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Designer Oligonucleotides for Probing DNA-Protein and Protein-Protein Interactions


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DESIGNER OLIGONUCLEOTIDES FOR PROBING DNA-PROTEIN AND PROTEIN-PROTEIN INTERACTIONS By ZEHUI CAO A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLOR IDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2004

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Copyright 2004 by Zehui Cao

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This work is dedicated to my family.

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ACKNOWLEDGMENTS I would like to thank my parents for the constant care and support they give me, and my wife, Qian Li, for her support and patience all the time. I would also like to thank my advisor, Dr. Weihong Tan, for his confidence in me and the encouragement he gave me to continue my study, Dr. James Winefordner for his kind help with my graduate study, and the members of my advisory committee for their helpful guidance and suggestions. I would like to thank Chih-Ching Huang for his helpful and enjoyable collaboration with me on many projects, and all the members of Tan research group for their help in my daily work and life. iv

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TABLE OF CONTENTS Page ACKNOWLEDGMENTS .................................................................................................iv LIST OF TABLES ...........................................................................................................viii LIST OF FIGURES ...........................................................................................................ix ABSTRACT ......................................................................................................................xii CHAPTER 1 INTRODUCTION ......................................................................................................1 Importance of Protein Detection and Protein Interaction Study ................................1 Analytical Techniques for Protein and Protein Interaction Study ..............................3 Fluorescence Techniques for Signal Transduction ....................................................6 Fluorescence Quenching and Fluorescence Energy Transfer ...........................9 Fluorescence Anisotropy ................................................................................13 Oligonucleotides as Probes for Protein Interactions ................................................17 Molecular Beacons for Nonspecific Protein Detection ..................................19 Aptamers for Specific Protein Detection Based on FRET .............................20 Aptamers for Protein Detection Using Fluorescence Anisotropy ..................26 2 MOLECULAR APTAMERS FOR REAL TIME PROTEIN-PROTEIN INTERACTION MONITORING ............................................................................28 Experimental Section ...............................................................................................32 Results and Discussion .............................................................................................35 FRET-Based Signaling Aptamer for Protein Binding. ...................................36 Dual-labeled aptamer for thrombin-protein binding study...................37 FRET-based 27mer aptamer for thrombin-protein binding..................40 Fluorescence Anisotropy (FA) Based Aptamer Probes for Protein Interactions. .......................................................................................41 Quick Evaluation of Binding Constants of Protein-Protein Interactions........45 Kinetics of Protein-Protein Interactions in Competitive Assays. ...................48 Conclusions ..............................................................................................................52 v

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3 MOLECULAR APTAMERS-BASED AFFINITY CAPILLARY ELECTROPHORESIS FOR PROTEIN-PROTEIN INTERACTIONS ..................55 Experimental Section ...............................................................................................58 Chemicals and Buffers ....................................................................................58 Apparatus ........................................................................................................59 Separation of Aptamer ....................................................................................60 Aptamer-Based ACE ......................................................................................60 PEG-Assisted Aptamer ACE ..........................................................................61 Results and Discussions ...........................................................................................62 Conformation of aptamer ................................................................................62 Quantification of Thrombin ............................................................................66 Determination of Dissociation Constant (Kd) ................................................68 Competitive Assay ..........................................................................................69 Effect of PEG on Aptamer-Thrombin Complex in CE ..................................72 Aptamer-Based Mobility Shift Assay for Thrombin-AHT Interaction ..........76 Conclusions ..............................................................................................................78 4 NUCLEASE-RESISTANCE OF TELOMERE-LIKE SINGLE-STRANDED OLIGONUCLEOTIDES MONITORED IN LIVE CELLS BY FLUORESCENCE ANISOTROPY IMAGING ......................................................................................80 Introduction ..............................................................................................................80 Fluorescence Techniques for Monitoring Intracellular Biointeractions.........80 Telomere and Its Presence in Live Cells ........................................................82 Experimental Section ...............................................................................................84 Fluorescence Anisotropy Imaging (FAI) System ...........................................84 The Image Processing Program ......................................................................87 Cell Culturing .................................................................................................87 Cell Injection Using Electroporation ..............................................................88 Materials .........................................................................................................88 Results and Discussions ...........................................................................................89 Measuring Anisotropy of Standard Samples ..................................................89 Effects of PMT Voltage on Anisotropy Data .................................................92 Monitoring ssDNA Digestion in Homogeneous Solutions ............................94 Digestion of ssDNA in Live Cells ..................................................................95 Stability of Telomere-Like Oligonucleotides in the Nuclei of Live Cells......99 Conclusions ............................................................................................................102 5 SUMMARY AND FUTURE DIRECTIONS .........................................................104 Summary of Oligonucleotide-Based Protein Interaction Study .............................104 Future Directions ....................................................................................................107 vi

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APPENDIX A PROGRAMMED DNA CONFORMATION CHANGE AND APTAMER ACTIVATION .......................................................................................................112 Introduction ............................................................................................................112 Nanomotors and Photo-Regulation of DNA Hybridization ...................................113 Photo-Regulated Aptamer Release and Activation ................................................116 B ENHANCED PROTEIN BINDING AND INHIBITION BY DUAL-APTAMERS .......................................................................................................122 Introduction to Dual-Aptamers ..............................................................................122 Design of Dual-Aptamers for Thrombin and Experimental Results ......................124 LIST OF REFERENCES .................................................................................................130 BIOGRAPHICAL SKETCH ...........................................................................................138 vii

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LIST OF TABLES Table page 2-1 Sequences of the fluorophore-labeled aptamers used in this work. .........................36 B-1 Inhibition of thrombin-fibrinogen reaction by various ologinucleotides. ..............125 viii

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LIST OF FIGURES Figure page 1-1 A typical Jablonski diagrm. ........................................................................................7 1-2 Absorption and fluorescence emission spectra of fluorescein in pH 9.0 buffer. ........7 1-3 Principle of nucleic acid detection using MB. ..........................................................12 1-4 Illustration of the principle of fluorescence anisotropy. ...........................................14 1-5 Measuring fluorescence anisotropy. .........................................................................16 1-6 Conformation change of thrombin-binding aptamer induced by thrombin. .............22 1-7 Conformation change of PDGF aptamer induced by PDGF. ...................................25 2-1 Dye-labeled protein-binding aptamers reporting protein-protein interactions. ........31 2-2 3-dimentional structure of human -thrombin in complex with 15Ap. ....................35 2-3 Human -thrombin binding induced relative fluorescence change of dual-labeled 15mer aptamer. .........................................................................................................37 2-4 Thrombin bound to FQ-15Ap interacts with other proteins. ....................................38 2-5 Dual-labeled 27mer aptamer for -thrombin/protein interactions. ...........................40 2-6 TAMRA-labeled 15Ap for -thrombin/protein interactions based on fluorescence anisotropy. ................................................................................................................42 2-7 Binding between -thrombin and anti-human thrombin (AHT) confirmed by gel electrophoresis on a 7.5% native Tris-HCl gel. .......................................................44 2-8 TAMRA-labeled 27Ap for -thrombin/protein interactions based on fluorescence anisotropy. ................................................................................................................45 2-9 Effect of the order of incubation with thrombin on thrombin-protein interaction...49 2-10 Rate-limiting step in aptamer-thrombin-protein interactions ...................................51 3-1 Schematics of the capillary electrophoresis setup. ...................................................59 ix

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3-2 Capillary electrophoresis traces of thrombin binding aptamer in the presence of 25 mM Tris, 192 mM glycine and various concentrations of KCl. ..............................63 3-3 Capillary electrophoresis traces of thrombin binding aptamer in the presence of 25 mM Tris, 192 mM glycine and various other metal ions. ........................................65 3-4 Detection of thrombin using aptamer-based ACE. ...................................................67 3-5 Analyses of AT III-thrombin interaction using aptamer-based ACE. ......................70 3-6 Using thrombin binding aptamer to monitor thrombin/AT III interaction. ..............71 3-7 Quantification of AT III-thrombin interaction. ........................................................71 3-8 Effect of PEG on the stability of G-Apt*Thrmb complex. .......................................74 3-9 Analyses of thrombin and thrombin-AT III interaction using PEG-containing sample matrix. ..........................................................................................................75 3-10 Binding between G-Apt*Thrmb and anti-human thrombin (AHT) confirmed by capillary electrophoresis. ..........................................................................................77 4-1 Schematics of the FAI system. .................................................................................85 4-2 Fluorescence images obtained from I VH (left) and I VV (right) channels of the FAI system. ......................................................................................................................90 4-3 Anisotropy of 200 nM TAMRA solutions with increasing concentration of glycerol.. ...................................................................................................................91 4-4 Effect of PMT voltage on anisotropy measurements. ..............................................93 4-5 Anisotropy change of 5-TAMRA-(TTAGGG) 6 -3 (TeloH) during digestion by DNase I in solution monitored by FAI. ....................................................................94 4-6 Digestion of Ctrl1 in live cells monitored by anisotropy change. ............................96 4-7 Anisotropy values obtained by averaging in the nucleus region of the anisotropy images of two cells injected with Ctrl1 and Ctrl1-S respectively. ...........................98 4-8 Digestion of TeloH, TeloT, TeloH2 and TeloCtrl in live cells monitored by anisotropy change. ..................................................................................................101 A-1 Structure of azobenzene is changed by lights of different wavelengths. ................114 A-2 Schematics of photo-regulated activity of thrombin aptamer. ................................117 A-3 Inhibition of thrombin by Ap-CS and Ap-MS monitored by measuring scattering light. ........................................................................................................................119 x

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B-1 Illustration of inhibition of thrombin by a single aptamer (top) and a dual aptamer (bottom). .................................................................................................................123 B-2 Inhibition of thrombin by 15Ap and DA-8S monitored by measuring scattering light. ........................................................................................................................127 xi

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Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy DESIGNER OLIGONUCLEOTIDES FOR PROBING DNA-PROTEIN AND PROTEIN-PROTEIN INTERACTIONS By Zehui Cao December 2004 Chair: Weihong Tan Major Department: Chemistry Proteins are responsible for carrying out most of the functions in living cells. As the result of their important roles in human life, changes in protein expression and properties are directly associated with many diseases. Sensitive detection of the disease-related proteins and understanding of their interactions with other biomolecules may be the first step to effective disease diagnoses as well as finding cures. As a good alternative to traditional antibody-based ligands, protein ligands based on short nucleic acids have advantages such as inexpensive production, easy labeling, good stability, and relatively small sizes. Oligonucleotide probes can not only target many DNA-binding proteins nonspecifically, but also bind to and sense target proteins with high selectivity thanks to recent developments in molecular aptamers. Both nonspecific and specific oligonucleotide probes can be easily labeled with fluorescent tags for easy and sensitive analyses of proteins of interest. We have explored the application of those fluorescently-labeled oligonucleotide probes for study of DNA-protein and protein-protein interactions xii

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in homogeneous solutions, under heterogeneous separation conditions, and even in real time in live cells. By combining protein-binding aptamers and signal transduction mechanisms such as fluorescence resonance energy transfer and fluorescence anisotropy, we were able to monitor protein-protein interactions in real time without labeling either of the two interacting proteins, posing minimum effects on the binding properties of the proteins. Our method has been shown to be simple and effective, with the capability of providing detailed information regarding binding sites and binding kinetics. We have also shown that fluorophore-labeled aptamers, with their small sizes and charged backbones, are ideal capturing and sensing agents in affinity capillary electrophoresis (ACE) for sensitive protein detection and protein-protein interaction analyses. The aptamer-based ACE should be valuable in many areas of protein research. Lastly, we combined the imaging capability of confocal microscopy with fluorescence anisotropy measurements to monitor interactions between fluorophore-labeled DNAs and cellular proteins in live cells. As a demonstration, DNA digestion by nucleases was studied inside cell nuclei in real time. We found that DNAs with telomere-like sequences were more resistant to cellular nucleases than other sequences we tested. xiii

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CHAPTER 1 INTRODUCTION Importance of Protein Detection and Protein Interaction Study Proteins are macromolecules that consist of one or more unbranched chains of amino acids. They are the driving force for proper function of cells and ultimately organisms in human life. A great variety of proteins work together or with other molecules to realize almost all important biological functions, such as catalysis of most biochemical reactions, regulation of cell growth and apoptosis, and controlling motion and locomotion of cells and organisms. Because of the deep involvement of proteins in life, diseases caused by disorders in functions of cells and organisms will always result in changes in expression level of certain proteins and disruptions of various interactions between proteins or between proteins and other molecules. The role of analytical chemistry in protein related studies would be the development of simple yet effective techniques for sensitive and selective detection of important proteins as well as their interactions with other biomolecules. These methods can then be used to greatly facilitate early disease diagnoses, disease mechanism understanding, and eventually drug discovery. With the better understanding of the genetic basis of many diseases such as various cancers, certain biomolecules may emerge as a new class of disease markers with greater sensitivity and selectivity. Take cancer as an example, traditionally, diagnosis of cancers have used biomarker molecules that are produced in higher than normal level either directly by tumor cells or by the response of the human body to the presence of cancers. 1

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2 Detection of the biomarkers in a patients body fluids can serve as the first step in cancer diagnosis and provide critical information to doctors as whether biopsy is needed. Some classic tumor markers include -fetoprotein (AFP), carcinoembryonic antigen (CEA), and prostate specific antigen (PSA). They are usually not very specific to a particular cancer as the level of one tumor marker can be elevated by more than one type of cancer. Another problem is that presence of cancer does not necessarily cause a detectable level of tumor markers, especially in the early stage of the cancer. Extra caution is thus needed in some cases to avoid false negatives. In contrast, changes induced by gene mutation can be more specific. The mutated genes in cancer cells can lead to expression of new proteins not found in normal cells, over-expression of certain proteins that promote cell growth and division such as growth factors and related proteins, or mutation of proteins that inhibit tumor cell proliferation. One example of proteins capable of indicating cancers is the human epidermal growth factor receptor 2 (HER2) usually found on cell membranes. HER2 has been shown to be over-expressed in about 25% of all breast cancer patients. 1 Therefore, tests for HER2 protein in tissue samples are recommended for breast cancer diagnosis. Over-expression of many other growth factors, including insulin-like growth factor-I (IGF-I), epidermal growth factor (EGF) and platelet-derived growth factor (PDGF), are also related with tumor progression. 2-4 Sensitive detection of these important proteins may serve as good indications of both the presence and stage of cancers. Since proteins often work together with many other molecules on a specific task, detection and analyzing of such interactions may help understand the cause of the diseases as well as find the cure. For instance, the p53 protein is a tumor suppressor that

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3 induces cell apoptosis under stressful conditions such as DNA damage by activate transcription of certain genes. 5 Mutation or inhibition of the wild type p53 makes p53 not able to prevent cells from growing out of control and eventually becoming tumors. It has been found that the protein product of MDM2 gene can bind to p53 and inhibit its transcriptional activity, thus contributing to cancer development. The revelation of this protein-protein interaction makes targeting MDM2 protein of great therapeutic interest. 6 Inhibition of the p53-MDM2 interaction will certainly help in early tumor suppression. Analytical Techniques for Protein and Protein Interaction Study Many analytical techniques have been used in protein research. Overall, they can be summarized into two major categories. One of them uses molecular ligands that selectively recognize the proteins of interest, while the other does not rely on such ligands. The latter approach includes many separation-based methods such as those based on electrophoresis and chromatography. These methods usually take advantage of differences in mass, size and charge between proteins. Among them, gel electrophoresis allows proteins to migrate at different rates on cross-linked polyacrylamide sieving support. When the detergent sodium dodecyl sulfate (SDS) is used, the proteins are detangled and negatively changed, and the separation is based only on size. Another related technique known as isoelectric focusing often uses polyamino-polycarboxylic acid ampholytes to form a solid support with a pH gradient. A protein with charges migrates on the support until it reaches the region where the pH equals the proteins isoeletric point (pI) at which the protein carries no charge. In this way, proteins with different pIs can be separated and analyzed. Introduced not long ago, another form of electrophoresis called capillary electrophoresis (CE), is conducted in open fuse-silica capillaries. 7 Unlike traditional electrophoresis, Joule heat generated during

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4 electrophoresis in CE can be dissipated very quickly because of the small capillary diameter. This allows ultrahigh voltage to be applied for the separation, leading to very rapid analyses and high efficiency with theoretical plates up to one million. 8 CE separation is based on both charge and size of the molecules, making it ideal for analyses of biopolymers such as nucleic acids and proteins. Filled-column based chromatography is also used for protein analyses. Besides the widely used HPLC with stationary phases based on hydrophobicity and hydrophilicity, one commonly used method is ion-exchange chromatography, in which resins with charged groups are present in the stationary phase to retain molecules with opposite charges. Molecules of the same charge as the resins come out first. Another type is the size-exclusion chromatography, where beads with pores of certain sizes are packed in the column. Molecules smaller than the pore size will get trapped more easily in the pores, and thus migrate more slowly than molecules of bigger sizes. One thing especially noteworthy is that recent advances in proteomic research have resulted in a powerful approach for large-scale protein analyses. The development of two-dimensional gel electrophoresis (2-D gel), combined with breakthroughs in mass spectrometry (MS), especially soft ionization techniques such as electrospray ionization (ESI) 9 and matrix-assisted laser desorption ionization (MALDI), 10 have made high-throughput protein analyses possible. 11 The separation-based techniques for protein analyses require no protein modifications and can often detect multiple targets. However, a few factors limit their application in some areas of protein study. First, most of the time, good separation may require specifically optimized buffer conditions, which may be very different from those

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5 of cellular environments. Proteins may likely behave very differently than in their native system, which makes it undesirable to study protein interactions. Furthermore, detection of proteins is done after the separation is completed. Thus it may take more time than protein assays that are capable of detection in real time. When it comes to large-scale protein analyses, the time difference may be even more significant. More importantly, detection in a non-real-time fashion makes it near impossible to study kinetics of protein functions or interactions. A very different approach for protein study than the separation techniques is the use of selective binding ligands for targeting proteins. The ligands can be proteins, nucleic acids and many other molecules. However, the most widely used protein ligands are a special category of proteins called antibodies. Antibodies are produced by the immune systems of human or animals upon the invasion of foreign molecules (antigens) such as proteins, carbohydrate polymers, and nucleic acids. 12 They usually contain two heavy polypeptide chains and two light chains, and have molecular weights ranging from 150 kDa to about 950 kDa. Antibodies are able to selectively bind to targets with very high affinity constants, between 10 5 and 10 10 M -1 As a result, they have been the preferred recognition agents for protein detection for the last decades. One of the commonly used antibody-based protein assays is the enzyme-linked immunosorbent assay (ELISA). 13 Although in many different forms, this technique is generally based on antibody-antigen recognition. In a simplified form of the ELISA assay, an antibody is immobilized on a solid surface as a capture agent for the antigen. Samples containing the target protein are incubated with the surface-bound antibodies. The targets will be captured by the antibody while other molecules can be washed away.

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6 The following step usually involves the use of another antibody that also binds to the antigen at a different site. Thus the two antibodies and the antigen form a sandwich-like complex on the surface. The second antibody has been linked to an enzyme previously, so that when solution containing substrate for the enzyme is added to the sandwich complex, the enzymatic reaction can take place. The enzyme and substrate pair is carefully chosen in a way that the colorless substrate will become colored after the enzymatic reaction. ELISA assay is very sensitive because the enzymatic reaction is actually a signal amplification process. The other major advantage is that sample does not need to undergo stringent purification before the test, which may save sample preparation time. ELISA has been routinely used to detect proteins in serum samples. Despite the popularity of this technique, one problem with ELISA is that the antibodies that can be used are limited. The assay may take hours for the enzymatic reaction to complete, so apparently it is not a real-time detection method. Fluorescence Techniques for Signal Transduction Fluorescence has been one of the most important tools for molecular sensing in many fields of research. Upon absorbance of the energy of the incident light, electrons of the fluorescent molecules can be excited to the excited singlet states. Return of the electrons to the ground state is accompanied by emission of photons, resulting in fluorescence. The process of fluorescence is illustrated by the Jablonski diagram in Figure 1-1. S 0 S 1 and S 2 stand for the singlet ground, first, and second electronic states respectively, while T stands for triplet state. As one can see, there is less emitted energy than absorbed energy, thus fluorescence always occurs at a longer wavelength than that of the incident light. The difference between a fluorescent molecules wavelength of

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7 maximum absorption and its wavelength of maximum emission is termed Stokes shift. Figure 1-2 shows typical absorption and emission spectra of a commonly used fluorophore known as fluorescein. Figure 1-1. A typical Jablonski diagrm. Figure 1-2. Absorption and fluorescence emission spectra of fluorescein in pH 9.0 buffer. 14

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8 Because fluorescent molecules have their distinct absorption and emission profiles, they can be selectively detected using related instruments. That is, by measuring fluorescent emission from an unknown sample, it is possible to tell if a certain fluorophore is present. Another major advantage of fluorescence techniques is its very high sensitivity. Since fluorescence is a photon-producing process, use of ultra-sensitive detectors capable of single-photon counting results in very sensitive fluorescence measurements, and even detection of single molecules is possible. 15 Even though there are many ways to utilize fluorescence for molecular recognition, the most basic approach is to tag fluorophores to the analytes that are not fluorescent by themselves for detection purpose. Once the fluorescent molecule is linked to the non-fluorescent target, it is possible to sense the target by measuring fluorescence from the fluorophore. There are two ways to link fluorophores to target molecules. First, in some applications, the fluorophore can be directly adsorbed onto the target through non-covalent forces. One example is the dye ethidium bromide for probing double-stranded DNA (dsDNA). When mixed with target sample, it can intercalate into the double strand of the double-helical DNA and give strong fluorescence. 16 The second approach to tag a target with a dye is more indirect. The fluorophore is first attached to a probe molecule, usually via covalent bonding. The probe should have good affinity selectively for the target molecule. So when the target and dye-labeled probe are mixed together, the target can be fluorescently tagged and later detected. For protein targets, antibodies are the most widely used probes. Fluorescent labels have replaced radioactive labels in antibody-based immunoassays for highly sensitive antigen detection. 17

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9 One thing that needs to be addressed when conducting fluorescence-based analyte detection is how to separate the fluorescence signal of the fluorophore linked to the analyte from that of the fluorophore in its free form. A simple way to do this is to use ELISA-like format to capture the target on a solid surface and then stain it with fluorophore. The unbound fluorophore in solution can be easily washed away later. In this way, any fluorescence coming from the solid support should be originated from target-bound fluorophore. Another way to isolate useful fluorescence signal is to combine dye-labeled protein probe with separation techniques. For example, after the analyte and the dye-labeled antibody are mixed together, separation can be carried out using CE or HPLC. The antibody-analyte complex is isolated from the free dye-labeled antibody due to their size and charge differences. Under fluorescence detection, two peaks representing each of them will be present, indicating the presence of the analyte. Any other species will not be recognized because of the lack of fluorescence. Both approaches can take advantage of the high sensitivity and selectivity of fluorescence measurements. However, the detection does not proceed until after the binding event has happened, leading to difficulties in kinetics study. As a result, a detection scheme is needed to produce signal change during the target recognition process. Fortunately, fluorescence based detection can adopt various designs to achieve such a goal. Two major approaches capable of real time target detection have been employed in our work. Fluorescence Quenching and Fluorescence Energy Transfer There are a variety of ways to decrease fluorescence emission of a fluorescent molecule. The process of decreasing fluorescence is termed quenching. 18 Quenching happens through two major mechanisms. One of them is collisional quenching. When a

