Abstract Georgianna, Wesleigh Fowler Edwards

ABSTRACT

GEORGIANNA, WESLEIGH FOWLER EDWARDS. Development of Novel Methodologies for the Photoregulation of Biological Processes. (Under the direction of Dr. Alexander Deiters.)

The flow of genetic information can be affected at various stages, ultimately leading to the up- or down-regulation of gene expression. Essential to elucidating gene function is first gaining precise external control over biological processes. Many techniques currently employed to investigate gene function may interfere with normal downstream processes or harm the organism of study, and their applications often lack specificity. Photochemical biology provides an alternative approach, in which a light-removable protecting group

(termed “caging group”) installed on a protein or nucleic acid inhibits its nascent function; non-damaging light irradiation removes the caging group and restores activity. Additionally, light-responsive moieties such as diazobenzenes can be harnessed as reversible photoregulated groups for controlling biological macromolecules. Since light-irradiation can be precisely regulated, decaging can be achieved with a high level of spatial and temporal control. The research described herein reports the development of novel tools to achieve spatio-temporal control of biological processes through the photoregulation of proteins and nucleic acids, both in vitro and in vivo. Specifically, our approaches involve (1) modulating enzymatic activity by directly installing a caging group on the protein of interest, (2) utilizing photocaged nucleotides to regulate the polymerase chain reaction, (3) investigating the effects of microwave irradiation on nucleic acids, and (4) controlling protein dimerization through the use of a photoregulated chemical inducer of dimerization (CID). Development of Novel Methodologies for the Photoregulation of Biological Processes

by Wesleigh Fowler Edwards Georgianna

A dissertation submitted to the Graduate Faculty of North Carolina State University in partial fulfillment of the requirements for the Degree of Doctor of Philosophy

Chemistry

Raleigh, North Carolina

2010

APPROVED BY:

Alexander Deiters, PhD Jonathan Lindsey, PhD Committee Chair

David Muddiman, PhD Lin He, PhD

Ralph Dean, PhD

DEDICATION

This work is dedicated to

My Mom and Dad, for their unconditional love and guidance,

Steve Buchau, an exemplary teacher and scientist who put me on the right path,

Asha Balakrishnan, for sharing piano lessons, tennis balls, and this amazing lifelong

friendship,

The wonderful friends and fellow graduate students I’ve met along the way,

And to my husband, Ryan Georgianna, for his friendship, understanding and love, and for

reminding me of what matters most in life.

BIOGRAPHY

The author, Wesleigh F. Edwards Georgianna, was born December 23, 1982 to parents Bill and Lisa Edwards in Virginia Beach, VA. She grew up at the beach and enjoyed playing tennis and volunteering at Sentara Leigh Hospital in the Emergency and Surgical

Recovery Rooms during the summers. After graduating from Norfolk Academy in 2001, she attended Virginia Polytechnic Institute and State University, and worked in the lab of Dr.

Timothy Long to investigate the use of poly(lactide)s as vascular sutures. Additionally, she was an Associate Justice with the Virginia Tech Undergraduate Honor System, and earned her Emergency Medical Technician license. She graduated from Virginia Tech in 2005,

receiving a B.S. in Biochemistry with a concentration in Biotechnology.

Wesleigh then entered the Chemistry graduate program at North Carolina State

University under the direction of Dr. Alexander Deiters, where she undertook five years of

doctoral research in biological chemistry. Wesleigh has accepted a position as a project

leader at the Duke Translational Research Institute, where she works with other researchers

to transition basic medical science discoveries into important clinical therapeutics.

iii

ACKNOWLEDGEMENTS

I express my sincerest appreciation to everyone who has made this work possible. I

would particularly like to acknowledge my advisor, Prof. Dr. Alex Deiters, for his research supervision and financial support during my graduate studies. I would also like to thank the former and current members of the Deiters Lab, especially Andrew, Yan, Jeane, Colleen, Jie,

Hank, Jesse, and Harry for their humor and advice. Thank you also to my advisory committee for their assistance in the completion of my doctoral research.

Additionally, thank you to those who have particularly contributed to my graduate studies through their guidance and indispensible fellowship. Doug, there are not enough words to tell you how grateful I am for your help over the last five years, and more importantly, your friendship. Virginia, thank you for your research counsel and for always being there to listen, both in graduate school and beyond. Thank you also to Elke, Rania, and

Jennifer, with whom I served on PLU and whose camaraderie I very much enjoyed.

I would also like to thank my Mom and Dad for the love and support I have received

from them; I am blessed to have parents who wholeheartedly encouraged me from the

beginning. Lastly, I would especially like to thank my husband, Ryan, who has kept me

motivated these last five years, and has been there for me with research advice,

understanding, laughter and companionship, and love.

TABLE OF CONTENTS

LIST OF TABLES………………………………………………………………………………. viii

LIST OF FIGURES……………………………………………………………………………..... ix

CHAPTER 1: INTRODUCTION TO PHOTOCHEMICAL BIOLOGY………………………… 1

CHAPTER 2: PHOTOCHEMICAL REGULATION OF THE POLYMERASE CHAIN

REACTION……………………………………………………………………………………… 22

2.1 Introduction to the polymerase chain reaction………………………………………… 22

2.2 Hybridization studies with photocaged DNA…………………………………………. 23

2.3 Light-activation of PCR……………………………………………………………….. 26

2.4 Light-deactivation of PCR…………………………………………………………….. 29

2.5 Concurrent up- and down-regulation of PCR…………………………………………. 32

2.6 Summary and outlook…………………………………………………………………. 32

2.7 Experimental methods…………………………….……………………………………34

CHAPTER 3: THE EFFECT OF MICROWAVE IRRADIATION ON OLIGONUCLEOTIDE

HYBRIDIZATION………………………………………………………………………………. 36

3.1 Introduction to DNA hybridization and microwave irradiation………………………. 36

3.2 Development of a fluorescence hybridization assay…………………………………... 37

3.3 Microwave irradiation of DNA……………………………………………………….. 40

3.4 Summary and outlook…………………………………………………………………. 48

3.5 Experimental methods………………………………………………………………….49

CHAPTER 4: LIGHT-REGULATION OF PROTEIN FUNCTION THROUGH

PHOTOCAGING………………………………………………………………………………… 52

4.1 Lysozyme………………………………………………………………………………. 52

4.1.1 Introduction to lysozyme……………………………………………………………... 52

4.1.2 PhotoPEGylation of lysozyme………………………………………………………... 54

4.1.3 In vitro activity assays of lysozyme function………………………………………… 58

4.1.4 Summary and outlook………………………………………………………………… 61

4.1.5 Experimental methods………………………………………………………………... 62

4.2 Cre Recombinase……………………………………………………………………... 64

4.2.1 Introduction to DNA recombination and Cre recombinase…………………………... 64

4.2.2 Expression and purification of wild type Cre………………………………………… 71

4.2.3 In vitro non-specific caging of lysine residues……………………………………...... 74

4.2.4 Unnatural amino acid mutagenesis …………………………………………………... 76

4.2.5 Expression and purification of CreONBY……………………………………………. 80

4.2.6 In vitro light-activation……………………………………………………………...... 82

4.2.7 In vivo light-activation to demonstrate spatiotemporal control of gene function…….. 83

4.2.8 Expression of TAT-modified Cre…………………………………………………...... 88

4.2.9 Summary and outlook………………………………………………………………… 90

4.2.10 Experimental methods………………………………………………………………... 91

4.3 Photochemical control over FimE-mediated DNA recombination in E. coli………… 99

4.3.1 Introduction to FimE………………………………………………………………..... 99

4.3.2 FimE expression and in vivo assays………………………………………………… 104

4.3.3 Summary and outlook……………………………………………………………….. 105

4.3.4 Experimental methods………………………………………………………………. 106

4.4 Photochemical control over Flpe-mediated DNA recombination in E. coli………… 108

4.4.1 Introduction to Flpe recombinase………………………………………………….... 108

4.4.2 Expression of wild type Flpe………………………………………………………... 110

4.4.3 The Flpe bacterial assay……………………………………………………………... 110

4.4.4 Summary and outlook……………………………………………………………….. 113

4.4.5 Experimental methods………………………………………………………………. 114

CHAPTER 5: LIGHT-REGULATION OF PROTEIN DIMERIZATION…………………….. 119

5.1 Introduction to natural and synthetic protein dimerizers……………………………… 119

5.2 Expression and purification of DHFR and In Vitro Dimerization Assays……………. 128

5.3 Expression and purification of GFP-DHFR and In Vitro Fluorescent Dimerization

Assays…………………………………………………………………………………. 132

5.4 In Vivo Dimerization Assays………………………………………………………….. 138

5.5 Summary and outlook…………………………………………………………………. 139

5.6 Experimental methods……………………………………………………………….... 140

CHAPTER 6: THE HALO-TAG AS A PROTEIN-BASED CAGING GROUP……………… 148

6.1 Introduction to the HaloTag protein…………………………………………………... 148

6.2 Expression and purification of HaloTag………………………………………………. 150

6.3 Summary and outlook…………………………………………………………………. 152

6.4 Experimental methods………………………………………………………………… 152

APPENDIX 1: RECIPES………………………………………………………………………. 155

REFERENCES…………………………………………………………………………………. 157

vii

LIST OF TABLES

Table 2.1. Melting temperatures of non-caged (P1) and caged (P2-P7) oligonucleotides before and after irradiation……………………...……………………………………… 25

Table 3.1. Sequences and melting temperature of DNA oligomers……………………. 38

Table 4.2. Percentage of red and green fluorescence in HEK-293T cells transfected with

pC-SL and/or caged Cre as detected by flow cytometry………………………………... 88

Table 4.3. Expression of FimE from pFIP and subsequent recombination……………. 105

viii

LIST OF FIGURES

Figure 1.1. Comparison between “forward” and “reverse” genetics approaches………… 2

Figure 1.2. Light-irradiation of a caged substrate………………………………………… 5

Figure 1.3. The RNAi pathway is initiated by dsRNA or precursor miRNA to down-

regulate gene expression…………………………………………………………………..7

Figure 1.4. Evaluation of RNAi in zebrafish embryos…………………………………...10

Figure 1.5. Light-irradiation of antisense activity through the incorporation of light-

removable caging groups………………………………………………………………... 12

Figure 1.6. A photocaged morpholino targeting the ntl gene in zebrafish embryos induces

the mutant phenotype upon UV irradiation……………………………………………... 13

Figure 1.7. Use of a light-activated anti-chordin negatively-charged PNA (ncPNA) to

control gene expression in zebrafish embryos…………………………………………... 15

Figure 1.8. Formation and light-activation of the PhotoMorph heteroduplex…………... 16

Figure 1.9. PhotoMorph efficacy in zebrafish embryos………………………………… 18

Figure 1.10. Renilla luciferase signal after PS-DNA transfection and ± UV irradiation

(365 nnm, 23 W, 5 min), normalized to the transfection of the non-caged PS-DNA

antisense agent…………………………………………………………………………... 19

Figure 1.11. Spatial regulation of Renilla luciferase expression in NIH-3T3 cells with

caged PS-DNA antisense agents………………………………………………………… 20

Figure 2.1. The polymerase chain reaction enables to rapid amplification of a DNA

template………………………………………………………………………………….. 22

Figure 2.2. Photodeprotection of the NPOM group (red) to yield thymidine…………... 24

Figure 2.3. PCR activation by light……………………………………………………... 27

Figure 2.4. Agarose gel of dsDNA amplified by light-regulated PCR………………….. 28

Figure 2.5. Time-course of a PCR, light-activated at cycle 15………………………….. 29

Figure 2.6. Light-deactivation of PCR………………………………………………….. 30

Figure 2.7. Time-course of a PCR light-deactivated at cycle 10………………………... 31

Figure 2.8. Light-switching between two different PCRs through simultaneous activation and deactivation of primers……………………………………………………………... 32

Figure 3.1. Schematic of the fluorescence quenching assay used to determine if

microwave irradiation (MW) is capable of melting dsDNA……………………………. 38

Figure 3.2. The CEM Discover microwave synthesizer………………………………… 40

Figure 3.3. Fluorescence signals arising from various conditions employed in the DNA melting experiments with D1F and D1R………………………………………………... 42

Figure 3.4. Representative power, pressure, and temperature profile graphs show typical reaction conditions at various microwave powers………………………………………. 43

Figure 3.5 Microwaved DNA sustains no damage as detected with denaturing PAGE… 44

Figure 3.6. Microwave melting curves indicate that microwave irradiation induces DNA melting at a specific power regardless of the duplex length……………………………. 46

Figure 4.1. Lysozyme-catalyzed hydrolysis of N-acetylmuramic acid and N-

acetylglucosamine………………………………………………………………………. 53

Figure 4.2. Structure and PEGylation of lysozyme……………………………………... 54

Figure 4.3. Structure of the Photo-PEG reagent………………………………………… 56

Figure 4.4. SDS-PAGE analysis of lysozyme PEGylation……………………………… 57

Figure 4.5. (NHS)-PEGylation of lysozyme results in partial to complete attenuation of

enzyme activity…...……………………………………………………………………... 58

Figure 4.6. Micrococcus lysodeikticus exposed to different lysozymes for 15 min,

followed by an optical density (OD450)measurement………………………………….. 59

Figure 4.7. Micrococcus lysodeikticus exposed to Photo-PEGylated lysozyme followed

by an optical density (OD450) measurement after 15 min……………………………… 60

Figure 4.8 Lysis assays performed with varied amounts of wild type lysozyme as compared to irradiated PhotoPEGylated enzyme allowed the calculation of Vmax values and subsequent determination of percent activity restoration to the PhotoPEGylated lysozyme………………………………………………………………………………… 61

Figure 4.9. Site-specific recombination occurs through formation and then resolution of the Holliday junction……………………………………………………………………. 65

Figure 4.10. Cre recombinase catalyzes the inversion or excision/integration of DNA

between two loxP sites…………………………………………………………………... 66

Figure 4.11. Heat shock induced Cre recombination in Zebrafish affords control over

DNA recombination and expression of the kRASG12D oncogene……………………... 69

Figure 4.12. Cre activity has been modulated by a mutated estrogen receptor ligand- binding domain fusion, Cre-ERT, that binds tamoxifen (ER ligand) …………………… 70

Figure 4.13. Expression of Cre recombinase from pET-21Cre yields pure protein, as determined by SDS-PAGE analysis…………………………………………………….. 72

Figure 4.14. In vitro assay of Cre activity…………………………….………………… 73

Figure 4.15. The NPOC photocage attenuates Cre activity until it is removed with 365 nm

UV light to yield active enzyme……………………………….………………………... 75

Figure 4.16. Crystal structure of the active site of Cre recombinase……………………. 77

Figure 4.17. A general approach for the site-specific incorporation of unnatural amino acids into proteins in vivo……………………………………………………………….. 80

Figure 4.18. Expression of Cre recombinase in its wild-type form and containing a caged tyrosine at position 324………………………………………………………………….. 81

Figure 4.19. Expression with our modified pEVOL produced a greater amount of

CreONBY than expression with pSupONBY…………………………………………… 82

Figure 4.20. Light irradiation of CreONBY restored about 70% activity compared to that of WT Cre……………………………………………………………………………….. 83

Figure 4.21. The Cre Stoplight plasmid (pC-SL)……………………………………….. 84

Figure 4.22. Spatial and temporal control of Cre recombinase in vivo…………………. 85

Figure 4.23. In the absence of CreONBY, UV irradiation (365nm, 23W, 5 min) does not affect the conversion of DsRed to GFP in cells transfected with pC-SL………………...86

Figure 4.24. Results of flow cytometry counting of transfected cells…………………... 87

xii

Figure 4.25. Expression of TATCre from pTriEx-HTNC yields highly pure protein, as

determined by SDS-PAGE analysis…………………………………………………….. 89

Figure 4.26. FimE mediated recombination inverts the promoter (Ptrc*) located between

the inverted repeat right (IRR) and inverted repeat left (IRL) of the FimS, leading to GFP

expression……………………………………………………………………………… 102

Figure 4.27. FimE recombination as measured by GFP expression levels, which differ

greatly between induced and uninduced cultures……………………………………… 103

Figure 4.28. Flpe recombinase catalyzes the inversion or excision/integration of DNA between two frt sites…………………………………………………………………… 109

Figure 4.29. Expression of Flpe recombinase from pET-21Flpe yields pure protein, as determined by SDS-PAGE analysis…………………………………………………… 110

Figure 4.30. The Flpe-mediated DNA recombination reporter assay…………………. 111

Figure 5.1. CID-mediated protein dimerization enables the bridging of two proteins or two split protein domains……………………………………………………………… 120

Figure 5.2. The yeast two-hybrid system……………………………………………… 121

Figure 5.3. Rapamycin-induced dimerization of FRB and FKBP in a yeast three-hybrid system leads to expression of a reporter gene…………………………………………. 122

Figure 5.4. Use of the yeast three-hybrid system with a novel DEX-MTX CID leads to

expression of β-galactosidase and a blue color upon successful dimerization………… 123

Figure 5.5. The antibiotic trimethoprim………………………………………………... 124

xiii

Figure 5.6. Selective chemical labeled of subcellularly targeted E. coli DHFR in CHO

cells…………………………………………………………………………………….. 125

Figure 5.7. Yeast three-hybrid assay of TMP-SLF activity…………………………… 126

Figure 5.8. Activation of FucT7 as determined by fluorescent anti-sLex……………… 127

Figure 5.9. The novel light-switchable Bis-TMP CID, constructed with a photoresponsive

DAB core flanked by two TMP molecules……………………………………………. 128

Figure 5.10. Expression of wild type E. coli DHFR from pET21-DHFR in BL21(DE3)

Gold cells yields high purity protein, as determined by 12% SDS-PAGE

analysis………………………………………………………………………………… 129

Figure 5.11. SDS-PAGE analysis of the DHFR activity assay using MTX-resin……... 130

Figure 5.12. The in vitro DHFR dimerization assay…………………………………... 131

Figure 5.13. SDS-PAGE analysis of resin-immobilized DHFR incubated with Bis-TMP

and irradiated 30 min (365 nm, 25 W)………………………………………………… 132

Figure 5.14. Expression of GFP-DHFR from pBAD+GFP-DHFR in BL21(DE3) Gold

cells yields high purity protein, as determined by 12% SDS-PAGE analysis…………. 133

Figure 5.15. Fluorescence levels in the in vitro DHFR assay using both DHFR and GFP-

DHFR…………………………………………………………………………………... 135

Figure 5.16. Structure of Bis-MTX-C9 and a schematic depiction of the dimerized

nanoring formed when two peptide-linked DHFR molecules are incubated with Bis-

MTX-C9………………………………………………………………………………... 136

xiv

Figure 5.17. SEC analysis of trimer, dimer and internal monomer formed upon incubation

of tethered DHFR with Bis-MTX-C9………………………………………………….. 136

Figure 5.18. SEC analysis of Bis-TMP induced DHFR dimerization…………………. 137

Figure 5.19. SEC analysis of Bis-TMP induced tethered DHFR dimerization in the presence of 100 mM NADPH…………………………………………………………. 138

Figure 6.1. The HaloTag protein forms a covalent bond with a ligand containing a

reactive chloroalkane linker……………………………………………………………. 149

Figure 6.2. SDS-PAGE analysis of HaloTag protein expression……………………… 151

Figure 6.3. The dansyl-chloroalkane HaloTag linker………………………………….. 151

CHAPTER ONE – INTRODUCTION TO PHOTOCHEMICAL BIOLOGY

Whole-genome sequencing of numerous organisms has created a wealth of data that

must now be annotated in order to better understand the information stored in an organism’s

genome. The field of functional genomics relies on the perturbation of gene function by interfering with transcription, translation, or protein function to further understand genetic processes and to associate them with their corresponding genes. Traditionally, two general approaches have been used to investigate gene function.[1] The first approach, termed

“forward genetics”, identifies a phenotype (an observable characteristic, trait, or morphology) of interest and traces it back to a genotype (the inherited instructions within the genetic code); conversely, the second approach, “reverse genetics”, involves first mutating and thus perturbing a gene followed by correlating the phenotypic result[2] (Figure 1.1).

Figure 1.1. Comparison between “forward” and “reverse” genetics approaches, as performed in yeast as a model organism; however, identical approaches can be applied in other organisms as well. A forward genetics approach is a classical genetic screen that seeks to determine the gene(s) responsible for an observed phenotype. In contract, reverse genetics screens study the phenotype effects produced by perturbing a known gene. From Stockwell, B. R. Chem. Rev. Genet. 2000, 1, 116.

Methods employed in gene perturbation include (1) mutagenesis of gene sequence by chemical means,[3, 4] (2) mutagenesis via insertion of known transgenes[5] or transposable elements,[6, 7] (3) targeted gene disruption through homologous recombination-based techniques,[8-10] and (4) RNA interference (RNAi).[11] Method (1) provides a simple and

2 rapid result in that both means efficiently create random point mutations within the DNA sequence; however, the phenotypic results may be difficult to distinguish from naturally occurring single nucleotide polymorphisms (SNPs),[12] and the mutational process is difficult to control which may thus irreparably damage the organism under study. Method (2) allows easy detection of the inserted sequences and subsequent phenotype identification, but shares the same limitations as Method (1).[13] Method (3) has been particularly useful, specifically in the generation of targeted gene knockouts in model organisms such as the mouse,[14] as a gene may be deleted or inserted through the use of recognition sites located engineered into the genome at specific positions. Of all the previously described methods, (4) is regarded as an especially powerful tool for perturbing gene function,[15] as RNAi can effectively silence a single target RNA transcript that is present in thousands of copies per cell, leading to a discernible phenotype directly correlated to the gene.[15]

None of the methods described above can be regulated with high spatial and temporal resolution. However, spatio-temporal control over gene function is necessary in order to study biological systems with a level of precision that is similar to that found in nature.

Additionally, a means to study real time events in cellular processing and gene expression would be invaluable in elucidating gene function, predicting protein interactions and pathways, and achieving precise external control over cellular events. Recently, the developing field of photochemical biology has provided new tools in functional genomics to affect and study gene function. Here, the perturbation of gene function is triggered by light irradiation, which has the advantage of being precisely regulated in a spatial and temporal

fashion. Thus, the use of light allows a fine level of control over biological processes to

subsequently enable robust studies of gene function and processes in vivo without damage to

the system. Additionally, as light irradiation is non-invasive, its utilization does not affect

natural interactions within the system. This is in contrast to aforementioned techniques,

whose introduction into the cell or organism can perturb multiple biological functions.

One approach to the development of light-activatable biological processes is the

installation of a light-removable protecting group, a so called “caging group”, on the

biomolecule involved in the process of interest. This caging group can be placed directly on a protein or small molecule to inhibit its nascent activity[16-19] (Figure 1.2, A). Exposure to

UV light of a non-damaging wavelength (365 nm)[20] removes the caging group and restores

the molecule’s biological function. As light is non-invasive and can be precisely regulated in

location, time, and amplitude, spatial and temporal control over the activity of the molecule

can be achieved.[19] A successful caging approach must meet the following criteria: (1)

caging should render the biomolecule completely inactive and the caging group needs to be

stable to hydrolysis, (2) decaging must occur at a wavelength of light that does not harm the

cell or organism, (3) the biomolecule must be released in high yield and at a fast rate, and (4)

any by-products of the photo-deprotection reaction should not be toxic or interfere otherwise

with the biological system.[21] Various caging groups have been developed, many based on the photolysis of the ortho-nitrobenzyl (ONB) moiety (Figure 1.2, B).[22-26] ONB-based

caging groups are advantageous due to their ease of synthesis, their light sensitivity, and their

compatability with a variety of functional groups, such as phosphates, carboxylates,

hydroxyls, and amines.[21] Several small molecules,[27-29] nucleic acids,[30-33] and proteins[34-37] have been photocaged successfully.

Figure 1.2. Light-irradiation of a caged substrate. A. Conceptual scheme for a decaging reaction. B. Generation of an active molecule from an inactive, caged molecule by removal of the o-nitrobenzyl derived caging group with UV light. R = H, CH3 or CO2H; X = O, N, or S.

In theory, photocaging groups have potential applications in the field of functional

genomics, notably through loss-of-function reverse genetics techniques, in which the deletion

or perturbation of a gene provides a phenotype that can be correlated to the function of the

disrupted gene.[38-40] However, to date, photocaging groups have only been used to

establish proof-of-concept and have not yet been applied to functional genomics studies, as

caging 1) requires chemical synthesis to make and install the caging group(s), which may not

be accessible in genomics labs; 2) is typically employed on a small scale not amenable to

large-scale genomic screens; and importantly, 3) requires specific foreknowledge of the

native pathway or process under study in order to strategically incorporate the caging

group(s) and successfully control biological function. Regardless, everal examples exist in

which caging strategies have been implemented to demonstrate their utility in achieving

spatiotemporal control over biological processes. The first such example utilizes RNA

interference (RNAi) to provide transient gene silencing or knock-down expression in cell

culture and model organisms.[41] RNAi is a naturally occurring cellular mechanism involved

in post-transcriptional eukaryotic gene regulation by degrading target mRNA, or inhibiting

its translation.[42-45] The mechanism involves small interfering RNAs (siRNAs), which are

typically introduced into the cell by injection or transfection of double-stranded RNA, or by

expression from a transfected plasmid.[43, 46, 47] To initiate RNAi, the enzyme Dicer

cleaves the double-stranded RNA to produce single-stranded, active siRNA.[48] These

nucleic acids, typically 21-23 nucleotides in length, are then loaded into the RNA-Induced

Silencing Complex (RISC),[49] where they bind to the protein Argonaute and act as a

template for mRNA sequence recognition.[50] Base-pair mismatches between the RISC complex and the mRNA lead to translational repression, while exact complementarity drives mRNA cleavage[44, 51, 52] (Figure 1.3).

Figure 1.3. The RNAi pathway is initiated by dsRNA to down-regulate gene expression. Formation of the RISC effector complex leads to mRNA degradation or steric blocking of translation. Adapted from Carthew, R. W. Curr. Opin. Genet. Dev., 2006, 16, 203.

While the RNAi mechanism is a highly effective regulator of gene expression, a

major drawback to its implementation in functional genomics is the susceptibility of dsRNA

to enzymatic degradation in vivo by Protein Kinase R,[53] which affects the efficacy and

duration of gene silencing.[54] In order to optimize RNAi, chemical modifications have been

designed to better stabilize dsRNA and prevent off-target effects.[55-58] Additionally, to

obtain a more precise level of spatiotemporal control over silencing, small molecule-

inducible promoters for dsRNA expression have been reported,[59, 60] as well as small

molecule effectors that regulate the RNAi pathway itself. [61, 62]

More recently, the use of photolabile protecting groups installed on siRNAs has enabled the regulation of gene knockdown in a temporal fashion.[63, 64] Caging groups

installed on the nucleobases or the phosphodiester backbone of RNA have been used to

prevent Dicer activity and the siRNA-RISC interaction, as well as inhibit scission of the

RNA target within the RISC. For example, Mikat and Heckel inserted nitrophenylpropyl

(NPP) caging groups at the O-4 and O-6 postions of thymidine and guanosine, respectively.[65] The NPP groups inhibit hydrogen bonding and prevent base pairing, forming a bulge between the siRNA and mRNA target within the RISC that attenuates cleavage. Prior to irradiation, the caged siRNA was unable to catalyze RNA scission in the

RISC. Upon UV exposure, the caging groups were removed and base pairing restored, leading to RISC-induced cleavage and successful RNAi.

