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Published in final edited form as: Curr Opin Chem Biol. 2021 Jul 29;65:109–117. doi: 10.1016/j.cbpa.2021.06.004

Chemical approaches for investigating site-specific protein S-fatty acylation

Emma H Garst 1,2, Tandrila Das 1,2, Howard C Hang 1,3,*
PMCID: PMC8671186  NIHMSID: NIHMS1721467  PMID: 34333222

Abstract

Protein S-fatty acylation or S-palmitoylation is a reversible and regulated lipid post-translational modification (PTM) in eukaryotes. Loss-of-function mutagenesis studies have suggested important roles for protein S-fatty acylation in many fundamental biological pathways in development, neurobiology and immunity that are also associated with human diseases. However, the hydrophobicity and reversibility of this PTM have made site-specific gain-of-function studies more challenging to investigate. In this review, we summarize recent chemical biology approaches and methods that have enabled site-specific gain-of-function studies of protein S-fatty acylation and the investigation of the mechanisms and significance of this PTM in eukaryotic biology.

Introduction

Lipid post-translational modifications (PTMs) perform many important functions in biology. Protein S-fatty acylation in particular is involved in the regulation of many important biological systems including development[1], neurobiology[2,3], immunity[4,5] and pathogenesis[6]. S-fatty acylation is the attachment of a long-chain fatty acid, frequently a palmitoyl group (C16:0), onto cysteine residues via a thioester bond on proteins. S-fatty acylation has been implicated in a number of important regulatory functions including modulating the cellular localization, membrane partitioning, oligomerization, conformation, stability and activity of proteins[7*]. This reversible fatty acid modification is found across eukaryotes and is regulated by DHHC-palmitoyl acyl transferases (DHHC-PATs) and acyl protein thioesterases (APTs)[7*] (Figure 1), which have been associated with a variety of human diseases[3,8].

Figure 1. Reversible protein S-fatty acylation.

Figure 1.

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S-palmitoylation is a reversible long-chain fatty acid modification that can be added to or removed from cysteine residues by DHHC-PATs and APTs, respectively.

While a significant body of work has been conducted to study the palmitoylation of Ras in vitro and in cells[9-12*], S-fatty acylation remains a challenging and underexplored modification when compared to other post translational modifications. Recent advances in chemical biology, biochemical methods, and proteomics have facilitated the detection and discovery of S-fatty acylated proteins and their sites of modifications[13-15], but the mechanistic and functional analysis of specific protein S-fatty acylation sites is still difficult due to physical properties such as hydrophobicity and reversibility. In this review, we explore the recent developments in chemical biology that have allowed for the study of site-specific S-fatty acylation in vitro and in living cells.

Direct chemical modification

Direct chemical modification is an attractive approach to explore the effects of lipidation on protein function. For example, maleimide-functionalized phospholipids have been used extensively to conjugate soluble proteins to lipid nanoparticles for bioavailability and therapeutic delivery[16,17]. These maleimide functionalized phospholipids have also been used to explore the biophysical properties of chemically lipidated proteins. Lin et al. used the maleimide-functionalized phospholipid 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine (DOPE) to tether H-Ras to a phospholipid bilayer at one or both of its S-palmitoylation sites, Cys181 and Cys184[18] (Figure 2A). Using this method, the researchers demonstrated H-Ras forms dimers with a Kd of approximately 1 x 103 molecules/μm2 on a membrane surface, while at comparable concentrations soluble H-Ras remains a monomer. While this study suggests lipid conjugation and membrane association have a clear effect on Ras oligomerization, a dual chain phospholipid does not closely resemble the native structure of S-palmitoylation. In order to better replicate a natural S-palmitoyl modification, recent work has focused on the direct modification of recombinant protein with single chain lipids.

Figure 2. Lipid analogs used for direct chemical modification in mechanistic studies of S-fatty acylation.

Figure 2.

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A) In order to characterize the biophysical properties of H-Ras monomers on a membrane surface, Lin et al. conjugated H-Ras to a maleimide functionalized phospholipid, MCC-DOPE, which had been incorporated into a lipid bilayer[18]. B) To structurally characterize lipid modified caveolin-3, Kim et al. used Ellman’s reagent followed by octanethiol coupling to functionalize and modify three naturally palmitoylated cysteines with an 8-carbon chain. This protein was then structurally analyzed using solution state NMR[19]. C) To structurally and biophysically characterize lipidated IFITM3, Garst et al. used the palmitoyl lipid analog maleimide palmitate to directly modify the free cysteines of recombinant protein under mild, reducing conditions. Using this method the researchers discovered local structural changes in the lipidated protein and measured the differences in membrane affinity of modified and unmodified protein[20**].

The generation of recombinant lipidated protein through direct chemical modification is challenging in part due to the hydrophobicity of such lipid mimics – outside of a reconstituted membrane environment, the small molecules used to mimic a natural S-palmitoyl modification may be insoluble even in relatively harsh detergents and require solvation in an organic solvent. One method to resolve this has been to focus on short and mid-chain fatty acid mimics that are less hydrophobic and therefore more chemically accessible. A recent study sought to replicate S-fatty acylation of caveolin-3 by cysteine conjugation to octanethiol (Figure 2B)[19]. This was accomplished through activation of three cysteines of interest by 5,5’-dithiobis(2-nitrobenzoic acid) (Ellman’s reagent) followed by coupling with octanethiol to mimic modification with an eight-carbon, medium chain fatty acid. While this method did not produce entirely homogenous lipidated caveolin-3, it did allow for structural analysis of lipidated caveolin-3 and revealed subtle structural changes around the sight of lipidation[19].

