development and optimization of bioconjugations to probe

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Development and Optimization ofBioconjugations to Probe and Modulate ProteinFunctionChristopher Travis

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DEVELOPMENT AND OPTIMIZATION OF BIOCONJUGATIONS TO PROBE AND

MODULATE PROTEIN FUNCTION

Christopher Richard Travis

Chester Springs, Pennsylvania

A Thesis Presented at the College of William & Mary in Candidacy for Departmental Honors

Department of Chemistry

College of William & Mary

December, 2018

ABSTRACT

Bioconjugate chemistry is a critical field with widespread applications to the visualization,

diagnosis, and treatment of various diseases. Thus, it is crucial to investigate and optimize present

bioconjugation methods, while continuing to develop novel bioconjugations to expand the scope

of the field and provide numerous chemical tools for various applications. This thesis describes

the development and optimization of bioconjugations using unnatural amino acid (UAA)

technology to prepare homogenous, well-defined macromolecular complexes. First, the utilization

of the Glaser-Hay bioconjugation to modulate protein function will be discussed. Next, an

investigation into the aqueous Glaser-Hay reaction mechanism and subsequent optimization of the

bioconjugation will be presented. A novel [2 + 2 + 2] cycloaddition bioconjugation reaction will

then be presented, followed by efforts to generate a multivalent bioconjugate. Finally, efforts to

assay and modulate the activity of Cas9 will be examined. This thesis aims to extend the chemical

toolbox to probe and control biological systems, with applications in the fields of medicine and

pharmaceuticals.

i

TABLE OF CONTENTS

Acknowledgements iii

Table of Figure iv

Table of Tables vi

Chapter 1: Introduction to Bioconjugates and Unnatural Amino Acids…………………… 1

Bioconjugates…………………………………………………………………... 1

Unnatural Amino Acids………………………………………………………… 3

Conclusion…………………………………………………………………....... 9

References…………………………………………………………………........ 10

Chapter 2: Utilization of Alkyne Bioconjugations to Modulate Protein Function………… 14

Introduction…………………………………………………………………………….. 14

Incorporation of UAAs into the Chromophore of GFP…………………………… 15

Investigation of the Impact of Glaser-Hay Bioconjugations on the

Fluorescence Profile of GFP……………………………………………………… 18

Conclusion……………………………………………………………………………… 21

Materials and Methods……………………………………………………………….. 22

References……………………………………………………………………………… 23

Chapter 3: Mechanistic Investigation into the Aqueous Glaser-Hay Bioconjugation……... 27

Introduction…………………………………………………………………………….. 27

Investigating the Aqueous Mechanism of the Glaser-Hay Coupling……………. 28

Optimization of the Biological Glaser-Hay Coupling…………………………….. 35

Streamlining the Glaser-Hay Bioconjugation……………………………………… 37

Conclusion………………………………………………………………………………. 39

Materials and Methods………………………………………………………………... 40

References………………………………………………………………………………. 44

Chapter 4: Synthesis of a Novel Dipropargyl Amine UAA and Development of a

Novel [2 + 2 + 2] Cyclotrimerization Bioconjugation……………………….. 48

Introduction…………………………………………………………………………….. 48

Development of Physiologically Compatible [2 + 2 + 2] Cyclotrimerization… 50

Synthesis of UAA with Dipropargyl Functionality………………………………… 50

Site-Specific Incorporation of pDPrAF……………………………………………... 51

Development and Optimization of Biological [2 + 2 + 2] Cyclotrimerization... 52

Investigation of the Versatility of pDPrAF…………………………………………. 55

Conclusion………………………………………………………………………………. 57


References………………………………………………………………………………. 62

ii

Chapter 5: Towards the Development of Multivalent Bioconjugates……………………... 66

Introduction…………………………………………………………………………….. 66

Towards the Development of a Biological 1,3-Dipolar Cycloaddition

of an Azide and a Diyne…………………………………………………………….. 69

Towards the Development of a Terminal Alkyne Addition to a Diyne

in Biological Settings………………………………………………………………... 71

Towards the Use of a Biological Sonogashira Reaction to Prepare a

Multivalent Bioconjugate…………………………………………………………… 72

Conclusion………………………………………………………………………………. 74


References………………………………………………………………………………. 79

Chapter 6: Utilization of Unnatural Amino Acids to Probe CRISPR/Cas9………………...81

Introduction…………………………………………………………………………….. 81

Previous Work………………………………………………………………………….. 86

Expression of Wild Type and Mutant Cas9 and dCas9…………………………… 87

Cleavage Assay………………………………………………………………………….89

Conclusion………………………………………………………………………………. 92


References ………………………………………………………………………………. 95

iii

ACKNOWLEDGEMENTS

I would first like to thank Dr. Doug Young for his mentorship and guidance over the past three

and a half years. Thank you for helping me develop as a scientist and figure out what I want to do

with the rest of my life. Thank you for always tolerating my negativity and for taking me to New

Orleans.

I am also grateful to Dr. Lisa Landino, Dr. Rob Hinkle, and Dr. Margaret Saha for serving on my

thesis committee and for teaching excellent courses.

I would also like to thank Johnathan Maza for teaching me my way around the lab when I was a

freshman and answering all of my terrible questions. I am also grateful to Zack Nimmo for his

friendship as well as his collaboration and leadership on several projects we worked on together.

Additionally, I would like to acknowledge the Young Lab as a whole for making my time in the

lab so memorable and full of good times. A special thank you to Gillian Gaunt, Lauren Mazur,

Christina Howard, and Emily Peairs for their collaboration on several of the projects detailed in

this thesis.

Thank you to Diya for always being there for me. You’re the GOAT.

iv

TABLE OF FIGURES

1.1 Nonspecific conjugation between an antibody and a drug 2

1.2 The twenty canonical amino acids 4

2.1 Overview of site-specific UAA incorporation 5

2.2 Mechanism to incorporate UAA into polypeptide chain site-specifically 7

1.5 Evolution of specific aaRS through double sieve selection 8

1.6 Library of UAAs 9

2.1 Structures of UAAs and site of incorporation into GFP chromophore 16

2.2 SDS-PAGE of UAAs incorporated into GFP 17

2.3 Fluorescence profiles of GFP mutants 17

2.4 Structure of GFP chromophore with UAAs incorporated 18

2.5 Glaser-Hay bioconjugations performed at GFP chromophore 18

2.6 SDS-PAGE of Glaser-Hay bioconjugations 19

2.7 Fluorescence profiles of GFP after undergoing Glaser-Hay couplings 20

2.8 Comparison of Glaser-Hay and Cadiot-Chodkiewicz biocojugations 21

3.1 Glaser-Hay reaction 27

3.2 Biological Glaser-Hay coupling 28

3.3 Proposed mechanisms for Glaser-Hay reaction in organic solvents 29

3.4 Dimerization of propargyl alcohol via Glaser-Hay coupling 29

3.5 13C NMR of aqueous Glaser-Hay reaction time course 30

3.6 Effects of varying propargyl alcohol concentration in an aqueous

Glaser-Hay coupling 32

3.7 Effects of varying copper(I) iodide concentration in an aqueous


3.8 Effects of varying TMEDA concentration in an aqueous


3.9 UV/Vis spectrum of aqueous Glaser-Hay coupling 34

3.10 Structures of ligands employed in Glaser-Hay bioconjugations 36

3.11 SDS-PAGE of Glaser-Hay bioconjugations investigating catalase and ligands 37

3.12 Overview of various starting points for Glaser-Hay bioconjugation 38

3.13 SDS-PAGE of Glaser-Hay bioconjugations during protein purification

and on cell lysate 39

4.1 Antibody-drug conjugates 48

4.2 Example of current bioconjugation employed 48

4.3 Recently developed aqueous [2 + 2 + 2] cycloaddion 49

4.4 Structure of dicarboxylated biphenyl ligand employed in bioconjugations 50

4.5 Physiologically compatible [2 + 2 + 2] cycloaddition 50

4.6 Synthesis of pDPrAF UAA 50

4.7 SDS-PAGE of mutant GFP containing pDPrAF 51

4.8 [2 + 2 + 2] cyclotrimerization bioconjugation 53

4.9 Time course of [2 + 2 + 2] cyclotrimerization bioconjugation 54

4.10 Effects of temperature on [2 + 2 + 2] cyclotrimerization bioconjugation 55

v

4.11 Effects of pH on [2 + 2 + 2] cyclotrimerization bioconjugation 56

4.12 SDS-PAGE demonstrating versatility of pDPrAF 56

5.1 Preparation of a multivalent bioconjugate 67

5.2 Potential reactivity of a biological 1,3-diyne 68

5.3 Published copper(I)-catalyzed reaction between an azide and a 1,3-diyne 69

5.4 Attempted copper(I)-catalyzed reaction between an azide and a 1,3-diyne 69

5.5 Glaser-Hay coupling to prepare a simple 1,3-diyne 70

5.6 Aqueous reaction between an azide and a 1,3-diyne with a

copper-iron catalyst system 70

5.7 Addition of terminal alkyne to internal alkyne via a copper-palladium

catalyst system in aqueous solution 71

5.8 Addition of terminal alkyne to a 1,3-diyne via a copper-palladium catalyst

system in aqueous solution 71

5.9 Biological addition of a terminal alkyne to a 1,3-diyne via a copper-palladium

catalyst system 72

5.10 Copper click reaction to afford bromotriazole-containing bioconjugate

and biological Sonogashira to generate a multivalent bioconjugate 73

6.1 Mechanism of CRISPR/Cas9 in bacteria 82

6.2 Crystal structure of Cas9 82

6.3 Structural depiction of clefts in Cas9 83

6.4 Mechanism to program Cas9 for site-specific endonuclease activity 84

6.5 Sites for site-specific mutagenesis for eventual UAA incorporation 86

6.6 Sequencing data of site-specific mutagenesis experiments 87

6.7 SDS-PAGE of wild type Cas9 and dCas9 expression 88

6.8 Structure of UAAs used for incorporation into Cas9 and dCas9 88

6.9 SDS-PAGE of mutant Cas9 containing pAzF in position 1265 88

6.10 Agarose gel of PCR amplification of EGFP-gRNA7 template DNA 90

6.11 Plasmid map of pIRG 90

6.12 Agarose gel of supercoiled and linearized pIRG 91

6.13 Agarose gel of wild type Cas9 cleavage assay 91

vi

TABLE OF TABLES

5.1 Conditions tested in biological Sonogashira reactions 73

6.1 Rationale behind selection of sites for UAA incorporation in Cas9 86

6.2 Conditions tested in efforts to express mutant Cas9 and dCas9 89

1

CHAPTER 1: INTRODUCTION TO BIOCONJUGATES AND UNNATURAL AMINO

ACIDS

Bioconjugates

Bioconjugate chemistry is a critical area of present research requiring the integration of

chemical tools into biological systems, with far-reaching applications in the fields of medicine,

pharmaceuticals, and materials. A bioconjugate is a covalent linkage between two molecules, at

least one of which is a biomolecule such as a protein, carbohydrate, or oligonucleotide.1,2 Most

often, these biomolecules are reacted with a fluorescent probe, surface, small molecule, or other

biomolecule to result in the bioconjugation product. This secondary reaction partner often confers

novel functionality to the biomolecule, such as fluorescence in instances of conjugation with a

fluorescent probe or potency towards inhibition of a target protein in instances of conjugation with

a small molecule drug. Biomolecules with enhanced properties through bioconjugation have

applications including therapeutics and diagnostics.3-7

Protein bioconjugates, in which at least one reaction partner is a protein, represent an

especially important class of bioconjugates, as they have been employed in a number of critical

applications, most notably for the preparation of antibody-drug conjugates.2 Antibody-drug

conjugates represent a key subset of current cancer treatments, as they combine the potency of a

small molecule inhibitor while utilizing the antibody’s specificity in helping to localize the drug

to its target for inhibition.8

The design and structure of a bioconjugate relate directly to its function.9,10 A variety of

methodologies are commonly employed to prepare bioconjugates. The naturally occurring

reactivity of thiol or amine functional groups in biomolecules are often utilized for bioconjugation

2

reactions.10 In protein bioconjugates, the native nucleophilic characteristics of lysine, serine, or

cysteine are exploited for reaction with a second molecule.

Despite the stability of the covalent bonds

that link together the two components of

bioconjugates, it is difficult to regulate the

precise location on the biomolecule where the

bioconjugation reaction will take place.1,2 This

is due to the fact that multiple sites within the

same protein are often capable of reacting in the

same fashion. For example, multiple lysine, serine, or cysteine residues in a protein could react,

which would afford a heterogenous mixture of bioconjugation products (Figure 1.1). This is

particularly problematic when working to prepare antibody-drug conjugates. Immunoglobulin G

(IgG) is the most common type of antibody found in the human body, and it is often employed in

the preparation of bioconjugates through conjugation at native lysine or cysteine sites on IgG.11

However, this antibody contains 80 lysine residues and 14 cysteine disulfide pairs, meaning that

bioconjugation results in a highly heterogenous product mixture, with the number and location of

conjugated drug molecules varying tremendously. This means that the prepared antibody-drug

conjugate complexes would vary significantly in potency, stability, and solubility, leading to

inconsistent and potentially dangerous treatment for patients. Purifying a heterogenous mixture of

the IgG-drug bioconjugates to yield a specific complex is possible, but difficult and time-

consuming.

Thus, research to develop mechanisms to regulate the bioconjugation site for covalent

linkage is at the forefront of the field, as this generates well-defined conjugates. Such regulation

Figure 1.1. Utilization of the nucleophilicity of native

lysine, serine, or cysteine residues to prepare

bioconjugates often results in heterogenous product

mixtures.

3

affords increased sample homogeneity, enhanced complex stability, and minimized perturbation

of biological function of the biomolecule.9,12-16 However, site-specific conjugations are often

difficult to achieve, as they require additional manipulation of the biomolecule. The conjugation

reaction must not only occur under physiological conditions (37°C, pH ~ 7), but also occur

bioorthogonally due to the wide range of chemical functionality of biomolecules, particularly

proteins.17 Bioorthogonal reactions are reactions that occur on biomolecules, but do not interfere

with biological processes. Additionally, the conjugation reaction must be site-specific, to ensure

that the site of the bioconjugation does not interfere with key sites on the biomolecule, such as the

active site of a protein. One excellent way to bioorthogonally prepare protein bioconjugates is to

utilize unnatural amino acid (UAA) technology, in which an UAA is incorporated site-specifically

into a protein.18 UAAs are not only site-specific, but also offer unique reactivity not found naturally

in proteins, as UAAs often introduce novel chemical moieties. This offers a unique, bioorthogonal

reactive site in proteins which can be utilized to prepare well-defined, homogenous, stable protein

bioconjugates. One commonly employed bioconjugation that utilizes the unique reactivity of

UAAs is the copper-catalyzed azide alkyne cycloaddition (CuAAC), or copper click reaction.10

Through this, a protein harboring an azide-containing UAA is reacted with an alkyne to generate

a bioconjugate.

Unnatural Amino Acids

Proteins are critical for cellular life, as they catalyze an incredible array of cellular

reactions.19 Aptly coined the workhorses of the cell, proteins are critical to cellular structure,

function, and regulation. Although there are many unique proteins catalyzing many unique

chemical reactions, the building blocks of all proteins are chemically simple. The twenty canonical

amino acids, which are those found naturally in cells, consist of just five different elements and

4

have very limited chemical functionality. The 20 canonical amino acids all consist of the same

backbone, containing an amine group, a chiral carbon, and a carboxylic acid group. Thus, these

amino acids differ only in their R group (Figure 1.2). Beyond several acidic and basic amino acid

R groups, there is limited potential for chemical reactivity in the canonical amino acids.

Given the prevalence of protein reactions in cells, expansion of the reactivity of amino acid

R groups is valuable. Unnatural amino acids (UAAs) are synthetic analogs of amino acids, and

can afford novel chemical functionality outside the realm of the twenty natural amino acids.20,21

UAAs are encoded at the DNA level, and allow the researcher to site-specifically dictate the exact

location where the UAA is incorporated in the protein sequence (Figure 1.3). This ensures optimal

UAA placement with minimized detrimental alterations to the protein’s structure and function.

Given the simple two carbon backbone of amino acids, UAA synthesis is relatively simple,

as it involves protection of the carboxylic acid functional group and amine functional group,

Figure 1.2. There are 20 amino acids found naturally in cells, but they have very limited chemical

functionality. Adapted from Penn State Dept. of Biology.