Alternatively, Nguyen et al. showed that a single di-methoxynitrophenylethyl

(DMNPE) caging group placed on the 5’-phosphate of a single-stranded siRNA was able to prevent its binding to the RISC before UV irradiation.[64] While fully active siRNAs were

obtained after irradiation, residual RNAi activity was still detected prior to irradiation. To eliminate this activity, Shah et al. modified the dsRNA by placing a phosphate group on the

5’ and 3’ ends of both the sense and antisense strands. Thus, by selectively installing one

DMNPE caging group on each terminal phosphate, the residual activity prior to irradiation could be eliminated. As such, this approach was shown to completely inhibit Dicer processing and subsequent RISC loading prior to irradiation, whereupon UV exposure led to removal of the caging groups and activation of the RNAi pathway. Additionally, this caging

8 strategy resulted in efficient removal of the DMNPE groups from the dsRNA, eliminating any phototoxicity associated with long periods of UV irradiation.[66]

Blidner et al. utilized a similar approach to achieve photoinduced RNA interference in cell culture and in zebrafish. The phosphate backbone of 2’-fluoro substituted nucleic acids (FNAs) was modified with the DMNPE caging group.[67] The caging group mediated

RNAi-induced knock-down of a GFP reporter gene in zebrafish upon exposure to 365 nm of light (Figure 1.4, A-C), while also stabilizing the FNAs to non-specific nuclease degradation

(Figure 1.4, D).

Figure 1.4. Evaluation of the light-activation of RNAi in zebrafish embryos. A. Injection analysis. Dextran tracking dye was observed throughout the injected region, demonstrating a successful injection (left panel), while delivery of a GFP expression plasmid showed mosaic expression, as expected (middle panel). Overlay of the dextran dye and GFP signals (right panel). B. Knock-down activity with no siRNA (left panel), with control siFNA (middle panel) and with irradiated control siFNA (right panel). Irradiated and nonirradiated control siFNA produced identical GFP knock-down and demonstrated that light has no effect on RNAi activity. C. Caged siFNA (middle panel) produces no GFP knock-down, resembling the GFP control (left panel); however, irradiation (365 nm, 40 J cm2, 7 hours post-ferzilization) resulted in silencing of gene expression (right panel). D. While the 2’fluoro-modification protected the nucleic acids from non-specific endonuclease degradation for about 5 min, the DMNPE modification on the FNA provided more than 80 min of protection. Adapted from Blidner, R. A.; Svoboda, K. R.; Hammer, R. P.; and Monroe, W. T. Mol. Biosyst. 2008, 4, 431.

Another example of a novel photocaging strategy that could be applied to functional genomics involves the use of caging groups to spatiotemporally regulate antisense techniques. This methodology provides an additional way to perturb gene function followed by phenotypic analysis.[68, 69]. Antisense agents are comprised of DNA or modified DNA,

such as phosphorothioate DNA (PS-DNA),[70] morpholinos (MOs),[71] locked nucleic acids

(LNAs),[72] and peptide nucleic acids (PNAs).[73] Each agent modulates gene expression at

the translational level to prevent protein production by inducing enzymatic degradation or

steric blocking of mRNA. In the case of DNA and PS-DNA, the enzyme RNaseH degrades

the RNA strand of the (PS-)DNA:RNA duplex, thereby degrading the target mRNA.[74]

However, RNaseH does not recognize MOs, LNAs, or PNAs, and these agents prevent

translation by blocking access of the mRNA to the ribosome.[75]

While antisense technology provides a powerful tool for the study of gene function, its use is limited by a lack of spatial and temporal control. Upon introduction to a developing embryo, the antisense agent is immediately active and may be distributed to daughter cells.[76] This is problematic when the knock-down gene target is involved in early

development and silencing leads to embryonal lethality. However, installation of a

photolabile protecting group on the antisense agent prevents hybridization to complementary

mRNA, inhibiting its activity; this is in contrast to the previously described light-activated

siRNA approaches, where installed caging groups did not fully abrogate RNA:RNA duplex

formation. UV irradiation then removes the caging group and allows the antisense agent to

hybridize.[20, 77-80] Subsequent RNaseH degradation or prevention of translation thus

occurs (Figure 1.5).

Figure 1.5. Light-regulation of antisense activity through the incorporation of light-removable caging groups (grey squares). Brief UV irradiation at 365 nm removes the caging groups, enables sequence-specific mRNA binding, and blocks translation or activates RNaseH catalyzed cleavage of the transcript. From Young, D. D.; Lusic, H.; Lively, M. O.; Yoder, J. A.; and Deiters, A. ChemBioChem. 2008, 1318, 2937.

Furthermore, the Chen group demonstrated light-controlled gene silencing in zebrafish embryos using a light-activated morpholino antisense agent that targeted the no tail

(ntl) gene.[78] A morpholino complementary to ntl mRNA was tethered to a short sequence of complementary oligonucleotides through a dimethoxynitrobenzyl-based photocleavable linker (Figure 1.6). Light irradiation cleaved the linker and activated the free morpholino.

Zebrafish embryos were injected at the one-cell stage with the ntl-targeting caged

morpholino and were either globally irradiated (360 nm, 10 sec bursts) four hours post-

fertilization or grown in the dark. Loss of ntl expression was noted in the irradiated embryos

by an abnormal tail phenotype, and Western blot analysis showed successful knock-down of

12 the ntl gene (Figure 1.6). The Chen group thus demonstrated the applicability of photocaged antisense agents to the temporal control of gene expression in a model organism.[78]

B. C.

D. E.

Figure 1.6. A photocaged morpholino targeting the ntl gene in zebrafish embryos induces the mutant phenotype upon UV irradiation.A. General scheme for the caged morpholino and its hybridization to an RNA target upon light-triggered cleavage and dissociation of the inhibitor. B. Wild type zebrafish embryo. C. Irradiation alone does not induce the ntl phenotype in the wild type zebrafish embryo. D. Embryos injected with the ntl caged morpholino (cMO) develop normally without UV light exposure. E. Irradiation at the sphere stage activates the cMO and results in the ntl mutant phenotype. F. Western blot analysis confirms loss of ntl expression in the presence of UV irradiation, while no effect is seen with the β-actin control. Adapted from Shestopalov, I. A.; Sinha, S.; and Chen, J. K. Nat. Chem. Biol. 2007, 3, 650.

Alternatively, photocontrol over gene expression in zebrafish was demonstrated by the Dmochowski group, who used light-activated, negatively-charged PNAs (ncPNAs) to target the chordin gene.[81] Hybridization of the 18-mer antisense ncPNA to its target mRNA was prevented by a complementary 12-mer 2’-OMe RNA sense strand attached to the ncPNA via a 1-(5-(N-maleimidomethyl)-2-nitrophenyl) ethanol N-hydroxysuccinimide ester photocleavable linker (Figure 1.7) that formed a hairpin structure. In the absence of light, no mutant phenotype was observed, as the antisense strand could not bind its target mRNA and block assembly of the ribosome; however, 8 min of irradiation, three hours post-fertilization removed the complementary sense strand and enabled the ncPNA to bind its target and inhibit the ribosome. This prevented chordin expression, as evidenced by an abnormal vertebral dorsum in the zebrafish embryo (Figure 1.7, A-D).

Figure 1.7. Use of a light-activated anti-chordin negatively-charged PNA (ncPNA) to control gene expression in zebrafish embryos. The antisense ncPNA strand is blocked by a 2’-OMe sense strand; the two strands are attached via a 1-(5-(N-maleimidomethyl)-2-nitrophenyl) ethanol N-hydroxysuccinimide ester photocleavable linker. In the absence of irradiation, the antisense strand cannot bind its mRNA target and the chordin gene is expressed. Upon irradiation and photolysis of the linker, the antisense ncPNA hybridizes to its target and inhibits translation. A-D. Transmitted light images of representative zebrafish embryos 24 h post-fertilization. A. The uninjected embryo exhibits normal development. B. In the absence of irradiation, injection with 0.5 mM ncPNA-2’-OMe-RNA results in a normal embryonic phenotype. C. In the absence of irradiation, injection with 1.0 mM ncPNA-2’-OMe-RNA results in a normal embryonic phenotype. D. An embryo injected with 0.5 mM ncPNA-2’-OMe-RNA and irradiated 8 min, 3 h post-fertilization, displays the mutant chordin phenotype with a small, underdeveloped backbone. Adapted from Tang, X.; Maegawa, S.; Weinberg, E. S.; Dmochowski, I. J. J. Am. Chem. Soc. 2007, 129, 11000.

More recently, the Mayer group utilized light-activatable morpholinos (PhotoMorphs) to control gene expression in zebrafish.[80] These PhotoMorphs were generated by hybridizing morpholinos to complementary 25-mer RNAs, which contained a photocleavable nitrophenyl linker in the phosphodiester backbone between bases 12 and 13 (Figure 1.8).

Thus, the complementary strand prevents the morpholino from hybridizing to its target mRNA. Upon light irradiation, the complementary strand is removed, enabling the morpholino to bind to its mRNA target and inhibit translation.

Figure 1.8. Formation and light-activation of the PhotoMorph heteroduplex. The morpholino and caging strand are hybridized to form the PhotoMorph. UV irradiation (365 nm) cleaves the caging strand at the photocleavable linker, causing the strand to dissociate and liberating the morpholino. From Tomasini, A. J.; Schuler, A. D.; Zebala, J. A.; Mayer, A. N. Genesis 2009, 47, 736.

A PhotoMorph targeting the GFP gene was first used as a proof-of-concept to evaluate the efficiency of the caged and decaged morpholino. Transgenic zebrafish gut larvae were injected with either a wild type GFP morpholino (MO) or the GFP PhotoMorph, then

irradiated (365 nm) four hours post-fertilization. Zebrafish injected with the MO displayed no GFP fluorescence, while those injected with the GFP PhotoMorph showed fluorescence in

the absence of UV irradiation (Figure 1.9, A). However, a 90% knock-down in fluorescence

was detected in irradiated embyos compared to the uninjected control, illustrating that the

PhotoMorph is an efficient, light-activated antisense technique.

To further demonstrate the efficacy of this approach, another PhotoMorph targeting

the ntl gene was injected into zebrafish embryos at the one-cell stage, then irradiated 1.5, 4,

or 9 hours post-fertilization (Figure 1.9, B). Zebrafish embryos injected with non-caged MO

exhibited the mutant, no tail phenotype, while embryos injected with the ntl PhotoMorph

showed no mutation in the absence of irradiation. However, upon light irradiation at 1.5

hours post-fertilization, a severe no tail phenotype was observed, which became less severe if

embyos were irradiated 4 hours post-fertilization. Irradiation at 9 hours post-fertilization did

not result in a mutant phenotype; embryos looked identical to the uninjected wild type

control. PhotoMorphs were thus successfully employed to achieve light-activatable control

over gene expression in zebrafish.

Figure 1.9. PhotoMorph efficacy in zebrafish embryos. A. The GFP PhotoMorph displays little activity prior to light-irradiation, as compared to the wild type, uninjected control, while embryos injected with non-caged MO showed no GFP fluorescence. Irradiaton of the PhotoMorph four hours post-fertilization resulted in a 90% decrease in fluorescence compared to the wild type. B. Embryos injected with the ntl PhotoMorph display no mutant phenotype prior to light-irradiation, while injection with non-caged MO results in the no tail phenotype. Embryos injected with the ntl PhotoMorph and irradiated 1.5 hours post-fertilization show a mutant phenotype similar to the non-caged MO control. This phenotype is less severe when irradiation occurs 4 hours post- fertilization. When irradiated 9 hours post-fertilization, the ntl PhotoMorph-injected embryos display a normal phenotype. Adapted from Tomasini, A. J.; Schuler, A. D.; Zebala, J. A.; Mayer, A. N. Genesis 2009, 47, 736.

Photocaged antisense agents were also investigated in the Deiters Lab by Dr. Douglas

Young.[77, 82] PS-DNA antisense agents were synthesized that contained three and four 6- nitropiperonyloxymethyl (NPOM) caging groups installed with standard DNA synthesis techniques and evenly spaced throughout the PS-DNA oligomer. The caged PS-DNA was then utilized to silence expression of a Renilla luciferase reporter gene in mammalian cell culture, demonstrating both spatial and temporal control over antisense activity. Caged and non-caged PS-DNA were transfected into NIH-3T3 cells, along with a luciferase expressing reporter plasmid. The sequence of the antisense agent was previously demonstrated to target

the Renilla luciferase gene.[83] A PS-DNA control with no silencing activity was also

transfected. Cells were then briefly irradiated at 365 nm and the luciferase signal quantified

(Figure 1.10). The non-caged PS-DNA successfully silenced luciferase expression, while the

PS-DNA control showed no silencing, as compared to non-transfected cells. Importantly, 3-

and 4-caged PS-DNA showed no activity until irradiation; the luciferase signal was then

decreased to that of the non-caged PS-DNA. In this manner, temporal control over gene

expression was achieved.

Figure 1.10. Renilla luciferase signal after PS-DNA transfection and ± UV irradiation (365 nnm, 23 W, 5 min), normalized to the transfection of the non-caged PS-DNA antisense agent. The signal for non-irradiated 3- and 4-caged PS-DNA is comparable to that of the control and non-transfected cells. However, upon irradiation and removal of the caging group, the antisense agents successfully knock-down gene expression, as evidenced by a decrease in luciferase signal similar to that of the non-caged control. From Young, D. D.; Lusic, H.; Lively, M. O.; Yoder, J. A.; Deiters, A. ChemBioChem. 2008, 9, 2937.

In order to achieve spatial control over the activity of the photocaged antisense PS-

DNA, NIH-3T3 cells expressing Renilla luciferase were transfected with or without caged

PS-DNA and the non-caged control. Cells were then irradiated (365 nm, 5 min, 23 W) in a specific location through the use of a mask. Only the irradiated population of cells

transfected with 3- or 4-caged PS-DNA displayed little to no luciferase activity (Figure 1.11)

In contrast, cells containing no caged PS-DNA and expressing luciferase showed no decrease

in gene expression as compared to cells harboring the non-irradiated 3- and 4-caged PS-

DNA. Thus, it was demonstrated that photocaged PS-DNA could effectively provide spatial and temporal control over gene expression, inhibiting RNaseH activity until UV irradiation removed the caging groups and restored active PS-DNA.

Figure 1.11. Spatial regulation of Renilla luciferase expression in NIH-3T3 cells with caged PS-DNA antisense agents. The cellular monolayer was irradiated only within the white dashed circle. A. Negative control with no luciferase expression. B. Cells expressing luciferase and harboring 3-caged PS-DNA. C. Cells expressing luciferase and harboring 4-caged PS-DNA. D. Positive control without PS-DNA. E. Positive control without PS-DNA and no irradiation. F. Cells expressing luciferase and transfected with inactive control PS-DNA. From Young, D. D.; Lusic, H.; Lively, M. O.; Yoder, J. A.; and Deiters, A. ChemBioChem. 2008, 1318, 2937.

In summary, photocaging groups provide a powerful tool to regulate gene function.

Their use affords a robust level of spatial and temporal control that surpasses traditional

biological techniques and is comparable to that found in nature. As such, the research presented herein integrates molecular biology with novel photocaging strategies, enabling

the use of techniques from both fields toward the development of methodologies to control

biological processes with light. To accomplish this, approaches were developed to

photoregulate the activity of proteins and nucleic acids, both in vitro and in vivo.

Specifically, this research has the following objectives:

(1) Light-regulation of the polymerase chain reaction (PCR) using photocaged

oligonucleotides.

(2) Investigation of the effects of microwave irradiation on nucleic acid hybridization.

(3) Light-regulation of enzymatic function by bionconjugating a protein with

photocleavable polyethylene glycol (photo-PEG) groups.

(4) Light-regulation of recombinase enzymes by installing a caging group site-

specifically on the protein to enable photochemical control over DNA recombination.

(5) Light-regulation of protein-dimerization by using a reversibly photoswitchable small

molecule dimerizer.

The resulting techniques facilitate the regulation of gene function, as well as the control of protein dimerization and DNA hybrdization, through the use of light. The developed light-regulatable tools provide powerful additions to the tools used by molecular and cell biologists.

CHAPTER TWO – PHOTOCHEMICAL REGULATION OF THE POLYMERASE CHAIN REACTION

2.1 Introduction to the Polymerase Chain Reaction

The polymerase chain reaction (PCR), developed by Kary Mullis,[84] is employed in the in vitro isolation and exponential amplification of specific DNA sequences.[85-87] The reaction utilizes a thermophilic DNA polymerase and requires two oligonucleotide primers designed to anneal to the DNA template via Watson-Crick base pairing, and concomitantly flank the region of interest. Repeat cycles of varying temperature allow for denaturation, primer annealing, and polymerase-mediated extension. In a short time, extremely small amounts of DNA can be exponentially enriched to substantial quantities (Figure 2.1).

Figure 2.1. The polymerase chain reaction enables rapid amplification of a DNA template.

Due to the sensitivity and fidelity of the DNA polymerase, the amplification product can be isolated without mutations. A variety of polymerases are commercially available with

differing levels of transcriptional fidelity, i.e. error rate. The commonly used Taq DNA

polymerase from Thermus aquaticus has a rate of 2.28×10-5 errors per nucleotide per cycle

(e/n/c),[88] while higher fidelity DNA polymerases such as Vent (NEB) from Thermococcus

litoralis and Phusion (Finnzymes) from Pyrococcus furiosus possess a rate of 3.32×10-6 and

4.40×10-7 e/n/c, respectively.[89, 90] In the years since its inception, PCR has revolutionalized the field of molecular biology, making possible whole genome sequencing and paving the way for genetic disease diagnosis and genetic fingerprinting.[91, 92]

Photochemical regulation over this important reaction could afford several advantages,

including time-dependent amplification of different genes and temporal control during real-

time quantitative PCR experiments. We have thus utilized photocaged oligonucleotide

primers to perform phototriggered DNA amplification through both light-activated and light-

deactivated PCR.

2.2 Hybridization Studies with Photocaged DNA

In the Deiters lab, Dr. Harry Lusic developed the novel NPOM caging group for N-

heterocyclic molecules,[93] which was applied to the specific caging of thymidine[94, 95]

and guanidine[96] nucleotides on their heterocyclic bases. Installation of the sterically-

demanding caging group on thymidine subsequently disrupted A-T base pairing, as

evidenced by the attenuated catalytic activity of a DNAzyme.[20] Brief irradiation with UV

light at 365 nm (25 W, handheld UV lamp, 2 min) removed the caging group and restored

DNAzyme activity (Figure 2.2).

Figure 2.2. Photodeprotection of an NPOM-caged thymidine (the caging group is shown in blue) to yield thymidine.

To implement the caging strategy toward the photoregulation of PCR, we first studied the ability of one or more caging groups to disrupt complementary base pairing. The corresponding caged thymidine phosphoramidite was incorporated into DNA oligomers using standard DNA synthesis equipment and protocols. The DNA oligomers P1-P7, consisting of 19 nucleotides, a typical length for PCR primers, and containing 0-4 caged thymidines were synthesized (Table 2.1). These primers were then analyzed for their annealing and melting properties in the presence of a complementary oligonucleotide.

Table 2.1. Melting temperatures of non-caged (P1) and caged (P2-P7) oligonucleotides before and after irradiation. T* denotes caged thymidine; melting temperatures (Mp) determined with the non-caged complement 5’ CGAACCTGGTCGAAATCAG 3’; ND = not detectable.

DNA Sequence Mp / o C − UV Mp / o C +UV P1 5’ CTGATTTCGACCAGGTTCG 3’ 65.3 ± 0.3 65.0 ± 0.8 P2 5’ CT* GATTTCGACCAGGTTCG 3’ 62.1 ± 0.7 64.1 ± 0.2 P3 5’ CTGATTT* CGACCAGGTTCG 3’ 54.3 ± 1.1 64.1 ± 0.8 P4 5’ CT* GATTT* CGACCAGGTTCG 3’ 55.5 ± 0.5 63.8 ± 0.3 P5 5’ CT* GATTT* CGACCAGGT T* CG 3’ NA 64.2 ± 0.2 P6 5’ CTGA T*T*T* CGACCAGGTTCG 3’ 50.0 ± 1.0 64.5 ± 0.5 P7 5’ CT* GA T* T T* CGACCAGGT T* CG 3’ NA 64.2 ± 0.3

Melting curves were measured on a BioRad MyiQ RT-PCR thermocycler by

conducting a sequence of three heating and cooling cycles (1µM of both primer and

complementary DNA with 12.5 µL iQ SYBR Green Supermix to a total volume of 25µL; 40

oC to 80 oC with a 0.5 oC/min ramp). The melting temperatures were determined to be 65.3

oC (P1), 62.1 oC (P2), 54.3 oC (P3), 55.5 oC (P4), and 50.0 oC (P6). P5 and P7 did not

hybridize and thus, there were no detectable melting temperatures. Our results indicate that

both the number of caging groups and the position of the caged thymidine residues affect

DNA hybridization. Installation of a single caging group results in a melting temperature

depression of 3.2 oC and 11.0 oC as seen in P2 and P3, respectively. This effect is less

pronounced in P2, most likely due to the close proximity of the caged T to the 5’ terminus,

which leads to a lower level of interference with the hybridization of neighboring

nucleotides. Very similar melting point depressions and positional variations have previously

25 been observed in T-mismatches.[97, 98] With the incorporation of additional caging groups in P4 and P6, melting temperatures generally decrease further. However, addition of a single caged thymidine close to the 5’ terminus of P3 had no effect in P4. A positional effect was observed with three caging groups, as seen in P5 and P6. The primer P6 contains three adjacent caged thymidines near its 5’ end; these caging groups disrupt 5’ hybridization, but do not interfere with the hybridization of nucleotides located at the 3’ end, leaving twelve noncaged nucleotides to effectively prime the template DNA. P5, whose melting temperature could not be detected, contains three caging groups distributed throughout the DNA oligomer, leading to a more effective disruption of hybridization. In order to ensure complete removal of the photolabile group, each primer was irradiated for 8 minutes at 365 nm, then analyzed in the same melting temperature assay. Irradiation fully restored DNA hybridization, as each primer displayed a comparable melting temperature to the non-caged analog P1. In summary, an NPOM caging group positioned every six to nine bases and evenly distributed throughout the oligonucleotide efficiently disrupts DNA:DNA hybridization. Identical observations were made for DNA:RNA hybridization.[77]

2.3 Light-Activation of the Polymerase Chain Reaction

Our experiments revealed that the presence of three caging groups distributed throughout a 19 nucleotide oligomer is sufficient to disrupt hybridization and to prevent annealing of a PCR primer to its cognate DNA template at the typical annealing temperature range of 50-65 oC.[98] As the stability of the caging group to PCR conditions was examined

with the monomeric caged thymidine and was found to be unaffected by the elevated

temperatures required for PCR, P5 was utilized in a light-activated PCR experiment (Figure

2.3). All reactions described herein employed the same reverse primer (5’

AGAGAGCTCGAGATCGCCATCTTCCAGCAGGCGCACCATTGCCCCTGT 3’).

Figure 2.3. PCR activation by light. Black line: template; single blue line: primer; double blue line: product; red circle: caged thymidine.

A control PCR reaction with non-caged and irradiated P1 produced approximately

140 ng of the expected ~0.6 kb PCR product (Figure 2.4, lane 5); however when caged P5

was employed in the absence of UV irradiation, only trace amounts of product were detected

(Figure 2.4, lane 6). After irradiation, the function of P5 was restored (Figure 2.4, lane 7),

leading to comparable amounts of PCR product, as found in the reaction with P1 (Figure 2.4,

lane 5). As such, this experiment represents the first example of a light-activated PCR.

Figure 2.4. Agarose gel of dsDNA amplified by light-regulated PCR. UV+ indicates reactions that were irradiated, while UV- denotes no irradiation; L = DNA ladder; lanes 1-4 show the results for PCRs performed with truncated wild type (P10), wild type (P8) or caged (P9) hairpin primers; lanes 5-7 show the results for PCRs performed with wild type (P1) or caged (P5) linear primers; lanes 8-10 show the simultaneous activation and deactivation of PCR with light using primers P5 and P9.

While we initially employed irradiation prior to the thermal cycling, temporal control

could be achieved by initiating the reaction through irradiation at a specific timepoint. Here,

an identical reaction with UV irradiation at cycle 15 was conducted, as shown in Figure 2.5.

Prior to irradiation, no PCR product can be detected in reactions conducted with the caged

primer P5. In contrast, the non-caged primer P1 leads to the expected exponential

amplification. Upon UV irradiation of the reaction with P5 at cycle 15, the amount of DNA

increases exponentially, while no amplification occurs in the corresponding non-irradiated

reaction.

Figure 2.5. Time-course of a PCR, light-activated at cycle 15. Error bars were obtained from three independent experiments.

2.4 Light-Deactivation of the Polymerase Chain Reaction

After achieving photochemical control over activation of PCR at a specific time

point, we examined the possibility to switch off PCR activity via light irradiation. To

accomplish this, a self-complementary primer was designed that is predisposed to form a

hairpin rather than act as a PCR primer. By installing caging groups on the complementary

portion of this sequence, it is possible to block self-hybridization, thus enabling the

polymerization reaction. The PCR is then stopped by removing the caging groups, leading to

hairpin formation and primer deactivation (Figure 2.6).

Figure 2.6. Light-deactivation of PCR. Black line: template; single green line: primer; double green line: product red circle: caged thymidine.

In order to achieve photochemical deactivation, an appropriate hairpin primer P8 was

designed (5’ GGTCAGTAAATTGTTTTTCAATTTACTGACCG 3’), and a photocaged

analog P9 was synthesized (5’ GGT*CAGT*AAAT*TGTTTTTCAATTTACTGACCG 3’.

The same reverse primer as in the light-activation reactions was again utilized in the light- deactivation experiments. PCR using the truncated primer P10 (5’ CAATTTACTGACCG

3’), which possesses only half of the hairpin and not its complement, led to DNA

amplification after 25 cycles (Figure 2.6 and Figure 2.4, lane 1). Conversely, the non-caged

hairpin primer P8 failed to amplify the DNA, leading to very little PCR product (Figure 2.4,

lane 2). However, in the absence of light irradiation the caged hairpin primer P9 successfully

acted as a primer, yielding 137 ng of PCR product (Figure 2.4, lane 3). This suggests that the

three installed caging groups prevented hairpin formation, and allowed the complementary

sequence to act as a PCR primer. If P9 was irradiated for 8 minutes at 365 nm, the caging

groups were removed, leading to hairpin formation and suppression of DNA amplification

(Figure 2.4, lane 4).

These light-regulatory mechanisms could also be employed in a temporally controlled fashion by irradiating the caged primer after 10 cycles of PCR (Figure 2.7). The non-caged primer P8 was used as a control, as it formed a hairpin immediately and blocked Taq DNA polymerase and any subsequent amplification. Because the caged primer P9 was able to bind to the DNA template, amplification occurred. The reaction containing P9 was irradiated at cycle 10, leading to removal of the caging groups, hairpin formation, and effective inhibition of DNA amplification. At the same point, amplification continued in the non-irradiated reaction with P9.

Figure 2.7. Time-course of a PCR light-deactivated at cycle 10. Error bars were obtained from three independent experiments.

