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Progress in Research Methods for Protein Palmitoylation
CHINESE JOURNAL OF ANALYTICAL CHEMISTRY Volume 42, Issue 4, April 2014 Online English edition of the Chinese language journal
Cite this article as: Chin J Anal Chem, 2014, 42(4), 616–622.
REVIEW
Progress in Research Methods for Protein Palmitoylation FANG Cai-Yun, ZHANG Xiao-Qin, LU Hao-Jie* Department of Chemistry and Institutes of Biomedical Sciences, Fudan University, Shanghai 200433, China
Abstract: Protein palmitoylation, as one of the most important lipid post-translational modifications, plays a key role in different cellular functions such as signal transduction, metabolism, apoptosis, and so on. Disorders of protein palmitoylation will lead to many human diseases, including cancer. Therefore, qualitative and quantitative analysis of protein palmitoylation is very important for underlying its biological functions. Here, the technologies and methods for palmitoylation in recent years were reviewed. Their advantages/disadvantages and future trend were also discussed. Key Words:
1
Palmitoylation; Mass spectrometry; Proteomics
Introduction
Protein post-translational modification (PTMs) plays very important roles and can make proteins more complex in structure and perfect in function. Lipid modification is one of the significant PTMs. Palmitate is the most abundant fatty acid and can be preferentially modified to proteins covalently. Depending on their attachment way, palmitoylation is divided into two types, N-palmitoylation (addition to Gly/Cys residue by amide linkage) and S-palmitoylation. S-palmitoylation, also known as S-acylation, essentially refers to the covalent attachment of a saturated 16-carbon palmitate moiety to a specific cysteine (Cys) residue through a thioester linkage[1] (Fig.1). It is a reversible and dynamic modification and has a major regulatory role in protein function. It was found that palmitoylation occurs on numerous membrane proteins and membrane associated cytosolic proteins, such as signaling proteins (including the key factor in cancer and synaptic signal interruption), cellular adhesion molecules, neurotransmitter receptors, and so on. Palmitoylation can modulate the activities and stability of proteins, protein-protein interactions and protein’s membrane association, thus it plays a significant role in a wide range of biological processes such as cell signal transduction, metabolism, apoptosis, and carcinogenesis[1,2]. In order to further understand the mechanisms of protein
palmitoylation, it is necessary to analyze the palmitoylated proteins qualitatively and quantitatively, characterize all of the related enzymes about palmitoylation and depalmitoylation, and monitor the dynamically the cycle of palmitoylation and depalmitoylation. Traditionally, site-directed mutagenesis approach is used, in which the candidate Cys residue is mutated to Ser or Ala, the mutant clone and wild type are then transfected into cells respectively and analyzed their functional differences. However, this approach cannot detect endogenous palmitoylated proteins directly, and the mutation of specific Cys may cause its adjacent Cys to be abnormal[3]. In addition, numerous inhibitors were widely used to inhibit palmitoylation in cells, such as 2-bromopalmitate (2-BP)[4], Cerulenin and Tunicamycin[5]. Although these methods contributed to our understanding of protein palmitoylation greatly, they were laborious and time consuming because of
Fig.1 Protein S-palmitoylation which consists in the attachment of a saturated C16 acyl chain to protein cysteine (Cys) via a thioester bond
Received 10 August 2013; accepted 16 December 2013 * Corresponding author. Email: [email protected] This work was supported by the National Natural Science Foundation of China (Nos. 21205018, 21025519), and the Shanghai Pujiang Program of China (No. 13PJD003). Copyright © 2014, Changchun Institute of Applied Chemistry, Chinese Academy of Sciences. Published by Elsevier Limited. All rights reserved. DOI: 10.1016/S1872-2040(13)60727-6
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insufficient known palmitoylated sites. Firstly, although a class of palmitoyl acyltransferases (PATs) sharing a conserved Asp-His-His-Cys (DHHC) motif within a cysteine-rich domain (CRD), as well as several deacylating enzymes, was recognized several years ago, the enzymology of protein S-acylation and deacylation remains poorly understood[6]. Secondly, there is no general consensus motif for palmitoylation site prediction[7]. Thirdly, palmitoylation is more frequently detected on transmembrane or membrane associated proteins[8], while the solubility and abundance of membrane proteins are relatively low. Attachment of long chain fatty acid further increases its hydrophobicity and makes it more difficult to be detected. Fourthly, unlike protein phosphorylation and glycosylation, no convenient method (e.g. S-palmitoylation-specific antibody or affinity group) is available to detect and enrich low abundant S-palmitoylated proteins. Therefore, understanding of the mechanisms and functions of reversible S-palmitoylation has progressed at a slow pace. With the development of mass spectrometry (MS) and proteomic techniques, more and more palmitoylated proteins and sites have been identified, which facilitates the understanding of protein palmitoylation. Here, the techniques and methods for palmitoylation analysis based on MS and proteomics in recent years are reviewed.
2
Bioinformatics approach
To some degree, palmitoylation site prediction by computational method can provide database for further experimental design. Several protein palmitoylation sites predictors were developed. CSS-Palm[9,10] and NBA-Palm[11] could predict the protein palmitoylation modification sites online. NBA-Palm (http://nbapalm.biocuckoo.org/) was constructed based on Naive Bayesian Algorithm (NBA). The training data set included 245 non-homologous sites from 105 proteins. CSS-Palm was based on clustering and scoring strategy (CSS) algorithm and Group-based Prediction System (GPS) algorithm. The training data set of CSS-Palm 3.0 (http://csspalm.biocuckoo.org/) included 439 palmitoylation sites from 194 proteins. In 2013, CSS-Palm 4.0 was released, which included a fourth-generation of GPS algorithm and the latest training data set containing 583 palmitoylation sites from 277 distinct proteins. In parallel, Deng et al[12] proposed prediction models CKSAAP-Palm (http://www.aporc.org/doc/ wiki/CKSAAP-Palm) and PPWMs[13]. They used the encoding scheme based on the composition of k-spaced amino acid pairs (CKSAAP) to represent the sequence fragments. Then support vector machine (SVM) was used as the predictor.