5

followed by reaction of one of the endogenous amino acids

to alter the R-group, thus forming an UAA. Through such

reactions, UAAs have been designed and synthesized to

contain a number of different functional groups, including

alkynes, azides, fluorophores, photocleavable groups,

cyanides, and a multitude of others.21

While these UAAs can be synthesized organically, their

incorporation into proteins proves more difficult. Ultimately,

UAAs can be site-specifically incorporated into proteins in

vivo, meaning that their incorporation must occur under

physiological conditions.20 The Central Dogma of biology

involves the transcription of DNA to form RNA, and the

translation of RNA to form proteins.19 DNA is the hereditary

material of the cell, and thus it contains the blueprints for each protein the cell synthesizes. DNA

has a double helix structure with two complementary strands, each of which contains a series of

nucleotides linked with phosphodiester bonds. There are four types of DNA nucleotides: adenine

(A), thymine (T), cytosine (C) and guanine (G). RNA nucleotides are the only differ in that Uracil

(U) replaces Thymine (T). RNA nucleotides are read and ultimately translated in groups of three,

with each codon (a unique set of three nucleotides) corresponding to a specific amino acid.19

Given that there are 64 possible codons, but only 20 endogenous amino acids, the genetic

codon is simultaneously specific and redundant. In this sense, each amino acid may be represented

by multiple codons (multiple sets of three RNA nucleotides), but each codon is specific to only

Figure 1.3. UAAs can be site-

specifically incorporated into proteins

co-translationally using a two plasmid

system and the cell’s endogenous

translational machinery.23

6

one amino acid. The degeneracy of the genetic code can be exploited in order to site-specifically

incorporate UAAs.

Three codons out of the 64 total are designated stop codons, meaning that they do not code

for the addition of an amino acid, but rather just signal the end of translation (the end of additional

amino acids being added to the polypeptide chain).19 Given the redundancy of the stop codon,

UAAs can be incorporated into proteins by hijacking the endogenous mechanism of protein

synthesis through the Schultz methodology. This process requires an orthogonal aminoacyl-tRNA

synthetase (aaRS), an orthogonal tRNA-codon pair, and an UAA.23,24

This methodology relies on the multiplicity of stop codons and the fact that they do not

code for a particular amino acid. In this, a particular stop codon (often the TAG codon) is

manipulated in order to encode for the UAA rather than the termination of translation. To do this

site-specifically, the DNA of the protein must be mutated to change a specific preexisting codon

to a stop codon at the exact location where the UAA is to be incorporated.23,25 However, for the

UAA to actually be incorporated at the suppressed stop codon, there must exist an aaRS/tRNA

pair that is capable of recognizing the suppressed codon and the UAA.

In cells, there are 20 aaRSs which are specific to each of the canonical 20 amino acids.

These aaRSs are responsible for the catalyzing the linkage of the specific amino acid to the

appropriate tRNA. There is no cross-reactivity between synthetases.19 Thus, if just the UAA was

added to cells, there would be no incorporation of the UAA into protein at the suppressed stop

codon, as there would be no aaRS/tRNA pair to recognize the UAA.23,25 In order to successfully

incorporate an UAA, an aaRS/tRNA pair must be imported and evolved from another organism,

in this case Methanocaldococcus jannaschii, an archaea.26 The exogenous orthogonal aaRS/tRNA

pair is analogous in structure and function but does not interfere with the endogenous mechanism,

7

as there is no cross reactivity. Instead, this orthogonal pair allows for complete translational read-

through of the suppressed TAG stop codon (also known as the amber stop codon) and the

subsequent site-specific incorporation of the UAA (Figure 1.4).20,27

The process of generating and

evolving a specific aaRS to recognize and

ultimately aid in the incorporation of a

particular UAA is complex.28-30 However,

specificity can ultimately be achieved using

a double-sieve selection.29 In this, the

natural aaRS that recognizes tyrosine found

in M. jannaschii undergoes genetic

mutation to produce a library of around 106

– 107 mutant aaRSs, which differ in

particular amino acid residues in the site

where the amino acid binds. These mutations alter the chemical environment of the binding pocket

and thus affect the binding affinity of different amino acids to the aaRS. To determine which aaRS

from the library is most effective at incorporating a particular UAA, a positive selection is

performed first. This selection involves the co-transformation of the mutated aaRSs and a

chloramphenicol-resistant enzyme altered to contain a TAG mutation in E. coli. Cells are then

plated in the presence of chloramphenicol and the particular UAA. Those cells which did not

undergo successful transformation fail to grow as they do not produce a chloramphenicol-resistant

enzyme and do not degrade the chloramphenicol. Thus the corresponding aaRSs are eliminated

from the library.

Figure 1.4. Site-specific UAA incorporation allows for the

introduction of novel chemical functional groups into

proteins. The Schultz Methodology takes advantage of the

redundancy of the genetic code to incorporate the UAA via

hijacking the TAG stop codon.12

8

However, cell growth at this stage does not indicate UAA incorporation, as it is possible

that the new aaRS binding site mutation has resulted in that its charging of a different canonical

amino acid to the suppressor tRNA.29 Thus, negative selection is performed to elucidate which

aaRSs selectively bind the UAA. This negative selection process begins with the isolation and

purification of plasmids from the cells grown during positive selection. These plasmids are then

co-transformed with the barnase gene, which is mutated to contain three TAG codons. Barnase is

a ribonuclease that degrades cellular RNA, thus preventing protein synthesis and causing cell

death.

Following growth of these transformed cells in the absence of UAA, some cells are seen

to still suppress the TAG codons in the barnase gene, thus producing the barnase protein and dying.

These cells must be suppressing the TAG codons through aaRSs that pair endogenous amino acids

to the suppressor tRNA. Therefore, the cells that do not die must contain aaRSs that do not pair

endogenous amino acids to the suppressor tRNA. These results coupled with the fact that these

aaRSs cleared the positive selection indicate that these synthetases specifically charge the UAA of

interest to the suppressor tRNA. Usually, multiple rounds of positive and negative selection are

Figure 1.5. A series of positive and negative selection steps results in the selection of a specific

aminoacyl-tRNA synthetase from a library of 106 -10

7 different mutations.

9

performed to ensure that the optimum aaRS/tRNA pair is chosen to selectively incorporate the

UAA (Figure 1.5).

As described above, UAA technology is highly specific and the site-specific incorporation

of UAAs allows for the introduction of novel chemical functional groups into proteins in controlled

locations. Large libraries of UAAs with diverse chemical handles have been synthesized and

incorporated into proteins (Figure 1.6). Site-specific UAA incorporation has been successfully

employed in many studies of protein structure and function, as well as in studies which altered and

enhanced protein function. Furthermore, UAA incorporation has shown value in the development

of novel diagnostic tools and therapeutics.28

Conclusion

Unnatural amino acid technology is a critical tool used in the preparation of protein

bioconjugates. By site-specifically incorporating synthetic UAAs with unique chemical handles

not found elsewhere in the protein, bioorthogonal conjugation reactions can be achieved,

Figure 1.6. The wide variety of synthesized UAAs allow scientists to incorporate novel

chemical functionalities that are not found in the 20 canonical amino acids into proteins.

Adapted from Liu, et al. (2010).

10

generating specific, homogenous bioconjugates. This thesis will describe the beneficial

applications of the recently developed Glaser-Hay bioconjugation, which utilizes an alkyne-

containing UAA. Then, the mechanism and kinetics of the aqueous Glaser-Hay reaction will be

investigated, and efforts made to optimize the bioconjugation to reduce protein oxidation. Next, a

novel bioconjugation reaction will be reported for the first time: a cyclotrimerization utilizing a

newly synthesized dipropargyl amine UAA. Then, first steps towards the preparation of

multivalent bioconjugates will be discussed. Lastly, work to probe the protein Cas9 with UAAs

will be discussed. All projects aim to provide advances in the fields of bioconjugate chemistry and

UAA technology, as work at the intersection of these fields has critical applications to therapeutics,

diagnostics, and more.

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21. Liu, C., & Schultz, P.G. (2010). Adding new chemistries to the genetic code. Annu. Rev.

Biochem., 79, 413-44.

22. Maza, J.C., Jacobs, T.H., Uthappa, D.M., & Young, D.D. (2016). Employing unnatural

amino acids in the preparation of bioconjugates. Synlett 27, A–I.

23. Cornish, V. & Schultz, P.G. (1995). Site directed mutagenesis with an expanded genetic

code. Annu. Rev. Biophys. Biomol. Struct., 24, 435-462.

24. Wang, L., Xie, J., and Schultz, P.G. (2006). Expanding the genetic code. Annu. Rev.

Biophys Biomol. Struct., 35, 225-249.

25. Martin, A. & Schultz, P.G. (1999). Opportunities at the interface of chemistry and biology.

Trends Cell Biol., 9, M24-M28.

26. Wang, L, Brock, A., Hererich, B., & Schultz, P.G. (2001). Expanding the genetic code of

E. coli. Science, 292, 498-500.

13

27. Mehl, R., et al. (2003). Generation of a bacterium with a 21 amino acid genetic code. J. Am

Chem. Soc., 125, 935-939.

28. Young, D.D. & Schultz, P.G. (2018). Playing with the molecules of life. ACS Chem. Biol.,

13, 854-870.

29. Wang, L. & Schultz, P.G. (2001). A general approach for the generation of orthogonal

tRNAs. Chem. Biol., 8, 883-890.

30. Wang, Q., Parrish, A., and Wang, L. (2009). Expanding the genetic code for biological

studies. Chemistry and Biology, 16, 323-336.

14

CHAPTER 2: UTILIZATION OF ALKYNE BIOCONJUGATIONS TO

MODULATE PROTEIN FUNCTION

Introduction

Protein engineering is a powerful tool for the development of new therapeutics, catalysts,

and biosensors.1-8 While many advances in the field have been made, designing novel protein

functionality is still a challenge, as it requires an intricate understanding of the subtle interplay

between protein structure and function. Current engineering techniques often focus on using

selections and screens to optimize or enhance existing protein functionality.2 While this method

has proved useful for optimizing existing function, generating new function where it does not exist

is still a hurdle.

By introducing novel chemical functionality to proteins, UAAs have allowed for the

development of unique protein function; however, the evolved proteins are often limited to a single

new function, depending on the UAA incorporated.2,9 In addition, the functionality is limited to

the UAA itself, which suffers from constraints, such as the requisite for an aminoacyl tRNA

synthetase capable of recognizing the UAA, the synthetic accessibility of the UAA, and the size

of the UAA which may preclude its uptake by a biological system.10 A more appealing strategy

would allow for the generation of a UAA-containing protein “template” upon which researchers

could synthetically introduce different chemical moieties that would in turn lead to altered protein

function depending on the moiety employed.

Bioorthogonal chemistry, which employs reactions that proceed to completion under

physiological conditions (pH ~7, 37C), offers a unique mechanism to add new chemical

functionality to proteins.11,12 Indeed, a variety of reactions have been developed that can add new

chemistry to living systems. In particular, the cycloaddition between azides and alkynes that is

15

either copper(I) mediated or strain promoted, has become a widespread technique to introduce new

chemistry to proteins.13-16 More recently, our group has developed a biorthogonal variant of the

Glaser-Hay reaction, which brings together two terminal alkynes to form a diyne on a protein in

the presence of copper(I) under physiological conditions.17-21 The resulting stable diyne linkage

has a well-defined linear geometry, and due to the abundance of commercial available terminal

alkynes, a variety of chemical moieties can be reacted onto a protein using this technique. As such,

we sought to utilize the power of this new chemistry to generate new and different protein function

dependent upon the alkyne reaction partner and not purely the UAA.

Incorporation of UAAs into the Chromophore of GFP

Specifically, we designed a proof-of-concept experiment to alter the function of green

fluorescent protein (GFP) via reaction of different terminal alkynes, onto the GFP chromophore.

GFP is a 27 kDa protein isolated from Aequorea victoria with photochemical properties arising

from an internal chromophore composed of Ser65-Tyr66-Gly67.22-24 New chemical properties

afforded by UAA introduction in place of Tyr66 have already been documented to alter GFPs

fluorescence profile.25 All UAAs incorporated were found to blueshift the fluorescence profile of

GFP, with more highly conjugated UAAs exhibiting a greater degree of spectral shifting. Based

on these results, the ability to modulate the conjugation of GFPs fluorophore using UAA

mutagenesis is apparent, and represents a convenient means to rationally design new protein

function.25,26 However, this mutagenesis approach is limited by the size and complexity of the

UAA. An alternative approach involves exploiting the chemical functionality in pre-existing

UAAs to serve as functional handles for biorthogonal reactions, acting as a “template” for the

chemical derivatization of new protein function.

16

This chemically

templated protein function

could be achieved via the

genetic incorporation of p-

propargyloxyphenyalanine

(pPrF, 1) or p-

ethynylphenylalanine (pEtF,

2) into residue 66 of GFP

(Figure 2.1).27 This provides a

terminal alkyne handle for

reaction with different

chemical moieties via the bioorthogonal Glaser-Hay reaction. The resulting diyne linkage is highly

conjugated and represents a prime candidate to introduce new photochemical properties into GFP’s

fluorophore without the need to evolve a new aaRS or express multiple versions of the protein

containing different UAAs. Moreover, the pEtF (2) is directly conjugated with the aromatic ring

of the UAA, allowing for a comparison of the conjugation between the different UAAs. We

hypothesized that the altered conjugation and chemical properties around the fluorophore will lead

to new photophysical properties, demonstrating the utility of a chemically programmable protein

engineering strategy. Herein we report our findings on utilizing the Glaser-Hay reaction on GFP’s

fluorophore to alter its fluorescent properties.

In order to obtain protein possessing an alkynyl moiety, a GFP plasmid harboring a TAG

mutation at position 66 was co-transformed with the polyspecific pCNF-aaRS/tRNA pair.28

Conveniently, this aaRS is capable of recognizing both 1 and 2 and for expression of an alkyne-

17

containing GFP.20 Incorporation was also confirmed by SDS-PAGE analysis of protein expression

in the presence and absence of the various UAAs (Figure 2.2). As the alkynyl UAA is incorporated

into position 66 in GFP’s fluorophore, the extended conjugation afforded by the UAA alters the

spectral properties of GFP. To assess that pPrF-GFPTAG66 was successfully produced, spectra for

the GFP-variant were compared to the wild type. These spectra exhibited blue-shift, in agreement

with the literature precedent, in the pPrF variant relative to the wild type (Figure 2.3). A similar

expression was performed using the pCNF aaRS/tRNA pair and pEtF to produce a separate GFP

mutant with a bioconjugation handle. Conveniently, due to the modularity of this approach, only

a single protein expression is

necessary and all functional

modification can be achieved

synthetically. This is in contrast

to previous experiments, which

required an individual protein

expression for each UAA in

order to modify function.

18

Investigation of Impact of Glaser-Hay Reactions on the Fluorescence Profile of GFP

With a pPrF-GFPTAG66 and pEtF-GFPTAG66 in hand,

we then sought to employ our previously reported

biorthogonal Glaser-Hay reaction to install new and varied

chemical functionality into the chromophore of GFP

(Figure 2.4). To investigate, we performed biorthogonal

Glaser-Hay reactions on the chromophore’s alkyne handle

to couple terminal alkyne-bearing aliphatic and aromatic

compounds with different chemical functionalities. Glaser-

Hay reactions were performed by using a working

concentration of 500 mM of CuI and TMEDA in the

presence of alkyne-UAA bearing GFPTAG66 and the cognate alkynyl partner (Figure 2.5). Reactions

proceeded for 4 hours at 4C and then purified via centrifugation with a molecular weight cut-off

column. The protein was

placed in phosphate-

buffered saline solution

(pH ~7.2) for analysis

using fluorescence

spectroscopy.

Gratifyingly, our initial

attempts with to couple the

terminal alkynes to the

fluorophore were

19

successful. Furthermore, the different characteristics of the alkyne moieties installed successfully

shifted the fluorescence profile away from the parental alkynyl-GFPTAG66 spectra, each in a unique

way. While it might be expected that the requisite for the reaction to occur within the -barrel of

GFP may hinder this reaction from occurring, we hypothesize that the hydrophobic nature of

interior of GFP actually aided in the hydrophobic alkyne localization, thereby facilitating the

reaction by increasing effective concentration. Additionally, SDS-PAGE analysis with Coomassie

revealed that the Glaser-Hay reaction only minimally altered protein concentration suggesting only

minimal protein degradation (Figure 2.6).

We found that our initial

Glaser-Hay reactions on pPrF-

GFPTAG66 had different effects

on the fluorescent profile of GFP

(Figure 2.7, A). Reacting 1-

hexyne (3) on the chromophore

caused a general broadening and quasi-red shift of the fluorescent spectra. Reacting propargyl

amine (4) on the chromophore caused a slight band broadening, as well as potential increase in

fluorescence intensity, perhaps due to the increased polarity of the introduced amine group.

Interestingly, coupling with an aromatic alkyne resulted in a dramatic red-shift of the fluorescence

to above that of wild-type GFP. Both ethynylanisole (5) and ethynylaniline (6), resulted in

excitation spectra maxima above 540 nm, dramatically altering the fluorescence of GFP.

We next sought to explore the effects of reacting terminal alkynes in direct conjugation

with the aromatic ring of residue 66. This was feasible with the GFP mutant harboring 2.

