Chemoselective and Bioorthogonal Ligation Reactions E-Book

• Herausgeber: Wiley-VCH Verlag GmbH & Co. KGaA
• Kategorie: Wissenschaft und neue Technologien
• Sprache: Englisch

Seitenzahl: 1415

Veröffentlichungsjahr: 2017

Beschreibung

This timely, one-stop reference is the first on an emerging and interdisciplinary topic. Covering both established and recently developed ligation chemistries, the book is divided into two didactic parts: a section that focuses on the details of bioorthogonal and chemoselective ligation reactions at the level of fundamental organic chemistry, and a section that focuses on applications, particularly in the areas of chemical biology, biomaterials, and bioanalysis, highlighting the capabilities and benefits of the ligation reactions. With chapters authored by outstanding scientists who range from trailblazers in the field to young and emerging leaders, this book on a highly interdisciplinary topic will be of great interest for biochemists, biologists, materials scientists, pharmaceutical chemists, organic chemists, and many others.

Leseprobe

Table of Contents

Cover

Title Page

Copyright

List of Contributors

Preface

Contents to Volume 1

Part I: Chemistries

Chapter 1: A Brief Introduction to Traditional Bioconjugate Chemistry

1.1 Introduction

1.2 Reactive Groups of Biomolecules

1.3 Traditional Bioconjugate Reactions

1.4 Cross-Linking Strategies

1.5 Challenges Associated with Traditional Bioconjugate Reactions

1.6 Conclusions

References

Chapter 2: [3+2]-Dipolar Cycloadditions in Bioconjugation

2.1 Introduction

2.2 Copper-Catalyzed Strategies

2.3 Strain-Promoted Cycloaddition

2.4 Future Directions

References

Chapter 3: Diels–Alder and Inverse Diels–Alder Reactions

3.1 Introduction

3.2 Diels–Alder Reaction

3.3 Inverse Diels–Alder Reaction

3.4 Summary and Outlook

References

Chapter 4: The Staudinger Ligation

4.1 Mechanism and Scope of the Classical Staudinger Reaction

4.2 Methodology and Mechanism of the Nontraceless Staudinger Ligation

4.3 Methodology and Mechanism of the Traceless Staudinger Ligation

4.4 Methodology and Mechanism of the Staudinger-Phosphite and Staudinger-Phosphonite Reaction

4.5 Applications of the Staudinger Ligation and its Variants as Bioorthogonal Tools

References

Chapter 5: Thiol–Ene Chemistry

5.1 Introduction

5.2 Mechanism and Stereochemistry

5.3 Reaction Kinetics

5.4 Chemoselectivity and Side Reactions

5.5 Applications and Representative Examples from the Literature

References

Chapter 6: Ligand-Directed Tosyl and Acyl Imidazole Chemistry

6.1 Introduction

6.2 Ligand-Directed Tosyl Chemistry

6.3 Ligand-Directed Acyl Imidazole Chemistry

6.4 Conclusions and Future Directions

References

Chapter 7: Bioorthogonal Labeling of Cellular Proteins by Enzymatic and Related Mechanisms

7.1 Introduction

7.2 Enzymatic Labeling

7.3 Self-Labeling Proteins and Peptides

7.4 Alternate Methods of Protein Labeling

7.5 Conclusions

Acknowledgments

References

Chapter 8: Metal-Mediated Bioconjugation

8.1 Selective Bond Formation on Biomolecules Using Organometallics

8.2 Oxidative Ligations at Tyrosine (Ni, Pd, Ru, Ce)

8.3 Indium-Mediated Ligations

8.4 Reductive Alkylation of Lysine (Ir)

8.5 Metal-Promoted Cysteine Alkylation (Au, Rh)

8.6 Ligations Featuring Rhodium Carbenoids

8.7 Tsuji–Trost Alkylation of Tyrosine (Pd)

8.8 Mizoroki–Heck Reaction (Pd)

8.9 Cross-Coupling at Alkynes (Pd, Cu)

8.10 Suzuki–Miyaura Cross-Coupling (Pd)

8.11 Olefin Metathesis (Ru)

8.12 Prospects in Metal-Mediated Ligations

References

Contents to Volume 2

Part II: Applications

Chapter 9: Protein and Antibody Labeling

9.1 Labeling Single Proteins to Study Intramolecular Conformational Changes

9.2 Monitoring Intermolecular Protein Interactions

9.3 Enzymes and Post-translational Protein Modifications

9.4 Cell Membrane Staining and Tumor Detection

9.5 Protein Labeling for Therapeutic Applications

9.6 Biosensing

9.7 Protein-Based Smart Materials

Summary

References

Chapter 10: Activity-Based Protein Profiling

10.1 Introduction

10.2 Bioorthogonal Chemistry in Activity-Based Protein Profiling

10.3 Selected Applications of Tandem ABPP

10.4 Conclusions and Future Outlook

Acknowledgments

References

Chapter 11: Nucleic Acid Labeling, Ligation, and Modification

11.1 Introduction

11.2 The CuAAC Reaction for Oligonucleotide Labeling

11.3 DNA and RNA Labeling in a Cellular Environment

11.4 Fluorescent Nucleoside Analogs

11.5 Chemical Issues in the Synthesis of Alkyne and Azide Oligonucleotides

11.6 Enzymatic Incorporation of Alkyne dNTPs into DNA

11.7 Artificial Triazole DNA and RNA Backbones

11.8 Enzymatic Methods for Adding Alkyne and Azide Groups into RNA for Labeling

11.9 The SPAAC Reaction on DNA and RNA

11.10 Conclusions

Acknowledgments

References

Chapter 12: Chemoselective Reactions for Glycan Labeling

12.1 Glycoconjugates: Diverse Biopolymers

12.2 Modification to Native Structures

12.3 Metabolic Labeling of Glycans

12.4 Applying Multiple Bioorthogonal Reactions to One System

12.5 Bacterial Glycan Labeling

12.6 Conclusions and Future Outlook

Acknowledgments

References

Chapter 13: Chemoselective Attachment of Lipids to Proteins

13.1 Introduction

13.2 Challenges in Selective Protein Lipidation

13.3 Natively Occurring Lipid Modification of Proteins

13.4 Introducing Phosphatidylethanolamine (PE) into Proteins: The Effects on LC3 and Its Role in Autophagy

13.5 Introducing Glycosylphosphatidylinositol (GPI) Anchors and Their Mimics into Proteins: The Impact of Membrane Attachment on Prion Protein Function

13.6 Lipidation of Small GTPases

13.7 Perspective

Acknowledgment

References

Chapter 14: In Vivo Applications of Bioorthogonal Chemistries

14.1 Introduction

14.2 Methods of Incorporating Bioorthogonal Functional Groups into Living Animals

14.3 Bioorthogonal Chemistries for Imaging and Labeling Biomolecules

In Vivo

14.4 The Future of

In Vivo

Bioorthogonal Chemistry

14.5 Conclusion

References

Chapter 15: Immobilization of Biomolecular Probes for Arrays and Assay: Critical Aspects of Biointerfaces

15.1 Introduction

15.2 Physical and Chemical Strategies for Protein Immobilization

15.3 Surface Parameters Affecting the Performance of Immobilized Proteins

15.4 Evaluation of the Bioimmobilization Process

15.5 Conclusion

References

Chapter 16: Chemical Ligations in the Design of Hydrogel Materials

16.1 Introduction

16.2 Ligation Reactions Suitable for Hydrogel Formation in Biologic Environments

16.3 Effect of Ligation Strategy on Hydrogel Physical Properties

16.4 Future Perspectives

References

Chapter 17: Nanoparticle Bioconjugates: Materials that Benefit from Chemoselective and Bioorthogonal Ligation Chemistries

17.1 Introduction

17.2 Nanoparticle Materials

17.3 Synthesis and Functionalization of Nanoparticles

17.4 Overview of the Bioconjugation of Nanoparticles

17.5 Traditional Bioconjugate Chemistries

17.6 Chemoselective and Bioorthogonal Chemistries

17.7 Conclusions

References

Chapter 18: Application of Engineered Viral Nanoparticles in Materials and Medicine

18.1 Introduction to Virus-Based Materials

18.2 Shape and Structure of Viruses

18.3 Production and Principles of Virus Modification

18.4 Applications of VNPs

18.5 Summary and Outlook

Acknowledgments

References

Index

End User License Agreement

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Guide

Cover

Table of Contents

preface

Begin Reading

List of Illustrations

Chapter 1: A Brief Introduction to Traditional Bioconjugate Chemistry

Figure 1.1 Peptide chains illustrating the structure of the 20 canonical

l

-amino acids, the N-terminus, and the C-terminus. The amino acid residues are linked by stable amide bonds and differ in the structure of their side chains. For each ionizable side chain, the predominant ionization state at pH 7.0 is shown, and the approximate p

K

a

value is listed. Most bioconjugate reactions target functional groups associated with the side chains.

Figure 1.2 Two examples of proteins, (a) human serum albumin (HSA) (Protein Data Bank ID 1AO6) and (b)

E. coli

acyl carrier protein (ACP) (Protein Data Bank ID 1T8K). The structures highlight the abundance of lysine residues (Lys, blue), aspartic acid (Asp, red), and glutamic acid (Glu, red) residues, as well as the scarcity of cysteine residues (Cys), particularly residues with available thiol groups (orange, circled) versus those that are part of disulfide bridges (Cys–Cys, yellow). Note that ACP has no cysteine residues.

Figure 1.3 Reduction of disulfide bonds to thiols using (a) TCEP or (b) DTT.