The photochemical deactivation of DNA hybridization via hairpin formation has

recently been applied to antisense agents in mammalian cell culture, and enabled the light- activation of gene function.[20]

2.5 Concurrent Up- and Down-Regulation of PCR

With the two caged primers P5 and P9 possessing opposing effects on the PCR reaction upon light irradiation, it was possible to use both primers simultaneously to stop the production of one PCR product, while also triggering the amplification of a different PCR product via irradiation with UV light (Figure 2.8). Thus, P5, P9, and the reverse primer were included in the PCR reaction mixture. A product band of ~1.0 kb was detected in the non- irradiated reaction after 20 cycles, which is attributed to the active, caged and non-hairpin forming P9, and the inhibited, caged P5 (Figure 2.4, lane 9). Alternatively, a product band of

~0.6 kb was observed in the irradiated reaction, as a result of the decaging of P5, and the decaging and subsequent hairpin formation of P9 (Figure 2.4, lane 10). As expected, both bands were observed in the control reaction using non-caged and non-hairpin primers (Figure

2.4, lane 8).

Figure 2.8. Light-switching between two different PCRs through simultaneous activation and deactivation of primers.

2.6 Summary and Outlook

In summary, a photochemical activation and deactivation of the polymerase chain reaction has been developed. This was accomplished through the incorporation of multiple caged thymidine phosphoramidites into oligonucleotide primers using standard DNA synthesis protocols. By effectively disrupting DNA hybridization with caging groups and then restoring it with light irradiation, it was possible to control activation and deactivation of

PCR in a temporal fashion. Moreover, by conducting a simultaneous activation and deactivation, light switching from one DNA amplification product to another was achieved.

Photocaged primers thus provide an alternative strategy to hot start PCR techniques

toward the reduction of non-specific amplification.[99-101][99, 102] Photocaged primers

could potentially be employed in overlap PCR,[103] where the first round of PCR could be

completed with noncaged primers and the second round could be performed with caged

primers after brief UV irradiation, eliminating the need to transfer product from one reaction

to another. Additionally, Dr. Douglas Young (Deiters Lab) demonstrated that the photocage

itself can be used to block DNA extension by the polymerase.[104] In this manner, “sticky

ends” may be formed in the final PCR product that, after irradiation at the conclusion of the

reaction, can then be ligated into a digested vector without the use of restriction enzymes to

cut the insert. This is especially attractive for smaller PCR products that would otherwise be

difficult to cut by traditional restriction digest.

2.7 Experimental Methods

PCR Conditions. Taq DNA polymerase, Taq reaction buffer and nucleotide triphosphates

(dNTPs) were purchased from New England Biolabs (NEB). pET-21Cre (see Chapter 4.2)

served as the DNA template in all reactions. 75 ng of template was incubated in the presence

of P5 and a reverse primer

(5’AGAGAGCTCGAGATCGCCATCTTCCAGCAGGCGCACCATTGCCCCTGT 3’, 1

μM each; the same reverse primer was used in all PCRs) with dNTPs (0.3 mM each), Taq

Reaction Buffer (10 mM Tris-HCl, 50 mM KCl, 1.5 mM MgCl2, pH 8.3) and water in a total

volume of 50 µL. Taq DNA Polymerase (0.3 μL) was then added to initiate the reaction. An initial denaturation at 95 °C was performed, followed by 40 cycles consisting of 95 °C (30 sec), 50 °C (30 sec), and 72 °C (1 min), with a final extension at 72 °C (2 min). The reactions were conducted in triplicate, then run on a 1% agarose gel. The gel was stained with ethidium

bromide (10 µL of 1 mg/100 ml ethidium bromide in water), visualized on a transilluminator

(254 nm), and quantitated by band integration of the ethidium bromide-stained agarose gels using Image Quant 5.2. Additional experiments conducted with photocaged primers may require optimization of annealing temperature, as demonstrated with the photocaged primers synthesized by TriLink Biotechnologies. With these primers, an increase in annealing temperature from 50 °C to 55 °C was necessary to prevent non-specific amplification.

Light Irradiation of Caged Primers. Prior to placement in a thermocycler, selected reactions were placed in a 200 µL PCR tube and either irradiated through the PCR tube on a

34 transilluminator (8 min, 365 nm, 25 W) or maintained in the dark by wrapping the PCR tube in aluminum foil.

CHAPTER THREE – THE EFFECTS OF MICROWAVE IRRADIATION ON DEOXYOLIGONUCLEOTIDE HYBRIDIZATION

3.1 Introduction to DNA Hybridization and Microwave Irradiation

Despite its biological importance, the effects of microwave irradiation[105-111]

on DNA structure, processing, and replication have been virtually unexplored. Of the few

studies conducted under microwave irradiation, two investigated the cellular effects of

irradiation leading to sterilization,[112, 113] and were performed in traditional household

microwave ovens. Two investigations were conducted on the genetic damage caused by

microwave irradiation.[114-116] Due to a general interest in the Deiters lab concerning

the effects of microwave irradiation on chemical transformations, the effects of

microwave irradiation on the hybridization of DNA oligomers were investigated.

Hybridization is a key property of DNA, and the ability to modulate hybridization with

microwave irradiation provides several applications, including studies involving DNA

hybridization probes, interactions with proteins, and experiments utilizing hybridization

for materials science purposes. Additionally, the susceptibility of DNA to microwave

irradiation due to its large dipole moment (approximately 1000 D)[117], as compared to that of the lambda repressor protein (60 D)[118], hen egg white lysozyme (50 D)[119],

and water (2.5 D),[120] has implications in microwave-mediated oligonucleotide

synthesis. This is similar to the effects observed in the synthesis of peptides,[121, 122] as

well as enzymatic reactions involving the processing of DNA, e.g. in a microwave-

assisted PCR.[123, 124]

3.2 Development of a Fluorescence Hybridization Assay

To investigate the effects of microwave irradiation, a viable assay to detect DNA hybridization was first established. While many real-time protocols[125-127] exist for the detection of double-stranded DNA (e.g. real-time PCR using intercalating fluorescent probes), these methods are not applicable to the closed environment of a microwave reactor.

Thus, an offline assay was required to determine if the DNA was melting or annealing while being subjected to microwave irradiation. In order to achieve this, an end-point assay was designed that relies upon fluorescence quenching to detect the melting and annealing of DNA in a microwave synthesizer (Figure 3.1). This assay uses two sets of double-stranded DNA.

One of the pairs possesses a 5’ fluorescein modification which hybridizes in close proximity to a 3’ dabcyl modification, and the other pair consists of the identical sequences with no modifications. Due to the proximity of the fluorophore and quencher on the hybridized modified oligomer pair, very little background fluorescence should initially be present.[128]

Upon microwave irradiation of the system and melting of the dsDNA, the ssDNA will subsequently anneal in a statistical fashion to either its previous partner or to the non- modified complement. If the fluorescein-modified strand anneals to its non-modified complement, the quencher is removed, leading to an increase in the fluorescence signal. A similar result is observed if the DNA melts, but does not re-hybridize. If microwave irradiation is not capable of inducing DNA melting, then no change in fluorescence should be observed.

heat or MW + + + + hν

cool + + + hν

fluorescence

Figure 3.1. Schematic of the fluorescence quenching assay used to determine if microwave irradiation (MW) is capable of melting dsDNA. Open circle = fluorophore (fluorescein); closed circle = fluorescence quencher (dabcyl).

In order to probe the viability of the assay, we designed dsDNA with a calculated

Tm >70 °C, and obtained the complementary fluorophore- and quencher-modified (D1F*

& D1R*) and unmodified oligomers (D1F & D1R)(Alpha DNA Technologies)(Table

3.1).

Table 3.1. Sequences and melting temperature of DNA oligomers. F* contains a fluorescein label at the 5’ end; R* contains a dabcyl label at the 3’ end.. Calculated Tm values were determined using the Oligo Analyzer 3.1 (IDT DNA). Actual (fluorescein-labeled and dabcyl-labeled DNA) and Non-labeled DNA Tm values were determined in triplicate on a BioRad MyiQ RT-PCR thermocycler.

DNA DNA Sequence Calculated Tm / oC Actual Tm / oC Non-labeled Tm / °C D1F* 5’ CGCACCCAGGCTTAGCTACAAACAT 3’ 61.4 81.5 ± 0.4 80.9 ± 0.3 D1R* 5’ GCGTGGGTCCGAATCGATGTTTGTA 3’ D2F* 5’ CCCAGGCTTAGCTACA 3’ 50.6 66.2 ± 0.3 66.0 ± 0.4 D2R* 5’ GGGTCCGAATCGATGT 3’ D3F* 5’ AGGCTTAGCTACA 3’ 39.9 51.5 ± 0.3 51.9 ± 0.3 D3R* 5’ TCCGAATCGATGT 3’

We first verified that the non-labeled and labeled oligo pairs had similar melting

points to ensure that there was no measurable effect of the dabcyl and fluorescein dyes on

the behavior of the hybridized pairs. The melting temperatures of all oligonucleotide

pairs (D1F*:D1R*, D2F*:D2R*, and D3F*:D3R*) were determined on a BioRad MyiQ

RT-PCR thermocycler by conducting a sequence of three heating and cooling cycles,

while measuring the increase of fluorescence due to the loss of the quencher in the

melting process. As expected, melting temperatures for the non-labeled oligos were found

to be identical, within the error margin of the experiment, to those obtained using the

fluorescently modified oligomers (Table 3.1). It is unclear why there is a large

discrepancy between the calculated and actual Tm values.

Initial hybridizations were conducted thermally (Eppendorf Mastercycler, 95 oC (5

min), 37 oC (30 min), 4 oC (30 min)) to generate the two sets of double stranded DNA. A

1:1 ratio of dabcyl-modified oligomer to fluorescein-modified oligomer was employed to sufficiently afford hybridization and effectively suppress the fluorescence signal in

D1F*:D1R* (Figure 3.2, column 1). A positive control experiment was conducted by hybridizing D1F* and D1R to determine the maximum attainable fluorescence signal

(Figure 3.2, column 2). Additionally, a thermal comparison control experiment was

performed in which we employed a ten-fold excess of the non-labeled dsDNA D1F:D1R

to the labeled dsDNA D1F*:D1R*. This ensured that an adequate amount of non-labeled

DNA was present for hybridization when D1F*:D1R* melted. The resulting fluorescence

signal (Figure 3.2, column 3) is lower than that of the positive control experiment

(D1F*:D1R), as upon melting and subsequent hybridization, the labeled DNA can

reanneal to its labeled complement forming D1F*:D1R* and thus quench fluorescence

(Figure 3.1).

3.3 Microwave Irradiation of DNA

To differentiate between microwave effects and thermal effects on the melting of

dsDNA, experiments were conducted in a CEM Discover microwave synthesizer (Figure

3.2) equipped with a jacketed reaction vessel enabling temperature control through the continuous flow of coolant while precisely measuring the temperature using a fiber optics probe (CEM Coolmate).

A. B.

Figure 3.2. The CEM Discover microwave synthesizer. A. In the CEM Discover single-mode microwave cavity, the sample is located in the center of the cavity and focused microwaves are generated and used to irradiate the sample. B. The jacketed reaction vessel allows robust cooling of the sample during microwave irradiation through the continuous circulation of cryogenic coolant around the sample compartment. Images obtained from www.cem.com.

By attempting to melt DNA at temperatures well below the calculated melting

point (e.g. –20 to 20 °C), we could demonstrate that the melting was a direct effect of the microwave irradiation, and not due to a thermal influence that could convolute the results. Thus, the pre-hybridized oligomers were mixed and subjected to various conditions to probe hybridization. The mixture of dsDNAs D1F*:D1R* and D1F:D1R

was either heated then cooled to hybridize the DNA strands under typical conditions

found in a hybridization protocol[129] ((95 oC (5 min), 37 oC (30 min), 4 oC (30 min)),

subjected to purely thermal conditions mimicking the microwave temperature profile (–

20 °C to 20 °C to –20 °C, 0 W), or microwave-irradiated (–20 °C to 20 °C to –20 °C) at

300 W while coolant at –60 °C was simultaneously circulated through the jacketed

reaction vessel. A temperature profile starting at –20 °C was necessary due to the non

microwave-transparent hybridization buffer which would otherwise reach high

temperatures in only a few seconds at a high power of 300 W. The reaction could not be

kept at –20 °C under microwave irradiation and was stopped once 20 °C (measured using

a fiberoptics sensor) was reached (typically 30 sec to 3 min). All reactions were

subsequently equillibrated at 4 °C to ensure hybridization. (Figure 3.3).

1400

1200

1000

800

600

400 relative fluorescence units

200

0 DNA: D1F*:D1R* D1F*:D1R D1F*:D1R* D1F*:D1R* D1F*:D1R* D1F:D1R D1F:D1R D1F:D1R Temp.: A A A B B MW: - - - - 300 W

Figure 3.3. Fluorescence signals arising from various conditions employed in the DNA melting experiments with D1F and D1R. All assays were conducted in triplicate and error bars represent standard deviations. Temperature profiles: A) 95 °C to 4 °C; B) –20 °C to 20 °C to –20 °C. D1F* = 5’ fluorescein-labeled D1F; D1R* = 3’ dabcyl-labeled D1R. Initially the D1F*/D1R* dsDNA was added, followed by the addition of non- labelled D1F/D1R to monitor the changes in hybridization under the various experimental conditions.

Additionally, power, pressure and temperature profiles were obtained from each reaction to verify that all hybridization reactions occurred under identical conditions.

Representative graphs are shown in Figure 3.4.

A 30 25 20 15 10 5 0 -5 0 20 40 60 80 100 120 140 -10 -15 -20 -25

B 80

0 0 10 20 30 40 50 -20

-40

C 120 100 80 60 40 20 0 -20 0 5 10 15 20 25 30 -40 Time/s

Figure 3.4. Representative power (green square), pressure (pink triangle), and temperature (blue diamond) profile graphs show typical reaction conditions at various microwave powers. While the pressure remained constant in all reactions, less time was required to raise the temperature of the reaction as the input power increased. A. At 25 W, the temperature of the reaction increased from –20 °C to 20 °C over approximately 3 min. B. At 75 W, the temperature of the reaction increased from –20 °C to 20 °C in about 1 min. C. At 100 W, less than 30 sec were required to raise the temperature of the reaction from –20 °C to 20 °C.

In order to ensure that no single-stranded DNA breaks occurred during microwave

irradiation, aliquots of the reaction mixture containing D2F*, D2R*, D2F, and D2R were run on a 13% denaturing PAGE gel. As expected, only one band was present for each experiment; no streaking or smears were evident, indicating that the DNA remained intact during irradiation at 5-200 W (Figure 3.5). We were unable to determine, however, whether the DNA sustained damage to individual bases.

M 1 2 3 4 5 6 7

Figure 3.5. Microwaved DNA sustains no damage as detected with denaturing PAGE. Aliquots of D2F*:D2R*:D2F:D2R hybridizations were subjected to microwave irradiation and analyzed by PAGE through ethidium bromide staining. Only a single band is visible in each lane. M: marker (hybridized D2F*:D2R* with no irradiation); 1: 0 W; 2: 5 W; 3: 75 W; 4: 112 W; 5: 115 W; 6: 175 W: 7: 200 W.

As expected, the thermal heating from –20 °C to 30 °C without microwave

irradiation (Figure 3.3, column 4) did not result in any increase of fluorescence,

delivering an identical readout as the initially hybridized D1F*:D1R* pair (Figure 3.3,

column 1). However, heating the dsDNA mixture from –20 °C to 20 °C using microwave

irradiation of 300 W (Figure 3.3, column 5) yielded a fluorescence signal identical with

the positive control under thermal conditons (95 °C, Figure 3.3, column 3). This indicates that microwave irradiation is capable of inducing complete DNA melting at temperatures well below the 81.5 °C melting point of the D1F:D1R DNA duplex.

Based on the apparent influence of microwave irradiation on DNA hybridization,

this phenomenon was further probed by employing different DNA/DNA duplexes based

on the same sequence, but shortened to afford lower melting temperatures of 66.2 °C

(D2F:D2R) and 51.5 °C (D3F:D3R) (Table 1). All oligomer pairs were subsequently used in the described melting assay following the identical temperature profile as before

(–20 °C to 20 °C to –20 °C), but varying microwave power (0, 25, 50, 75, 100, 125, 150,

175, and 200 W) in order to generate microwave melting curves in an analogous fashion to thermal melting curves (Figure 3.6). At 300 W, the reaction is over very rapidly (less than ~5 sec) and as such, the power was not increased above 200 W in the following experiments. The reaction times decreased with increasing power from 3 min to 30 sec.

3500

3000

2500

2000

relative fluorescence units 1500 D1 D2 D3 1000

0 50 100 150 200

power / W Figure 3.6. Microwave melting curves indicate that microwave irradiation induces DNA melting at a specific power regardless of the duplex length. Oligomers were combined and subjected to a variety of microwave powers under similar thermal profiles (–20 °C to 20 °C to –20 °C). All assays were conducted in triplicate and error bars represent standard deviations.

Interestingly, all of the DNA sequences have similar inflection points at 111 W

(D1), 117 W (D2), and 100 W (D3), as determined by differentiation of the sigmoidal curves shown in Fig. 3. Although each oligo pair possesses a different thermal melting temperature, they all melt at virtually the same microwave power (109 ± 8 W). This result was unexpected and indicates that the microwave irradiation specifically interacts with the DNA. The energy transfer into the oligonucleotide increases with the number of nucleotides, thus maintaining the melting temperature at the same microwave power despite the increased degree of hybridization in longer dsDNA sequences. We speculate that the microwave irradiation is capable of disrupting ions associated with the

DNA,[130] while simultaneously interacting with the DNA leading to hydrogen bond disruption and DNA melting. Investigations into the dielectric permittivity of DNA have shown that DNA dispersion and melting can be correlated with counterion fluctuations on short sections of the double helix.[131, 132] The microwave irradiation could disrupt cation binding to the negatively charged DNA, as cations and anions align with the electromagnetic field, thus moving them in opposite directions.[109].

Thus, at a specific microwave power the cations become completely dissociated from the anionic phosphodiester backbone, leading to the repulsion of the negative charges on the two DNA strands. This may destablize the duplex and facilitate DNA melting to minimize the negative charge repulsion.[133, 134] Finally, it is known that the dielectric permittivity of single stranded DNA is greater than that of dsDNA.[135]

Consequentially, when the double-stranded DNA begins to unravel, forming single strands, a greater coupling with the electromagnetic field occurs, providing increasing energy transfer to overcome the hydrogen bonding interactions of the remaining dsDNA.

The observed effects may be one or a combination of these possibilities. This is in contrast to thermal heating, where the bulk solvent initially absorbs the energy, delivering a thermal melting point that is dependent on the number of base pairs providing hydrogen bonding interactions.[136]

3.4 Summary and Outlook

In conclusion, we have demonstrated a new mechanism for disrupting DNA hybridization at temperatures much lower than necessary under purely thermal heating by employing microwave irradiation. Interestingly, the microwave irradiation yields a

different hybridization disruption than thermal heating, as complementary DNA

oligomers with varying lengths all melted at identical microwave powers. This finding

suggests a specific interaction of microwave irradiation with deoxyoligonucleotides.

Microwave systems enabling online analyses of DNA melting will be required to

elucidate the true mechanism. It is now clear that microwave irradiation contributes to the melting of dsDNA at a specific power (109 ± 8 W), regardless of duplex Tm. As dsDNA

absorbs less light than ssDNA, coupling of a UV spectrophotometer to the microwave

could allow determination of the precise point at which the dsDNA begins to melt based

on UV absorption. Time-resolved studies could be performed to further investigate the microwave effect. If microwave irradiation causes cation dissociation and subsequent melting at specific locations within the duplex, the DNA would instantaneously re-

hybridize once the irradiation ceases. However, if the hybridized DNA directly interacts

with the electromagnetic field to induce rapid and complete melting of the duplex, then

the strands will only slowly reanneal after the irradiation has been turned off. These studies could provide insight into the mechanism for microwave-induced DNA melting,

and subsequent discoveries have important implications in the future development of

microwave-assisted DNA technologies, particularly in the area of high-throughput

screening and genome analysis.

3.5 Experimental Methods

General. All hybridization experiments were perfomed in a CEM Discover microwave

reactor equipped with a Coolmate system, which circulated chilled (–60 oC), microwave transparent fluid through a jacket reaction vessel to allow the maintenance of low temperatures during microwave irradiaton. The CEM Discover system is a single-mode

cavity allowing for focused delivery of microwave irradiation (Figure 4.4). All reactions

were conducted at a constant microwave frequency of 2450 MHz.[137]

Initial Hybridizations. Dabcyl- and fluorescein-modified DNA oligomers were purchased

from Alpha DNA and rsuspended in distilled water to a stock concentration of 200 μM.

Initial hybridization reactions contained 7 μM of each labeled oligomer in 20 μL of 10X

hybridization buffer (100 mM Tris-HCl, pH 8.3, 500 mM KCl, 15 mM MgCl2) and distilled

water to a final volume of 200 μL. Reactions were heated in an Eppendorf Mastercycler to 95

°C for 5 minutes, 37 °C for 30 minutes, and then cooled to 4 °C to ensure complete

hybridization. The resulting fluorescence of each reaction was then measured using a

Molecular Devices Gemini EM microplate spectrofluorimeter (λex = 494 nm/λem = 521 nm).

Microwave Hybridizations. Non-labeled complementary DNA oligomers were purchased from IDT DNA and suspended in distilled water to a stock concentration of 200 μM. 40 μL of the initial hybridization reactions were mixed with 7 μM of each complement oligomer in

50 μL of 10X hybridization buffer and distilled water to a final volume of 500 μL. Each reaction was transferred to a microwave reaction vial containing a stir bar. Samples were cooled to –20 °C, irradiated at the appropriate power until the temperature of the reaction reached 20 °C, then cooled to 0 °C, with typical reaction times from 30 sec to 5 min.

Thermal Hybridizations. Following initial hybridizations, thermal hybridization reactions were set up with non-labeled DNA complements as described for microwave hybridizations, then returned to the Eppendorf Mastercycler and heated to 95 °C for 5 minutes, 37 °C for 30 minutes, and then cooled to 4 °C to ensure complete hybridization.

Control Hybridizations. Control hybridizations were performed to mimic the temperature profile of the microwave experiment. Initial hybridizations were performed, and then complement oligomers were added as described above. Measuring the temperature with a thermocouple probe, reactions were chilled in a dry ice/ethanol bath to 20 °C, then placed in warm water until 20 °C was reached (30 sec), and were then returned to the dry ice/ethanol bath until 4 °C was reached.

Determining Melting Temperatures. Actual Tms were measured for the non-labeled, and

the fluorescein-labeled:dabcyl-labeled oligo pairs using a BioRad MyiQ RT-PCR

thermocycler and performing three heating and cooling cycles (a total volume of 25 µL; 30

°C to 80 °C with a 0.5 °C/min ramp). All samples were measured in triplicate and averaged to determine the Tm.

CHAPTER FOUR – LIGHT-REGULATION OF PROTEIN FUNCTION THROUGH PHOTOCAGING

4.1 PhotoPEG, a New Reagent for the Photochemical Control of Protein Function

4.1.1 Lysozyme as a Model Protein

Hen egg white lysozyme is a 14.7 kDa protein that catalyzes the hydrolysis of 1,4-β- linkages within peptidoglycan and chitodextrin, thus destroying the cell walls of gram positive bacteria. Specifically, lysozyme is a member of the family of N-acetylmuramide glycanhydrolases that hydrolyse the glycosidic bond between N-acetylmuramic acid and N- acetylglucosamine (Figure 4.1).[138] Lysozyme is present in mucus, tears and saliva, and thus plays a key role in the prevention of bacterial infection.[139, 140] For this reason, lysozyme has been extensively studied as a classic model of protein structure-function relationships,[141-144] its crystal structure has been solved,[144] and assays to assess its enzymatic activity have been developed.[138, 145, 146]

Figure 4.1. Lysozyme-catalyzed hydrolysis of N-acetylmuramic acid (NAM) and N-acetylglucosamine (NAG) the components of gram-positive bacterial cell walls. Adapted from Kuroki, R.; Weaver, L. H.; and Matthews, B. W. Proc. Natl. Acad. Sci. 1999, 96, 8949.

We sought to modulate lysozyme activity through the bioconjugation of light- removable PEG groups that inhibit lysis of a bacterial substrate until UV irradiation removes

the PEG groups and restores enzyme function. In addition to its N-terminus, lysozyme

contains six solvent-accessible lysine residues (Figure 4.2, A) whose amine groups readily

serve as reactive sites.[145]

A. B.

365 nm

Figure 4.2. Structure and photochemical dePEGylation of lysozyme. A) Lysozyme contains six solvent- accessible lysine residues (yellow) and an N-terminus that are readily PEGylated. B) PEGylation completely inhibits enzymatic activity until the PEG units are removed with light irradiation.

4.1.2 PhotoPEGylation of Lysozyme

PEG groups have been used extensively in biological and pharmaceutical applications

to improve the pharmacokinetic properties of biomolecules.[145, 147-149] PEG-

bioconjugates provide enhanced in vivo stability and are nontoxic and non-

immunogenic.[150-152] They are typically generated from proteins through a process termed

“PEGylation”, which involves the surface installation of PEG groups through a covalent

reaction with amino functionalities, e.g. the ε-NH2 of lysine. PEGylation greatly enhances the

solubility, half-life, cellular uptake, and bioavailability of proteins in cells and

organisms.[147, 153, 154] A number of PEGylated therapeutics have been approved by the

FDA and are currently on the market,[155-157] demonstrating the growing interest in

PEGylation chemistry toward drug delivery and protein stabilization.

However, the PEGylation of proteins can negatively impact their biological activity

and lead to complete inhibition of protein function, e.g., through steric blocking of the active

site of an enzyme.[158] The bulk of the PEG polymer wraps around the protein, thus providing the desired protective effects by sterically blocking it without perturbing the

conformation and dynamics of the protein native fold.[159, 160] For this reason, PEGs

containing an ester linkage for tethering to the biomolecule have been developed that are

susceptible to spontaneous hydrolysis and provide a slow release pathway for protein

delivery.[158, 161] While hydrolyzable linkers facilitate general release of the active

therapeutic over time, a means to obtain precise spatial and temporal control over bioavailability without compromising stability would be advantageous.

We reasoned that the installation of a photolabile PEG directly onto a protein would provide a “PEG photocage” that can inhibit protein activity (Figure 4.2, B). Upon irradiation

with non-phototoxic UV light, the PEG photocage will be removed from the protein, thus

restoring its native activity. Since UV irradiation can be precisely controlled with high spatial

and temporal resolution, this methodology provides a fundamentally new approach to

achieve spatio-temporal control over protein activity. We thus utilized the first

photocleavable PEG reagent (Photo-PEG) in the photochemical control of a Photo-

PEGylated lysozyme.

We designed a lysozyme-photocleavable linker-PEG conjugate (Figure 4.3) that is

stable to hydrolysis and, like previously used PEGylation reagents,[147, 154] wholly

abrogates lysozyme activity. However, unlike these reagents, activity is restored with light, as brief UV irradiation at 365 nm cleaves the linker and releases free, active lysozyme. The photolabile linker is covalently attached to PEG (MW 5000 Da) and is comprised of an

ortho-nitrobenzyl core that confers light sensitivity (see Chapter 1, Figure 1.2). The Photo-

PEG reagent also bears a terminal N-hydroxysuccinimide (NHS) ester that makes the PEG- linker moiety susceptible to nucleophilic attack by a primary amine. Lysozyme was thus reacted with the novel Photo-PEG that was synthesized by Dr. Harry Lusic (Deiters Lab).