More recently, our laboratory employed N-1-hexadecylmaleimide (maleimide palmitate) as a palmitic acid mimic to directly chemically modify the antiviral protein Interferon-induced transmembrane protein 3 (IFITM3) at two conserved cysteines, Cys72 and Cys105 (Figure 2C)[20**]. Because maleimide palmitate is soluble in an aqueous environment (90% water, 10% DMSO) chemical coupling could be achieved under bioorthogonal conditions. Furthermore, this chemical modification was revealed to be an effective S-palmitoylation mimic in molecular dynamics studies[20**]. Using this lipid mimic, the researchers determined that lipidation of IFITM3 changes its local structure and increases the affinity of the evolutionarily conserved amphipathic domain with the lipid bilayer[20**]. While direct chemical modification has proven to be a strong technique in exploring the effect of lipidation on recombinant protein, there are a few notable drawbacks to this method. Because these chemical coupling methods are cysteine dependent, cysteines that are not lipid modified must be removed through mutagenesis. This may be an issue in larger proteins, or proteins that require disulfide bonds for stable folding. Furthermore, the conjugation site must be accessible to chemical modification and cannot be buried in the protein or a lipid membrane. In order to address these issues, the development of more specific methods to explore S-fatty acylation is desirable.

Semi-synthesis of lipidated proteins

Due to the limitations of direct chemical modification, more involved methods for site-specific protein modification are necessary to explore a diverse array of S-fatty acylated proteins of interest. Native chemical ligation (NCL) has been an important tool to selectively incorporate and probe the effect of various post-translational modifications since its development in the mid-nineties[21,22]. Native chemical ligation utilizes the nucleophilic attack of an N-terminal cysteine on a C-terminal thioester to conjugate two peptide fragments with a native peptide bond. Over the last twenty years, there have been many examples of the selective incorporation of lipid PTMs using NCL or the closely related expressed protein ligation (EPL)[23-26]. However, the incorporation of S-fatty acyl modifications is not amenable to NCL due to the labile thioester bond, which can compete with the C-terminal thioester leaving group resulting in a mixture of products (Figure 3A). In order to overcome this limitation, researchers have previously used non-native linkage methods to chemically attach S-fatty acylated lipo-peptides to a protein of interest[27-29]. In a groundbreaking study of palmitoylation regulation and turnover in the cell, S-palmitoylated and N-farnesylated lipo-peptides mimicking the C-terminal domain of N-Ras were synthesized and attached to recombinant protein using cysteine-selective maleimide conjugation (Figure 3B), after which the semisynthetic protein was injected into cells for the study of N-Ras localization and turnover[11**,12*]. Using this method, the researchers were able to uncover a generic system of spatial organization that allows protein acyl transferases (PATs) to redirect proteins via the post-Golgi sorting apparatus. However, in these experiments the native protein is not chemically accessible due to the limitations of NCL. The expansion of protein semi-synthesis with methods like α-keto acid-hydroxylamine (KAHA) ligation[30,31], diselenide-selenoester ligation (DSL)[32,33], and serine/threonine ligation (STL)[34,35] has opened the door to a broader array of protein targets, including S-fatty acylated proteins.

Figure 3. Development of S-fatty acylated proteins using new protein synthesis methods.

Figure 3.

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A) Although NCL is a common method in protein synthesis, the labile thioester linkage in S-fatty acylation can be targeted by the N-terminal cysteine nucleophile, resulting in a mixture of protein products. B) Previously, the Waldmann lab has developed methods to chemically lipidate Ras by conjugating recombinant Ras with a maleimide functionalized S-palmitoylated, N-farnesylated lipo-peptide[27-29]. This method has been used to explore the biology of lipidated Ras in vitro and in cells[9-12*]. Schematic adapted from Rocks et al.[12*] C) The development of a new 1,3 propanedithiol-protected salicylaldehyde (SALPDT) ester tolerates the sequential use of NCL and STL without undesired cross-reactions. This methodology allows for the synthesis of proteins that are S-fatty acylated in the C-terminal domain. Reaction scheme adapted from Huang et al.[38**] D) RBM has previously been developed to solubilize hydrophobic peptides in NCL. However, it is not compatible with STL and the incorporation of a S-palmitoyl modification. In order to overcome this limitation, Huang et al. developed a γ-aminobutyric acid modified RBM group that can be activated under mild acidic conditions[39**]. This chemistry was utilized in the synthesis of natively lipidated sarcolipin, M2, and IFITM3. Reaction scheme adapted from Huang et al.[39**]