Figure 1.4 (a) The structures and symbolic notations for (i) glucose (Glc), (ii) glucosamine (GlcN), (iii)

N

-acetylglucosamine (GlcNAc), (iv) glucuronic acid (GlcA), and (v) 6-

O

,2-

N

-disulfated glucosamine (GlcNS6S). (b) The structures of (i) galactose (Gal), (ii) mannose (Man), (iii) xylose (Xyl), (iv) fucose (Fuc), (v) iduronic acid (IdoA), and (vi)

N

-acetylneuraminic acid (Neu5AC, a sialic acid). (c) Structures of (i) lactose, which has Gal and Glc residues joined by a β-1 → 4 glycosidic linkage, and (ii) isomaltose, which has two Glc residues joined by an α-1 → 6 glycosidic linkage. (d) Structures of two glycosaminoglycans, (i) chondroitin and (ii) heparin, and (iii) an example of an N-linked glycan.

Figure 1.5 (a) The reducing end of a carbohydrate exists in equilibrium between cyclic hemiacetal and aldehyde forms. This equilibrium is shown for a glucose residue. The aldehyde group can react with amine nucleophiles (not shown; see Section 1.3.1). (b) The reducing end can be modified to an amine using ammonium carbonate.

Figure 1.6 Oxidation of the vicinal diols in a carbohydrate to aldehyde groups using sodium periodate. Nucleophiles can react with the aldehyde groups (see Section 1.3.1). In this case, the periodate oxidation of a glucose residue is shown.

Figure 1.7 Activation of hydroxyl groups with cyanogen bromide to (a) amine-reactive cyanate esters and (b) amine-reactive cyclic imidocarbonates.

Figure 1.8 Chemical structures of (a) DNA and (b) RNA strands. The structural model in panel (a) shows a ball-and-stick model of a double-stranded DNA helix that is 20 base pairs in length. Hydrogen atoms have been omitted for clarity. Two complementary strands of nucleic acid align antiparallel to one another and hybridize through Watson–Crick base pairing to form the double helix.

Figure 1.9 (a) General structure of a glycerophospholipid. The R group varies between different lipids: (i) phosphatidic acid, (ii) phosphatidylethanolamine (PE), (iii) phosphatidylglycerol (PG), (iv) phosphatidylcholine (PC), (v) phosphatidylserine (PS), and (vi) phosphatidylinositol (PI). (b) Space-filling model of a PC lipid bilayer. Hydrogen atoms have been omitted for clarity. (c) General structure of a sphingolipid. The R

1

groups are fatty acid residues and the R

2

headgroups are similar to those found for phospholipids (e.g., choline, carbohydrates).

Figure 1.10 Various reactive derivatives of fluorescein, a popular fluorescent dye: (i) carboxyfluorescein, which can be activated for reaction with amines; (ii) amine-reactive fluorescein succinimidyl ester; (iii) amine-reactive fluorescein isothiocyanate (FITC); (iv) fluoresceinamine, which can be coupled with activated carboxylic acids; (v) thiol-reactive fluorescein maleimide; and (vi) thiol-reactive fluorescein iodoacetamide.

Figure 1.11 (a) Reaction between a carbonyl (aldehyde or ketone) and a primary amine to form an unstable imine, followed by (b) reduction to a stable secondary amine with sodium cyanoborohydride. (c) Reaction between a carbonyl and a hydrazide to form a hydrazone bond. (d) Reaction between an carbonyl and aminooxy group to form an oxime.

Figure 1.12

N

-(3-Dimethylaminopropyl)-

N

′-ethylcarbodiimide (EDC) activates a carboxylic acid to an

O

-acylisourea intermediate that can react with (a) a primary amine to yield an amide or (b)

N

-hydroxysuccinimide (NHS) or sulfo-NHS to yield a more stable but still reactive succinimidyl ester. (c) The succinimidyl ester reacts with an amine to yield an amide.

Figure 1.13 Reaction between an isothiocyanate and a primary amine to form a thiourea.

Figure 1.14 Reaction between an amine with (a) a cyanate ester to form an isourea and (b) a cyclic imidocarbonate to form an N-substituted imidocarbonate.

Figure 1.15 Michael addition between a maleimide and a thiol to form a stable thioether linkage.

Figure 1.16 Reaction between an iodoacetamide and a thiol to yield a stable thioether.

Figure 1.17 (a) Activation of a thiol group with 2,2′-dipyridyl disulfide. (b) Thiol–disulfide exchange reaction between a pyridyl disulfide derivative and a thiol to form a new disulfide linkage and pyridine-2-thione as a by-product.

Figure 1.18 Reversible formation of a cyclic boronic ester from the reaction between a boronic acid and a

cis

-1,2-diol. The ionization equilibrium for each species is shown. The boronic ester is favored at basic pH.

Figure 1.19 (a) Representative examples of homobifunctional cross-linker structures and reactions: (i) disuccinimidyl glutarate (DSG) and (ii) bismaleimidotriethyleneglycol (BM-PEG

3

). (b) Structure of succinimidyl 4-(

N

-maleimidomethyl)cyclohexane-1-carboxylate (SMCC), a representative example of a heterobifunctional cross-linker, and its cross-linking reaction.

Figure 1.20 Conversion of a carboxylic acid group to (a) an amine group and (b) a thiol group after reduction.

Figure 1.21 Conversion of an amine group to a carboxylic acid group using succinic anhydride.

Figure 1.22 Conversion of an amine group to a thiol group using 2-iminothiolane.

Figure 1.23 (a) Tetrameric ribbon structure of streptavidin bound to four biotin ligands. (b) Close-up view of the biotin-binding pocket of avidin. (c) Structure of (i) biotin and its derivatives: (ii) amine, (iii) hydrazide, (iv) succinimidyl ester, and (v) maleimide. (d) Direct conjugation between biotin and (strept)avidin-modified biomolecules. (e) Indirect conjugation between two biotinylated biomolecules using (strept)avidin.

Chapter 2: [3+2]-Dipolar Cycloadditions in Bioconjugation

Figure 2.1 General pericyclic mechanism for [3+2] dipolar cycloadditions.

Figure 2.2 Thermal and Cu(I)-catalyzed azide–alkyne cycloaddition.

Figure 2.3 The first proposed mechanism of the Cu(I)-catalyzed azide–alkyne cycloaddition.

Figure 2.4 Dinuclear mechanism for the Cu(I)-catalyzed azide–alkyne cycloaddition.

Figure 2.5 Stoichiometric reactions with Cu(I) acetylides.

Figure 2.6 Proposed competing catalytic cycles involving mono- and dinuclear acetylides.

Figure 2.7 Single-electron processes involved in Cu(I)/Cu(II) oxidative cycling.

Figure 2.8 Difference in CuAAC rate due to chelating azides. Conversion reported after 30 min [55].

Figure 2.9 Accelerating ligands for CuAAC applied to bioconjugation.

Figure 2.10 Cell surface labeling using Cu(II)–

l

-histidine.

Figure 2.11 Unwanted disulfide formation during peptide functionalization.

Figure 2.12 Cu(I)-catalyzed coupling between sulfonyl azides and alkynes.

Figure 2.13 Cu(I)-catalyzed sulfonyl azide–alkyne coupling or click sulfonamide reaction (CSR).

Figure 2.14 Alternative synthetic products arising from capture of the ketenamine intermediate.

Figure 2.15 C-terminal protein labeling using native chemical ligation.

Figure 2.16 Labeling of lysine residues using diazo transfer.

Figure 2.17 Regioisomeric products from the SPAAC.

Figure 2.18 Distortion/interaction energy model to explain barrier height.

Figure 2.19 Deconvolution of energy contributions for [3+2] cycloaddition. Energy reported in kcal mol

−1

.

Figure 2.20 Mutually orthogonal reactivity of azides or tetrazines.

Figure 2.21

In vivo

labeling of surface glycans in mice.

Figure 2.22 Potential side reactions of cyclooctynes.

Figure 2.23 General structures of commonly employed SPAAC reagents.

Figure 2.24 Alternative dipoles for strain-promoted bioconjugation reactions.

k

rel

relative to azide dipoles with comparable cyclooctynes.

Figure 2.25 Sequential [3+2]–retro [4+2] cycloaddition with sydnones.

Figure 2.26 Generation of nitrile oxide

in situ

and decomposition pathway via nucleophiles.

Figure 2.27 Condensation equilibria for nitrones and stable cyclic nitrones.

Figure 2.28 Metabolic incorporation of mannosyl-functionalized dipoles.

Figure 2.29 Photoinducible reagents for bioorthogonal labeling.

Figure 2.30 Photo-click reaction of tetrazoles gives a fluorescent product.

Chapter 3: Diels–Alder and Inverse Diels–Alder Reactions

Scheme 3.1 Diels–Alder reaction mechanism.

Figure 3.1 Initial approach for covalent biomodification of oligonucleotides by Diels–Alder cycloaddition in aqueous media.

Figure 3.2 Frontier orbital model of Diels–Alder cycloadditions.

Figure 3.3 Examples of electron-rich dienes used in Diels–Alder bioconjugation.

Figure 3.4 Examples of strained alkenes that are involved in Diels–Alder cycloaddition reactions.

Figure 3.5 Stereoselectivity of Diels–Alder cycloaddition.

Figure 3.6 Diels–Alder bioconjugation and immobilization of diene-modified oligonucleotides.

Figure 3.7 Diels–Alder-mediated approach for immobilizing ligands to benzoquinone-based monolayers.

Figure 3.8 Immobilization of diene-functionalized streptavidin on maleimide-functionalized glass slides.