Figure 4.3. A. The PhotoPEG reagent; B. Structure of the lysine-Photo-PEG conjugate on lysozyme. Only one PEGylated site is shown.

Hen egg white lysozyme (EMD) was first incubated with increasing equivalents of a

non-light cleavable N-hydroxysuccinimide NHS-PEG5000 (Jenkem Technology) overnight at

4 °C in phosphate buffered saline (PBS), pH 7.5. One equivalent of NHS-PEG (Figure 4.4, lane 2) produced a singly PEGylated protein; however, a substantial amount of non-

PEGylated wild-type enzyme remained.

M 1 2 3 4 5 6 7 8 175 80 58 46

30 25

Figure 4.4. SDS-PAGE analysis of lysozyme PEGylation. M: protein maker (in kDa); lane 1: wild-type lysozyme; lanes 2-6: PEGylation with 1, 5, 10, 25, and 50 equiv. of NHS-PEG, respectively; lane 7: PEGylation with 50 equiv. Photo-PEG; lane 8: Photo-PEG modified lysozyme from lane 7 after UV exposure (365 nm, 25 W, 30 min).

When five equiv. were added (Figure 4.4, lane 3), a small amount of non-PEGylated lysozyme was detected. While increasing the amount of (NHS)-PEG from 10 to 25 equiv. led to a total consumption of wild-type lysozyme (Figure 4.4, lanes 4-5), multiple PEGylated protein species were produced which still displayed some enzymatic activity (Figure 4.5).

PEGylation with 50 equiv. of NHS-PEG (Figure 4.4, lane 6) completely abolished lysozyme

activity (Figure 4.5). Thus, we employed 50 equiv. of Photo-PEG in the PEGylation of

lysozyme (lane 7), followed by dialysis into PBS (pH 7.5) using a dialysis membrane with a

cutoff of 50 kDa in order to remove excess PEG reagent. Irradiation of the Photo-PEG modified enzyme with UV light of 365 nm (transilluminator, 25 W, 30 min,) releases all PEG groups, regenerating wild type lysozyme as determined by SDS-PAGE analysis (Figure 4.4, lane 8).

0.7

0.6

0.5

0.4 no lysozyme

450 5eq NHS-PEG 0.3

OD 10eq NHS-PEG 25eq NHS-PEG 0.2 50eq NHS-PEG

0.1

0 0:00 4:48 9:36 14:24 19:12

Time

Figure 4.5. (NHS)-PEGylation of lysozyme results in partial (5 to 25 eqiv) to complete (50 equiv) attenuation of enzyme activity, as measured by a decrease in optical density at 450 nm.

4.1.3 In Vitro Activity Assays of Lysozyme Function

Lysozyme assays were performed as previously reported.[146] The amount of

purified PEGylated lysozyme was determined by a standard Bradford assay. The lyophilized

gram-positive substrate Micrococcus lysodeikticus (Sigma) was suspended in 66 mM sodium

phosphate, pH 6.24. To achieve a starting optical density (OD450) of approximately 1.0,

lysozyme or PEGylated lysozyme in 100 µL PBS was added to 100 µL of cell suspension for

a final enzyme concentration of 3.0 µM. The OD450 was measured after a 20 min reaction

time. Active wild-type lysozyme induces cell lysis, leading to a reduction in optical density

(Figure 4.6). In contrast, the lysozyme PEGylated with the classical NHS-PEG reagent (from

Figure 4.4, lane 6), and the lysozyme modified with the PhotoPEG (from Figure 4.4, lane 7)

were completely inactive, since the same OD450 was observed as in the absence of any lysozyme (Figure 4.6).

no lysozyme

wt lysozyme

NHS-PEG lysozyme

Photo-PEG lysozyme

0.0 0.2 0.4 0.6 0.8 1.0 1.2

OD450

Figure 4.6. Micrococcus lysodeikticus exposed to different lysozymes for 20 min, followed by an optical density (OD450) measurement. Wild-type lysozyme leads to complete cell lysis. In contrast, lysozymes PEGylated with the NHS-PEG or Photo-PEG are completely inactive, thus inducing no cell lysis. Error bars represent standard deviations from three independent experiments.

We next investigated the photochemical activation of enzyme function through a light

induced removal of the PEG groups. The completely inactive Photo-PEG modified lysozyme

was irradiated for 0-30 min, and enzymatic activity was determined as described above. As expected, in the absence of UV irradiation (0 min) no enzymatic activity was observed resulting in an OD450 of 1.1 after 15 min, identical to the OD450 at the beginning of the assay.

However, UV irradiation for 5, 10, 15, and 20 min led to increasing enzymatic activity

resulting in the reduction of the optical density of the cell suspension due to cell lysis.

Optimal light-activation of the enzyme was observed after irradiation for 25 and 30 min,

since the measured OD450s were identical to the application of wild-type lysozyme (Figure

4.7). This observation is in agreement with the gel-based assessment of complete photochemical PEG removal after 30 min of UV irradiation (Figure 4.4, lane 8).

1.0

0.8

0.6 450 OD 0.4

0.2

0.0 0 5 10 15 20 25 30 wt light irradiation / min

Figure 4.7. Micrococcus lysodeikticus exposed to Photo-PEGylated lysozyme followed by an optical density (OD450) measurement after 15 min. The Photo-PEGylated lysozyme is completely inactive before irradiation (0 min); however, complete activity could be restored through UV irradiation (365 nm; 25 W; 5-30 min).

Lastly, the percent activity restoration of irradiated PhotoPEGylated lysozyme was

investigated. Lysis assays were performed with decreasing percentages of wild type

lysozyme and the Vmax values were compared to the Vmax value of irradiated Photo-

PEGylated lysozyme. A450 values were measured every 12 seconds for 20 minutes (Figure

4.8), and the Vmax values calculated from the initial slope of each reaction (first 72 sec; linear

regression resulted in R > 0.998) for wild-type lysozyme and PhotoPEGylated lysozyme

activated by the optimal UV irradiation (365 nm, 25 W, 30 min). The obtained velocities

–5 –1 –5 –1 were: Vmax(wild type) = (263 ± 6)×10 s and Vmax(Photo-PEG+UV) = (132 ± 4)×10 s .

Thus, the light-activation of Photo-PEGylated lysozyme was found to be 50% of wild-type

lysozyme activity. An incomplete restoration of enzymatic activity is not uncommon in the

light-activation of biological processes,[19] and can be easily augmented by increasing the amount of the caged biological molecule and/or extending the time of the process under study. Importantly, the Photo-PEG reagent fulfills the specific criterion of complete inactivity prior to irradiation. Thus, we successfully demonstrated the application of light- removable PEG groups in the photochemical regulation of enzyme activity.

1.0 −UV +UV (5 min) +UV (10 min) 0.8 +UV (15 min) +UV (20 min) +UV (25 min) 0.6 +UV (30 min) 450 wild-type OD 0.4

0.2

0.0 00:00 04:00 08:00 12:00 16:00 20:00 time / minutes

Figure 4.8. Lysis assays performed wild-type lysozyme and irradiated Photo-PEGylated enzyme allowed the calculation of Vmax values and subsequent determination of percent activity restoration to the Photo-PEGylated lysozyme. Error bars represent standard deviations from three independent experiments.

4.1.4 Summary and Outlook

In summary, we have developed a photocleavable PEG reagent (Photo-PEG) and

applied it successfully in the generation of a light-activated enzyme. This light-removable

PEGylation represents the first example of a literal protein photocage, as the polyethylene

glycol chains surround the enzyme and prohibit any interaction with its substrate. Irradiation

with UV light of 365 nm removes the otherwise completely stable PEG groups and restores

enzymatic activity. Lysine PEGylation with the developed reagent is rapid and facile, and the

installation of Photo-PEG groups on the protein can be easily assessed by standard gel electrophoresis, which, if necessary, facilitates purification of the caged protein. This methodology combines the advantages of protein PEGylation (increased protein solubility, intracellular stability, and circulation time) with the advantages of light-activation of protein function (spatial and temporal activity control with high precision). Due to the simplicity of the necessary manipulations, the developed methodology has a wide range of potential applications in the study of protein function in cell culture and whole organisms, as well as activation of protein therapeutics, with the only requirement being the presence of a suitable number of reactive amino groups.

4.1.5 Experimental Methods

Lysozyme PEGylation. Lyophilized hen egg white lysozyme was dissolved in phosphate buffered saline (PBS, pH 7.5) to a final concentration of 300 µM. To 100 µL of the lysozyme solution, 50 equivalents of NHS-PEG5000 or the photocleavable Photo-PEG reagent were

added as a solid and the reaction mixture was incubated overnight at 4 °C. The solution was

then dialyzed (SpectraPor, 50 kDa MWCO) over two days into PBS (pH 7.5) at 4 °C with

one change of buffer. To determine the amount of PEGylated protein present after dialysis, a

standard Bradford assay was performed.[162]

Lysozyme Assays. The lyophilized lysozyme substrate Micrococcus lysodeikticus (Sigma) was suspended in 66 mM sodium phosphate, pH 6.24, to an optical density at 450 nm of approximately 2.0, providing a 2X cell suspension stock. Wild-type or PEGylated lysozyme

(100 µL of 6 µM enzyme solution in PBS) was added to the cell suspension (100 µL) for a final enzyme concentration of 3 µM. Prior to the assay, the Photo-PEG lysozyme was kept in the dark or was selectively irradiated in one well of a 96-well plate on a transilluminator (365 nm, 25 W, 5-30 min) pre-chilled to 0 °C with an ice pack. Wild type lysozyme was also irradiated in the same manner and showed no decrease in activity, indicating that irradiation causes no discernible photodamage to the enzyme. A450 values were measured every 12 seconds for 20 minutes (SpectraMax Plus384 UV plate reader, Molecular Devices), and all assays were performed in triplicate.

SDS-PAGE Analysis. To ensure complete cleavage of Photo-PEG upon irradiation,

PEGylated lysozyme was analyzed by denaturing polyacrylamide electrophoresis (SDS-

PAGE). 100 ng wild type and PEGylated lysozyme (non-irradiated and irradiated) were mixed with 6X SDS loading buffer and separated on a 14% SDS-PAGE running gel overlaid with a 4% stacking gel at 8 mA until the buffer loading dye was clear of the stacking gel, then at 14 mA for 45 minutes. The low amperage was determined to provide more compact protein bands and reduce streaking of the highly PEGylated proteins. The gel was then stained with rapid coomassie stain for two hours, followed by incubation in the destaining solution for at least two hours.

4.2 Photochemical Control over Cre-mediated DNA Recombination

4.2.1 Introduction to DNA Recombination and Cre Recombinase

Site-specific recombinase proteins catalyze DNA strand exchange between defined target sequences on each of two DNA segments.[163] The recombinases are subdivided into

two categories, dependent upon the nucleophile involved in the reaction. The resolvase-

invertase family[164] of recombinases (e.g. Hin invertase[165] and Tn3 transposase[166])

relies on a serine nucleophile to catalyze concerted cleavage and rejoining of double-stranded

DNA at nucleotides separated by two bases. Conversely, the integrase family[167] (e.g. λ

integrase,[168] FimE, Flp recombinase,[169] and Cre recombinase[170]) utilizes a tyrosine

nucleophile to catalyze sequential strand exchanges that occur six to eight base pairs apart. In

both instances, a covalent DNA-protein intermediate, essential to recombination-site base

pairing and strand exchange, is formed. Recombination proceeds through a four-way

branched intermediate termed a “Holliday junction”,[171, 172][171] which forms when a

single DNA strand from each of the two involved DNA duplexes is exchanged with another

within a region of identical sequence to yield a branched structure (Figure 4.9). The catalytic

amino acid hydroxyl attacks a specific phosphodiester bond within the conserved double-

stranded DNA recognition site, cleaving the first strand and forming, in the case of a catalytic

tyrosine nucleophile, a 3’-phosphotyrosine intermediate and a free 5’-hydroxyl; this free

hydroxyl attacks the neighboring phosphotyrosine linkage and completes the first round of

DNA strand exchange. The Holliday junction allows branch migration through regions of

sequence homology so that the process can then be repeated with the remaining DNA

64 strands. Resolution of the final nucleoprotein intermediate completes the recombination event and results in two DNA duplexes with exchanged DNA sequences.

Figure 4.9. Site-specific recombination occurs through formation and then resolution of the Holliday junction. In the case of a tyrosine recombinase, four monomers adhere to each strand of DNA at the recognition sites of two DNA duplexes. The catalytic tyrosines of two monomers then cleave the DNA, forming the Holliday junction and two phosphotyrosine intermediates. The two free hydroxyl residues attack the nucleoprotein complex and catalyze strand exchange. The Holliday junction migrates to allow isomerization of the DNA and the process is repeated to create two new DNA duplexes. From Gopaul, D. N.; Guo, F.; and Van Dyne, G. D. EMBO J. 1998, 17, 4175.

Cre (Cyclization recombination) recombinase is a well-characterized member of the integrase family that has been used extensively in genome modification. Abremski and

Hoess[173] were the first to isolate and purify the 38 kDa protein from Bacteriophage P1.

Cre catalyzes the recombination of DNA located between two palindromic loxP (locus of

crossing over for P1) sites, each consisting of two 13 base pair repeats flanking an eight base

pair asymmetric core, thus forming the recombinase binding element (RBE). Orientation of

the core determines deletion, insertion, or inversion of the intervening DNA (Figure

4.10).[174] To facilitate recombination, a single Cre monomer binds to each RBE, thereby

forming a tetramer that joins the loxP sites so that cleavage and strand exchange may occur

(Figures 4.9 and 4.10).[175] The catalytic domain of Cre is comprised of Arg173, His289,

Arg292, and Tyr324, which form the essential “R-H-R-Y” motif conserved among members of the integrase family.[176]

Figure 4.10. Cre recombinase catalyzes the inversion or excision/integration of DNA between two loxP sites. Orientation of the 34bp loxP sites determines whether the intervening DNA will be inverted or removed/integrated. The loxP site is composed of two 13 bp inverted repeat elements to which the Cre protein binds and a spacer region (green) separating them, where the DNA breakage and reunion occurs.

Cre has a wide range of applications, from basic vector construction to silencing or

inducing gene expression, and has been an invaluable tool for engineering knockouts and

conditional alleles.[177-180] Cre has been used extensively in functional genetics for several

reasons:

• Efficacy of the Cre/loxP system in creating gene knockouts and transgenes has

been demonstrated in a variety of organisms, from E. coli and yeast,[181] to

mouse,{{86 Brault,V. 2007}} zebrafish,[182] and drosophila,[183] as well as

tobacco,[184] wheat,[185] and maize.[186]

• Unlike other members of the integrase family that require a non-linear substrate

for catalysis, Cre-mediated recombination occurs regardless of DNA substrate

conformation (linear, supercoiled, or circular).

• Cre needs no external energy source and uses only Mg2+ as cofactor.

• Successful recombination using the Cre/loxP system has been shown both in vitro

and in vivo.

While wide-spread use of the Cre/loxP system is due to its simplicity and effectiveness, there are some limitations, particularly in more complex organisms such as the mouse, that remain to be addressed:

• Temporal control over recombination events is difficult to achieve; organisms

harboring a Cre-encoding plasmid or transgenic gene express the enzyme

constitutively and thus attempts to modulate activity rely on regulation of the

promoter, e.g., through an engineered small molecule or a tissue-specific

promoter.

• Spatial control is difficult to obtain; to circumvent this limitation, Cre must be

expressed under a tissue-specific promoter and can no longer be used

ubiquitously.

In an effort to address these limitations, Feil et al. fused Cre to a mutated ligand- binding domain of the human estrogen receptor (ER) that binds the small molecule tamoxifen

[187]; recombination between two loxP sites occurred only with the addition of tamoxifen.[188] Furthermore, Cre activity was regulated through a split Cre construct,[189] in which the Cre gene was divided between amino acids 59 and 60, and each fragment was fused to the mutated ligand-binding domain of the ER. These two constructs were expressed on separate plasmids and neither individual Cre fragment was able to catalyze recombination.

Upon tamoxifen induction, the entire active Cre protein was reconstituted to enable DNA recombination. A similar, rapamycin inducible split-Cre system was generated as well.[190]

However, both systems again required small molecule induction.

As such, a means to acquire spatio-temporal control over Cre activity would be an invaluable tool in molecular biology. Providing tighter control over recombination events would also have great impact on the field of therapeutics, potentially enabling the use of the

Cre/loxP system toward the treatment of human disease. Cre activity has been mediated through the use of special heat shock promoters and by a ligand-binding event. Le et al[191] facilitated heat-shock inducible recombination by placing the Cre gene under control of the hsp70 promoter in hsp70-Cre transgenic zebrafish. The zebrafish also harbored a floxed

EGFP gene upstream of a kRASG12D oncogene that was expressed in muscle cells (Figure

4.11, A). A 37 °C heat shock was applied to the entire organism by uniform heating to obtain temporal control over DNA recombination, or to individual cells through the use of a laser, achieving spatial recombination control. However, while spatio-temporal control of

recombination was achieved without the use of tissue-specific promoters and recombination ocurred within four hours of induction (Figure 4.11, B), Cre activity was noted in the absence of heat shock (Figure 4.11, B and C), which the authors attributed to leaky expression of the hsp70 promoter.

B C

Figure 4.11. Heat shock induced Cre recombination in Zebrafish affords control over DNA recombination and expression of the kRASG12D oncogene. A. The Cre reporter construct contains a β-actin promoter for expression in muscle cells and a floxed EGFP gene with stop codon upstream of the kRASG12D gene. B. Within four hours of heat shock, recombination is apparent; however, recombination is also present after 12 hours withour heat shock. C. Over 100 cells/embryo express kRASG12D 12 hours after heat shock; additionally, nearly 20 cells/embryo show kRASG12D expression at 16 hours in the absence of heat shock. Adapted from Le, X., et al. Proc. Natl. Acad. Sci. 2007, 104, 9410.

The Koh group further mediated Cre activity by photocaging tamoxifen, the small

molecule necessary for activation of Cre when fused to the mutated ligand binding domain of

the human estrogen receptor.[192] While this was the first report of spatio-temporal control of recombination through a photoinducible system (Figure 4.12), the method required multiple light irradiations to decage the small molecule and activate recombination.

Additionally, recombination activity was low; only 26% of the activity of a β-galactosidase reporter system was reached.

Figure 4.12. Cre activity has been modulated by a mutated estrogen receptor ligand-binding domain fusion (Cre-ERT that binds tamoxifen (ER ligand). Addition of tamoxifen activates Cre, which excises neomycin (Neo) and its stop codon from between two loxP sites. Subsequent expression of the LacZ reporter gene produces β- galactosidase, which cleaves the substrate X-gal and results in blue colored cells. Wild type HEK-293T cells do not exhibit any endogenous β-galactosidase activity in the absence (A) or the presence (B) of tamoxifen. In the presence of photocaged tamoxifen, HEK-293T cells do not express β-galactosidase (C). However, upon irradiation, the cells express β-galactosidase as evidenced by their blue color (D). Adapted from Link, K. H., Shi, Y., and Koh, J. T. J. Am. Chem. Soc. 2005, 127, 13088.

Therefore, photoregulating the enzyme and not the small molecule would obviate the need for small molecule induction, and bypass difficulties with cellular uptake. Without the steric bulk of the fused ligand-binding domain, the recombination event should proceed at

100% efficiency. In order to solve these problems and provide highly stringent spatial and temporal control over Cre activity, we thus sought to photocage the enzyme directly.

As mentioned in Chapter 1, photocaging affords a high level of spatial and temporal

control over biological molecules. We therefore utilized a light-responsive caging group

through two approaches to: 1) non-specifically cage the lysine residues in Cre in vitro, and 2)

directly cage the essential tyrosine in vivo within the catalytic site of the enzyme itself.

4.2.2 Expression and Purification of Wild Type Cre Recombinase

The Cre gene was PCR amplified from P1 Phage genomic DNA.The Cre gene was

then inserted into pET-21a (Novagen) under control of the T7 promoter, generating pET-

21Cre, and yielding hexahistidine-tagged wild type Cre recombinase upon expression in E. coli. The hexahistidine tag provides not only an affinity handle for easy purification, but it has also been shown to further enhance the cellular uptake of Cre.[193] The expected vector sequence was confirmed by DNA sequencing. BL21(DE3) E. coli cells were transformed with pET21-Cre; cultures were induced with 1 mM IPTG when an OD600 of 0.6 was reached,

then expressed for four hours at 37 °C. The resulting 6xHis-tagged protein was purified by

Ni-NTA metal affinity chromatography, using a commercially available agarose resin

coupled to nitrilotriacetic acid (Ni-NTA; Qiagen). Stringent washing of the resin with

increasing concentrations of imidazole yielded the protein with up to 95% homogeneity

(Figure 4.13).

Figure 4.13. Expression of Cre recombinase from pET-21Cre yields pure protein, as determined by SDS-PAGE analysis.

Purified Cre was dialyzed into a storage buffer containing 1 M Tris-HCl, 300 mM

NaCl, 500 mM EDTA and 50% glycerol, pH 8.0, then stored at –20 °C. Between 0.5 and 1.0 mg/L of Cre were produced per expression, as determined by either UV/Vis spectroscopy using a molar extinction coefficient of 1.17x108 M-lcm-l[194] or through a standard Bradford

Assay using the Bradford Protein Reagent (BioRad). Cre was assayed for activity using

pLox2+, a commercially available linear DNA substrate containing two loxP sites bordering an ampicillin resistance gene and an origin of replication (New England Biolabs).[195]

During recombination, Cre excises the intervening DNA as a circular plasmid that confers antibiotic resistance to E. coli; enzymatic activity is subsequently determined by counting the number of resulting transformants (Figure 4.14, A).

Figure 4.14. In vitro assay of Cre recombinase activity. A. Active Cre excises an AmpR plasmid from the substrate; transformation gives rise to many colonies on selective media. B. Caged Cre is inactive and few colonies arise from transformation of the linear substrate until light-irradiation; Cre can then recombine the substrate, as visualized by the resulting number of colonies.

Each assay was performed as per published recombination protocols.[195] Cre was

incubated at 37 °C with 200 ng of pLox2+ in reaction buffer containing 50 mM Tris-HCl, 33

mM NaCl, and 10 mM MgCl2, pH 8.0 for 30 minutes. To complete the reaction, the enzyme

was then inactivated by heating to 70 °C for 10 minutes. 10 μL of cells were transformed

with 1/15 of the reaction mixture. Here, NovaBlue E. coli cells (Novagen) are required for

transformations, as they lack the recA gene that repairs nicks in cellular DNA; this genotype

ensures that colonies do not arise from transformation and intracellular ligation of the linear

substrate, but from plasmids generated through successful Cre recombination. Additionally,

because the linear substrate is generated by restriction digest, a small percentage of the

substrate remains uncut; therefore, control transformations with pure substrate must be

performed to determine background colony numbers. The activity of our expressed Cre was

compared to that of commercially available Cre recombinase (Novagen); following the

assays described above, and both enzymes produced over 1000 transformants.

4.2.3 In Vitro Non-Specific Caging of Lysine Residues

We first chose to regulate recombinase activity by photocaging Cre in vitro. We

hypothesized that non-specifically caging any of the thirteen lysine residues present in the

enzyme would attenuate its activity (Figure 4.15, B), because the steric bulk of the caging

groups would (1) inhibit proper formation of the tertiary structure necessary for catalysis and

(2) prevent DNA binding, as the crystal structure of Cre-loxP has revealed that conserved

residue Lys201 interacts with the minor groove of the DNA substrate.[172] To cage the ε-

amino group of lysine, we chose a derivative of 6-nitroveratryl chlorocarbamate (NVOC), the

NH2-specific photoprotection group developed in 1970 by the Woodward lab.[196]

Previously, Marriott reported successful photocaging of an essential lysine in the protein G-

actin using the NVOC caging group.[197] To make use of more readily available starting

materials, we utilized the nitropiperonyl chloroformate (NPOC) caging group (Figure 4.15),

in which the veratryl moiety of NVOC-Cl is replaced with a piperonyl group. The caging reaction was performed at physiological pH and the resulting carbamate linkage can be photolyzed by UV light of 365 nm.

Figure 4.15. The NPOC photocage may attenuate Cre activity until it is removed with 365 nm UV light to yield active enzyme.

Purified Cre was dialyzed into a suitable buffer (25 mM, pH 8.0) to perform the caging reaction, e.g. a buffer lacking primary amino groups that would subsequently cage in lieu of the protein. Several buffers were first tested to ensure solubility and enzymatic activity, including sodium phosphate, borate, carbonate, and PBS. Once wild type Cre activity was verified in each buffer, the caging reaction was performed with the addition of

1000 equivalents of caging group. All caging reactions were performed at 4 °C for 24-48 hours. Dialysis of the caging reaction was performed to remove excess caging groups, but repeated attempts of dialysis on the small scale required for protein caging failed to yield more than trace amounts of protein, as determined by SDS-polyacrylamide gel electrophoresis (PAGE). We then employed buffer exchange via microconcentration, but failed to elute active protein; a native Cre control reaction generated no colonies above background following recombination and NovaBlue transformation.

We therefore shifted our approach to caging of Cre immobilized on a Ni-NTA resin, making use of the 6xHis tag ligated to Cre. Purified Crewas incubated with the Ni-NTA

resin, then 1000 molar equivalents of caging group, based on the 13 lysine residues, were

added to the resin-bound Cre and the caging reaction was performed 24 hours at 4 °C.

Subsequent assays demonstrated only moderate recombinase activity with a low number of colonies.

Additionally, treating the substrate with alkaline phosphatase greatly reduced the number of background colonies. However, the photocaged irradiated (365 nm light, 10 minutes, hand-held lamp) and the photocaged non-irradiated assays generated the same number of colonies as the native Cre control. We were thus unable to ascertain if (1) Cre was successfully caged, but the caging groups did not affect tertiary structure as initially thought, or (2) Cre remained uncaged. We thus decided to photocage Cre in vivo using unnatural amino acid mutagenesis.

4.2.4 Unnatural Amino Acid Mutagenesis for the Photocaging of Cre Recombinase

Since all caging/decaging attempts in vitro were unsuccessful, we turned our attention to caging in vivo. Cre utilizes a nucleophilic tyrosine at position 324 to catalyze sequential strand exchange amongst its cognate loxP sites via formation of a covalent protein-DNA intermediate (Figure 4.16).[171, 198] As such, we hypothesized that the presence of a caging group on the nucleophilic hydroxyl group of Tyr324 would render Cre completely inactive, and thus inhibit recombination. Light irradiation would then remove the caging group and

restore Cre activity.

Figure 4.16. Crystal structure of the active site of Cre recombinase. Cre cleaves DNA strands by thenucleophilic attack of Tyr324 onto the DNA phosphodiester backbone, forming a covalent bond between the 3’-O of guanosine with the Tyr-OH group (PDB 1Q3V).