Among these alternative methods, STL has proved the most promising for the incorporation of S-fatty acylation analogs. STL was developed after the observation that the 1,2-hydroxylamine functional group from an N-terminal Ser or Thr residue can selectively react with a salicylaldehyde moiety to form an N,O-benzylidene acetal-linked intermediate, followed by acidolysis and formation of a native amide bond[34,35]. Due to its thiol-independent chemistry and the relative abundance of Ser and Thr in nature, STL can be used in conjunction with NCL to synthesize larger proteins and access new, challenging targets[36,37]. More significant to the study of lipid PTMs, STL can be used to conjugate lipo-peptides containing thioester linkages without undesired side reactions. This has recently been demonstrated in the total synthesis of several S-palmitoylated proteins including S-palmitoylated sarcolipin, S-palmitoylated matrix-2 (M2) ion channel, and S-palmitoylated (IFITM3)[38**,39**]. In order to access these S-fatty acylated proteins of interest, a number of new chemical strategies were used, including the development of a new 1,3 propanedithiol-protected salicylaldehyde (SALPDT) ester which allowed for an N-to-C sequential NCL and STL method to ligate three to four peptide fragments (Figure 3C)[38**]. Protection of the second peptide’s C-terminus with an inert SALPDT group allowed for the clean ligation of fragment 1 and 2 using NCL before the removal of the protecting group and subsequent coupling of the new C-terminal SAL moiety with S-palmitoylated fragment 3 through STL (Figure 3C). Using this method, Huang et al. successfully synthesized natively Cys50-S-palmitoylated M2 and Cys105-S-palmitoylated IFITM3 in milligram quantities[38**].

Another technical hurdle in the synthesis of S-palmitoylated proteins has been the physical properties of lipid-modified proteins – namely, the increased hydrophobicity and poor solubility of one or more of the peptide fragments under typical ligation conditions. In order to access lipo-peptides with poor aqueous solubility, Huang et al. developed methods for removable-backbone-modification (RBM)-assisted Ser/Thr ligation[39**]. The RBM method was previously developed to make hydrophobic protein synthesis chemically accessible via NCL by conjugating a hydrophobic peptide to a solubilizing hydrophilic motif such as a charged lysine or arginine polypeptide[40-42]. However, these methods rely on nucleophilic attack to activate the RBM through deacetylation, which can also decompose the palmitoyl thioester linkage and the SAL ester necessary for STL. In order to access the synthesis of proteins containing insoluble, hydrophobic S-palmitoylated lipo-peptide fragments, Huang et al. developed a new strategy where a γ-aminobutyric acid group was added to the RBM, allowing for autocyclization and removal of the acetyl group under weak acidic conditions that do not threaten to decompose the SAL ester or the S-palmitoyl thioester linkage[39**] (Figure 3D). The combination of these chemistries has allowed for the total synthesis of S-palmitoylated sarcolipin and M2 proteins, and was utilized in the synthesis of S-palmitoylated IFITM3 described above. Together, the development of these chemistries allows for the study of natively S-palmitoylated proteins in vitro and opens the door to many structural and mechanistic studies of S-fatty acylation on a molecular level. However, these new protocols have yet to be applied to the mechanistic exploration of S-palmitoylation biology in vitro or in vivo. Further work on these natively modified protein targets will surely reveal new insights into the effect of S-palmitoylation on protein structure and function.

Site-specific lipidation of proteins in living cells

There is a lack of accessible tools for rigorous in cell mechanistic analysis of the role of site-specific S-palmitoylation of proteins. Historically, this modification has been interrogated with loss of function studies by mutagenesis of specific cysteine residues or the knockout or inhibition of DHHC-PATs and depalmitoylases. However, the dynamic nature of the modification makes it more challenging to dissect the effect of site-specific S-palmitoylation in cells. Previously, lipo-protein semi-synthesis and microinjection have been used to study the localization and turnover of S-palmitoylated Ras in cells[11**,12*]. While these pioneering studies of S-palmitoylated proteins have allowed for the exploration of basic protein lipidation biology, the method is limited to proteins that can be introduced to the cell exogenously. S-palmitoylated transmembrane proteins or proteins that are incorporated into larger functional complexes may not be accessible using this method. Furthermore, the use of cellular microinjection is a specialized technique with a relatively high barrier to entry. Considering these limitations, genetic code expansion coupled with copper-free bioorthogonal reactions provides a unique opportunity for stable protein modification in live cells.

Direct chemical modification of proteins and generation of lipo-proteins helped to make significant advances in mechanistic understanding of post translational lipid modifications, but we still lacked methods to make site-specifically modified proteins in cells. Genetic code expansion and bioorthogonal chemistry have been explored in recent years for site specific post translational modification of proteins. Genetic code expansion uses the cellular protein synthesis machinery to generate proteins with unnatural amino acids incorporated site specifically. Genetic code expansion involves using an orthogonal aminoacyl-tRNA synthetase/tRNA pair to incorporate the desired unnatural amino acid (UAA) at a specific site on the protein of interest generally in response to the amber stop codon (UAG) on mRNA[43]. This method has been successfully performed in bacteria, yeast, mammalian cells, worms, flies and mice[44,45]. Genetic code expansion allowed generation of homogenous protein population with site specific incorporation of post translationally modified amino acids[46]. This has been extensively used to study serine and tyrosine phosphorylation, sulfation, and lysine modifications in E. coli and mammalian cells[47,48]. In addition, this method has been used to incorporate short chain fatty acid modified amino acids in proteins[49]. In a recent study, stable incorporation of butyryl-lysine analogs in a chromosomally expressed key transcriptional regulator in S. typhimurium decoded the role of short chain fatty acid acylation in regulating microbial pathogenesis in mice[49].