Figure 3.9 Diels–Alder bioconjugation of oligodeoxyribonucleotides to a cell-penetrating peptide.

Scheme 3.2 Inverse Diels–Alder reaction mechanism.

Figure 3.10 Estimated relative reactivities for dienophiles (top) and tetrazines (bottom) used in inverse Diels–Alder reactions.

Figure 3.11 Frontier orbital model of inverse Diels–Alder cycloadditions.

Scheme 3.3 Synthetic methodologies for tetrazines.

Figure 3.12 Development of novel highly strained bicyclic

trans

-cyclooctenes for tetrazine inverse Diels–Alder additions with large rate constants.

Figure 3.13 Stable cyclopropene mini-tags for inverse Diels–Alder cycloaddition with tetrazines.

Figure 3.14 Mutual orthogonality of inverse Diels–Alder and Michael thiol–ene addition.

Figure 3.15 Radiolabeled biomarker AZD2281-

18

F.

Figure 3.16 Methylcyclopropene tags as small dienophiles for tetrazine inverse Diels–Alder bioconjugation. (a) Live-cell imaging by labeling cyclopropene-modified phospholipids. (b) Metabolic imaging of unnatural mannosamine derivatives on live-cell surfaces.

Figure 3.17 Post-synthetic modification of DNA by inverse Diels–Alder reaction.

Figure 3.18 Oligonucleotide-templated fluorogenic tetrazine inverse Diels–Alder ligation.

Chapter 4: The Staudinger Ligation

Scheme 4.1 General reaction mechanism of the Staudinger reaction.

Scheme 4.2 General mechanism of the nontraceless Staudinger ligation highlighting a range of phosphines investigated.

Scheme 4.3 General mechanism of traceless Staudinger ligation employing phosphine derivatives of type

15

.

Scheme 4.4 General mechanism of the Staudinger-phosphite/Staudinger-phosphonite reaction.

Scheme 4.5 Sequential coupling of two different azide compounds by CuAAC and Staudinger reaction using borane-protected alkyne phosphonites [31].

Scheme 4.6 Incorporation of nonnatural modified monosaccharides into glycans via biosynthetic pathway and subsequent biotin labeling by Staudinger ligation [3].

Scheme 4.7 Specific fluorescence activation

via

Staudinger ligation of (a) coumarin–phosphine fluorogenic probe

34

[37] and (b) quenched phosphine fluorophore

36

[38].

Scheme 4.8 (a) Semisynthesis of RNase A combining the traceless Staudinger ligation and expressed protein ligation [41] and (b) chemoselective synthesis of a cyclic peptide by the traceless Staudinger ligation [21a].

Scheme 4.9 Phosphorylation [22] (a) and site-specific PEGylation [24, 44] (b) site-specific PEGylation [24,42] of proteins using the chemoselective Staudinger-phosphite reaction.

Chapter 5: Thiol–Ene Chemistry

Figure 5.1 Chemical structures of common commercially available thiol and ene monomers.

Scheme 5.1 Thiol–ene addition products for monomers with varying functionality. Monothiol and monoene monomers result in the dimerization product. Dithiol and diene monomers form linear oligomer and/or polymer chains. Monomers with average functionality greater than two result in cross-linked networks.

Scheme 5.2 Initiation and propagation mechanisms for thiol-Michael addition reactions. Base (B)-catalyzed initiation results from a base abstracting a proton from the thiol group generating a thiolate anion and the conjugate acid. Nucleophile (N)-catalyzed initiation results from the acrylate undergoing nucleophilic attack from the phosphine generating a phosphonium-enolate intermediate. The enolate abstracts hydrogen from a thiol group generating the thiolate anion. The propagation mechanism results from the thiolate anion attacking the electron-deficient ene generating an intermediate carbanion or enolate in the case of an acrylate group. The carbon-centered anion subsequently abstracts a proton from another thiol group forming the addition product and regenerating the thiolate anion.

Scheme 5.3 Initiation, propagation, and termination mechanism for thiol–ene radical step growth polymerization. Initiation results from absorption of light by the photoinitiator (I) and subsequent cleavage into primary radical species . Propagation results from cyclical addition of thiyl radicals across ene groups generating the addition product and a carbon-centered radical. The carbon radical abstracts hydrogen from another thiol group regenerating the thiyl radical and forming the thiol–ene addition product. Termination primarily occurs by radical–radical recombination of thiyl radicals, carbon radicals, or thiyl and carbon radicals.

Scheme 5.4 Propagation mechanism for thiol–ene radical polymerizations with homopolymerizable ene functional groups. Thiyl radical addition to the ene functional group and subsequent hydrogen abstraction in the step growth cycle results in the thiol–ene addition product. Carbon radical homopolymerization with another ene functional group results in the chain growth homopolymerization product.

Scheme 5.5 Propagation mechanism for thiol–yne radical polymerizations. Thiyl radical addition to the yne functional group generates a vinyl sulfide radical. Subsequent hydrogen abstraction results in the vinyl sulfide addition product. Thiyl radical addition to the vinyl sulfide group generates a carbon radical and subsequent hydrogen abstraction results in the thiol–vinyl sulfide addition product.

Figure 5.2 Functional group conversion versus time for (a) PETMP and TMPDAE (thiol —, allyl ether −−) polymerized with 1 wt% benzophenone and UV light and (b) PETMP and diethylene glycol diacrylate (thiol —, acrylate −−) polymerized with 0.5 wt% Irgacure 651 and UV light.

Figure 5.3 Chemical structures of common thiol functional groups.

Figure 5.4 Chemical structures of common ene functional groups.

Figure 5.5 Ene functional group conversion versus irradiation time for a 1 : 1 stoichiometric mixture of PETMP and hexanediol dinorbornene (−−), a 1 : 1 stoichiometric mixture of PETMP and DVE (---), and a 1 : 2 stoichiometric mixture of PETMP and hexanediol diacrylate (—). Formulations contained 0.1 wt% Irgacure 651. The thiol–norbornene formulation was irradiated at , the thiol–vinyl ether formulation was irradiated at , and the thiol–acrylate formulation was irradiated at .

Scheme 5.6 Mechanism for reaction of oxygen with the thiol–ene polymerization. Oxygen reacts with the carbon-centered radical forming a peroxy radical. The peroxy radical subsequently abstracts hydrogen from a thiol functional group, generating an alkylhydroperoxide and a reactive thiyl radical.

Figure 5.6 Acrylate functional group conversion versus irradiation time for hexanediol diacrylate and PETMP. Samples are in thickness, are open to ambient oxygen, and contain 0 (—), 25 (−−), and 50 (---) wt% PETMP. All samples contain 5.0 wt% Irgacure 651 and are irradiated at UV light.

Figure 5.7 Loss tangent versus temperature for tricyclodecane dimethanol diacrylate with 0 (—), 30 (−−), and 40 (---) wt% PETMP. Formulations contained 0.1 wt% Irgacure 651 and were irradiated at UV light.

Scheme 5.7 Addition–fragmentation mechanism for allyl sulfides and trithiocarbonates. Thiyl radicals attack the (a) allyl sulfide or (b) trithiocarbonate double bond forming a radical intermediate that subsequently cleaves to reform a new allyl sulfide or trithiocarbonate double bond and thiyl radical. The cleavage and reformation process enables stress relaxation within the network without affecting the overall cross-link density or material properties.

Scheme 5.8 Addition of thiol to maleimide functional groups.

Chapter 6: Ligand-Directed Tosyl and Acyl Imidazole Chemistry

Figure 6.1 The specific labeling of natural proteins. (a) Photoaffinity labeling methods and (b) affinity labeling methods.

Figure 6.2 The general reaction schemes of (a) P-PALM and (b) P-ALM.

Figure 6.3 The general reaction scheme of LDT chemistry. Nu denotes a nucleophilic residue on the protein surface.

Figure 6.4 LDT chemistry-based labeling of CA labeling [24]. (a) Chemical structures of labeling reagents for CA. (b) Purified hCAII was incubated with

1

or

2

in buffer without or with either EZA or GSH at 37 °C. In lane 7, a 1 : 1 conjugate of CAII and Dc dye was used as a standard marker to determine the labeling yields. Samples were subjected to SDS-PAGE and analyzed by in-gel fluorescence imaging (FL) and Coomassie Brilliant Blue (CBB) staining. (c) The labeling reaction in human RBC was detected by SDS-PAGE combined with fluorescence imaging and CBB staining. (d) For

ex vivo

labeling, blood taken from a mouse was incubated with

3

or

4

in buffered saline (pH 7.4) with or without EZA for 20 h. For

in vivo

labeling, mice were intravenously injected with

3

or

4

. Blood was taken from the tail-vein and analyzed by Western blot using SAv–HRP (right) and anti-mouse CAII antibody (right).

Figure 6.5 LDT chemistry-based FKBP12 labeling. (a) Chemical structures of labeling reagents for FKBP12. (b) Time course of labeling yields and (c) initial rates (M h

−1

) by

6

(•),

7

(), and

7+FK506

(♦). ND, not detected. (d) The general reaction scheme of the photo-cross-linking by the photoreactive FKBP12. (e) Photo-cross-linking of endogenous FKBP12 with its interacting protein (transiently expressing FRB-HA) in HeLa cells. All biotinylated proteins were captured and purified with NeutrAvidin beads and subjected to Western blotting analysis.

Figure 6.6 (a) Chemical structure of the quenched LDT (Q-LDT) reagent

9

and (b) the schematic illustration of the Q-LDT-based biosensor.