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10 fluorophore is excited by an incident light, it leaps onto the excited state and will stay there for a very short period of time, usually in the nanosecond range, before coming back to the ground state. The time it stays on the excited state is called fluorescence lifetime. During its lifetime, the excited fluorophore could collide with other molecules in the solution. The collision may cause energy loss of the fluorophore, which means that the absorbed energy is dissipated in a way other than fluorescence emission resulting in fluorescence quenching. The decrease in fluorescence intensity can be described using the Stern-Volmer equation: F 0 /F = 1+K [Q] = 1+ k q 0 [Q] where K is the Stern-Volmer quenching constant, k q is the bimolecular quenching constant, 0 is the unquenched lifetime, and [Q] is the quencher concentration. 18 Many molecules can be collisional quencher, including oxygen, halogens, amines and acrylamide. The other type of quenching is called static quenching, where the quencher can form non-fluorescent complex with the fluorophore. This quenching happens even before the fluorophore is excited to the excited state. In addition to fluorescence quenching, another related important fluorescence process is the fluorescence resonance energy transfer (FRET). This process involves an energy donor which should be fluorescent, and an energy acceptor. FRET happens when the emission spectrum of the donor overlaps with the absorption spectrum of the acceptor. Because of the overlapping, the photon energy may be able to transfer between the donor-acceptor pair. One important thing to note about FRET is that the donor does not emit photons for the energy transfer to occur. The donor and acceptor have to be

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11 coupled by a dipole-dipole interaction. 18 For this interaction to take place, the two molecules need to be very close, typically less than 10 nm. In fact, the efficiency of the energy transfer is highly dependent on the distance between the donor and acceptor. The relationship can be described using the following equation: E = R 0 6 / (R 0 6 + r 6 ) where E is the energy transfer efficiency, r is the distance between the donor and acceptor, and R 0 is the Frster distance of the donor-acceptor pair. The Frster distance is defined as the distance between the specific donor-acceptor pair at which the FRET efficiency is 50%. The Frster distance is in the range of 3-6 nm, within the size of many macromolecules. The acceptors do not have to be fluorescent. When a non-fluorescent acceptor is used, the result of the energy transfer is quenching of the donor fluorescence. Some non-fluorescent acceptors can act as dark quenchers, meaning they can effectively quench a broad range of fluorophores whose emission spectra overlap with their absorption spectra. Examples of dark quenchers include dabcyl, Black Hole Quenchers TM (BHQ TM -1 and BHQ TM -2, Biosearch Technologies, Inc., Novato, CA). Since FRET is highly dependent on distance between the donor and acceptor, if there is a way to design a probe such that with and without target molecules, there would be a change in the donor and acceptor distance, then the fluorescence change can directly report the presence of the target. One important example of utilizing this idea is the development of molecular beacons (MBs) for detection of nucleic acids in real time. 19 Figure 1-3 illustrates the principle of how a MB recognizes its target nucleic acid:

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12 Figure 1-3. Principle of nucleic acid detection using MB. The MB is a synthetic oligonucleotide, generally ranging from 25-50 bases in length. The molecule contains a stem and loop structure (Figure 1-3). The single-stranded loop region consists of a probe sequence complementary to the intended target nucleic acid. The sequences directly flanking the loop region are complementary to each other but unrelated to the target. Thus, these flanking sequences anneal to form the MB stem. Signal transduction in MBs is accomplished by FRET. A fluorophore is covalently attached to one end of MB, and a quencher, often dabcyl, is covalently coupled to the other end. The fluorescent dye acts as the energy donor, and the quencher acts as a non-fluorescent acceptor. When the stem sequences are hybridized with each other, these two moieties are kept in close proximity to each other, causing the fluorescence of the donor to be quenched by energy transfer. In the presence of target DNA (tDNA), however, the loop region forms a hybrid that is longer and more stable than that of the stem. This forces the MB to undergo a spontaneous conformational change that forces the stem apart. With the quencher no longer positioned near the fluorophore, fluorescence is restored, thus signaling the binding of the MB to its target. Since signal is generated only in the presence of the target DNA, there is no need to separate the hybrid and the MB itself. As a result, continuous monitoring of the fluorescence signal can be used for real-time tDNA detection as well as for hybridization kinetics study.

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13 Another report used a second fluorophore as the acceptor instead of a quencher. 20 In such a design, the energy transferred from the donor will excite the acceptor which then emits photons. If we assign I D and I A as emission intensity of the donor and acceptor respectively, the result of FRET between a closely located donor and acceptor is the decrease of I D and increase of I A In fact, for the MB labeled with two dyes, the ratio of I A / I D is used to sense the presence of tDNA, which provides better sensitivity than simply measuring I A or I D Besides being widely employed for real-time nucleic acid detection, FRET can also be suitable for analyzing proteins. One example is the detection of a DNA-binding protein, catabolite activator protein (CAP), using DNA as the probe. 21 CAP binds to a sequence-specific short piece of double-stranded DNA (dsDNA). To construct the probe for CAP, the specific dsDNA was broken into halves from the middle of the sequence. The two pieces had short complementary overhangs labeled with a fluorophore and quencher respectively. The overhangs could help the two pieces come back and anneal to form the protein-binding dsDNA. However, the length of the overhang was designed to be short enough that in the absence of CAP, the two fragments would not be able to anneal. Only when CAP was present, could the annealing be stabilized because the protein binding favored the formation of the dsDNA. As the two half probes came close to each other, quenching of fluorescence was induced by FRET between the fluorophore and quencher. In this way, rapid protein detection was successfully conducted in homogeneous solutions. Fluorescence Anisotropy Like FRET, fluorescence anisotropy (FA) is another choice for real-time analyte detection. FA is related to the phenomenon that upon excitation with polarized light,

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14 fluorescent molecules often give depolarized emission. Anisotropy can be considered to describe the extent of such depolarization. Figure 1-4 shows an illustration of the principles of ansitropy. Figure 1-4. Illustration of the principle of fluorescence anisotropy. The subscripts V and H in Figure 1-4 refer to the orientation (vertical or horizontal) of the polarization for the intensity measurements, with the first subscript indicating the direction of the polarization of the excitation and the second for the polarization of the fluorescence emission. When excited by a polarized light, the fluorescent molecules that have absorption transition moments oriented along the electric vector of the incident light are preferentially excited. 18 During the lifetime on the excited state, usually nanoseconds, those molecules may still rotate to other directions before returning from the excited state to the ground state and emitting light. As a result, the emission contains not only components with the same polarization state as the excitation source, but also light of other polarization directions. To define this depolarization quantitatively, the concept of anisotropy (r) is introduced and described using the following equation: r = (I VV -I VH ) / (I VV +2I VH )

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15 Apparently, the extent of the depolarization is related to the fluorescence lifetime of the fluorophore and how fast the fluorophore can rotate in its microenvironments. Also illustrated in Figure 1-4, larger fluorescent molecules rotate more slowly on the excited state, resulting in a smaller I VH component in the emission and consequently a higher anisotropy value. There are other factors that can influence rotation of molecules and their anisotropy as well, e.g., the viscosity of the solution. Measurement of anisotropy is usually carried out with two polarizers. One of them is placed in front of the light source to generate vertically polarized excitation. The other polarizer is located somewhere between the sample and the detector. By rotating the emission polarizer to the direction parallel or perpendicular to that of the excitation polarizer, the two polarization components of the emission I VV and I VH can be obtained. Figure 1-5 shows a simplified setup for anisotropy measurements. The I VV and I VH values obtained are then applied to an equation for anisotropy calculation, r = (I VV -GI VH ) / (I VV +2GI VH ) Compared to the previous equation for anisotropy, one may notice the addition of a G parameter, which is called G-factor. The introduction of G-factor is because the optical components and the detector in the instrument may have different response to different polarization states. For example, even though the sample generates same amount of I VV and I VH after all the optical path and detector readout, the final result may be that the two values become very different. In such cases, the G-factor is needed to calibrate and make up for the instrument caused error. Determination of G-factor is usually done by measuring I HH and I HV and calculating using G = I HV / I HH

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16 Figure 1-5. Measuring fluorescence anisotropy. The connection between molecular weight of the fluorescent molecule and fluorescence anisotropy makes anisotropy an ideal method for detection of macromolecules and biomolecular interactions. A probe molecule can be labeled with a dye and used to interact with its protein target. The change in the size of the probe caused by the interaction can lead to a detectable anisotropy change, thus reporting the target molecule. Dye-labeled DNA molecules have been employed to study interactions with proteins under different conditions. 22 Alternatively, protein can also be labeled with fluorophores to study protein-DNA and protein-protein interactions. 23;24 In theory, as long as the biological interaction induces changes in the rotation of the fluorescently labeled probe, anisotropy should be applicable for real-time analyzing of such interactions. On the other hand, other commonly used fluorescence techniques including FRET often require precise conformational change of the probe molecule upon target binding to correctly report the interaction. Other advantages of fluorescence anisotropy include: 1) FA is able to use only one dye label to report target molecules in real time and in homogeneous solutions; 2) FA is a

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17 ratiometric approach, thus less affected by sample fluorescence fluctuation and photo-bleaching. One important thing needs to be taken into account when analyzing anisotropy data is that measured anisotropy from a solution sample is the sum of the contribution from all fluorescent species in the solution. For example, a probe P is labeled with a fluorophore F to give PF. In the given solution, PF has an anisotropy value of r PF When analyte A containing sample is added to the solution of the PF, the probe will bind to the analyte to form a complex A-PF. Anisotropy of the complex, r A-PF is higher that r PF However, the anisotropy measured for this mixed solution is determined by the following equation, r = (1-) r PF + r A-PF where is the fraction of the total PF that has been converted to the complex. It is clear that the more A is added to form more A-PF, the higher the measured anisotropy will be. When A is in excess and all PF is bound to the analyte, the highest anisotropy is achieved, which equals to r A-PF This constructs the basis for quantitative detection of target using fluorescence anisotropy. However, a disadvantage of fluorescence anisotropy is also revealed here, which is the limited dynamic range of detection, in this case, between r PF and r A-PF Oligonucleotides as Probes for Protein Interactions As discussed before, antibodies have been the preferred protein-binding ligands for decades. However, some of their intrinsic properties may limit their applications in certain protein research fields. First, the production of antibodies involves using animal hosts and is a rather time-consuming process. Second, like any other proteins, antibodies are also sensitive to their surrounding environments and may be denatured by changes in pH and salt concentrations. Nonspecific adsorption of antibodies to many solid surfaces

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18 can sometimes destabilize them, which limited their application on a solid supports. Moreover, the larger molecular weight of the antibodies, over 150 kD, is not practical for many assays such as the fluorescence anisotropy where a significant size difference between the probe and the target is desired. In an addition, antibodies lack the ability to have a built-in signal transduction mechanism. Consequently, the detection of target proteins requires either fluorescent labeling of the targets or a second signaling probe. Recent development in antibody fragments uses only the antigen-binding fragments of the antibody for target capturing. 25 Antibody fragments improve upon antibodies in the areas that they can be produced inexpensively in large scale and are much smaller. But the stable F ab fragment is about 50 kD, still comparable to or larger than many common proteins. On the other hand, nucleic acid based protein-binding ligands may be excellent candidate for studying protein-DNA and protein-protein interactions for a couple of reasons. First, nucleic acid probes can be small and flexible, and they are routinely synthesized using DNA synthesizers. Second, there are many ways to attach functional groups and fluorescent dyes to any position of the sequence. Third, nucleic acids, especially DNAs, can withstand harsher environments than proteins and are less sensitive to many buffer conditions. Immobilization of DNAs onto solid surfaces is also relatively easy, with multiple methods to choose from. DNA ligands can be used to study proteins that have non-specific affinity for DNA, such as nucleases. Proteins like transcription factors bind to specific DNAs thus can be studied using the DNAs with the right sequences. Recent advances in selecting protein-specific DNA ligands make it possible to analyze any protein of interest with nucleic acid probes.

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19 A few approaches have been adopted to analyze proteins in real time and in homogeneous solution, such as FRET and anisotropy based assays. Molecular Beacons for Nonspecific Protein Detection Although molecular beacons were originally designed for binding and recognition of specific nucleic acids, these probes can also lead to increased fluorescence upon binding of certain proteins. The protein recognition ability of MBs was first demonstrated with a single-stranded DNA binding protein (SSB). 26 SSB is a 75.6 kDa tetrameric protein that acts as an accessory protein in DNA replication, recombination, and repair. A tetramethylrhodamine (TAMRA)/dabcyl molecular beacon was used, and SSB concentrations as low as 20 nM could be detected using a conventional fluorescence spectrophotometer. Monitoring fluorescence over time shows that the SSB-MB interaction is rapid, reaching equilibrium within 10 s. In fact, the MB binds with the SSB much more quickly than with its complementary DNA. The MB-based SSB assay is, however, not particularly specific. SSB leads to a fluorescence enhancement nearly equal to that of the complementary DNA, but other proteins can also bind with the MB and cause a fluorescence increase. For example, histone and RecA proteins have both been demonstrated to bind with MB and augment fluorescence. The selectivity of MBs for protein detection was further examined through detailed binding studies of the ssDNA-binding enzyme lactate dehydrogenase (LDH). 27 LDH occurs as five distinct isoenzymes, and MBs were used to elucidate how minor structural changes in the protein affect its ability to bind ssDNA. Two LDH isoenzymes from three different species were assayed using MBs, and the resulting fluorescence varied by as much as 80% between the various samples.

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20 Together, these results demonstrate that while MBs are sensitive and somewhat selective to DNA-binding proteins, they are not specific enough to be capable of distinguishing a particular protein. However, the utility of MBs for studying non-specific DNA-binding proteins in enzyme cleavage assays has already been noted. Aptamers for Specific Protein Detection Based on FRET In order to overcome the limitation of non-specific DNA probes for proteins and be competitive with antibodies, nucleic acids ligands that selectively recognize target proteins need to be developed. As a result of great efforts in this direction, aptamers have emerged and gained great attention from researchers. Aptamers are nucleic acids that have high affinity and selectivity for their target molecules. By using the systematic evolution of ligands by exponential enrichment (SELEX) process, 28;29 oligonucleotide sequences can be isolated to recognize virtually any class of molecules. 30 The SELEX process begins with a library of synthesized oligonucleotides usually containing 10 14 to 10 15 random sequences. This library is then incubated with the target molecule of interest under certain conditions. The sequences that interact with and bind to the target molecules are isolated for the next round of incubation. This process is repeated until a sequence that binds to the target with the highest affinity and selectivity is determined. Compared to antibodies, aptamers also have high affinity and selectivity for proteins. 31;32 Advantages of aptamers over antibodies roots from the easy production, easy labeling, and chemical stability of nucleic acid molecules in comparison to protein molecules. In order to employ aptamers for real-time target detection, FRET has been chosen by many researchers as the signal transduction mechanism. In the FRET assay, the aptamer is labeled with a fluorophore and a quencher similar to an MB assay. When the aptamer specifically binds to its target, a consequent conformational change of the

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21 aptamer may take place. This change usually results in a change in the distance between the fluorophore and the quencher, leading to changes in the quenching efficiency as well as in the measured fluorescence. By carefully designing the positions of the two labels, a large difference in fluorescence intensity before and after aptamer-target binding can be achieved. Therefore, very small amounts of target molecules are detected using highly sensitive fluorescence measurements. The first MB-like aptamer probe was reported for a small molecule. Stojanovic et al. developed such a sensor based on a previously reported aptamer for cocaine detection. 33 The cocaine-binding aptamer sequence was labeled at the two ends with a 6-FAM dye and a dabcyl respectively. The presence of cocaine would bring the two termini together to form a stable secondary structure. The fluorophore and quencher were also brought close so that the quenched fluorescence could be used to report cocaine molecules. A similar approach was reported for the detection of the Tat protein of HIV-1. 34 The RNA aptamer was also split into two subunits. One of them was labeled with both fluorophore and quencher in a MB-like structure, so that fluorescence was quenched in the absence of the target. The presence of the Tat protein would stabilize the hybridization between the two subunits and open the MB-like structure of the first subunit. A fluorescence enhancement as high as 14-fold could be obtained for sensitive detection of the Tat protein. Two groups developed aptamer beacons for human -thrombin in about the same time period. 35;36 Thrombin is a serine protease that works with fibrinogen and Factor 13 to help stop bleeding. The first reported aptamer for thrombin contains a 15-nucleotide

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22 consensus sequence, 5-GGTTGGTGTGGTTGG-3. NMR and X-ray diffraction studies have demonstrated that this sequence can adopt a compact unimolecular conformation termed a quadruplex containing two G-quartet structures. 37-39 When bound to thrombin, the aptamer exists primarily in its quadruplex form, while in free solution, it can adopt a more relaxed linear conformation, depending in part upon the ionic strength and temperature (Figure 1-6). Low salt concentrations favor the linear state of the aptamer at room temperature. Figure 1-6. Conformation change of thrombin-binding aptamer induced by thrombin. When bound to thrombin, the 15-base aptamer forms a quadruplex conformation with the protein binding primarily in the base 4-12 region. Fluorescence is quenched upon thrombin binding as the fluorophore and quencher moieties are pulled close together. Based on this conformational change, Li et al. developed an MB-like aptamer probe. 36 When the aptamer is labeled with a fluorophore-quencher pair, the distance between the two labels changes as the aptamer binds to thrombin and takes the compact quadruplex form. To maximize this distance change, and thus to obtain the largest fluorescence decrease upon thrombin binding, a 6-FAM dye and a dabcyl quencher were

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23 linked to the two ends of the aptamer. The fluorescence of 6-FAM was seen to decrease instantly when excess thrombin was added to the aptamer probe. Furthermore, a titration of the probe by thrombin revealed a detection limit of 370 pM (S/N>3). The selectivity of the aptamer-based probe was well demonstrated by mixing the aptamer with different proteins of interest. The results clearly showed that other proteins, including the closely-related -thrombin, did not display much quenching of 6-FAM. In order to obtain even better sensitivity, the fluorophore-quencher pair was replaced by a two-fluorophore pair, coumarin and 6-FAM. The absorption spectrum of 6-FAM overlaps with emission spectrum of coumarin, making FRET between the two dyes possible. Binding of the two-fluorophore aptamer probe to thrombin resulted in decreased coumarin fluorescence and increased 6-FAM intensity. When the ratio of the two intensities was used to build a calibration curve, a lower detection limit of thrombin was achieved at ~112 pM. The approach Hamaguchi et al. took to detect thrombin was to add a few bases to one end of the thrombin aptamer that were complementary to the sequence at the other end. 35 With a fluorophore-quencher pair labeled at the two ends, this sensor gave low fluorescence in the absence of thrombin because of the double helix stem formed between the two complementary termini. Upon binding with thrombin, a more compact quadruplex structure of the aptamer would be favored and the two stem ends were separated. Thus, an enhanced fluorescence was the indication of the presence of the target. FRET-based aptamer probe was later used for real-time detection of cancer-related proteins. One of the often targeted oncoproteins is platelet-derived growth factor (PDGF), a dimeric protein composed of a combination of subunit A and B. Out of the three

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24 isoforms of the protein (PDGF-AA, PDGF-AB and PDGF-BB), the BB form, in particular, has been implicated in tumor growth and progression. While generally undetectable in normal cells, PDGF is overexpressed in a variety of human tumors including gliboblastomas and sarcomas. Aptamers that are selective to the PDGF-B chain have been previously isolated. 40 When bound to the protein, the consensus secondary structure motif of the PDGF aptamers is a three-way helix junction with a conserved single-stranded loop at the branch point. However, without PDGF, only two of the helixes are stable in physiological conditions while the third helix containing the two ends of the aptamer is separated into two strands (Figure 1-7). This conformational difference forms the basis for the construction of a FRET-based PDGF probe. By labeling the two ends of the aptamer with a fluorophore-quencher pair, the fluorescence of the fluorophore is expected to decrease when the aptamer is bound to PDGF. Such a PDGF probe was designed by labeling the 5-end of the aptamer with fluorescein and the 3-end with a dabcyl quencher. 41 To test the capability of this probe for real-time quantitation of PDGF, a series of titration experiments were conducted in a physiological buffer at 37C. The fluorescence of fluorescein was found to decrease as the PDGF concentration increased. The lowest PDGF concentration that could be detected was determined at 0.11 nM. The dissociation constant, K d for PDGF-aptamer complex was calculated to be 3 nM. The very high affinity of aptamers for their targets ensures the probes capability to quantitate trace amounts of target proteins in homogeneous solutions. This aptamer-based assay showed a good ability to differentiate between PDGF-BB and other proteins. The addition of a 5-fold excess of unrelated

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25 proteins including lysozyme, BSA, hemoglobin, and myoglobin had no observable effect on the fluorescence. Unrelated peptide growth factors such as epidermal growth factor and insulin-like growth factor 1 also did not appear to bind with the aptamer probe. Figure 1-7. Conformation change of PDGF aptamer induced by PDGF. Binding of PDGF causes the aptamer to change from a relatively unconstrained structure to a tightly packed one containing a 3-way helical junction with a conserved single-stranded loop. In the same report, the authors continued their work and applied this FRET-based assay to detect PDGF in real cellular samples. For a cell line that was known to secret PDGF, Serial dilutions of protein preparations from each cell line were incubated with a fixed amount of the aptamer probe in a 96-well microtiter plate to obtain dose response curves. Presence of PDGF-BB gave a dose response curve with sharp slope. In comparison, PDGF-AB and AA gave less steep slopes. All the above mentioned examples have demonstrated that by combining the FRET signal transduction mechanism, and the excellent affinity and selectivity of the aptamers for their target proteins, easy yet sensitive assays can be built to analyze proteins of interest in real time without need for any separation, even in relatively complex biological samples.

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26 Aptamers for Protein Detection Using Fluorescence Anisotropy The primary advantage of using aptamers as anisotropy probes over many other molecules, especially antibodies, is their relatively small sizes. They are usually not only much smaller than monoclonal antibodies, but also smaller than their target proteins in most cases. This makes aptamer-based anisotropy probes ideal for protein detection. With the aptamer conveniently labeled with a fluorophore, the anisotropy probe will report binding with a target protein via the increase in the anisotropy of the fluorophore. Unlike the FRET-based assays, where the conformational change of the aptamer is essential for target detection, the aptamer anisotropy assay is not as heavily dependent on the structure of the aptamer probe, and thus may be highly useful for applications where the understanding of the aptamer structure is limited. In an effort to explore the application of an aptamer anisotropy probe for protein analysis, the PDGF aptamer was labeled with a single fluorescein dye at the 5-end. 42 PDGF in the nM range, when added to the aptamer solution, caused a significant anisotropy increase due to the overall larger molecular weight of the complex. Controls were done using pure fluorescein dye mixed with PDGF and no perceptible change in anisotropy was observed. The detection limit of this assay was determined to be 2 nM. In conclusion, the selectivity of the anisotropy probe was tested to be as good as the FRET-based assays. Among a variety of control proteins, not only did unrelated proteins produce no observable changes in the anisotropy of the aptamer, but also the three isoforms of PDGF could be differentiated by the anisotropy assay, with highest selectivity for PDGF-BB. The fluorescence anisotropy increases measured with the PDGF-AB and PDGF-AA isoforms were only ~40% of that resulting from PDGF-BB binding.