Further modification of proteins with long chain fatty acids required coupling of genetic code expansion with bioorthogonal chemistry. This involves generation of proteins with unnatural amino acids containing functional groups that can be subsequently labeled with reactive long chain fatty acids using bioorthogonal chemistry (Figure 4A)[50,51]. Bioorthogonal reactants involve chemical moieties that are non-toxic to cells, metabolically stable, and a unique reactive pair in cells. Tremendous efforts in the last decades have led to development of several such reactions which are now being used regularly in vitro, in cells and animals for a plethora of applications[51,52]. The first example of chemical palmitoylation using genetic code expansion and bioorthogonal chemistry is palmitic acid modified recombinant GFP. p-Ethynylphenylalanine was site-specifically incorporated in GFP and further reacted to an azide-containing fatty acid derivative using copper-catalyzed alkyne-azide cycloaddition (CuAAC) reaction[53*]. This method has since been expanded to replicate S-fatty acylation via the conjugation of a fatty acid to the model therapeutic protein urate oxidase (Uox), which is used as a drug for hyperuricemia[54*]. Fatty acid modified Uox has increased binding capacity to Human serum albumin (HSA) which thereby increases its serum half-life in vivo. Conjugation of fatty acid to Uox is not amenable to CuAAC since copper can significantly reduce the enzymatic activity of Uox. Therefore, Cho et al. used strain-promoted azide-alkyne cycloaddition (SPAAC) reaction for the conjugation of a palmitic acid mimic (Figure 4B)[55]. p-azido-L-phenylalanine (AzF) was incorporated in Uox and reacted to a palmitic acid analog containing dibenzocyclooctyne group (DBCO-Pal) to generate fatty acid modified Uox (Uox-112Pal) (Figure 4B). Uox112Pal showed similar specific activity as Uox but higher HSA binding capacity[54*]. While the efficient incorporation of a desired UAA at specific sites can be challenging, this method can be expanded to generate other fatty acid conjugated therapeutic proteins with enhanced serum half-life in vivo.

Figure 4. Genetic code expansion and bioorthogonal labeling for site-specific lipidation of protein of interest.

Figure 4.

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A) Genetic code expansion can be used to site-specifically incorporate an unnatural amino acid (UAA) with unique reactivity. In brief, an orthogonal tRNA/tRNA synthetase pair which can incorporate a specific UAA is developed by directed evolution. This tRNA is specific for a stop codon (generally amber codon). When that codon is inserted into a protein sequence, the unnatural amino acid is incorporated during translation at the specific site. A number of these unnatural amino acids have been utilized including endo/exo-BCNK[20**,56**], 2’-aTCOK[20**,56**], and 4’-a/eTCOK[56**]. The protein of interest with UAA incorporated site specifically can further undergo a chemical coupling reaction with a complementary partner to attach some functional moiety such as a lipid. B) Using genetic code expansion, Cho et al. incorporated the unnatural amino acid p-azido-L-phenylalanine into urate oxidase (Uox)[54*]. This protein was then purified and chemically coupled to a lipid mimic, DBCO-palmitate, using strain-promoted azide-alkyne cycloaddition (SPAAC)[59]. Schematic adapted from Cho et al.[54*] C) In Li et al., genetic code expansion was utilized to express N-Ras in eukaryotic cells with the unnatural amino acid exo-BCNK, which contains a strained alkyne group amenable to in cell bioorthogonal coupling[56**]. To site-specifically lipidate N-Ras, a lipid analog containing a tetrazine reactive group (Tz-6) was applied to the cells which undergoes IEDAC with the protein-incorporated strained alkyne. This gain-of-function lipidation of N-Ras in cells allowed the researchers to explore N-Ras localization and signaling. D) In Garst et al., the methods developed in Li et al. were expanded to the gain-of-function lipidation of transmembrane proteins[20**]. In brief, the UAA exo-BCNK was incorporated into the antiviral protein IFITM3 and coupled to Tz-6 to mimic S-palmitoylation in cells. Site specifically lipidated IFITM3 was then characterized in a number of ways including quantification of the protein activity through viral challenge and imaging protein localization through fluorescent microscopy[20**].

Recent studies from Peng and Hang labs have expanded on previous methods of site-specific protein modification in cells to incorporate lipid modifications on proteins mimicking S-palmitoylation. In order to accomplish this, genetic code expansion was coupled with Inverse-electron-demand Diels–Alder cycloaddition (IEDAC) to label proteins site specifically with palmitic acid mimics in live cells (Figure 4C)[20**,56**]. The extension of these methods to a cellular system involves a number of challenges including stability of a UAA, as well as the solubility, membrane permeability, subcellular distribution, and accumulation of the bioorthogonal fatty acid derivatives. These studies use bioorthogonal coupling reactions with fast second-order rate constants like IEDAC or tetrazine ligation between strained alkynes or alkenes and tetrazines, as they are the most suitable for in cell reaction[57,58]. To optimize this chemistry for in cell usage, Li et al. incorporated different strained alkyne or alkene containing UAAs in N-Ras and used a panel of fatty acid tetrazine derivatives of different lengths[56**]. Tetrazine ligation was the most efficient for strained alkene containing UAA, exo-BCNK and tetrazine derivative, Tz-6 (Figure 4C). Chemical lipidation of N-Ras with Tz-6 increased its membrane association and downstream signaling in cells as evidenced by increased ERK phosphorylation. In comparison, a compound consisting of a tetrazine reactive group without the long chain fatty acid did not increase localization to the cell membrane or ERK phosphorylation, indicating the gain of activity is due to N-Ras lipidation rather than the non-physiological product of the bioorthogonal tetrazine reaction. This study is the first example of gain-in-function on chemical lipidation of proteins in cells and provides an orthogonal way for stable incorporation of a lipid modification to understand its function in cells.