Figure 6.7

19

F-NMR hCAI biosensor based on LDT chemistry [29]. (a)

19

F-NMR spectra change of purified

19

F-labeled hCAI (100 μM) following addition of AAZ (0, 80, 160 μM). , −62.0 ppm,

19

F-labeled hCAI with a free ligand-binding pocket; •, −63.0 ppm,

19

F-labeled hCAI + AAZ. (b) Magnified images of the active sites of

19

F-labeled hCAI (magenta) and

19

F-labeled hCAI + AAZ (green).

Figure 6.8 (a) The general reaction scheme of ligand-directed acyl imidazole (LDAI) chemistry. (b) The illustrated scheme of one-step construction of caged enzymes by LDAI chemistry and photo-uncaging reaction.

Figure 6.9 LDAI-based hCAI labeling

in vitro

[34]. (a) Chemical structures of labeling reagents for hCAI. (b) In-gel fluorescence analysis of the labeling reaction. hCAI was mixed with

10

,

11

,

12

in HEPES buffer (50 mM, pH 7.2) at 37 °C. In lane 9, a 1 : 1 conjugate of hCAII and Dc dye was used as a standard marker to determine the labeling yields. (c) MALDI-TOF mass analyses of hCAI labeling by

10

or

13

in HEPES buffer (50 mM, pH7.2) at 37 °C for 7 h. , native hCAI; ♦, Dc-labeled hCAI; •, FB-labeled hCAI. (d) Relative labeling ratio of the labeled amino acids of hCAI in the reaction with

10

. (e) Relative labeling ratio of the labeled amino acids of hCAI in the reaction with

13

.

Figure 6.10 One-step construction of caged hCAI [34]. (a) Chemical structure of a caged labeling reagent. (b) Labeling of hCA in human RBC lysate for proof of protein selectivity. The lysate (20-fold diluted with HEPES-buffered saline (HBS, pH 7.4)) was mixed with

10

and incubated at 37 °C for 24 h in the presence or absence of EZA. The left and right image shows the CBB staining and fluorescence image of the SDS-PAGE gel, respectively. (c) MALDI-TOF mass analyses of hCA labeling by

14

and uncaging in RBC lysate (20-fold diluted with HEPES-buffered saline). , hCA; ▾, caged hCA. (d) Enzyme activities of hCA, caged hCA, and uncaged hCA.

Figure 6.11 Endogenous FR labeling and construction of a fluorescent biosensor using LDAI chemistry. (a) Chemical structures of LDAI- and LDT-based labeling reagents. (b) Endogenous FR labeling on live cells. KB cells were treated with

15

or

17

in RPMI 1640 (folate-free, 10% FBS) at 37 °C for 24 h with or without FA. (left) Biotin-blotting analysis using SAv–HRP; (right) Western blotting analysis using a mouse anti-FR antibody and anti-mouse IgG-HRP conjugate. (c) Time-lapse fluorescence images (0 or 480 s) of Fl-modified FR labeled by

16

after addition of FA. Scale bars, 20 µm.

Chapter 7: Bioorthogonal Labeling of Cellular Proteins by Enzymatic and Related Mechanisms

Figure 7.1

Biotinylation of target proteins.

Biotinylation of target proteins is an ATP-dependent reaction in which the BPL facilitates the covalent attachment of the biotin cofactor to a lysine of the target protein. The lysine targeted for biotinylation is identified by the surrounding amino acids, a sequence referred to as the biotin acceptor peptide.

Figure 7.2

Proximity biotinylation of target proteins.

(a) The BPL/BAP system was used to examine protein–protein interactions between FKBP and FRB. Target proteins were expressed with either the BPL (structure A – blue) fusion or the BAP sequence (structure B – purple). (b) In the presence of rapamycin, the BAP-containing protein was labeled with biotin and could subsequently be identified using a fluorophore-conjugated streptavidin reporter. (Fernández-Suárez 2008 [16]. Reproduced with permission of American Chemical Society.)

Figure 7.3

Simultaneous labeling of biotinylated proteins.

BPL proteins from different organisms recognize unique BAP sequences, allowing for simultaneous labeling of two protein targets. Here fluorescent proteins anchored to the outer membrane are labeled with either a yeast or bacterial BAP and then differentially targeted with QDs for visualization. (Chen 2007 [21]. Reproduced with permission of American Chemical Society.)

Figure 7.4

Enzymatic labeling and immobilization via farnesyltransferase.

(A) Schematic of attachment of substrate farnesyl pyrophosphate to a protein bearing the signal sequence, CAAX, recognized by FTase. (B) Immobilization and release of aldehyde-modified GFP with hydrazine-modified agarose beads. (i) Immobilization completed within 45 min in the presence of aniline via hydrazone formation. (ii) Release of modified GFP upon addition of hydroxylamine via oxime ligation after approximately 3 h. (iii) Control ligation using unmodified GFP-CVIA. Images in the top row are bright-field images and images at the bottom are fluorescence microscopy images. Scale bars represent 200 µm. (Rashidian 2012 [26]. Reproduced with permission of American Chemical Society.) (C) GST-CVIA-alkyne (a, b) and GST-CVIA-farnesyl (c, d) immobilization onto PEG-azide-treated glass slides. Spots treated with Alexa Fluor-modified anti-GST antibody and imaged. (Viswanathan 2013 [27]. Reproduced with permission of American Chemical Society.)

Figure 7.5

Formylglycine-generating enzyme.

(a) Schematic of protein modification using FGE. (b) Targeting of anti-HLA-conjugated AAV2

Ald13

nanoparticles to receptor-bearing cells. (a) Schematic representation of the method used to conjugate AAV2

Ald13

nanoparticles with an anti-HLA antibody. (b) Transduction of 293T and HepG2 cells by AAV2

WT

, AAV2

Ald13

, and AAV2

Ald13

conjugated with an isotype control antibody (AAV2

Ald13

-ctrlAb) or AAV2

Ald13

conjugated with an anti-HLA antibody (AAV2

Ald13

-αHLA) to express GFP. The percentage of GFP

+

cells was analyzed by flow cytometry. Error bars represent the standard deviation of the mean from experiments conducted in triplicate (

*

p

< 0.05). (c) GFP fluorescence microscopy images of 293T cells infected by indicated particles. Upper: Bright-field image. Lower: GFP fluorescence. Scale bar represents 50 µm. (Liu 2013 [45]. Reproduced with permission of Wiley.)

Figure 7.6

LplA facilitated labeling of target protein.

(a) Schematic showing both the

wt

lipoylation of target proteins (upper) and the modified reaction in which an alkyl azide chain is added to the target protein (lower). (b) LplA-mediated labeling of a membrane-anchored CFP is shown. Cy3 labeling of the LAP by LplA shows only minimal background in control samples. (Panels (a) and (b): Fernández-Suárez, Baruah 2007 [55]. Reproduced with permission of Nature Publishing Group.) (c) Labeling of cellular proteins using an optimized lipoic acid acceptor peptide (LAP2). NES = nuclear export sequence. CAAX = prenylation tag. NLS = nuclear localization sequence. MAP2 = microtubule-associated protein. (Panel (c): Uttamapinant 2010 [56]. Reproduced with permission of PNAS.)

Figure 7.7

Peroxidase-mediated cross-linking.

(a) Schematic of HRP-catalyzed cross-linking of protein tyrosine side chains. (b) Immobilization of cells to tyrosine cross-linked hydrogels. Photomicrographs of L929 cells seeded to Gela-Ph, Alg-Ph, Alb-Ph, Gela-Ph + Alg-Ph, and Alb-Ph + Alg-Ph gels 4 h after seeding. Density of seeded cells was measured at two time points: 5.0 × 10

5

(4 h) and 5.0 × 10

4

cells/well (72 h). Bars: 100 µm. (Sakai 2010 [67]. Reproduced with permission of American Chemical Society.)

Figure 7.8

PPTase-mediated labeling of target proteins.

(a) Diagram of the PPTase-mediated transfer of a phosphopantheine group of CoA to a serine residue of a target protein. (b) Sequential labeling of

S. cerevisiae

cells expressing a PCP fusion protein at the cell surface were sequentially labeled with CoA-Cy3, CoA-5, and CoA-fluorescein facilitated by the PPTase AcpS. Fluorescence micrographs were taken at defined time intervals after which the cells were washed and allowed to grow under optimal conditions. (Vivero-Pol 2005 [71]. Reproduced with permission of American Chemical Society.)

Figure 7.9

Sortase tethering.

(a) Schematic showing target protein tethering to the cell wall by sortase. Protein containing sorting signal sequence is cleaved by sortase, resulting in a thioester linkage that is subsequently attacked by the terminal amine of a pentaglycine cross-bridge in the peptidoglycan layer resulting in the tethered product and regeneration of the sortase active site. (b) Gly

3

-modified polystyrene beads modified with EGFP in the presence and absence of sortase A. Modification was dependent on the concentration of sortase A present, as well as the presence of Ca

2+

. Observed fluorescence of modified beads was approximately 70-fold higher than unmodified beads or those modified in the absence of sortase. (Parthasarathy 2007 [85]. Reproduced with permission of American Chemical Society.)

Figure 7.10

Sortase labeling and protein modification.

(a) Time-course labeling of surface-expressed ODF-LPETG on HEK 293T cells. Cells were modified with G3-Bt followed by treatment with Sav-Alexa Fluor 488 (top) and G5-EGFP (bottom). For complete Figure showing negative controls and Western blot analysis, see Tanaka

et al

. [98]. (Tanaka 2008 [98]. Reproduced with permission of Wiley.) (b) Schematic demonstrating the modification of target proteins using hydrazinolysis mediated by sortase. (Li 2014 [99]. Reproduced with permission of Wiley.)