3
Identification of palmitoylation sites and lipid moieties by mass spectrometry MS is a fundamental tool for biologists and biochemists
because of its high sensitivity and low sample consumption. Target proteins are tryptically purified by chromatography or immunoprecipitation, digested and identified the species of fatty acid by mass shift of these modification-containing fragment ions in MS after hydroxylamine treatment[14]. Hemagglutinin (HA) is a major envelope glycoprotein which can mediate viral and cell membrane fusion. HA is anchored in the viral envelope by a light HA2 chain containing one transmembrane domain and a cytoplasmic tail. Three Cys residues in the C-terminal region, one in the transmembrane domain and the other two in the cytoplasmic tail, are highly conserved and potentially palmitoylated in all HA subtypes. Kordyukova et al[15] analyzed the species of fatty acids in influenza virus protein HA by matrix assisted laser ionization time of flight mass spectrometry (MALDI-TOF MS). They found that influenza B virus HA possessing two cytoplasmic Cys contained palmitate, whereas influenza C virus HA with one transmembrane Cys was stearoylated. HA of influenza A virus having one transmembrane and two cytoplasmic Cys contained both palmitate and stearate. Serebryakova et al[16] assessed the primary structures of HA2 C-terminal anchoring segments of A/X-31(H3 subtype), A/Puerto Rico/8/34 (H1 subtype) and A/FPV/Weybridge/34 (H7 subtype) by using MALDI-TOF MS. They found that these peptides isolated from the viruses were acylated not only by palmitic, but also by stearic acid residues. The palmitate/stearate ratio in the three strains studied was different, and the A/FPV/Weybridge/ 34 strain had a priority to stearate binding. Ozols et al[17] reported a [3H]-palmitate labeling method to identify the palmitoylation sites of tubulin. Purified porcine brain α-tubulin containing [3H]-palmitate was cleaved with cyanogen bromide, digested with trypsin and Lys-C protease and then subjected to gas/liquid-phase sequencing. It was shown that C376 was the primary site for palmitoylation compared with residues C20, C213 and C305. Bizzozero et al[18] found that palmitoylation was the major PTM of native myelin proteolipid protein (PLP), and the majority of PLP and minor myelin proteolipid protein (DM-20) molecules in the central nervous system were fully acylated. Gas chromatography-mass spectrometry (GC-MS) is widely used to separate and analyze the complicated samples due to high resolution of GC and high sensitivity of MS, which is suitable for analyzing low molecular weight and volatile compounds. Sorek et al determined the lipid moieties of AtROP6[19] and CBL1[20] by GC-MS. The fatty acids were cleaved from proteins by hydrogenation with platinum (IV) oxide, which caused an acid transesterification of the acyl groups, adding an ethyl group to the carboxyl head of the fatty acid. The addition of the ethyl group reduced the polarity of fatty acids, allowing their efficient separation by GC. The analysis could be carried out with as little as 1 μg of purified proteins and allows chemical identification and, potentially, quantification of the acyl
FANG Cai-Yun et al. / Chinese Journal of Analytical Chemistry, 2014, 42(4): 616–622
moieties[21]. Another approach termed isotope-coded fatty acid transmethylation (iFAT) was developed for quantitative comparison of acyl moieties in different cell states in vitro[22], which was suitable to handle hydrophobic membrane proteins. Identity of acyl moieties were identified by database searching and comparison of chromatographic retention time with that of standard fatty acid methyl esters, while stable isotope labeling was used as internal standard for quantitation. However, numerous palmitoylated proteins are membrane proteins with strong hydrophobicity, and it is not easy to obtain pure protein sample. So this kind of method has its limitation to some extent.
4
“Palmitate-centric” approach
4.1
Metabolic labeling with radioactive palmitate
S-palmitoylated proteins were originally detected by metabolic labeling with radioactive 3H-palmitate and 125 I-palmitate followed by autoradiography of acyl moieties or labeled proteins. While 3H-palmitate was the most widely used radioactively labeled with palmitate[23]. However, these approaches were typically tedious, hazardous and expensive owing to lengthy film exposure times, high level of radioactive palmitate and lack of any straightforward means to enrich and identify radiolabeled proteins. In addition, they were only used to detect the high abundant palmitoylated proteins in living cells due to their low sensitivity and could not provide protein’s stoichiometric information. 4.2
Metabolic labeling with non-radioactive derivatives of palmitate
Azido or Alkynl fatty acid probes are very sensitive and can afford new opportunity to detect and enrich palmitoylated proteins in cells. Azide- or alkynyl-palmitate analogs are similar with their natural fatty acid counterparts in their structure and function, and can be incorporated metabolically into cellular proteins. And then the proteins modified with these probes are selectively conjugated to modified biotin or a modi-fied fluorophore that reacts selectively with the azido or alkynyl group via a Staudinger ligation[24] or a click reaction[25], so that the palmitoylated proteins can be enriched and/or detected. The advantages of these strategies are that they are nonradioactive, have short detection time and high degree of sensitivity in comparison with radioactive methods. They also enable the enrichment of palmitoylated proteins in cellular mixtures for either affinity purification or detection by fluorescence imaging. 4.2.1
Approaches with azido fatty acid probe as chemical reporter
Organic azides are functional groups of sufficiently high chemical reactivity to be easily modified, but are metabolically inert in cellular environments. Hang et al[26] developed a series of ω-azido fatty acids that contained 12-, 14-, 15- and 16-carbon chains as non-radioactive probes, which could be efficiently metabolized by mammalian cells. Proteins incorporating azido fatty acids were then subjected to Staudinger ligation with biotin phosphine reagent followed by western blotting or enriched by streptavidin beads to detect palmitoylated proteins rapidly. Their results indicated that ω-azido C-12 (Az-C12) selectively targeted N-myristoylated proteins, whereas the longer ω-azido fatty acids such as Az-C15 primarily labeled S-acylated proteins. Kostiuk et al[27] used Az-C14-CoA as chemical probe and identified 21 palmitoylated proteins such as HMGCS from rat liver mitochondria by MS based method. Ching et al[28] found that WntD did not undergo lipid modification and was secreted at high levels by metabolic labeling, which was different from other Wnt proteins. Martin et al[29] studied the myristoylated proteins during apoptosis using a bio-orthogonal azidomyristate analog. They reported that this method represented over a million-fold signal amplification in comparison to using radioactive labeling methods. 4.2.2