Garst et al. expanded the scope of this method to transmembrane proteins in cells (Figure 4D)[20**]. The Hang lab has investigated the role of IFITM3 Cys72 S-Palmitoylation in its antiviral activity by loss in function studies. They applied this method to chemically lipidate IFITM3 at position 72 by incorporating exo-BCNK and labeling with Tz-6. Chemically lipidated IFITM3 has similar perinuclear localization as endogenous IFITM3 in cells. Moreover, cells expressing chemically lipidated IFITM3 showed increased resistance to Influenza virus infection showing gain in function on lipidation. These studies show that amber suppression coupled with bioorthogonal chemistry can be readily applied to interrogate the site-specific role of protein lipidation in cells, particularly for proteins with multiple sites of modification. With this method, new cellular targets inaccessible by previously developed protocols can be lipid modified and analyzed. Although this method is a strong technique for the site-specific lipid modification of proteins in cells, a significant non-physiological synthetic linkage is incorporated in the process, which could potentially introduce unwanted experimental artifacts. While in the future it may be desirable to improve the chemical mimicry of this synthetic modification, these studies provide an exciting new technology to study the function of protein lipidation in live cells.

Conclusion

S-fatty acylation is an integral part of protein function and regulation across a broad range of eukaryotic proteins. Recent advances in the detection of S-palmitoylation through chemical proteomics and other methods have revealed many proteins with this specific lipid PTM. However, in the past the mechanistic study of S-fatty acylation has been limited by the tools available to study this system. The development of new chemistries has allowed for the exploration of S-palmitoylation in many different contexts.

In recent years, researchers have developed many new tools and techniques to probe S-fatty acylation using chemical biology. In this review we have covered advances in three major areas – direct chemical modification, lipo-protein synthesis, and genetic code expansion paired with bioorthogonal chemistry. These methods have allowed researchers to explore new areas of lipid PTM biology, including the structure of lipidated proteins[19,20**] by direct chemical modification and the activity of lipidated proteins in cells[20**,56**] by gain-of-function lipidation via genetic code expansion and bioorthogonal chemistry. Furthermore, newly developed methods in lipo-protein synthesis have led to the generation of a number of natively S-palmitoylated proteins in vitro, including M2, sarcolipin, and IFITM3[38**,39**]. While these studies did not proceed to functional studies on the S-fatty acylated proteins, the chemical methods developed provide new opportunities to explore the biophysical and biochemical characteristics of S-fatty acylated proteins in vitro.

The development of these tools and techniques has provided new avenues for the exploration of S-palmitoylation biology. While much of the previous work utilizing chemical biology to study S-palmitoylation has focused on Ras, these new approaches allow for the expansion of targets to different and challenging S-fatty acylated proteins including transmembrane proteins and larger proteins. In the future, the expansion of hese methods and their application in novel biological systems will greatly enhance our understanding of lipid post translational modification biology.

ACKNOWLEDGMENTS

E. H. Garst and T. Das were supported by the Tri-Institutional Chemical Biology program through the NIH Chemistry-Biology Training Grant T32 GM115327. This work was supported by the grant NIH-NIGMS R01GM087544. Some schematics were made with the assistance of BioRender.com.