Figure 7.11

Transglutaminase-based PEGylation.

(a) General reaction catalyzed by TGase. (b) Site-specific PEGylation of sCT using microbial TGase (mTGase). Altering the ratio of cosolvents ethanol and water resulted in increased control over mTGase labeling specificity. When the reaction was carried out in 50% (v/v) ethanol, mono-PEGylated sCT was obtained with PEG attachment only at Gln20. (Panel (b): Mero 2011 [115]. Reproduced with permission of Elsevier.)

Figure 7.12

Transglutaminase-mediated labeling.

(a) Site-specific conjugation of (DOTA)

n

-decalysine to chCE7agl. Two recognition sites for TGase are located in close proximity in the antibody. However, it was shown that only one DOTA-decalysine was conjugated and was bound to the antibody through both glutamines. (http://journals.plos.org/plosone/article?id=10.1371/journal.pone.0060350 Created under creative commons license: CC BY-SA:3.0 https://creativecommons.org/licenses/by-sa/3.0/) (b) Imaging studies of HeLa cells modified with Alexa 668 cadaverine using gpTGase. HeLa cells were engineered such that a transmembrane moiety (TM) was fused to the C-terminal end of CFP, resulting in CFP being displayed on the surface. Additionally, short peptide Q-tags were fused to the N-terminal end of CFP as gpTGase substrates. A cell suspension was then reacted with gpTGase in the presence of amine-containing Alexa 568 cadaverine substrate. Fluorescence imaging demonstrated cofluorescence of CFP and Alexa 568 cadaverine. (Lin 2006 [103]. Reproduced under permission of American Chemical Society.)

Figure 7.13

Immunoblot imaging using SNAP/CLIP-tag fusions.

(a) Mechanism of the SNAP-tag fusion. (b) Dual labeling of recombinant proteins with SNAP- and CLIP-tag fusions. Two recombinant proteins were labeled

in vitro,

following purification (upper gels) or after the direct separation of cell lysates via SDS-PAGE (lower gels). (Panel (b): Gautier 2008 [135]. Reproduced with permission of Elsevier.)

Figure 7.14

Simultaneous labeling of SNAP/CLIP-tag labeled cells.

(A) Simultaneous labeling of Chinese hamster ovary (CHO) cells with nuclear-localized SNAP-tag fusions (red) and membrane-localized CLIP-tag fusions (green). (B) To differentiate between old and new proteins, the

S. cerevisiae

protein Aga2p was expressed as either a SNAP- or CLIP-tag in separate cell cultures. Self-labeling of fusion proteins was first conducted with Cy3- or Cy5-labeled substrates and then, following a period of growth, a second labeling with fluorescein and Cy3-labeled substrates. Newly synthesized Aga2p protein to be differentiated from older Aga2p proteins labeled with an alternate fluorescent dye in parental cells. (Gautier 2008 [135]. Reproduced with permission of Elsevier.)

Figure 7.15

Microcontact printing of GFP to a gold surface.

Self-assembled monolayers were reacted with benzyl guanosine to functionalize a gold surface. A GFP-SNAP-tag fusion was then captured to the surface in a distinct pattern that had been printed to a gold surface. (Engin 2010 [160]. Reproduced with permission of American Chemical Society.)

Figure 7.16

HaloTag labeling with fluorescent dyes and QDs.

(a) Labeling of HeLa cells with HaloTag system. Cells transiently expressing a p65–HaloTag fusion were incubated with the fluorescently labeled substrates, Panels 1–3. Panels 4–6 are the corresponding bright-field image. (Reproduced from Ref. [164] with permission.) (b) Schematic showing the conjugation of luciferase HaloTag fusions to QDs using halogenated substrates that could readily be synthesized. (Zhang 2006 [166]. Reproduced with permission of Wiley.)

Figure 7.17 Targeted degradation of a luciferase HaloTag fusion. (a) Schematic diagramming the targeted degradation of proteins in the proteasome using hydrophobic HaloTag substrates that mimic a partially denatured protein. (b) Using this technique and the substrates shown, researchers showed a decrease in the luminosity of luciferase. (Neklesa 2011 [170]. Reproduced with permission of Nature Publishing Group.)

Figure 7.18

Affinity-based protein profiling.

(a) Principle of ABPP in which a probe is attached to a chemical tag that has downstream application. (b) Bioorthogonal labeling for ABPP involves an initial probe reaction

in vivo

followed by

in vitro

labeling using either click chemistry or a modified Staudinger ligation reaction to add the probe molecule.

Figure 7.19 Biarsenical dyes and their applications. (A) Overview of biarsenical dyes. Subpanel (a) shows representative labeling of target proteins with the FlAsH and ReAsH dyes whose chemical structures are shown in subpanel (b). Subpanel (c) shows the affinity of the fluorescent dyes for the target tetracysteine sequence based on the sequence of the intervening amino acids. (Scheck 2011 [196]. Reproduced with permission of American Chemical Society.) (B) Target proteins were labeled with the FlAsH tag and then resolved on SDS-PAGE. The addition of the dye molecule did not significantly perturb migration through the gel and could still be visualized. One of the limitations of the biarsenical dyes can also be observed in this Figure as dyes show nonspecific interactions with nontarget proteins in some instances. (Adams 2002 [195]. Reproduced with permission of American Chemical Society.)

Figure 7.20 SpyCatcher/SpyTag and the structure of the CnaB domains. (a) The crystal structure and topology diagram of the

S. pyogenes

protein Spy108 and the initially characterized pili protein that contains an isopeptide bond. The isopeptide bonds are shown as black bars. The N-terminus is colored blue while the C-terminus is red. (Reprinted from Ref. [222] with permission. Copyright AAAS.) (b) The SpyCatcher/SpyTag system uses the CnaB2 domain of the FbaB protein from

S. pyogenes.

The protein is divided into the 13-amino acid SpyTag and 139-amino acid SpyCatcher, both of which can be expressed as fusions to target proteins. (Reprinted from Ref. [221] with permission.)

Figure 7.21

Labeling cell surface proteins with the SpyCatcher/SpyTag system.

HeLa cells expressing a recombinant GFP-ICAM-1-SpyTag fusion (a) were subsequent labeled with the SpyCatcher protein conjugated to Alexa Fluor 555. The individual panels of (b) show GFP localization and labeling with Alexa Fluor 555. The lower panels correspond to controls in which SpyCatcher is not labeled. (Zakeri 2012 [221]. Reproduced with permission of PNAS.)

Figure 7.22

Controlling macromolecular topology.

Positioning the SpyCatcher and SpyTag proteins at specific positions within an elastin-like protein allowed researchers to combine peptides

in vivo

and

in vitro

to form unique protein structures. (Zhang 2013 [225]. Reproduced with permission of American Chemical Society.)

Figure 7.23

Split inteins.

(a) Recombinant proteins with complementary intein sequences at either termini can recombine i

n vivo

or

in vitro

to form a single fusion product. (b) A split-luciferase reporter system for monitoring inducer presence/concentration in living cells was developed using an intein (DnaE) to restore luciferase activity. In the presence of the inducer, the intein excised as the luciferase reformed. The researchers demonstrated that the luciferase signal directly correlated to inducer concentrations. (Ozawa 2001 [231]. Reproduced with permission of American Chemical Society.)

Figure 7.24

Split protein systems.

The upper schematic illustrates the basic concept of the split protein system for examining protein–protein interactions. The target proteins are expressed as fusions to the N- (purple) and C-terminal (yellow) portions of the split protein. As proteins are brought within proximity, the split protein reforms typically serving as a reporter molecule. The proteins at the bottom are some of those reported as successful split protein structures. The N- and C-terminal portions of the protein are shaded as stated earlier. (Shekhawat 2011 [257]. Reproduced with permission of Elsevier.)

Figure 7.25

Coiled-coil interactions.

(a) Helical wheel diagrams for the E/K coils that utilize a repeating heptad sequence on complementary coils. (Crescenzo 2003 [261]. Reproduced with permission of American Chemical Society.) (b) Through engineering of the amino acid sequence of complementary helices, it is possible to induce strong interactions through the formation of covalent bonds or disulfide linkages. (Wang 2014 [262]. Reproduced with permission of American Chemical Society.)

Chapter 8: Metal-Mediated Bioconjugation

Scheme 8.1 Oxidative cross-linking of multimeric proteins.

Scheme 8.2 Oxidative ligation at aromatic residues.

Scheme 8.3 Indium-mediated allylation of aldehydes for bioconjugation.

Scheme 8.4 Reductive alkylation of amines in bioconjugation.

Scheme 8.5 Gold-promoted reaction of thiols and allenes.

Scheme 8.6 Thiol alkylation with rhodium carbenoids.

Scheme 8.7 Kirmse–Doyle reaction in bioconjugation.

Scheme 8.8 Indole alkylation using rhodium carbenoids.

Scheme 8.9 Directed protein modification using metallopeptides.

Scheme 8.10 Tsuji–Trost allylation in bioconjugation.

Scheme 8.11 The Mizoroki–Heck reaction in bioconjugation.

Scheme 8.12 General mechanism for Pd-catalyzed cross-coupling reactions used in bioconjugation.

Scheme 8.13 Copper-free Sonogashira coupling for bioconjugation.

Scheme 8.14 Arylation of terminal alkynes using preformed aryl-Pd(II) reagents.

Scheme 8.15 Suzuki–Miyaura cross-coupling for bioconjugation.

Scheme 8.16 Ru-catalyzed olefin metathesis.