Approaches with alkynyl fatty acid probe as chemical reporter
Although azide fatty acid analogues have made great progress in detection of protein acylation, appending an azide group to an alkyl fatty acid chain may interfere with the hydrophobicity of fatty acid and the mechanism, by which it inserts into lipid membranes. While alkyne tag can maintain the hydrophobicity of carbon chain and minimize interference with the hydrophobic lipid environment. They are metabolically inert (Fig.2). In addition, it is also reported that click chemistry is more efficient at coupling a rhodamine azide to alkyne-labeled proteins than either the reverse click
Fig.2
Diagrammatic sketch of palmitoylated proteins/peptides labelling with alkynyl fatty acid analogue
Alkynyl fatty acid analogue was incorporated into endogenous sites of palmitoylation via metabolic labelling during cell culture. After sufficient labelling, the cells were lysed, coupled with azide-conjugated materials by click chemistry and then detected by fluorescence detecting or LC-MS analysis
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chemistry format or a Staudinger ligation of alkyne-biotin to azido-labeled proteins. ω-Alkynyl fatty acids offer an advantage over their corresponding azido-modified counterparts, resulting in higher sensitivity and overall improved detection[30]. Fluorescent detection of palmitoylated proteins with chemical reporters was used to investigate dynamic and complex biological processes[31]. Quantitative fluorescence microscopic method was used for the studies of intracellular targeting and trafficking of GFP-tagged palmitoylated proteins in living cells[32,33]. Thus, Hannoush et al[34] reported a suite of novel chemical probes based on ω-alkynyl fatty acids for biochemical detection and cellular imaging of lipid-modified proteins. They found that ω-alkynyl fatty acids of varying chain length could be metabolically incorporated onto cellular proteins. After the alkynyl group incorporated onto acylated proteins was chemoselectively ligated to azide-tagged biotin or fluorophore by a Cu1-catalyzed alkyne-azide [3 + 2] cycloaddition reaction, the conjugated proteins were separated by gel electrophoresis and analyzed by Western blot using streptavidin-linked horseradish peroxidase. Using fluorescence imaging, this methodology could describe the subcellular distribution of lipid-modified proteins across a panel of different mammalian cell lines and during cell division. Zhang et al[35] described a tandem labeling and fluorescence imaging method using two orthogonal chemical reporters to simultaneous monitor palmitate cycling rates and protein turnover, in which one chemical reporter was for detecting the S-palmitoylation and the other was for analyzing protein turnover. They also found that shorter fatty acid analogs (az-12 and alk-12) preferentially labeled N-myristoylated protein, while the longer fatty acid derivatives (az-15 and alk-16) were incorporated onto S-palmitoylated proteins[36]. Yount et al[37,38] used alk-16 as chemical reporter and analyzed S-palmitoylation proteome in mouse dendritic cell line. They selectively identified 157 proteins with different cellular functions (60 and 97 proteins assigned to high- and medium-confidence lists, respectively). They found that S-palmitoylation of IFITM3 on membrane-proximal Cys regulated its clustering in membrane compartments and its antiviral activity against influenza virus. Yap et al[39] reported that ω-alkynyl palmitate analog could be readily and specifically incorporated into GAPDH or mitochondrial 3-hydroxyl-3-methylglutaryl-CoA synthase in vitro and reacted with an azido-biotin probe or the fluorogenic probe 3-azido-7-hydroxycoumarin using click chemistry for rapid detection of protein palmitoylation. Martin et al[40] used commercially available palmitic acid analog 17-octadecynoic acid (17-ODYA) as a bioorthogonal, click chemistry probe for in situ labeling. They identified 125 high-confidence and about 200 medium-confidence palmitoylated proteins from Jurkat T cells, including G protein, receptors and a family of uncharacterized hydrolases whose plasma membrane localization depended on palmitoylation. It was the first global
inventory of acylated proteins from human cells using a bioorthogonal labeling strategy for the enrichment and identification of palmitoylated proteins. Subsequently, they reported the metabolic incorporation of 17-ODYA in combination with stable-isotope labeling with amino acids in cell culture (SILAC) and pulse-chase methods to generate a global quantitative map of dynamic protein palmitoylation events in cells. They confidently identified and quantified more than 400 palmitoylated proteins in mouse T-cell hybridoma cells, and distinguished stably palmitoylated proteins from those that turned over rapidly by using this approach[41]. In general, “palmitate-centric” approach has its own merits and weaknesses. The metabolic labeling method cannot be readily applied to analyze tissue samples and body fluids because radioactive or chemically modified palmitate analogs should be incorporated metabolically into cellular proteins. This method is also potentially difficult to study protein S-acylation in cancer cells, in which overexpression and hyperactivity of fatty-acid synthase dramatically decrease the intake of exogenous long-chain fatty acids[42]. In comparison with those more stably palmitoylated proteins, “palmitatecentric” approach may be likely bias for proteins that undergo rapid palmitate turnover. Once palmitate analogues are introduced to cellular proteins, it can only be used to analyze protein palmitoylation other than shorter, longer or unsaturated lipid chains. Physiological effects caused by the structural differences between natural fatty acids and fatty acid analogues are not very clear. Most importantly, the metabolic pathway in eukaryotes is extremely complicated, so it may probably result in unpredictable biological effects due to the introduction of palmitate analogues. 5
“Cysteine-centric” approach
Diagrammatic sketch of “cysteine-centric” approach is shown in Fig.3. Acyl-biotinyl exchange (ABE) was first proposed by Drisdel’ group in 2004[43]. In this approach, after free thiols present in proteins were completely blocked by thiol reactive reagents such as idoacetamide (IAA) or N-ethylmaleimide (NEM), the palmitoyl thioester linkage was selectively cleaved by neutral hydroxylamine (HA) and resulted in new free sulfhydryl groups. Consequently, S-acylated proteins could be detected by autoradiography via radioactive NEM labeling, analyzed by the immunoblotting methods, or purified and enriched via avidin/streptavidinbiotin interactions after the proteins were marked with thiol specific reactive reagents such as biotin-conjugated 1-biotinamido-4-[4’-(maleimidomethyl)cyclohexanecarboxamido] butane (Btn-BMCC)[44] or biotin-HPDP (N-[6-(biotinamido)hexyl]3’-(2’-pyridyldithio) propionamide)[45]. Wan et al[46] used biotin-HPDP as biotinylation reagent to purify palmitoylated proteins. Fifty palmitoyl proteins in yeast were identified by