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27 Overall, the nucleic acid based protein probes may be able to replace antibodies in many fields of protein research, especially with current trend of proteomics aimed at large-scale and high-throughput analyses of whole cell proteins. While much of the research with nucleic acid probes is simply focused on protein detection, we have applied those probes for protein function study by investigating protein-DNA and protein-protein interactions

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CHAPTER 2 MOLECULAR APTAMERS FOR REAL TIME PROTEIN-PROTEIN INTERACTION MONITORING The functions of living cells are mostly executed and regulated by proteins. The important roles of proteins are often realized through interactions between two or more proteins. As an example, growth factor proteins interact with their receptors on the cell membrane to regulate the proliferation of the cells. In another example, serine protease thrombin works with fibrinogen and Factor 13 to help stop bleeding. In order to understand how cells fulfill their functions and how they react to changes in the environments, it is necessary to gain insight into how proteins interact with each other under different conditions. The function of proteins is regulated by their structure, which is necessary for correct interaction with other molecules small (e.g., the substrate for an enzyme protein) or large (e.g., the chromosomal DNA for DNA and RNA polymerases). The less the proteins are perturbed, the truer information one can obtain about protein-protein interactions. It is well known that proteins fold into certain tertiary structures through a variety of bonds and interactions such as hydrogen bonds and hydrophobic effects. The functions of proteins in biological systems are highly dependent on their tertiary structures. As a result, chemical modifications to proteins such as dye labeling may cause a reduction or even a loss of protein activities by either directly blocking the active binding sites or affecting the three-dimensional folding of the proteins. Therefore, it is highly desirable to avoid any modifications of proteins when monitoring protein-protein interactions in order 28

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29 to obtain the most true-to-life information. However, it remains challenging to accomplish label-free protein interaction monitoring in real time. Bioanalytical techniques based on molecular separation such as gel electrophoresis and capillary electrophoresis (CE) can provide label-free protein-protein binding detection in a relatively complex system, 43 but they lack the ability of real-time analysis in homogeneous solutions. Surface plasmon resonance (SPR) is another technique used to probe two interacting proteins. It requires no protein labeling but usually needs one of the proteins to be immobilized on a sensor silica surface. More recent development in protein-protein interactions is the yeast two-hybrid system that was first reported in 1989. 44 In this system, two proteins are fused to the DNA-binding domain (DNA-BD) and the transcription activation domain (TA) of a yeast transcription factor respectively in the yeast nucleus. The interaction between the two proteins will bring DNA-BD and TA in close proximity which results in the expression of the reporter gene. This method has been widely used to study protein-protein interactions and recently it has been adapted to map protein interactions on a proteome-wide scale. 45;46 However, several factors may limit the application of the two-hybrid system in certain areas. First, it consists of rather time-consuming and labor-intensive procedures compared to some other techniques when used on a small number of proteins. Second, the interactions have to take place in the yeast nuclei, which may be problematic for proteins that do not function well in the nuclei such as the membrane proteins. Moreover, transcription factors and some other proteins can activate the reporter gene expression by themselves, leading to false positives. Another commonly used technique for protein-protein interactions is based on fluorescence resonance energy transfer (FRET), where

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30 two dye-labeled proteins interact with each other to trigger a fluorescence signal change of the fluorophores due to the energy transfer. It is simple and can be easily adapted for most proteins. Still, some proteins may lose their biological activities after they are modified or labeled with dye molecules. Here we describe a convenient and versatile method for real-time protein-protein interactions based on a competitive assay using protein-binding aptamers. Aptamers possess affinity and selectivity, comparable to those of antibodies, for their intended protein targets. At the same time, as discussed previously, aptamers have many inherent advantages over antibodies in the field of real-time protein analyses. Despite being excellent molecular probes for proteins, aptamers have not been used extensively to study the interactions between their target proteins and other proteins. It was reported that aptamers were used for protein-binding small molecule screening and radioactive isotope labeling was used for detection. 47 More recently, an assay was constructed for protein interactions based on protein-dependent ribozymes combined with aptamers. 48 Here we have developed a new competitive assay using protein-binding aptamers directly for protein-protein interactions. Based on the highly stable and flexible structures of aptamers, two signal transduction strategies were established to detect the binding events between the aptamer-binding proteinbait protein, and a second protein prey protein. As illustrated in Figure 2-1, in one approach, the aptamer was labeled with a fluorophore and a quencher to form internal FRET. Binding of the aptamer to the bait protein caused a quenched fluorescence while the binding of the prey protein to the bait protein may displace the aptamer and result in a restoration of fluorescence. In the other approach, the aptamer was labeled with only one fluorophore and the

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31 fluorescence anisotropy of the aptamer was monitored in real time. Binding of the aptamer to the much larger bait protein molecules resulted in an increased anisotropy. Further change in the fluorescence anisotropy of the aptamer could be triggered by the interaction between the bait and prey proteins. Figure 2-1. Dye-labeled protein-binding aptamers reporting protein-protein interactions. (A) Aptamer is dual-labeled with a fluorophore and a quencher. The folded form of the aptamer when it binds to the bait protein results in a quenched fluorescence. The bait-prey protein interaction causes release of aptamer from the bait protein, leading to a restored fluorescence; (B) Aptamer is labeled with only one dye. When bound to the much larger bait protein, the aptamer displays slow rotational diffusion. The interaction between bait and prey proteins displaces the aptamer. The unbound aptamer has much faster rotational diffusion. The change in the rotation rate is reported by fluorescence anisotropy of the dye molecule. Neither approach requires labeling of the interacting proteins, allowing the real interaction between the two proteins to be revealed based on their unaffected biological activities. Both methods enabled monitoring of protein-protein interactions in real-time

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32 and homogeneous solutions with great ease and effectiveness. While they excel in different aspects of protein interaction study, we found that the combination of these two methods capable of providing detailed and solid knowledge about the kinetics of the protein-protein binding as well as mechanism and binding site information of the interactions, which is not possible or easily obtainable with many other techniques. Experimental Section Materials. Dye-labeled aptamers were obtained from Integrated DNA Technologies, Inc. (Coralville, IA). The sequences of the 15mer and 27mer thrombin-aptamer are 5-GGT TGG TGT GGT TGG-3, and 5-ACC CGT GGT AGG GTA GGA TGG GGT GGT-3 respectively. For FRET-based assays, both aptamers were dual-labeled with 6-FAM at the 5 end and Dabcyl at the 3 end to form FQ-15Ap and FQ-27Ap respectively. For fluorescence anisotropy assays, both aptamer sequences were labeled with only TAMRA at the 3 end to make T-15Ap and T-27Ap respectively. A control 15mer aptamer was labeled with only 6-FAM at the 3 end (F-15Ap). All aptamers were purified with HPLC. Human -thrombin (M.W.~36.7 kDa), human antithrombin III (AT3) (M.W.~58 kDa) and a monoclonal antibody anti-human thrombin (AHT) (M.W.~150 kDa) were obtained from Haematologic Technologies Inc. (Essex Junction, VT). Bovine serum albumin (BSA) (M.W.~67 kDa) and a sulfated fragment 54-65 of protein hirudin, Gly-Asp-Phe-Glu-Glu-Ile-Pro-Glu-Glu-Tyr(SO 3 H)-Leu-Gln (HirF) (M.W.~1.5 kDa), were from Sigma-Aldrich, Inc. (St. Louis, MO). All tests were performed in a 20 mM Tris-HCl buffer with a pH of 7.6 that contained 50 mM NaCl and 5% (V/V) glycerol. All reagents for the buffer were obtained from Fisher Scientific Company L.L.C. (Pittsburgh, PA).

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33 Fluorescence measurements. Fluorescence measurements were done on a Fluorolog-3 spectrofluorometer (Jobin Yvon Inc., Edison, NJ). For FRET-based assays, fluorescence of 6-FAM was monitored with excitation wavelength of 490 nm and emission wavelength of 520 nm. For anisotropy-based experiments, Fluorescence of TAMRA was monitored with 555 nm as excitation and 580 nm as emission wavelength. Slit widths were varied to yield best signals. All measurements were carried out in a 100 L cuvette. In the aptamer/thrombin binding experiments, very small volume of -thrombin with high concentration was added to an aptamer solution in the cuvvette to make molar ratio of aptamer and thrombin 1:1, and the fluorescence signals were recorded before and after the addition. For protein-protein binding reaction, aptamer/thrombin mixture at 1:1 molar ratio was placed in the cuvette, very small volume of the second protein solution with high concentration was added to the mixture to make a desired prey protein concentration. All dilution effects caused by addition of samples to the original solutions were corrected during data analysis. Anisotropy measurements. Anisotropy measurements were based on the following equation: Anisotropy VHVVVHVV I G I IGIr 2 where the subscripts V and H refer to the orientation (vertical or horizontal) of the polarizers for the intensity measurements, with the first subscript indicating the position of the excitation polarizer and the second for the emission polarizer. G is the G-factor of the spectrofluorometer, which is calculated as G = I HV /I HH by the instrument. G-factor is dependent on the emission wavelength. For a certain dye, the G-factor would be measured and used throughout the experiments that used the same dye. Then the

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34 spectrofluorometer would keep the excitation polarizer vertical and rotate the emission polarizer from vertical to horizontal position to measure the intensities for anisotropy calculation. For TAMRA, all intensities were measured at emission wavelength of 580 nm with excitation wavelength of 555 nm. Time based measurements were carried out by continuously monitoring the anisotropy readout. With an integration time of 0.5 second, each anisotropy measurement would take about 4.1 seconds. Kinetic Studies. Experiments were done in a 100uL cuvette in the spectrofluorometer. While the detection system was running, the reaction samples were quickly mixed together. Data were recorded from the point of mixing to when the signal reached plateau and stabilized. The reaction was regarded completed when the signal was at the plateau. Detections were either done using steady state anisotropy measurements for AT3 and AHT study with T-15Ap, or steady state fluorescence measurements for HirF study with FQ-15Ap. Study with AT3 was conducted at room temperature while AHT and HirF experiments were done at 5 C to enable monitoring of the otherwise too fast reactions by our instrument. The temperatures of reactions were maintained using a RTE-111 water bath/circulator (Neslab Instruments, Inc., Newington, NH). Gel electrophoresis. Gel electrophoresis was performed on a Mini-Protean 3 precast gel system (Bio-Rad Laboratories, Inc., Hercules, CA). Samples loaded on a 7.5% resolving Tris-HCl native gel (Bio-Rad Laboratories, Inc., Hercules, CA) were run at 150 V for 150 minutes. The gel was then taken out, rinsed with ultra-pure water and stained with Coomassie blue stain reagent (Fisher Scientific Company L.L.C., Pittsburgh, PA) for 1 hour. A digital camera was used to image the stained gel.

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35 Results and Discussion Human -thrombin (-thrombin) and its aptamers were used as a model system to demonstrate the capability of aptamers to probe protein-protein interactions. Human -thrombin is a serine proteinase which has two positive-charged sites termed Exosite I and II on the opposite sides of the protein. 49 Exosite I was found to bind to fibrinogen 50 and hirudin 51 while Exosite II binds to heparin. Two different aptamers have been identified that have high affinity and selectivity for -thrombin. The first one is a 15mer single-stranded DNA aptamer which was reported to bind to the fibrinogen-binding site of -thrombin (Figure 2-2), 52 namely Exosite I. The other DNA aptamer, with a 27mer backbone length, was determined to bind to the Exosite II of -thrombin. 53 Both aptamers were found to adopt a G-quadruplex structure when bound to -thrombin. A 15mer Exosite I binding aptamer (15Ap, Table 2-1) and a 27mer Exosite II binding aptamer (27Ap, Table 2-1) with similar thrombin-binding affinity were chosen to study the interactions of -thrombin with other proteins. Figure 2-2. 3-dimentional structure of human -thrombin in complex with 15Ap. 54 15 Ap is shown in purple and blue at the bottom.

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36 Table 2-1. Sequences of the fluorophore-labeled aptamers used in this work. Oligo name Oligo sequence FQ-15Ap 5-(6-FAM)-GGT TGG TGT GGT TGG-(Dabcyl)-3 T-15Ap 5-GGT TGG TGT GGT TGG-(TAMRA)-3 FQ-27Ap 5-(6-FAM)-ACC CGT GGT AGG GTA GGA TGG GGT GGT-(Dabcyl)-3 T-27Ap 5-ACC CGT GGT AGG GTA GGA TGG GGT GGT-(TAMRA)-3 FRET-Based Signaling Aptamer for Protein Binding. We previously reported a molecular beacon aptamer for -thrombin detection based on the 15Ap. 36 Here a slightly modified aptamer (FQ-15Ap, Table 2-1) has been used that incorporates a 6-carboxyfluorescein (6-FAM) at the 5 end of the DNA as the donor and a Dabcyl at the 3 end as the quencher. The quenching of 6-FAM emission is caused by energy transfer between 6-FAM and Dabcyl in the protein-binding induced G-quartet structure where the two labels are in close proximity. When excess -thrombin was added to an FQ-15Ap solution at room temperature, the fluorescence of 6-FAM dropped about 55 percent (Figure 2-3). It is known that high metal ion concentrations, especially the presence of K + can promote the formation of G-quartet, 55;56 which will result in a much lower fluorescence signal change upon aptamer/-thrombin binding. However, using a buffer without any metal ions was found to inhibit protein-protein interactions. By keeping a 50 mM NaCl concentration in the buffer, we were able to sustain the protein activities and get relatively high fluorescence quenching induced by protein binding to the aptamer. When -thrombin was added into a control 15mer aptamer that was labeled only with 6-FAM, no significant fluorescence change was observed (Figure 2-3), indicating that the fluorescence decrease in the FQ-15Ap-thrombin binding experiment was due to the binding-induced conformational change of

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37 the aptamer rather than a direct quenching of the dye 6-FAM by -thrombin. We did not observe the quenching (i) under conditions where thrombin would not bind the aptamer beacon, and (ii) with a scrambled aptamer beacon to which thrombin does not bind. This result was consistent with reported molecular beacon aptamer study. 36 0.00.20.40.60.81.01.2FQ-15ApF-15ApRelative Fluorescence 0.00.20.40.60.81.01.2FQ-15ApF-15ApRelative Fluorescence 0.00.20.40.60.81.01.2FQ-15ApF-15ApRelative Fluorescence 0.00.20.40.60.81.01.2FQ-15ApF-15ApRelative Fluorescence Figure 2-3. Human -thrombin binding induced relative fluorescence change of dual-labeled 15mer aptamer. On the left, 6-FAM florescence intensity of 100 nM FQ-15Ap aptamer in physiological buffer before (1, blue column) and after (white column) the addition of 500 nM -thrombin. One the right, a control 15mer aptamer labeled with only 6-FAM (F-15Ap) underwent the same experiment and the relative fluorescence before (1, blue column) and after (white column) the addition of 100 nM -thrombin was measured. Dual-labeled aptamer for thrombin-protein binding study. The 1:1 molar ratio FQ-15Ap/-thrombin solution (bait solution) was used to identify interactions of -thrombin with other proteins. When a second protein (prey protein) binds to the same site of -thrombin as the FQ-15Ap, the aptamer is expected to be displaced and the freed aptamer will shift back to a more relaxed conformation,

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38 resulting in restored 6-FAM fluorescence. A sulfated fragment of hirudin that contained the C-terminal 13-residue 51 (HirF) instead of hirudin was used for binding -thrombin. The addition of HirF to the FQ-15Ap bait solution caused a sharp fluorescence increase (Figure 2-4), which was expected since both HirF and FQ-15Ap bound to the same site of -thrombin. Control experiments showed that there was no fluorescence change when HirF was added to a FQ-15Ap in the absence of thrombin, indicating that there was no direct interaction between the aptamer and HirF. The time course results showed that this competitive binding reaction was fast as the aptamer departed within seconds after HirF was added to the aptamer-thrombin complex solution. 050010001500200025003000125130135140145150155160165170 Fluorescence (A.U.)Time (s) Figure 2-4. Thrombin bound to FQ-15Ap interacts with other proteins. In a solution of mixed 100 nM FQ-15Ap and 100 nM -thrombin, 200 nM AT3 (), 500 nM HirF () or 300 nM AHT () was added at 0 sec. and fluorescence of 6-FAM was continuously monitored.

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39 Several other proteins were also investigated for interactions with -thrombin using the FQ-15Ap bait solution. The addition of an antibody, anti-human thrombin (AHT), caused no significant change in the fluorescence of 6-FAM (Figure 2-4). While this result indicates that AHT does not compete with the aptamer for the Exosite I of -thrombin, we can not exclude the possibility that AHT still binds to -thrombin but at a different site of -thrombin. More experiments were done to address this issue (results are presented later in this paper). A serine protease inhibitor antithrombin III (AT3) was also tested in the bait solution. A slow-signal increasing trend was observed for AT3 (Figure 2-4). Addition of excess AT3 further increased the 6-FAM fluorescence, but the fluorescence intensity never exceeded that of the FQ-15Ap solution in the absence of -thrombin. This result could be explained in that the binding of AT3 to -thrombin may have caused a conformational change in -thrombin that rendered the binding with the aptamer at Exosite I unstable. 57 The slow reaction rate of AT3 was probably due to the fact that its interaction with the active site of serine proteinases is a multi-step, covalent-bond-forming process. 58 Bovine serum albumin (BSA) was used as a control protein for interaction with -thrombin. No fluorescence change was observed for BSA. Another set of control experiments were conducted by adding the prey proteins to be tested to an FQ-15Ap buffer solution without -thrombin. None of the proteins affected fluorescence of the aptamer, meaning they did not interact with either the aptamer or the fluorophore. It is also possible to quantify the amount of prey protein that is interacting with thrombin using different level of signal change. We found that at higher thrombin to aptamer ratio such as 2:1, it took more prey protein to cause similar quantity of signal

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40 change, thus diminishing the sensitivity of this assay. For that reason, 1:1 ratio of thrombin and aptamer was used in all our experiments. FRET-based 27mer aptamer for thrombin-protein binding. The sequence of the Exosite II-binding 27mer aptamer was adopted from a previous report. 53 050010001500200025003000708090100110120130140150 Fluorescence (A.U.)Time (s) Figure 2-5. Dual-labeled 27mer aptamer for -thrombin/protein interactions. In a solution of mixed 100 nM FQ-27Ap and 100 nM -thrombin, 300 nM AT3 (), 500 nM HirF () or 300 nM AHT () was added at 0 sec. and fluorescence of 6-FAM was continuously monitored. This aptamer was labeled with 6-FAM and Dabcyl similar to FQ-15Ap. With the addition of -thrombin, FQ-27Ap also displayed decreased 6-FAM fluorescence because 6-FAM and Dabcyl at the two ends of the aptamer were brought closer in the quadruplex structure. The relative fluorescence decrease was found to be a little larger than that in the

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41 FQ-15Ap experiments (Figure 2-3). Compared to the noise level, the absolute fluorescence difference between the bound and the unbound FQ-27Ap provided adequate sensitivity for our thrombin-protein interaction study. Different proteins were investigated in a FQ-27Ap/-thrombin bait solution in a similar way as in the FQ-15Ap based assay. The results for HirF and AHT showed slightly decreased signals (Figure 2-5), indicating no displacement of FQ-27Ap took place. The fluorescence reduction could be caused by interactions of thrombin with those two molecules. In contrast, antithrombin III still displayed a gradual increase in 6-FAM fluorescence, meaning that, contrary to a previous report, 57 binding between thrombin and the serpin antithrombin III can also destabilize binding at Exosite II. Again, the slow interaction between AT3 and thrombin caused rather gradual displacement of FQ-27Ap. Fluorescence Anisotropy (FA) Based Aptamer Probes for Protein Interactions. To address some of the unresolved problems in FRET experiments such as how AT3 really binds to -thrombin and what happens between AHT and -thrombin, we developed a complementary strategy based on fluorescence anisotropy. Fluorescence anisotropy is widely used for studying the interactions of biomolecules due to its capability of sensing changes in molecular size or molecular weight. We labeled the thrombin aptamers with only one TAMRA dye at the 3 end to create anisotropy aptamer probes, the 15mer T-15Ap and the 27mer T-27Ap (Table 2-1). The T-15Ap was first investigated for its ability to probe protein interactions. When T-15Ap/-thrombin (1:1) solutions were mixed together, the anisotropy of T-15Ap increased more than 30%. This bait solution was then tested with different prey proteins (Figure 2-6). The anisotropy dropped within seconds upon addition of HirF to the bait solution and remained almost constant after that. This result correlates well with the

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42 result from the FRET-based experiment and may be explained as a quick displacement of the aptamer by HirF at the Exosite I binding site of -thrombin. The anisotropy decreased as a result of the increased concentration of unbound aptamer which had a much lower molecular weight than that of the aptamer-protein complex. The reaction was rapid, indicating a simple binding between HirF and -thrombin through non-covalent bonds. 050010001500200025000.110.120.130.140.150.160.170.180.19 AnisotropyTime (s) Figure 2-6. TAMRA-labeled 15Ap for -thrombin/protein interactions based on fluorescence anisotropy. In a solution of mixed 100 nM T-15Ap and 100 nM -thrombin, 200 nM AT3 (), 500 nM HirF () or 300 nM AHT () was added at 0 sec. and anisotropy of TAMRA was recorded in real time. The AT3 curve showed a different decreasing trend with time. It was rather slow and gradual, similar to the FRET-based result. However, in the FRET assay, while it clearly illustrated that the aptamer was displaced, it did not provide much information about how this displacement took place. There could be several pathways that the AT3/-thrombin interaction might have taken. One of them is that the AT3 molecules would

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43 quickly bind to the active site of -thrombin, and a slow conformational change of -thrombin induced by AT3 binding then caused FQ-15Ap to leave Exosite I. In another pathway, AT3 would slowly attack thrombin and such interaction would force the aptamer to leave thrombin. The FRET-based method could not differentiate between these two mechanisms. On the other hand, using fluorescence anisotropy, if the AT3/-thrombin interaction underwent the first pathway, the increased molecular weight through the binding of AT3 to -thrombin/aptamer complex in the first step would introduce an initial anisotropy increase. Then, the anisotropy would slowly decrease from that point on as the T-15Ap slowly became unbound. However, the real time anisotropy detection of the AT3/-thrombin interaction (Figure 2-6) demonstrated no such initial anisotropy jump. Combined with the result from FQ-27Ap, the anisotropy experiments seemed to better support the second pathway as the mechanism for this protein-protein interaction. The anisotropy approach is shown here to be able to provide insight into the kinetics and mechanisms of the targeted interactions, which will be highly useful in understanding proteins functions. It is our belief that site-directed aptamers enable real-time, sensitive studies on protein-protein interaction. It is interesting to observe that AHT caused an immediate anisotropy increase of T-15Ap when added to the aptamer/-thrombin bait solution (Figure 2-6). While the lack of a decreased anisotropy correlated with the FRET-based result that showed AHT had no effect on binding between the 15mer aptamer and -thrombin, the anisotropy increase suggested the presence of a binding between AHT and -thrombin. Furthermore, this binding happened at a different site than Exosite I, which added extra weight to the aptamer/-thrombin complex. The binding of AHT and -thrombin was further

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44 confirmed using gel electrophoresis (Figure 2-7). One advantage of the anisotropy-based method over the FRET-based method and many other techniques might be that it can differentiate interactions at different binding sites. Figure 2-7. Binding between -thrombin and anti-human thrombin (AHT) confirmed by gel electrophoresis on a 7.5% native Tris-HCl gel. Left lane contained 50 pmole -thrombin. Middle lane had 32 pmole AHT. Right lane had mixture of 32 pmole AHT and 50 pmole -thrombin. Bait solutions containing T-27Ap and -thrombin were also used to probe protein-protein interactions at the Exosite II of -thrombin (Figure 2-8). HirF caused a slightly lower anisotropy change even though it binds to Exosite I. Considering HirF is a rather small molecule (M.W. =~1.5 KDa), the small anisotropy decrease was likely caused by HirF displacing T-27Ap. However, this displacement was much smaller compared to that of T-15Ap. AT3 displayed a gradually decreasing anisotropy as it slowly displaced T