Scheme 8.17 Olefin metathesis in bioconjugation.

Chapter 9: Protein and Antibody Labeling

Figure 9.1 Single protein labeling to interrogate conformational changes. (a) Schematic representation of intramolecular bipartite tetracysteine display of aPP. (b) Structural dependence of bipartite tetracysteine binding sites. Crystallographic B-factor putty rendering for bipartite tetracysteine mutants of aPP (PDB 2BF9).

Figure 9.2 FRET in CaM/peptide complexes. (a) Binding of thioamide-labeled peptides (green) to Cnf-labeled CaM (blue) in the presence of Ca

2+

. (b) An image showing Cnf (pink) at position 100 in calmodulin (CaM

F

F*

100

) and thiophenylalanine (yellow sulfur atom) at position 1 in the pOCNC peptide (pOCNC-F′

1

) in the presence of Ca

2+

ions (purple spheres) with an average 18.9. Å separation of the center of the thioamide from the center of the ε-carbons of the Cnf ring. (c) Fluorescence emission spectra of solutions of 10 μM CaM

F

F*

100

in the presence of increasing concentrations of pOCNC-F′

1

and relative fluorescence of complexes of CaM

F

F*

100

/pOCNC-F′

1

(F

Thio

) to an oxoamide control CaM

F

F*

100

/pOCNC (F

Oxo

).

Figure 9.3 Specific cross-linking (S-CROSS) of interacting proteins. (a) Mechanism of S-CROSS: Cross-linking of CLIP-tag and/or SNAP-tag fusion proteins with bifunctional molecules in which the substrates of the two tags are connected via a fluorophore (either Cy5 or Cy3). (b) Experimental protocol: Pairs of proteins are fused to SNAP-tag and CLIP-tag and coexpressed in mammalian cells. Cells are lysed in the presence of fluorescent bifunctional molecules, and the resulting lysate is then analyzed after SDS-PAGE by in-gel fluorescence imaging. (c) Results of hetero- and homotypic interactions by SDS-PAGE and in-gel fluorescence scanning. (d) Coomassie staining of gel (c). (e) Cross-linking efficiency (mean SD of three independent experiments) determined in (c) versus rapamycin concentration fitted with a sigmoidal dose-response equation.

Figure 9.4 Intermolecular protein labeling to interrogate conformational changes. (a) Schematic representation of intermolecular bipartite tetracysteine display. (b) Monomeric bipartite tetracysteine variants of GCN4 (L20P) bind FlAsH and ReAsH (25 nM) with diminished affinities relative to the wild-type bipartite tetracysteine dimer.

Figure 9.5 Detection and identification schematic of AMPylated substrates with

N

6

-propargyl adenosine-5′-triphosphate (

N

6

pATP).

Figure 9.6 Imaging strategies: (A) Preconjugate method; Con A-PLP-FTZ was performed in solution and purified before adding to the cells. (B)

In vitro

labeling method; cells were first treated with Con A-PLP and then with AF488 hydrazide.

Figure 9.7 Covalin-dependent bioconjugations. (a) General scheme for covalin-dependent bioconjugations. An object represents synthetic probes, biomolecules, beads, or cells. (b) General scheme for the bioconjugation of molecular probes via covalin to cell surfaces. In the first step, cell surfaces could be derivatized either with the SNAP-tag substrate or with the HaloTag substrate, but for reasons of clarity only one of the two orientations is shown.

Figure 9.8 Drawing of amino acids and sites in a protein that can be modified by PEGylation.

Figure 9.9 Sites on human growth hormone that can be modified by either a chemical or enzymatic PEGylation approach.

Figure 9.10 Liposome structures and modifications. (a) Common structural arrangements for lipid-based drug delivery vehicles. (b) Schematic of the structure of a typical liposome with several of the potential covalent modifications depicted.

Figure 9.11 A fluorescent sensor protein for sulfonamides and for Zn

2+

. (a) A biosensor for sulfonamides. In the absence of sulfonamide, the biosensor is in a closed conformation. In the presence of sulfonamide, the equilibrium is shifted toward an open conformation. (b) A biosensor for Zn

2+

. The binding of the intramolecular ligand to HCA is Zn

2+

dependent. In the absence of Zn

2+

, the biosensor is in an open conformation. In the presence of Zn

2+

, the equilibrium shifts toward a closed conformation. This shift should lead to a more efficient FRET between the two fluorophores.

Figure 9.12 Protein-cross-linked hydrogels for the detection and sequestration of heavy metal ions. (a) Pea metallothioneins (PMTs) capture toxic metal ions by condensing to form binding pockets. When these proteins are incorporated as the sole cross-links of a polymeric gel, this conformational change results in a bulk contraction of the material. (b) Comparison of gel size before and after exposure to Cd

2+

ions. This can be used to estimate metal ion concentration and indicate when regeneration of the material is necessary. (c) The gel can be subjected to multiple rounds of reuse using inexpensive chelators (such as EDTA).

Chapter 10: Activity-Based Protein Profiling

Figure 10.1 Activity-based probes and their applications. A probe consisting of a reactive group (triangle), a detection tag (star), and a spacer (line) site-specifically labels a purified enzyme or a targeted fraction of enzymes within a whole proteome, cell, or organism. Subsequently, different techniques can be used for analysis. (a) Straightforward gel-based detection facilitates profiling of sets of enzymes, for example, in different life or disease stages. (b) Pretreatment of the samples with potential inhibitors provides a means for screening against purified enzymes or complex mixtures. (c) Isolation of probe-labeled targets by streptavidin–biotin enrichment allows mass spectrometry-based identification. (d) The use of fluorophores or radiotracers as tags enables visualization by fluorescent microscopy or nuclear tomographic imaging.

Figure 10.2 Schematic picture of the ABPP tandem labeling strategy: a probe with a bioorthogonal handle (half circle) covalently reacts with an enzyme target. In a next step, a tag (star) is chemoselectively introduced by means of the bioorthogonal reaction partner (ball).

Figure 10.3 The Staudinger–Bertozzi ligation in ABPP: an azide-labeled ABP–enzyme complex is treated with a tag–phosphine conjugate. The intermediate aza-ylide rearranges to an amide, resulting in a stably tagged enzyme.

Figure 10.4 Examples of ABPs used in combination with the Staudinger–Bertozzi ligation. (a) The proteasome inhibitor

1

and its azide derivative

2

. Both contain a vinyl sulfone electrophile that reacts with the nucleophilic N-terminal threonine residue of the catalytically active proteasome subunits β1, β2, and β5. (b) The cysteine cathepsin targeting ABP

3

and the natural product E-64 (

4

).

Figure 10.5 Copper-catalyzed click chemistry, in which an azide and an alkyne undergo a 1,3-dipolar cycloaddition under influence of Cu(I). Due to the instability of Cu(I), it is usually generated from CuSO

4

and a reducing agent and stabilized by a ligand. In this example, the azide is part of the ABP and the alkyne is part of the tag, but this can also be the other way around.

Figure 10.6 Nondirected or reactivity probes. These probes only contain a reactive group and a ligation handle. (a) Sulfonyl esters

5

and

6

target enzymes with a variety of different mechanisms. (b) Iodoacetamide probe

7

for reactive cysteine residues and sulfonyl fluoride probe

8

for reactive tyrosine and serine residues.

Figure 10.7 The principle of click chemistry ABPP and 2D-DIGE: after labeling of different samples with the same ABP, two different fluorophores are introduced by tandem labeling. Next, samples are mixed and subjected to 2D gel electrophoresis: isoelectric focusing in one dimension and SDS-PAGE in the other dimension. Enzymes with the same abundance and activity state will yield overlay of fluorescence, while enzymes with differences (indicated by arrowheads) give a higher fluorescence intensity of one or the other fluorophore.

Figure 10.8 (a) The natural product rugulactone (

9

) and the four alkyne-conjugates

10–13

based on its structure. Only probes with an intact α,β-unsaturated ketone (

10

and

11

) showed antibacterial activity. (b) Parts of sequences of ThiD kinase from different bacterial species. The active site cysteine C213 is not modified by the rugulactone probes. Instead, C110 that only occurs in bacteria susceptible to rugulactone forms a covalent adduct with the α,β-unsaturated ketone.

Figure 10.9 Enzyme-generated electrophiles. (a) Upon oxidation of the ethynyl substituent of the naphthalene in ABP

14

by a cytochrome P450 enzyme, a reactive ketene is formed, which reacts with a nucleophile in the P450. Subsequent click chemistry allows detection of the covalent complex. (b) Oxidation of ABP

15

by a monoamine oxidase (MAO) results in a Michael acceptor, which links to the FAD cofactor in the MAO.

Figure 10.10 Photocrosslinking ABPs used in combination with click chemistry. (a) Libraries of hydroxamates with a benzophenone photocrosslinker as a side chain of an amino acid residue. (b) The HDAC inhibitor SAHA (

18

) and an analog containing a benzophenone photocrosslinker (

19

).

Figure 10.11 Pre-clicking of electrophilic ABPs can influence their selectivity toward enzyme targets. (a) The natural product and glycosidase inhibitor cyclophellitol

20

and its probe derivatives

21

and

22

. The latter is clicked onto a BODIPY fluorophore, making it highly selective and sensitive toward glucocerebrosidase. (b) Probe

22

docked into a crystal structure of glucocerebrosidase. The BODIPY moiety makes contacts with a hydrophobic patch near the active site.