Interestingly, this strategy resulted in an even greater blue shift of the pEtF-GFPTAG66 compared to

20

both the pPrF and wild-type

variants, likely due to the

increased conjugation of the

direct attachment of that

terminal alkyne on the phenyl

ring. As a result of this shift, a

different excitation wavelength

was necessary, as 395 nm was

found to no longer excite the

pEtF-containing chromophore.

Based on absorption

experiments, we selected 280

nm as the wavelength to excite

the pEtF-GFPTAG66 and all its

Glaser-Hay derivatives. In the

same fashion as the pPrF, the

biorthogonal Glaser-Hay was

performed on pEtF-GFPTAG66 using the same reaction partners (Figure 2.7, B). Once again, 3 was

found to broaden the fluorescence spectra. Interestingly, 4 had a drastic red-shift relative to the

pEtF parent chromophore. We believe this helps validate our initial speculation that the polarity

of the amine has a drastic impact on the fluorescent properties of the chromophore, as in this

instance the whole system is in direct conjugation. Interestingly, when employing the aromatic

alkynes in the fluorophore modulation, a less dramatic effect was observed than with the pPrF

21

mutants. Reaction with 5 only slightly red-shifted the spectra; however, 6 had a more significant

impact both on the intensity and the red-shifting of the fluorophore. Additionally, attempts to

repeat the experiments using the bromo-alkyne derivative of 2 under Cadiot-Chodkiewicz coupling

conditions resulted in the identical spectra, but were performed under more mild reaction

conditions (Figure 2.8). These results were expected as the final products of both the Glaser-Hay

or Cadiot-Chodkiewicz reactions are identical.20 This represents a viable alternative reaction to

these protein modification approaches.

Conclusion

In conclusion, we have extended our work on the biological Glaser-Hay to utilize the

biorthogonal chemistry to modulate protein function. Using two previously reported alkyne-

containing UAAs within the chromophore of GFP (position 66), we have successfully performed

the Glaser-Hay reaction on the chromophore of GFP. The resulting diyne linkage alters the

fluorescence profile of GFP depending on the moiety attached to the terminal alkyne. Our future

work seeks to extend the reaction to aromatic containing alkynes, which we hope will have a

greater impact on GFP fluorescence due to the increased conjugation found in an aromatic system.

22

Our findings highlight the potential of biorthogonal chemistry, particularly diyne forming

chemistries, to modulate protein function without the need for tedious selections and screens.

Materials and Methods

General

Solvents and reagents were obtained from either Sigma-Aldrich or Fisher Scientific and used

without further purification, unless noted. Reactions were conducted under ambient atmosphere

with non-distilled solvents. Unnatural amino acids were prepared according to literature

protocols.20 NMR data was acquired on an Agilent 400 MHz. All GFP proteins were purified

according to manufacturer’s protocols using a Qiagen Ni-NTA Quik Spin Kit. Fluorescence data

was measured using a PerkinElmer LS 55 Luminescence Spectrometer.

General Biological Glaser-Hay Protocol

To 1.5 mL Eppendorf tube was added 3 µL of a 500 mM CuI solution in water and 3 µL 500 mM

TMEDA solution in water. This mixture was then incubated at 4oC for 10 mins. Following the

incubation, 1xPBS (22 µL) was added, followed by the alkyne containing GFPTAG66 (10 µL) in

PBS (~0.5mg/mL) and a 40 mM solution (4 µL) of the cognate alkyne in DMSO. The mixture was

allowed to react at 4°C for 4 hr.

Protocol for Fluorimetry Scans

After reacting for 4 hr, 10 µL of the reaction mixture was added to a quartz cuvette and diluted up

to 3 mL with PBS. This was then excited at either 395 nm (for the pPrF-containing chromophore)

or 280 nm (for the pEtF-containing chromophore) with a 10 nm slit width for the excitation and

23

emission wavelengths. Slit widths were increased or decreased as necessary depending on the

intensity of the reaction product’s signal. The scan speed was set at 500 nm/min.

General Protocol for Biological Glaser-Hay with Aromatic Reaction Partners

To 1.5 mL Eppendorf tube was added 3 µL of a 500 mM CuI solution in water and 3 µL 500 mM

TMEDA solution in water. This mixture was then incubated at 4 oC for 10 mins. Following the

incubation, 1xPBS (22 µL) was added, followed by the alkyne containing GFPTAG66 (10 µL) in

PBS (~0.5mg/mL) and a 40 mM solution (4 µL) of the cognate alkyne in DMSO. The mixture was

allowed to react at 4°C for 4 hr. Following this, the unreacted aromatic alkyne was washed away

using a 10 MWCO spin column (Corning) and rinsing with 50 µL portions of PBS 8 times The

solution was then concentrated to ~25 µL, as indicated on the spin column. Of this cleaned

solution, 10 µL was placed into a quartz cuvette and diluted to 2 mL with PBS for fluorescence

analysis.

References

1. Wu A. & Senter P. (2005). Arming antibodies: prospects and challenges for

immunoconjugates. Nat Biotechnol., 23, 1137–1146.

2. Brustad E.M. & Arnold F.H. (2011). Optimizing non-natural protein function with directed

evolution. Curr Opin Chem Biol., 15, 201–210.

3. Zhu H. & Snyder M. (2003). Protein chip technology. Curr. Opin. Chem. Biol., 7, 55–63.

4. Tan W., Sabet L., Li Y., et al. (2008). Optical protein sensor for detecting cancer markers in

saliva. Biosens. Bioelectron., 24, 266–271.

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5. Link A.J., Mock M.L., & Tirrell D.A. (2003). Non-canonical amino acids in protein

engineering. Curr. Opin. Biotechnol., 14, 603–609.

6. Banghart M., Volgraf M., & Trauner D. (2006). Engineering light-gated ion channels.

Biochemistry, 45, 15129–15141.

7. Steen Redeker, E., Ta D.T., Cortens D., Billen B., Guedens W., & Adriaensens, P. (2013).

Protein engineering for directed immobilization. Bioconjug. Chem., 24, 1761– 1777.

8. Presta L. (2006). Engineering of therapeutic antibodies to minimize immunogenicity and

optimize function. Adv. Drug. Deliv. Rev., 58, 640–656.

9. Young T.S. & Schultz P.G. (2010). Beyond the canonical 20 amino acids: expanding the


10. Liu C. & Schultz P.G. (2010). Adding new chemistries to the genetic code. Annu. Rev.

Biochem., 79, 413–444.

11. Sletten E.M. & Bertozzi C.R. (2009). Bioorthogonal chemistry: fishing for selectivity in a sea

of functionality. Angew. Chem. Int. Ed., 48, 6974–6998.

12. Maza J.C., Jacobs T.H., Uthappa D.M., Young D.D. (2016). Employing unnatural amino acids

in the preparation of bioconjugates. Synlett. 27, 805-813.

13. Rostovtsev V.V., Green L.G., Fokin V.V., & Sharpless K.B. (2002). A stepwise huisgen

cycloaddition process: copper(I)-catalyzed regioselective ‘‘ligation” of azides and terminal

alkynes. Angew. Chem. Int. Ed. Engl., 41, 2596–2599.

14. Wang Q., Chan T.R., Hilgraf R., Fokin V.V., Sharpless K.B., & Finn M.G. (2003).

Bioconjugation by copper(I)-catalyzed azide-alkyne [3+2] cycloaddition. J. Am. Chem. Soc.,

125, 3192–3193.

25

15. Agard N.J., Prescher J.A., & Bertozzi, C.R. (2004). A strain-promoted [3+2] azide-alkyne

cycloaddition for covalent modification of biomolecules in living systems. J. Am. Chem. Soc.,

126, 15046–15047.

16. Baskin J.M., Prescher J.A., Laughlin S.T., et al. Copper-free click chemistry for dynamic in

vivo imaging. Proc. Natl. Acad. Sci. USA, 104, 16793–16797.

17. Glaser C. (1896). Ber. Dtsch. Chem. Ges., 2, 422–424.

18. Shi W. & Lei A. (2014). 1,3-Diyne chemistry: synthesis and derivations. Tet. Lett., 55, 2763–

2772.

19. Maza J.C., McKenna J.R., Raliski B.K., Freedman M.T., & Young D.D. (2015). Synthesis

and incorporation of unnatural amino acids to probe and optimize protein bioconjugations.

Bioconj. Chem., 26, 1884-1889.

20. Maza J.C., Nimmo Z.M., & Young D.D. (2016). Expanding the scope of alkyne-mediated

bioconjugations utilizing unnatural amino acids. Chem. Commun., 52, 88–91.

21. Lampkowski J.S., Villa J.K., Young T.S., & Young D.D. (2015). Development and

optimization of Glaser-Hay bioconjugations. Angew. Chem. Int. Ed., 54, 9343-9346.

22. Pakhomov A. & Martynov V. (2008). GFP family: structural insights into spectral tuning.

Chem. Biol., 15, 755–764.

23. Ormö M., Cubitt A.B., Kallio K., Gross L.A., Tsien R.Y., & Remington S.J. Crystal structure

of the Aequorea victoria green fluorescent protein. Science, 273, 1392–1395.

24. Craggs T. (2009). Green fluorescent protein: structure, folding and chromophore maturation.

Chem. Soc. Rev., 38, 2865–2875.

25. Young D.D., Jockush S., Turro N., Schultz P.G. (2011). Synthetase polyspecificity as a tool

to modulate protein function. Bioorg. Med. Chem. Lett., 21, 7502–7504.

26

26. Maza J.C., Villa J.K., Landino L.M., Young D.D. (2016). Utilizing unnatural amino acids to

illustrate protein structure-function relationships: an experiment designed for an

undergraduate biochemistry laboratory. J. Chem. Edu., 93, 767–771.

27. Deiters A. & Schultz P.G. (2005). In vivo incorporation of an alkyne into proteins in

Escherichia coli. Bioorg. Med. Chem. Lett., 15, 1521–1524.

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aminoacyl-tRNA synthetase with atypical polysubstrate specificity. Biochemistry, 50, 1894–

1900.

27

CHAPTER 3: MECHANISTIC INVESTIGATION INTO THE AQUEOUS GLASER-

HAY BIOCONJUGATION

Introduction

As previously discussed, protein bioconjugates, in which a protein is conjugated to another

molecule, represent a critical area of research with widespread applications, most notably in the

development of strategies for site-specific drug delivery and improved cellular imaging

strategies.1-5 Unnatural amino acid (UAA) technology introduces novel chemical moieties into

proteins and allows for the preparation of well-defined, homogenous protein bioconjugates, which

have been demonstrated to have therapeutic advantages over the heterogenous bioconjugate

mixtures afforded through reactivity of native amino acid residues.6-10

The Glaser-Hay coupling

represents an attractive reaction

to develop as a bioconjugation, given its many chemical advantages, among them the formation of

a highly stable carbon-carbon bond in a well-defined linear 1,3 diyne as well as the tolerance of

the reaction to a wide range of functional groups.11 (Figure 3.1). Our previous work reported the

first successful biological Glaser-Hay coupling in a full-length protein under mild, aqueous

reaction conditions.12 This bioconjugation relies on the incorporation of an UAA containing a

terminal alkyne: p-propargyloxyphenylalanine (pPrF, 1) (Figure 3.2). The diyne product of the

Glaser-Hay reaction has many useful downstream applications, including its use as the starting

point for many cycloaddition reactions yielding multivalent carbo- and heterocycle products with

diverse biological, photochemical, and optoelectronic properties.13-17

+

Figure 3.1. The Glaser-Hay reaction generates a stable linear diyne and

is tolerant of many functional groups.

28

While the biological Glaser-Hay coupling is effective, significant protein oxidation has

been observed after around 6 hours of reaction time.12 Our previous work reported two optimized

conditions for the Glaser-Hay bioconjugation, the use of the traditional TMEDA ligand in a pH

6.0 reaction to afford faster coupling as well as the use of a carboxylated biphenyl ligand in a pH

8.0 reaction to minimize protein degradation.18 Both optimized reactions were performed at room

temperature. Despite the utility of the Glaser-Hay bioconjugation and this previous work, we

hypothesized there was a potential for further optimization, which could be facilitated via an

enhanced understanding of the reaction mechanism in aqueous solution.

Investigating the Aqueous Mechanism of the Glaser-Hay coupling

The mechanism of the Glaser-Hay coupling in organic solution has been studied in detail.

In 1964, Bohlmann et al. reported the formation of a dicopper(II)-diacetylide complex as the rate-

limiting step in the observed second-order kinetics.19 Later work suggested that the mechanism

progresses through a dicopper(III) intermediate (Figure 3.3).20

Most recently, Vilhelmsen et al. reported the reaction as zero-order in the alkyne starting

material.21 This study utilized UV/Vis spectroscopy as well as 13C NMR to study reaction

29

progression under various conditions. They reported that increased copper(I) and/or increased

TMEDA resulted in an increased reaction rate. However, they also notably reported that the

reaction changed to a slower zero-order kinetics reaction after some time, due to the buildup of

water in the reaction through its absorption from the air. Given the detrimental effects of water on

the rate of the organic Glaser-Hay coupling, we sought to investigate the kinetics of the aqueous

Glaser-Hay coupling, as the reaction pathway may proceed by a different reaction pathway in

water than previously observed organic solvents.

For this study, we employed UV/Vis

spectroscopy as well as both 13C and 1H NMR to monitor

the Glaser-Hay coupling of the dimerization of propargyl

alcohol in an aqueous solution (Figure 3.4). For NMR experiments, deuterated water (D2O) was

employed as the solvent in place of water to facilitate an NMR lock and for clarity of NMR spectra.

Notably, reaction progress was tracked via relative integration of 13C product and reactant peaks.

Although such a method for quantitative kinetics studies is abnormal, it has been proven effective

in quantitative analyses, specifically in kinetic studies of in vivo and in vitro processes.22-27 Further,

Vilhlemsen et al. successfully utilized integration of 13C NMR to track the kinetics Glaser-Hay

Figure 3.3. (A) Glaser-Hay mechanism progressing through a dicopper(II) diacetylide complex, as proposed by

Bohlmann.19

(B) Computationally determined Glaser-Hay mechanism progressing through a dicopper (III)

intermediate, as proposed by Fomina.20

B A

Figure 3.4. Dimerization of propargyl

alcohol via the Glaser-Hay coupling.

30

coupling in organic solution.21 While poor signal to noise ratio and the effects of the nuclear

Overhauser effect are cited as pitfalls of integrating 13C NMR spectra, the strength of the 400 MHz

NMR instrument used coupled with the deuterated solvent used appears to be sufficient to

overcome these limitations.21

The reaction shown in Figure 3.4 was prepared in a vial and air was bubbled through the

solution for 10 minutes. The vial was then sealed and allowed to stir at 80°C. At specific

timepoints, a portion of the reaction was removed and added directly to a flame-dried NMR tube.

Following NMR acquisition, this volume was added back to the reaction mixture. The relatively

low equivalencies of copper(I) iodide (2.4 mol%) and TMEDA (4.0 mol%) were chosen as the

31

baseline as a reflection of the low concentrations of catalyst and ligand when the reaction is

conducted on a biological system. Given these starting conditions, our results are applicable to the

previously developed Glaser-Hay bioconjugation.

In order to probe the reaction kinetics of the aqueous Glaser-Hay coupling, we varied the

concentrations of reaction components, including propargyl alcohol, copper(I) iodide, and

TMEDA. Each trial was then compared as a function of time for the rate of formation of a product

peak observed at 50 ppm in the 13C NMR relative to the corresponding starting material peak at

approximately 49.3 ppm for each set of reaction conditions (Figure 3.5). The 50 ppm peak

corresponds to the methylene peak in the diyne-containing product. The peak seen at

approximately 49.3 ppm corresponds to the methylene peak in the terminal alkyne-containing

starting material.

Under all reaction conditions tested, we observed the instantaneous disappearance of the

terminal alkyne peak in the 1H NMR along with the immediate formation of a triplet peak in the

13C NMR. This indicates the instantaneous formation of a copper acetylide intermediate during the

aqueous Glaser-Hay coupling.

Through varying the amount of propargyl alcohol in the reaction, we observed the reaction

did not increase in rate when levels of starting material were higher. Thus, relative integration

displays that a higher proportion of starting material is converted when smaller amounts of

propargyl alcohol are used. (Figure 3.6). In other terms, the reaction does not appear to progress

at a faster rate when more propargyl alcohol is added.

32

Varying the amount of copper(I) iodide in the reaction yielded surprising results. We

observed that increases in the level of copper(I) in the reaction resulted in decreased reaction rates

and greater conversion of starting material to product, even at ten times (24 mol%) the original

copper (I) iodide amount (Figure 3.7). This finding is significant, as it suggests that the aqueous

Glaser-Hay coupling may not progress through a dicopper acetylide intermediate, as had been

reported as the mechanism of the reaction in organic solution. This is logical given the high

coordination of water as the solvent, as it will act as a ligand to bind copper, making it less likely

to form a dicopper complex. Instead, these results suggests that copper coordinates to just a single

alkyne at a time to generate a copper(I) acetylide, which has been reported to be fairly stable.28 We

hypothesize that the Glaser-Hay coupling occurs between a copper acetylide and a free alkyne.