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45 27Ap. In contrast, AHT induced an instant anisotropy increase similar to what was found with T-15Ap, suggesting that AHT does not affect binding at Exosite II and probably binds to a third site of -thrombin other than Exosite I and II. 05001000150020002500300035000.130.140.150.160.170.180.190.200.21 AnisotropyTime (s) Figure 2-8. TAMRA-labeled 27Ap for -thrombin/protein interactions based on fluorescence anisotropy. (A) In a solution of mixed 100 nM T-27Ap and 100 nM -thrombin, 200 nM AT3 (), 500 nM HirF () or 300 nM AHT () was added at 0 sec. and anisotropy of TAMRA was recorded in real time. Quick Evaluation of Binding Constants of Protein-Protein Interactions. In many research areas and biological applications, it is important to not only identify a protein-protein interaction, but also determine how strong the affinity is between the two proteins. The binding affinity of a protein-protein interaction can be represented by the dissociation constant (K d ) of the binding reaction. Typically, for a newly found interaction, no matter what detection technique is used, it is often necessary

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46 to build a calibration curve using various analyte concentrations. K d of the interaction can then be derived for the calibration curve. In a competitive assay, such as described in this work, the interaction of aptamer and its target protein is a known system. Addition of the prey protein may shift the equilibrium of the aptamer/bait protein binding reaction and cause a new signal from the aptamer when a new equilibrium is reached. We found that, based on the known aptamer/-thrombin interaction and equilibrium conditions, it was possible to theoretically calculate the K d of -thrombin/prey protein binding reaction using the signal change occurred when the prey protein was added to the aptamer/-thrombin complex solution. Assume C A molar of T-15Ap aptamer and C T molar of -thrombin are mixed together. When C P molar of prey protein is added to the mixture, it will displace T-15Ap and result in a decreased anisotropy value of r new r new can be represented using the following equation: r A x + r AT (1 x) = r new where r A and r AT are anisotropies of the two fluorescent species of the solution, T-15Ap and T-15Ap/-thrombin complex respectively, and x is fraction of the unbound T-15Ap aptamer. Since r A and r AT are known properties of the aptamer/-thrombin system and r new is the measured new anisotropy, it is easy to find out that: ATAATnewrrrrx Then the concentrations of unbound and bound T-15Ap are: [T15Ap] = C A x [T-15Ap/-thrombin] = C A (1-x) Because the dissociation constant of aptamer/-thrombin reaction (K d/AT ) is already known, then:

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47 [-thrombin] = ]15Ap-T[]thrombin-15Ap/-T[/ATdK Since C T = [-thrombin] + [T-15Ap/-thrombin] + [prey/-thrombin], [prey/-thrombin] = C T [-thrombin] [T-15Ap/-thrombin] Similarly, C P = [prey/-thrombin] + [prey protein], so [prey protein] = C P [prey/-thrombin] Finally, the dissociation constant of -thrombin/prey protein binding reaction (K d/TP ) is given by the following equation: K d/TP = ]thrombin-prey/[]thrombin-[]proteinprey [ It is clear that theoretically, the aptamer signal change induced by competitive binding can be easily applied to elucidate affinity of the protein-protein binding interaction. There is no requirement to take multiple measurements using different prey protein concentrations since aptamer/-thrombin serves as a good reference system. Using a simple computer program, it is possible to routinely calculate protein-protein binding affinity using data obtained from the aptamer-based competitive assay for protein-protein interactions. We demonstrated this capability by calculating K d of -thrombin/HirF binding reaction to be ~190 nM using a reported 15mer aptamer-thrombin K d/AT of 75 nM. 59 This K d is close to what other people have observed (150 nM). 53 Despite the quick and easy evaluation of K d using the above described method, it is important to note that errors in sample handling and fluorescence signal measurements might lead to a considerable amount of uncertainty in the calculated dissociation constants. In this case, multiple measurements may be required. We also notice that there is no information about the stoichiometry of binding and the potential of co-operativity

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48 in any of the binding reactions in our estimation. This simple method for evaluating binding constant can at least be used as a quick estimation in protein-protein interaction studies. Kinetics of Protein-Protein Interactions in Competitive Assays. While the thermodynamic properties of the protein-protein interactions will probably not be affected by the competitive binding of the aptamer, the reaction rates are most likely still dependent on the kinetics of aptamer-protein binding. The detection of protein-protein interactions where the aptamer is displaced consists of two major steps, the dissociation of the aptamer and thrombin, and the association of thrombin and the prey protein. k 1 k -1 Apt + Thr Apt-Thr k -2 k 2 Thr + P Thr-P where Apt, Thr and P are designated to aptamer, thrombin and the prey protein, respectively. The affinities of the aptamer and the prey protein for thrombin can be represented by their binding constants: K Apt-Thr = k 1 /k -1 K Thr-P = k 2 /k -2 One situation that should be considered in the aptamer-based competitive assay is that even though the two binding constants could be very close, there could still be large differences between k -1 and k -2 and k 1 and k 2 In the cases where k -1 << k -2 namely the off rate of aptamer is vastly smaller than that of the prey protein, it may take an

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49 enormously long time to detect a signal change even though thermodynamically the protein should be able to displace the aptamer from thrombin. 0204060801000.00.20.40.60.81.0 Completion of ReactionTime (s) Figure 2-9. Effect of the order of incubation with thrombin on thrombin-protein interaction. 500 nM HirF was first incubated with 100 nM thrombin () and then 100 nM FQ-15Ap was added at time 0 to replace HirF. In another case (), 100 nM FQ-15Ap was incubated with 100 nM thrombin first and 500 nM HirF was added later at time 0. Completion of reactions was monitored using fluorescence changes of 6-FAM. In order to evaluate the possibilities of such false negatives in our assays, it is necessary to compare the reaction rates of aptamer and prey proteins with thrombin. One direct way to do the comparison is to change the order aptamer and the prey protein are incubated with thrombin. In one experiment, the prey protein was incubated with thrombin first and then the aptamer was used to displace the prey protein from the thrombin/protein complex. In another experiment, the order of adding aptamer and prey protein to thrombin was reversed. By comparing kinetic profiles of these two

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50 experiments, it is possible to find out if aptamer binding makes interaction between thrombin and prey protein difficult to take place. HirF was tested along with FQ-15Ap in this way because they compete for the same Excsite I on thrombin. It was found that aptamer replacing HirF was even slower than HirF replacing aptamer (Figure 2-9), meaning off rate of aptamer would not be so slow as to affect thrombin/HirF interaction. Another indirect method was also used to study the effects of aptamer on thrombin/protein interactions. If aptamer binding to and dissociation from thrombin was a much slower process than thrombin/protein interaction, one would expect that changing prey protein concentration would not change the observed rates of thrombin/protein binding in the aptamer-based assay since the prey protein was not in the rate-limiting step of the two steps mentioned earlier. On the other hand, changing aptamer concentration should greatly affect the observed rates since aptamer was in the rate-limiting step. Experiments were conducted to study the thrombin/AT3 interaction. Different concentrations of AT3 were added to thrombin/T-15Ap incubation solution and the anisotropy of the aptamer was monitored as AT3 would displace T-15Ap. The results show a clear dependence of thrombin/AT3 kinetics on AT3 concentration (Figure 2-10A), which contradicts with the assumption that aptamer binding was the rate limiting step. In another study with AHT, T-15Ap concentration was varied to see if the aptamer had any effects on the observed rates of thrombin/AHT reaction even though T-15Ap and AHT were found to bind to different parts of thrombin (Figure 2-10B). The results show no noticeable change in the kinetics, indicating aptamer had no effects on thrombin/AHT binding either.

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51 Figure 2-10. Rate-limiting step in aptamer-thrombin-protein interactions. (A) 100nM T-15Ap and 100 nM thrombin were first incubated. Various amounts of AT3 were added at time 0: () 100 nM; () 200 nM; () 300 nM. (B) Various concentrations of T-15Ap were incubated with 100 nM thrombin: () 50 nM; () 100 nM; () 200 nM. Then 300 nM of AHT was added at time 0. Completion of reactions was monitored using anisotropy changes. 0100200300400500 0.0 0.2 0.4 0.6 0.8 1.0 1.2 Completion of ReactionTime (s)0500100015002000250030003500 -0.1 0.0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1.0 1.1 Completion of ReactionTime (s) A B0100200300400500 0.0 0.2 0.4 0.6 0.8 1.0 1.2 Completion of ReactionTime (s)0500100015002000250030003500 -0.1 0.0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1.0 1.1 Time (s) A B

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52 The protein-protein interactions we studied here have shown to be not affected by the aptamer binding to its target. Compared to two interacting proteins, aptamers are usually much smaller than their target proteins and tend to bind to the targets only through non-covalent forces. They are also less likely to cause induced conformational changes of the target proteins than in protein-protein interactions. Thus it is not surprising that aptamers interacting with their targets would not kinetically interfere with protein-protein detection in our competitive assay. Conclusions Aptamers have great potential in molecular recognition due to their excellent structural stability and exceptional flexibility with various intra-molecular modifications. While most previous work has been focused on using aptamers as probes for direct detection of their target molecules, this work has opened novel applications for aptamers in areas where understanding of the interactions between known proteins and other molecules bears great significance. Many aptamers can be easily labeled with a fluorophore and a quencher to form intra-molecular FRET. Folded conformations of many aptamers have shown to be stabilized by binding to their target molecules. 33;39;40;53 This makes possible a fluorescence signal change of the fluorophore induced by FRET when the aptamer binds to its target. In some cases, target-binding induced FRET can cause up to 90% fluorescence quenching, making FRET-based detection very sensitive. In an alternative approach, FRET can be formed within an aptamer even if the aptamer lacks the necessary conformational changes accompanying the binding to the target molecules. 35 The other detection strategy used in this work, the fluorescence anisotropy, relies on the relatively smaller sizes of aptamers compared to proteins. Aptamers have shown to

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53 be suitable for fluorescence anisotropy based protein studies and detections. 42;60 It is demonstrated here that aptamer based anisotropy probes can provide sufficient signal change for protein-protein interaction study. Both above-mentioned methods allow real time monitoring of protein-protein interactions without any modifications to the interacting proteins. In a recent report, aptamers were used for a similar purpose, RNA ribozymes were required to produce a fluorescence signal change. 48 Aptamers complexed with another molecule may have affected binding affinity toward their targets. As a result, more target protein is necessary to cause enough initial signal change for protein-protein binding study. In fact, a thrombin concentration of 20 times of the ribozyme/aptamer complex was used, 48 compared to the 1:1 molar ratio of -thrombin and aptamer used in this work. With excess bait protein in a competitive assay, a considerable amount of prey protein would be necessary to significantly affect the signal of the aptamer, which may easily lead to false negatives. Using assays directly based on aptamers could preserve aptamers affinity to the proteins and monitor protein-protein interactions with high sensitivity. Our results have shown that two detection methods complement each other. The FRET-based assay relies on direct measurements of sample fluorescence, which makes it highly sensitive and selective. It can also be easily adapted for binding site-specified high throughput protein interaction screening in an array format. On the other hand, fluorescence anisotropy has shown to offer a large amount of information about protein-protein binding that is not readily available using many other techniques including FRET. The aptamer-based competitive assay should be useful for finding protein-binding targets with comparable affinities in a large array of compounds. When detection of

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54 weaker protein-protein binding is desired, it is possible to lower the aptamers affinity towards the target protein by adding, removing or changing bases of the aptamer that are not directly involved in the aptamer/protein binding. This flexibility or tenability makes aptamers more appealing for competitive assays than antibodies. The aptamer competitive assay should also hold the potential for studying interactions between proteins and other molecules such as small organic molecules, DNAs and RNAs. With aptamers being rapidly developed for a growing number of proteins, it is possible to build a large array of aptamers in protein-drug candidate interactions for large scale drug discovery, or in whole cell protein-protein interactions for disease diagnosis and functional proteomics.

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CHAPTER 3 MOLECULAR APTAMERS-BASED AFFINITY CAPILLARY ELECTROPHORESIS FOR PROTEIN-PROTEIN INTERACTIONS We have demonstrated the capability of oligonucleotide aptamers serving as protein probes for real-time protein interaction study. However, except its competence in homogeneous solution systems, aptamer couple with fluorescence detection can also excel in protein analyses using separation-based techniques, particularly capillary electrophoresis (CE). Electrophoresis has been an established method for protein analyses. Traditionally, the mobility shift gel electrophoresis has been used to study DNA-protein interactions, until recent, development of the capillary electrophoresis mobility shift assay and affinity capillary electrophoresis (ACE). 61-63 ACE refers to a collection of techniques in which high-affinity binding probe is used in conjunction with capillary electrophoresis (CE) separation to determine analytes. When coupled with fluorescent labels and laser-induced fluorescence detection (LIF), this immunoassay technique has shown advantages such as high mass sensitivity, rapid separations, simultaneous determination of multiple analytes, and compatibility with automation. 64-66 Antibody-antigen (Ab-Ag) interaction is widely employed in ACE. Both competitive and non-competitive immunoassays canbe used to detect the antigen. 67-69 In a competitive immunoassay, antigen (Ag) is mixed with a fluorophore-labeled antigen (Ag*) and a limiting concentration of antibody (Ab). CE-LIF analysis yields two zones that correspond to the Ag* and the Ab-Ag* complex. The relative intensities of the Ag* and Ab-Ag* allow the quantification of the original 55

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56 concentration of Ag. In the non-competitive assay, a dye-labeled Ab (Ab ) is mixed with Ag. The Ag can be quantified by determining Ag-Ab* after CE separation. Though simple in design, antibody-based ACE has a few limitations. First, it is not always easy to uniformly label either Ab or Ag with fluorophores. Second, for protein analytes, antibodies may not be the ideal ligand in electrophoresis. Compared to many proteins, antibodies are much larger in size, making it difficult to separate Ab* from Ab*-Ag complex due to a small electrophoretic mobility difference between Ab* and Ab*-Ag. Furthermore, charge density on the antibody molecule may very well be similar to that on the protein antigen, which does not help the separation in CE. One way to overcome these disadvantages of the antibody-based ACE is to develop alternative ligands that not only possess the high binding strength to the analyte, but also are easy to label and separate. Aptamers have been successfully used in capillary electrochromatography and affinity chromatography. 70;71 Aptamers, especially DNA aptamers possess several advantages in ACE assays. 72-74 Size of aptamers is considerably smaller than antibodies, and often smaller than the protein analyte. The small size ensures a better separation between the aptamer and the aptamer-protein complex in CE. In addition, aptamers have predictable behavior in electrophoresis as a result of their uniform charge-to-size ratios. Aptamers hold negative charges in a wide range of pH owing to the negative phosphate backbone of nucleic acids. This makes it possible to fine tune the pH of the running buffer to render the protein target neutral or positive for even better separation of the aptamer and complex. The advantages of DNA aptamers that apply to the homogeneous assays are also applicable here in the ACE, such as inexpensive syntheses, easy labeling, and chemical stability for long-time storage.

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57 Kennedy and coauthors have successfully applied DNA aptamers to the quantitative analysis of two proteins (IgE and thrombin) in CE. 72 In their experiments, the fluorophore labeled aptamer and protein mixtures were injected and separated by CE. The peak areas of free and protein-bound aptamer were used for the quantification of the proteins. In this way, the detection limits of IgE and thrombin were 46 pM and 40 nM, respectively. The authors pointed out that the binding constant between aptamer and thrombin was significantly weaker than that of aptamer and IgE, resulting in the higher detection limit of thrombin relative to that obtained for IgE. It was presumably believed that lower binding affinity caused a significant loss of the complex of protein-aptamer by dissociation during the electrophoresis process. Unstable aptamer-protein complexes may completely dissociate during the separation, leading to a very broad peak or no peak corresponding to the complex, which in turn makes it difficult to use aptamers for the quantitative analyses of proteins. In a more recent work, they further investigated the electrophoresis conditions required to successfully detect aptamer-ligand complexes. 74 In their report, the tris(hydroxyamino)-methane-glycine-potassium (TGK) buffer at pH 8.4, minimal column length and a high electric field were required for the successful detection of aptamer-protein complexes. They concluded that these results showed potential for aptamer based ACE. In another study, Krylov and coauthors proposed a new method that allows for the use of low-affinity aptamers as affinity probes in the quantitative analyses of proteins. 73 The method is based on the nonequilibrium capillary electrophoresis of the equilibrium mixture (NECEEM), which allows for the accurate quantitative analysis of proteins even when the aptamer-protein complexes may completely decay during the separation. 75

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58 In this work, we have investigated the effects of an electrophoresis buffer containing different metal ions on the conformation of the 15mer thrombin-binding aptamer. Furthermore, we found that adding an appropriate concentration of poly(ethylene glycol) (PEG) to the aptamer and thrombin mixture might stabilizing the complex of aptamer and thrombin. Finally, we further studied the protein-protein interactions of thrombin with a few anti-thrombin proteins using the fluorophore-labeled aptamers. These studies should provide useful information on using molecular aptamers for protein-protein interactions in ACE. Experimental Section Chemicals and Buffers The 6-carboxyfluorescein (6-FAM) was labeled at the 5 end of aptamer (5-(6-FAM)-GGT TGG TGT GGT TGG-3) obtained from Integrated DNA Technologies, Inc. (Coralville, IA). Thrombin (M.W. ~ 36.7 kDa), human anti-thrombin III (AT III, M.W. ~ 58 kDa) and a monoclonal antibody anti-human thrombin (AHT, M.W.~150 kDa) were obtained from Haematologic Technologies Inc. (Essex Junction, VT). A sulfated hirudin fragment 54-65, Gly-Asp-Phe-Glu-Glu-Ile-Pro-Glu-Glu-Tyr(SO 3 H)-Leu-Gln (HirF, M.W. 1.5 kDa), fluorescein, PEG (M.W. 8 kDa) and poly(N-vinyl-2-pyrrolidone) (PVP, 1.3 MDa) were from Sigma-Aldrich, Inc. (St. Louis, MO). The electrophoresis buffer consisted of 25 mM tris(hydroxy-amino)methane (Tris), 192 mM glycine and 0-10 mM KCl, LiCl, MgCl 2 and BaCl 2 at pH 8.4 was used for separating aptamer. The electrophoresis buffer consisted of 10 mM Tris-HCl pH 8.4 and 15 mM KCl was for quantifying thrombin and studying protein-protein interactions. All solutions were made in the electrophoresis buffer including the stock solutions of protein, aptamer, PEG and

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59 fluorescein (internal standard). All reagents for the buffers were obtained from Fisher Scientific Company L.L.C. (Pittsburgh, PA). Apparatus The basic design of the separation system has been previously described (see Figure 3-1). 76 + High voltage power supplyFocus lensLaserBuffer reservoirFilterObjectivePinholePMT Capillary + High voltage power supplyFocus lensLaserBuffer reservoirFilterObjectivePinholePMT Capillary Figure 3-1. Schematics of the capillary electrophoresis setup.

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60 Briefly, a high-voltage power supply (Gamma High Voltage Research Inc., Ormond Beach, FL) was used to drive electrophoresis. The entire detection system was enclosed in a black box with an HV interlock. The high-voltage end of the separation system was put in a laboratory-made plexiglass box for safety. A 2.5-mW Ar ion laser with 488 nm output (Spectra Physics, Mountain View, CA) was used for excitation. The emission was collected with a 20 X objective (numeric aperture = 0.25). One 520-nm interference filter was used to block scattered light before the emitted light reached the photomultiplier tube (Hamamatsu R928, Hamamatsu Photonics K.K., Hamamatsu, Japan). The amplified currents were transferred directly through a 50-k resistor to a 24-bit A/D interface at 10 Hz (AT-MIO-16, National Instruments, Austin, TX) and stored in a personal computer. Capillaries (Polymicro Technologies, Phoenix, AZ) 50 m i.d. and 365 m o.d. were used for electrophoresis separations with or without PVP coating. Separation of Aptamer Sample prepared in a separation buffer and consisted of 200 nM aptamer and 10 nM fluorescein (internal standard). Samples were injected into the capillary (total length, 35 cm; effective length, 10 cm) hydrodynamically (h = 5 cm) for 10 s. Separation buffer were constituted of 25 mM Tris, 192 mM glycine and 0 -10 mM KCl at pH 8.4. The electrophoresis separation was carried out with an electric field of 285 V/cm. Aptamer-Based ACE In thrombin quantification, the aptamer was mixed with thrombin and fluorescein (internal standard) in the electrophoresis separation buffer (10 mM Tris-HCl pH 8.4 and 15 mM KCl) and incubated for 60 min at room temperature. The final concentrations of aptamer, fluorescein and thrombin were 200 nM, 10 nM and 0-1.0 M, respectively. The

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61 resulting samples injected into the capillary (total length, 50 cm; effective length, 25 cm) hydrodynamically (h = 10 cm) for 10 s. The electrophoresis separation was carried out with an electric field of 350 V/cm. For the quantification of anti-thrombin proteins (AT III, HirF and AHT), aptamers were mixed with thrombin in the electrophoresis separation buffer (10 mM Tris-HCl pH 8.4 and 15 mM KCl) and incubated for 60 min at room temperature. The desired concentrations of anti-thrombin proteins were mixed with aptamer-thrombin complex solutions and incubated for another 60 min. Fluorescein was added to the resulting samples as an internal standard to 10 nM. The final concentrations of aptamer, thrombin and anti-thrombin proteins were 200 nM, 200 nM and 0-10.0 M, respectively. The samples were injected into the capillary (total length, 40 cm; effective length, 25 cm) hydrodynamically (h = 10 cm) for 10 s. The electrophoresis separation was carried out with an electric field of 500 V/cm. At the end of each run, the capillary was rinsed with 0.5 N NaOH for 10 min to remove the protein adsorbed on the capillary. PEG-Assisted Aptamer ACE To study the effect of PEG, aptamers were mixed with thrombin in 10 mM Tris-HCl pH 8.4, 15 mM KCl and 0-10% PEG and incubated for 60 min at room temperature. The final concentrations of aptamer and thrombin were 200 nM. The electrophoresis buffer consisted of 10 mM Tris-HCl pH 8.4 and 15 mM KCl. The resulting samples were injected into the PVP coated capillary (total length, 15 cm; effective length, 5 cm) hydrodynamically (h = 1.5 cm) for 20 s. The electrophoresis separation was carried out with an electric field of 666 V/cm. Similarly, to quantify thrombin and study the interactions of thrombin and anti-thrombin proteins (AT III, AHT and HirF), the desired concentrations of thrombin and anti-thrombin proteins were mixed with aptamer and

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62 aptamer-thrombin complex solutions, respectively. The resulting samples were injected and separated in a PVP coated capillary. The electrophoresis buffer consisted of 10 mM Tris-HCl pH 8.4, 15 mM KCl and the separation was carried out with an electric field of 666 V/cm. At the end of each run, the capillary was rinsed with 5% PVP for 10 min. Results and Discussions Conformation of aptamer It has been reported that a 15-mer thrombin binding aptamer adopts an intramolecular G-quadruplex structure (Figure 1-6) in the presence of K + 77-80 Its affinity for thrombin has been associated with the inhibition of thrombin-catalyzed fibrin clot formation. 81;82 Studies with circular dichroism, temperature-dependent UV spectroscopy, differential scanning calorimetry, isothermal titration calorimetry, capillary electrophoresis, NMR, and mass spectrometry have revealed intramolecular G-quadruplex structures of the 15-mer aptamer in the presence of various metal ions. 77-80;83-85 The G-quadruplex aptamer (G-Apt) is stable both kinetically and thermodynamically because of the tight association of cations with quinine residues. 86-88 The stability of the G-Apt is a very important factor in the detection of thrombin in ACE because thrombin will only bind to the Gquadruplex form of the aptamer. 37;82 Here, we evaluated the stability of the G-Apt in the presence of several metal ions (K + Li + Ba 2+ and Mg 2+ ). Figure 3-2 shows the electropherograms for the aptamer with increasing concentrations of KCl. The aptamer was separated into two peaks when we first used a K + -containing running buffer. That was when we started to think that the two conformations of the aptamer might be able to be separated in CE even though a few related reports did not mention that. 72;74