Figure 10.12 Pre-clicking of electrophilic ABPs to enable different applications. (a) 4-Chloro-isocoumarin derivative

23

is an ABP for intramembrane serine proteases of the rhomboid family and can be used for profiling with in-gel detection. (b) ABP

24

, a pre-clicked version of

23

, enables exactly timed measurements and has been used to determine the labeling kinetics in different micelle environments.

Figure 10.13 ABPP using strain-promoted click chemistry. Cyclooctyne derivatives can react with azides in the absence of Cu(I). The driving force of the reaction is the release of ring strain in the cyclooctyne. Thiol–yne coupling can occur as a side reaction, which can be suppressed by blocking thiols with reagents such as iodoacetamide.

Figure 10.14 ABPP using the Diels–Alder ligation. A diene (here on the probe–enzyme complex) and a dienophile (here a maleimide derivative) undergo a coupling reaction. It is important to pre-block cysteine residues when using a maleimide as a dienophile.

Figure 10.15 ABPP with combinations of bioorthogonal ligations. (a) Dual labeling of different proteasome subunits. A whole proteome is first incubated with a selective β1 ABP and then with a pan-selective proteasome ABP. In the tagging step, a Staudinger–Bertozzi reagent and a maleimide derivative are reacted with the labeled proteome. (b) Triple labeling of different proteasome subunits. A whole proteome is reacted with ABPs selective for the different proteasome subunits. Tagging is done in two steps: first by incubation with a Staudinger–Bertozzi reagent and a tetrazine derivative, removal of the reagents, then Cu(I)-catalyzed click chemistry with an azide–fluorophore derivative.

Figure 10.16 ABPs incorporating two reactive groups for probing the microenvironment of the active site of enzymes. (a) A vinyl sulfone electrophile targeting the catalytic N-terminal threonine of active proteasome subunits is connected through a spacer with an aromatic azide photocrosslinker. An azide handle enables detection via tandem labeling. (b) A carbamate electrophile that targets the active site serine of fatty acid amide hydrolase is connected through a spacer with a benzophenone photocrosslinker. An alkyne handle can be used for click chemistry-mediated detection.

Figure 10.17 Detection of intracellular probe targets by click chemistry in combination with a cleavable linker. Probe-labeled cells or tissues are lysed and subjected to click chemistry with a biotin tag that incorporates a cleavable linker and a bioorthogonal reaction partner, here an alkyne. Enrichment using a resin with immobilized streptavidin also leads to binding of highly abundant proteins and endogenously biotinylated proteins. Selective cleavage of the ABP-bound target (path A) followed by protease digestion and LC–MS/MS leads to a lower background than standard on-bead digestion. Alternatively, an on-bead digestion (path B) can be performed, followed by selective elution of the ABP-modified peptide to determine the site of modification.

Figure 10.18 Examples of clickable cleavable linkers. (a) A proteasome-targeting epoxyketone is conjugated to a Lev-like cleavable linker (boxed) and an azide, which can be clicked onto a biotin derivative for enrichment and chemoselective release by hydrazine. (b) A vicinal diol cleavable linker (boxed) connected to a biotin on one end and an azide or alkyne on the other end enables enrichment and chemoselective release of ABP targets by treatment with periodate. (c) A cleavable trifunctional tag that incorporates an azide and a rhodamine fluorophore on one end and a biotin on the other end of a diazobenzene cleavable linker (boxed). This linker was used to detect the targets of the heat instable hydroxyderricin-based ABP

31

.

Chapter 11: Nucleic Acid Labeling, Ligation, and Modification

Figure 11.1 (a) Examples of alkyne-modified nucleosides used for incorporation into oligonucleotides by solid-phase synthesis. (b) Examples of azide labels that have been used in click labeling of oligonucleotides by the CuAAC reaction [7, 30–35].

Figure 11.2 Labeling oligonucleotides at the 2′-position of the ribose sugar with fluorescent dyes that are unstable to the conditions of oligonucleotide deprotection [32].

Figure 11.3 (a) For TO (or CyIQ) as the 2′-modification, the U–A base pair blocks excitonic interactions (green, TO or CyIQ; red, TR). (b) TO (green) and TR (red) as two base surrogates allow excitonic interactions that interfere with energy transfer (ET).

Figure 11.4 “Click–click–click” labeling of oligonucleotides. (a) Differential protection of alkynes. (b) Azide-functionalized labels used in this work; benzyl, biotin, fluorescein, dabcyl, and galactose azides [39].

Figure 11.5 Click chemistry to synthesize oligonucleotide probes with various dyes attached internally at the 2′-amino position of 2′-amino-LNA monomers.

Figure 11.6 Synthesis of oligoribonucleotide anandamide conjugates for cell uptake studies.

Figure 11.7 Reverse click labeling using oligonucleotide containing 2′-

O

-mesyl-dT and an alkyne Cy-dye, for example, Cy3B. Conversion of 2′-mesyloxyethyl ribothymidine to 2′-azidoethyl ribothymidine. Conditions: (i) NaN

3

, DMF, 65 °C, 20 h (ii) lkyne Cy-dye: CuSO

4

, sodium ascorbate, tris-hydroxypropyl triazole ligand, DMSO, 55 °C, 2 h to give dye-labeled oligonucleotide.

Figure 11.8 A method for click DNA and RNA labeling in cells and whole animals using 5-ethynyl dU or 5-ethynyl U and a fluorescent azide.

Figure 11.9 Click RNA labeling using T7 RNA polymerase, 5-azidopropyl UTP, and a fluorescent alkyne.

Figure 11.10 Fluorescent nucleobase analogs. (a) Pyrrolo- and furanopyrimidines with low fluorescence quantum yields. (b) Triazole-linked pyrrolopyrimidines. (c) Base pairing properties of fluorescent benzyl triazole-linked pyrrolopyrimidine.

Figure 11.11 Side reaction during oligonucleotide synthesis and deprotection caused by Markovnikov hydration of the ethynyl group of C, 7-deaza-G, and U.

Figure 11.12 (a) Alkynyl dUTP and dCTP that are incorporated efficiently into DNA by KlenTaq. (b) Biotin azide. (c) dT

spin

TP lacks the aromatic group and is incorporated much less efficiently into DNA.

Figure 11.13 Synthesis and PCR amplification of DNA containing an artificial triazole linkage and its biocompatibility in both bacterial and mammalian cells.

Figure 11.14 G-clamp nucleobase compensation for decreased duplex stability caused by the triazole linkage.

Figure 11.15 (a) Intra-strand DNA/RNA ligation. (b) Biocompatible triazole backbone in RNA. (c) Inter-strand click ligation.

Figure 11.16 Synthesis of azide- and alkyne-functionalized RNA and click RNA ligation to produce artificial RNA backbone linkages at the ligation sites.

Figure 11.17

O

-Propargylguanosine-5′-monophosphate for incorporation into RNA at the 5′-end by

in vitro

transcription and ethynyluridine-5′,3′-diphosphate for 3′-RNA labeling after conjugation to FAM azide.

Figure 11.18 RNA 5′- and internal labeling strategy based on the efficient incorporation of an alkyne dinucleotide during transcription.

Figure 11.19 RNA 3′- and internal labeling strategy based on the incorporation of 2′-azide NTPs by a PAP enzyme.

Figure 11.20 (a) Dibenzocyclooctyne (DIBO) oligonucleotide labeling reagents: DIBO

p

-nitrophenyl carbonate and phosphoramidite monomer for 5′-oligonucleotide addition. (b) Click linkage between DNA strands formed from oligonucleotides with 3′- azide and 5′-DIBO. Both regioisomers of the dibenzocyclooctyl triazole are shown [68].

Figure 11.21 (a) Addition of cyclooctyne to RNA during solid-phase synthesis and on-column labeling by SPAAC reaction with azides. (b) Equivalent chemistry using a nitrile oxide labeling reagent and SPNOAC chemistry.

Figure 11.22 Synthesis of BCN reagents for oligonucleotide labeling.

Figure 11.23 Synthesis of aminopropyladenosine monomer for incorporation into oligonucleotides and subsequent labeling with azide or BCN.

Figure 11.24 (a) DIBO reagents for oligonucleotide labeling. (b) BCN and azide reagents for oligonucleotide labeling. (c) DNA cross-linking by the SPAAC reaction using the reagents in (a) and (b) above. (d) Site-specific orthogonal fluorescent labeling of DNA using amide bond formation and SPAAC chemistry.

Chapter 12: Chemoselective Reactions for Glycan Labeling

Figure 12.1

Common glycan structures.

N-linked glycans are attached to Asn residues and include a core structure formed from two GlcNAc and three mannose (Man) residues (enclosed by dashed line); further elaboration produces a diverse set of extended and branched N-linked structures. Mucin-type O-linked glycans are formed by extension from GalNAc residues attached to Ser or Thr residues. Modification of proteins on Ser or Thr residues by GlcNAc results in the O-GlcNAc modification. Gangliosides result upon glycosylation of ceramide residues in lipids and are ultimately capped by sialic acid.

Figure 12.2

Labeling glycans using PAL.

Terminal sialic acid residues are oxidized using periodate to introduce an aldehyde functional group into the sugar. Further reaction through hydrazine or oxime ligation (shown) allows labeling with fluorophores or biotin.

Figure 12.3

Labeling glycans using GAL.

An aldehyde is introduced onto terminal Gal or GalNAc residues through oxidation by galactose oxidase. The aldehyde is then subjected to hydrazine or oxime ligation to label with detection or purification tags.

Figure 12.4

Chemoenzymatic labeling with GalT(Y289L).

Recombinant GalT(Y289L) enzymatically adds Gal or GalNAc analogs containing chemical handles to terminal GlcNAc residues. This reaction can be applied to either O-GlcNAc-modified proteins or GlcNAc-terminated N-glycans.