Footnotes

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Declaration of interests

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

REFERENCES

  • 1.Zhang MM, Hang HC: Protein S-palmitoylation in cellular differentiation. Biochem Soc T 2017, 45:275–285. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.Fukata Y, Fukata M: Protein palmitoylation in neuronal development and synaptic plasticity. Nat Rev Neurosci 2010, 11:161–175. [DOI] [PubMed] [Google Scholar]
  • 3.Zaręba-Kozioł M, Figiel I, Bartkowiak-Kaczmarek A, Włodarczyk J: Insights Into Protein S-Palmitoylation in Synaptic Plasticity and Neurological Disorders: Potential and Limitations of Methods for Detection and Analysis. Front Mol Neurosci 2018, 11:175. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4.Das T, Yount JS, Hang HC: Protein S-palmitoylation in immunity. Open Biol 2021, 11:2004–11. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Lin H: Protein cysteine palmitoylation in immunity and inflammation. Febs J 2021. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.Sobocińska J, Roszczenko-Jasińska P, Ciesielska A, Kwiatkowska K: Protein Palmitoylation and Its Role in Bacterial and Viral Infections. Front Immunol 2018, 8:1–19. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • *7. Chamberlain LH, Shipston MJ: The Physiology of Protein S-acylation. Physiol Rev 2015, 95:341–376. Comprehensive overview of protein S-acylation biology.
  • 8.Ko P, Dixon SJ: Protein palmitoylation and cancer. Embo Rep 2018, 19. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.Meister A, Nicolini C, Waldmann H, Kuhlmann Ju, Kerth A, Winter R, Blume A: Insertion of Lipidated Ras Proteins into Lipid Monolayers Studied by Infrared Reflection Absorption Spectroscopy (IRRAS). Biophysj 2006, 91:1388–1401. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 10.Güldenhaupt J, Adigüzel Y, Kuhlmann J, Waldmann H, Kötting C, Gerwert K: Secondary structure of lipidated Ras bound to a lipid bilayer. Febs J 2008, 275:5910–5918. [DOI] [PubMed] [Google Scholar]
  • **11. Rocks O, Peyker A, Kahms M, Verveer PJ, Koerner C, Lumbierres M, Kuhlmann Ju, Waldmann H, Wittinghofer A, Bastiaens PIH: An acylation cycle regulates localization and activity of palmitoylated Ras isoforms. Science (New York, NY) 2005, 307:1746–1752. First introduction of exogenous lipidated protein via protein semi-synthesis and microinjection to control S-palmitoylation in cells.
  • *12. Rocks O, Gerauer M, Vartak N, Koch S, Huang Z-P, Pechlivanis M, Kuhlmann Ju, Brunsveld L, Chandra A, Ellinger B, et al. : The Palmitoylation Machinery Is a Spatially Organizing System for Peripheral Membrane Proteins. Cell 2010, 141:458–471. Continuation of in cell lipidation work achieved in Rocks et al. (2005). Using microinjection of semi-synthetic S-palmitoylated Ras, researchers demonstrate a generic system for the spatial organization of peripheral membrane proteins in cells.
  • 13.Hannoush RN, Sun J: The chemical toolbox for monitoring protein fatty acylation and prenylation. Nature Chemical Biology 2010, 6:498–506. [DOI] [PubMed] [Google Scholar]
  • 14.Hang HC, Linder ME: Exploring Protein Lipidation with Chemical Biology. Chemical Review 2011, 111:6341–6358. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Thinon E, Hang HC: Chemical reporters for exploring protein acylation. Biochem Soc T 2015, 43:253–261. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Roth A, Drummond DC, Conrad F, Hayes ME, Kirpotin DB, Benz CC, Marks JD, Liu B: Anti-CD166 single chain antibody-mediated intracellular delivery of liposomal drugs to prostate cancer cells. Mol Cancer Ther 2007, 6:2737–2746. [DOI] [PubMed] [Google Scholar]
  • 17.Takahara M, Kamiya N: Synthetic Strategies for Artificial Lipidation of Functional Proteins. Chem European J 2020, 26:4645–4655. [DOI] [PubMed] [Google Scholar]
  • 18.Lin WC, Iversen L, Tu HL, Rhodes C, Christensen SM, Iwig JS, Hansen SD, Huang WYC, Groves JT: H-Ras forms dimers on membrane surfaces via a protein-protein interface. Proceedings of the National Academy of Sciences 2014, 111:2996–3001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Kim J-H, Peng D, Schlebach JP, Hadziselimovic A, Sanders CR: Modest Effects of Lipid Modifications on the Structure of Caveolin-3. Biochemistry 2014, 53:4320–4322. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • **20. Garst EH, Lee H, Das T, Bhattacharya S, Percher A, Wiewiora R, Witte IP, Li Y, Peng T, Im W, et al. : Site-Specific Lipidation Enhances IFITM3 Membrane Interactions and Antiviral Activity. ACS Chem Biol 2021, 5:844–856. Development of the S-palmitoyl analog, maleimide-palmitate, allows for the direct chemical modification of free cysteines and structural and biochemical analysis of lipid-modified IFITM3 in vitro. Furthermore, genetic code expansion and bioorthogonal chemistry is used to chemically lipidate IFITM3 in cells for gain-of-function antiviral activity assays.