Thus, increased copper in the aqueous reaction leads to much of the alkyne being tied up in copper

acetylide, leaving little free terminal alkyne starting material to react with the copper acetylide to

33

form the dimer product. Further, the addition of more copper may lead to the formation of larger

copper acetylide clusters, which may limit reactivity of the catalyst in an aqueous solution.29

Varying the amount of TMEDA in the reaction demonstrated kinetics similar to those

found in organic solution. Increases in the amount of TMEDA led to increased reaction rates,

suggesting increases in nitrogenous ligand concentration help facilitate quicker reaction and more

efficient coupling (Figure 3.8). Results indicate that the reaction is approximately first order in

TMEDA.

UV/Vis spectroscopy indicated a shift in absorbance immediately after the addition of

propargyl alcohol to the catalyst mixture in aqueous solution. When just copper(I) iodide and

TMEDA were in an aqueous solution, the solution was blue and had a UV/Vis absorbance peak at

a wavelength just above 600 nm (Figure 3.9). When the propargyl alcohol was added to the

34

reaction, the color nearly immediately changed to a green-yellow, and the UV/Vis absorbance peak

shifted to a wavelength of just above 400 nm. Taking specific timepoints throughout the reaction

showed that the maximum wavelength of absorption did not change over the course of the reaction.

35

Optimization of the Biological Glaser-Hay coupling

With our newly improved understanding of the kinetics of the aqueous Glaser-Hay

coupling, we sought to further optimize the aqueous Glaser-Hay bioconjugation to reduce protein

oxidation and improve coupling efficiency. We previously reported that the addition of the radical

scavengers ascorbic acid, cysteine, and oleic acid did not afford better coupling additions and that

ascorbic acid actually increased protein oxidation and inhibited coupling.18

Taking these results along with aforementioned observations of the instantaneous

formation of the copper acetylide and the lack of formation of a dicopper intermediate, we

hypothesized that the oxidative damage to protein in the aqueous Glaser-Hay bioconjugation could

be due to damage from hydrogen peroxide (H2O2) generation, rather than free radicals.

This hypothesis is also supported by our previous report that ascorbic acid causes increased

protein oxidation and inhibits coupling. It has been reported that ascorbic acid can be cytotoxic

due to its production of hydrogen peroxide, which is a reactive oxidative species (ROS) that

damages proteins and other cellular components.30,31 Further, it has been demonstrated that the

presence of the enzyme catalase in cells can mitigate the cytotoxicity of ascorbic acid through the

breakdown of hydrogen peroxide. Given this, we sought to investigate whether catalase would

decrease protein oxidation and improve coupling efficiency in the Glaser-Hay bioconjugation.

Finally, it has also been reported that hydroxyl radicals, which directly damage proteins,

are generated through the interaction of Cu2+ ions and hydrogen peroxide in phosphate-buffered

solution, which is the solution in which the aqueous Glaser-Hay bioconjugation is conducted.29

Additionally, it has been suggested that the level of radical formation is at least partially dependent

on the ligand chelating Cu2+ in solution. Given this as well as our previous report of a carboxylated

biphenyl ligand affording better coupling than the originally reported TMEDA, we sought to

36

investigate whether other ligands could be employed to minimize protein degradation in the

Glaser-Hay bioconjugation.18

In order to probe the effects of catalase and various nitrogenous ligands, we performed a

250 mL expression of GFP containing pPrF in position 151. Following purification, the protein

was buffer exchanged into PBS and concentrated to a standard concentration of 1.0 mg/mL to

remove variability in protein expression. With the GFP/pPrF in hand, we prepared a series of

reactions to test the effectiveness of catalase and different nitrogenous ligands in a coupling

reaction between the mutant protein and an Fluor-488 alkyne dye.

For the ligand, the traditionally

used 2 as well as the recently reported 3

were selected for investigation, along

with biquinoline (4), a dimethylated

bipyridyl (5) and terpyridine (6).

(Figure 3.10). In the interest of solubility,

for 2, a 500 mM solution was prepared in

H2O. For 3, the solution was prepared in

1 M NaOH. For 4, 5, and 6, the solutions were prepared in DMSO. When conducting the reaction

with 2, GFP/pPrF of pH = 6.0 was employed for 4 hours, based on the previously optimized

conditions.18 Similarly, in a reaction with 3, GFP/pPrF of pH = 8.0 was employed for 8 hours, in

accordance with previously optimized conditions. For reaction with 4, 5, and 6, the same reaction

conditions as 3 were employed due to similarities in ligand structure.

A total of ten reactions were prepared, as the effectiveness of each ligand was tested with

and without the addition of catalase. Gratifyingly, the addition of catalase improved coupling and

Figure 3.10. Nitrogenous ligands selected for investigation.

37

reduced protein oxidation for each of the ligands examined (Figure 3.11). Reactions with ligands

2 and 4 demonstrated little protein oxidation. However, reaction with ligand 5 demonstrated

significantly more protein oxidation, especially in the absence of catalase. These results indicate a

likelihood that hydrogen peroxide is a byproduct of the Glaser-Hay coupling in aqueous solution.

This peroxide generation is likely the source of protein oxidation during the aqueous

bioconjugation. Here, we demonstrate that the addition of catalase as well as appropriate ligand

selection can greatly reduce protein degradation and improve coupling in the Glaser-Hay

bioconjugation.

Streamlining the Glaser-Hay Bioconjugation

Bioconjugation reactions such as the Glaser-Hay on proteins containing UAAs can be time-

consuming and involve multiple purification steps. We sought to streamline the process through

attempting the Glaser-Hay bioconjugation reaction on the lysate from purification as well as during

the protein purification process (Figure 3.12). Each of these two novel reaction pathways was

tested on ligands 2, 3, and 4 in Glaser-Hay bioconjugations with Fluor-488 alkyne dye. Preparation

of ligand solutions and duration of reaction time were the same as the aforementioned conditions.

38

To prepare to test the Glaser-Hay bioconjugation during the protein purification process,

the protein was purified, with purification facilitated by the hexa-histidine tag present at the end

of the protein. Purification was performed using a Qiagen Ni-NTA Quik Spin Kit according to

manufacturer’s protocol up until the elution step. At this point, the GFP/pPrF was bound to the Ni-

NTA resin, and reactants were added in efforts to perform the Glaser-Hay bioconjugation with the

protein bound to the purification matrix.

To test the Glaser-Hay bioconjugation on cell lysate, protein expressions were spun down

and lysed using commercially available BugBuster for 20 minutes. After centrifugation, the lysate

was used directly in the Glaser-Hay bioconjugation.

39

Both reaction preparation strategies were successful, as indicated by the presence of

fluorescence bands on the SDS-PAGE (Figure 3.13). Binding the mutant protein to the nickel

purification resin prior to conducting the Glaser-Hay bioconjugation afforded better coupling than

conducting the reaction on the lysate. Nevertheless, each of these approaches allows for increased

applicability of the Glaser-Hay bioconjugation through streamlining the reaction process and

rendering no need for additional purification steps.

Conclusion

Overall, our results suggest that the aqueous Glaser-Hay reaction does not progress through

a dicopper intermediate, which is mechanistically different from the organic Glaser-Hay reaction.

Furthermore, we successfully demonstrated that the addition of catalase to the Glaser-Hay

bioconjugation improves coupling and reduces protein degradation, suggesting that hydrogen

peroxide is formed as a byproduct of the aqueous Glaser-Hay reaction. Additionally, we further

investigated the impact of various nitrogenous ligands on the Glaser-Hay bioconjugation,

presenting biquinoline (4) as a promising ligand for future use. Finally, we demonstrated the

feasibility and applicability of a streamlined approach to conducting the Glaser-Hay

bioconjugation through carrying out the reaction on the cell lysate or during protein purification.

40


General. Reactions were conducted under ambient atmosphere with non-distilled solvents. NMR

data was acquired on a Varian Gemini 400 MHz. All GFP proteins were purified according to

manufacturer’s protocols using a Qiagen Ni-NTA Quik Spin Kit.

Dimerization of Propargyl Alcohol

The following was used as the standard to monitor the aqueous Glaser-Hay coupling. Amounts of

Cu(I), TMEDA, and propargyl alcohol were varied to analyze the effect of each on the reaction

kinetics. To a flame-dried vial, 8 mg of copper(I) iodide (2.4 mol %) was added to 2 mL D2O,

along with 10 µL of TMEDA (4.0 mol %). Next, 100 µL of propargyl alcohol (1.717 mmol, 1 eq)

was added, and air was bubbled through the reaction for 10 minutes. The reaction vial was sealed

and was stirred at 80°C for 12 hours. Timepoints were taken at 1 hour, 3 hours, 6 hours, 8 hours,

10 hours, and 12 hours. At each timepoint, a sample was removed from the vial and directly added

to an NMR tube for 1H and 13C NMR analysis.

Synthesis of p-dipropargylaminophenylalanine (pPrF)

Synthesis of p-propargyloxyphenylalanine (pPrF): Boc-Tyrosine-OMe (114 mg, 2 eq, 0.385

mmol) was added to a flame-dried vial. Cesium carbonate (254 mg, 3 eq, 0.578 mmol) was then

added, followed by dry DMF (3 mL). This mixture was stirred at 100°C for 20 mins. 5- Bromo-1-

pentyne (20 µL, 1 eq, 0.193 mmol) was then added to the mixture, as well as a catalytic potassium

iodide. The reaction was stirred overnight at 100°C, then cooled to room temperature and extracted

with brine (10 mL) and diethyl ether (10 mL). The ether layer was then washed with brine (10 mL

x 3). The brine layer was then back-extracted with ether (10 mL). The organic layers were

41

combined, dried with magnesium sulfate, filtered, and excess solvent was removed in vacuo.

Column chromatography (silica gel, 5:1 hexanes/ethyl acetate) was performed to yield the

protected product. This was then dissolved in 1,4-dioxane (2 mL). Then, 1 M lithium hydroxide

(2 mL) was added and the reaction was stirred at room temperature for 2 hours. 1,4-dioxane was

then removed in vacuo and the resulting water solution was acidified through the dropwise addition

of 6 M HCl. The reaction was then extracted into ethyl acetate and the organic layer dried with

magnesium sulfate and filtered. Excess solvent was removed in vacuo to yield a colorless oil. This

oil was dissolved in dichloromethane (DCM, 1.5 mL). Trifluoroacetic acid (TFA, 0.5 mL) was

added and the reaction was stirred at room temperature for 1 hour. Excess solvent was removed in

vacuo to yield pPrF as a white crystal (22 mg, 0.061 mmol, 31.6% yield). 1H NMR (400 MHz,

CDCl3): δ 7.02 (d, J = 12 Hz, 2 H), 6.82 (d, J = 12 Hz, 2 H), 4.95 (d, J = 8 Hz, 1 H), 4.53 (d, J = 8

Hz, 1 H), 4.03 (t, J = 4 Hz, 2 H), 3.71 (s, 3 H), 3.02 (m, J = 8 Hz, 1 H), 2.39 (t, J = 4 Hz, 2 H), 1.97

(m, J = 8 Hz, 2 H), 1.55 (s, 1 H), 1.41 (s, 9 H). 13C NMR (400 MHz, CDCl3): δ 172.4, 157.9, 130.3,

127.9, 114.5, 83.5, 79.9, 68.8, 66.0, 54.5, 52.2, 37.4, 28.3, 28.2, 21.1, 15.1.

Expression of pPrF-containing GFP-151

Escherichia coli BL21(DE3) cells were co-transformed with a pET-GFP-TAG-151 plasmid (2.0

µL) and a pEvol-pCNF plasmid (2.0 µL) using an Eppendorf electroporator. Cells were then plated

on LB-agar plates supplemented with ampicillin (50 mg/mL) and chloramphenicol (34 mg/mL)

and grown at 37°C. After 16 hours, a single colony was used to inoculate LB media (10 mL)

supplemented with ampicillin and chloramphenicol. The culture was grown to confluence at 37°C

over 16 hours. This culture was then used to begin an expression culture in LB media (250 mL) at

OD600 = 0.1, then incubated at 37°C until it reached an OD600 of between 0.7 and 0.8. At this point,

42

mutant protein expression was induced through the addition of 1 M ITPG (250 µL) and 20%

arabinose (250 µL), as well as 100 mM pPrF (2.5 mL). Induced cells were grown for an additional

16 hours at 30°C, then harvested via centrifugation (10 mins, 5000 rpm). The media was decanted,

and the cell pellet was stored in a -80°C freezer for 20 minutes. Mutant GFP was then purified

using commercially available Ni-NTA spin columns according to the manufacturer’s protocol.

Protein yield and purity was then assessed via SDS-PAGE and spectrophotometrically via a

Nanodrop spectrophotometer. Protein was then transferred into phosphate buffered saline solution

(PBS) using 10k MWCO spin columns prior to use in bioconjugation reactions.

Glaser-Hay Bioconjugation Protocol

To a sterile 1.5 mL eppendorf tube, the following were added: 5 µL of a vigorously shaken solution

of CuI (500 mM in H2O) and 5 µL of nitrogenous ligand (500 mM). The two solutions were

thoroughly mixed by pipetting. Next, 30 µL of GFP containing a terminal alkyne UAA (GFP/

pPrF; pH = 8.0, 1.04 ± 0.03 mg/mL) and 20 µL of Fluor-488 Alkyne (1 mM in DMSO) were added

to the tube. Finally, 5 µL of catalase (10 mg/mL in H2O) was added to the tube. For control

reactions, 5 µL of PBS at the appropriate pH was added in place of catalase. The reaction was

incubated at room temperature (22°C) for the appropriate reaction duration. Excess reactants were

then removed by buffer exchange using 10k MWCO concentrator columns. The reaction was

washed with PBS (8 × 200 µL) to a final volume of 50 µL. The reaction was analyzed by SDS-

PAGE and imaged immediately to analyze fluorescence. Fluorescence intensity indicated the

effective coupling reaction as the GFP is denatured and no longer fluorescent, while the coupling

to the fluorophore re-establishes a fluorescent signal. The gel was then stained for 3 hours using

Coomassie Brilliant Blue, then destained overnight using a methanol solution (60% deionized

43

H2O, 30% MeOH, 10% acetic acid). The gel was then analyzed again on the gel imager to indicate

protein presence and relative degradation.

Protocol for Glaser-Hay bioconjugation during protein purification

Expression of GFP/pPrF was spun down and cells were lysed using commercially available

BugBuster™. 250 µL of lysate was added to 100 µL of Ni-NTA resin and allowed to bind and was

washed according to manufacturer’s protocol. The resin was then washed with PBS (5 x 200 µL).

Then, 75 µL of PBS was added to wet the resin. Next, 10 µL of a premixed 1:1 solution of CuI

(500 mM in H2O) and nitrogenous ligand (500 mM) was added. Finally, 40 µL of Fluor-488

Alkyne (1 mM in DMSO) was added. The reaction was incubated at room temperature (22°C) for

the appropriate reaction duration. The nickel resin was then washed with PBS (8 x 200 µL) and

wash buffer (3 x 200 µL) and protein was eluted following manufacturer’s protocol. The reaction

was analyzed by SDS-PAGE and imaged immediately to analyze fluorescence. Fluorescence

intensity indicated the effective coupling reaction as the GFP is denatured and no longer

fluorescent, while the coupling to the fluorophore re-establishes a fluorescent signal. The gel was

then stained for 3 hours using Coomassie Brilliant Blue, then destained overnight using a methanol

solution (60% deionized H2O, 30% MeOH, 10% acetic acid). The gel was then analyzed again on

the gel imager to indicate protein presence and relative degradation.

Protocol for Glaser-Hay bioconjugation on cell lysate

Expression of GFP/pPrF was spun down and cells were lysed using commercially available

BugBuster. The lysed cells were centrifuged again and the lysate was decanted. 20 µL of a

premixed 1:1 solution of CuI (500 mM in H2O) and nitrogenous ligand (500 mM) was added to

44

250 µL of lysate. Next, 40 µL of Fluor-488 Alkyne (1 mM in DMSO) was added. The reaction

was incubated at room temperature (22°C) for the appropriate reaction duration. The lysate was

then bound to Ni-NTA resin and protein was purified according to manufacturer’s protocol. The

reaction was analyzed by SDS-PAGE and imaged immediately to analyze fluorescence.

Fluorescence intensity indicated the effective coupling reaction as the GFP is denatured and no

longer fluorescent, while the coupling to the fluorophore re-establishes a fluorescent signal. The

gel was then stained for 3 hours using Coomassie Brilliant Blue, then destained overnight using a

methanol solution (60% deionized H2O, 30% MeOH, 10% acetic acid). The gel was then analyzed

again on the gel imager to indicate protein presence and relative degradation.