Direct caging of Cre was achieved through the use of unnatural amino acid mutagenesis. Site-directed missense mutation, in which a single nucleotide substitution within a codon leads to the incorporation of a different amino acid into a protein, has been used in protein characterization to identify essential catalytic residues,[199, 200] determine tertiary structure,[201, 202] and enhance native activity.[203] While this is an effective means to investigate protein function, mutagenesis is limited to the 20 naturally occurring amino acids, as opposed to the various chemical and structural modifications possible for a small molecule. As such, a method to include unnatural amino acids in an organism’s genetic repertoire while also controlling their site of incorporation greatly enhances the studies of protein structure and function.[204]

Incorporation of unnatural amino acids can be accomplished in four ways: (1) solid- phase chemical synthesis of small proteins, (2) ligation of chemically synthesized peptides to an expressed protein, (3), in vitro unnatural amino acid incorporation through chemically

acylated tRNAs, or (4) site-specific in vivo incorporation of the unnatural amino acid directly

from the growth medium. Method (1), developed by Merrifield,[205] is used to synthesize

proteins de novo, and various amino acid analogues, such as fluorinated leucine or valine, as

well as D-amino acids, can be incorporated into a protein.[206] Synthesis is limited, however, to peptides up to 100 amino acids in length. To circumvent this problem, method (2) was developed, joining together peptide blocks synthesized by (1) and enabling the synthesis of larger proteins.[207] A drawback to (2) is the requirement that mutually reactive side groups be placed on each peptide block.[208] Method (3) provides a decent platform for protein modification; however, the tRNA must first be chemically aminoacylated with the desired unnatural amino acid prior to the in vitro coupled transcription/translation reaction, and protein yield is low, typically less than 100 µg/mL.[204] Method (4) is not encumbered by any of the above issues associated with chemical synthesis; instead, unnatural amino acids can be incorporated into proteins of any size by harnessing the biomachinery of the cell.[209-

211] We therefore adopted this strategy to insert a photocaged tyrosine residue at position

324 of Cre recombinase, since Tyr324 provides the catalytic hydroxyl nucleophile involved

in DNA strand cleavage during recombination.

The technology to facilitate site-specific incorporation of amino acids into a protein in

vivo was developed by the Schultz group.[212-215] The methodology utilizes a nonsense

mutation to insert an unnatural amino acid in response to an amber stop codon. First, a

suppressor tRNA was generated that recognizes the amber stop codon (UAG) and inserts an

unnatural amino acid at this position; because of the poor interspecies recognition of non-

78 endogenous tRNAs, a suppressor tRNATyr library was constructed from the tyrosyl-tRNA

(tRNATyr) of the archaebacterium Methanocaldococcus jennaschii for use in an E. coli expression system. Importantly, the M. jennaschii tRNATyr must suppress the stop codon sufficiently so that protein biosynthesis can occur, while remaining uncharged by any endogenous E. coli aminoacyl tyrosyl-tRNA synthetases (TyrRS) (Figure 4.17).[215, 216]

The suppressor library was then screened to identify those tRNAsTyr able to incorporate tyrosine in the presence of their cognate M. jennaschii TyrRS. Next, the M. jennaschii TyrRS was evolved to recognize an unnatural amino acid and use it to charge its cognate tRNATyr, as demonstrated with the incorporation of O-methyl-L-tyrosine into the dihydrofolate reductase protein. The iterative evolution of a mutant TyrRS enables the charging of a suppressor tRNATyr with almost any unnatural amino acid, as shown by the successful incorporation of photocaged amino acids, to yield light-regulated proteins.[217-220]

Figure 4.17. A general approach for the site-specific incorporation of unnatural amino acids into proteins in vivo. From Wang, Q.; Parrish, A. R.; Wang, L. Chem. Biol. 2009, 16, 323.

4.2.5 Expression and Purification of CreONBY

We thus utilized unnatural amino acid mutagenesis[221-223] with the protein

biosynthetic machinery of E. coli to selectively incorporate a photocaged ortho-nitrobenzyl

tyrosine (ONBY)[217] at position Tyr324. A wild type Cre expression vector (pET-21Cre)

was constructed, and Tyr324 was mutated to an amber stop codon TAG324 (pET-

21CreY324TAG). Expressions were performed with pET-21CreY324TAG and the plasmid

pSupONBY, which encodes six copies of the supressor tRNATyr and one copy of the

TyrRS.[217] In the presence of ONBY, photocaged CreONBY is produced, while expression

in the absence of ONBY did not result in any Cre protein (Figure 4.18).

Figure 4.18. Expression of Cre recombinase in its wild type form and containing a caged tyrosine at position 324. TAG = amber stop codon. Structure of ortho-nitrobenzyl tyrosine (ONBY).

In order to increase the yield of CreONBY, expressions were performed in lieu of

pSupONBY with a modified version of the plasmid pEVOL (Schultz Lab)[220] that was

recently constructed to allow higher yields of proteins containing unnatural amino acids. In

contrast to pSupONBY whose TyrRS and tRNATyr were constitutively active, pEVOL

contains two copies of an aminoacyl tRNA synthetase and one copy of a tRNATyr, all under control of the araC promoter and thus inducible with arabinose. The new pEVOL is less toxic to cells, as evidenced in our lab by a decrease in time to reach optimal optical density and a higher number of doubly transformed E. coli cells growing on agar plates. To generate the modified pEVOL used to express CreONBY, Dr. Chungjung Chou (Deiters Lab) replaced the aminoacyl tRNA synthetase for with TyrRS. To express caged Cre, E. coli were

transformed with the modified pEVOL and pET-21CreY324TAG. As with pSupONBY, expression in the presence of 1 mM ONBY yielded CreONBY, while expression in the absence of the amino acid did not produce protein. In comparison to expression with pSupONBY, the yield of CreONBY was markedly increased (Figure 4.19).

Figure 4.19. Expression with our modified pEVOL produced a greater amount of CreONBY than expression with pSupONBY. M: protein marker; lane 1: wild type Cre; lane 2: pEVOL expression with no ONBY; lane 3: pSup expression with ONBY; lane 4: pEVOL expression with ONBY.

4.2.6 In Vitro Light-Activation

To verify that our photocaged Cre recombinase is inactive until irradiated, assays

were again performed with 200 ng of the substrate pLox2+ and 50 ng of Cre or CreONBY to

assure that the substrate was not limited and that recombined DNA was not due to the action

of only a small quantity of decaged enzyme. In separate reactions, wild type Cre and

CreONBY were kept in the dark or irradiated with an LED fiber optic lamp (Prizmatix

BLCC-02, 30 W, 365 nm, 20 min), then incubated at 37 °C for 30 minutes, followed by heat inactivation and transformation. A timecourse was performed to determine the irradiation conditions that would restore maximum Cre activity as compared to wild type (Figure 4.20).

Wild type Cre and CreONBY irradiated for twenty minutes both yielded >1000 colonies

upon transformation, while both non-irradiated CreONBY and the pLox2+ control yielded

<15 colonies. This confirmed that photocaged CreONBY is completely inactive prior to

removal of the caging group and that light irradiation restores recombination activity to

~70% of that of wild type Cre. Irradiation times between 25 and 60 min did not lead to increased recombination and thus 20 min of irradiation was used for the in vitro assays.

6000

5000

4000

3000

2000 Number of Colonies of Number 1000

0 plox2+ no UV 2 min 5 min 10 min 20 min WT contol Figure 4.20 Light irradiation timecourse of CreONBY. Irradiation for 20 min restored about 70% activity compared to that of wild type Cre (365nm, fiber optic lamp, 37 °C). The wild type Cre reaction was also irradiated to ensure that neither protein nor substrate was damaged by irradiation.

4.2.7 In Vivo Light-Activation and Spatial and Temporal Control of Gene Function

In order to demonstrate photochemical regulation of Cre activity in mammalian cell culture, HEK293T cells were transfected with the Cre Stoplight plasmid (pC-SL; Figure

4.21)[224] and CreONBY. pC-SL encodes DsRed and a transcription termination region, both flanked by loxP sites and located upstream of a GFP gene. Prior to Cre-mediated recombination, cells exclusively express DsRed under control of a CMV promoter; after recombination and the ensuing excision of DsRed and its terminator, cells express GFP.

Figure 4.21. The Cre Stoplight plasmid (pC-SL). Adapted from Yang, Y. S.; and Hughes, T.E. Biotechniques 2001, 31, 1036.

In the absence of Cre recombinase, cells produce DsRed and exhibit red fluoresence

(Figure 4.22, A). As expected, transfection of wild-type Cre led to in vivo recombination and

observation of GFP expression. (Figure 4.22, B). Importantly, cells transfected with caged

CreONBY exclusively showed DsRed expression, verifying the complete inactivity of the

caged Cre protein in mammalian cells (Figure 4.22, C). However, a brief UV irradiation with

a handheld UV lamp (365 nm, 23 W, 5 min) almost exclusively yielded GFP expression

(Figure 4.22, D). UV light alone had no effect on the conversion of DsRed to GFP (Figure

4.23).

Figure 4.22. Spatial and temporal control of Cre recombinase in vivo. A-D) Photochemical control of Cre- catalyzed DNA recombination in 293T cells transfected with the Cre-Stoplight plasmid. In the absence of wild type Cre (A), only DsRed is expressed, while in the presence of wild type Cre, the DsRed gene is excised, and GFP is expressed (B). Non-irradiated caged Cre leads to the expression of DsRed (C). Irradiated caged Cre is activated and thus leads to the expression of GFP and the silencing of DsRed (D). (E-F) Spatial control of caged Cre: recombination of the Cre-Stoplight plasmid occurred only in irradiated cells (within the dashed circle), yielding GFP expression (E); this is further seen in the overlaid image of GFP and DsRed after spot irradiation (F). Irradiation was performed for 20 sec with an epi-fluorescence inverted microscope equipped with a 100 W mercury lamp and a DANSA filter cube (330-400 nm), followed by imaging with a Lecia DM5000B microscope 48 hours post-transfection, 100× magnification.

In order to obtain spatial control over DNA recombination in mammalian cell culture, a mono-layer of HEK-293T cells co-transfected with pC-SL and CreONBY was irradiated in a small area for 20 sec with an epi-fluorescence inverted microscope equipped with a 100 W mercury lamp and a DANSA filter cube (330-400 nm). While red fluorescence was detected in all cells, green fluorescence was only observed in the irradiated area of the cell monolayer

(dashed circle) demonstrating spatial control over the DNA recombination event (Figure

4.22, E-F). We thus confirmed that DNA recombination in HEK293T cells can be effectively triggered with light in a spatiotemporal fashion.

Figure 4.23. In the absence of CreONBY, UV irradiation (365nm, 23W, 5 min) does not affect the conversion of DsRed to GFP in cells transfected with pC-SL. Only DsRed is expressed. Imaged with Lecia DM5000B microscope 48 hours post-transfection, 100× magnification.

To quantitate the observed recombination activity of CreONBY, flow cytometry data was collected and analyzed with a two-dot plot (Figure 4.24 and Table 4.2). As expected, cells transfected with only pC-SL exhibited a high level of red fluorescence (84.0 %), with

17.1 % of cells exhibiting both red and green fluorescence. Only 1 % of cells exhibited solely green fluorescence. It is important to note that DsRed matures through a green intermediate stage,[225] accounting for green fluorescence in the absence of Cre.

Figure 4.24. Results of flow cytometry counting of transfected cells. The number of cells expressing DsRed and GFP were counted and a significant amount of recombination was observed in the irradiated cells containing caged Cre. A) HEK293T cells; B) no caged Cre; C) caged Cre –UV; D) caged Cre +UV.

The levels of red and green fluorescence after CreONBY transfection, but prior to light

irradiation, remained consistent with those of transfected cells not exposed to Cre (80.4% red

fluorescence; 18.3% of cells expressed green and red fluorescence; 1.3% of cells exhibited

green fluorescence). However, upon irradiation of CreONBY, a marked decrease in red fluorescence was observed (67.2%), with an increase in the number of cells expressing both green and red fluorescence (27.1%). An increase in cells expressing only green fluorescence was also detected (~6%), indicating activation of the Cre protein and subsequent recombination.

Table 4.2. Percentage of red and green fluorescence in HEK-293T cells transfected with pC-SL and/or caged Cre as detected by flow cytometry. Similar levels of fluorescence are noted in cells not exposed to caged Cre or not irradiated. A statistically significant decrease in red fluorescence and simultaneous increase in green fluorescence is detected for irradiated cells containing caged Cre.

% Red % Red and Green % Green Fluorescence Fluorescence Fluorescence no caged Cre 84.0 17.1 1.0 caged Cre –UV 80.4 18.3 1.3 caged Cre +UV 67.2 27.1 5.7

4.2.8 Expression of TAT-Modified Cre

In an effort to improve the DNA recombination rate of CreONBY in mammalian cells

by obviating the use of inefficient transfection reagents, we began investigating the cellular

uptake of TAT-modified Cre and CreONBY. When expressed as a fusion, the nine amino

acid TAT transduction sequence of HIV[226], as well as other transduction domains,[227-

229] enables its partner protein to cross the cell membrane via protein transduction. TAT-

Cre has been shown to provide an effective means of cell-permeant uptake and subsequent

DNA recombination.[193, 230-232] While the exact mechanism of protein cell permability is

heavily debated, it is thought that the nine amino acid TAT sequence interacts

electrostatically with the cell membrane; this disrupts the membrane and enables the fusion

protein to pass into the cell.[233] We thus hypothesized that TATCreONBY would be readily

taken up by a cell from its standard growth medium, and thus greatly expand the utility of

light-activated DNA recombination. We attempted to insert the TAT sequence upstream of

the Cre gene in both pET-21Cre and pET-21CreY324TAG through the use of restriction

digest and hybridization of TAT sense and antisense strands (IDT DNA). However, this

88 resulted in multiple insertions of the TAT sequence that caused a frameshift in the Cre gene.

Additionally, Jie Zhang (Deiters Lab) constructed pBAD-TATCre, but was not able to express protein. We thus obtained the plasmid pTriEx-HTNC-Cre[230](Addgene) that contains a 5’-6xHis tag, a 5’-TAT sequence, and a 5’-nuclear localization sequence (NLS) from the SV40 large T antigen that targets the protein to the nucleus via the nuclear pore complex.[234] NovaBlue (DE3) E. coli cells were transformed with TriEx-HTNC and cultures were induced with 1 mM IPTG when an OD600 of 0.6 was reached. The resulting protein was purified by Ni-NTA metal affinity chromatography with up to 95% homogeneity

(Figure 4.25) and then dialyzed into Cre storage buffer. A Bradford assay was performed to measure the concentration of TATCre, which was determined to be 110 ng/µL.

M TATCre 62

48 33

Figure 4.25. Expression of TATCre from pTriEx-HTNC yields highly pure protein as determined by SDS- PAGE analysis.

TATCre was assayed for activity using our standard in vitro assay with 100 ng of protein and 200 ng of pLox2+ substrate; however, the assay resulted in only 50 colonies as compared to over 500 colonies from wild type Cre and no colonies on the control plate. As previous reports of TATCre give comparable activity to wild type Cre, TATCre will need to be expressed again and assayed to ensure robust activity prior to further testing in vivo. Once

activity is improved, Tyr324 of the protein will be mutated to an amber stop codon for expression of TATCreONBY from pTriEx-HTNC.

4.2.9 Summary and Outlook

In summary, we have expressed Cre recombinase and shown that its activity can be stringently regulated both spatially and temporally in vitro and in mammalian cell culture through the use of a light-cleavable caging group installed directly on the tyrosine residue essential for catalysis. The described photocaged Cre/lox system has numerous applications toward the creation of knock-in and knock-out organisms in a spatio-temporal fashion.

Additionally, the insertion of the TAT domain at the N-terminus of Cre and CreONBY should allow cells to take up the protein directly from their medium, eliminating the need for less efficient chemical transfection. Some limitations with this system still exist, including the difficulty in expressing CreONBY, as well as the inability to activate CreONBY in a location that is not accessible to light. The use of CreONBY in model organisms will most likely be facilitated by microinjection[235] to introduce the protein or by transitioning the components necessary for ONBY incorporation (synthetase and tRNA) into eukaryotic cells and multicellular organisms. It would also be interesting to investigate the use of both wild- type and caged Cre in extracted tissue samples, e.g. brain tissue from a mouse harboring a floxed gene. Additionally, the use of caged Cre to trigger RNA interference from a floxed dsRNA plasmid is cell culture could provide an alternative stragety for the photoregulation of gene function.

4.2.10 Experimental Methods

pET-21Cre Vector Construction. Top-10 cells (Invitrogen) were used for cloning and sub-

cloning experiments. The Cre gene was PCR amplified (95 °C (2 min), then 30 cycles of 95

°C (30 sec), 55 °C (30 sec) and 72 °C (1.5 min)) from bacteriophage P1 (a gift from Dr. Eric

Miller at NCSU) using the primers 5’-

AGAGAGGCTAGCTCCAATTTACTGACCGTACACCAA-3’ and 5’-

AGAGAGCTCGAGATCGCCATCTTCCAGCAGGCGCACCATTGCCCCTGT-3’ which contain NheI and XhoI restriction sites, respectively. The Cre gene was then inserted into pET-21a(+) as an NheI-XhoI fragment, generating pET-21Cre, and yielding hexahistidine-

tagged wild type Cre recombinase upon expression in E. coli. The hexahistidine tag provides

not only an affinity handle for easy purification, but it has also been shown to further

enhance the cellular uptake of Cre.[193] The expected vector sequence was confirmed by

DNA sequencing using the internal sequencing primers 5’-CTGATTTCGACCAGGTTCGT-

3’ and 5’-GCTAACCAGCGTTTTCGTTC-3’.

pET-21CreY324TAG Vector Construction for Unnatural Amino Acid Mutagenesis.

Following the manufacturer’s protocol (Stratagene), QuikChange mutagenesis was employed to mutate Tyr324 into an amber stop codon (TAT→TAG) with primers 5’-

GGACCAATGTAAATATTGTCATGAACTAGATCC-3’ and 5’-

ATCCAGGTTACGGATCTAGTTCATGACAATATT-3’, creating pET-21CreY324TAG.

Successful amber stop codon mutation was confirmed by DNA sequencing using the internal

Cre sequencing primer 5’-CTGATTTCGACCAGGTTCGT-3’.

Wild Type Cre Expression and Purification. To express wild type Cre, BL21(DE3) Gold cells (Stratagene) were transformed with pET-21Cre, inoculated 1:100 mL in 25 mL LB medium containing 50 µg/mL ampicillin, and were grown at 37 °C, 250 rpm. At an OD600 of

0.6, expression was induced with IPTG (0.5 mL of a 1 M stock solution) to a final concentration of 0.5 mM for four hours at 37 °C, followed by 15 hours at 25 °C. Freezing the cell pellet and storing it overnight at 80 °C resulted in a substantial loss of enzyme activity, thus cells were pelleted, lysed, and the protein was immediately purified in a 4 °C cold room.

Ni-NTA resin (Qiagen) was used to purify both wild type and caged Cre according to the manufacturer’s instructions. Briefly, 1 mL of Qiagen lysis buffer and 25 mg of lysozyme were added to the cell pellet resulting from a 25 mL culture (~ 250 µL), and cells were resuspended and incubated one hour on ice. For complete lysis, cells were vortexed and sonicated (Branson Sonifier 450) with a microtip attachment (2 min continuously on ice, 60

W, 50 % duty cycle). Cellular debris was pelleted in an Eppendorf refridgerated centrifuge at

4°C, 13,200 rpm, 30 min, then the lysate was decanted and 25 µL of Ni-NTA resin was added to approximately 1 mL of cleared lysate in a 1.5 mL eppendorf tube. The resin was placed on a rocker in the cold room for one hour, then pelleted (3000 rpm, 15 sec) and washed with five times with 1 mL Qiagen wash buffer. 50 µL of Qiagen elution buffer was added to the resin repeatedly until electrophoretic analysis showed no protein (typically three-four total elutions). Protein production and purity were verified by 12% SDS-PAGE.

The samples were run 12 mA to allow the loading dye to clear the stacking gel (about 15-20 min), then electrophoresed at 24 mA for 45-60 min or until the prestained protein marker

(NEB) was adequately separated. Finally, elution fractions containing Cre were pooled and dialyzed (Fisherbrand regenerated cellulose dialysis tubing; 6000-8000 nominal MWCO) at 4

°C into storage buffer (20 mM Tris-HCl, pH 7.8, 300 mM NaCl, 1 mM EDTA, and 50% glycerol) over 24 hours with two changes of buffer. Dialyzed Cre was stored at –20 °C. The enzyme concentration was determined by a Bradford assay to be 250 ng/µL (Figure 4.7), with an overall protein yield of 2 mg/L. No loss of activity was noted after one year.

In Vitro Photocaging of Cre. For non-specific, in vitro photocaging of lysine residues within Cre, a 100X caging group solution was made with 51 mg NPOC-Cl dissolved in 700

μL DMSO. To 100 ng of Cre that had been dialyzed into sodium phosphate, borate, carbonate, or PBS buffer (50 μL), all pH 7.8, 1 μL of the 100X caging group solution was added and incubated 24 to 48 hours at 4 °C. Photocaged Cre was irradiated in the presence of pLox2+ substrate at 365 nm for 20 minutes at 37 °C with a fiberoptic LED system (365 nm,

20 min), then incubated an additional 30 minutes at 37 °C, followed by heat inactivation and transformation as described in the In Vitro DNA Recombination section below.

Transformants were counted the following day to determine recombination efficiency.

CreONBY Expression and Purification with pSupONBY. To produce caged Cre,

BL21(DE3) Gold cells were transformed with both pET-21CreY324TAG and pSupONBY

(Schultz lab), the vector encoding the Methanococcus jennaschii tRNA/tyrosyl-tRNA

synthetase pair, then grown in LB medium containing 50 µg/mL ampicillin, 30 µg/mL chloramphenicol, and enriched with 1 mM ONBY. An ONBY stock solution (100mM,

100X) was made by dissolving 32 mg ONBY in 50 µL hot (80 °C) DMSO, followed by the addition of hot water (80 °C, 950 µL) to a total volume of 1 mL. The appropriate amount of

ONBY was added to 37 °C pre-warmed LB medium dropwise with intense swirling to achieve a 1 mM concentration; the pH of the media was measured, and if necessary adjusted to pH 7.0 prior to inoculation. A control expression was also performed without ONBY to verify that no protein was produced in the absence of the unnatural amino acid, and to allow determination of the correct optical density at which to induce protein expression without optical interference from the ONBY precipitate. The OD600 of the control culture was

determined spectroscopically. Expression was again induced at an OD600 of 0.6, and the protein expressed and purified as described above. Protein production and purity were verified by 12% SDS-PAGE. Finally, like Cre, CreONBY was dialyzed into storage buffer

(20 mM Tris-HCl, pH 7.8, 300 mM NaCl, 1 mM EDTA, and 50 % glycerol), and stored at

–20 °C. A Bradford assay was performed and the concentration determined to be 75 ng/µL, with an overall protein yield of 1 mg/mL. No loss of activity was noted after one year of storage.

CreONBY Expression and Purification with pEVOL-ONBY. To produce caged Cre with

pEVOL-ONBY, BL21(DE3) Gold cells were transformed with both pET-21CreY324TAG

and pEVOL-ONBY (modified by Chungjung Chou, Deiters Lab), then grown in LB medium

containing 50 µg/mL ampicillin, 30 µg/mL chloramphenicol, and enriched with 1 mM

ONBY. An ONBY stock solution (100 mM, 100X) was made by dissolving 32 mg ONBY in

50 µL hot (80 °C) DMSO, followed by the addition of hot water (80 °C, 950 µL) to a total

volume of 1 mL. The appropriate amount of ONBY was added to 37 °C pre-warmed LB

medium, dropwise with intense swirling to achieve a 1 mM final concentration; the pH of the

media was measured, and if necessary adjusted to pH 7.0 prior to inoculation. The starter was

inoculated 1:100 into the LB medium. A control expression was also performed without

ONBY to verify that no protein was produced in the absence of the unnatural amino acid, and

to allow determination of the correct optical density at which to induce protein expression

without interference from the ONBY precipitate. The OD600 of the control culture was

determined spectroscopically. After approximately four to five hours, expression of the

TyrRS was induced at OD600=0.6 with 0.02 % arabinose and expression of CreONBY was induced with 0.5 mM IPTG for four hours at 37 °C, followed by 15 hours at 25 °C. The

protein was isolated and purified as described above. Protein production and purity were

verified by 12% SDS-PAGE. The samples were run 12 mA to allow the loading dye to clear

the stacking gel (about 15-20 min), then electrophoresed at 24 mA for 45-60 min or until the

prestained protein marker (NEB) was adequately separated. Finally, like Cre, CreONBY was

dialyzed into storage buffer (20 mM Tris-HCl, pH 7.8, 300 mM NaCl, 1 mM EDTA, and 50

% glycerol), and was stored at –20 °C. A Bradford assay was performed and the

concentration determined to be 90 ng/µL, with an overall protein yield of 1.5 mg/mL. This

was a slightly higher yield than that obtained with pSupONBY.

In Vitro DNA Recombination. In vitro Cre recombination assays were performed for 30

minutes at 37 °C in 30 µL recombination buffer (50 mM Tris-HCl, 33 mM NaCl, 10 mM

MgCl2, pH 7.5) with 200 ng pLox2+ substrate and 100 ng Cre. Cre was heat inactivated at 70

°C for five minutes prior to transformation into ultracompetent recA– NovaBlue cells

(Novagen) following the manufacturer’s protocol. Briefly, 10 µL of cells were thawed on ice

for 15 minutes, then 1 µL of the Cre recombination reaction was added. Cells were incubated

on ice with the DNA for 30 mintues, then heat shocked 40 sec at 42 °C. The cells were

returned to ice for 2 min and 90 µL of SOC medium were added. The cells were then plated

on LB agar containing 50 µg/mL Amp and left inverted, overnight, at 37 °C. To determine

the number of background colonies, control transformations were also performed solely with

the DNA substrate. CreONBY was irradiated in the presence of substrate at 365 nm for 20

minutes at 37 °C with a fiberoptic LED system (365 nm, 20 min), then incubated an

additional 30 minutes at 37 °C, followed by heat inactivation and transformation as described

above. An irradiation timecourse was performed to determine the optimum irradiation time

(20 min, restoring 70% of wild type Cre activity)

Cellular Viability Assays. In order to assess the cellular toxicity of UV irradiation,

HEK293T cells were either irradiated (hand held UV lamp, 365 nm, 23 W, 5 min) or not

irradiated (three wells each in a 96-well plate) at 80 % confluency. Cultures were incubated for 24 hours, and then assayed using Cell Titer Glo Assay (Promega). A positive control

experiment with dead cells was achieved via desiccation of three wells. To each well 20 uL

of Cell Titer Glo reagent was added and incubated with the cells for four hours. Fluorescence measurements (λex = 530 nm/λem = 590 nm) were performed on a Labsystems Fluoroskan

Ascent FL plate reader. Non-irradiated cells: 384 ± 45 RFU; irradiated cells: 366 ± 28 RFU;

dead cells: 24 ± 1 RFU. Thus, no measurable cytotoxicity was observed as a direct effect of

UV irradiation.