FANG Cai-Yun et al. / Chinese Journal of Analytical Chemistry, 2014, 42(4): 616–622
Fig.3 Diagrammatic sketch of “cysteine-centric” approach Disulfide bonds of proteins were reduced firstly, and free cysteine residues were alkylated. After proteins were digested into peptides, the sample was treated with hydroxylamine to selectively cleave thioesters. The newly exposed cysteine residues were disulfide-bonded to affinity beads and were purified. The eluate was then analyzed by LC-MS
MS including many SNARE proteins, amino acid permeases and signaling proteins. Thioester bond between palmitic acid and protein was sensitive to hydroxylamine treatment, so most palmitoylated peptide ions were markedly increased in abundance after HA treatment, while the MS signal of non-palmitoylated peptide ions had no significant change before and after HA treatment. Thus, by comparing the spectral counts of identified peptides between the +HA sample and -HA sample, we could evaluate the degree of credibility of each palmitoylation site. Acyl-biotinyl exchange chemistry and streptavidin agarose affinity purification were applied to enrich palmitoylated peptides from tubulin by Zhao et al[47]. The results indicated that 11 cysteine residues of the α/β tubulin heterodimer were palmitoylated by using nano-LCMS/MS. Hemsley et al[48] also reported that α-tubulin and AGG2 were palmitoylated in plant. Kang et al[49] identified 68 known and >200 candidate S-acylated proteins from whole rat brain, purified rat synaptosomes and cultured embryonic rat neurons by ABE approach, which included both hydrophobic and hydrophilic proteins, such as neurotransmitter receptors, adhesion molecules, scaffold proteins, and so on. Yang et al[50] reported a proteomics approach (palmitoyl protein identification and site characterization, PalmPISC). In combination with label-free spectral counting quantitation, PalmPISC led to the identification of 67 known and 331 novel candidate S-acylated proteins as well as the localization of 25 known and 143 novel candidate S-acylation sites. Emmer et al[51] identified 124 palmitoylated proteins in Trypanosoma brucei by ABE approach. Except label-free quantitative method, isobaric labeling approaches were also used to distinguish S-acylated proteins from contaminants, in which all samples were pooled and analyzed simultaneously owing to the isobaric tags. The analysis time was reduced, as well as the repeatability and reproducibility were improved. Thus, isobaric labeling
approach was widely used in proteomic research. Zhang et al[52] presented a palmitoylcysteine isolation capture and analysis (or PICA) method, to screen substrates of human DHHC2 in HeLa cells. They quantified the protein profiles between ZDHHC2 knockdown and control HeLa cells by using isotope-coded affinity tag (ICAT) approach. A total of 57 sites were identified from 50 proteins, and CKAP4/p63 was an important substrate of DHHC2, a putative tumor suppressor. After the proteins were blocked by methyl methanethiosulfonate (MMTS), hydrolysed by hydroxylamine, and purified by thiopropyl sepharose, Forrester et al[53] identified the S-palmitoylated proteins in HEK293 cells by Isobaric tags for relative and absolute quantitation (iTRAQ) to differentiate S-palmitoylated peptides and “contaminated peptides”. In total, 93 putative sites of S-acylation on 88 peptides were identified. ABE approach does not need in vivo metabolic labeling, so it can be readily applied to analyze cells (including cancer cells), tissue and body fluid samples. Although great progress of ABE has been made in detecting palmitoylated proteins, there are several problems existing in ABE approach: (1) ABE would result in higher false discovery rates. ABE-based method is thioester bond-specific but not S-palmitoylationspecific, the false discovery rates derived from other modifications, which contains relatively high thioester bonds. Thus, it is critical to block all free sulfhydryl groups in the proteins that would otherwise react with the labeling reagents, so the type, amount and incubation time of the blocking reagents should be empirically tested. The reaction time, concentration of hydroxylamine and pH of the solution should be tested to make sure of complete cleavage of palmitate with hydroxylamine. Complete labeling of the free sulfhydryl groups after palmitate cleavage is very important. To minimize the false-positive results, control experiment (-HA) is usually required. Signal development with streptavidin
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often results in the detection of a small pool of endogenously biotinylated proteins, so appropriate control experiments should be carried out. Otherwise, these factors can result in nearly one-third of the identified proteins being designated as false positive results[8,54]. In comparison with ABE approach, click chemistry produced a significantly lower number of false positives[55]. (2) S-acylated proteins can be labeled with fatty acids other than palmitate, while all fatty acids are removed after ABE chemistry. So the identities of long chain fatty acids attached to proteins of interest cannot be determined directly. However, this problem can potentially be addressed by GC-MS, which allows direct identification of lipids attached to proteins.
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Given the technical difficulties hindering palmitoylation analysis, this modification’s function has been underestimated. In recent years, there is continuously increasing interest and significant progress has been made in protein palmitoylation, especially with the development of MS-based proteomic approaches (such as ABE approach and metabolic labeling with palmitate analogue), so more and more palmitoylated sites have been identified. However, current researches mainly focus on the identification of palmitoylated sites-qualitative analysis, and all methods have their own limitations. So more qualitative and quantitative approaches for protein palmitoylation, especially effective and accurate approaches, needs to be further developed. For example, protein palmitoylation is often found to modify transmembrane or membrane associated proteins, thus the techniques of membrane proteomics can be used to improve proteins’ solubility. Existing methods require typically complicated procedures, cause easily sample loss (especially for low abundant proteins), and lead to relatively high false positive results. To solve these problems, we need to develop new approaches with a simplified experimental procedure or by designing new suitable detergents, new matrices to enrich palmitoylated proteins with high selectivity and efficiency, or new materials for qualitative and quantitative analysis simultaneously. The development of new fluorescent labeling reagents with high sensitivity and selectivity or fatty acid analogues with high metabolic efficiency is also critical to obtain more reliable information for palmitoylated proteins and sites. Live cell imaging may also provide further a perspective tool for dynamic monitoring of complex palmtoylation and insights into its biological importance and function.