References




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17. Pigulski, B., Mecik, P., Cichos, J., & Szafert, S. (2017). Use of stable amine-capped

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48

CHAPTER 4: SYNTHESIS OF A NOVEL DIPROPARGYL AMINE UAA AND

DEVELOPMENT OF A NOVEL CYCLOTRIMERIZATION BIOCONJUGATION

Introduction

A wide array of bioconjugation reactions are presently employed in the fields of medicine,

pharmaceuticals, and materials. 1-3 Protein bioconjugates are especially critical for drug delivery,

as targeted localization of drugs via the specificity of antibodies allows for reduced side effects in

chemotherapy patients, as lower doses can provide the same therapeutic effect as high quantities

of the unconjugated small molecule drug

alone.4-7

Unnatural amino acid (UAA)

technology represents a means to develop

bioorthogonal well-defined, homogenous

protein bioconjugate products, which have been

shown to have therapeutic advantages over

heterogenous bioconjugates.8-12 One common reaction employed to prepare a well-defined,

homogenous protein bioconjugate is the copper-catalyzed azide alkyne cycloaddition (CuAAC),

known as the “copper click” reaction, in which an UAA with an alkyne group is reacted with an

azide-containing reaction partner.9-11 However, there are a limited number of bioconjugation

reactions available for use with UAAs to generate protein bioconjugates (Figure 4.2).5 Thus, it is

49

critical to develop novel bioconjugation reactions to provide numerous chemical tools for different

applications and expand the scope of bioconjugation reactions.

The [2 + 2 + 2] cyclotrimerization reaction can be catalyzed by a variety of transition metals

and relies on three terminal alkyne groups reacting to generate polycyclic compounds, in a fashion

similar to a Diels-Alder reaction.13 This cycloaddition is widely used and is a key organic

methodology to generate a stable, polysubstituted benzene ring.14 To the best of our knowledge, a

biological [2 + 2 + 2] cyclotrimerization has never been reported, despite the prominence of this

reaction in organic synthesis. Recently, Wang et al. reported the success of a rhodium-catalyzed

cyclotrimerization in aqueous conditions in air at 60°C, demonstrating the potential success of this

reaction under

milder conditions

that may be feasible

under physiological

environments (Figure 4.3).15 We sought to implement principles from this reaction into a biological

system to afford a novel bioconjugation reaction, adding to the chemical biology toolbox to assist

in probing diseases and improving drug delivery systems.

Herein, we report a novel cyclotrimerization bioconjugation utilizing a newly synthesized

and site-specifically incorporated dipropargyl amine UAA. This work has key applications

towards the preparation of specific, homogenous protein bioconjugates with applications in

medicine, pharmaceuticals, and materials.

50

Development of Physiologically Compatible [2 + 2 + 2] Cyclotrimerization

Given the aqueous cycloaddition demonstrated by Wang and colleagues, we sought to test

whether this reaction could proceed successfully at temperatures lower than 60°C using the same

dicarboxylated biphenyl ligand that has previously been demonstrated

to provide effective chelation in the Glaser-Hay bioconjugation

(Figure 4.4).16 As a proof-of-concept experiment, we attempted to

dimerize propargyl ether under similar conditions as previously

reported (Figure 4.5). Gratifyingly, when performed at room temperature, product was able to be

detected by thin layer chromatography (TLC) and 1H NMR (29.8% yield).

Synthesis of UAA with Dipropargyl Functionality

In order for this reaction to be useful in an amino acid context, a dipropargyl functionality

is essential. Thus, we synthesized an UAA containing the dipropargyl amine functionality base on

a substituted phenylalanine backbone, 1, which we named pDPrAF. Para-amino-phenylalanine

was substituted with propargyl bromide in a similar fashion to the alkyne-containing UAA pPrF

synthesis (Figure 4.6). Though intending for addition of the alkyne-containing propargyl groups

to react solely at the amine, we also observed reaction at the unprotected carboxylic acid as well.

However, we realized that the standard deprotection protocol would convert the ester back to the

51

desired carboxylic acid while retaining the desired dipropargylamine functionality. Thus, we chose

not to initially protect the carboxylic acid group to reduce the overall number of steps in the

synthesis.

Our two-step synthesis afforded the desired product 1 with an overall yield of 47.1%. The

first step, the SN2-like addition of three alkyne-containing propargyl groups to the molecule,

resulted in the formation of the intermediate with good yield (47.9%). The intermediate was then

deprotected in a series of acid/base reactions to afford the desired product in excellent yield (98%).

Site-specific Incorporation of pDPrAF

Following the synthesis of 1, the next requisite is the genetic encoding of the new UAA.

While this typically requires the previously described double-sieve selection, previous research

reports that several pre-existing aaRS/tRNA pairs demonstrate a degree of polyspecificity towards

multiple UAAs. Consequently, we attempted to identify an appropriate aaRS capable of both

recognizing 1 and charging it onto the appropriate tRNA. We first investigated several synthetases

for testing due to either known polyspecificity or due to their incorporation of structurally similar

UAAs.17 Plasmids encoding both the aaRS and tRNA were co-transformed into BL21(DE3) E.

coli with a pET-GFP-TAG-151

plasmid, harboring GFP with a

TAG codon at position 151.

Following protein expression, GFP

mutants were purified using a Ni-

NTA resin and analyzed by SDS-PAGE to determine incorporation of 1. Gratifyingly, the

promiscuous pCNF aaRS was shown to effectively incorporate 1 into GFP at position 151, in

similar yields to its incorporation of pPrF and pBrPrF, two commonly employed alkyne-containing

52

UAAs (Figure 4.7). Thus, the previously evolved pCNF aaRS was utilized to express mutant GFP

harboring 1 at position 151.

Development and Optimization of Biological [2 + 2 + 2] Cyclotrimerization

Next, we moved to work to exploit the functionality of 1 in order to develop a biological

cyclotrimerization. As is the case in other bioconjugations, the amount of transition metal catalyst

added to the reaction should be minimized, as such metals can be cytotoxic and can negatively

impact biological systems. However, the toxicity of rhodium has been demonstrated to be lower

than that of other transition metals, including platinum, palladium, cadmium, nickel and

chromium.18 More specifically, in assays testing the impact of transition metals on oxidative

damage in epithelial cells, rhodium was demonstrated to be the least cytotoxic of the metals tested.

Further, rhodium complexes have also been integrated into proteins to generate a stable

organometallic protein containing rhodium.19 Thus, it is reasonable to attempt to employ rhodium

in bioconjugation reactions, provided low concentrations and relatively short reaction times.

To test the viability of this reaction, we sought to couple the pDPrAF mutant GFP with

Fluor 488 Alkyne (Figure 4.8). To do this, we mixed 5 µL of a rhodium (I) dimer complex,

[Rh(cod)Cl]2 (250 mM in DMSO), with 5 µL of carboxylated biphenyl ligand (500 mM in H2O).

Then, 30 µL of pDPrAF mutant GFP (pH = 7.4, ~1.0 g/mL) and 20 µL of Fluor-488 alkyne (1

mM in DMSO) were added. This aqueous reaction ran for 12 hours at 4°C. A lower temperature

than in the organic test of this aqueous coupling was employed in order to limit protein degradation

by rhodium. Gratifyingly, the reaction was successful, as indicated by the presence of a fluorescent

band on the SDS-PAGE gel at the appropriate molecular weight. Due to the denaturing gel, the

GFP protein should be denatured, and the fluorescent signal is the result of a direct conjugation

53

between the protein and the fluorophore. Unfortunately, in initial reactions, significant levels of

protein oxidation were also observed.

We hypothesized that the observed protein oxidation was due to the formation of radicals

at some point in the reaction process. To test this hypothesis, we employed catalase and sodium

ascorbate as radical scavengers to reduce protein oxidation and minimize protein degradation. As

previously described in Chapter 3, our lab has previously employed catalase to reduce protein

degradation in the aqueous Glaser-Hay bioconjugation.20 Further, sodium ascorbate has a long

history of use reducing protein oxidation in bioconjugation reactions, namely in the copper click

reaction and in biological Sonogashira couplings.10,21 Both catalase and sodium ascorbate resulted

in significantly reduced protein degradation, suggesting that damaging radicals are likely formed

at some point in the reaction. The addition of sodium ascorbate was demonstrated to be the most

effective in preventing protein degradation. Overall, we successfully performed a novel

bioconjugation reaction between GFP harboring 1 in position 151 and Fluor 488 alkyne through a

[2 + 2 + 2] cyclotrimerization. The resulting product was confirmed via SDS-PAGE as described,

and samples have been sent to mass spectral analysis for validation of product formation.

54

We then sought to further optimize the reaction. Initially, a time course was conducted,

investigating reaction times of 0 hours, 2 hours, 4 hours, 8 hours, 12 hours, and 24 hours with all

other conditions kept constant. It was elucidated that protein degradation increased fairly linearly

with time, and the maximum ratio of coupling to form the bioconjugate product occurred after a

reaction time of 2 hours (Figure 4.9).

Then, we sought to test the effect of temperature on the reaction, evaluating reaction

temperatures of 4°C, 22°C (room temperature), and 37°C. These reactions were conducted for 2

hours with all other conditions held constant. It was observed that the 22°C reaction afforded the

best coupling with no more protein degradation than was observed in the 4°C reaction. temperature

Reaction at 37°C led to significant protein degradation and reduced coupling (Figure 4.10).

55

Finally, we sought to determine the effect of different pH solutions on the reaction. In our

original proof of concept, GFP harboring 1 in a phosphate-buffer solution with pH 7.4 was

employed. Consequently, the cyclotrimerization reaction was conducted for 2 hours at 4°C on

reactions containing protein in solution of pH 6.0, pH 7.4, and pH 8.0. While pH did not appear to

have an effect on coupling, it did have minor impact on protein degradation, with the pH 7.4

reaction affording the least protein degradation.

Investigation of the Versatility of pDPrAF

Finally, versatility of the novel UAA 1 was assessed through examining its reactivity in

alkyne/azide 1,3-dipolar bioconjugations as well as Glaser-Hay bioconjugations. For the

56

alkyne/azide 1,3-dipolar

(copper click) bioconjugation,

the terminal alkyne groups of 1

within GFP were reacted with

an azide-containing

fluorophore to form the stable

triazole complex. For the

Glaser-Hay bioconjugation,

the terminal alkyne group in 1

in GFP was reacted with an

alkyne-containing fluorophore

in the presence of a

Cu(I)/TMEDA catalyst to

generate a linear, stable diyne functional group. For each of these reactions, we also hypothesized

that these reactions could occur on both terminal alkynes within the same protein, which would

afford a bioconjugate with two fluorophores attached to GFP at position 151. Gratifyingly, SDS-

PAGE demonstrated that 1 incorporated into proteins is capable of being employed in both copper

click and Glaser-Hay bioconjugations. Samples have been sent for mass spectrometry to confirm

these results and to determine whether attachment of two fluorophores is observed. Combined with

its use in our newly developed cyclotrimerization bioconjugation, there are at least three distinct

bioconjugation reactions in which pDPrAF can participate, making it a valuable amino acid

(Figure 4.12).

57

Conclusion

Overall, we present the synthesis and incorporation of a novel, dipropargyl amine UAA

with unique functionality capable of undergoing both the alkyne/azide 1,3-dipolar cycloaddition

and Glaser-Hay bioconjugations. Further, this novel UAA was utilized in a novel

cyclotrimerization bioconjugation. This bioconjugation affords a highly stable, polysubstituted

benzene ring as part of the conjugate, generating a highly stable covalent linkage between the two

reaction partners. This novel biological reaction has applications in medicine, pharmaceuticals,

and materials.


General

Reactions were conducted under ambient atmosphere with non-distilled solvents. NMR data was

acquired on a Varian Gemini 400 MHz. All GFP proteins were purified according to

manufacturer’s protocols using a Qiagen Ni-NTA Quik Spin Kit.

Synthesis of p-dipropargylaminophenylalanine (pDPrAF)

p-Aminophenylalanine-OMe (0.500 g, 1 eq, 1.784 mmol) was added to a flame-dried vial.

Potassium carbonate (1.232 g, 5 eq, 8.918 mmol) was added, followed by DMF (7 mL). This

mixture was stirred at room temperature for 5 minutes. Propargyl bromide (0.781 mL, 5 eq, 8.918

mmol) was then added and the reaction was stirred at 80°C for 96 hours. The reaction was then

cooled to room temperature and extracted with DCM and brine. The organic layers were combined,

dried with magnesium sulfate, filtered, and excess solvent was removed in vacuo. The reaction

was purified via flash chromatography (silica gel, 3:1 hexanes:ethyl acetate) to yield the desired

58

product as a yellow oil (0.337 g, 0.855 mmol, 47.9% yield). 1H NMR (400 MHz, CDCl3): δ 7.06

(d, J = 9 Hz, 2 H), 6.87 (d, J = 9 Hz, 2 H), 4.96 (d, J = 8 Hz, 1 H), 4.70 (q, J = 18 Hz, 2 H), 4.56

(d, J = 8 Hz, 1 H), 4.08 (s, 4 H), 3.02 (t, J = 8 Hz, 2 H), 2.51 (s, 1 H), 2.24 (s, 2 H), 1.41 (s, 9 H).

13C NMR (400 MHz, CDCl3): δ 171.4, 155.3, 147.0, 130.3, 126.8, 115.9, 80.1, 79.4, 75.6, 72.9,

54.5, 52.7, 40.6, 37.2, 28.5. M/Z = 395.3.

59

This product was then dissolved in 1,4-dioxane (2 mL). Then, 1 M lithium hydroxide (2 mL) was

added and the reaction was stirred at room temperature for 2 hours. 1,4-dioxane was then removed

in vacuo and the resulting water solution was acidified through the addition of 6 M HCl. The

reaction was then extracted into ethyl acetate and the organic layer dried with magnesium sulfate

and filtered. Excess solvent was removed in vacuo to yield a light brown oil. This oil was dissolved

in DCM (1.5 mL). Trifluoroacetic acid (TFA, 0.5 mL) was added and the reaction was stirred at

room temperature for 1 hour. Excess solvent was removed in vacuo to yield pDPrAF as a brown

solid (0.215 g, 0.840 mmol, 98% yield). 1H NMR (400 MHz, MeOD): δ 7.18 (d, J = 9 Hz, 2 H),

6.98 (d, J = 9 Hz, 2 H), 4.13 (s, 4 H), 3.30 (s, 1 H), 3.24 (dd, J = 9 Hz, 1 H), 3.07 (dd, J = 9 Hz, 1

H), 2.58 (s, 2 H). 13C NMR (400 MHz, MeOD): δ 147.6, 129.9, 124.6, 116.0, 78.9, 72.7, 39.8,

35.3. M/Z = 257.1. Overall yield 47.1%.

60

Expression of pDPrAF-containing GFP-151

Escherichia coli BL21(DE3) cells were co-transformed with a pET-GFP-TAG-151 plasmid (2.0

µL) and a pEvol-pCNF plasmid (2.0 µL) using an Eppendorf electroporator. Cells were then plated

on LB-agar plates supplemented with ampicillin (50 mg/mL) and chloramphenicol (34 mg/mL)

and grown at 37°C. After 16 hours, a single colony was used to inoculate LB media (10 mL)

supplemented with ampicillin and chloramphenicol. The culture was grown to confluence at 37°C

over 16 hours. This culture was then used to begin an expression culture in LB media (20 mL) at

OD600 = 0.1, then incubated at 37°C until it reached an OD600 of between 0.7 and 0.8. At this point,

mutant protein expression was induced through the addition of 1 M ITPG (20 µL) and 20%

arabinose (20 µL), as well as 100 mM pDPrAF (200 µL). Induced cells were grown for an

additional 16 hours at 30°C, then harvested via centrifugation (10 mins, 5000 rpm). The media

was decanted, and the cell pellet was stored in a -80°C freezer for 20 minutes. Mutant GFP was

then purified using commercially available Ni-NTA spin columns according to the manufacturer’s

protocol. Protein yield and purity was then assessed via SDS-PAGE and spectrophotometrically

61

via a Nanodrop spectrophotometer. Protein was then transferred into phosphate buffered saline

solution (PBS) using 10k MWCO spin columns prior to use in bioconjugation reactions.

Biological Cyclotrimerization protocol

To a sterile 1.5 mL Eppendorf tube, the following were added: 5 µL of [Rh(cod)Cl]2 (250 mM in

DMSO) and 5 µL of 2,2’-Bipyridine-4,4’-dicarboxylic acid (500 mM in DI H2O). The two

solutions were mixed thoroughly by pipetting until a dark red color was achieved. Next, 30 µL of

GFP containing pDPrAF at position 151 (GFP/pDPrAF; pH=7.4; ~1.0 mg/mL) and 20 µL of Fluor-

488 Alkyne (1 mM in DMSO) were added to the tube. Finally, 5 µL of sodium L-ascorbate (200

mM in DI H2O) was added to the tube. The reaction was incubated at 4°C. After 2 hours, excess

reactants were removed via buffer exchange using 10k MWCO spin columns. The reaction was

washed with phosphate buffered saline solution (pH 7.4 PBS, 8 x 200 µL) to a final volume of 50

µL. The reaction was analyzed by SDS-PAGE and imaged using a SYPRO Ruby scan to analyze

fluorescence. The gel was stained for 3 hours using Coomassie Brilliant Blue, then destained

overnight using a methanol solution (60% deionized water, 30% methanol, 10% glacial acetic

acid). The gel was then imaged using a Coomassie scan protocol.