Figure 12.5

Structures of naturally occurring monosaccharides.

Metabolic labeling approaches often employ analogs of naturally occurring sugars, particularly ManNAc, GalNAc, GlcNAc, fucose, and the most common form of sialic acid,

N

-acetylneuraminic acid (NeuAc).

Figure 12.6

Keto sugars.

The ManNAc analog ManLev (a) incorporates a ketone into the

N

-acyl side chain of ManNAc, which is converted into SiaLev upon metabolism in the cell. The GalNAc analog keto-Gal (b) incorporates a ketone at the C-2 position. Keto-Gal is the sugar component of the nucleotide donor UDP-keto-Gal, which was later developed by the Hsieh-Wilson group for use in chemoenzymatic labeling experiments (see Figure 12.4).

Figure 12.7

Metabolism of sialic acid analogs.

In cells, ManNAc is converted to sialic acid through sequential action by the cytosolic enzymes ManNAc 6-kinase, sialic acid-9-phosphate synthase, and sialic acid-9-phosphate phosphatase. Sialic acid is then activated to CMP–sialic acid. CMP–sialic acid is transported to the Golgi apparatus, where sialic acid is transferred to glycans by sialyltransferases. Analogs may enter the pathway either in place of ManNAc or in the place of sialic acid before conversion to CMP–sialic acid analogs.

Figure 12.8

Metabolism of GalNAc analogs.

Modified GalNAc is metabolized through the GalNAc salvage pathway. In this pathway, GalNAc is incorporated into glycoconjugates through its conversion to the corresponding UDP-GalNAc analog, which is then transferred to serines or threonines of proteins through the action of a polypeptide GalNAc-transferase (ppGalNAcT).

Figure 12.9

Structures of azide-modified monosaccharides.

Azide-modified sugars include, from left to right, the ManNAc analog ManNAz, the GalNAc analog GalNAz, the GlcNAc analog GlcNAz, and the fucose analog azido-fuc.

Figure 12.10

Incorporation of GlcNAc analogs through hexosamine metabolism.

GlcNAc can be metabolized through one of two pathways. In the first pathway (bottom), GlcNAc is converted to UDP-GlcNAc and transferred onto proteins. Alternatively, GalNAc may be converted to GlcNAc through the GalNAc salvage pathway (top). In this pathway, GalNAc is converted to UDP-GalNAc followed by conversion to UDP-GlcNAc through the action of UDP-GlcNAc 4-epimerase. UDP-GalNAz and UDP-GlcNAz are readily interconverted by UDP-GlcNAc 4-epimerase, but UDP-GalNAlk and UDP-GlcNAlk are not.

Figure 12.11

Chemistries for labeling azidosugars.

Three chemistries have been employed to label azide groups incorporated into glycans. (a) The Staudinger ligation involves the reaction between the azidosugar and phosphine probe. (b) The copper-catalyzed azide–alkyne cycloaddition involves reacting azidosugars with an alkyne-labeled probe in the presence of copper, producing a triazole. (c) The copper-free Huisgen [3+2] cycloaddition eliminated the need for copper catalyst by replacing the alkyne with a ring-strained cyclooctyne, resulting in a noncytotoxic reaction for labeling azidosugars.

Figure 12.12

Structures of cyclooctyne variants used in copper-free click chemistry.

A number of variations on the traditional cyclooctyne have been made to improve reaction kinetics of the ring-strain-catalyzed cycloadditions, including DIFO, DIBO, S-DIBO, and BARAC.

Figure 12.13

Alkynyl-modified monosaccharides.

Two classes of alkynyl sugars have been made by appending sugars with either a terminal alkynyl group or a Poc group. Alkynyl analogs include ManNAlk, GlcNAlk, and alkynyl-fuc. Poc analogs include ManPoc, GlcPoc, and GalPoc.

Figure 12.14

The inverse Diels–Alder (DARinv) reaction for alkenes.

(a) In the DARinv reaction, an alkene reacts with a tetrazine-containing probe to form a stable adduct in a reaction orthogonal to the azide–alkyne chemistries. (b) ManNAc has been modified to include an alkene group on its

N

-acyl side chain.

Figure 12.15

Cyclopropene sialic acid analogs.

Two versions of cyclopropene-modified sugars have been developed. In these sugars, methylcyclopropene is incorporated into the C-9 position in sialic acid to make 9-Cp-sialic acid (a) or on the

N

-acyl side chain of ManNAc to make ManNCyc (b).

Figure 12.16

Structures of isonitrile-substituted monosaccharides.

Three monosaccharides containing a primary isonitrile have been synthesized, consisting of the ManNAc analog ManN-

n

-Iso (a), the GlcNAc analog GlcN-

n

-Iso (b), and the GalNAc analog GalN-

n

-Iso (c).

Figure 12.17

Labeling isonitrile sugars.

Isonitrile-containing monosaccharides react with a tetrazine-conjugated probe through a [4+1] cycloaddition to form a stable adduct.

Figure 12.18

Thiol-modified sugars for metabolic labeling.

(a) Thiol-modified ManNAc (ManNTGc) is metabolized to thiol-modified sialic acid (Neu5TGc). (b) 6-Thio-fucose has been used to metabolically tag glycosylated antibodies.

Figure 12.19

A dual-function sialic acid analog.

Sialic acid is modified with azide on the

N

-acyl side chain as well as an alkynyl group at the C-9 position to create one sugar with two modifiable functional groups.

Figure 12.20

Modified bacterial monosaccharides.

Analogs of the bacterial sugars Pse and KDO have been synthesized to include an azide functional group, resulting in AzPse (a) and azide-KDO (b).

Chapter 13: Chemoselective Attachment of Lipids to Proteins

Figure 13.1 Natively occurring lipid modifications of proteins. The most frequent acyl chains are depicted here. (POI – Protein of Interest).

Figure 13.2 Natively occurring C-terminal lipid modifications of proteins. For clarity only a core GPI structure is shown here and for the peptidoglycan a precursor (lipid II) is depicted. (POI – Protein of Interest.)

Figure 13.3 Semisynthesis of a key protein in authophagosome assembly: phosphoethanolamine-anchored LC3. Here two strategies are depicted that eventually lead to very similar LC3-variants that only differ in fatty acid chain length (C

16

vs. C

18

). Both rely on recombinant expression of LC3 (aa 1-114) fused to an intein as well as on the solid phase peptide synthesis (SPPS) of LC3 (aa 115-120) linked to 1,2-dipalmitoyl-

sn

-glycero-3-phosphoethanolamine (DPPE) or 1,2-distearoyl-

sn

-glycero-3-phosphoethanolamine (DSPE, shown above). Differences are highlighted in the grey box that indicates N-terminal fusion to maltose-binding protein (MBP) in order to ensure efficient, soluble expression of the LC3-intein fusion construct by Yang

et al.

and in red, which indicates a light-cleavable solubilization tag for the lipidated synthetic peptide in order to facilitate handling as described by Huang

et al.

[5, 26]. In both cases the additional tags are tracelessly removed either enzymatically or by irradiation with UV light.

Figure 13.4 Two semisynthetic strategies for membrane-anchored PrP. A) Protein trans-splicing generates lipidated PrP upon spontaneous assembly of a functional intein complex from a recombinant N-terminal fusion protein and a synthetic C-terminal fusion part. B) Expressed protein ligation employs a recombinantly produced N-terminal PrP-α-thioester for ligation with the synthetic lipidated peptide carrying an N-terminal cysteine residue.

Figure 13.5 PrP variants carrying a peptidic membrane anchor mimicking a GPI (a) and PrP carrying a synthetic GPI anchor (b) have been analyzed by CD spectroscopy (middle) to determine protein folding. Both variants are predominatly a-helical as expected for cellular PrP. No influence on secondary structure of the lipid modifications is observed in a direct comparison to non-lipidated PrP (triangles). The thioflavin T (ThT) assay indicates aggregation into amyloid fibrils by a change of the fluorescence emission maximum of a dye binding to the fibrils. Both lipidated PrP variants in the presence of liposomes show an increased lag phase and much slower formation of amyloid structures when compared to non-lipidated PrP (black trace).

Figure 13.6 Maleimide ligation of an S-palmitoylated and S-farnesylated synthetic peptide carrying an N-terminal maleimide and C-terminal methyl ester to the small GTPase Ras.

Figure 13.7 S-Palmitoylation dependent Ras trafficking in eukaryotic cells. Thioesterase activity removes palmitoyl from Ras at the plasma membrane. Only farnesylated Ras can rapidly dissociate from any cellular membrane as found in the endoplasmic reticulum ER and become trapped at the Golgi where palmitoylation activity is concentrated within the cell. From there secretory vesicles transport palmitoylated Ras to the inner leaflet of the plasma membrane where Ras works as an important signaling switch. Signaling depends on the nucleotide status of Ras, which is not considered here but a constitutively active Ras:GTP variant is shown. At the plasma membrane the de- and re-palmitoylation cycle starts again.

Figure 13.8 Expressed protein ligation (EPL) of a double geranlygeranylated synthetic peptide carrying an N-terminal cysteine residue with a recombinantly produced Rab-α-thioester.

Figure 13.9 Transport of Rab proteins between cellular membranes depends on their GTPase cycle and interaction with a variety of effector proteins such as guanine nucleotide exchange factor (GEF), GTPase activating protein (GAP) and GDP dissociation inhibitors (GDI).

Chapter 14: In Vivo Applications of Bioorthogonal Chemistries