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Cite this article as: Chin J Anal Chem, 2014, 42(4), 616–622.
REVIEW
Progress in Research Methods for Protein Palmitoylation FANG Cai-Yun, ZHANG Xiao-Qin, LU Hao-Jie* Department of Chemistry and Institutes of Biomedical Sciences, Fudan University, Shanghai 200433, China
Abstract: Protein palmitoylation, as one of the most important lipid post-translational modifications, plays a key role in different cellular functions such as signal transduction, metabolism, apoptosis, and so on. Disorders of protein palmitoylation will lead to many human diseases, including cancer. Therefore, qualitative and quantitative analysis of protein palmitoylation is very important for underlying its biological functions. Here, the technologies and methods for palmitoylation in recent years were reviewed. Their advantages/disadvantages and future trend were also discussed. Key Words:
1
Palmitoylation; Mass spectrometry; Proteomics
Introduction
Protein post-translational modification (PTMs) plays very important roles and can make proteins more complex in structure and perfect in function. Lipid modification is one of the significant PTMs. Palmitate is the most abundant fatty acid and can be preferentially modified to proteins covalently. Depending on their attachment way, palmitoylation is divided into two types, N-palmitoylation (addition to Gly/Cys residue by amide linkage) and S-palmitoylation. S-palmitoylation, also known as S-acylation, essentially refers to the covalent attachment of a saturated 16-carbon palmitate moiety to a specific cysteine (Cys) residue through a thioester linkage[1] (Fig.1). It is a reversible and dynamic modification and has a major regulatory role in protein function. It was found that palmitoylation occurs on numerous membrane proteins and membrane associated cytosolic proteins, such as signaling proteins (including the key factor in cancer and synaptic signal interruption), cellular adhesion molecules, neurotransmitter receptors, and so on. Palmitoylation can modulate the activities and stability of proteins, protein-protein interactions and protein’s membrane association, thus it plays a significant role in a wide range of biological processes such as cell signal transduction, metabolism, apoptosis, and carcinogenesis[1,2]. In order to further understand the mechanisms of protein
palmitoylation, it is necessary to analyze the palmitoylated proteins qualitatively and quantitatively, characterize all of the related enzymes about palmitoylation and depalmitoylation, and monitor the dynamically the cycle of palmitoylation and depalmitoylation. Traditionally, site-directed mutagenesis approach is used, in which the candidate Cys residue is mutated to Ser or Ala, the mutant clone and wild type are then transfected into cells respectively and analyzed their functional differences. However, this approach cannot detect endogenous palmitoylated proteins directly, and the mutation of specific Cys may cause its adjacent Cys to be abnormal[3]. In addition, numerous inhibitors were widely used to inhibit palmitoylation in cells, such as 2-bromopalmitate (2-BP)[4], Cerulenin and Tunicamycin[5]. Although these methods contributed to our understanding of protein palmitoylation greatly, they were laborious and time consuming because of
Fig.1 Protein S-palmitoylation which consists in the attachment of a saturated C16 acyl chain to protein cysteine (Cys) via a thioester bond
Received 10 August 2013; accepted 16 December 2013 * Corresponding author. Email: [email protected] This work was supported by the National Natural Science Foundation of China (Nos. 21205018, 21025519), and the Shanghai Pujiang Program of China (No. 13PJD003). Copyright © 2014, Changchun Institute of Applied Chemistry, Chinese Academy of Sciences. Published by Elsevier Limited. All rights reserved. DOI: 10.1016/S1872-2040(13)60727-6
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insufficient known palmitoylated sites. Firstly, although a class of palmitoyl acyltransferases (PATs) sharing a conserved Asp-His-His-Cys (DHHC) motif within a cysteine-rich domain (CRD), as well as several deacylating enzymes, was recognized several years ago, the enzymology of protein S-acylation and deacylation remains poorly understood[6]. Secondly, there is no general consensus motif for palmitoylation site prediction[7]. Thirdly, palmitoylation is more frequently detected on transmembrane or membrane associated proteins[8], while the solubility and abundance of membrane proteins are relatively low. Attachment of long chain fatty acid further increases its hydrophobicity and makes it more difficult to be detected. Fourthly, unlike protein phosphorylation and glycosylation, no convenient method (e.g. S-palmitoylation-specific antibody or affinity group) is available to detect and enrich low abundant S-palmitoylated proteins. Therefore, understanding of the mechanisms and functions of reversible S-palmitoylation has progressed at a slow pace. With the development of mass spectrometry (MS) and proteomic techniques, more and more palmitoylated proteins and sites have been identified, which facilitates the understanding of protein palmitoylation. Here, the techniques and methods for palmitoylation analysis based on MS and proteomics in recent years are reviewed.
2
Bioinformatics approach
To some degree, palmitoylation site prediction by computational method can provide database for further experimental design. Several protein palmitoylation sites predictors were developed. CSS-Palm[9,10] and NBA-Palm[11] could predict the protein palmitoylation modification sites online. NBA-Palm (http://nbapalm.biocuckoo.org/) was constructed based on Naive Bayesian Algorithm (NBA). The training data set included 245 non-homologous sites from 105 proteins. CSS-Palm was based on clustering and scoring strategy (CSS) algorithm and Group-based Prediction System (GPS) algorithm. The training data set of CSS-Palm 3.0 (http://csspalm.biocuckoo.org/) included 439 palmitoylation sites from 194 proteins. In 2013, CSS-Palm 4.0 was released, which included a fourth-generation of GPS algorithm and the latest training data set containing 583 palmitoylation sites from 277 distinct proteins. In parallel, Deng et al[12] proposed prediction models CKSAAP-Palm (http://www.aporc.org/doc/ wiki/CKSAAP-Palm) and PPWMs[13]. They used the encoding scheme based on the composition of k-spaced amino acid pairs (CKSAAP) to represent the sequence fragments. Then support vector machine (SVM) was used as the predictor.