Biological Glaser-Hay protocol

To a sterile 1.5 mL Eppendorf tube, the following were added: 5 µL of a vigorously shaken CuI

solution (500 mM in DI H2O) and 5 µL of tetramethylethylenediamine (500 mM in DI H2O). The

two solutions were thoroughly mixed by pipetting. Next, 30 µL of GFP containing pDPrAF at

position 151 (GFP151/pDPrAF; pH = 6; ~1.0 mg/mL) and 20 µL of Fluor-488 Alkyne (1 mM in

DMSO) were added to the tube. The reaction was incubated at room temperature (22°C). After 4

62

hours, excess reactants were removed via buffer exchange using 10k MWCO concentrator

columns. The reaction was washed with phosphate buffered saline solution (pH 6 PBS, 8 x 200

µL) to a final volume of 50 µL. The reaction was analyzed by SDS-PAGE and imaged using a

SYPRO Ruby scan to analyze fluorescence. The gel was stained for 3 hours using Coomassie

Brilliant Blue, then destained overnight using a methanol solution (60% deionized water, 30%

methanol, 10% glacial acetic acid). The gel was then imaged using a Coomassie scan protocol.

Biological copper click protocol

To a sterile 1.5 mL Eppendorf tube, the following were added: 2 µL of CuSO4 solution (50 mM in

DI H2O) and 2 µL of TCEP (50 mM in DI H2O). The two solutions were thoroughly mixed by

pipetting. Next, 20 µL of GFP containing pDPrAF at position 151 (GFP151/pDPrAF; pH = 7.4;

~1.0 mg/mL) and 10 µL of Fluor-488 Azide (1 mM in DMSO) were added to the tube. Finally, 10

µL of TBTA (5 mM in DMSO) was added to the tube. The reaction was incubated at 4°C. After

16 hours, excess reactants were removed via buffer exchange using MWCO concentrator columns.

The reaction was washed with phosphate buffered saline solution (pH 7.4 PBS, 8 x 200 µL) to a

final volume of 50 µL. The reaction was analyzed by SDS-PAGE and imaged using a SYPRO

Ruby scan to analyze fluorescence. The gel was stained for 3 hours using Coomassie Brilliant

Blue, then destained overnight using a methanol solution (60% deionized water, 30% methanol,

10% glacial acetic acid). The gel was then imaged using a Coomassie scan protocol.

References

1. Kalia, J., & Raines, R.T. (2010). Advances in bioconjugation. Curr. Org. Chem., 14, 138-

147.


63



4. Sievers, E.L. & Senter, P.D. (2013). Antibody-drug conjugates in cancer therapy. Annu.

Rev. Med., 64, 15-29.

5. Agarwal, P. & Bertozzi, C.R. (2015). Site-specific antibody−drug conjugates: the nexus

of bioorthogonal chemistry, protein engineering, and drug development. Bioconj. Chem.,

26, 176-192.

6. Jaiswal, J.K., Mattoussi, H., Mauro, J.M., & Simon, S.M. (2003). Long-term multiple

color imaging of live cells using quantum dot bioconjugates. Nat. Biotechnol., 21, 47.

7. Gao, X., Cui, Y., Levenson, R.M., Chung, L.W., & Nie, S. (2004). In vivo cancer

targeting and imaging with semiconductor quantum dots. Nat. Biotechnol., 22, 969.

8. Stephanopoulos, N. & Francis, M.B. (2011). Choosing an effective protein

bioconjugation strategy. Nat. Chem. Biol., 7, 876.



10. Young, D.D. & Schultz, P.G. (2018). Playing with the molecules of life. ACS Chem.

Biol., 13, 854-870.

11. Young, T.S. & Schultz, P.G. (2010). Beyond the canonical 20 amino acids: expanding the

genetic lexicon. J. Biol. Chem., 285, 11039-11044.

12. Hallam, T.J., Wold, E., Wahl, A., & Smider, V.V. (2015). Antibody conjugates with

unnatural amino acids. Mol. Pharm., 12, 1848-1862.

13. Kotha, S., Brahmachary, E., & Lahiri, K. (2005). Transition metal catalyzed [2+ 2+ 2]

cycloaddition and application in organic synthesis. Euro. JOC, 22, 4741-4767.

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14. Yamamoto, Y. (2005). Recent advances in intramolecular alkyne cyclotrimerization and

its applications. Curr. Org. Chem., 9, 503.

15. Wang, Y.H., Huang, S.H., Lin, T.C., & Tsai, F.Y. (2010). Rhodium (I)/cationic 2, 2′-

bipyridyl-catalyzed [2+ 2+ 2] cycloaddition of α, ω-diynes with alkynes in water under

air. Tetrahedron, 66, 7136-7141.

16. Nimmo, Z.M., Halonski, J.F., Chatkewitz, L.E., & Young, D.D. (2018). Development of

optimized conditions for Glaser-Hay bioconjugations. Bioorg. Chem., 76, 326-331.

17. Young, D.D., Young, T.S., Jahnz, M., Ahmad, I., Spraggon, G., Schultz, P.G. (2011).

Biochemistry, 50, 1894-1900.

18. Schmid, S., Zimmerman, S., Krug, H.F., & Sures, B. (2007). Influence of platinum,

palladium and rhodium as compared with cadmium, nickel and chromium on cell

viability and oxidative stress in human bronchial epithelial cells. Environ. Int., 33, 385-

390.

19. Satake, Y., Abe, S., Okazaki, S., Ban, N., Hikage, T., Ueno, T., Nakajima, H., Suzuki, A.,

Yamane, T., Nishiyama, H., & Watanabe, Y. (2007). Incorporation of a phebox rhodium

complex into apo-myoglobin affords a stable organometallic protein showing

unprecedented arrangement of the complex in the cavity. Organometallics, 26, 4904-

4908.

20. Travis, C.R., Mazur, L.E., Peairs, E.M., & Young, D.D. (2018). Mechanistic

investigation into the biological Glaser-Hay reaction. Org. Biomol. Chem. (in

preparation)

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21. Li, N., Lim, R., Edwardraja, S., and Lin, Q. (2011) Copper-Free Sonogashira Cross-

Coupling for Functionalization of Alkyne-Encoded Proteins in Aqueous Medium and in

Bacterial Cells. J. Am. Chem. Soc., 133, 15316−15319.

66

CHAPTER 5: TOWARDS THE DEVELOPMENT OF MULTIVALENT

BIOCONJUGATES

Introduction

As previously described, bioconjugates have widespread applications in various fields,

including therapeutics, diagnostics, and materials.1 However, each of the bioconjugation reactions

previously discussed, as well as nearly all the bioconjugation reactions utilized employ just two

reaction partners to form a divalent complex.2,3 With only two reactants, the bioconjugate product

is essentially limited to containing two distinct functionalities. For instance, an antibody-drug

conjugate possesses the localization functionality of the antibody and the therapeutic functionality

of the small molecule drug. Despite the great value of divalent bioconjugates across nearly all

fields, the preparation of a multivalent bioconjugate, in which three or more reaction partners

(where at least one is a biomolecule) are conjugated, could provide even more powerful and

applicable bioconjugates. For instance, a fluorescent probe could be conjugated with both an

antibody and a drug. This antibody-drug-probe conjugate would be capable of localization,

therapeutic efficacy, and visualization of the delivery and treatment all from the same molecule.

As previously described, unnatural amino acid (UAA) technology has proved especially

valuable in the preparation of protein bioconjugates, as the presence of UAAs with unique

chemical functionality in the protein can ensure well-defined, homogenous products.4 This offers

particularly important advantages in the use of antibody-drug conjugates to treat cancer, as

homogenous bioconjugates have been shown to be more effective in treatment than heterogenous

mixtures of bioconjugates, where the number and location of addition of drug molecules to the

antibody vary.4

67

Multivalent conjugates have been prepared via the incorporation of two distinct UAAs with

different reactivities into the same protein.5-7 Specifically, Wan et al. and Xiao et al. demonstrated

incorporation of two distinct UAAs into proteins via suppression of both the TAG (amber) stop

codon as well as the TAA (ochre) stop codon.5-6 This was accomplished via the introduction of

two separate orthogonal aminoacyl tRNA synthetase (aaRS)/tRNA pairs into cells. Xiao et al.

demonstrated successful preparation of a multivalent conjugate using this methodology through

the incorporation of a ketone-containing UAA (pAcF) and an azide containing UAA (AzK) into

an anti-HER2 IgG antibody. The distinct reactivities of these two groups was exploited to first

conjugate auristatin (nAF), a drug molecule, onto the ketone and then to conjugate a fluorophore

onto the azide via copper-free click chemistry

(Figure 5.1). The resulting trivalent

bioconjugate covalently linked an antibody, a

drug, and a fluorophore, offering three

distinct functionalities in a single

bioconjugate.

Neumann et al. took a different approach towards the preparation of multivalent protein

bioconjugates.7 This work demonstrated the potential to evolve a “quadruplet-decoding” ribosome

that successfully incorporates UAAs in response to quadruplet codons. Through this technology,

azide and alkyne UAAs were incorporated into specific sites into calmodulin, and an internal

copper click [copper(I)-catalyzed azide alkyne cycloaddition] was successfully executed to alter

protein function.

While these studies are incredible and have widespread value in the field of bioconjugate

chemistry, these processes are often tedious and complicated. Suppression of two stop codons with

68

systems of multiple aaRS/tRNA pairs can become complicated, and the evolution of a ribosome

capable of reading quadruplets is quite difficult.

Our lab has sought to perform two bioconjugation reactions on the same unnatural amino

acid, which would yield a multivalent protein bioconjugate without the need for the incorporation

of multiple UAAs into the same protein or the evolution of a quadruplet-decoding ribosome. Given

the stability of the triazole complex formed in the azide-alkyne “click” cycloaddition, past efforts

have focused on exploiting the potential reactivity of the linear 1,3-diyne generated in the Glaser-

Hay bioconjugation as well as the Cadiot-Chodkiewicz bioconjugation.8 Thus, a variety of

reactions aiming to add a third conjugation partner to a divalent bioconjugate containing a 1,3-

diyne have previously been explored (Figure 5.2).

Many organic reactions between a 1,3-diyne and substituted tetrazines, azides, amines,

thiols, silyls, and nitriles have been reported under conditions which would not be compatible in

biological systems, due to the use of organic solvents as well as high reaction temperatures.9-15

Thus, the aim of this research is to translate these reactions to physiological settings for the

preparation of protein multivalent conjugates in aqueous solution. To do this, reactions were first

tested under the reported organic conditions to confirm the feasibility of literature procedures, and

69

then investigated to find optimized conditions that would be compatible with biological

macromolecules.

Despite past efforts, all aforementioned approaches to react various functional groups with

a 1,3-diyne were unsuccessful in generating multivalent bioconjugates. Herein, I report on my

work in novel efforts to generate protein multivalent conjugates

Towards the Development of a Biological 1,3-Dipolar Cycloaddition of an Azide and a Diyne

First, the 1,3-dipolar cycloaddition and carbocyclization of an azide and a diyne to form a

napthotriazole was investigated (Figure 5.2, b). Mandadapu et al. previously reported successful

reaction of a 1,3-diyne and an azide in acetonitrile at 90°C in 2-4 days (Figure 5.3).14

Previous work

demonstrated the

feasibility of this

copper(I)-catalyzed

reaction in organic

solvents at 90°C to couple diphenyldiacetylene with benzyl azide to generate both the non-

carbocyclized and carbocyclized products (Figure 5.4). Despite the fact that those conditions were

not physiological, we hypothesized that the non-carbocyclized product could form at lower

temperatures in aqueous solvent. This would still form a trivalent protein complex, making it a

multivalent reaction. However, translation to biological conditions was unsuccessful.

70

Consequently, research

returned to organic conditions in

efforts to explore the feasibility of

different catalyst systems performing

the same cycloaddition but in aqueous solution. To do so, a Glaser-Hay reaction to generate a

homodimer containing a 1,3-diyne (1) was performed (Figure 5.5). Compound 1 was then reacted

with benzyl azide, with a Cu and FeCl3 catalyst system in water at room temperature for 48 hours

(Figure 5.6). After extraction, thin layer chromatography and 1H NMR suggested potential

formation of either carbocyclized or non-carbocyclized product via the appearance of a new spot

and the shifting of aromatic peaks in the 1H NMR relative to the benzyl bromide starting material,

respectively. Encouraged by the potential success of this reaction in aqueous solution, but

cognizant of the 48 hours needed for reaction, attempts were made to translate the reaction to a

physiological setting.

Two different biological reactions were examined. First, a 1,3-diyne heterodimer of

GFP151/pPrF and Biotin Alkyne was reacted with Fluor-488 Alkyne at 4°C for 12 hours using a

Cu and FeCl3 catalyst system. A successful reaction could be visualized by a fluorescence band on

the SDS-PAGE gel at the appropriate molecular weight. However, the reaction was unsuccessful

and no such product was achieved. Second, a 1,3-diyne heterodimer of GFP151/pPrF and Fluor-

488 Alkyne was reacted with Biotin Alkyne under the same conditions. Biotin has been reported

to have high affinity for streptavidin resin, which can be exploited to identify functional

71

multivalent conjugates.16 Thus, following reaction, the reaction as well as a control (in which no

catalyst was present) could be immobilized to a streptavidin resin and washed to remove non-

biotinylated protein. A successful reaction would be indicated by differences in fluorescence

between the reaction and the control. A differential was not seen as neither the reaction nor control

showed detectable fluorescence. This indicates that either the multivalent conjugation was

unsuccessful or afforded too low a yield for detection via immobilization on a streptavidin resin.

Towards the Development of a Terminal Alkyne Addition to a Diyne in Biological Settings

Efforts were then shifted to utilize a different reaction to prepare a trivalent bioconjugate.

Chen et al. reported the copper-palladium catalyzed addition of terminal alkynes to activated

alkynes in water at room temperature (Figure 5.7).17 We hypothesized that the presence of another

electron-rich

alkyne could

lead to the

diyne

possessing similar reactivity to the internal alkynes utilized by Chen. Thus, organic conditions

were tested to elucidate if a terminal alkyne could add to a 1,3-diyne under these conditions (Figure

5.8). Employing a copper-palladium catalyst system, 1 was reacted with phenylacetylene in water

at room temperature for 12 hours. Thin layer chromatography and 1H NMR indicated formation

of a novel product, through the appearance of a new spot and the shifting of aromatic 1H peaks

72

relative to the phenylacetylene starting material. Based on these preliminary results, attempts were

made to employ this reaction in a biological environment.

Using catalyst working concentrations of 500 mM CuI and 20 mM PdCl2(PPh3)2, a 1,3-

diyne heterodimer of GFP151/pPrF and Biotin Alkyne with Fluor-488 Alkyne were reacted at

room temperature for 4 hours. A successful reaction could be visualized by a fluorescence band

on the SDS-PAGE gel at the appropriate molecular weight. However, the reaction was

unsuccessful and no such product was achieved. Then, a 1,3-diyne heterodimer of GFP151/pPrF

and Fluor-488

Alkyne was reacted

with Biotin Alkyne

under the same

conditions (Figure

5.9). However, the presence of fluorophore was not observed on the streptavidin resin, indicating

the multivalent conjugation was unsuccessful. SDS-PAGE showed significant levels of protein

degradation during this reaction. To counter this, the same two reactions were performed at 4°C

for 4 hours. Protein degradation was again observed. The reaction was then attempted with a

working concentration of 100 mM CuI, which resulted in decreased protein degradation; however,

no products were observed.

Towards the Use of a Biological Sonogashira Reaction to Prepare a Multivalent Conjugate

With approaches to utilize the reactivity of the 1,3-diyne biologically proving difficult,

different reactive sites found within bioconjugates were investigated. The triazole complex

generated by the copper click reaction is highly stable and unreactive.2 However, previous research

demonstrated the formation of a bromotriazole complex through 1,3-dipolar azide-bromoalkyne

73

cycloaddition. Efforts were made to exploit the reactivity of this functional group through a

Sonogashira reaction to generate a trivalent bioconjugate (Figure 5.10). To do this, an array of

reactions between a bromotriazole-containing divalent bioconjugate and various groups containing

a terminal alkyne were performed under a variety of conditions (Table 5.1). Conditions for an

74

aqueous, physiologically compatible Sonogashira reaction were derived from literature

conditions.18

None of the conditions investigated clearly confirmed the creation of the desired trivalent

bioconjugate. However, SDS-PAGE of 12 indicated potential attachment of the fluorophore, but

in extremely low yields. This result is now being further investigated. Additionally, efforts are

currently underway to perform this Sonogashira reaction using a single palladium catalyst with no

copper, as is reported to be successful in physiological conditions.19

Conclusion

Overall, three distinct approaches towards the preparation of a multivalent bioconjugate

have been explored, but none have been successful to this point. The preparation of a multivalent

conjugate through reacting a terminal alkyne with a bromotriazole via a Sonogashira reaction

presently appears most promising, and investigation into this area will continue.