  • 21.Dawson P, Muir T, Clark-Lewis I, Kent S: Synthesis of proteins by native chemical ligation. Science 1994, 266:776–779. [DOI] [PubMed] [Google Scholar]
  • 22.Agouridas V, Mahdi OE, Diemer V, Cargoët M, Monbaliu J-CM, Melnyk O: Native Chemical Ligation and Extended Methods: Mechanisms, Catalysis, Scope, and Limitations. Chem Rev 2019, 119:7328–7443. [DOI] [PubMed] [Google Scholar]
  • 23.Grogan MJ, Kaizuka Y, Conrad RM, Groves JT, Bertozzi CR: Synthesis of Lipidated Green Fluorescent Protein and Its Incorporation in Supported Lipid Bilayers. J Am Chem Soc 2005, 127:14383–14387. [DOI] [PubMed] [Google Scholar]
  • 24.Gottlieb D, Grunwald C, Nowak C, Kuhlmann J, Waldmann H: Intein-mediated in vitro synthesis of lipidated Ras proteins. Chem Commun 2005, 260–262. [DOI] [PubMed] [Google Scholar]
  • 25.Reulen SWA, Brusselaars WWT, Langereis S, Mulder WJM, Breurken M, Merkx M: Protein–Liposome Conjugates Using Cysteine-Lipids And Native Chemical Ligation. Bioconjugate Chem 2007, 18:590–596. [DOI] [PubMed] [Google Scholar]
  • 26.Palà-Pujadas J, Albericio F, Blanco-Canosa JB: Peptide Ligations by Using Aryloxycarbonyl-o-methylaminoanilides: Chemical Synthesis of Palmitoylated Sonic Hedgehog. Angewandte Chemie Int Ed 2018, 57:16120–16125. [DOI] [PubMed] [Google Scholar]
  • 27.Brunsveld L, Kuhlmann J, Alexandrov K, Wittinghofer A, Goody RS, Waldmann H: Lipidated Ras and Rab Peptides and Proteins—Synthesis, Structure, and Function. Angewandte Chemie Int Ed 2006, 45:6622–6646. [DOI] [PubMed] [Google Scholar]
  • 28.Kuhn K, Owen DJ, Bader B, Wittinghofer A, Kuhlmann J, Waldmann H: Synthesis of Functional Ras Lipoproteins and Fluorescent Derivatives. J Am Chem Soc 2001, 123:1023–1035. [DOI] [PubMed] [Google Scholar]
  • 29.Mejuch T, Waldmann H: Synthesis of Lipidated Proteins. Bioconjugate Chemistry 2016, 8:1771–1783. [DOI] [PubMed] [Google Scholar]
  • 30.Pattabiraman VR, Ogunkoya AO, Bode JW: Chemical Protein Synthesis by Chemoselective α-Ketoacid–Hydroxylamine (KAHA) Ligations with 5-Oxaproline . Angewandte Chemie Int Ed 2012, 51:5114–5118. [DOI] [PubMed] [Google Scholar]
  • 31.Harmand TJ, Murar CE, Bode JW: Protein chemical synthesis by α-ketoacid–hydroxylamine ligation. Nat Protoc 2016, 11:1130–1147. [DOI] [PubMed] [Google Scholar]
  • 32.Mitchell NJ, Malins LR, Liu X, Thompson RE, Chan B, Radom L, Payne RJ: Rapid Additive-Free Selenocystine–Selenoester Peptide Ligation. J Am Chem Soc 2015, 137:14011–14014. [DOI] [PubMed] [Google Scholar]
  • 33.Kulkarni SS, Watson EE, Premdjee B, Conde-Frieboes KW, Payne RJ: Diselenide–selenoester ligation for chemical protein synthesis. Nat Protoc 2019, 14:2229–2257. [DOI] [PubMed] [Google Scholar]
  • 34.Li X, Lam HY, Zhang Y, Chan CK: Salicylaldehyde Ester-Induced Chemoselective Peptide Ligations: Enabling Generation of Natural Peptidic Linkages at the Serine/Threonine Sites. Org Lett 2010, 12:1724–1727. [DOI] [PubMed] [Google Scholar]
  • 35.Zhang Y, Xu C, Lam HY, Lee CL, Li X: Protein chemical synthesis by serine and threonine ligation. Proc National Acad Sci 2013, 110:6657–6662. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36.Lee CL, Liu H, Wong CTT, Chow HY, Li X: Enabling N-to-C Ser/Thr Ligation for Convergent Protein Synthesis via Combining Chemical Ligation Approaches. J Am Chem Soc 2016, 138:10477–10484. [DOI] [PubMed] [Google Scholar]
  • 37.Zheng J-S, Tang S, Qi Y-K, Wang Z-P, Liu L: Chemical synthesis of proteins using peptide hydrazides as thioester surrogates. Nat Protoc 2013, 8:2483–2495. [DOI] [PubMed] [Google Scholar]
  • **38. Huang D, Li Y, Liang J, Yu L, Xue M, Cao X-X, Xiao B, Tian C-L, Liu L, Zheng J-S: The New Salicylaldehyde S , S -Propanedithioacetal Ester Enables N-to-C Sequential Native Chemical Ligation and Ser/Thr Ligation for Chemical Protein Synthesis. J Am Chem Soc 2020, 142:8790–8799. Development of new salicylaldehyde S,S-propanedithioacetal ester protecting group allows for the combination of native chemical ligation and Ser/Thr ligation for the synthesis of S-palmitoylated proteins such as M2 and IFITM3.
  • **39. Huang D, Montigny C, Zheng Y, Beswick V, Li Y, Cao X, Barbot T, Jaxel C, Liang J, Xue M, et al. : Chemical Synthesis of Native S-Palmitoylated Membrane Proteins through Removable-Backbone-Modification-Assisted Ser/Thr Ligation. Angewandte Chemie Int Ed 2020, 59:5178–5184. In order to solubilize hydrophobic peptide fragments during protein synthesis, Huang et al. developed new STL compatible Removable-Backbone-Modification chemistry. Using this method, they synthesized the S-palmitoylated proteins sarcolipin and M2. This chemistry allows for the application of STL protein synthesis to more hydrophobic lipidated and transmembrane proteins.