3
Identification of palmitoylation sites and lipid moieties by mass spectrometry MS is a fundamental tool for biologists and biochemists
because of its high sensitivity and low sample consumption. Target proteins are tryptically purified by chromatography or immunoprecipitation, digested and identified the species of fatty acid by mass shift of these modification-containing fragment ions in MS after hydroxylamine treatment[14]. Hemagglutinin (HA) is a major envelope glycoprotein which can mediate viral and cell membrane fusion. HA is anchored in the viral envelope by a light HA2 chain containing one transmembrane domain and a cytoplasmic tail. Three Cys residues in the C-terminal region, one in the transmembrane domain and the other two in the cytoplasmic tail, are highly conserved and potentially palmitoylated in all HA subtypes. Kordyukova et al[15] analyzed the species of fatty acids in influenza virus protein HA by matrix assisted laser ionization time of flight mass spectrometry (MALDI-TOF MS). They found that influenza B virus HA possessing two cytoplasmic Cys contained palmitate, whereas influenza C virus HA with one transmembrane Cys was stearoylated. HA of influenza A virus having one transmembrane and two cytoplasmic Cys contained both palmitate and stearate. Serebryakova et al[16] assessed the primary structures of HA2 C-terminal anchoring segments of A/X-31(H3 subtype), A/Puerto Rico/8/34 (H1 subtype) and A/FPV/Weybridge/34 (H7 subtype) by using MALDI-TOF MS. They found that these peptides isolated from the viruses were acylated not only by palmitic, but also by stearic acid residues. The palmitate/stearate ratio in the three strains studied was different, and the A/FPV/Weybridge/ 34 strain had a priority to stearate binding. Ozols et al[17] reported a [3H]-palmitate labeling method to identify the palmitoylation sites of tubulin. Purified porcine brain α-tubulin containing [3H]-palmitate was cleaved with cyanogen bromide, digested with trypsin and Lys-C protease and then subjected to gas/liquid-phase sequencing. It was shown that C376 was the primary site for palmitoylation compared with residues C20, C213 and C305. Bizzozero et al[18] found that palmitoylation was the major PTM of native myelin proteolipid protein (PLP), and the majority of PLP and minor myelin proteolipid protein (DM-20) molecules in the central nervous system were fully acylated. Gas chromatography-mass spectrometry (GC-MS) is widely used to separate and analyze the complicated samples due to high resolution of GC and high sensitivity of MS, which is suitable for analyzing low molecular weight and volatile compounds. Sorek et al determined the lipid moieties of AtROP6[19] and CBL1[20] by GC-MS. The fatty acids were cleaved from proteins by hydrogenation with platinum (IV) oxide, which caused an acid transesterification of the acyl groups, adding an ethyl group to the carboxyl head of the fatty acid. The addition of the ethyl group reduced the polarity of fatty acids, allowing their efficient separation by GC. The analysis could be carried out with as little as 1 μg of purified proteins and allows chemical identification and, potentially, quantification of the acyl
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moieties[21]. Another approach termed isotope-coded fatty acid transmethylation (iFAT) was developed for quantitative comparison of acyl moieties in different cell states in vitro[22], which was suitable to handle hydrophobic membrane proteins. Identity of acyl moieties were identified by database searching and comparison of chromatographic retention time with that of standard fatty acid methyl esters, while stable isotope labeling was used as internal standard for quantitation. However, numerous palmitoylated proteins are membrane proteins with strong hydrophobicity, and it is not easy to obtain pure protein sample. So this kind of method has its limitation to some extent.
4
“Palmitate-centric” approach
4.1
Metabolic labeling with radioactive palmitate
S-palmitoylated proteins were originally detected by metabolic labeling with radioactive 3H-palmitate and 125 I-palmitate followed by autoradiography of acyl moieties or labeled proteins. While 3H-palmitate was the most widely used radioactively labeled with palmitate[23]. However, these approaches were typically tedious, hazardous and expensive owing to lengthy film exposure times, high level of radioactive palmitate and lack of any straightforward means to enrich and identify radiolabeled proteins. In addition, they were only used to detect the high abundant palmitoylated proteins in living cells due to their low sensitivity and could not provide protein’s stoichiometric information. 4.2
Metabolic labeling with non-radioactive derivatives of palmitate
Azido or Alkynl fatty acid probes are very sensitive and can afford new opportunity to detect and enrich palmitoylated proteins in cells. Azide- or alkynyl-palmitate analogs are similar with their natural fatty acid counterparts in their structure and function, and can be incorporated metabolically into cellular proteins. And then the proteins modified with these probes are selectively conjugated to modified biotin or a modi-fied fluorophore that reacts selectively with the azido or alkynyl group via a Staudinger ligation[24] or a click reaction[25], so that the palmitoylated proteins can be enriched and/or detected. The advantages of these strategies are that they are nonradioactive, have short detection time and high degree of sensitivity in comparison with radioactive methods. They also enable the enrichment of palmitoylated proteins in cellular mixtures for either affinity purification or detection by fluorescence imaging. 4.2.1
Approaches with azido fatty acid probe as chemical reporter
Organic azides are functional groups of sufficiently high chemical reactivity to be easily modified, but are metabolically inert in cellular environments. Hang et al[26] developed a series of ω-azido fatty acids that contained 12-, 14-, 15- and 16-carbon chains as non-radioactive probes, which could be efficiently metabolized by mammalian cells. Proteins incorporating azido fatty acids were then subjected to Staudinger ligation with biotin phosphine reagent followed by western blotting or enriched by streptavidin beads to detect palmitoylated proteins rapidly. Their results indicated that ω-azido C-12 (Az-C12) selectively targeted N-myristoylated proteins, whereas the longer ω-azido fatty acids such as Az-C15 primarily labeled S-acylated proteins. Kostiuk et al[27] used Az-C14-CoA as chemical probe and identified 21 palmitoylated proteins such as HMGCS from rat liver mitochondria by MS based method. Ching et al[28] found that WntD did not undergo lipid modification and was secreted at high levels by metabolic labeling, which was different from other Wnt proteins. Martin et al[29] studied the myristoylated proteins during apoptosis using a bio-orthogonal azidomyristate analog. They reported that this method represented over a million-fold signal amplification in comparison to using radioactive labeling methods. 4.2.2
Approaches with alkynyl fatty acid probe as chemical reporter