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63 Figure 3-2. Capillary electrophoresis traces of thrombin binding aptamer in the presence of 25 mM Tris, 192 mM glycine and various concentrations of KCl. Separation buffer was composed of 25 mM Tris, 192 mM glycine and 0 mM (A), 0.25 mM (B), 0.5 mM (C), 1.0 mM (D), 5.0 mM (E), 10.0 mM (F) KCl at pH 8.4. Samples were prepared in a separation buffer and contained a final concentration of 200 nM aptamer and 10 nM fluorescein (internal standard). The samples were injected into the capillary (total length, 35 cm; effective length, 10 cm) hydrodynamically (h = 5 cm) for 10 s and an electric field of 285 V/cm was applied to drive the separation. To confirm this hypothesis, running buffers without K + or Na + were tested. It can be seen in Figure 3-2 that the aptamer came out a single peak at the time where the first

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64 peak showed up in the K + -containing buffers. Since G-quadruplex structure is not stable without metal ions, the single peak was believed to be primarily the linear form of the aptamer. We suspected that there might have been an unstable G-Apt based on the slight tailing of the single aptamer peak. The first and second peak except the internal standard peak, in Figure 3-2 B to F, corresponded to the linear aptamer (L-Apt) and the G-Apt, respectively. The peak area ratio of the G-Apt to L-Apt form increased with the increased concentration of K + This result agreed with other reports that the stability of the G-Apt was dependent upon the concentration of K + 78;79;83 Furthermore, the increase in the K + concentration was found to improve the resolution between the L-Apt and G-Apt peak. Figure 3-3 shows that other metal ions, such as Li + Mg 2+ and Ba 2+ were not capable of stabilizing the G-Apt or effectively separating the G-Apt from the L-Apt. It was reported that cations with an ionic radius in the range of 1.3-1.5 fit well within the two G-quartets of the complex. 79 Our results were in agreement with the reports that K + Rb + NH + Sr 2+ and Ba 2+ are able to form stable cation-Apt complexes, while Li + Na + Cs + Mg 2+ and Ca 2+ only form weak complexes. Among the four metal ions (K + Li + Mg 2+ and Ba 2+ ) chosen in our experiments, the aptamer displayed a stable G-Apt peak with K + or Ba 2+ present. As shown in Figure 3-3, as little as 2.5 mM Ba 2+ could effectively stabilize the peak of the G-Apt. However, similar to Mg 2+ Ba 2+ associates with high affinity to the phosphate backbone of oligonucleotides and decreases the mobility of the aptamer. This was demonstrated by the change in the aptamer peak position relative to the fluorescein peak. On the other hand, the decrease of the mobility of fluorescein indicated a severely reduced electroosmotic flow (EOF), which led to peak broadening, increased separation time and decreased resolution. For this reason, Ba 2+ was

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65 certainly not suitable for the study of protein-DNA and protein-protein interactions in ACE. Instead, KCl was chosen to be added to sample matrix and electrophoresis buffer in further experiments. Figure 3-3. Capillary electrophoresis traces of thrombin binding aptamer in the presence of 25 mM Tris, 192 mM glycine and various other metal ions. Separation buffer were composed of 25 mM Tris, 192 mM glycine, and 10 mM LiCl (A), 2.5 mM MgCl2 (B), 2.5 mM BaCl2 (C) at pH 8.4. Samples prepared in a separation buffer and contained a final concentration of 200 nM aptamer and 10 nM fluorescein (internal standard). Other experimental conditions same as in Figure 3-2.

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66 Quantification of Thrombin As mentioned previously, literature reports had pointed out that the unstable aptamer-protein complex would undergo dissociation in the electrophoresis process. 72-74 This caused broadening or disappearance of the complex peak unless hydrodynamic flow was used to force the complex through the cappillary. 72 Although the flow-assisted method allows quantification of the aptamer-protein complex, it is inconvenient and may eliminate some of the advantages of CE. A relatively long capillary (effective length 25 cm) was used in our experiments. The advantage of a long capillary was that there was less interference with the peak of the G-Apt by the aptamer-thrombin peak (Apt*Thrmb) because the complex of aptamer-thrombin would almost completely decay in a long capillary. Therefore, we were able to focus on the peak of G-Apt for quantification. Figure 3-4A and B compare the electropherograms obtained for 200 nM aptamer without and with 200 nM thrombin in the sample. Fluorescein was used as an internal standard (IS) to correct variations in the injection volume. The electrophoresis buffer was further optimized from before (25 mM Tris, 192 mM glycine and 10 mM KCl at pH 8.4, Figure 3-2F) to 10 mM Tris-HCl added 15 mM KCl at pH 8.4. In Figure 3-4A, the peak area ratio (G-Apt/L-Apt) was calculated to be 2.97, which was significantly higher than that in Figure 3-2F (1.09). The electropherogram with thrombin (Figure 3-4B) shows a large decrease in the free G-Apt peak which can be attributed to the binding of the G-Apt to the thrombin. A calibration curve was constructed (Figure 3-4C) based on the peak area of the free G-Apt as a function of thrombin concentration. The curve is linear up to 200 nM thrombin and a 1:1 binding ratio between the aptamer and thrombin is clearly demonstrated. The limit of detection (LOD), which was calculated as the concentration yielding a signal change 3 times the peak to peak noise, was 9.8 nM, and was lower than

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67 previous reported 40 nM. 72 We suspect that the main reason is that the quantification in the literature did not differentiate between G-Apt and L-Apt, while we focused on the peak area change of the G-Apt, which might have resulted in a higher sensitivity. Figure 3-4. Detection of thrombin using aptamer-based ACE. Electropherograms obtained for 200 nM aptamer with 0 (A) and 200 nM thrombin (B). Calibration curve constructed using samples containing 200 nM aptamer with various concentrations of thrombin (C). The electrophoresis separation buffer was 10 mM Tris-HCl and 15 mM KCl at pH 8.4. Samples prepared in separation buffer and consisted of a final concentration of 200 nM aptamer, 10 nM fluorescein (internal standard), and 0-1.0 M thrombin. The samples were injected into the capillary (total length, 50 cm; effective length, 25 cm) hydrodynamically (h = 10 cm) for 10 s and an electric field of 350 V/cm was applied to drive the separation. Peak areas were corrected for variations in injection volume by dividing by the area of the internal standard peak.

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68 Determination of Dissociation Constant (Kd) The value of K d was determined under the conditions of the experiments. The equilibrium concentration of free G-Apt is proportional to the area of the G-Apt peak (A g eq ). 73 [G-Apt] eq = cA g eq (1) where c is a constant. The equilibrium concentration of the complex (G-Apt*Thrmb) is equal to: [G-Apt*Thrmb] eq = [G-Apt] o [G-Apt] eq = c(A g o A g eq ) (2) where A g o is the peak area G-Apt without thrombin. The ratio R of the two equilibrium fractions: R = [G-Apt] eq / [G-Apt*Thrmb] eq = A g eq / (A g o A g eq ) (3) The knowledge of ratio R is sufficient for the determination of K d : K d = {[thrombin] o (1+R)-[G-Apt] o } / (1 + 1/R) (4) where [thrombin] o and [G-Apt] o are the initial concentrations of thrombin and G-Apt, respectively. The fluorescence intensities of aptamer were found to be unchanged on a fluorometer under different concentrations of KCl, indicating the conformational change of the aptamer did not affect the fluorescence intensity of the labeled 6carboxyfluorescein. As a result, the [G-Apt] o could be calculated from Figure 3A by the areas of peak L-Apt (A o ) and G-Apt (A g o ) [G-Apt] o = [Apt] o A g o / (A o + A g o ) (5) Based on six experiments with different concentrations of thrombin and aptamer, the value of K d (G-Apt*Thrmb) was calculated using Equations 4 and 5 to be 20 nM. This value is smaller than those obtained by Kennedy et al. (450 nM) and Krylov et al.

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69 (240 nM). This can be explained by the fact that the binding affinity measured here is between the G-Apt and thrombin, which is supposed to be stronger than that between thrombin and the overall aptamer (G-Apt + L-Apt). Competitive Assay Thrombin has two positive-charged sites termed Exosite I and II on the opposite sides of the protein. 89 Exosite I was found to bind to fibrinogen 50 and hirudin 90-92 while Exosite II binds to heparin and a monoclonal antibody 93 Two different aptamers have been identified that have high affinity and selectivity for thrombin. The first one is a 15-mer single-stranded DNA aptamer (in this work) which was reported to bind to the fibrinogen-binding site of -thrombin, namely Exosite I. 52 The other DNA aptamer, with a 27-mer backbone length, was determined to bind to the Exosite II of -thrombin. 53 Both aptamers were found to adopt a G-quartet structure when bound to -thrombin. 53;94 In the previous chapter, we used the complex of aptamer-thrombin to probe thrombin-protein interactions in a competitive assay where the binding of the aptamer to thrombin was altered by a second protein that interacts with thrombin. Two signal transduction strategies, fluorescence energy transfer and fluorescence anisotropy, have been designed to study the interactions of thrombin with different proteins using two aptamers specific for two binding sites on -thrombin. 95 Here, we have further demonstrated the results by aptamer based ACE. As shown in Figure 3-5, the increasing concentration of AT III caused an increase in the G-Apt peak area as expected. This result agreed with another report that the binding of AT III to thrombin may cause a conformational change in thrombin that rendered the binding with the aptamer at Exosite I unstable. 57 In Figure 3-6, the reaction between AT III and thrombin was monitored in a close-to-real-time fashion. The reaction of thrombin and AT III was completed within 10 min, and the result agreed

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70 with our previous work. 95 A calibration curve was made based on the peak area of free G-Apt vs. AT III concentration (Figure 3-7). The curve is almost linear up until 200 nM AT III and a 1:1 binding ratio between the thrombin and AT III was displayed. Based on these data, the LOD was calculated to be 2.1 nM. Figure 3-5. Analyses of AT III-thrombin interaction using aptamer-based ACE. Electropherograms obtained for 200 nM aptamer with 200 nM thrombin and varies concentrations of AT III. In electropherograms A-H, AT III concentrations were 0, 5, 10, 20, 50, 100, 200 and 400 nM respectively. Aptamers were mixed with thrombin and incubated for 60 min at room temperature. The desired concentrations of AT III were mixed with aptamer-thrombin complex solutions. The resulting samples were added fluorescein as an internal standard to 10 nM and incubation another 60 min. Separation was carried out at a constant electric field of 500 V/cm. Other conditions as in Figure 3-4.

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71 Figure 3-6. Using thrombin binding aptamer to monitor thrombin/AT III interaction. In a solution of mixed aptamer and thrombin incubated in electrophoresis buffer for 60 min. Then AT III were mixed with the aptamer/thrombin complex solution and incubated for another 60 min. The final concentration of aptamer, thrombin and AT III were 200 nM. After the resulting sample incubation for different time, rapid inject into capillary and the separation were carried out at a constant electric field of 500 V/cm. Other conditions the same as in Figure 3-4. Figure 3-7. Quantification of AT III-thrombin interaction. Calibration curve constructed using samples containing 200 nM aptamer, 200 nM thrombin and various concentrations of AT III (0-1.0 M). Peak area of G-Apt was corrected for variations in injection volume by dividing by the area of the internal standard peak. Other conditions as in Figure 3-4 and Figure 3-5.

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72 The addition of another antibody, 400 nM AHT (2 times concentration of thrombin), caused no significant change in the peak of the G-Apt (data not shown). While this result indicates that AHT does not compete with the aptamer for the Exosite I of thrombin, it does not mean that AHT does not bind to thrombin at a different site. More experiments were done to demonstrate this point, and the results are presented later in this paper. A sulfated fragment of hirudin that contained the C-terminal 13-residue (HirF) instead of hirudin was used for studying binding with thrombin. Although the K d of HirF-thrombin (150 nM) at pH 7.4 is similar to the reported K d of aptamer-thrombin (200 nM) and both HirF and aptamer bind to the same site (Exosite I) of thrombin, HirF caused no significant change in the peak of the G-Apt even when the added concentration was as high as 10 M (50 times concentration of thrombin). It has been reported that binding of thrombin to hirudin (65 amino acids) and some derived hirudin fragments strongly depends on pH. 96 The optimum pH for the interaction between hirudin and thrombin was found to be between pH 7.5 and pH 8.0. The K d value increased at higher pH values, and the plot of log K d against pH displayed an asymptotic slope of -2 in the alkaline pH range. As a result, the sample and electrophoresis buffer at pH 8.4 in our experiments may also cause a much higher K d between HirF and thrombin than reported in literatures. On the other hand, the relative lower salt concentration of buffer used in this experiment may also have an impact on the binding affinity of HirF. Effect of PEG on Aptamer-Thrombin Complex in CE Unlike AT III, which can displace G-Apt from a G-Apt*Thrmb complex and cause changes in the area of the G-Apt peak, the thrombin antibody AHT would not affect G-Apt/thrombin binding. It is necessary to observe the peak of the G-Apt*Thrmb complex

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73 and analyze its mobility shift to study the interaction between AHT and thrombin. To resolve the problem of complex dissociation during CE, a short PVP coated capillary (effective length 5 cm) was used to shorten the analysis time and prevent the protein adsorption on the capillary. However, with a total separation time less than 50 sec, peak broadening was still not improved (Figure 3-8A). There have been reports suggesting that linear polymers can promote the cage effect which stabilizes protein complexes during electrophoresis. 97-99 In addition, polymers in electrophoresis buffers may aid the separation by interacting with solutes and capillary walls to prevent adsorption. Moreover, polymer in the region where the complexes have dissociated, may hinder further separation of the two components, and lead to an enhanced probability for a re-association. The dissociation step may also be slowed down by the polymer, if the dissociation requires complex-complex interactions. Finally, polymers may cause macro-crowding effect that increases local concentrations of analytes that cause lower level of dissociation of complex. Given all the background information, we tried adding PEG to the sample matrix. As shown in Figure 3-8 B-F, addition of PEG clearly revealed the peak of the G-Apt*Thrmb complex. The L-Apt and G-Apt can not be separated due to the fact that the experiments were done in a very short time. However, it did not effect the quantification of thrombin and AT III by the complex peak of G-Apt*Thrmb. The optimum concentration of PEG was found to be 2.0%. A higher concentration of PEG would cause peak broadening, probably because that the collisions of aptamer and protein molecules into the polymer network may have induced distribution of the migration rates of those molecules.

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74 Figure 3-8. Effect of PEG on the stability of G-Apt*Thrmb complex. Aptamers were mixed with thrombin in 10 mM Tris-HCl at pH 8.4, 15 mM KCl and 0% (A), 0.5% (B), 1.0% (C), 2.0% (D), 5.0% (E) and 10% (F) PEG, incubated for 60 min at room temperature. The final concentrations of aptamer and thrombin were 200 nM. The electrophoresis buffer was 10 mM Tris-HCl, 15 mM KCl at pH 8.4. The resulting samples were injected into the PVP coated capillary (total length, 15 cm; effective length, 5 cm) hydrodynamically (h = 1.5 cm) for 20 s. The electrophoresis separation was carried out with an electric field of 666 V/cm.

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75 Figure 3-9. Analyses of thrombin and thrombin-AT III interaction using PEG-containing sample matrix. Each sample contained a final concentration of 200 nM aptamer and 0 (A), 50 (B), 200 nM (C) thrombin. In figure E-G, each sample contained a final concentration of 200 nM aptamer, 200 nM thrombin, and 50 nM (E), 100 nM (F), 200 nM (G) AT III. Figure D and H are the calibration curves constructed with various concentrations of thrombin and AT III respectively. The sample matrix consisted of 10 mM Tris-HCl, 15 mM KCl and 2% PEG at pH 8.4. The electrophoresis buffer was 10 mM Tris-HCl, 15 mM KCl at pH 8.4. Other conditions as in Figure 3-5 and Figure 3-8.

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76 Figures 3-8 A-C show the increased G-Apt*Thrmb to free aptamer peak area ratio with increasing thrombin. The calibration curve in Figure 3-9 D has a linear range up to 200 nM thrombin. The LOD of thrombin is 10.9 nM. In a similar way, AT III was quantified using the G-Apt*Thrmb based competitive assay (Figure 3-9 E-H). The electropherograms with increasing AT III have shown a decreased peak of G-Apt*Thrmb and an increased peak of free aptamer. These results once again confirmed the displacement of G-Apt by AT III in CE. The LOD of AT III was estimated at 21.2 nM. Aptamer-Based Mobility Shift Assay for Thrombin-AHT Interaction The substrate specificity of thrombin is regulated by binding of macromolecular substrates and effectors to Exosites I and II. 92 Exosites I and II have been reported to be linked allosterically, such that binding of a ligand to one exosite results in nearly total loss of affinity for ligands at the alternative exosite, whereas other studies support the independence of the interactions. 92 Previous results in bare capillaries revealed that AHT had no effect on binding between the aptamer and thrombin. In order to study the interaction between thrombin and AHT in ACE, it might be helpful to analyze the mobility change of G-Apt*Thrmb complex with the addition of AHT. Using the PEG-assisted ACE, electropherograms were obtained (Figure 3-10 A-D) and clearly showed changes in the migration time of the G-Apt*Thrmb complex as AHT concentration was varied. We further optimize the concentration of PEG in this experiment, and found 2% PEG was again the best. As shown in Figure 3-10 B-D, this mobility shift can be attributed to the increased overall molecular mass of the G-Apt*Thrmb*AHT binding complex. Control experiments revealed that no mobility shift was observed when 200 nM aptamer was mixed with only 400 nM AHT.

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77 Figure 3-10. Binding between G-Apt*Thrmb and anti-human thrombin (AHT) confirmed by capillary electrophoresis. In a solution of mixed aptamer and thrombin incubated in 10 mM Tris-HCl, 15 mM KCl and 2% PEG at pH 8.4 for 60 min. Then AHT were mixed with the aptamer/thrombin complex solution and incubated for another 60 min. Each sample contained a final concentration of 200 nM aptamer, 200 nM thrombin and 0 nM (A), 50 nM (B), 100 nM (C) and 200 nM (D) AHT. Other conditions as in Figure 3-8.

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78 Figure 3-10B (aptamer 200 nM, thrombin 200 nM and AHT 50 nM) displays an unresolved small peak (arrow point and the migration time of that peak is equal to that of the second peak in Figure 3-10D (aptamer 200 nM, Thrmb 200 nM and AHT 200 nM). This result reveals that only a small fraction of G-Apt*Thrmb binding with AHT when the concentration of AHT is lower than G-Apt*Thrmb. The highly broadened peaks in Figure 3-10 B-C indicate the shift from the G-Apt*Thrmb complex to the G-Apt*Thrmb*AHT complex. In addition, the dissociation constant of AHT-thrombin is not very low at 14 nM, 100 which attributed to the dissociation of G-Apt*Thrmb*AHT complex at low AHT concentrations. Even though quantification of AHT is difficult, the interaction between thrombin and AHT, and the fact that AHT and G-Apt bind to different sites of thrombin have been clearly revealed using this mobility shift assay in ACE. Conclusions In this work, we have demonstrated that the 15-mer thrombin-binding DNA aptamer adopts two different forms in the presence of K + or Ba 2+ and only the G-quadruplex form can bind thrombin to form a complex. Binding between aptamer and proteins is thus highly dependent on the conformation of the molecular aptamers. The presence of thrombin and Antithrombin III only affected the G-Aptamer peak in affinity capillary electrophoresis. The G-Aptamer based CE analysis showed a higher binding affinity between G-Aptamer and thrombin. As a result, a better detection limit of thrombin could be achieved. The aptamer-based competitive affinity capillary electrophoresis assay has been also applied to quantify Thrombin/Antithrombin III interaction and to monitor this reaction in real time. We have also shown that a mobility shift based affinity capillary electrophoresis assay, using poly(ethylene glycol) in the

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79 sample matrix, can be used to study the interactions between thrombin and proteins that do not displace G-Aptamer binding property with Exosite I site of the thrombin. We believe that oligonucleotide aptamers possess advantages over other protein ligands in affinity capillary electrophoresis, and the aptamer-based ACE assay can be an effective alternative approach for studying protein-protein interactions and for analyzing binding site information and binding constants.