Synthesis of 1

Propargyl alcohol (2.05 g, 36.54 mmol, 2.11 mL) was added to THF (24 mL) in a flame dried

round-bottom flask. Copper iodide (160 mg) was dissolved in TMEDA (240 μL) and THF (8 mL).

This mixture was then combined with the propargyl alcohol and stirred for 24 hours while bubbling

air through at 60°C. The reaction was then purified via column chromatography in 1:1 Hexanes:

Ethyl Acetate. 1H NMR (400 MHz, D2O): δ 4.19 (s, 4H) ppm.

75

Organic 1,3-dipolar cycloaddition and carbocyclization of an azide and a diyne to form a

napthotriazole

To a flame-dried vial, 50 mg of 1 (0.455 mmol, 1 eq) wad added and dissolved in H2O. Next, 93

µL of benzyl azide was added, followed by a spatula tip each of granular copper (Cu) and iron(III)

chloride (FeCl3). The reaction was allowed to stir at room temperature for 48 hours. Formation of

products was evaluated by thin layer chromatography and crude 1H NMR spectroscopy.

Expression of Protein Containing Unnatural Amino Acid

A pET-GFP-TAG plasmid (0.5 μL, for GFP synthesis) was co-transformed with the polyspecific

pEVOL-pCNF aminoacyl tRNA synthetase plasmid (0.5 μL) into Escherichia coli BL21 (DE3)

competent cells using an Eppendorf Eporator electroporator. The cells were then plated (100 μL)

on LB agar supplemented with ampicillin (50 μg/mL) and chloramphenicol (34 μg/mL). The plates

were incubated 16 hours at 37°C. One colony was used to inoculate LB media (10 mL) containing

ampicillin (50 μg/mL) and chloramphenicol (34 μg/mL). The culture was shaken overnight at 37°C

and used to initiate an expression culture (250 mL media, ampicillin 50 μg/mL, chloramphenicol

34 μg/mL) at an OD600 = 0.1. The cultures were incubated at 37°C until OD600 = 0.6 was reached.

Protein expression was induced by addition of 20% arabinose (250 μL), 0.8 mM isopropyl-β-D-1-

thiogalactopyranoside (IPTG, 250 μL), and the unnatural amino acid (2.5 mL, 100 mM). Cultures

were incubated at 30°C overnight, then pelleted by centrifugation (5,000 rpm, 10 min). Pelleted

cells were stored at -80°C until purification. The cell pellet was resuspended with 500 μL of

Bugbuster (Novagen), and 200 μL of cell lysis buffer and incubated for 20 minutes at 37°C.

Cellular debris was pelleted out by centrifugation at 5,000 rpm for 10 minutes and the supernatant

was added to an equilibrated Ni-NTA resin (200 μL). GFP was purified according to

76

manufacturer’s protocol before being analyzed by SDS-PAGE (BioRad 10% precast gels, 150 V,

1.5 hours). Gels were stained using Coomassie Brilliant Blue, and destained (60% H2O, 30%

MeOH, 10% acetic acid). The gel was analyzed using the Coomassie protocol on the gel imager.

Protein was used without further purification.

General Procedure for Biological Glaser-Hay Coupling

To a sterile 1.5 mL eppendorf tube, the following were added: 5 μL of a vigorously shaken solution

of CuI (500 mM in H2O) and 5 μL of bidentate nitrogenous ligand (TMEDA, 500 mM in H2O).

The two solutions were thoroughly mixed by pipetting. Next, reaction partners 1 and 2 were added

to the mixture and mixed by pipetting. Reaction partners consisted of GFP/pPrF (30 μL, ~1 mg/mL

in PBS), AlexaFluor 488 Alkyne (20 μL, 1 mM in DMSO), or biotin alkyne (20 μL, 1 mM in

DMSO). The reaction was incubated at room temperature for 4 hours. The reaction was stopped

by removing excess reactants via buffer exchange using 10k MWCO concentrator columns, and

then subsequently analyzing the products by SDS-PAGE. This reaction afforded heterodimers

containing 1,3-diynes.

Biological 1,3-dipolar cycloaddition and carbocyclization of an azide and a diyne to form a

napthotriazole

To a sterile 1.5 mL eppendorf tube, the following were added: 5 μL of a vigorously shaken solution

of Cu (500 mM in H2O) and 5 μL of iron(III) chloride (FeCl3, 500 mM in H2O). The two solutions

were thoroughly mixed by pipetting. Next, 30 μL of the 1,3-diyne heterodimer of GFP151/pPrF

and Biotin Alkyne was added. Finally, 20 μL of the AlexaFluor 488 Alkyne (1 mM in DMSO)

was added. Reaction partners consisted of GFP/pPrF (30 μL, ~1 mg/mL in PBS), AlexaFluor 488

77

Alkyne (20 μL, 1 mM in DMSO), or biotin alkyne (20 μL, 1 mM in DMSO). The reaction was

incubated at 4°C for 12 hours. The reaction was stopped by removing excess reactants via buffer

exchange using 10k MWCO concentrator columns, and then subsequently analyzing the products

by SDS-PAGE. No multivalent conjugates were observed.

Organic Copper-Palladium catalyzed addition of a terminal alkyne to a 1,3-diyne

To a flame-dried vial, 60 mg of 1 (0.545 mmol, 1 eq) was added and dissolved in H2O. Next, 120

µL of phenylacetylene (1.091 mmol, 2 eq) was added, followed by 6 mg of CuI (5 mol%) and 2

mg of PdCl2(PPh3)2 (2.5 mol%). The reaction was allowed to stir at room temperature for 12

hours. Formation of products was evaluated by thin layer chromatography and crude 1H NMR.

Biological Copper-Palladium catalyzed addition of a terminal alkyne to a 1,3-diyne

To a sterile 1.5 μL Eppendorf tube, the following were added and mixed: 5 μL of palladium (II)

acetate (Pd(OAc)2, 20 mM in 1:1 H2O/DMSO) or bis(triphenylphosphine)palladium(II) dichloride

(PdCl2(PPh3)2, 20 mM in DMSO) and 5 μL of copper(I) iodide (CuI, 500 mM in H2O). Next, 30

μL of the 1,3-diyne heterodimer of GFP151/pPrF and Biotin Alkyne was added. Finally, 20 μL of

the AlexaFluor 488 Alkyne (1 mM in DMSO) was added. Reaction conditions varied as described

in the text. Following reaction, excess reactants were removed via buffer exchange using 10k

MWCO concentrator columns. The products were heated to 98°C for 10 minutes and analyzed by

SDS-PAGE. No multivalent product was detected.

78

General Procedure for Biological 1,3-Dipolar Azide-(Bromo)alkyne Cycloaddition

To a sterile 1.5 mL eppendorf tube, the following were added: 2 μL of CuSO42 (50 mM in H2O),

2 μL of TCEP (50 mM in H2O). Next, reaction partners 1 and 2 were added to the mixture and

mixed by pipetting. Reaction partners consisted of GFP151 containing either azide or bromoalkyne

UAA (20 μL in PBS), Ub48/pAzF (15 μL in PBS), or AlexaFluor 488 Azide (10 μL, 1 mM in

DMSO). Lastly, 10 μL of TBTA (5 μM in DMSO) was added, followed by 20 μL PBS. In reactions

involving linkers, 5 μL of 100 μM linker dissolved in DMSO was added. The reaction was

incubated at 4°C for 16 hours. The reaction was stopped either by performing SDS-PAGE

immediately or by removing excess reactants via buffer exchange using Spin-X UF concentrator

columns, 73 and then subsequently analyzing the purified products by SDS-PAGE.

General Procedure for Biological Sonogashira Coupling Reaction18

To a sterile 1.5 μL Eppendorf tube, the following were added and mixed: 3 μL of palladium (II)

acetate (Pd(OAc)2, 20 mM in 1:1 H2O/DMSO) and 3 μL of triphenylphosphine-3,3′,3′′-trisulfonic

acid trisodium salt (TPPTS, 100 mM in H2O). Then, 2 μL of copper (II) triflate (CuOTf, 12 mM

in DMSO) was added. Next, 10 μL of a bromotriazole-containing dimer of GFP151/pBrPrF and

AlexaFluor 488 Azide. Next, 10 μL of the third reaction partner, consisting of either AlexaFluor

488 Alkyne (1 mM in DMSO), biotin alkyne (1 mM in DMSO), or AlexaFluor 680 Alkyne (1 mM

in DMSO), was added. Reaction time and temperature varied as shown in Table 5.1. Following

reaction, excess reactants were removed via buffer exchange using 10k MWCO concentrator

columns. The products were heated to 98°C for 10 minutes and analyzed by SDS-PAGE. No

multivalent product was detected.

79

References






4. Agarwal, P. & Bertozzi, C.R. (2015). Site-specific antibody−drug conjugates: the nexus

of bioorthogonal chemistry, protein engineering, and drug development. Bioconj. Chem.,

26, 176-192.

5. Wan, W., et al. (2010). A facile system for genetic incorporation of two different

noncanonical amino acids into one protein in Escherichia coli. Angew. Chem. Int. Ed., 49,

3211-3214.

6. Xiao, H., et al. (2013). Genetic incorporation of multiple unnatural amino acids into

proteins in mammalian cells. Angew. Chem., 125, 14330-14333.

7. Neumann, H., et al. (2010). Encoding multiple unnatural amino acids via evolution of a

quadruplet-decoding ribosome. Nature, 464, 441.

8. Kolb, H.C. & Sharpless, K.B. (2003). The growing impact of click chemistry on drug

discovery. Drug Discov. Today, 8, 1128-1137.

9. Shi, W. & Lei, A. (2014). 1,3-Diyne chemistry: synthesis and derivations. Tetrahedron

Lett., 55, 2763-2772.

10. Yang, L. & Hua, R. (2013). Cycloaddition of 1,4-diaryl-1,3-butadiynes with nitriles: an

atom-economic approach to benzo[f]quinazolines. Chem. Lett., 42, 769-771.

80

11. Nizami, T.A. & Hua, R. (2014). Cycloaddition of 1,3-butadiynes: efficient synthesis of

carbo- and heterocycles. Molecules, 19, 13788-13802.

12. Fairbanks, B.D., Scott, T.F., Kloxin, C.J., Anseth, K.S., & Bowman, C.N. (2008). Thiol-

yne photopolymerizations: novel mechanism, kinetics, and step-growth formation of

highly cross-linked networks. Macromolecules, 42, 211-217.

13. Sun, H., Wu, X., & Hua, R. (2011). Copper (I)-catalyzed reaction of diaryl buta-1,3-diynes

with cyclic amines: an atom-economic approach to amino-substituted naphthalene

derivatives. Tetrahedron Lett., 52, 4408-4411.

14. Mandadapu, A.K., et al. (2011). Unprecedented Cu-catalyzed coupling of internal 1,3-

diynes with azides: one-pot tandem cyclizations involving 1,3-dipolar cycloaddition and

carbocylization furnishing napthotriazoles. Org. Lett., 13, 3162-3165.

15. Blackman, M.L., Royzen, M., & Fox, J.M. (2008). Tetrazine ligation: fast bioconjugation

based on inverse-electron-demand Diels-Alder reactivity. J. Am. Chem. Soc., 130, 13518-

13519.

16. Weber, P.C., Ohlendorf, D.H., Wendoloski, J.J., & Salemme, F.R. (1989). Structural

origins of high-affinity biotin binding to streptavidin. Science, 243, 85-88.

17. Chen, L. & Li, C.J. (2004). Facile and selective copper–palladium catalyzed addition of

terminal alkynes to activated alkynes in water. Tet. Lett., 45, 2771-2774.

18. Kodama, K., et al. (2007). Site-specific functionalization of proteins by organopalladium

reactions. ChemBioChem, 8, 232-238.

19. Li, N., Lim, R.K., Edwardraja, S., & Lin, Q. (2011). Copper-free Sonogashira cross-

coupling for functionalization of alkyne-encoded proteins in aqueous medium and in

bacterial cells. J. Am. Chem. Soc., 133, 15316-15319.

81

CHAPTER 6: UTILIZATION OF UNNATURAL AMINO ACIDS TO PROBE

CRISPR/CAS9

Introduction

CRISPR/Cas9 technology is a powerful and facile tool in genome engineering, due in large

part to its ability to selectively alter DNA by only targeting specific foreign strands entering the

bacteria without perturbing the host genome. CRISPR stands for clustered regularly interspersed

palindromic repeats and is an immune system that has evolved naturally in bacteria to protect them

from phage infections.1 This system relies on the activity of Cas9, which is an RNA-guided

endonuclease which detects and cleaves foreign viral DNA to prevent a virus from overtaking a

host cell. CRISPR/Cas9 is a revolutionary technique due to its incredible specificity, as it is capable

of recognizing and cleaving exact DNA sequences.2 Presently, researchers aim to exploit this

technology from bacteria to use in other systems to edit DNA segments containing harmful

mutations, thus ultimately restoring the DNA to its healthy state. Most notably, CRISPR/Cas9

research is underway to develop methods to genomically edit embryos to remove deleterious

genetic mutations.3

CRISPR/Cas9 acts quickly in a well-understood mechanism to target and destroy foreign

DNA in bacterial cells (Figure 6.1).3 Upon entering a cell, a virus releases double stranded DNA.

Nearly instantly, Cas9 reads the foreign DNA strand and creates a novel “spacer” into the CRISPR

locus of the bacterial genome.4 Transcription of the spacer in the CRISPR gene results in the

production of CRISPR-associated RNA (crRNA). The crRNA pairs with a tracrRNA to form a

complex, which binds both the Cas9 protein as well as the foreign viral DNA. This action brings

the Cas9 into the proper orientation and alignment to cleave the viral DNA. Following this, the

spacer remains the CRISPR gene, allowing for cleavage of the same foreign viral DNA if it again

82

enters the bacterial cell. This means that CRISPR/Cas9 offers long-term immunity, as the CRISPR

locus can contain a library of crRNAs that recognize DNA from viruses that have previously

infected the bacteria.2

The CRISPR immune system has been shown to be both highly prevalent and highly

conserved, as it appears in 90% of archaea and 40% of bacteria.5 In several recent studies,

CRISPR/Cas9 has been demonstrated to possess the

capability to disable the genomes of Heptatits B

virus, human papilloma virus (HPV), and human

immunodeficiency virus (HIV).6 Thus, research to

study Cas9 and probe its function as a part of the

CRISPR/Cas system is critical towards future efforts

to cure and prevent diseases.