In Vivo DNA Recombination. The following experiments were conducted by Dr. Douglas

Young (Deiters Lab). HEK-293T cells were passaged into two 4-well chamber slides, and

grown to 40% confluence. The cells were then transfected with the Cre stoplight construct (1

μg) alone (2 wells per slide), or with either wild type Cre recombinase (3 µg, 1 well per slide)

or caged Cre recombinase (10 µg, 1 well per slide) using DOTAP transfection reagent (5:2

DOTAP/DNA; Roche Biomedicals) in OptiMEM medium (Invitrogen). The transfection was

incubated at 37 oC for 4 hours, followed by replacement of tranfection media in one of the

non-protein treated Cre stoplight wells, followed by a subsequent transfection with wild type

Cre recombinase (3 µg) using the DOTAP transfection reagent. This and the remaining wells

were then incubated for 24 hours, followed by the removal of the OptiMEM transfection

medium, irradiation of one slide (2 min, 365 nm, 23 W handheld UV lamp), and replacement

of medium with standard growth media (Dulbecco’s modified Eagle’s medium; Hyclone

with 10% Fetal Bovine serum (Hyclone) and 10% streptomycin/ampicillin (MP

o Biomedicals). The cells were then incubated at 37 C (5% CO2) for 24 hours then imaged on

a Lecia DM5000B to assess GFP and DsRed expression.

Spatial Regulation of Cre Recombinase Activity. The following experiments were

conducted by Dr. Douglas Young (Deiters Lab). HEK-293T cells were passaged into a 4- well chamber slide, and grown to 40% confluence. The cells were then transfected with the

Cre stoplight construct (1 μg) and caged Cre recombinase (10 µg, 1 well per slide) using

DOTAP transfection reagent (5:2 DOTAP/DNA; Roche Biomedicals) in OptiMEM medium

o (Invitrogen). The transfection was incubated at 37 C, 5% CO2 for 24 hours, followed by the

removal of tranfection medium and spot irradiation using a Jenco Epi-fluorescence inverted

microscope equipped with a 100W mercury lamp and a DANSA filter cube (330-400 nm

excitation) for 30 seconds. The medium was then replaced with standard growth medium,

and the cells were incubated 48 hours then analyzed on a Lecia DM5000B compound

microscope for expression of GFP and DsRed reporter proteins.

Flow Cytometry. The following experiments were conducted by Dr. Douglas Young

(Deiters Lab). HEK-293T cells were passaged into 2 6-well culture plate, and grown to 40%

confluence. The cells were then transfected with a no transfection control well, a Cre

stoplight plasmid control well, a wild type Cre and Cre stoplight plasmid well, and a caged

Cre and Cre stoplight well using the previously described protocol. After 24 hours the

98 transfection medium was removed and one 6-well plate was irradiated for 2 minutes with a hand-held UV lamp (365 nm, 23 W). The media was then replaced with standard growth mediuum and the cells were incubated at 37 oC for 48 hours. The growth medium was then removed, and the cells were trypsinized (500 µL, HyClone Trypsin, VWR International), and transferred to culture tubes. Cells from each well were then counted on a Becton-Dickinson

FACSCalibur cell counter for 20,0000 events monitoring GFP (525 nm) and DsRed (600 nm) emission wavelengths.

4.3 Photochemical Control over FimE-mediated DNA Recombination in E. coli

4.3.1 Introduction to FimE

After achieving photochemical control over DNA recombination in mammalian cell

culture with photocaged Cre, we further sought to obtain such robust control in E. coli by

developing a photocaged FimE recombinase. FimE is naturally active in bacteria, whereas

Cre is most active in eukaryotic systems,[236, 237] and we have not been able to observe any

Cre recombinase activity in E coli. The inactive recombination is most likely due to a cellular

concentration of Mg2+ that is too low[238], even when the growth medium was supplemented

with additional Mg2+. A member of the integrase family, FimE is involved in the phase-

variable growth and regulation of type 1 fimbriae in gram-negative bacteria.[239] These

fringe-like appendages are required for cellular adhesion and convey virulence to E. coli.[240] Expression is regulated through site-specific inversion of the Fim switch

(FimS),[241][242][241] a 314 bp stretch of DNA containing the promoter for the fimbrial structural genes bound by two nine base-pair inverted repeats, which form the recombinase binding elements.{{601 Leathart,J.B. 1998}} In bacteria, the promoter is active only in one orientation, and recombination is thought to require critical cellular cofactors which results in inactivity of purified FimE in vitro.[243] Unlike Cre, which catalyzes DNA recombination bi-directionally, FimE can only invert the FimS gene unidirectionally; a second enzyme,

FimB, works to return the switch to its original orientation.[244] Thus, no eqilibrium

between switched and non-switched genes exists in the absence of FimB, as opposed to the

equilibrium established during Cre recombination. While FimE has not yet been utilized in

100

molecular biology to the extent of Cre, the ability of FimE to singly invert a promoter upstream of any target gene has great potential in the engineering of a cellular switch for

controlled gene expression.[245]

The catalytic domain of FimE is comprised of Arg41, His136, Arg139, and Tyr171,

[246] forming the essential “R-H-R-Y” motif conserved among members of the integrase family; DNA recombination is thus mediated by the catalytic tyrosine at position 171.[246]

Therefore, we reasoned that a photochemical strategy similar to the one we employed for the

photoregulation of Cre recombinase could be implemented to control FimE activity in E.

coli.

The Keasling lab recently developed an inducible gene expression system utilizing

FimE. [245] The FimE gene was cloned into a bacterial expression vector under control of

the araC promoter; upon addition of arabinose to the system, FimE is expressed. The GFP

gene was cloned downstream of FimS in the “off” orientation such that active FimE would

invert the promoter and drive subsequent expression of GFP (Figure 4.26).

101

Figure 4.26. FimE mediated recombination inverts the promoter (Ptrc*) located between the inverted repeat right (IRR) and inverted repeat left (IRL) of the FimS gene, leading to GFP expression. From: Ham, T. S.; Lee, S. K.; Keasling, J. D.; and Arkin, A. P. Biotechnol. Bioeng. 2006, 94, 1.

Special BLR(DE3) E. coli cells (Invitrogen) with a fim- genotype were required for

this assay to ensure that recombination and GFP expression were not due to native FimE activity. Additionally, cells were grown on LB agar medium containing 0.5% glucose to repress leaky expression from the araC promoter. Fluorescence was then quantified to

determine expression levels and thus recombination efficiency (Figure 4.27). Cells that were

not induced exhibited minimal GFP fluorescence, most likely attributable to cellular

autofluorescence.[247] As such, the recombination-catalyzed inversion of the promoter and subsequent GFP expression demonstrated the applicability of FimE to inducible gene expression.

102

1400

1200 600 1000

800

600

uninduced 400 induced GFP Fluorescence/OD GFP 200

0 0 2 4 6 8 10 Time (h)

Figure 4.27. FimE recombination as measured by GFP expression levels, which differ greatly between induced (red square) and uninduced (blue diamond) cultures. FimE recombination was induced at 0 h with 5 mM arabinose and subsequent GFP fluorescence was quantified every hour for eight hours. The uninduced control produced only minimal fluorescence, while the level of fluorescence increased six-fold for the induced culture. Adapted from Ham, T. S.; Lee, S. K.; Keasling, J. D.; and Arkin, A. P. Biotechnol. Bioeng. 2006, 94, 1.

We chose to use the Keasling GFP expression system as a reporter for photocaged

FimE, as the FimE and GFP genes are encoded by the same plasmid and would be amenable

to co-transformation. Mutation of Tyr171 to the amber stop codon and subsequent

transformation with pEVOL into E. coli should yield inactive, photocaged FimE

(FimEONBY). The FimEONBY could then be activated by irradiating living bacteria.

Successful decaging and recombination would be measured over time by GFP fluorescence,

providing the first example of in vivo photocontrol over DNA recombination in E. coli.

103

4.3.2 FimE Expression and In Vivo Assays

We initially sought to clone the FimE gene by PCR amplifying it from Top-10 cells and inserting it into pBAD-33mycHis for 6XHis tag-mediated purification and generation of pBAD-FimE. While FimE has been shown to be active only in vivo or in cell lysate,[243] our

purpose was to demonstrate successful expression by SDS-PAGE and mass spec analysis

prior to assaying recombination in E. coli. DNA sequencing showed that the FimE gene was

correctly inserted into the vector, but repeated expressions with varying conditions (see

experimental protocols) yielded no protein.

The empty pFIP vector that encoded both FimE and the inverted promoter (See

Section 4.3.1) [245] was obtained from Addgene. We thought that the GFP gene had been

cloned into the vector as a reporter downstream of the promoter region, such that FimE-

induced recombination would lead to GFP expression in the same manner as described by T.

S. Ham, et al. However, arabinose induction did not lead to higher fluorescence levels than

non-induction (Table 4.3). We thus determined, after contacting Dr. Hahn, that no ribosome

binding site (RBS) was present on the vector and it had to be inserted along with a selected

reporter gene such as GFP. Additionally, Addgene had not correctly propagated the plasmid

on 0.5% glucose and thus leaky FimE expression had already inverted the pFIP promoter.

104

Table 4.3. Expression of FimE from pFIP and subsequent recombination, as measured by GFP fluorescence. Relative fluorescence units (RFU) were nearly identical between arabinose induced and uninduced cultures. Standard deviations were obtained from three independent experiments.

Hours Post-Induction Arabinose Induction No Induction 0 5550 ± 351 RFU 4022 ± 344 RFU 5 7447 ± 2058 RFU 6654 ± 1163 RFU 15 12424 ± 1334 RFU 10858 ± 2124 RFU

Thus, the unmodified pFIP plasmid was directly obtained from the Keasling Lab, and a cloning strategy to insert the RBS and GFP DNA sequences from pGFPuv (Clontech) into pFIP was designed, but not yet conducted. Once these sequenced have been cloned into pFIP it will enable us to express GFP upon FimE-catalyzed DNA recombination. We can then mutate the essential Tyr171 to the amber stop codon and express FimEONBY in the presence of modified pEVOL. Direct irradiation of the growing E. coli cells would decage and activate the protein, leading to inversion of the promoter and resulting in GFP fluorescence. As such, we can demonstrate the first example of light-induced DNA recombination in E. coli.

4.3.3 Summary and Outlook

We have begun work to create a light-activatable DNA recombination system in E. coli. The photocaged FimE can be both expressed and assayed in vivo without the additional steps of protein isolation and purification. Importantly, in vivo photoregulation of FimE recombinase provides an additional level of control over DNA recombination that, unlike Cre recombinase, can be employed in E. coli. A light-induced genetic switch based on DNA

105

recombination in E. coli will add a very useful part to the growing repertoire of synthetic

biology.

4.3.4 Experimental Methods

pBAD-FimE Vector Construction. The 780 bp FimE gene was PCR amplified (95 °C (2 min), then 30 cycles of 95 °C (30 sec), 55 °C (30 sec) and 72 °C (1.5 min)) from E. coli

BL21(DE3) Gold cells added to the PCR reaction mixture using the primers 5’-

GCCCCATGGAAATGAAGAATAAGGCTGA-3’ and 5’-

CCCAAGCTTTAAAACAGCGTGACGCT-3’ which contain NcoI and HindIII restriction

sites, respectively. The FimE gene was then inserted into pBAD-mycHis (Invitrogen) as an

NcoI-HindIII fragment, generating pBAD-FimE. The expected vector sequence was confirmed by DNA sequencing of the entire FimE gene using the sequencing primer 5’-

ATGCCATAGCATTTTTATCC-3’.

Expression and Purification of FimE from pBAD-FimE. E. coli Top-10 cells were transformed with pBAD-FimE, inoculated 1:100 mL in 25 mL LB medium containing 50

µg/mL ampicillin, and were grown at 37 °C, 250 rpm. At an OD600 of 0.6, expression was induced with arabinose (250 µL of a 20% stock solution) to a final concentration of 0.2%.

Several induction times were attempted, for four, six, eight or 15 hours at 37 °C, to the same times followed by 15 hours at 25 °C, then cells were pelleted, lysed, and purified with Ni-

NTA resin according to the manufacturer’s instructions. Briefly, 1 mL of Qiagen lysis buffer

106

and 25 mg of lysozyme were added to the cell pellet resulting from a 25 mL culture (~ 250

µL), and cells were resuspended and incubated one hour on ice. To lyse, cells were vortexed and sonicated in a VWR sonication bath, model 150D (six cycles of 15 sec of sonication followed by 15 sec on ice). Cellular debris was pelleted in an Eppendorf refridgerated

centrifuge at 4 °C (13200 rpm, 20 min), then 50 µL of Ni-NTA resin was added to approximately 1 mL of cleared lysate. The resin was placed on a rocker in the 4 °C cold room for one hour, then washed four times with 1 mL of Qiagen wash buffer and eluted twice in 75 µL each of Qiagen elution buffer. Aliquots were then analyzed by 12% SDS-

PAGE; however, regardless of the different expression conditions investigated (see above),

no 23.5 kDa FimE protein was detected.

Conditions for Propagation of pFIP in E. coli. pFIP was sent directly from the Keasling

Lab. We received three BLR(DE3) E. coli colonies harboring pFIP on an LB agar plate

containing 50 µg/mL ampicillin and 0.5 % glucose (dextrose). Subsequent liquid cultures

were grown from one of the colonies in LB media supplemented with Amp and 0.5% glucose

to ensure that the FimS remained uninverted. A 100X (50%) glucose stock solution was

made from 0.5 g D-glucose in 1 mL of sterile water. The solution was heated to 70 °C to

dissolve the glucose, and then stored in the dark at room temperature for up to one week.

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4.4 Photochemical Control over Flpe-mediated DNA Recombination in E. coli

4.4.1 Introduction to Flpe Recombinase

As we are interested in obtaining photochemical control over recombination in E. coli, we are working to develop a tandem in vivo system that enables expression of a photocaged recombinase in a cell harboring a reporter plasmid. Light irradiation will activate recombination and lead to expression of the reporter gene. We thus sought to achieve photochemical control over DNA recombination via photocaging another member of the integrase family, the 48.6 kDa Flippase recombinase (Flpe) of Saccharomyces cerevisiae,[248] since it shows activity in bacterial cells. Flpe is naturally encoded on the yeast 2µ plasmid and is responsible for maintaining the correct copy number of the 2µ plasmid within the cell, by recombining two plasmids into one and vice versa.[249, 250] Like

Cre and FimE, Flpe catalyzes DNA cleavage and strand exchange by an essential tyrosine within the conserved recombinase “R-H-R-Y” catalytic tetrad of Arg191, His305, Arg308, and Tyr343.[251] This recombination occurs between two palindromic 34 bp inverted repeats, termed frt (Flippase recognition target) sites that include an eight base-pair asymmetric core that confers directionality (Figure 4.28).[252]

108

Figure 4.28. Flpe recombinase catalyzes the inversion or excision/integration of DNA between two frt sites. Orientation of the 34bp loxP sites determines whether the intervening DNA will be inverted or removed/integrated. The frt site is composed of two 13 bp inverted repeat elements to which the Cre protein binds and an eight bp spacer region (blue) separating them, where the DNA breakage and reunion occurs.

Flpe recombinase has an identical mechanism to Cre recombinase, in which a tyrosine nucleophile catalyzes sequential strand exchanges.[253] This exchange proceeds through the formation of a Holliday junction between the covalent DNA-protein intermediates.[254, 255][255] Resolution of the final nucleoprotein structure completes the recombination event and results in two DNA duplexes with exchanged DNA sequences.

Flpe recombination has been utilized in bacteria[256, 257] (in contrast to Cre recombinase) and in mammalian systems,[258-261] and has found a particular niche in plant biotechnology.[262-265] However, as with other site-specific recombinases (see previous sections), precise external control of Flpe activity cannot be achieved. However, light- activation as a means to obtain robust spatio-temporal control over DNA recombination would increase the applicability and effectiveness of Flpe in plant and animal systems. We

109 therefore sought to regulate Flpe activity through the installation of a photocaging group directly in its active site. Thus, DNA recombination would not occur until brief UV irradiation removed the caging group and restored activity.

4.4.2 Expression of Wild Type Flpe

The Flpe gene was PCR amplified from Saccharomyces cerevisae and inserted into the pET-21a vector for T7-mediated and 6xHis tagged bacterial expression, creating pET21-

Flpe. The expected vector sequence was confirmed by DNA sequencing. E. coli BL21(DE3)

Gold cells were transformed with pET21-Flpe and cultures were induced with 1 mM IPTG when an OD600 of 0.6 was reached, then expressed for four hours at 37 °C. The resulting Flpe protein was purified with Ni-NTA resin and analyzed by 10% SDS-PAGE (Figure 4.29).

Flpe

58 46

Figure 4.29. Expression of Flpe recombinase from pET-21Flpe yields pure protein, as determined by SDS- PAGE analysis.

4.4.3 The Flpe Bacterial Assay

The Voziyanov Lab previously developed a novel Flpe reporter system for use with

E. coli that consists of a pBAD33-Flpe expression plasmid and a pBUI reporter plasmid that encodes the α-LacZ gene located between two frt sites (Figure 4.30).[266, 267] In this system, cells produce the alpha fragment of β-galactosidase. This fragment and the omega

110

fragment, which must be expressed by the host cell, reassemble in vivo and subsequently hydrolye X-gal, as indicated by a blue colony color. Flpe-mediated recombination excises the

LacZ gene from pBUI, inhibiting production of galactosidase; the bacteria can no longer hydrolyze X-gal, resulting in a white colony color (Figure 4.30). Thus, transformation of both plasmids into E. coli Top-10 cells provides an arabinose-inducible reporter system to visually determine Flpe-mediated DNA recombination. Additionally, in vitro recombination can be assayed in the same manner as the activity of Cre recombinase (see 4.2.2 using purified Flpe and pBUI in Flpe reaction buffer for 30 min at 37 °C. An aliquot can then be transformed into E. coli, and the resulting colonies inspected for a white color, indicative of

Flpe-mediated β-galactosidase excision.

- Flpe +Flpe

Figure 4.30. The Flpe- mediated DNA recombination reporter assay. Introduction of Flpe either in vitro or through arabinose-induced expression in vivo catalyzes recombination of the pBUI substrate, which encodes the

LacZ gene (blue rectangle) between two frt sites (purple arrows). In the absence of Flpe, colonies hydrolyze X- gal to produce a blue color. Flpe-mediated recombination leads to white colonies no longer able to hydrolyze X- gal. Figure adapted from Ma, C. H.; Kwiatek, A.; Bolusani, S.; Voziyanov, Y.; and Jayaram, M. J. Mol. Biol. 2007, 368, 183.

111

We obtained pBAD33-Flpe and pBUI from the Voziyanov Lab. Our expressed Flpe was first assayed for activity in vitro with pBUI in Flpe reaction buffer for 30 min at 37 °C.

Top-10 cells were transformed with 1 µL of the 30 µL reaction, then plated on LB agar containing Amp and with X-gal spread on top. A control recombination reaction was

performed without Flpe and was also transformed into E. coli. After 24 hours at 37 °C, the

control reaction exhibited over 300 blue colonies, with no white colonies identified. In contrast, the Flpe assay yielded over 300 colonies, but with over 150 white colonies. An equal amount of white and blue colonies was expected, as recombination has been shown to reach an equilibrium between excised and inserted DNA. We thus demonstrated that we could successfully utilize pBUI as a reporter plasmid for Flpe-mediated recombination in vitro.

We next sought to validate the Flpe in vivo assay in order to test the light-activation of caged Flpe expressed in bacterial cells. Top-10 cells were co-transformed with pBAD33-

Flpe and pBUI, then plated on X-gal with and without arabinose. As expected, in the absence of arabinose, no Flpe expression occured and only blue colonies were observed. However, in the presence of arabinose over 300 white colonies and only five blue colonies were observed.

With successful in vitro and in vivo assays for Flpe activity, we next sought to express

FlpeONBY from pBAD33-Flpe. We therefore used QuikChange mutagenesis to mutate the catalytic Tyr343 to the TAG amber stop codon in Flpe in both pET21-Flpe and pBAD33-

Flpe. DNA sequencing verified the expected result for both plasmids. BL21(DE3) Gold cells were transformed with pET21-Flpe343TAG and pSupONBY. A 25 mL expression was

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performed in LB medium containing 1 mM ONBY. Cells were induced with 1 mM IPTG at

an OD600 of 0.6, then grown four hours at 37 °C followed by 15 hours at room temperature.

Cells were then pelleted and the protein was purified. However, SDS-PAGE analysis showed

that no FlpeONBY was produced, while the wild type Flpe control was expressed.

Expressing FlpeONBY from pBAD33-Flpe was not attempted since both pBAD33-

Flpe343TAG and pSupONBY harbor the same chloamphenicol resistence marker.

4.4.4 Summary and Outlook

We have begun developing a photoinducible Flpe system for use in E. coli to achieve

light-control over DNA recombination. As expression from pSupONBY did not produce

photocaged Flpe, expression from our modified pEVOL should be investigated. However,

the chloramphenicol resistence marker must be altered in either pEVOL or pET21-Flpe to

enable propagation of both plasmids in the cell. The photocaged Flpe can then be both expressed and assayed in vivo without the additional steps of protein isolation and purification. Importantly, in vivo photoregulation of Flpe recombinase has the potential to provide spatial and temporal control over DNA recombination in E. coli. In contrast to FimE, the Flpe recombinase could also be used in cell culture without the need for additional cofactors. This system thus provides another molecular biology tool for the spatiotemporal control of DNA recombination with light.

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4.4.5 Experimental Methods

pET21-Flpe Vector Construction. The 1400 bp Flpe gene was PCR amplified directly from

S. cerevisiae cells added to the PCR (95 °C (2 min), then 30 cycles of 95 °C (30 sec), 56 °C

(30 sec) and 72 °C (1.5 min)) with the primers 5’-

TGTTCGCTAGCATGCCACAATTTGGTATAT-3’ and 5’-

TGGTGCTCGAGTATGCGTCTATTTATGTAG-3’ which contain NheI and XhoI restriction sites, respectively. The Flpe gene was then inserted into pET-21a(+) (Novagen) as an NheI-XhoI fragment, generating pET21-Flpe. The expected vector sequence was confirmed by DNA sequencing using the primers 5’-TAATACGACTCACTATAGGG-3’ and 5’-AAGGGGTAGGATCGATCCA-3’ which together provided complete sequence coverage of the gene, from upstream of the start codon to downstream of the 6xHis tag.

Expression and Purification of Flpe from pET21-Flpe. E. coli BL21(DE3) Gold cells were transformed with pET21-Flpe, inoculated 1:100 in 25 mL LB medium containing 50

µg/mL ampicillin, and were grown at 37 °C, 250 rpm. At an OD600 of 0.6, expression was induced with IPTG (25 µL of a 1 M stock solution) to a final concentration of 1 mM IPTG.

Cells were grown for six hours, then pelleted, lysed, and purified with Ni-NTA resin

according to the manufacturer’s instructions. Briefly, 1 mL of Qiagen lysis buffer and 25 mg

of lysozyme were added to the cell pellet resulting from a 25 mL culture (~ 250 µL), and

cells were resuspended and incubated one hour on ice. Cells were vortexed and sonicated in a

VWR sonication bath, model 150D (six cycles of 15 sec of sonication followed by 15 sec on

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ice). Cellular debris was pelleted in an Eppendorf refrigerated centrifuge at 4 °C (13,200

rpm, 20 min), then 50 µL of Ni-NTA resin was added to approximately 1 mL of cleared

lysate. The resin was placed on a rocker in a 4 °C cold room for one hour, then washed four

times with 1 mL of Qiagen wash buffer and eluted twice in 100 µL each of Qiagen elution

buffer. Protein production and purity was verified by 12% SDS-PAGE analysis to identify

the 48 kDa protein. Samples were run 12 mA to allow the loading dye to clear the stacking

gel (about 15-20 min), then electrophoresed at 24 mA for 45-60 min or until the prestained

protein marker (NEB) was adequately separated. Finally, Flpe was dialyzed into storage

buffer (20 mM Tris-HCl, pH 7.8, 300 mM NaCl, 1 mM EDTA, and 50% glycerol), and

stored at 20 °C. A Bradford Assay was performed to determine protein concentration (135

ng/µL).

In Vitro Flpe Activity Assays. The pBUI (Amp resistant) substrate plasmid used for in vitro

and in vivo activity assays with E. coli was obtained from the Voziyanov Lab. In vitro Flpe

recombination assays were performed for 30 minutes at 37 °C in recombination buffer (50

mM Tris-HCl, 33 mM NaCl, 10 mM MgCl2, pH 7.5) with 200 ng pBUI substrate and 100 ng

Flpe expressed from pET21-Flpe (30 µL total reaction volume). Flpe was heat inactivated at

70 °C for five minutes, followed by transformation into ultracompetent Top-10 cells

following the manufacturer’s protocol using 25 µL of cells recovered in 100 µL LB medium.

To prepare the selective growth plates, 25 mL of LB agar containing 30 µg/µL

chloramphenicol was poured into a 10 × 1.5 cm petri dish and the agar allowed to harden

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(about 15 min). The surface of the agar was then spread with 40 µL X-gal dissolved in DMF

(see Appendix 1 for recipe) and allowed to dry (approximately 5 min). All of the recovered

cells were then plated and incubated over night at 37 ºC.. Successful in vitro recombination

resulted in ~50 % white colonies (about 480 white colonies to 525 blue colonies), while the

control reaction containing no Flpe and only pBUI exclusively yielded blue colonies (about

1000 colonies each).

In Vivo Flpe Activity Assays. The pBAD33-Flpe plasmid (Cam resistant) was obtained

from the Voziyanov Lab (Louisiana Tech University). As this plasmid does not contain a

6xHis tag, it was used only with in vivo assays. Top-10 cells (25 µL) were co-transformed

with pBAD33-Flpe and pBUI, recovered in 250 µL SOC medium, then plated on LB agar

containing 0.2% arabinose and both 50 µg/µL ampicillin (for pBUI propagation) and 30

µg/µL chloramphenicol (for pBAD33-Flpe propagation). Control plates containing no

arabinose were also prepared. Arabinose is necessary to induce Flpe expression and thus 250

µL were added to 25 mL of molten LB agar during preparation. To prepare the selective

growth plates, 25 mL of LB agar containing 30 µg/µL chloramphenicol was poured into a 10

× 1.5 cm petri dish and the agar allowed to harden (about 15 min). The surface of the agar was then spread with 40 µL X-gal dissolved in DMF (see Appendix 1 for recipe) and allowed to dry (approximately 5 min). 75 µL of cells were plated, then incubated ~18 hours at 37 °C, then placed at 4 °C to better develop the blue color of recombination deficient colonies.

White colonies were counted to determine recombination efficiency, with typically all

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colonies white. Control plates were also visually inspected to ensure that all colonies were

blue.

Unnatural Amino Acid Mutagenesis. Following the manufacturer’s protocol (Strategene),

QuikChange mutagenesis was employed to mutate Tyr343 into an amber stop codon

(TAT→TAG) with primers 5’-GCCAGGACAACGTAGACTCATCAGATAACAGC-3’ and

5’-GCTGTTATCTGATGAGTCTACGTTGTCCTGGC-3’ in pET21-Flpe, generating

pET21-FlpeY343TAG. Successful amber stop codon mutation was confirmed by DNA

sequencing with the primer 5’-AAGGGGTAGGATCGATCCA-3’, which provided coverage

from the N-terminus to ~50 bp downstream of the TAG mutation.