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CHAPTER 4 NUCLEASE-RESISTANCE OF TELOMERE-LIKE SINGLE-STRANDED OLIGONUCLEOTIDES MONITORED IN LIVE CELLS BY FLUORESCENCE ANISOTROPY IMAGING Introduction Fluorescence Techniques for Monitoring Intracellular Biointeractions One of the major challenges of life science is to understand roles of the vast amount of biomolecules in cells. An ideal way to study cell functions would be monitoring interactions between biomolecules in live cells. Most cell imaging techniques are based on detection of fluorescence signals generated from fluorescent tags that are linked to the molecules of interesting. The development of fluorescent proteins, such as green fluorescent protein (GFP) has made it easy to fluorescently tag proteins for intracellular imaging. 101-103 GFP has been used to monitor cellular gene expression 102 and protein traffiking and localization. 104;105 To detect intracellular interactions between proteins, variants of GFP that have different fluorescence spectra are used to form a fluorophore pair for fluorescence resonance energy transfer (FRET). 106 The interaction between proteins causes changes in efficiency of energy transfer and thus in the ratio of acceptor to donor fluorescence signals. Although FRET-based techniques are powerful tools in imaging of cellular bio-interactions, they have certain limitations. Compared to assays using a fluorophore and quencher pair, FRET based on GFPs tends to have higher fluorescence background, thus makes small signal changes caused by low level of target less appreciable. To overcome this problem, careful selection or design of the GFP pair needs to be done to obtain low 80

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81 acceptor background. In addition, photo-bleaching of the two fluorophore at different rates may cause false change in the ratio of acceptor to donor emissions. Lastly, for protein-protein interaction study, often the two interacting proteins need to be fused with GFP and both host protein and GFP should remain functional. Thus, GFP-based FRET assay should serve better for biological systems where two interacting proteins are molecularly well characterized. 107 As a complementary technique to GFP-based FRET assays, we think that fluorescence imaging based on fluorescence anisotropy (FA) may hold great potential for intracellular bio-interaction study. The fluorescence anisotropy of a fluorophore reflects the molecules ability to rotate in its micro-environments. FA is dependent on things that affect the rotational diffusional movements of the fluorophore such as the size and mass of the molecule the fluorophore is attached to, viscosity of the solution and the fluorescence lifetime of the fluorophore. Since most biological events inside cells induce changes in molecular weight of involved molecules, e.g. binding of two proteins results in a heavier complex, it is theoretically possible to label one molecule with a fluorophore and monitor its interaction using anisotropy. Traditionally, measurements of anisotropy have been done mainly in homogeneous solutions where no localized anisotropy information is available. By combining florescence imaging and anisotropy measurements, bio-interactions in heterogeneous samples, such as live cells, can be monitored. Fluorescence anisotropy imaging has been reported in study of cells 108 and single molecules 109 based on conventional fluorescence microscope. Here we describe an anisotropy imaging system built upon on a confocal microscope. Besides commonly known advantages of confocal microscopes over conventional ones, such as better

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82 resolution and 3-D imaging capability, the multi-channel image acquisition capability of a confocal microscope enables simultaneous acquisition of two images representing two perpendicular polarization states of the fluorescence emission. In addition, the pixel-to-pixel scanning scheme of confocal makes alignment of the two polarization images much less of an issue for future anisotropy image calculation. During the development of our setup, a few groups also chose confocal microscopes to build their anisotropy imaging systems on, 110-112 probably due to the same reason. We used our anisotropy imaging setup to monitor digestion of single-stranded DNAs by nucleases inside live cells. Our interest lies in finding out how compositions and structures of DNAs affect their ability of resisting cellular nuclease degradation. Telomere and Its Presence in Live Cells One type of DNA sequence that belongs to something called telomeres is particularly interesting in our study of nuclease-DNA interaction. Telomeres are the end of eukaryotic chromosomes. The length of the telomeres varies greatly between different species, from 20bp to 150kbp per telomere. 113 They are composed of simple and highly repetitive DNA sequences, with one of the two telomeric DNA strands a GT-rich sequence. The GT-rich repeats are not the same for different species, for example, human and mouse have a TTAGGG repeating unit, while it is TTGGGG for Tetrahymena. 114 Telomeres have been found to carry important biological functions. One of them is believed to be the protection of chromosomes. During DNA replication, short RNA primers are employed to initiate DNA synthesis. Removal of the terminal primers at the end of the replication always leaves a small region of single-stranded DNA (ssDNA) that is not replicated. 115 It has been shown that length of telomeric DNA in human fibroblasts shortened during cell aging in vitro. 116 Had it not been the repetitive telomere sequence,

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83 the genome DNA would have been damaged from the very first round of replication. On the other hand, normal cells can only replicate certain times until the telomere end is completely lost. In this case, the telomeric sequence ensures that cells can not be immortal. Compared to normal cells, cancer cells can replicate indefinitely, thus grow out of control. The reason behind this is believed to be an unusual DNA polymerase called telomerase. 117 In one report, activity of the telomerase enzyme was detected in most human tumor samples and in none of the normal tissues. 118 In another report, introduction of telomerase-encoding genes into telomerase-negative cells evidently extended their normal life-span. 119 This clear relationship between telomere, telomerase, and cancers has inspired great interest in this line of research. One of the early telomere papers showed that the G-strand of the telomeric DNA was longer than the C-strand in Oxytricha, creating a G-rich 3-end overhang. 120 The size of the overhang was found to be about 50 nucleotides in mouse and human telomeres. To understand why the single G-strand can be maintained without inducing DNA-damage response such as degradation by nucleases, different models have been proposed. The classical view is that the G-strand is protected by proteins that bind tightly to single-stranded DNA. 114 A new model found that the G-rich single strand can tuck back into the double strand region of the telomere to form a closed loop structure called T-loop, which avoids exposure of the DNA end to cellular enzymes. 121 The T-loop model explains very well why the termini of telomeres are stable in cell nuclei. However, one may ask why the repeat unit of telomeric DNA is always G-rich instead of any random sequence that can also form the T-loop. It is well known that G-rich telomeric DNAs in many species form parallel four-stranded structure called G

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84 quadruplex by bonding guanines to each other. 122;123 In this work, we tried to study whether the special structure of the telomere-like single-stranded G-rich oligonucleotides contributes to the stability of telomeric overhang in cell nuclei. Fluorescence anisotropy was chosen to study the digestion process over other approaches such as FRET. A FRET design may be able to generate a fluorescence increase when the dual-labeled probe is cleaved by the nucleases. However, further digestion of the cleaved pieces will not induce any more signal change, thus FRET can only monitor the first cut. Moreover, many non-specific bind of proteins to the FRET probe might also cause a conformational change of the DNA, leading to false positives. In contrast, digestion of a DNA molecule can be monitored by anisotropy until the smallest fragments. These is also likely no other process that can decrease the anisotropy of the dye-labeled DNA. Lastly, anisotropy is a ratiometric measurement that is less affected by photo-bleaching and system variations. Experimental Section Fluorescence Anisotropy Imaging (FAI) System The FAI system was built on an Olympus FV500-IX81 confocal microscope (Olympus America Inc., Melville, NY) (See Figure 4-1). The emission of fluorescent samples would go through a few dichroic mirrors before entering a detection channel. A flat polarization beam splitter (PBS) (MOXTEK, Inc., Orem, UT) replaced a dichroic mirror in a way that emission of one polarization state would be reflected in one detection channel while the emission of the other perpendicular polarization state would go though and be reflected by a mirror into the next channel (Figure 4-1). Two bandpass filters for emission of TAMRA dye were placed in the two channels right before the PMTs. One of them was a BP560-600 filter that came with the microscope, and the other was a

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85 580DF30 from Omega Optical, Inc. (Brattleboro, VT). The two filters have similar transmission profiles. Polarization beam splitterMirrorIIPinholePMT 1PMT 2ImgImg Filter ObjectiveLaserSample Anisotropy imageScanning mirrorsExcitation beam splitter Polarization beam splitterMirrorIIPinholePMT 1PMT 2ImgImg Filter ObjectiveLaserSample Anisotropy imageScanning mirrorsExcitation beam splitter Figure 4-1. Schematics of the FAI system.

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86 A 5 mW 543 nM He-Ne laser was the excitation for TAMRA throughout the experiments. Usually only 20-30% of the laser power was used. The objective used for all the experiments was a PLAPO60XO3PH 60x oil immersion objective with a numerical aperture of 1.40 from Olympus (Melville, NY). Scanning of the samples was controlled by the Olympus FluoView program. Images of 512x512 pixels were taken simultaneously in the two polarization channels. The scanning rate was chosen at slow level (~2 s per image) to obtain lower noise level. Noise was further reduced by taking four images for each measurement and recording the average of the four. This could be programmed on the computer. The dynamic range of the confocal signal was 0 to 4095. The signal level could be controlled by a few parameters such as voltage of the PMT, PMT gain (multiplying signal by a number), and laser power. We avoided using the latter two parameters because the gain would increase noise and high laser power could cause photo-bleaching of the dye. The two voltages applied on the two PMTs were always identical. During the monitoring of dye-labeled DNA inside live cells, when there was a drop in the intensity level, the both PMT voltages were increased to keep the signal level in the range of 2000 to 3500. There would be slight increase of background but that could be easily deducted during image processing. When to acquire images during monitoring of the cell was manually controlled. Time between two acquisitions could range from 2 to 30 minutes. For different cells, the time of the data points were different. In an effort to determine the G-factor of the FAI system in calculation of anisotropy, a pure TAMRA solution was used. Since TAMRA is a small organic

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87 molecule (M.W.=430.45), its anisotropy in water should be very close to zero. Two polarization images of the TAMRA solution was taken on the FAI system and the intensities were averaged separately and designated I VV and I VH Considering the equation for anisotropy calculation: r = (I VV -GI VH ) / (I VV +2GI VH ) Apply r = 0, and the I VV and I VH values to the equation, we obtained a G-factor value of 0.61 for our FAI setup. This G-factor was used for all the experiments. The Image Processing Program All the raw images obtained from the Olympus software were then processed in the ImageJ program ( http://rsb.info.nih.gov/ij ) to calculate anisotropy images using a homemade software plug-in. What the plug-in did was to read the two polarization images, find the background and noise levels, and subtract the images with the background value. Then pixel by pixel, the plug-in would try to calculate an anisotropy value for each pixel using data from the two raw images. Anisotropy would be calculated only when the intensity from both images at that pixel was at least 3 times higher than the noise level, otherwise, an anisotropy of zero was given to that pixel. After doing this for the whole image, an anisotropy image was produced and recorded. With this plug-in we programmed, the whole process of calculating anisotropy images could be easily done at a click of a button. Cell Culturing The breast cancer cell line MDA-MB-231 (ATCC, Manassas, VA) was cultured according the providers instructions. Cells for injection experiments were cultured in the glass bottom culture dishes from MatTek Corporation (Ashland, MA). The thin glass bottom would not interfere with fluorescence measurements as plastic would.

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88 Cell Injection Using Electroporation The electroporation system was built upon a Grass S44 stimulator (Astro-Med. Inc., West Warwick, RI) which was capable of msec voltage pulses. Two regular electrical wires were connected to the two termini on the stimulator. The ends of the two wires were linked to platinum wires as two electrodes. A Eppendorf Femtotip from Brinkmann Instruments, Inc. (Westbury, NY) was used to actually get close enough to the cell membrane for the injection. The very end of the tip was less than 1 m in diameter. The tip could be screwed in a homemade hollow pipe which was secured on a PCS-6000 motorized micromanipulator from EXFO Burleigh Products Group Inc. (Victor, NY) which wascapable of less than 1 m movement. The micromanipulator was stationed by the stage of the microscope and could move the attached pipe and Femtotip precisely to where the cells were located. Before putting on the pipe, the Femtotip was loaded with 3-5 L of DNA samples to be injected. When the tip was screwed into the pipe, the platinum wire attached to the cathode was introduced into the tip and immersed in the sample solution through the pipe. The platinum wire of the anode was simply place in the cell dish. After all these procedures, the injection tip was moved close to the cell of interest using the micromanipulator under the 60x objective. Once the tip was in contact with the cell membrane, we would manually switch on the stimulator for a short second. The voltage pulses would produce small pores on the membrane and guide DNA molecules into the cell. Concentration of DNA sample was usually in the range of 5-10 M. Materials All DNA samples were synthesized and purified by GenoMechanix, LLC (Gainesville, FL). A buffer that mimics physiological conditions was used for solution

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89 tests. It contained 20 mM Tris-HCl at pH 7.5, 150 mM NaCl, 5 mM KCl, 1 mM MgCl 2 1 mM CaCl 2 and 5% (V/V) glycerol. All the chemicals were from Sigma-Aldrich (St. Louis, MO) unless otherwise specified. Results and Discussions Measuring Anisotropy of Standard Samples The Olympus FV500-IX81 confocal microscope with polarized laser source was modified using a polarization beam splitter (PBS). While most commercially available PBSs are made into a cube form, they can not fit into the holder designed for emission dichroic mirrors in the scanning unit. Instead, a flat PBS was placed in the holder and properly aligned so that lights with polarization states same as and perpendicular to that of the excitation laser can be separated into two detection channels. Two band pass filters were chosen for TAMRA dye (excitation=560 nm, emission=580 nm). They were placed in front of the PMTs in the two channels. This configuration was designed to measure anisotropy of only TAMRA. As the sample is scanned by the scanning unit, two images containing different polarization states of the emission can be obtained simultaneously. These images are then processed in the ImageJ computer program ( http://rsb.info.nih.gov/ij ) using a homemade plug-in. The plug-in uses the following equation for anisotropy calculation in pixel level, r = (I VV -GI VH ) / (I VV +2GI VH ) where the subscripts V and H refer to the orientation (vertical or horizontal) of the polarization state, with the first subscript indicating the position of the excitation polarization and the second for the emission polarization. G is the G-factor of the confocal system, which represents the ratio of the responses of the optical components to vertically and horizontally polarized lights. Calculation of the two whole images results

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90 in an anisotropy image with the same size. Anisotropy can then be read out from one pixel or as average value from a region of interest. AB AB Figure 4-2. Fluorescence images obtained from I VH (left) and I VV (right) channels of the FAI system. A) Images from 200 nM TAMRA solution containing 20% (V/V) glycerol. B) Images from 200 nM TAMRA solution containing 60% (V/V) glycerol. The area on the upper right corner of the images was used for anisotropy calculation. To test if the FAI system had any response to anisotropy changes, TMARA solutions with different concentrations of glycerol were analyzed. Glycerol was intended to increase the viscosity of the dye solutions and consequently the anisotropy. TAMRA

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91 was made into 200 nM solutions with 0, 20, 40, 60, and 80% (V/V) glycerol. Then those solutions were dropped on a microscope glass slide. The images were obtained via a 20x objective at a PMT voltage of 650 V and laser power level of 25%. Figure 4-2 shows images of TAMRA with 20% (A) and 60% (B) glycerol. Note that for each glycerol concentration, the image on the left represents the I VH channel while the one on the right is from I VV channel. It is clear that from 20 % to 60%, the fluorescence intensity I VV turned from being lower than I VH to higher, indicating an increase in anisotropy of the TAMRA dye. The actual anisotropy was calculated from images of each glycerol concentration by selecting a rectangular area and taking the average intensity. The curve of anisotropy vs. glycerol concentration shown in Figure 4-3 has an obvious trend of increasing anisotropy. 0204060800.000.020.040.060.080.100.120.140.16 AnisotropyGlycerol concentration (%) Figure 4-3. Anisotropy of 200 nM TAMRA solutions with increasing concentration of glycerol.

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92 Effects of PMT Voltage on Anisotropy Data During the process of monitoring fluorescence from live cells, due to reasons like photo-beaching, the intensity may gradually decrease. When the signal drops to a very low level, there are a few ways to amplify signal, such as increase PMT voltage or gain, and increase laser power. As discussed in the Experimental Section, we mostly used PMT voltage to control the signal level due to its low impact on noise level and fluorophore stability. However, since we had two PMTs in two channels, even if the voltage was increased to the same level for both PMTs, they could still have different response to such voltage increase. This difference could result in a change in the calculated anisotropy. Therefore, it was desirable to study whether the same sample would have different anisotropy at different PMT voltage level. To do this, a 15mer ssDNA with a TAMRA label at the 3-end was made into 100nM solution. Then we put it on a glass slide on the microscope. While laser power was kept at 25% and PMT gain at 1, voltage on both PMTs was adjusted. Intensities as well as calculated anisotropies were recorded at each voltage and eventually plotted against voltage to get Figure 4-4. The I VH and I VV almost went up in parallel as PMT voltage was brought up. However, careful examination of the curves shows that the I VH began to max out over 850 V since the maximum readout from the PMT was 4095. While I VH was close to saturated, I VV kept going up, which resulted in a sudden jump in the anisotropy value after 850 V. Besides this jump, anisotropy remained rather consistent throughout the lower voltage range. The conclusion we could draw from this results was that the voltage of the PMTs probably would have much impact on the anisotropy measurements, especially at lower than 850 V. In the mean time, it was advisable to keep the signal at a high level yet away from saturation.

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93 050010001500200025003000350040004500675725775825875PMT voltage (V)Fluorescence intensity 0.000.020.040.060.080.100.120.140.16675725775825875925PMT voltage (V)Anisotropy 050010001500200025003000350040004500675725775825875PMT voltage (V)Fluorescence intensity 0.000.020.040.060.080.100.120.140.16675725775825875925PMT voltage (V)Anisotropy Figure 4-4. Effect of PMT voltage on anisotropy measurements. Top: measured fluorescence intensity vs. PMT voltage change, with for IVH and for IVV. Bottom: PMT voltage vs. anisotropies calculated using intensity data from the top graph.

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94 Monitoring ssDNA Digestion in Homogeneous Solutions Digestion of TAMRA-labeled ssDNA was first monitored on the FAI system in homogeneous solutions before moving to cellular samples. A 36mer ssDNA with the 6 repeat units of human telomeric sequence, 5'-(TTAGGG) 6 -3', was labeled with a TAMRA dye at the 5 end. 4.5 L of 2.5 M of this DNA (TeloH) was dropped on a thin glass slide placed above a 60x objective on the confocal. Then 0.5 L of 10 mg/mL DNase I was added to the drop. Images from the VV and VH channels were recorded every 5 minutes for 1 hour. Those images were calculated into 13 anisotropy images and the average anisotropy value from the same region of interest on each anisotropy image was taken and plotted against time. The result is shown in Figure 4-5. 01020304050600.050.060.070.080.090.100.11 AnisotropyTime (min.) Figure 4-5. Anisotropy change of 5-TAMRA-(TTAGGG) 6 -3 (TeloH) during digestion by DNase I in solution monitored by FAI. Monitoring started after 4.5 L of 2.5 M TeloH was mixed with 0.5 L of 10mg/mL DNase I.

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95 A clear trend of decreasing anisotropy is displayed, indicating digestion of DNA can be monitored using anisotropy. This anisotropy drop is similar to what we obtained from digestion of the same DNA by DNase I on a spectrofluorometer capable of anisotropy measurements. Control experiment on the confocal showed that without the DNase I, there was no change in the anisotropy of the dye-labeled DNA. Digestion of ssDNA in Live Cells A breast cancer cell line MDA-MB-231 (ATCC, Manassas, VA) was used to study DNA digestion in live cells. Cells in a glass-bottom dish were washed off culture media with a HEPES buffer (10 mM HEPES, 10 mM D-glucose, 150 mM NaCl, 5 mM KCl, 1 mM MgCl 2 2 mM CaCl 2 pH=7.3) for a few times. Then the dish was moved to a microincubator on the microscope stage at 37C controlled by a TC-202A temperature control (Harvard Apparatus, Inc., Holliston, MA). The cells were covered with HEPES buffer, then light mineral oil to prevent evaporation. Injection of DNA samples into single cells was done using electroporation according to a previous report. 124 Slightly modified conditions were used for our cells. A Grass S44 stimulator was employed to produce voltage pulses. Once the microinjection tip was brought in contact with the cell membrane, voltage pulses were applied. Those pulses were able to disrupt the cell membrane structure and form small pores for a short period of time. 124 At the same time, the negatively changes DNA molecules quickly diffused out because of the negative charges on the cathode and penetrated the cells through the small pores. Voltage was applied in the frequency of 3~5 pulses per second, with a duration time of 20~30 ms. Magnitude of voltage ranged from 5 V to 15 V. The amount of DNA injected into the cell using electroporation varied from time to time. By comparing the intensity of injected dye-labeled DNA in the cells to that of standard DNA solutions, the cellular

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96 concentration of the injected DNA was estimated to be around 100 nM to 1 M. Considering the cell volume was about 2-3 pL (~50 m length, ~10 m diameter), the concentration corresponds to 2.5~25 5 DNA molecules in one cell. 0.25 0.08 Figure 4-6. Digestion of Ctrl1 in live cells monitored by anisotropy change. Anisotropy images of a cell injected with Ctrl1 at 2, 7, 12, 17, 23 and 33 minute after the injection, with the scale bar of anisotropy on the bottom.

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97 Once the DNA was injected into the cell, emission from the TAMRA label was separated and recorded in the two polarization detection channels every few minutes. These images would later be converted to a series of anisotropy images. Anisotropies from different areas of the cell, such as cytoplasm and nucleus, were usually obtained by taking an average in the specific region in the anisotropy image. As a test of the cell based DNA digestion assay, a TAMRA-labeled DNA with a sequence of 5-CGT CGG CTA GCC GAG CTA GTA AAC CAC CAT GGT CCG ACG-TAMRA-3 (Ctrl1) was investigated in live cells. Anisotropy images were obtained as described above for each cell injected. Figure 4-6 shows a series of anisotropy images acquired from one cell at different time after the DNA injection. It can be seen that both the nucleus and cytoplasm areas of the cell displayed gradual decrease in anisotropy as the experiment proceeded. This could indicate certain level of nuclease activity in cytoplasm. One thing worth noting and typical of all the cells we observed is that the anisotropy in the cytoplasm region was always lower than that in the nucleus, with the cytoplasm value closer to anisotropy of the DNA in solution samples. It is possible that the high density of macromolecules in the nucleus makes it more viscous which in turn causes higher anisotropy. Another possible reason is that some basic proteins such as histone, may non-specifically bind to the dye-labeled DNA, increasing its molecular weight. Since we were interested in the relative anisotropy decrease of the DNA induced by nuclease digestion, we averaged the anisotropy values only in the nucleus region on each image and plotted it against time. As shown in Figure 3, the anisotropy dropped to the lowest level after about 40 min. For most cells that had been monitored, anisotropy would stop going down within 2 h. The absolute anisotropy

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98 decrease (~0.03) in Figure 4-7 was less than what had been observed for DNA digestion in homogeneous solutions (~0.06-0.09). The possible reason is that once the dye-labeled DNA molecules were cleaved down to very small fragments, they became much easier to escape via the nucleus membrane and eventually the cell membrane into the extracellular environment. As a result, smaller DNA fragments contributed less to the anisotropy obtained from the images than the bigger fragments. This was supported by two experimental results. First, pure TAMRA dye injected into the cells was seen to completely diffuse out within a minute. Second, during the monitoring of the digestion of Ctrl1 inside live cells, both detection channels showed decreasing fluorescence intensity. Since photo-bleaching was less a problem because laser was shone on the sample for just a few seconds every few minutes, it was most likely some dye-linked DNA fragments diffused out of the cell. 01020304050600.1250.1300.1350.1400.1450.1500.1550.1600.1650.1700.1750.180 AnisotropyTime (min.) Figure 4-7. Anisotropy values obtained by averaging in the nucleus region of the anisotropy images of two cells injected with Ctrl1 and Ctrl1-S respectively. Then they are plotted against time. () for Ctrl1 and () for Ctrl1-S.

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99 A control DNA (Ctrl1-S) was synthesized with the same sequence as Ctrl1 using nuclease-resistant phosphorothioate modified DNA bases. In these bases, one of the two nonbridging oxygen atoms in the phophodiester bond is replaced by sulfur. Cells injected with Ctrl1-S showed barely any anisotropy decrease within the 1-2 h monitoring time period. Quantitative result for one cell is shown in Figure 4-7. The anisotropy remained constant throughout the experiment. The slightly higher anisotropy might be due to the fact that phosphorothioate-backboned DNAs are more subject to non-specific binding with proteins than regular DNAs are. 125 Reduction of fluorescence intensity in the detection channels was still observed, but to a less extent, indicating a slower diffusion of the non-cleaved Ctrl1-S. The results from the control DNA shows that it is practical to use localized anisotropy change to monitor DNA digestion by nucleases in live cells. A few other oligonucleotides and their phosphorothioate-modified counterparts were also tested in live cells. Results from those molecules confirmed that regular ssDNA could be easily degraded while the modified oligonucleotides hardly showed any anisotropy change. It was always observed that the modified oligonucleotides had higher initial anisotropies than the regular ones. Stability of Telomere-Like Oligonucleotides in the Nuclei of Live Cells In order to study stability of single-stranded telomeres in the cells, a few TAMRA-labeled oligonucleotides with 6 telomeric repeat units were synthesized. One of them with the human telomeric sequence, 5-TAMRA-(TTAGGG) 6 -3, was named TeloH, while aother with Tetrahymena sequence, 5-TAMRA-(TTGGGG) 6 -3, was termed TeloT. Both sequences were long enough to form intramolecular G-quardruplex in Na + or K + containing solutions even though the detailed conformation may differ with sequence. 126-128 The reason the dye was labeled at the 5 end was because the single

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100 stranded telomere tip in cells has an open 3 end. Another DNA was synthesized with a slightly different sequence than TeloH--5-TAMRA-A(GGGTTA) 6 -3. This DNA (TeloH2) was used to see if position of the dye label in the telomeric sequence would have any impact on the DNA digestion. A control DNA sequence (TeloCtrl), which replaced all guanine bases in TeloH with cytosine bases, was used to compare with all the G-quardruplex forming oligonucleotides. All the above ssDNAs were injected into single cells separately and monitored for anisotropy changes using procedures described previously. For each DNA sample, over 10 cells were tested. Anisotropy images were obtained and the anisotropy in the nucleus region was averaged and plotted against time. Our results demonstrated that almost all the cells injected with TeloH, TeloT or TeloH2 showed quite stable anisotropy inside the nuclei for up to 2 hours, while most cells with TeloCtrl gave degraded anisotropy at different rate, just like what happened to Ctrl1. The rate of DNA digestion was probably related to the stage of the cell in the cell cycle since nuclease activity might vary at different stage of cell cycle. The amount of DNA injected should have also contributed to the varying rate of anisotropy change of TeloCtrl. Another observation was that for TeloH, TeloT and TeloH2, there was less drop in the fluorescence intensity during the experiments, while significant fluorescence decrease sometimes happened during monitoring with TeloCtrl. This might be explained by reasons described before which account the diffusion of cleaved DNA fragments for the intensity drop. Figure 4-8A shows the anisotropy change from four typical cells injected with the 4 ssDNAs respectively. It can be seen that for the three G-quardruplex

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101 forming oligonucleotides, there was slight anisotropy increase within experimental error, instead of any decrease. For TeloCtrl, there was clear decrease. 0204060800.100.110.120.130.140.150.160.17 AnisotropyTime (min.) -0.010.000.010.020.030.040.05TeloHTeloH2TeloTTeloCtrlAnisotropy decrease0204060800.100.110.120.130.140.150.160.17 AnisotropyTime (min.) -0.010.000.010.020.030.040.05TeloHTeloH2TeloTTeloCtrlAnisotropy decrease Figure 4-8. Digestion of TeloH, TeloT, TeloH2 and TeloCtrl in live cells monitored by anisotropy change. (A) Anisotropy values obtained by averaging in the nucleus region of the anisotropy images of 4 cells injected with TeloH (), TeloT (), TeloH2 ( ) and TeloCtrl () respectively. (B) Anisotropy decrease between the start and end of the monitoring of the cell. The value is the average from 4 cells for each of TeloH, TeloT, TeloH2 and TeloCtrl.