Cas9 is a crescent-shaped protein with a

distinct bilobed architecture, containing a nuclease

lobe (Figure 6.2, colored) and an α-helical lobe

(Figure 6.2, gray).1 The two lobes are connected via

83

two linking segments, one of which is formed by a region rich in arginine. The nuclease lobe

contains both the HNH and RuvC domains, each of which is responsible for the cleavage of one

strand of the double-stranded DNA target. The HNH domain consists of a two-stranded antiparallel

β-sheet with four neighboring α-helices, which are critical to its role to cleave the DNA

complementary to the guide crRNA:tracrRNA complex.7 The RuvC domain consists of a six-

stranded β-sheet surrounded by four α-helices, which all contribute to the nuclease domain’s active

site, which serves to cleave the noncomplementary strand. Cas9 has two prominent clefts on its

surface, which are ultimately critical in dictating its dual function: it is a DNA-binding protein as

well as an endonuclease (Figure 6.3, A).1 On the nuclease lobe, there is a deep, narrow groove

with the RuvC domain lying at the bottom. The α-helical lobe features a wider groove, which is

positively charged along its surface due to its arginine residues (Figure 6.3, B). This cleft binds

the crRNA:tracrRNA complex, through arginine residues 69, 70, 71, and 75. Binding of the RNA

complex to the α-helical lobe induces a conformational change in Cas9 such that the RuvC domain

84

and the HNH are aligned exactly opposite each other in the cleft on the nuclease lobe, thus creating

a main channel where target DNA is aligned and cleaved.1

While bacterial CRISPR/Cas9 systems employ a crRNA:tracrRNA complex to bind target

viral DNA as well as Cas9, this approach is altered in molecular biology and gene editing

research.2,8 Instead, a chimeric single-guide RNA (sgRNA) is commonly used. The sgRNA is

prepared through the addition of a linker loop to fuse the 3’ crRNA sequence to the 5’ tracrRNA

sequence (Figure 6.4). Use of an RNA-guided wild-type Cas9 endonuclease results in a double-

stranded DNA cleavage. Mutated forms of Cas9 are also often employed in research, offering the

specific DNA binding of Cas9 without double-stranded DNA cleavage.2 Mutation of aspartic acid

to alanine at position 10 eliminates the activity of the RuvC domain while mutation of histidine to

alanine at position 840 eliminates the activity of the HNH domain. When both of these mutations

are present, catalytically dead Cas9 (dCas9) is generated. The dCas9 protein is capable of

interacting with RNA complexes to bind, target DNA sequences, but it is incapable of DNA

cleavage. Applications of dCas9 include the transcriptional activation or repression of genes as

well as the labeling of chromosomal loci.9-11 Further, dCas9 is capable of isolating DNA regions

and their associated proteins through the pull-down of specific genetic loci.12

85

Many therapeutic applications of CRISPR/Cas9 have been developed, both to counter viral

infections as well as to edit out deleterious genetic mutations in cells and embryos. As previously

mentioned, CRISPR/Cas9 technology has been used to disable the genomes of Hepatitis B, HPV,

and HIV in vitro.6 In the case of HIV, SpyCas9 has shown the ability to both block de novo

infection by cleaving the viral intermediate prior to host genome integration and to eradicate pre-

existing infections by targeting sequences in the long terminal repeat of the virus. Further, in

intestinal stem cells, CRISPR/Cas9 has been demonstrated to correct a DNA mutation that causes

cystic fibrosis by eliminating CTFR gene function.13 Genome editing via this technique was shown

to restore gene function. Current research has sought to employ CRISPR/Cas9 technology to

correct genetic mutations on embryos. The capability of CRISPR/Cas9 to achieve precise gene

targeting in one-cell stage monkey embryos has been demonstrated, indicating its ability to remove

deleterious mutations without off-target mutation occurring.14 Despite its effectiveness in many

studies, the ethics of genomic editing via Cas9 remain heavily contentious, as some might wish to

use it to give offspring preferred genes, rather than just eliminating potentially harmful ones.2

Despite success in genomic approaches, much work remains to be done to probe the

function and activity of the Cas9 protein itself. Presently, only one study has studied the

incorporation of UAAs into Cas9, demonstrating the incorporation of a photocaged UAA to

eliminate function followed by ultraviolet irradiation to restore Cas9 endonuclease function.15

Further probing of Cas9 and dCas9 via the incorporation of UAAs has the potential to enhance

and refine control of the powerful CRISPR/Cas9 technology. Here, we seek to incorporate UAAs

into Cas9 and dCas9 to explore a variety of applications, among them the use of photosensitive

UAAs in Cas9 as well as the immobilization of dCas9 onto a solid support for DNA pull-down

applications.

86

Previous Work

Previous work selected key sites for mutation within Cas9 and the inactive dCas9, choosing

sites near or in key sites of the protein, including the active sites for cleavage as well as the

arginine-rich region where the sgRNA is believed to bind (Figure 6.5, Table 6.1).

87

Site-directed mutagenesis was performed on each of these sites in both Cas9 and dCas9

plasmids. For the Cas9 plasmid, sequencing results demonstrated successful insertion of the TAG

codon at all sites except the Y72 position (Figure 6.6). For the dCas9 plasmid, sequencing has not

yet been performed, but comparison to negative control reactions indicates successful insertion of

the TAG codon at all sites except the Y72 position.

Expression of Wild Type and Mutant Cas9 and dCas9

With wild type and mutant plasmids in hand, the expression of Cas9 and dCas9 with and

without UAA incorporation was attempted. Optimization of wild type expression was attempted

prior to attempting expression of mutant Cas9. Gratifyingly, we were successfully able to express

Cas9 and dCas9 in 100 mL expression cultures, which were induced at 18°C overnight and then

88

purified via nickel resin-based affinity chromatography (Figure 6.7). Optimization of wild type

Cas9 and dCas9 expression demonstrated higher protein yield after a 48 hour induction period.

Expression of mutant, UAA-

containing Cas9 and dCas9 was

then attempted. Mutant plasmids

Cas9-D10TAG, Cas9-Y450TAG,

Cas9Y1131TAG, and Cas9-Y1265TAG were each co-transformed with a plasmid containing

pCNF, a polyspecific orthogonal aaRS/tRNA

capable of introducing a wide range of

tyrosine-derived UAAs in response to the

TAG codon. Expression was attempted with

the azide-containing UAA pAzF and the

alkyne-containing UAA pPrF (Figure 6.8).

Protein expression and purification were conducted in exactly the same manner as they

were for the wild type protein. However, expression trials under these conditions did not result in

the mutant Cas9 or dCas9 protein containing pAzF or pPrF at positions 10, 450, or 1131. However,

incorporation of pAzF at position 1265 in Cas9 was successful (Figure 6.9). A number of other

expression conditions were explored in painstaking fashion in efforts to express mutant Cas9 or

dCas9 protein

with UAA at

positions 10, 450,

or 1131 (Table

6.2). Additionally,

89

a number of expression sizes were explored, ranging from 20 µL to 250 µL. Unfortunately, as seen

by SDS-PAGE, none of these attempts was successful.

Alterations in purification technique were also investigated. Neither the use of the

traditional nickel affinity resin nor the use of a cobalt affinity resin for purification yielded mutant

protein in the elutions. Altering the incubation time of the Cas9 or dCas9 with the resin also had

no effect. Performing purification steps in a refrigerated (4°C) centrifuge also had no effect.

A possible explanation for these results is that the UAAs cannot be incorporated into

positions 10, 450, and 1131 with high fidelity due to the position on the interior of the protein of

these residues, as well as their roles in post-translational processing and folding. Position 1265 is

a surface-exposed residue, which suggests why its expression was easier to achieve, as this residue

likely plays a lesser role in protein folding dynamics. The development of effective conditions to

generate Cas9 and dCas9 mutants at positions 10, 450, and 1131 is necessary.

Cleavage Assay

To test the nuclease function of our purified wild type Cas9 samples, we performed a

cleavage assay on a target DNA plasmid. This is important, as it is necessary to verify that our

90

expressed Cas9 protein can indeed cleave DNA prior

to attempting to investigate the role of UAAs in photo-

caging or immobilization applications.

To assay the cleavage activity of Cas9, three

components are needed: an sgRNA strand, a target

DNA strand capable of hybridizing to the sgRNA, and

the WT Cas9 itself. A 121-base sgRNA (EGFP-

gRNA7) was designed based on a reported experiment

involving Cas9 in the literature.15 This sgRNA was

obtained as a single-stranded DNA oligomer, which was then amplified by PCR and analyzed by

agarose gel electrophoresis (Figure 6.10). The plasmid pIRG was selected for use as the target

DNA to be cleaved, as it is capable of hybridizing to the synthetic sgRNA (Figure 6.11). The

restriction enzyme XhoI was used to linearize the plasmid pIRG. While in plasmid form, pIRG

most likely takes on several unique supercoiled formations that appear as bands in different

locations (appear to be different sizes) when analyzed by agarose gel electrophoresis. After

successful linearization, pIRG appeared as a

single band of the correct molecular weight

(Figure 6.12).

With amplified sgRNA in DNA form,

linearized pIRG, and wild type Cas9 in hand,

cleavage reactions were performed to verify that

Cas9 would selectively cleave the DNA target.

First, the sgRNA was transcribed from DNA to

91

RNA using an in vitro transcription kit (HiScribe T7

High Yield RNA Synthesis Kit) according to

manufacturer’s protocol, with the exception of the

DNase step. Using DNase to remove the DNA

template at the end of transcription is effective but

proved problematic as residual DNase degraded the

target DNA in the cleavage assay. Therefore, DNase

was not used at any point during transcription. Wild

type Cas9 or deionized water was added to the

sgRNA, followed by linearized or plasmid pIRG, and the reaction was incubated at 37°C

overnight. Successful cleavage of linearized pIRG by Cas9 would be confirmed by two bands on

the agarose gel, and cleaved plasmid

pIRG would appear as a single band

rather than a mixture of several

supercoiled plasmid bands.

Analysis by agarose gel

electrophoresis revealed total

degradation of linearized or plasmid

pIRG when Cas9 was present, likely

due to off-site and nonspecific

cleavage by Cas9 (Figure 6.13).

However, it is also possible that

impurities in the purified wild type

92

Cas9 contain a DNase or other protein capable of DNA degradation. Present research is working

to evaluate the potential off-site cleavage by Cas9 through running the assay for a range of different

times and comparing results. If a shorter run time does not appear to have total degradation via

agarose gel electrophoresis, it would suggest that Cas9 is acting off-site and nonspecifically.

Further optimization may also include redesigning a new sgRNA to facilitate only specific

cleavage.

Conclusion

Overall, expression of UAA-containing Cas9 and dCas9 mutants has proven extremely

difficult, as has optimization of the assay for wild type Cas9. However, this project still has

excellent potential to probe Cas9 using UAAs to allow for photocontrol and immobilization

applications.


Optimized Expression of WT or UAA-containing Cas9 and dCas9

A plasmid containing WT or mutant protein sequences (pET-28b-Cas9-His or pET-dCas9-VP64-

6xHis or mutated plasmids, 2.0 µL) was co-transformed with a pEVOL-pCNF plasmid (2.0 µL)

into Escherichia coli BL21(DE3) cells using an Eppendorf eporator. The cells were then plated

and grown on LB agar in the presence of chloramphenicol (34 µg/mL) and, depending on plasmid-

encoded resistance, either kanamycin (100 µg/mL) or ampicillin (50 µg/mL) at 37°C overnight.

One colony was then used to inoculate 2xYT media (10 mL) containing both chloramphenicol

and, depending on plasmid resistance, kanamycin or ampicillin. The culture was incubated at 37°C

overnight and used to inoculate an expression culture (100 µL 2xYT media, 34 µg/mL

chloramphenicol and either 100 µg/mL kanamycin or 50 µg/mL ampicillin) at an OD600 = 0.1. The

93

culture was incubated at 37°C until it reached an OD600 = 0.7. For expressions of UAA-containing

protein, expression was induced by the addition of UAA (1 mL, 100 mM in H2O) as well as 0.8

mM isopropyl β-D-1-thiogalactopyranoside (IPTG, 100 µL) and 20% arabinose (100 µL). The

cultures were incubated at 18°C for 48 hours and then centrifuged at 5,000 rpm for 20 minutes and

stored at -80°C. To purify the protein, the cell pellet was resuspended using 500 µL of Bugbuster

(Novagen) containing lysozyme and 200 µL lysis buffer and incubated at room temperature for 30

minutes. The solution was then centrifuged at 13,000 rpm for 10 mins. The supernatant was added

to an equilibrated spin column containing HisPur Nickel Resin (Thermo Fisher Scientific). The

initial incubation of supernatant and resin was performed for 1.5 hours on ice with shaking. Cas9

was then purified according to manufacturer’s protocol. Purified Cas9 was analyzed by SDS-

PAGE and employed without further purification.

Cleavage Assay

PCR Amplification

PCR amplification was performed in a Bio-Rad iCycler 96 well reaction module thermocycler.

The protocol was adapted from the KAPA HiFi Polymerase PCR amplification protocol. A 1x

master mix was created by adding 5 µL KAPA HiFi Buffer, 0.75 µL 10 mM dNTPs, 0.75 µL of

10 mM of T7 forward primer dissolved in sterile deionized water, 0.75 µL of 10 mM of T7 reverse

primer also dissolved in sterile deionized water, and 12.25 µL of sterile deionized water. To this

solution was added 1 ng EGFP-gRNA7 template DNA (5 µL of 0.2 ng/µL). The tubes was then

loaded into the thermocycler using the following protocol: 1) initial denaturation at 95°C for 3

mins; 2) 20 cycles of: denaturation at 98°C for 20 secs, annealing at 52.3° for 15 secs, and

extension at 72°C for 15 secs; 3) final extension at 72°C for 1 min; 4) the thermocycler completed

94

the run by holding at 4°C. The samples were then PCR cleaned and concentrated (Zymo Research

DNA clean and Concentrator Kit) and eluted in 20 µL of sterile deionized water. The concentration

was determined using a NanoDrop 2000 Spectrophotometer (Thermo Fischer Scientific), and the

sample 95 was also run on a 1.5% agarose gel to assess PCR amplification efficiency. Sequences

for the EGFP-gRNA7 template and PCR primers are displayed in the table below.15

Strand Sequence Tm (°C)

EGFP-gRNA7 5’-taatacgactcactatagggagatagctagtctaggtcgatgcgttttagagctaga

aatagcaagttaaaataaggctagtccgttatcaacttgaaaaagtggcaccgactggg

tgctt-3’

-

T7 forward

primer

5’-taatacgactcactataggg-3’ 47.5

T7 reverse

primer

5’-aaagcaccgactcggtgcca-3’ 62.1

Plasmid Linearization

To a PCR tube, the following was added: 1 µg of pIRG plasmid, 5 µL of 10X CutSmart Buffer

(New England BioLabs), and diH2O up to 49 µL. One microliter of restriction enzyme XhoI (New

England BioLabs) was added (for a 50 µL total reaction volume), and the solution was thoroughly

mixed by pipetting. The tube was incubated at 37°C for 1 hour and then at 65°C for 20 minutes to

inactivate the enzyme. The sample was then PCR cleaned and concentrated (Zymo Research DNA

Clean & Concentrator Kit) and eluted in 10 µL of Cas9 activity buffer (20 mM HEPES, 150 mM

KCl, 0.5 mM DTT, 0.1 mM EDTA, 10 mM MgCl2, pH 7.4). The product was run on a 1% agarose

gel containing ethidium bromide to assess linearization efficiency.

Cleavage Reactions

The EGFP-gRNA7 DNA template (100 ng) was transcribed to RNA using the HiScribe T7 high

Yield RNA Synthesis Kit (New England BioLabs) according to the manufacturer’s protocol,

95

excluding the addition of DNase. The cleavage reactions were created by adding the following to

a PCR tube: 1 µL (12,000-16,000 ng) of unpurified synthetic gRNA, 2 µL of TAE/Mg2+ buffer

(40 mM tris-acetate, 1 mM EDTA, 12.5 mM magnesium acetate), and 40 µL WT Cas9. For control

reactions, WT Cas9 was replaced with diH2O. Next, 750 ng of target DNA (pIRG, 5 µL of 150

ng/µL, either linearized or supercoiled) in Cas9 activity buffer (20 mM HEPES, 150 mM KCl, 0.5

mM DTT, 0.1 mM EDTA, 10 mM MgCl2, pH 7.4) was added. The tubes were incubated at 37°C

for 16 hours, then at 72°C for 20 minutes to denature protein, and finally held at 4°C. The products

were run on a 0.8% agarose gel containing ethidium bromide to assess DNA cleavage.15

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2. Doudna, J. A., & Charpentier, E. (2014). The new frontier of genome engineering with

CRISPR-Cas9. Science, 346, 1258096.

3. Zhao, Y., Ying, Y., & Wang, Y. (2014). Developing CRISPR/Cas9 technologies for

research and medicine. MOJ Cell Sci. Report, 1, 6.

4. Mali, P., Esvelt, K. M., & Church, G. M. (2013). Cas9 as a versatile tool for engineering

biology. Nat. Methods, 10, 957.

5. Horvath, P. & Barrangou, R. (2010). CRISPR/Cas, the immune system of bacteria and

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6. Kennedy, E. M., & Cullen, B. R. (2015). Bacterial CRISPR/Cas DNA endonucleases: A

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7. Nishimasu, H., Ran, F.A., Hsu, P.D., Konermann, S., Shehata, S.I., Dohmae, N., Ishitani,

R., Zhang, F. and Nureki, O., 2014. Crystal structure of Cas9 in complex with guide RNA

and target DNA. Cell, 156(5), pp.935-949.

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9. Tsui, C., et al. (2018). dCas9-targeted locus-specific protein isolation method identifies

histone gene regulators. Proc. Natl. Acad. Sci. U. S. A., 201718844.

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dCas9‐based transcription factors. Plant Biotechnol. J., 13(4), 578-589.

11. Fujita, T., & Fujii, H. (2014). Identification of proteins associated with an IFNγresponsive

promoter by a retroviral expression system for enChIP using CRISPR. PLOS One, 9,

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12. Fujita, T., & Fujii, H. (2015). Isolation of specific genomic regions and identification of

associated molecules by engineered DNA-binding molecule-mediated chromatin

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13. . Schwank, G., et al. (2013). Functional repair of CFTR by CRISPR/Cas9 in intestinal stem

cell organoids of cystic fibrosis patients. Cell Stem Cell, 13, 653-658.

14. Niu, Y., Shen, B., Cui, Y., Chen, Y., Wang, J., Wang, L., Kang, Y., Zhao, X., Si, W., Li,

W., & Xiang, A.P. (2014). Generation of gene-modified cynomolgus monkey via

Cas9/RNA-mediated gene targeting in one-cell embryos. Cell, 156, 836-843.

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