Flpe-ONBY Expression and Purification with pSupONBY. To produce caged Flpe,

BL21(DE3) Gold cells were transformed with both pET21-FlpeY343TAG and pSupONBY,

then grown in LB medium containing 50 µg/mL ampicillin, 30 µg/mL chloramphenicol, and

enriched with 1 mM ONBY. A 100X ONBY stock solution (32 mg) was made up in 50 µL

hot DMSO, and then hot water was added to achieve a total volume of 1 mL. The appropriate

amount of ONBY was added to 37 °C pre-warmed LB medium (25 mL) dropwise with intense swirling to achieve a 1X concentration (1 mM); if necessary, the pH of the medium was then adjusted to pH 7.0 prior to inoculating the overnight culture 1:100. A control expression was also performed without ONBY to verify that no protein was produced in the absence of the unnatural amino acid, and to allow determination of the correct optical density

117 at which to induce protein expression without interference from the ONBY precipitate. The

OD600 of the control culture was determined spectroscopically, and cells were again induced at an OD600 of 0.6 after approximately 4 hours. The protein was expressed five hours at 37

ºC, then 15 hours at room temperature. Purification was performed as described above.

However, no photocaged protein was detected by 12% SDS-PAGE analysis. Unfortunately, repeated attempts to express FlpeONBY from pET21-FlpeY343TAG and pSupONBY have been unsuccessful to date.

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CHAPTER FIVE – LIGHT-REGULATION OF PROTEIN DIMERIZATION

5.1. Introduction to Natural and Synthetic Protein Dimerizers

Protein-protein interactions are integral to cellular function and signalling, and within the cell form the basis of the interactome, which encompasses all of the cell’s protein-protein interactions..[268, 269] Protein dimerization is involved in the regulation of enzymes, ion

channels, surface receptors, and transcription factors, whereby monomeric poteins undergo

non-covalent, reversible interactions to ellicit a specific cellular response.[270] Additionally,

it has been shown that dimerizing two monomeric proteins within the cell can be of

therapeutic importance, as studies of the immunosuppresants FK506 and rapamycin

demonstrate that their modes of action entail the dimerization of two proteins that would not

otherwise interact.[271, 272] Synthetic methods to induce or control dimerization are

important to studying gene function and modulating enzyme activity, and have lead to the

development of [273]chemical inducers of dimerization (CIDs).[274] CIDs are small

molecules comprised of two end moieties that interact with a protein target and are tethered

by a non-reactive linker; they elicit a particular cellular response by dimerizing two

monomeric proteins or two split protein domains that have an affinity for opposite ends of

the dimerizer (Figure 5.1).[275]

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Protein (or domain) CID-binding Two domain

Protein (or domain) One

Figure 5.1. CID-mediated protein dimerization enables the bridging of two proteins or two split protein domains. Adapted from Pollock, R.; and Clackson, T. Curr. Opin. Biotechnol. 2002, 13, 459.

Several CIDs have been applied to modulate protein-protein interactions in vivo,[276-

281] and many of these systems utilize CID-mediated transcription, where successful dimerization is visualized by subsequent expression of a reporter gene.[282-284] CID- induced gene expression is based on the yeast two-hybrid model developed by S. Fields and

O. Song to detect protein-protein interactions.[285] In the yeast Saccharomyces cerevisiae, the transcriptional activator Gal4 is encoded by both an N-terminal domain that binds to a specific upstream DNA activation sequence (UAS), and a C-terminal domain that activates transcription by recruiting the necessary protein components; expression of genes related to galactose metabolism occurs only when both domains interact.[285] To probe unknown protein interactions, the proteins of interest are expressed as hybrids, with one fused to the N- terminal DNA binding domain (DBD) and the other fused to the C-terminal transcription activating domain (TA) of Gal4. Proteins that dimerize reconstitute the Gal4 activation

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complex and enable transcription of a reporter gene (Figure 5.2, A). Transcription is stopped,

however, if the proteins do not interact (Figure 5.2, B).

C. D.

Figure 5.2. A-B. The yeast two-hybrid system. A. Transcription occurs when dimerization of two proteins (orange, purple) brings the N-terminal (red) and C-terminal (green) domains of GAL4 in proximity. B. The non- interacting proteins (orange, blue) cannot reconstitute GAL4, and no transcription occurs. C-D. The yeast three- hybrid system. C. Dimerization of proteins fused to either the GAL4 N- or C- termini occurs through the use of a CID. Successful CID-mediated dimerization leads to transcription. D. If one of the proteins cannot bind the CID, GAL4 is not reconstituted, and transcription does not occur.

The yeast-two hybrid system utilizes the natural dimerization of interacting proteins

to enable transcription. As such, transcription through CID-mediated protein dimerization

between two otherwise non-interacting proteins or protein domains has been termed a yeast

three-hybrid system[283, 286-288] (Figure 5.2, C-D). The Schreiber group developed a yeast

three-hybrid system in mammalian cells based on the binding of the cytosolic protein FKBP

to the previously mentioned immunosuppresant rapamycin.[283] Rapamycin also binds to

the phosphoinositide 3-kinase homolog FRAP (rapamycin-associated protein; also known as

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mTOR, or mammalian target of rapamycin). FKBP and FRAP have no natural interaction;

however, upon introduction of rapamycin into the cell, dimerization of FKBP and FRAP

occurs. Schreiber and his colleagues fused three copies of FKBP to the DBD of Gal4, and a

mutated FRAP binding domain termed called FRB to the Gal4 TA. Addition of rapamycin

dimerized both proteins at one of three FKBP fusions and activated transcription (Figure 5.3)

Figure 5.3. Rapamycin-induced dimerization of FRB and FKBP in a yeast three-hybrid system leads to expression of a reporter gene. Three FKBP genes were fused to the DBD of Gal4, which binds to the UAS sequence. The FRB gene was fused to the Gal4 TA (brown rectangle). Dimerization of FRB and FKBP led to successful transcription. From Liberles, S. D.; Diver, S. T.; Austin, D. J.; and Schreiber, S. L. Proc. Natl. Acad. Sci. 1997, 94, 7825.

Several CIDs have since been based on rapamycin.[289-291] In addition, the Cornish

group developed a novel CID for use in a yeast three-hybrid system based on the affinity of

the enzyme dihydrofolate reductase (DHFR) for the antibiotic methotrexate (MTX), and the

affinity of a glucocorticoid receptor for dexamethasone (DEX).[292] DHFR is a ubiquitous

24 kDa protein that catalyzes the reduction of dihydrofolate to tetrahydrofolate, the essential

building block for thymine biosynthesis in both eukaryotic and prokaryotic organisms.[293]

DHFR was fused to the DBD of the transcription factor LexA, while a rat glucocorticoid

122 receptor (GR) was fused to the TA domain of the B42 protein (Figure 5.4, A). The DEX-

MTX heterodimer was added to the yeast growth medium and a lacZ reporter gene was transcribed, producing β-galactosidase which was detected through conversion of X-gal into a blue pigment (Figure 5.4, B).

Figure 5.4. Use of the yeast three-hybrid system with a novel DEX-MTX CID leads to expression of β- galactosidase and the development of a blue color upon successful dimerization. A. DHFR fused to the DBD of LexA dimerizes in the presence of the DEX-MTX CID with the GR-B42 TA fusion to express the reporter gene. B. Successful dimerization produces blue colonies. Columns 1 and 2 contain yeast expressing known protein interactions; yeast in column 3 express both the LexA-DHFR and B42-GR fusions; yeast in column 4 express LexA-DHFR and B42; yeast in column 5 express LexA and B42-GR; and yeast in column 6 express LexA and B42. Cells were plated on 1 µM CID. From Lin, H.; Abida, W. M.; Sauer, R. T.; and Cornish, V. W. J. Am. Chem. Soc. 2000, 122, 4247.

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More recently, the Cornish group has developed an alternative CID to label cells in vivo that is based on the affinity of DHFR for the antibiotic trimethoprim (TMP) (Figure

5.5).[294] TMP was utilized in place of MTX to provide a stronger DHFR-dimerizer interaction, and to eliminate background from the interaction between endogenous DHFR and the MTX-based dimerizer; only E. coli DHFR, and not human DHFR, interacts with

5 TMP. Interestingly, DHFR has a KI of 1.3 nM for inhibition by TMP,[295] which binds 10

more tightly to E. coli DHFR than to eukaryotic DHFR.[296]

Figure 5.5. The antibiotic trimethoprim.

DHFR was thus fused to cyan fluorescent protein (CFP), and the use of TMP over

MTX prevented background fluorescence by inhibiting endogenous DHFR from binding the

new fluorescently labeled TMP. TMP was chemically labeled with the fluorescent dye Texas

red (BTR); as a result, only one protein interaction was required to affect successful labeling,

enabling the fluorescent TMP ligand to function as a conjugate tag. Expression vectors that

targeted E. coli DHFR to either the cell membrane or the nucleus were tranfected into CHO

cells, and after 24 hours, the TMP-BTR ligand was added to the medium (Figure 5.6).

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Figure 5.6. Selective chemical labeled of subcellularly targeted E. coli DHFR in CHO cells. A-C. Confocal micrographs: left column, excitation at 458 nm; middle column, excitation at 568 nm; right column, differential image contrast. A. CHO cells transfected with DNA encoding membrane-targeted DHFR-CFP. Cells were then incubated with 10 nM TMP-BTR and images were taken after 5 min. Both blue and red fluorescence are co- localized to the plasma membrane, indicating successful conjugation. B. CHO cells transfected with DNA encoding nucleus-targeted DHFR-CFP. Cells were then incubated with 10 nM TMP-BTR and images were taken after 5 min. Both blue and red fluorescence are co-localized to the nucleus, indicating successful conjugation. C. CHO cells tranfected as in B., then incubated with 1 µM TMP for 15 min, followed by 10 nM TMP-BTR. No red fluorescence can be seen in the nucleus, as the binding sites of DHFR-CFP were blocked with TMP. From Miller, L. W.; Cai, Y.; Sheetz, M. P.; and Cornish, V. W. Nat. Methods 2005, 2, 255.

The Cornish group then demonstrated that a MTX-conjugate could be used to fluorescently label proteins in vivo and image them in real time. Due to the high affinity of

DHFR for TMP, the labelling is practically irreversible.

More recently, the Cornish group synthesized TMP-SLF, a CID capable of dimerizing DHFR with FKBP, and utilized it to activate transcription of the lacZ reporter gene in a yeast three-hybrid system (Figure 5.7).[297] SLF is a cell-permeable small

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molecule that has no immunosuppressive activity and binds FKBP with nanomolar

affinity.[298] The DBD of LexA was fused to the N-terminus of DHFR, while the B-42 TA domain was fused to the C-terminus of FKBP. Cells expressing both fusion proteins were grown for three days on X-gal enriched medium with or without 1 µM TMP-SLF. Only cells grown in the presence of the CID produced a blue color, indicating that TMP-SLF was capable of dimerizing the DBD and TA fusions to reconstitute transcription.

Figure 5.7. Yeast three-hybrid assay of TMP-SLF activity. A. TMP-SLF heterodimerizes the LexA DNA binding domain-DHFR fusion protein and the FKBP-transcriptional activator fusion protein. Dimerization thus reconstitutes the transcriptional machinery and β-galactosidase is expressed. B. Only yeast grown on X-gal and treated with 1 µM TMP-SLF produce a blue color, indicating successful dimerization-mediated transcription. From Czlapinksi, J. L.; Schelle, M. W.; Miller, L. W.; Laughlin, S. T.; Kohler, J. J.; Cornish, V. W.; Bertozzi, C. R. J. Am. Chem. Soc. 2008, 130, 13186.

Furthermore, TMP-SLF was utilized in mammalian cell culture to activate

fucosyltransferase VII (FucT7), a glycosyltransferase localized in the Golgi apparatus. FucT7 plays a critical role in the immune system by producing the sialyl Lewis x (sLex) epitope; to

regulate it, a biocompatible, non-immunosuppresive CID is required, such as TMP-SLF.

FucT7 is comprised of a catalytic domain (Cat) and a localization domain (Loc), and both are

required for cellular function.[299, 300] As such, the FucT7 Cat domain was fused to the

DHFR gene, while the Loc domain was fused to FKBP. These fusion proteins were

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expressed in CHO cells, which lack endogenous FucT7 activity, in the presence of absence of

1 µM TMP-SLF. Only cells grown in the presence of TMP-SLF exhibited glycosyltransferase activity, as evidenced by the presence of cell-surface sLex that was

detected immunohistochemically with a fluoresecent antibody (Figure 5), indicating

successful in vivo dimerization.

Figure 5.8. Activation of FucT7 by TMP-SLF as determined by fluorescent anti-sLex. The mean fluorescent intensity was measure for all live cells. Open bars: vehicle-treated cells lacking TMP-SLF; filled bars: cells treated with 1 µM TMP-SLF.

We thus sought to adopt a CID-mediated cellular labelling strategy by designing a light-regulated CID with a photoreversibly switchable diazobenzene (DAB) core. Thus, we could utilize light irradiation to switch the conformation of the CID and affect the ability of the target proteins to dimerize. In this manner we could provide the first example of photoreversible protein dimerization in mammalian cell culture.

For the synthesis of the CID, a DAB core was modified with two TMP moieties, as shown in Figure 5.9. As approximately 95% of the DAB moiety exists under ambient light in

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the trans configuration, the Bis-TMP CID should provide a reasonably linear ligand to bind

two molecules of DHFR. Upon irradiation with 365 nm of light, the majority of the Bis-TMP

would switch to the cis configuration, whereupon one DHFR molecule would be displaced due to steric interactions. The Bis-TMP would then revert over time or through irradiation with UV light of 420 nm to the trans configuration and the displaced DHFR molecule could once again bind (Figure 5.9)

Figure 5.9. The novel light-switchable Bis-TMP CID constructs, containing a photoresponsive DAB core flanked by two TMP molecules. Bis-TMP exists in the trans configuration and can bind two molecules of DHFR. Irradiation at 365 nm switches the DAB core to the cis configuration, displacing one of the DHFR molecules. Bis-TMP1: n = 1; Bis-TMP2: n = 2; Bis-TMP3: n = 3, Bis-TMP4: n = 4.

5.2 Expression and Purification of DHFR and In Vitro Dimerization Assays

The wild-type DHFR gene was PCR amplified from E. coli genomic DNA and

ligated into the pET-21a expression vector, expressing DHFR under control of T7 RNA

polymerase and encoding a 6xHis tag, thus creating pET21-DHFR. The expected gene sequence was confirmed by DNA sequencing. BL21(DE3) Gold E. coli cells were transformed with pET21-DHFR; cultures were induced with 1 mM IPTG when an OD600 of

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0.6 was reached, and expressed for four hours at 37 °C. The resulting 6xHis-tagged protein was purified by Ni-NTA metal affinity chromatography, using Ni-NTA resin (Qiagen) and

verified by SDS-PAGE analysis (Figure 5.10). Stringent washing of the resin with increasing

concentrations of imidazole yielded the protein with up to 95% homogeneity. DHFR was

then dialyzed into storage buffer and protein concentration was determined by a Bradford

Assay to be 500 ng/µL, with a total protein yield of 10 mg/L culture.

Figure 5.10. Expression of wild-type E. coli DHFR (wtDHFR) from pET21-DHFR in BL21(DE3) Gold cells yields high purity protein, as determined by 12% SDS-PAGE analysis.

DHFR was then assayed for activity using a commerically available MTX resin (Sigma).

Dialyzed DHFR was incubated with the MTX resin for one hour at room temperature in DHFR

activity buffer. The resin was then washed until no DHFR was present in the flow-through, then 1 mM TMP was added. As E. coli DHFR has a higher affinity for TMP than MTX, DHFR was eluted from the resin and thus determined to be active in binding TMP (Figure 5.11). This activity assay was also utilized with DHFR fusions farther along in the project to verify activity prior to any CID- mediated dimerization assays. Alternatively, enzymatic assays for DHFR activity could also be performed.[301, 302]

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•M 1 2 3 4 5

Figure 5.11. SDS-PAGE analysis of the DHFR activity assay using an MTX-resin. M: protein marker; lane 1: DHFR flow-through; lanes 2-4: washes; lane 5: TMP elution.

We next designed an in vitro assay to investigate the light-responsive dimerization

potential of the Bis-TMP1 CID (Figure 5.12). Excess 6xHis-tagged DHFR was immobilized

on the Ni-NTA resin to ensure that all coordination sites on the resin were occupied. The

resin was then washed to remove the excess protein (7.5 µg / µL resin), and incubated with

Bis-TMP for one hour at room temperature in DHFR reaction buffer with gentle shaking to ensure that the resin would not settle. The resin was then washed and an additional excess of

DHFR (7.5 µg / µL resin) was added and incubated as before. The resin was again washed and then split into two 0.2 mL PCR tubes. One tube was irradiated 30 min on a

transilluminator (365 nm, 25 W) prechilled to 0-4 °C with an icepack, and the flow-through

was removed. The flow-through from the non-irradiated tube was also removed, and both samples were analyzed on a 12% SDS-PAGE gel along with aliquots of the previous DHFR additions and washes.

130

Figure 5.12. The in vitro DHFR dimerization assay. Resin-immobilized DHFR is incubated with Bis-TMP, then a second aliquot of DHFR is added and incubated with the resin. Upon irradiation at 365 nm of light, the Bis-TMP switches to the cis configuration, displacing the non-immobilized DHFR. The Bis-TMP CID then thermally returns to the trans configuration over time. Theoretically, this cycle is reversible if performed without removing any supernatant, as displaced DHFR can then bind again to the Bis-TMP.

Gratifyingly, SDS-PAGE analysis showed the utility of our assay, and that Bis-TMP1

was capable of dimerizing two molecules of DHFR, as a DHFR band was present in the

irradiated sample, and not in the non-irradiated sample (Figure 5.13). DHFR bands were also

present in the lanes corresponding to protein addition and the first washes. No DHFR bands

were present in subsequent washes, showing that the eluted protein was not from any

remaining DHFR in the reaction. However, while supporting the hypothesis shown in Figure

5.12, this assay was not reproducible.

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Figure 5.13. SDS-PAGE analysis of resin-immobilized DHFR incubated with Bis-TMP1 and irradiated 30 min (365 nm, 25 W). DHFR: Aliquots of DHFR added to the resin; Wash: the third wash after each addition of DHFR; –UV: flow-through collected from the non-irradiated reaction contains no DHFR; +UV: flow-through collected from the irradiated reaction contains DHFR.

In an effort to optimize the in vitro assay and obtain reproducible results, we assayed

three additional dimerizers: Bis-TMP2, Bis-TMP3 and Bis-TMP4 (Figure 5.9), which

contain 4, 6 or 8 more carbons than Bis-TMP1, respectively. We thought that the increased

linker length could potentially affect dimerization. Assays were performed as described

above with Bis-TMP1, and aliquots were subsequently analyzed with SDS-PAGE. However, there appeared to be no difference in light-elution of DHFR amongst all four dimerizers; thus, experiments were continued with Bis-TMP1.

5.3 Expression and Purification of GFP-DHFR and In Vitro Fluorescent Dimerization

Assays

We next designed a GFP-DHFR fusion protein for visualization of dimerization both in vitro and in mammalian cell culture. The DHFR gene from pET21-DHFR was cloned downstream of GFP in the vector pGFPuv. The GFP-DHFR fusion was then cloned into pBAD-mycHis to create the vector pBAD-GFP-DHFR. Additionally, a TEV cleavage site

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was inserted upstream of the His tag to facilitate its removal after purification with TEV

protease. Removal of the His tag ensures that addition of GFP-DHFR to the reaction will not

displace or compete with the immobilized Ni2+-coordinated DHFR. The expected vector sequence was verified by DNA sequencing. BL21(DE3) Gold cells were transformed with pBAD-GFP-DHFR; cultures were induced with 1 mM IPTG when an OD600 of 0.6 was

reached, and expressed for four hours at 37 °C. The resulting 6xHis-tagged 52 kDa GFP-

DHFR was again purified with a Ni-NTA resin and analyzed by SDS-PAGE analysis (Figure

5.14). GFP-DHFR was then dialyzed into DHFR storage buffer and protein concentration was determined by a Bradford Assay to be 300 ng/µL, with a total protein yield of 9.6 mg/L

culture. The fluorescence of GFP-DHFR was confirmed visually using a handheld UV lamp

(365 nm, 25 W) in a 1.5 mL eppendorf tube.

GFP-DHFR

175 83 54 48

Figure 5.14. Expression of GFP-DHFR from pBAD+GFP-DHFR in BL21(DE3) Gold cells yields high purity protein, as determined by 12% SDS-PAGE analysis.

Due to the fluorescence of GFP, results of the in vitro assay could be determined

spectroscopically with a UV plate reader. Thus, DHFR was again immobilized on the Ni-

NTA resin, then incubated with Bis-TMP. GFP-DHFR was added to the reaction and

incubated as before. The resin was again split, with one aliquot irradiated (30 min, 365 nm,

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25 W) on the pre-chilled transilluminator. Additionally, 1 mM TMP was added to both the

irradiated and non-irradiated reactions to remove DHFR from the resin. We expected a high amount of fluorescence in the aliquots corresponding to addition of GFP-DHFR and UV

irradiation. However, while fluorescence levels were high for these conditions, fluorescence

was also high in the non-irradiated sample (Figure 5.15). We also added 1 mM TMP to the

irradiated and non-irradiated reactions. The addition of TMP should have removed any

remaining dimer from the resin; thus, we expected fluorescence to be high in the non-

irradiated sample, as the dimer would have been intact. However, we saw nearly identical

fluorescence when TMP was added the non-irradiated sample as when TMP was added to the

irradiated sample (Figure 5.15). Upon repeating the assay, similar fluorescent readings were

obtained, indicating that some of the DHFR was successfully eluted upon irradiation of Bis-

TMP, but a substantial portion remained bound to the resin.

134

300 254.725 237.974 250

200

150 118.1525

100

Relative Fluorescent Units 23.6505 14.272 14.8595 15.698 20.274

Figure 5.15. Fluorescence levels in the in vitro DHFR assay using both DHFR and GFP-DHFR. +UV: irradiated resin (365 nm, 25 W); –UV: non-irradiated resin; +UV +TMP: 1 mM TMP added to irradiated resin; –UV +TMP: 1 mM TMP added to non-irradiated resin. Fluorescence was high upon addition of GFP-DHFR, and in the irradiated and non-irradiated samples. Relative fluorescence values are written above each sample.

As our initial in vitro assays using wild type DHFR displayed more promising results

than the in vitro fluorescent assays, and thus we sought alternative analysis methods to

quantify possible Bis-TMP1 CID-mediated dimerization. The Wagner group at the

University of Minnesota has developed a novel way to synthesize homogenous protein

polygons, termed nanorings, using two DHFR molecules fused together by a peptide

chain.[303, 304] They found that the linked DHFR molecules would spontaneously form

macrocycles upon treatment with the dimerizer Bis-MTX-C9 (two MTX molecules linked by

a nine-carbon chain)[303] (Figure 5.16). Subsequent nanoring formation could be analyzed by size exclusing chromatography (SEC).

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O OH HO O O O H H N N NH2 N N NH2 H H N O O N N N N N

H2N N N N N NH2

Figure 5.16. Structure of Bis-MTX-C9 and a schematic depiction of the dimerized nanoring formed when two peptide-linked DHFR molecules (blue) are incubated with Bis-MTX-C9 (green). Image provided by Dr. Adrian Fegan (Wagner Lab).

In collaboration with Dr. Adrian Fegan (Wagner Lab), DHFR dimerization and nanoring formation in the presence of our Bis-TMP1 CID or their Bis-MTX-C9 control were analyzed by SEC. Incubation of 7.5 µM Bis-MTX- C9 with tethered DHFR (two protein molecules linked by 13 amino acids) enables the formation of trimer or dimer, as well as

internal monomer, when the dimerizer binds only one tethered DHFR (Figure 5.17).

mAU Internal monomer 10

0 2 4 6 8 10 12 14 16 18 20 22 24 26 28 3 Minutes Figure 5.17. SEC analysis of trimer (left peak), dimer (middle peak) and internal monomer (right peak) formed upon incubation of tethered DHFR with Bis-MTX-C9. Image provided by Dr. Adrian Fegan (Wagner Lab).

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Assays were performed with 2.5-45 µM dimerizer and 2.5 µM monomeric DHFR or tethered DHFR, then analyzed by SEC. Initial results showed that the Bis-TMP1 was not forming a stable enough dimer with the monomeric DHFR to visualize by SEC, due to the peaks eluting at the same time (Figure 5.18, A). Dimerization in the presence of the tethered

DHFR was then investigated; however, no dimer formation was detected with up to 45 µM

Bis-TMP1 (Figure 5.18, B).

A. 20

15 mAU 10

0 5 10 15 20 25 30 35 Minutes B. 20

15 mAU 10

0 5 10 15 20 25 30 35 Minutes

Figure 5.18. SEC analysis of Bis-TMP1 induced DHFR dimerization. A. Overlay of incubation with monomeric DHFR and 0 µM, (black line) 2.5 µM, (green line) and 7 µM (blue line) Bis-TMP1. No change in retention time is detected, indicating little to no dimer formation. B. Overlay of incubation with tethered DHFR and 0 µM, (black line) 7.5 µM, (green line) 15 µM, (blue line) 22.5 µM, (pink line) and 45 µM (red line) Bis- TMP1. As in A, no change in retention time is detected, indicating little to no dimer formation. Images provided by Dr. Adrian Fegan (Wagner Lab).

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The second dimerization experiment was repeated with Bis-TMP and tethered DHFR

in the presence of 100 mM NADPH, which has been shown to increase binding of MTX and

TMP to DHFR.[303] SEC analysis showed that in the presence of NADPH, Bis-TMP1 was

able to form a small amount of dimer (Figure 5.19).

12.5

10.0

7.5 mAU

5.0

2.5

0.0

-2.5

-5.0 0 2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 34 36 Minutes Figure 5.19. SEC analysis of Bis-TMP1 induced tethered DHFR dimerization in the presence of 100 mM NADPH (red line). A small peak (left) denotes dimer formation. The addition of tethered DHFR (blue line) resulted in a single peak. Image provided by Dr. Adrian Fegan (Wagner Lab).

The results from the Wagner Lab indicated that Bis-TMP1 was capable of dimerizing

a small amount of DHFR. Thus, while we continued to try to demonstrate successful CID- mediated dimerization with the in vitro fluorescent assay, we also attempted to utilize Bis-

TMP1 to label the cell membrane of HEK-293T cells in vivo.

5.4. In Vivo Dimerization Assays

In order to express DHFR on the surface of HEK-293T cells, the DHFR gene was cloned into the vector pDisplay (Invitrogen), a 5.3 kb mammalian expression vector. The

138 resulting vector was termed pDisplay-DHFR. Expressed DHFR is fused at its N-terminus to the murine Ig κ-chain leader sequence that directs proteins to the cell secretory pathway, and at its C-terminus to a growth factor receptor transmembrane domain that anchors the DHFR in the plasma membrane.[305] The DHFR is thus displayed on the extracellular side of the cell membrane and is consequently accessible to the CID.

293T cells were transfected with pDisplay-DHFR and after 24 hours, cells were incubated with 100 nM CID, a 10-fold higher concentration than the Cornish Lab used to successfully label CHO cells. After four hours, the medium was changed to remove excess

CID, then GFP-DHFR was incubated with the cells overnight at 37 °C. Cells were then imaged, but no GFP fluorescence was detected on the surface of the cell.