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102 To compare the results based on multiple cells, we picked anisotropy data from 4 relative healthy cells for each of the 4 DNA sequences, and calculated the anisotropy decrease between the time of injection and the end of the monitoring. Then we took the average of the anisotropy decreases for each DNA and compared the results from the four in Figure 4-8B. It is clear that telomere-like sequences were more resistant to nuclease activity inside the cell nucleus than sequences that could not form G-quardruplex. It is also noteworthy that the stability of the telomere-like oligonucleotides is not heavily dependent on the detailed sequence since the Tetrahymena telomere sequence was as stable as human telomere sequence in the human breast cancer cells. The formation of the G-quardruplex is probably a more determining factor. Protein binding of the telomeric DNA may be able to protect the DNA from being degraded as reported previously. However, in our experiments, the telomere-like oligonucleotides injected into the cells were at the level of 10 5 copies and they could be quite stable. Assuming protein-DNA binding ratio of only 1:1, then 10 5 copies of telomere-binding proteins would be needed to protect those DNA molecules, which is quite unlikely in a single nucleus. The G-quardruplex structure, on the other hand, may have an inherent advantage over regular DNA structure in terms of resistance to nucleases in cell nuclei. Conclusions In this work, we have developed a fluorescence anisotropy imaging system based on a confocal microscope for monitoring DNA digestion inside live cells by localized anisotropy detection. We used this setup to study the nuclease-resistant capability of telomere-like ssDNAs in nuclei of human breast cancer cells and found that those oligonucleotides were clearly more stable than regular DNA sequences during our

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103 experiments. We conclude that the G-quardruplex structure of the telomere-like ssDNA may make it inherently more stable in intracellular environments than non-G-quardruplex structures. This may help understand why the G-quardruplex forming telomere sequences were adopted by eukaryotic cells in the very beginning to protect the end of chromosome. Other reports showed the stability of oligonucleotides with G-quardruplex structure in bovine serums using radio labeling and gel electrophoresis. 129 In contrast, our method based on fluorescence anisotropy provides a way to directly monitor DNA digestion in any region of live cells in real time. One application of our assay may be evaluation of nucleic acid based molecules for therapeutic purposes. While antisense agents can be important drug candidates for gene-related diseases, 130 aptamers can target disease-related proteins. Using our setup, one would be able to easily assess whether those agents can be stable enough against intracellular degradation for real applications.

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CHAPTER 5 SUMMARY AND FUTURE DIRECTIONS Summary of Oligonucleotide-Based Protein Interaction Study We have explored the applications of oligonucleotide-based protein ligands in protein research under different conditions. When used in homogeneous solutions, protein-binding aptamers can report protein-protein interactions in real time. The most important strong point of this assay is that neither of the proteins needs to be modified in any way. Thus it is believed that more reliable protein interaction information can be provided than the techniques that do require protein modification. Second, compared to direct protein-protein study, the indirect competitive assay can offer knowledge about the binding site involved in the interaction. On the other side, even when the prey protein does not compete with the aptamer for the bait protein, the fluorescence anisotropy based signal transduction mechanism still allows for the detection of co-binding of the aptamer and the prey protein on the bait protein. This distinctive capability of our assay is not achievable via many other techniques. Lastly, the ability of our method for real-time protein interaction monitoring offers great potential for high-throughput functional protein research. Real-time capability means the interaction is reported as it is happening. No extra steps are needed, which not only simplifies the sample handling and processing, but also greatly reduces the time needed for the tests. The aptamer-based assay is perfect for high-throughput protein function study in standard 96or 384-well plates on readily available instruments. The significance of this new approach may ultimately lie in its applicability in drug discovery. Biologically important protein systems can be selected as 104

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105 the target for such drug development, for example, the p53-MDM2 interaction discussed in Chapter 1. Because MDM2 binds to p53 to inhibit its function of arresting damaged cells, disruption of this interaction may lead to the recovery of p53s activity of controlling tumor proliferation. To find drug candidates out of thousands of organic molecules, a fast assay can be proposed based on the concept of oligonucleotide-based probes. First, an aptamer needs to be isolated for p53 that can compete with MDM2 for the same p53 binding site. The aptamer can then be modified to incorporate a signal transduction mechanism such as FRET or anisotropy. On a standard multi-well plate, each well will contain the aptamer probe, p53, and MDM2. Organic molecules are added to each well separately. If the organic molecule is able to disturb the p53-MDM2 binding, preferentially by interacting with MDM2, the MDM2 binding site on p53 will be left open. As more aptamer probes can bind to p53, a change in fluorescence signal will be detectable. In this way, molecules that can hinder MDM2s capability of inhibiting p53 can be rapidly identified as drug candidates and collected for further evaluation. This simple example clearly shows that, compared to most other techniques, a great deal of time can be saved in the early stage of drug discovery. Many more protein systems can be targeted in a similar way for various diseases. The use of oligonucleotide probes seems to be a better choice over antibodies in capillary electrophoresis (CE), considering many advantages of aptamers. Aptamers can be easily and uniformly labeled at designated internal locations with fluorophores for sensitive detection. In addition, the negative backbone of aptamer is beneficial for separation of aptamer-protein complex and aptamer itself. One problem with antibodies in CE is that the large antibodies, like most proteins, tend to be non-specifically adsorbed

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106 on the fused-silica inner wall of the capillary. As a result, not only the accuracy of the analyses becomes unreliable, the electrophoresis is also deteriorated due to the blocking of the inner surface of the capillary. Coating of the inside of the capillary with polymers can alleviate this problem in short term, but not in the long run. Replacing antibodies with aptamers apparently helps lighten the adsorption problem. The combination of oligonucleotide probes and CE can be used for quick discovery or confirmation of protein interactions. Being able to visualize the formation or deformation of the complex peak can serve as direct and reliable evidence for the presence of DNA-protein or protein-protein interactions. The fluorescence anisotropy imaging technique we developed and used in this work is a relatively new method for studying intracellular bio-interactions. Oligonucleotides can be easily labeled with fluorophores for intended protein interactions. Currently, anisotropy measurements are more accurate by averaging the anisotropy on a region of the image, than just taking data from single pixels. This is because fluorescence anisotropy has limited dynamic range (0-0.4). Neglectable variations in fluorescence intensity from pixel to pixel may result in significant anisotropy differences between pixels, especially considering that intensity data from two images need to be used for anisotropy calculation. However, average of multiple pixels is still a reliable indication of the anisotropy in a confined region. Our study presented here did not rely on single pixel measurements, so it was still practical to use anisotropy imaging. The FAI approach is a unique cellular imaging technique in the sense that anisotropy is mostly dependent on the size of the fluorescent molecules. Thus molecular conformation is of less concern. The other advantage is that anisotropy is related to the ratio of two components of the

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107 fluorescence emission. Therefore, fluctuation in the emission, either due to unstable light source or bleaching of the fluorophore, should pose minimum effect on the average anisotropy. In conclusion, fluorescence anisotropy imaging has its unique features as a tool for cellular study. Careful selection of both the probe-target system and the experimental design may yield interesting results concerning bio-interactions in or out of live cells. Future Directions Even though our work has been focused on the applications of oligonucleotides for detection and monitoring of protein interactions under different conditions using various approaches, future directions could be shifted to selection of new aptamers for important proteins and development of aptamers as therapeutically active reagents. Each cycle of aptamer selection usually contains incubation of nucleic acid library with target, separation of bound complex from free nucleic acids, and dissociation of complex and PCR amplification of the selected sequences. While the first and third steps are often kept unchanged in different laboratories, variants of the separation mechanism have been proposed. One of the latest developments is the CE-based aptamer selection. 131;132 Separation of nucleic acid sequences and the protein-binding complex in traditional aptamer SELEX is usually carried out using either nitrocellulose filtration or affinity chromatography. In nitrocellulose filtration, the DNA or RNA library incubated with protein target is filtered through nitrocellulose film. Since nitrocellulose is only sticky to protein molecules, the binding complex is retained on the film while the free nucleic acids are removed. Later, denaturing reagent will be used to dissociate the bound nucleic acid sequences from the complex for the next step. In affinity chromatography, the target protein is immobilized on insoluble resin. By running the library through the

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108 column, those sequences that have affinity for the target will migrate more slowly than the non-binding sequences. The fraction with protein-binding affinity will be collected and used for the PCR amplification step. Both methods involve separation on a solid support, which may, first, affect the interaction between the protein and nucleic acids, second, cause nonspecific adsorption of non-binding sequences. The former will limit the number of selected aptamers, while the latter introduces false positives and causes low selection efficiency. To solve these problems, CE-SELEX was developed. The basic idea is to separate the free nucleic acids and the protein complex in CE. Since the separation is carried out completely in solution, the above mentioned problems facing conventional SELEX are circumvented. The excellent separation efficiency of CE makes it possible to collect DNA or RNA fractions with highest protein affinity, resulting in less selection rounds. Features of CE such as short separation time and very low sample consumption make CE-SELEX even more appealing for aptamer development. With our current equipment and experience in aptamer and CE, CE-SELEX would be a very attractive way to isolate aptamer for proteins of interest. Another interesting development in SELEX is the use of magnetic microbead-aided separation. 133-135 In this method, the target protein is immobilized on the magnetic microbeads and incubated with the nucleic acid library. Unbound nucleic acids can be easily removed by washing the beads for a few times. The microbeads linked with the target and bound oligonucleotides can be manipulated and collected using magnets. The advantage of this technique is the simplified separation process. There is no need to run through a column or filter, and the beads with bound nucleic acids can be used directly for PCR amplification. This new method has helped us generate a new idea of high

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109 throughput aptamer selection for proteomic research. Proteomics target at analyses of expressions and functions of thousands of proteins in the whole cell. One of the major approaches to conduct proteomic research is by using protein microarray. 136 Similar to DNA microarrays, protein microarrays need to use immobilized capturing agents to bind and detect protein targets. However, current number of protein-binding ligands is very limited in terms of whole-cell protein study. One promising way of generating the needed ligands would be the selection of aptamers. Nonetheless, currently, the SELEX process is mostly dealing with one protein at a time, which would require tremendous amount of time for thousands of proteins. Our proposal for solving this problem would be to immobilize multiple proteins at one time on a solid support such as a glass slide. Then incubate this slide with the DNA or RNA library. Unbound nucleic acids are first washed off. And the bound sequences are consequently separated from the surface proteins and amplified by PCR. This process is repeated a few rounds until reasonable affinities of the isolated sequences are obtained for those target proteins. After the last incubation and washing, bound sequences would be obtained from individual protein spot, and be identified using DNA sequencing for each protein. One issue that needs to be addressed is the non-specific binding of nucleic acids on the glass slide. But this problem can be alleviated by a variety of surface coating chemistries. This proposed idea not only allows for high-throughput aptamer selection, it also offers a very simple separation strategy which only requires a few times washing. Future work on aptamer development could be carried out in this direction. Besides the use of aptamers for detection purposes and the development of aptamers, another important aspect of the aptamer research is the application of aptamers

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110 for therapeutic purposes. Aptamers were originally intended to be inhibitors of important biological functions of the target proteins. Their inherent low toxicity makes them ideal drug candidates targeting disease-related proteins. Even though antibody can also be used as protein inhibitor, their large sizes pose two problems for in vivo applications: slow tissue penetration and long blood residence, 25 which prevent the antibodies from getting to the target efficiently. Aptamers, on the other hand, do not have such concerns. One problem that needs to be considered for the therapeutic applications of aptamers is the stability of the aptamers in the nuclease-present environments. Many strategies have been developed to increase nuclease resistance of the oligonucleotides, some of which involve chemical modifications to the phosphate backbone or sugar moiety. For example, one report exploited phosphorothioate-modified aptamers for targeting at an immune receptor on T cells in vitro and in vivo. 137 Another group reported that by substituting 2 position of the sugar ring with O-methylor fluorogroup on the bases of an PDGF aptamer, and replacing the non-binding portions of the aptamer sequence with hexaethylene glycol spacers, a much longer half-life in serum was achieved while the binding affinity of the aptamer for PDGF was not altered much. 138 External labels were also evaluated for increased nuclease-resistance of aptamers. 139 It was found that aptamers linked to a biotin label at the 3-end showed increased stability in vitro, while a biotin-streptavidin group at the 3-end protected the aptamer from nucleases even in vivo. Besides the efforts in post-SELEX modifications for enhanced aptamer stability, some study has turned to directly using modified DNA or RNA bases in the SELEX process. 140 Aptamers isolated in this way would have built-in ability to resist nuclease digestion.

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111 By combining various aptamer modification strategies with our imaging capability, aptamer-target interaction can be monitored in live cells to study not only the localization of the target in cells, but also the impact of aptamer-binding on cell growth and proliferation. This would allow us to obtain the first-hand information regarding potential of aptamers for therapeutic applications.

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APPENDIX A PROGRAMMED DNA CONFORMATION CHANGE AND APTAMER ACTIVATION Introduction The potential of aptamers as therapeutic reagents has been outlined in the Chapter 5. One particular consideration that has been discussed in detail is the stability of the aptamers in biological fluid. However, there are many other aspects of the therapeutic applications of aptamers that need to be well thought out, just like with any traditional medicines. One of the more advanced issues concerning drug effectiveness is whether the drug candidate can be delivered to the diseased cells and bind to the targeted molecules. Drug delivery has been a field gaining great interest from both scientists and pharmaceutical industry. The ultimate goal of drug delivery is to delivery the drug safely to the specific target at the right time and at the right level. 141 However, site-specific drug delivery is still very difficult. Traditional delivery methods, including oral, nasal, and injection, may cause delivery of some of the drugs to healthy organs. As a result, serious side effects could happen while the targeted organ can not get sufficient drug molecules. One direct way to improve specificity of drug delivery is to use drug molecules highly selective for the targets in human body, and at the same time make sure the target is specific for the disease to be treated. This approach may require much more efforts in drug developments and study of disease mechanisms. To bypass those requirements, one alternative method could be controlling where the drug is delivered by using external 112

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113 forces or resources. The realization of this step would ensure that the drug molecules, such as aptamers, could be released or activated whenever and wherever they are needed. Nanomotors and Photo-Regulation of DNA Hybridization Our thinking of developing controlled aptamer release and activation was originated from previous research done in this laboratory on controlled DNA motion. 142 An oligonucleotide with a special sequence could take a G-quadruplex structure in the presence of metal ions. Part of the sequence of another oligonucleotide ( strand) was complementary to the G-strand. The addition of would induce the hybridization of these two and cause the G-strand to stretch out. Then, a third DNA ( strand), which was complementary to the whole strand sequence, was added to the same solution. Since could pair with more bases of strand than G-strand, it was able to hybridize with and release G-strand. As a result, the freed G-strand would come back to the folded G-quadruplex structure. During the above process, the stretch-shrink motion of the G-strand was driven by adding DNA fuel, namely and strands. By labeling the G-strand with a fluorophore and quencher, the DNA motion could be monitored via fluorescence change. A series of adding and in turn revealed cycling of fluorescence change as expected. The continuous change between the stretch and shrink states of the G-strand might provide a certain amount of work at single molecular level, which was why the G-strand was given the name nanomotor. The programmed movements of the nanomotor may be very useful for the manipulation of nano-scale elements. The above nanomotor work demonstrated the concept of using external resources to control the structure of DNA molecules. One problem with this model is that the need for DNA fuel may make it inconvenient to run the motor in some situations. Furthermore, the

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114 accumulation of the DNA fuel might disturb the nanomotor system and would eventually put a stop to the nanomotor. Compared to chemical fuel, external physical forces, such as electromagnetic fields, may be more suitable for powering molecular motors. Promise for such a nanomotor system has emerged from recent developments related to an organic molecule known as azobenzene. Azobenzene contains two benzene rings connected by a N=Nbond (Figure A-1). The unique feature of azobenzene is that its conformation is sensitive to lights of different wavelengths. Light of ~350nm stabilizes the cisform of the N=Nbond, while lights of >400nm would change the molecule to the tranform (Figure A-1). 300<<400nm>400nmtrans-azobenzenecis-azobenzene 300<<400nm>400nmtrans-azobenzenecis-azobenzene Figure A-1. Structure of azobenzene is changed by lights of different wavelengths. An interesting application of azobenzene was discovered when it was incorporated into a DNA strand. 143 The azobenzene was covalently linked to the oligonucleotide side chain, replacing a sugar moiety and the base, to form a synthetic DNA base. When this artificial base was in a double-stranded DNA (dsDNA), it was found that the isomerization of the azobenzene affected the stability of the hybridization. Under 350 nm UV light, the cisform of the azobenzene base rendered the hybridization more unstable than under the light of >400 nm. The stability difference of the dsDNA at the two azobenzene states was indicated by differences in melting temperature (Tm) of the

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115 dsDNA. With optimized synthesis and the use of multiple azobenzene bases, Tm of a dsDNA was as high as 20 C when irradiated with different lights. 144 This result implies that under fine tuned conditions, it is possible to use light to control the hybridization and de-hybridization of two complementary DNA stands at room temperature. As a consequence, the previously described DNA nanomotor could be redesigned to use light as the energy source. In the new design, instead of having two DNA strands ( and ) to change conformation of the G-strand, an oligonucleotide complementary to the nanomotor could be modified with multiple azobenzene bases. Under the irradiation of the light with a wavelength longer than 400 nm, the azobenzene strand would be able to hybridize with the G-strand and stretch the nanomotor. Changing the light to 350 nm would destabilize the dsDNA so that the G-strand can fold back to form stable G-quadruplex structure. Similar to the original nanomotor report, cycling the two different lights would make the G-strand change continuously between stretch and shrink states, providing work to the surrounding environments. Besides working as a nanomotor, the G-strand could be labeled with two molecules at the two ends so the stretch and shrink movements could bring the two molecules away and close as a way of switching on and off certain reactions. The advantages of using an external physical force are obvious, including noninvasive operations, remote-control capability, and reproducibility. Another design of photo-regulated nanomotor makes use of the hairpin structure of molecular beacons (MBs). The azobenzene base could be placed in the stem region of the beacon so that changing light would render the beacon to open and close as another form

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116 of nanomotor. This uni-molecular approach may have some advantages over the bimolecular nanomotor. Both strategies are under investigation and development. Photo-Regulated Aptamer Release and Activation The concept of photo-regulated DNA conformation can be well suited to control aptamers function and activity. As discussed in previous chapters, aptamers conformation is very important for their target-binding capability. Usually only one particular structure of the aptamer has the affinity for the target, e.g., the G-quadruplex structure of thrombin aptamers. With the ability of the synthetic azobenzene base to affect the stability of DNA double strands, it is possible to integrate those bases into the aptamer sequence and consequently control whether the aptamer can take the necessary conformation for target binding. One simple design would be similar to the bi-molecular nanomotor strategy, in which the complementary DNA (cDNA) of the aptamer would be modified with one or more azobenzene bases. Controlling the hybridization of the cDNA to the aptamer would indirectly decide whether the aptamer can bind to the target. Another uni-molecular approach requires the addition of a few bases at one end of the aptamer. Those bases would be complementary to bases at the other end of the aptamer so that the new oligonucleotide could form a hairpin structure. One example of implementing this design is for the thrombin aptamer, shown in Figure A-2.

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117 300<<400 nm>400 nm Inhibition of thrombin 300<<400 nm>400 nm Inhibition of thrombin Figure A-2. Schematics of photo-regulated activity of thrombin aptamer. The blue line represents the added bases that are complementary to part of the aptamer sequence (orange line). The solid dark green dot in the stem of the oligonucleotide on the far left is the synthetic azobenzene base. In the pro-hybridization form of the azobenzene base, the modified thrombin aptamer would take a stem-loop structure which prevents the formation of the G-quadruplex that is necessary for thrombin binding. Once the azobenzene base is switched to the con-hybridization state by proper light irradiation, the stem would become unstable, thus a G-quadruplex is favored. The uni-molecular approach for controlled aptamer activity is appealing because of its simplicity and no need to optimize the cDNA to aptamer concentration ratio for better controllability. Initial tests were done to test if the uni-molecule idea would work. Two oligonucleotides both containing the thrombin aptamer sequence were used to mimic the effect of the azobenzene base. One of the oligonucleotides could form a complementary stem (Ap-CS, 5-GC GGTTGGTGTGGTTGG ACCGC-3), while the other had the same sequence except that the middle of the stem had a mismatched base pair (Ap-MS, 5-GC GGTTGGTGTGGTTGG ACAGC-3). In the sequences shown here, the underlined portions represent the actual aptamer sequence and the bold bases form the stems. The goal of the tests was to see whether difference in the stem stability between Ap-CS and Ap-MS had any effect on their ability to inhibit thrombin. A commonly used test for

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118 thrombin activity is based on its ability to cleave a protein called fibrinogen to produce fragments known as fibrin. The crosslinking between fibrin molecules can then develop into a polymer network, often resulting in a white insoluble. The 15mer aptamer that was first isolated for thrombin was found to inhibit thrombins protease activity and slow down the formation of fibrin precipitate. Direct observation of a white insoluble is often used as a way to assess thrombins activity. Here a different method was applied which involved measuring scattering light from the sample. As a precipitate was forming in a clear solution, the solid would begin to scatter incident light into all directions. Measuring the intensity of the scattered light would be a good indication of the formation of the polymer network in real time. Fibrinogen was obtained from Sigma-Aldrich, Inc. (St. Louis, MO), and human -thrombin was from Haematologic Technologies Inc. (Essex Junction, VT). All DNAs were synthesized and purified by GenoMechanix, LLC (Gainesville, FL). Other chemicals were from Sigma-Aldrich (St. Louis, MO). Fibrinogen was made into 800 nM solutions in a physiological buffer (20 mM Tris-HCl buffer at pH 7.4, 150 mM NaCl, 5 mM KCl, 1 mM MgCl 2 1 mM CaCl 2 and 5% V/V glycerol). Ap-CS or Ap-MS was added to the solution to make a 4 M final concentration. The aptamer was first incubated with the substrate of thrombin instead of thrombin because both our experimental results and a previous report showed that the order of incubation did not affect much on thrombin inhibition. 145 This mixture solution was then placed in a 100 L quartz cuvette which sat in a Fluorolog-3 spectrofluorometer (Jobin Yvon Inc., Edison, NJ). The excitation and emission wavelengths on Fluorolog-3 were both set to 650 nm. Slit widths were 1 nm. The aptamer-fibrinogen incubation

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119 solution was monitored for scattering light continuously. After a few minutes, concentrated thrombin solution was added to make a 40 nM final concentration. Monitoring continued for another 15 to 20 minutes. The results of scattering intensity change for both Ap-CS and Ap-MS are shown in Figure A-3. 020040060080010001200020000400006000080000100000120000 IntensityTime (second) Figure A-3. Inhibition of thrombin by Ap-CS and Ap-MS monitored by measuring scattering light. for Ap-CS and for Ap-MS. Thrombin was added at near 200 second. It is clearly shown that Ap-MS, with the unstable stem, was able to slow down the formation of the fibrin insoluble more than Ap-CS. This indicates that by changing the conformation of the modified aptamer, a difference in thrombin inhibition capability can indeed be obtained. Current efforts are focused on optimizing synthesis of azobenzene

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120 containing oligonucleotides. Once that is completed successfully, similar results of thrombin inhibition are expected when the aptamer conformation is controlled by lights of different wavelengths. Our design of photo-regulated activation of aptamers represents a novel approach to achieving controlled specific drug delivery. Multiple strategies can be adopted to turn this design into real application of aptamer delivery in human body. One way might be hybridizing aptamers with their azobenzene-containing cDNAs on the tip of a fiber, and then delivering the fiber to the targeted organ and applying proper light to release the aptamers. Another way for controlled aptamer delivery could take advantage of a two-photon laser system. In such a system, the highly focused high power laser of longer wavelength causes the molecule to absorb two photons almost simultaneously so that the energy absorbed equals to that from a single photon of shorter wavelength. 146 For example, a 700 nm two-photon laser will act like a 350 nm light source when used as an excitation light. Because lights of long wavelengths (infrared or near infrared) can penetrate biological tissues much better than short wavelengths with much less damage, excitation and imaging of fluorescent molecules even in live animals is possible. 147 With the capability of two-photon excitation, we would be able to directly irradiate targeted organs in a patients body from outside. The modified aptamers that contain azobenzene bases and were previously delivered to the patients body will get released only in the irradiated region. In summary, aptamer-based drugs can potentially be delivered to targets in a specific and controlled manner. Integration of azobenzene into aptamer drug design may allow release and activation of aptamers whenever and wherever they are needed via

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121 external physical forces. The benefit gained from the controlled drug delivery by using our designs might be valuable for effective treatment of diseases such as various cancers. Immediate research plan at current state should be optimization of both the synthesis of azobenzene oligonucleotides and the design of aptamers. Thrombin and its aptamer would then be tested as a model system in vitro for controlled thrombin activity.

