylated glycans of mammalian type are resistant to Endo H treatment under native conditions. Glycosyltransferases deficient cell lines such as CHO Lec3.2.8.1 [309] or HEK293S GnTI− [310] that both produce proteins with perfectly homogeneous Man5GlcNAc2 N-glycans are well established in protein crystallography [311,312]. These cell lines have been recently successfully used in difficult structural studies of G-protein coupled receptor rhodopsin [313] or enzyme receptor protein tyrosine phosphatases [314,315]. Alternatively, chemical inhibitors of N-glycosylation pathways such as kifunensine or swainsonine are able to modify oligosaccharides also in other cell lines and produce Endo H sensitive low-mannose type glycans [306,312]. For a protein to crystallize, its molecules have to interact with each other in some way in order to form a repeating ordered space pattern as a nuclei from which the crystal might begin to grow. Charged residues on the surface of the protein contribute to these interactions by forming salt bridges, however, small motions such as those due to a flexible, solvent-exposed amino acid side chains can be equally disruptive to a well-ordered crystal lattice. One possible approach to reducing surface entropy used in structural biology that is inspired by naturally occurring PTM is an in vitro reductive methylation of lysine residues. It is well known that hydrophobic interactions of methylated lysines drive the formation of certain protein complexes, notably in the interaction of histone tails with chromodomains [316]. Indeed, reductive methylation of free amino groups has emerged as very efficient method for improving protein crystallizability in several large scale structural proteomics/genomics research programs [317,318] and helped to solve the structures of enzymes like Bacillus anthracis alanine racemase (EC 5.1.1.1) or Murray Valley encephalitis virus methyltransferase (EC 2.1.1.56) [317]. From yet another, even more practical point of view, proper PTMs on recombinantly expressed enzymes are absolutely necessary when used as a therapeutic agents in human clinical medicine. In 2010, there were approximately 200 therapeutic proteins approved in the United States and Europe, many hundreds in clinical trials and even larger number in preclinical development [319]. Majority of therapeutical proteins require posttranslational processing to obtain full biological function, only glycosylation itself is associated with 40–70% of all approved products [319,320]. In contrary to structural biology the goal here is different — to achieve PTM profile as much human-like as possible. Without engineering, PTMs of heterologously expressed proteins might be even immunogenic in human, therefore great efforts in recent years were dedicated to e.g. development of cellular technologies enabling production of therapeutic proteins with humanized N-glycosylation in various expression hosts [11]. For example, the sialylation of glycoprotein biotherapeutics is important to maintain long

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residence time in circulation and prevent clearance by asialoglycoprotein receptors in the liver [321]. Increased terminal sialylation can thus prolong the serum half-life of glycoproteins and in turn might decrease necessary dosage of the biotherapeutics or prolong intervals of their administration. This has been achieved by developing yeast and insect cell lines expressing genes of human glycosylation cascade [322,323] or by overexpression of α-2,6-sialyltransferase (EC 2.4.99.1) in CHO cells to promote sialic acid correct linkage [324]. While these glycosylation modification technologies were developed primarily with respect to production of therapeutic monoclonal antibodies, they are also applicable to production of recombinant enzymes. Out of twelve recombinant enzyme therapeutics marketed in 2010, only one was produced in yeast, eight in CHO cell lines and three in other human cell lines [319]. Therefore we might expect that recently developed sialylation technologies will have impact on the market of these recombinant enzymes, too.

5.

Conclusions and perspectives

Because of their key role in regulation of enzymes' activity, structure, and assembly, PTMs of enzymes and other key cellular proteins have been the subject of numerous investigations. Historically, the identification of these modifications in most abundant proteins produced a long list of individual modifications involving both the cleavage, reassembly, and branching of the polypeptide chain and modifications of the side chain groups. Today, the problem of enzymes' PTMs has been approached using modern technologies of contemporary proteomics assisted by both high throughput and high resolution experimental techniques. One direction of modern proteomics has been the global analysis of individual PTMs in an attempt to provide a “complete” list of proteins possessing the individual modifications. The second complementary direction is the use of modern high resolution mass spectrometry in order to detect all PTMs within the most important cellular enzymes and proteins. In the future, one might envisage the use of robust high throughput analytical techniques that will be able to detect multiple PTMs on a global scale of individual proteomes from a number of carefully selected cells in the way that has been used recently for the functional interrogation of human genome [324]. Meanwhile, continuing investigations in the field are opening a fascinating picture of functional space created by coordinated enzymes' PTMs and their mutual interplays on one hand, and specific chemical environment defining nonenzymatic protein modifications and noncovalent modulations of enzymes on the other [325]. In this respect, both the detailed mechanistic investigation using the methods of structural biochemistry and the global functional interrogations of individual key cellular enzyme systems using systems biology approach will most probably continue to provide the most critical breakthrough finding in the near future [81,146].

Acknowledgments Research in authors' laboratories has been supported by Charles University in Prague (UNCE 204025/2012) and by

Czech Science Foundation (P207/10/1040, 303/09/0477, 305/09/ H008, and P504/11/0394).

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