5.5 Summary and Outlook

We have thus begun to develop a photoreversible protein dimerization system for use in vitro and in mammalian cell culture, with the goal of utilizing our novel light-responsive

Bis-TMP1 CID to externally label cells. Induced protein dimerization on cells could be employed to translocate dyes and label cells in a light-dependent fashion. Alternatively, Bis-

TMP1 can be used intercellularly in a modified yeast three-hybrid system similar to that developed by the Cornish Lab, where DHFR is fused to a TA and DBD. However, the

Cornish system lacks spatial control, and the binding event is permanent. These limitations would be overcome with the use of our photoreversible Bis-TMP1 dimerizer. Further

139 investigation of our system is thus required, and the initial in vitro dimerization assays should be performed in the presence of NADPH, as indicated by findings in the Wagner Lab.

In addition, we have constructed a DHFR-transmembrane domain fusion for displaying DHFR on the surface of HEK-293T cells. A GFP-DHFR fusion was also expressed and purified in E. coli. We reasoned that cells expressing membrane-bound DHFR could be incubated with Bis-TMP1. After washing the cells to remove excess Bis-TMP1, addition of DHFR-GFP to the medium would enable dimerization and subsequent localization of fluorescence to the outside of the cell. We attempted this in parallel with in vitro assays using DHFR-GFP and 6xHis tagged DHFR. While these assays initially showed promising results, to date, we have been unable to utilize our photoreversible Bis-TMP1 CID to affect protein dimerization and subsequent membrane labelling in mammalian cell culture.

Once the in vitro assays are optimized, cellular DHFR expression from pDisplay-

DHFR should be verified by Western blot analysis prior to incubation with Bis-TMP1.

Additionally, Bis-TMP2, 3, and 4 should be investigated in in vivo studies to dimerize GFP-

DHFR.

5.6 Experimental Methods pET21-DHFR Vector Construction. The DHFR gene was PCR amplified (95 °C (2 min), then 30 cycles of 95 °C (30 sec), 55 °C (30 sec) and 72 °C (1.5 min)) from E. coli

BL21(DE3) Gold cells using the primers 5’-TGATTCGCTAGCATTAGTCTGATTGCGGC

-3’ and 5’-GTCTCGAGTTACCGCCGCTCCAGAATCT-3’ which contain NheI and XhoI

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restriction sites, respectively. The DHFR gene was then inserted into pET-21a(+) (Novagen) as an NheI-XhoI fragment, generating pET21-DHFR and yielding hexahistidine-tagged wild type DHFR upon expression in E. coli. The expected vector sequence was confirmed by

DNA sequencing using the sequencing primers 5’-GCTAGTTATTGCTCAGCGG-3’ and 5’-

GTCTCGAGTTACCGCCGCTCCAGAATCT-3’.

pBAD-GFP-DHFR Vector Construction. The DHFR gene was PCR amplified (95 °C (2 min), then 30 cycles of 95 °C (30 sec), 55 °C (30 sec) and 72 °C (1.5 min)) from pET21-

DHFR cells using the primers 5’-ATATGAGCTCAGTCTGATTGCGGC-3’ and 5’-

GCATACTAGTTCAGTGGTGGTGGT-3’ which contain SacI and SpeI restriction sites, respectively. The 6xHis-tagged DHFR gene was then inserted as a SacI-SpeI fragment into pGFPuv (Clontech) on the 3’ end of the GFP gene, generating pGFP-DHFR and yielding the

6xHis-tagged GFP-DHFR fusion upon expression in E. coli. However, in pGFPuv the GFP-

DHFR gene was under control of a constitutive lacUV promoter, thus there was no control over protein expression. The GFP-DHFR gene was then PCR cloned into pBAD-myc-His

(Invitrogen) using the primers 5’- ACGTCACTGCAGAAATGAGTAAAGGAGAAG-3’ and 5’-ATGAGATAAGCTTTCAGTGGTGGTGGT-3’ and yielding pBAD-GFP-DHFR.

The expected vector sequence was confirmed by DNA sequencing using the sequencing primer 5’-ATGCCATAGCATTTTTATCC-3’

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DHFR and GFP-DHFR Expression and Purification. To express DHFR and GFP-DHFR,

BL21(DE3) Gold cells were transformed with pET21-DHFR or pBAD+GFP-DHFR,

inoculated 1:100 mL in 25 mL LB medium containing 50 µg/mL ampicillin, and were grown

at 37 °C, 250 rpm. At an OD600 of 0.6, expression was induced with IPTG (0.5 mL of a 1 M

stock solution) to a final concentration of 0.5 mM for four hours at 37 °C, followed by 15

hours at 25 °C. The cell pellets could be frozen and stored overnight at 80 °C with no loss of

enzyme activity. The cell pellets were then thawed on ice, lysed, and the proteins purified

witha Ni-NTA resin according to the manufacturer’s instructions. Briefly, 1 mL of Qiagen

lysis buffer and 25 mg of lysozyme were added to the cell pellet resulting from a 25 mL

culture (~ 250 µL), and cells were resuspended and incubated for one hour on ice. For

complete lysis, cells were vortexed and sonicated (Branson Sonifier 450) with a microtip

attachment (2 min continuously on ice, 60 W, 50 % duty cycle). Cellular debris was pelleted

in an Eppendorf refridgerated centrifuge at 4°C, 13,200 rpm, 30 min, then the lysate was

decanted and 25 µL of Ni-NTA resin was added to approximately 1 mL of cleared lysate in a

1.5 mL eppendorf tube. The resin was placed on a rocker in the cold room for one hour, then pelleted (3000 rpm, 15 sec) and washed with five times with 5x1 mL Qiagen wash buffer.

500 µL of Qiagen elution buffer was added to the resin repeatedly until electrophoretic

analysis showed no protein (typically two or three total elutions). Protein production and

purity were verified by 12% SDS-PAGE. The samples were run at 12 mA to allow the

loading dye to clear the stacking gel (about 15-20 min), then electrophoresed at 24 mA for

45-60 min or until the prestained protein marker (NEB) was adequately separated. Finally,

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elution fractions containing DHFR and GFP-DHFR were dialyzed (Fisherbrand regenerated

cellulose dialysis tubing; 6000-8000 nominal MWCO) at 4 °C into DHFR storage buffer (1

mL 0.5M EDTA, pH 8.0, 635 µL ethanethiol, 10 mL 0.5 M KH2PO4, pH 7.0, 1.86 g KCl,

100 mL glycerol, and H2O to a final volume of 500 mL over 24 hours with two changes of buffer (500 mL each). Dialyzed protein was stored at 20 °C. The enzyme concentration was determined by a Bradford assay to be 500 ng/µL for DHFR (10 mg/L total protein concentration) and 480 ng/µL for GFP-DHFR (9.6 mg/L total protein concentration).

In Vitro DHFR Dimerization and Fluorescent Dimerization Assays. The in vitro DHFR dimerization and fluorescent dimerization assays were performed in an identical manner, with the exception that in the fluorescent dimerization assay, instead of DHFR, GFP-DHFR was added to the reaction after initial incubation of immobilized DHFR with Bis-TMP. All assays were performed at room temperature or at 4 °C, with no discernible difference between the two temperatures, as analyzed by SDS-PAGE. 20 µL of Ni-NTA resin (Qiagen) were equilibrated with 1X DHFR reaction buffer (50 mM Tris-HCl, pH 7.5, 0.01% ethanethiol, and 0.01% Triton-X) in a 1.5 mL clear eppendorf tube. The resin was then pelleted in an Eppendorf desktop centrifuge at 3000 rpm for 15-30 sec, and the flow-through was carefully removed from the tube with a pipet, with great precaution taken to avoid disturbing the resin at the bottom. 25 µg of DHFR in 1X activity buffer for total volume of

500 µL were then added to the resin. The reported binding capacity of the resin (from the

Qiagen Ni-NTA protein purification handbook) is 10 µg/µL; thus, the resin could bind 200

143

µg. We used 350 µg to ensure that all of the Ni2+ coordination sites would be occupied.

DHFR was incubated with the resin for one hour at room temperature with constant shaking

to prevent the resin from settling. The resin was then pelleted for 15-30 sec at 3000 rpm, the

flow-through was removed and saved for analysis, and the resin was washed with 10x1 mL

of 1X DHFR activity buffer. 150 µM Bis-TMP (5 µL of 30 µM stock) in 1X DHFR activity

buffer with a total volume of 500 µL was then added to the resin and incubated at room

temperature with constant shaking. After one hour, the resin was again pelleted and washed

as described above. To the resin, 350 µg of DHFR or GFP-DHFR were then added in 500 µL total volume and incubated one hour at room temperature with constant shaking. The resin was again pelleted and washed as described above, then resuspended in 50 µL of 1X DHFR activity buffer. The resin was split between two 200 µL PCR tubes so that one tube could be

irradiated and the other kept in the dark. Prior to resin irradiation, the transilluminator was

chilled to approximately 4 ºC for ~45 min with a frozen ice pack to ensure that the

temperature would not increase and thus denature the protein from the resin, leading to a

false positive result on the SDS-PAGE gel. One of the resin aliquots was placed directly on

the transilluminator and irradiated for 30 min at 365 nm. The ice pack was placed on top of

the PCR tube to maintain a low temperature. After 30 min, the flow-through was removed

from both the irradiated and the nonirradiated reactions. The resin was washed once with 1X

DHFR activity buffer, then remaining resin-bound DHFR was eluted with 1 mM TMP. All

fractions collected prior to irradiation were evaporated to dryness (Savant OligoPrep OP120),

and resuspended in 25 µL of water for SDS-PAGE analysis to ensure that all reactions has an

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equal volume. To 12 µL of each fraction, 4 µL of 4X protein loading dye were added and

heated 5 min at 99 °C. Each aliquot was then run on a 12% SDS-PAGE gel. The samples were run 12 mA to allow the loading dye to clear the stacking gel (about 15-20 min), then electrophoresed at 24 mA for 40 min or until the prestained protein marker (NEB) was adequately separated. The gel was removed and stained two hours in Coomassie staining solution, then one hour in destaining solution. The remaining 13 µL aliquots were transferred to individual wells in a 96-well plate. The volume was brought to 100 µL with 1X DHFR reaction buffer, then the plate was read on a Molecular Devices Gemini EM microplate spectrofluorimeter (λex = 395 nm / λem = 525 nm).

Bis-TMP Purification. All Bis-TMP CIDs were synthesized by Dr. Douglas Young,

followed by purification on a Hewlett Packard Series 1100 HPLC using a C-18 reverse phase

column. 100 µL of sample was injected and a gradient program run, in which the aqueous solvent (0.1% TFA in H2O, pH 2.2) was decreased from 90% to 40% to 90% in acetronitrile

over 35 min, with a flow rate of 0.5 mL/min. Fractions were collected at approximately 55

% aqueous solvent, as indicated by a bright yellow color. Fractions were then pooled and

evaporated to dryness (Savant OligoPrep OP120). Approximately 1 mg of Bis-TMP was

purified from four runs. The CID was suspended in 30 µL DMSO to make a 30 mM stock solution, and then stored at 4 ºC. The final working concentration of Bis-TMP in the in vitro assays was 150 µM.

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pDisplay-DHFR Vector Construction. The E. coli DHFR gene was PCR amplified from

pET21-DHFR using the primers 5’-CCAGCCGGCCAGATCTATGATCAGTCTGATT-3’

and 5’-GGATCCCGGGAGATCTGCCGCCGCTCCAGAAT-3’, and cloned into pDisplay

(Invitrogen) using CloneTech InFusion PCR technology. This commerically available system

uses a proprietary recombinase to insert a gene of interest into a linear vector without the use

of restriction enzymes to cut the PCR product. A single enzyme may be used to linearize the

vector; thus, we cut pDisplay with BglII, then mixed 150 ng vector with 2 molar equivalents

(31 ng) of the DHFR PCR product in 10 µL of water, according to the reaction molar ratio calculator provided by CloneTech (http://bioinfo.clontech.com/infusion/molarRatio.do). The mixture was then added to an aliquot of dried-down InFusion mixture and incubated 15 min at 37 °C followed by 15 min at 50 °C. 40 µL of TE buffer were added to the mixture and 2

µL were transformed into InFusion-specific cells. The colonies were then PCR screened to determine the presence of the DHFR gene, and the expected vector sequence was verified by

DNA sequencing using the primer 5’-TAATACGACTCACTATAGGG-3’.

In Vivo Dimerization Assays. HEK-293T cells were passaged into a 96-well plate and

grown to 80% confluence. The cells were then transfected with 1 μg of pDisplay-DHFR

using the XtremeGene transfection reagent (3:1, 3:2 and 6:1 XtremeGene/DNA; Roche

Biomedicals) in OptiMEM medium (Invitrogen). The transfection was incubated at 37 oC for four hours, followed by replacement of tranfection medium with DMEM standard growth medium and addition of 100 nM of the Bis-TMP1 CID. As a control, three wells that were

146

not transfected with pDisplay-DHFR were also incubated with Bis-TMP1. After another four hours, the DMEM was replaced to ensure removal of excess Bis-TMP1 and 3 µg of GFP-

o DHFR were added. Cells were again incubated at 37 C (5% CO2) for 24 hours, washed with

fresh DMEM, then imaged on a Lecia DM5000B to assess GFP expression. However, to

date, no GFP fluorescence was observed.

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CHAPTER SIX – THE HALOTAG AS A PROTEIN-BASED DIMERIZER

6.1 Introduction to the HaloTag Protein

Genetic fusions have been invaluable tools in the study of cellular processes.[306,

307] Fusions to fluorescent proteins allow visualization of cellular events such as protein

localization and protein interaction.[308-311] The landmark works of Osamu

Shimomura,[312] Roger Tsien,[313] and Martin Chalfie[314] toward the expression and

enhancement of green fluorescent protein (GFP) from the jellyfish Aequorea victoria have

enabled GFP to become the standard for in vivo cell imaging. Today, GFP fusions are widely utilized due to the ease of GFP expression, its small protein size, and its high tolerance to N-

or C-terminal modifications;[315] additionally, GFP derivatives such as YFP and DsRed have been created that provide a range of colorimetric options for imaging multiple proteins

within a single cell.[315][316] Genetic fusions such as the 6xHis tag or GST tag are also

employed to isolate and purify proteins.[317] The fusion protein must be genetically encoded

up- or downstream of its partner, and one drawback is that these methodologies are mututally

exclusive, i.e. the 6xHis tag is useful only for purification and not cellular imaging, while

GFP is appropriate for visualization, but not protein isolation. As such, it would be beneficial

to combine both functionalities into one fusion to limit the amount of cloning and gene

manipulation that is otherwise required.

Accordingly, the HaloTag system (Promega) was recently developed.[318, 319] The

HaloTag is a 34 kDa haloalkane dehalogenase from Rhodococcus rhodochrous whose

hydrolytic His289 residue was mutated to Gln to allow stable formation of a covalent bond

148

with any synthetic ligand that contains a specific chloroalkane linker (Figure 6.1).[319] This

linker acts as a multifunctional handle, where the remainder of the ligand is comprised of a

fluorescent dye, affinity tag or solid support. In this manner, only a genetic fusion between

the HaloTag and a protein of interest is required. Various synthetic ligands are commercially

available, or can be readily synthesized.[320] The HaloTag technology thus provides a

modular approach to protein labelling, cellular visualization, and purification (in conjunction

with a TEV protease site located between the protein of interest and the Halotag fusion

protein), both in bacterial and mammalian cell culture.[321]

A. B.

Figure 6.1. A. The HaloTag protein forms a covalent bond with a ligand containing a reactive chloroalkane linker. B. The TMR- and FAM-labeled, as well as resin-immobilibed ligands are commercially available (Promega). Adapted from Los, G. V., et al. ACS Chem. Biol. 2008, 3, 373.

We are interested in utilizing the HaloTag protein to afford light-regulation of protein dimerization. A photocleavable HaloTag ligand can be synthesized that contains on one end a chloroalkane linker, and on the other end a compound such as TMP that binds DHFR (See

Section 5.1). A modified yeast three hybrid system could then be constructed in which the

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HaloTag protein is fused to a DNA binding domain and DHFR is fused to a transcriptional activator. In the absence of light irradiation, the two proteins would dimerize and thus activate transcription of a reporter gene such as β-galactosidase. Photocleavage of the

HaloTag linker upon irradiation would halt transcription, thereby demonstrating a new method of obtaining photocontrol over protein dimerization, and its application to the regulation of gene function.

6.2 Expression and Purification of HaloTag

The HaloTag protein was expressed from modified pFN18 (Promega), which allows

T7 phage-driven expression of HaloTag in E. coli. Due to the proprietary presence of the lethal barnase gene downstream of the HaloTag gene, attempts to transform the commercially available pFN18 into E. coli did not result in any colonies. We thus designed primers to amplify the entire vector while excluding the barnase gene. A 6xHis tag was then inserted at the C-terminus of the protein to facilitate Ni-NTA affinity purification, yielding the modified pFN18 plasmid, pHT-His.

In collaboration with Laura Gardner (Deiters Lab), BL21(DE3) cells were transformed with pHT-His. A small scale initial expression was performed and cells were induced with 0.5 mM IPTG upon reaching an OD600=0.65. Protein was expressed for four hours at 37 °C, then purified. Analysis by 10% SDS-PAGE verified that the HaloTag protein was successfully expressed (Figure 6.2).

150

Figure 6.2. SDS-PAGE analysis of HaloTag protein expression. M: protein maker (in kDa); lane 1: lysate; lane 2: flow-through; lanes 3-6: washes; lanes 7-8: HaloTag elutions.

We next assayed the activity of our expressed HaloTag. The protein was incubated for one hour at room temperature in the presence of 1, 10, and 100 equiv. of a dansyl- modified chloroalkane ligand (Figure 6.3). Aliquots were then run on a 12% SDS-PAGE gel until the pre-stained protein ladder indicated sufficient run time. We expected active HaloTag to bind to the ligand and produce a blue-green fluorescent band on the gel when exposed to

UV light of 365 nm. We thus imaged the gel on a transilluminator, followed by conventional

Coomassie blue staining; however, while later Coomassie blue staining revealed the HaloTag band, no dansyl fluorescence was originally detected on the gel.

Figure 6.3. The dansyl-chloroalkane HaloTag linker.

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6.3 Summary and Outlook

We have successfully expressed the HaloTag protein and are currently working on the synthesis of novel ligands that can be used to affect protein dimerization, such as with a modified yeast three hybrid system, upon light irradiation. This new methodology would provide a new approach to the photoregulation of gene function.

6.4 Experimental methods

Cloning of the HaloTag Gene. pFN18 was purchased from Promega. Due to the presence of the lethal barnase gene directly downstream of the HaloTag gene, the plasmid could not be propagated in E. coli. To solve this problem, the entire plasmid was PCR amplified ((95 °C

(2 min), then 12 cycles of 95 °C (30 sec), 62 °C (30 sec) and 72 °C (8 min)) with the primers

5’-ATCGCGAAGCTTAAACGAATTCGGGCTCGGTA-3’, which annealed at the 3’ end of the barnase gene and ran in the 5’-3’ direction, and 5’-

ACTGCGAAGCTTGTGCCATGTGGATTCCTTAC-3’, which annealed at the 3’ end of the

HaloTag gene and ran in the 3’-5’ direction, thus amplifying a major portion of the plasmid.

Both primers contained a HindIII restriction site. PCR was performed for 30 cycles with the high fidelity Phusion DNA polymerase. The resulting linear DNA was digested with HindIII, then ligated and transformed into NovaBlue cells that were plated on Ampicillin. As cells containing the parent pFN18 could not grow, no background was obtained. Additionally, a

6xHis tag was inserted at the 3’ end of the HaloTag gene using an NcoI restriction site and the phosphorylated oligos 5’-CATGGTGGTGGTGGTGGTGGTGATC-3’ and 5’-

152

CATGGATCACCACCACCACCACCAC-3’. These oligos each contained a simulated cut

NcoI site and were annealed (95 °C for five minutes, followed by slow cooling to 4°C over

30 minutes) then ligated into the modified pFN18. NovaBlue cells were transformed and

colonies were PCR screened with the primers 5’-TAATCTCGAGCGTCGACAGCCAG-3’

and 5’-TAATCTCGAGCGTCGACAGCCAG-3’ to identify successful insertion of the 6xHis

tag. Positive hits were isolated to create the final expression plasmid, pHT-His. The expected

vector sequence was confirmed by DNA sequencing with the primer 5’-

CTGAATCTGCTGCAAGAAGAC-3’.

Expression and Purification of the HaloTag Protein. pHT-His was transformed into

BL21(DE3) cells. One colony was selected and grown overnight in 2 mL LB medium

containing 50 µg/mL Amp. 25 mL of LB medium was then inoculated 1:100 with 250 µL of

overnight culture and grown at 37 °C, 250 rpm to an OD600=0.65. Cells were then induced with 0.5 mM IPTG (final concentration) and grown an additional four hours at 37 °C. Lastly, cells were pelleted and stored overnight at –80 °C. Ni-NTA resin (Qiagen) was used to purify the HaloTag protein according to the manufacturer’s instructions. Briefly, 1 mL of Qiagen lysis buffer was added to the cell pellet resulting from 25 mL of culture. Cells were resuspended and incubated one hour on ice with approximately 25 mg of lysozyme. Cells were vortexed and sonicated in a VWR sonication bath, model 150D (six cycles of 15 sec of sonication followed by 15 sec on ice,). Cellular debris was pelleted in an Eppendorf refridgerated centrifuge at 4 °C, 13,200 rpm, 20 min, and 50 µL of Ni-NTA resin was added

153

to approximately 1 mL of cleared lysate. The resin was placed on a rocker in the 4 °C cold

room for one hour, then washed with four times with 1 mL of Qiagen wash buffer and eluted

twice in 75 µL each of Qiagen elution buffer. Protein production and purity were verified by

10% SDS-PAGE. Finally, HaloTag was dialyzed into storage buffer (50 mM Tris-HCl, pH

7.8, 150 mM NaCl, 1 mM EDTA and 30% glycerol), and stored at –20 °C.

HaloTag Activity Assay. 300 ng of HaloTag protein was incubated at room temperature with 1, 10, and 100 molar equivalents (equal to 0.0039, 0.039, and 0.39 ng, respectively) of a

dansyl chloroalkane ligand in 30 µL total volume of PBS, pH 7.5. After one hour, 10 µL of

the reaction were then mixed with 2 µL of 6X protein loading dye and run on a 10% SDS-

PAGE gel at 12 mA to clear the stacking gel, then 24 mA until adequate separation of the

protein marker was observed (~40 min). The gel was imaged on a transilluminator at 365 nm

for dansyl fluorescence, but no fluorescence was observed. The gel was then stained for two

hours with Coomassie dye and de-stained overnight to image the HaloTag protein band,

which was clearly visible.

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APPENDIX – RECIPES

A1.1 General Reagents

X-Gal. 20 mg/mL X-Gal in DMF. Store at –20 °C.

1 M IPTG. 1.192 g in 5 mL H2O. Store at –20 °C.

20% Arabinose. 0.2 g/mL in H2O. Store at –20 °C.

20% SDS. 0.2 g/mL in H2O. Store at room temp.

1000X Ampicillin. 50 mg/mL in H2O. Filter sterilize, then store at –20 °C.

1000X Chloramphenicol. 30 mg/mL in H2O. Filter sterilize, then store at –20 °C.

1X PBS. 8 g NaCl, 0.2 g KCl, 1.44 g Na2HPO4, 0.24 g KH2PO4, H2O to 1 L, then pH to 7.4. Autoclave and store at room temp to 4 °C.

TE Buffer. 0.5 mL 1 M Tris-HCl, pH 7.8, 0.1 mL 0.5M EDTA, pH 8, H2O to 50 mL. Store at room temp.

A1.2 Protein Analysis Reagents

10X SDS Running Buffer. 30 g Tris base, 144 g glycine, 10 g SDS, H2O to 1 L, then pH to 8.3. Store at room temp.

10-12% SDS-PAGE Gel Solution. 1.25 mL (10% PAGE)/1.5 mL (12% PAGE) 40% acrylamide:bisacrylamide, 1.25 mL 1.5 M Tris-HCl, pH 8.8, 25 µL 20% SDS, 25 µL TEMED, 6 µL APS, H2O to 5 mL. Use immediately.

4% SDS-PAGE Gel Stacking Stock Solution. 6.25 mL 40% acrylamide:bisacrylamide, 6.25 mL 1M Tris-HCl, pH 6.8, 250 µL 20% SDS, H2O to 50 mL. Store at room temp.

6X SDS-PAGE Sample Loading Dye. 100 µL glycerol, 62.5 mM Tris-Cl, pH 6.8, 200 µL SDS, 0.01 mg bromophenol blue, 50 µL ethanethiol, H2O to 1 mL. Store at –80 °C.

Rapid Coomassie Stain. 0.006 g coomassie brilliant blue, 100 mL glacial acetic acid, H2O to 1L. Store at room temp.

Rapid Coomassie Destain. 100 mL glacial acetic acid, H2O to 1L. Store at room temp.

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A1.3 DNA Analysis Reagents

10X TBE Buffer. 108 g Tris base, 55 g boric acid, 9.3 g EDTA, H2O to 1L. Store at room temp.

6X DNA Sample Loading Dye. 600 µL glycerol, 6 mg bromophenol blue, 60 µL 1M Tris- HCl, pH 7.9, 20 µL 0.5M EDTA, pH 8.0, H2O to 1 mL. Store at room temp.

1% Agarose Gel Solution. 1 g agarose in 100 mL 1X TBE buffer. Store at room temp; microwave ~1 min to melt and then pour into gel cast.

A1.4 Protein Storage and Activity Buffers

1X DHFR Storage Buffer. 1 mL 0.5M EDTA, pH 8.0, 635 µL ethanethiol, 10 mL 0.5 M KH2PO4, pH 7.0, 1.86 g KCl, 100 mL glycerol, H2O to 500 mL. Store at room temp to 4 °C. Once protein is dialyzed into buffer, store protein at –20 °C.

10X DHFR Activity Buffer. 0.5 M Tris-HCl, pH 7.5, 0.1 % ethanethiol, 0.1 % Triton-X. Store at room temp to 4 °C.

1X Flpe Dialysis Buffer. 25 mL 1M HEPES, pH 7.0, 36 µL ethanethiol, 1 mL 0.5 M EDTA, pH 8.0, 8.76 g NaCl, 75 mL glycerol, H2O to 500 mL. Store at room temp to 4 °C. Once protein is dialyzed into buffer, store protein at –20 °C.

10X Flpe Activity Buffer. 12.5 mL 1 M Tris-HCl, pH 7.8, 0.1 mL 0.5 M EDTA, 2.92 g NaCl, 25 mg PEG6000. Store at room temp to 4 °C.

1X Cre Storage Buffer. 10 mL 1M Tris-HCl, pH 7.8, 8.75 g NaCl, 1 mL 0.5M EDTA, pH 8.0, 250 mL glycerol, H2O to 500 mL. Store at room temp to 4 °C. Once protein is dialyzed into buffer, store protein at –20 °C.

10X Cre Activity Buffer. 330 mM NaCl, 500 mM Tris-HCl, 100 mM MgCl2

HaloTag Storage Buffer. 25 mL 1 M Tris-HCl, pH 7.8, 1 mL 0.5 M EDTA, 4.38 g NaCl, 150 mL glycerol, H2O to 500 mL. Store at room temp to 4 °C. Once protein is dialyzed into buffer, store protein at –20 °C.

10X DNA Hybridization Buffer. 100 mM Tris-HCl, pH 8.3, 500 mM KCl, 15 mM MgCl2

156

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