Although azide fatty acid analogues have made great progress in detection of protein acylation, appending an azide group to an alkyl fatty acid chain may interfere with the hydrophobicity of fatty acid and the mechanism, by which it inserts into lipid membranes. While alkyne tag can maintain the hydrophobicity of carbon chain and minimize interference with the hydrophobic lipid environment. They are metabolically inert (Fig.2). In addition, it is also reported that click chemistry is more efficient at coupling a rhodamine azide to alkyne-labeled proteins than either the reverse click
Fig.2
Diagrammatic sketch of palmitoylated proteins/peptides labelling with alkynyl fatty acid analogue
Alkynyl fatty acid analogue was incorporated into endogenous sites of palmitoylation via metabolic labelling during cell culture. After sufficient labelling, the cells were lysed, coupled with azide-conjugated materials by click chemistry and then detected by fluorescence detecting or LC-MS analysis
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chemistry format or a Staudinger ligation of alkyne-biotin to azido-labeled proteins. ω-Alkynyl fatty acids offer an advantage over their corresponding azido-modified counterparts, resulting in higher sensitivity and overall improved detection[30]. Fluorescent detection of palmitoylated proteins with chemical reporters was used to investigate dynamic and complex biological processes[31]. Quantitative fluorescence microscopic method was used for the studies of intracellular targeting and trafficking of GFP-tagged palmitoylated proteins in living cells[32,33]. Thus, Hannoush et al[34] reported a suite of novel chemical probes based on ω-alkynyl fatty acids for biochemical detection and cellular imaging of lipid-modified proteins. They found that ω-alkynyl fatty acids of varying chain length could be metabolically incorporated onto cellular proteins. After the alkynyl group incorporated onto acylated proteins was chemoselectively ligated to azide-tagged biotin or fluorophore by a Cu1-catalyzed alkyne-azide [3 + 2] cycloaddition reaction, the conjugated proteins were separated by gel electrophoresis and analyzed by Western blot using streptavidin-linked horseradish peroxidase. Using fluorescence imaging, this methodology could describe the subcellular distribution of lipid-modified proteins across a panel of different mammalian cell lines and during cell division. Zhang et al[35] described a tandem labeling and fluorescence imaging method using two orthogonal chemical reporters to simultaneous monitor palmitate cycling rates and protein turnover, in which one chemical reporter was for detecting the S-palmitoylation and the other was for analyzing protein turnover. They also found that shorter fatty acid analogs (az-12 and alk-12) preferentially labeled N-myristoylated protein, while the longer fatty acid derivatives (az-15 and alk-16) were incorporated onto S-palmitoylated proteins[36]. Yount et al[37,38] used alk-16 as chemical reporter and analyzed S-palmitoylation proteome in mouse dendritic cell line. They selectively identified 157 proteins with different cellular functions (60 and 97 proteins assigned to high- and medium-confidence lists, respectively). They found that S-palmitoylation of IFITM3 on membrane-proximal Cys regulated its clustering in membrane compartments and its antiviral activity against influenza virus. Yap et al[39] reported that ω-alkynyl palmitate analog could be readily and specifically incorporated into GAPDH or mitochondrial 3-hydroxyl-3-methylglutaryl-CoA synthase in vitro and reacted with an azido-biotin probe or the fluorogenic probe 3-azido-7-hydroxycoumarin using click chemistry for rapid detection of protein palmitoylation. Martin et al[40] used commercially available palmitic acid analog 17-octadecynoic acid (17-ODYA) as a bioorthogonal, click chemistry probe for in situ labeling. They identified 125 high-confidence and about 200 medium-confidence palmitoylated proteins from Jurkat T cells, including G protein, receptors and a family of uncharacterized hydrolases whose plasma membrane localization depended on palmitoylation. It was the first global
inventory of acylated proteins from human cells using a bioorthogonal labeling strategy for the enrichment and identification of palmitoylated proteins. Subsequently, they reported the metabolic incorporation of 17-ODYA in combination with stable-isotope labeling with amino acids in cell culture (SILAC) and pulse-chase methods to generate a global quantitative map of dynamic protein palmitoylation events in cells. They confidently identified and quantified more than 400 palmitoylated proteins in mouse T-cell hybridoma cells, and distinguished stably palmitoylated proteins from those that turned over rapidly by using this approach[41]. In general, “palmitate-centric” approach has its own merits and weaknesses. The metabolic labeling method cannot be readily applied to analyze tissue samples and body fluids because radioactive or chemically modified palmitate analogs should be incorporated metabolically into cellular proteins. This method is also potentially difficult to study protein S-acylation in cancer cells, in which overexpression and hyperactivity of fatty-acid synthase dramatically decrease the intake of exogenous long-chain fatty acids[42]. In comparison with those more stably palmitoylated proteins, “palmitatecentric” approach may be likely bias for proteins that undergo rapid palmitate turnover. Once palmitate analogues are introduced to cellular proteins, it can only be used to analyze protein palmitoylation other than shorter, longer or unsaturated lipid chains. Physiological effects caused by the structural differences between natural fatty acids and fatty acid analogues are not very clear. Most importantly, the metabolic pathway in eukaryotes is extremely complicated, so it may probably result in unpredictable biological effects due to the introduction of palmitate analogues. 5
“Cysteine-centric” approach
Diagrammatic sketch of “cysteine-centric” approach is shown in Fig.3. Acyl-biotinyl exchange (ABE) was first proposed by Drisdel’ group in 2004[43]. In this approach, after free thiols present in proteins were completely blocked by thiol reactive reagents such as idoacetamide (IAA) or N-ethylmaleimide (NEM), the palmitoyl thioester linkage was selectively cleaved by neutral hydroxylamine (HA) and resulted in new free sulfhydryl groups. Consequently, S-acylated proteins could be detected by autoradiography via radioactive NEM labeling, analyzed by the immunoblotting methods, or purified and enriched via avidin/streptavidinbiotin interactions after the proteins were marked with thiol specific reactive reagents such as biotin-conjugated 1-biotinamido-4-[4’-(maleimidomethyl)cyclohexanecarboxamido] butane (Btn-BMCC)[44] or biotin-HPDP (N-[6-(biotinamido)hexyl]3’-(2’-pyridyldithio) propionamide)[45]. Wan et al[46] used biotin-HPDP as biotinylation reagent to purify palmitoylated proteins. Fifty palmitoyl proteins in yeast were identified by
FANG Cai-Yun et al. / Chinese Journal of Analytical Chemistry, 2014, 42(4): 616–622