  • 40.Zheng J-S, He Y, Zuo C, Cai X-Y, Tang S, Wang ZA, Zhang L-H, Tian C-L, Liu L: Robust Chemical Synthesis of Membrane Proteins through a General Method of Removable Backbone Modification. J Am Chem Soc 2016, 138:3553–3561. [DOI] [PubMed] [Google Scholar]
  • 41.Li J-B, Tang S, Zheng J-S, Tian C-L, Liu L: Removable Backbone Modification Method for the Chemical Synthesis of Membrane Proteins. Accounts Chem Res 2017, 50:1143–1153. [DOI] [PubMed] [Google Scholar]
  • 42.Zheng J-S, Yu M, Qi Y-K, Tang S, Shen F, Wang Z-P, Xiao L, Zhang L, Tian C-L, Liu L: Expedient Total Synthesis of Small to Medium-Sized Membrane Proteins via Fmoc Chemistry. J Am Chem Soc 2014, 136:3695–3704. [DOI] [PubMed] [Google Scholar]
  • 43.Xie J, Schultz PG: A chemical toolkit for proteins — an expanded genetic code. Nat Rev Mol Cell Bio 2006, 7:775–782. [DOI] [PubMed] [Google Scholar]
  • 44.Liu W, Brock A, Chen S, Chen S, Schultz PG: Genetic incorporation of unnatural amino acids into proteins in mammalian cells. Nat Methods 2007, 4:239–244. [DOI] [PubMed] [Google Scholar]
  • 45.Ernst RJ, Krogager TP, Maywood ES, Zanchi R, Beránek V, Elliott TS, Barry NP, Hastings MH, Chin JW: Genetic code expansion in the mouse brain. Nat Chem Biol 2016, 12:776–778. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46.Liu CC, Schultz PG: Adding New Chemistries to the Genetic Code. Annual Review of Biochemistry 2010, 79:413–444. [DOI] [PubMed] [Google Scholar]
  • 47.Beránek V, Reinkemeier CD, Zhang MS, Liang AD, Kym G, Chin JW: Genetically Encoded Protein Phosphorylation in Mammalian Cells. Cell Chem Biol 2018, 25:1067–1074.e5. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 48.Italia JS, Peeler JC, Hillenbrand CM, Latour C, Weerapana E, Chatterjee A: Genetically encoded protein sulfation in mammalian cells. Nat Chem Biol 2020, 16:379–382. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 49.Zhang ZJ, Pedicord VA, Peng T, Hang HC: Site-specific acylation of a bacterial virulence regulator attenuates infection. Nature Chemical Biology 2019, 16:95–103. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 50.Lang K, Chin JW: Cellular incorporation of unnatural amino acids and bioorthogonal labeling of proteins. Chemical Reviews 2014, 114:4764–4806. [DOI] [PubMed] [Google Scholar]
  • 51.Lang K, Chin JW: Bioorthogonal Reactions for Labeling Proteins. Acs Chem Biol 2014, 9:16–20. [DOI] [PubMed] [Google Scholar]
  • 52.Sletten EM, Bertozzi CR: Bioorthogonal Chemistry: Fishing for Selectivity in a Sea of Functionality. Angewandte Chemie Int Ed 2009, 48:6974–6998. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • *53. Lim SI, Mizuta Y, Takasu A, Hahn YS, Kim YH, Kwon I: Site-specific fatty acid-conjugation to prolong protein half-life in vivo. J Control Release 2013, 170:219–225. Initial report on using click chemistry for site-specific chemical lipidation of GFP.
  • *54. Cho J, Lim SI, Yang BS, Hahn YS, Kwon I: Generation of therapeutic protein variants with the human serum albumin binding capacity via site-specific fatty acid conjugation. Scientific Reports 2017, 7:18041. The protein urate oxidase (Uox) was chemically lipidated in vitro through recombinant expression of Uox containing the unnatural amino acid p-azido-L-phenylalanine and conjugation to DBCO-palmitate. This protein was analyzed for enzymatic activity and binding to human serum albumin.
  • 55.Debets MF, van Berkel SS, Dommerholt J, Dirks A (Ton) J, Rutjes FPJT, van Delft FL: Bioconjugation with Strained Alkenes and Alkynes. Accounts Chem Res 2011, 44:805–815. [DOI] [PubMed] [Google Scholar]
  • **56. Li Y, Wang S, Chen Y, Li M, Dong X, Hang HC, Peng T: Site-specific chemical fatty-acylation for gain-of-function analysis of protein S -palmitoylation in live cells. Chem Commun 2020, 56:13880–13883. Seminal report on site-specific chemical lipidation of Ras in mammalian cells using genetic code expansion and tetrazine ligation for gain-of-function studies.
  • 57.Šečkutė J, Devaraj NK: Expanding room for tetrazine ligations in the in vivo chemistry toolbox. Curr Opin Chem Biol 2013, 17:761–767. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 58.Selvaraj R, Fox JM: trans-Cyclooctene — a stable, voracious dienophile for bioorthogonal labeling. Current Opinion in Chemical Biology 2013, 17:753–760. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 59.Dommerholt J, Rutjes FPJT, van Delft FL: Strain-Promoted 1,3-Dipolar Cycloaddition of Cycloalkynes and Organic Azides. Top Curr Chem 2016, 374:16. [DOI] [PMC free article] [PubMed] [Google Scholar]

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