Fig.3 Diagrammatic sketch of “cysteine-centric” approach Disulfide bonds of proteins were reduced firstly, and free cysteine residues were alkylated. After proteins were digested into peptides, the sample was treated with hydroxylamine to selectively cleave thioesters. The newly exposed cysteine residues were disulfide-bonded to affinity beads and were purified. The eluate was then analyzed by LC-MS
MS including many SNARE proteins, amino acid permeases and signaling proteins. Thioester bond between palmitic acid and protein was sensitive to hydroxylamine treatment, so most palmitoylated peptide ions were markedly increased in abundance after HA treatment, while the MS signal of non-palmitoylated peptide ions had no significant change before and after HA treatment. Thus, by comparing the spectral counts of identified peptides between the +HA sample and -HA sample, we could evaluate the degree of credibility of each palmitoylation site. Acyl-biotinyl exchange chemistry and streptavidin agarose affinity purification were applied to enrich palmitoylated peptides from tubulin by Zhao et al[47]. The results indicated that 11 cysteine residues of the α/β tubulin heterodimer were palmitoylated by using nano-LCMS/MS. Hemsley et al[48] also reported that α-tubulin and AGG2 were palmitoylated in plant. Kang et al[49] identified 68 known and >200 candidate S-acylated proteins from whole rat brain, purified rat synaptosomes and cultured embryonic rat neurons by ABE approach, which included both hydrophobic and hydrophilic proteins, such as neurotransmitter receptors, adhesion molecules, scaffold proteins, and so on. Yang et al[50] reported a proteomics approach (palmitoyl protein identification and site characterization, PalmPISC). In combination with label-free spectral counting quantitation, PalmPISC led to the identification of 67 known and 331 novel candidate S-acylated proteins as well as the localization of 25 known and 143 novel candidate S-acylation sites. Emmer et al[51] identified 124 palmitoylated proteins in Trypanosoma brucei by ABE approach. Except label-free quantitative method, isobaric labeling approaches were also used to distinguish S-acylated proteins from contaminants, in which all samples were pooled and analyzed simultaneously owing to the isobaric tags. The analysis time was reduced, as well as the repeatability and reproducibility were improved. Thus, isobaric labeling
approach was widely used in proteomic research. Zhang et al[52] presented a palmitoylcysteine isolation capture and analysis (or PICA) method, to screen substrates of human DHHC2 in HeLa cells. They quantified the protein profiles between ZDHHC2 knockdown and control HeLa cells by using isotope-coded affinity tag (ICAT) approach. A total of 57 sites were identified from 50 proteins, and CKAP4/p63 was an important substrate of DHHC2, a putative tumor suppressor. After the proteins were blocked by methyl methanethiosulfonate (MMTS), hydrolysed by hydroxylamine, and purified by thiopropyl sepharose, Forrester et al[53] identified the S-palmitoylated proteins in HEK293 cells by Isobaric tags for relative and absolute quantitation (iTRAQ) to differentiate S-palmitoylated peptides and “contaminated peptides”. In total, 93 putative sites of S-acylation on 88 peptides were identified. ABE approach does not need in vivo metabolic labeling, so it can be readily applied to analyze cells (including cancer cells), tissue and body fluid samples. Although great progress of ABE has been made in detecting palmitoylated proteins, there are several problems existing in ABE approach: (1) ABE would result in higher false discovery rates. ABE-based method is thioester bond-specific but not S-palmitoylationspecific, the false discovery rates derived from other modifications, which contains relatively high thioester bonds. Thus, it is critical to block all free sulfhydryl groups in the proteins that would otherwise react with the labeling reagents, so the type, amount and incubation time of the blocking reagents should be empirically tested. The reaction time, concentration of hydroxylamine and pH of the solution should be tested to make sure of complete cleavage of palmitate with hydroxylamine. Complete labeling of the free sulfhydryl groups after palmitate cleavage is very important. To minimize the false-positive results, control experiment (-HA) is usually required. Signal development with streptavidin
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often results in the detection of a small pool of endogenously biotinylated proteins, so appropriate control experiments should be carried out. Otherwise, these factors can result in nearly one-third of the identified proteins being designated as false positive results[8,54]. In comparison with ABE approach, click chemistry produced a significantly lower number of false positives[55]. (2) S-acylated proteins can be labeled with fatty acids other than palmitate, while all fatty acids are removed after ABE chemistry. So the identities of long chain fatty acids attached to proteins of interest cannot be determined directly. However, this problem can potentially be addressed by GC-MS, which allows direct identification of lipids attached to proteins.
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Given the technical difficulties hindering palmitoylation analysis, this modification’s function has been underestimated. In recent years, there is continuously increasing interest and significant progress has been made in protein palmitoylation, especially with the development of MS-based proteomic approaches (such as ABE approach and metabolic labeling with palmitate analogue), so more and more palmitoylated sites have been identified. However, current researches mainly focus on the identification of palmitoylated sites-qualitative analysis, and all methods have their own limitations. So more qualitative and quantitative approaches for protein palmitoylation, especially effective and accurate approaches, needs to be further developed. For example, protein palmitoylation is often found to modify transmembrane or membrane associated proteins, thus the techniques of membrane proteomics can be used to improve proteins’ solubility. Existing methods require typically complicated procedures, cause easily sample loss (especially for low abundant proteins), and lead to relatively high false positive results. To solve these problems, we need to develop new approaches with a simplified experimental procedure or by designing new suitable detergents, new matrices to enrich palmitoylated proteins with high selectivity and efficiency, or new materials for qualitative and quantitative analysis simultaneously. The development of new fluorescent labeling reagents with high sensitivity and selectivity or fatty acid analogues with high metabolic efficiency is also critical to obtain more reliable information for palmitoylated proteins and sites. Live cell imaging may also provide further a perspective tool for dynamic monitoring of complex palmtoylation and insights into its biological importance and function.
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