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APPENDIX B ENHANCED PROTEIN BINDING AND INHIBITION BY DUAL-APTAMERS Introduction to Dual-Aptamers From previous discussion on aptamers, one can see that even though the SELEX process is capable of producing aptamers with very high affinity for the targets, there is no guarantee that every aptamer has expected binding strength. One example of this situation is the 15mer thrombin aptamer. With its dissociate constant in the 100 nM range, this aptamer could hardly maintain a stable complex with thrombin in CE (Chapter 3). Aptamers with higher affinity for thrombin were later isolated. 53 However, these aptamers bind to Exosite II of thrombin and do not inhibit the blood clotting activity of thrombin. 53 The 15mer aptamer, being the only aptamer that can inhibit thrombins biological function, does a poor job in that matter, mostly due to its weak binding to thrombin. In order to increase the affinity of the 15mer aptamer, post-SELEX modification should be considered. However, while adding, removing, and changing bases can easily reduce an aptamers affinity, it is difficult to predict what modifications on which bases can enhance the affinity, unless every modification is tested on each base. To solve this problem, we propose to use spacers to link two of thrombins aptamers (15Ap and 27Ap in Chapter 2), to form a dual-aptamer for enhanced thrombin binding and inhibition. A short poly(ethylene) glycol (PEG) chain with 18 atoms on the backbone was chosen to be the spacer. This PEG chain is idea for a DNA linker because it is linear, hydrophilic and relatively inert. It is also commercially available as a phosphoramidite 122

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123 for easy synthesis on a DNA synthesizer. The length of the spacer is estimated to be about 2.2 nm. The principle of using linked aptamers to achieve greater thrombin inhibition is illustrated in Figure B-1. SubstrateThrombin 15Ap27Ap SubstrateThrombin 15Ap27Ap Figure B-1. Illustration of inhibition of thrombin by a single aptamer (top) and a dual-aptamer (bottom). The aptamer-thrombin reaction can be described with the following equation. Apt-Thr k ON k OFF Apt + Thr The dissociate constant K d is related to the two reaction rates, k ON and k OFF K d = k OFF / k ON When a single 15Ap is used to inhibit thrombin, the aptamer needs to compete with the substrate for thrombin. However, once the 15Ap comes off the thrombin and diffuses into the solution, it does not have much advantage over the substrate molecules in the

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124 same solution in terms of coming back to the binding site. This situation is changed, however, when the 15Ap is linked to the 27Ap. Because both aptamers have affinity for the thrombin, there is much less chance for both aptamers to fall off the thrombin at the same time compared to one aptamer coming off. As an example, consider only 15Ap has left the binding site. Since 27Ap is still on the thrombin, the 15Ap moiety is still very close to the thrombin. Compared to the substrates that are far away in the solution, the 15Ap has a much better chance to come back and bind to the Exosite I again. In terms of kinetics, it is clear that even though the k OFF of 15Ap will probably not change because it is related to the type and strength of the non-covalent forces between 15Ap and thrombin, the k ON of the linked 15Ap could be greatly enhanced because of the close proximity, or, from another angle, the high local concentration of 15Ap around the thrombin. The net result of this change is a lower dissociate constant, representing an improved binding affinity and better inhibition. Design of Dual-Aptamers for Thrombin and Experimental Results To test this idea, experiments were carried out to see if it was possible for both aptamers to bind to thrombin at the same time. A FRET assay similar to the one in Chapter 2 was conducted. 15Ap labeled with a fluorescein and dabcyl quencher was incubated with thrombin, and then excess 27Ap was added to see if there was any fluorescence change. In another experiment, excess 15Ap was added to thrombin/dual-labeled 27Ap mixture. Both tests showed no fluorescence increase, meaning no competition took place between the two aptamers. Linked aptamers were synthesized according to this sequence, 5'-GGT TGG TGT GGT TGG T-(S) n -T ACC CGT GGT AGG GTA GGA TGG GGT GGT-3'. S stands for spacer. Oligonucleotide bases on the 5 end formed 15Ap sequence while the 3 end

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125 had the 27Ap sequence. A few dual-aptamers were tested with different numbers of spacers, including 4, 6, 8, 10, and 12, which represented distances ranging from ~8 nm to ~25 nm. Other oligonucleotides tried included 15Ap and 27Ap. Inhibition of thrombin was evaluated based on the fibrinogen test discussed in Appendix A. Fibrinogen was from Sigma-Aldrich, Inc. (St. Louis, MO), and human -thrombin was from Haematologic Technologies Inc. (Essex Junction, VT). All DNAs were synthesized and purified by GenoMechanix, LLC (Gainesville, FL). Other chemicals were from Sigma-Aldrich (St. Louis, MO). 200 L physiological buffer (20 mM Tris-HCl buffer at pH 7.4, 150 mM NaCl, 5 mM KCl, 1 mM MgCl 2 1 mM CaCl 2 and 5% V/V glycerol) was added to a disposable transparent plastic cuvette. Then 0.5 L of 100 M oligonucleotide and 0.5 L of 10 M thrombin were added and incubated for 15 minutes. After that, 4 L of 20 mg/mL fibrinogen was mixed with the solution. Sample in the cuvette was examined continuously for the formation of a gel-like substance. The time when the sample was not fluidic any more was recorded. Multiple oligonucleotides were tested in parallel in several cuvettes. Typical results from one of several such tests are shown in Table B-1. Table B-1. Inhibition of thrombin-fibrinogen reaction by various ologinucleotides. DA-nS represent dual-aptamer. The number before S is the number of spacers in the linker. Oligonucleotide(s) in test solution Time till formation of fibrin gel None < 5 min 15Ap ~13 min 25Ap < 5 min 15Ap + 27Ap ~20 min

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126 DA-4S < 5 min DA-6S < 5 min DA-8S ~70 min DA-10S ~30 min DA-12S ~38 min A few observations of Table B-1 are interesting. First, 27Ap alone did not show any thrombin inhibition. The mixture of 15Ap and 27Ap did not improve the inhibition much compared to with 15Ap alone. The two dual-aptamers, DA-4S and DA-6S, actually displayed declined activity. The explanation would be that the lengths of 4 and 6 spacers were not long enough for two aptamers to bind comfortably to the two opposite sites of thrombin. As a result, some dual-aptamers would bind to Excsite I and others to Exosite II, thus the reduced inhibition. In contrast, eight spacers seemed to be ideal for enhanced thrombin binding. DA-8S demonstrated an impressive prolonged time for gel formation. Since 27Ap did not help with thrombin inhibition, the improved aptamer functionality could be well explained with the model we proposed shortly ago. Something not that clear is that DA-10S and DA-12S had worse performance than DA-8S, though still better than 15Ap. This may indicate that an optimized length is needed to link two aptamers for best binding affinity. Experiments similar to those in Appendix A were done to measure scattering light during the oligo-thrombin-fibrinogen incubation. Only 15Ap and DA-8S were tested in this case. Concentrations and other conditions were the same as in the gel-observation tests above except that everything was mixed in a 100 L quartz cuvette on a Fluorolog-3

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127 spectrofluorometer (Jobin Yvon Inc., Edison, NJ). The excitation and emission wavelengths on Fluorolog-3 were both set to 650 nm. Excitation and emission slit widths were 1 nm. The results of scattering light measurements are shown in Figure B-2. 0100020003000400050001000020000300004000050000600007000080000 IntensityTime (second) Figure B-2. Inhibition of thrombin by 15Ap and DA-8S monitored by measuring scattering light. for 15Ap and for DA-8S. Thrombin was added at near 500 second. Figure B-2 shows a much slower increase of scattering intensity for DA-8S, which is in agreement with previous results. This result further demonstrated the greatly enhanced thrombin inhibition by linking two aptamers. One interesting point about this approach is that only one of the aptamers needs to have the protein-inhibition activity.

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128 The other aptamer simply serves as an anchor on the protein for easy access to the thrombin by the active aptamer. More experiments were done to prove that the two aptamers in DA-8S did bind to thrombin simultaneously. The idea was to label DA-8S with fluorescein and TAMRA at the two ends. In its free form, the DA-8S had the linker longer than 16 nm, which was too far for efficient fluorescence resonance energy transfer (FRET) between the two dyes. However, the size of thrombin was likely about 3-4 nm, so when both aptamers bound to thrombin, enhancement in FRET should be observed since the two dyes would be brought close around the protein. Our results from the titration of thrombin to dual-labeled DA-8S revealed an increased TAMRA to fluorescein emission intensity ratio, indicating FRET indeed took place due to the simultaneous binding of thrombin by the two aptamers. Further experiments will be carried out to explore the application of the enhanced aptamer affinity. One idea is to use the dual-aptamer for thrombin detection in CE. The greater binding strength of the dual-aptamer is expected to be able to maintain a stable complex peak in CE. The other direction could be to study whether the new aptamer probe has better selectivity for the target and can distinguish between closely related proteins in complex biological samples. One implication of this new method for improving binding affinity of aptamers would be that in the SELEX process, the DNA library does not have to contain very long random sequences that exceed 50 bases. The longer the random sequence, the better chance one can select aptamers with high affinity. However, instead of improving binding strength on one site of the target protein, it might be a good alternative to select

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129 short aptamers with good affinity for multiple binding sites. Then these short sequences can be easily and cheaply linked and synthesized for very strong binding of the target. However, this statement is based on assumption at the moment. More comparison tests need to be conducted to draw any solid conclusions.

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BIOGRAPHICAL SKETCH Zehui Cao is from Zhuzhou, Hunan Province, China. He spent 18 years in that same province before he traveled almost 1000 miles to Nanjing, a city in eastern China, for college study. He got his B.S. degree in chemistry from Nanjing University in 1998. He spent one more year at the graduate school of Nanjing University studying organic chemistry before he traveled again over 10000 miles to University of Florida to pursue a Ph.D. degree in analytical chemistry under the supervision of Dr. Weihong Tan. 138


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Permanent Link: http://ufdc.ufl.edu/UFE0008333/00001

Material Information

Title: Designer Oligonucleotides for Probing DNA-Protein and Protein-Protein Interactions
Physical Description: Mixed Material
Copyright Date: 2008

Record Information

Source Institution: University of Florida
Holding Location: University of Florida
Rights Management: All rights reserved by the source institution and holding location.
System ID: UFE0008333:00001

Permanent Link: http://ufdc.ufl.edu/UFE0008333/00001

Material Information

Title: Designer Oligonucleotides for Probing DNA-Protein and Protein-Protein Interactions
Physical Description: Mixed Material
Copyright Date: 2008

Record Information

Source Institution: University of Florida
Holding Location: University of Florida
Rights Management: All rights reserved by the source institution and holding location.
System ID: UFE0008333:00001


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Full Text












DESIGNER OLIGONUCLEOTIDES FOR PROBING DNA-PROTEIN AND
PROTEIN-PROTEIN INTERACTIONS

















By

ZEHUI CAO


A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA


2004

































Copyright 2004

by

Zehui Cao

































This work is dedicated to my family.















ACKNOWLEDGMENTS

I would like to thank my parents for the constant care and support they give me,

and my wife, Qian Li, for her support and patience all the time. I would also like to thank

my advisor, Dr. Weihong Tan, for his confidence in me and the encouragement he gave

me to continue my study, Dr. James Winefordner for his kind help with my graduate

study, and the members of my advisory committee for their helpful guidance and

suggestions. I would like to thank Chih-Ching Huang for his helpful and enjoyable

collaboration with me on many projects, and all the members of Tan research group for

their help in my daily work and life.
















TABLE OF CONTENTS
Page


A C K N O W L E D G M E N T S .................................................................... ......... .............. iv

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

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

ABSTRACT .............. ..................... .......... .............. xii

CHAPTER

1 IN TRODU CTION .................................. .. ... ......... ... ...............

Importance of Protein Detection and Protein Interaction Study ...............................
Analytical Techniques for Protein and Protein Interaction Study..............................3
Fluorescence Techniques for Signal Transduction .........................................6
Fluorescence Quenching and Fluorescence Energy Transfer..........................9
Fluorescence A nisotropy ........................................... ......................... 13
Oligonucleotides as Probes for Protein Interactions ............................................. 17
Molecular Beacons for Nonspecific Protein Detection ..............................19
Aptamers for Specific Protein Detection Based on FRET ...........................20
Aptamers for Protein Detection Using Fluorescence Anisotropy ..................26

2 MOLECULAR APTAMERS FOR REAL TIME PROTEIN-PROTEIN
INTERACTION M ONITORING ........................................ ........................ 28

Experim mental Section ........................................ ................... .. ...... 32
R results and D iscussion............................................................ ................. ......35
FRET-Based Signaling Aptamer for Protein Binding..................................36
Dual-labeled aptamer for thrombin-protein binding study .................37
FRET-based 27mer aptamer for thrombin-protein binding .................40
Fluorescence Anisotropy (FA) Based Aptamer Probes for Protein
Interaction s .................. ...... .............. .... .. ....................... ... 4 1
Quick Evaluation of Binding Constants of Protein-Protein Interactions........45
Kinetics of Protein-Protein Interactions in Competitive Assays ..................48
C on clu sion s ........................................................................... 52









3 MOLECULAR APTAMERS-BASED AFFINITY CAPILLARY
ELECTROPHORESIS FOR PROTEIN-PROTEIN INTERACTIONS ..................55

E x p erim mental S section .................................................................... .....................5 8
C hem icals and B uffers......................................................... ............... 58
A apparatus .............................................................................................. ........59
Separation of A ptam er........................................................ ............. 60
A ptam er-B ased A C E ............................................. ............................. 60
PEG-Assisted Aptamer ACE .................................... ................ 61
R esu lts an d D iscu ssion s ........................................ ............................................62
Conform ation of aptam er...................... ................................ ............... 62
Quantification of Throm bin ........................ .......... ................................ 66
Determination of Dissociation Constant (Kd) ...........................................68
Com petitive A ssay ......... ..... ...... ...................... .. 69
Effect of PEG on Aptamer-Thrombin Complex in CE ..................................72
Aptamer-Based Mobility Shift Assay for Thrombin-AHT Interaction ..........76
C onclu sions ........................................... ............................ 78

4 NUCLEASE-RESISTANCE OF TELOMERE-LIKE SINGLE-STRANDED
OLIGONUCLEOTIDES MONITORED IN LIVE CELLS BY FLUORESCENCE
A N ISO TR O PY IM A G IN G ........................................................... .....................80

Introduction ................. ... .... ........ ........ .......... ....... .................. 80
Fluorescence Techniques for Monitoring Intracellular Biointeractions.........80
Telomere and Its Presence in Live Cells ................................................82
E xperim mental Section .................................................... ...... .......... .......... 84
Fluorescence Anisotropy Imaging (FAI) System ..........................................84
The Image Processing Program .............................................. ............... 87
C ell C ulturing ................................................. ................. 87
Cell Injection U sing Electroporation........................................ .................88
M a te ria ls ..................................................... ................ 8 8
R results and D discussions ............... .. .......................... .................. ............... 89
Measuring Anisotropy of Standard Samples ...............................................89
Effects of PMT Voltage on Anisotropy Data ...........................................92
Monitoring ssDNA Digestion in Homogeneous Solutions ..........................94
D igestion of ssD N A in Live Cells....................................... ............... .... 95
Stability of Telomere-Like Oligonucleotides in the Nuclei of Live Cells......99
C on clu sion s ........................................... ................... .. ............... 102

5 SUMMARY AND FUTURE DIRECTIONS.................................... ................ 104

Summary of Oligonucleotide-Based Protein Interaction Study .............................104
F u tu re D irectio n s ............... ................ ................................ .................. ... 10 7









APPENDIX

A PROGRAMMED DNA CONFORMATION CHANGE AND APTAMER
ACTIVATION .......... .... ........................................ ..... ........ 112

Introduction ........................ ......... .................... .................... ... ........... 112
Nanomotors and Photo-Regulation of DNA Hybridization............................... 113
Photo-Regulated Aptamer Release and Activation............................. .............116

B ENHANCED PROTEIN BINDING AND INHIBITION BY DUAL-
A PTAM ER S ...................................................... .......... .. ............ 122

Introduction to D ual-A ptam ers ........................... ........ ... ............... .... 122
Design of Dual-Aptamers for Thrombin and Experimental Results................124

L IST O F R E FE R E N C E S ........................................................... ......... .......................130

BIOGRAPHICAL SKETCH ............................................................. ............... 138
















LIST OF TABLES


Table page

2-1 Sequences of the fluorophore-labeled aptamers used in this work. .......................36

B-l Inhibition of thrombin-fibrinogen reaction by various ologinucleotides.............25
















LIST OF FIGURES


Figure p

1-1 A typical Jablonski diagrm ...... ........................... ........................................ 7

1-2 Absorption and fluorescence emission spectra of fluorescein in pH 9.0 buffer.........7

1-3 Principle of nucleic acid detection using MB............... .................12

1-4 Illustration of the principle of fluorescence anisotropy ................. ....................14

1-5 M easuring fluorescence anisotropy. ........................................................................16

1-6 Conformation change of thrombin-binding aptamer induced by thrombin .............22

1-7 Conformation change of PDGF aptamer induced by PDGF ................................25

2-1 Dye-labeled protein-binding aptamers reporting protein-protein interactions.........31

2-2 3-dimentional structure of human a-thrombin in complex with 15Ap...................35

2-3 Human a-thrombin binding induced relative fluorescence change of dual-labeled
15m er ap tam er ..................................................... ................ 3 7

2-4 Thrombin bound to FQ-15Ap interacts with other proteins..................................38

2-5 Dual-labeled 27mer aptamer for a-thrombin/protein interactions............................40

2-6 TAMRA-labeled 15Ap for a-thrombin/protein interactions based on fluorescence
anisotropy ............................................................................42

2-7 Binding between a-thrombin and anti-human thrombin (AHT) confirmed by gel
electrophoresis on a 7.5% native Tris-HCl gel. ........................................... ............ 44

2-8 TAMRA-labeled 27Ap for a-thrombin/protein interactions based on fluorescence
anisotropy ............................................................................45

2-9 Effect of the order of incubation with thrombin on thrombin-protein interaction. ..49

2-10 Rate-limiting step in aptamer-thrombin-protein interactions..............................51

3-1 Schematics of the capillary electrophoresis setup. ................................................59









3-2 Capillary electrophoresis traces of thrombin binding aptamer in the presence of 25
mM Tris, 192 mM glycine and various concentrations of KC. ........................63

3-3 Capillary electrophoresis traces of thrombin binding aptamer in the presence of 25
mM Tris, 192 mM glycine and various other metal ions .............................65

3-4 Detection of thrombin using aptamer-based ACE ............... ............. ...............67

3-5 Analyses of AT III-thrombin interaction using aptamer-based ACE.....................70

3-6 Using thrombin binding aptamer to monitor thrombin/AT III interaction ..............71

3-7 Quantification of AT III-thrombin interaction. .............................. ......... ...... .71

3-8 Effect of PEG on the stability of G-Apt*Thrmb complex................. ........... 74

3-9 Analyses of thrombin and thrombin-AT III interaction using PEG-containing
sam ple m atrix. ........................................................................75

3-10 Binding between G-Apt*Thrmb and anti-human thrombin (AHT) confirmed by
capillary electrophoresis .................. ........................... ........... ........ .... 77

4-1 Schematics of the FAI system ............ ................................. ............ ... 85

4-2 Fluorescence images obtained from IVH (left) and Ivy (right) channels of the FAI
sy ste m ...................................... .................................................... 9 0

4-3 Anisotropy of 200 nM TAMRA solutions with increasing concentration of
glycerol ........... ............................................. ....................... ......... 9 1

4-4 Effect of PMT voltage on anisotropy measurements. ............. ....... ..............93

4-5 Anisotropy change of 5'-TAMRA-(TTAGGG)6-3' (TeloH) during digestion by
DNase I in solution monitored by FAI.......................................... ................... 94

4-6 Digestion of Ctrl 1 in live cells monitored by anisotropy change.............................96

4-7 Anisotropy values obtained by averaging in the nucleus region of the anisotropy
images of two cells injected with Ctrl and Ctrl1-S respectively...........................98

4-8 Digestion of TeloH, TeloT, TeloH2 and TeloCtrl in live cells monitored by
anisotropy change............ ................. .... .. ......... .. ........ .............. .. 101

A-i Structure of azobenzene is changed by lights of different wavelengths...............14

A-2 Schematics of photo-regulated activity of thrombin aptamer ..............................17

A-3 Inhibition of thrombin by Ap-CS and Ap-MS monitored by measuring scattering
lig h t ...................................... .................................................... 1 1 9









B-l Illustration of inhibition of thrombin by a single aptamer (top) and a dual aptamer
(bottom ). ............................................................................ 123

B-2 Inhibition of thrombin by 15Ap and DA-8S monitored by measuring scattering
lig h t ...................................... .................................................... 1 2 7















Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy

DESIGNER OLIGONUCLEOTIDES FOR PROBING DNA-PROTEIN AND
PROTEIN-PROTEIN INTERACTIONS

By

Zehui Cao

December 2004

Chair: Weihong Tan
Major Department: Chemistry

Proteins are responsible for carrying out most of the functions in living cells. As the

result of their important roles in human life, changes in protein expression and properties

are directly associated with many diseases. Sensitive detection of the disease-related

proteins and understanding of their interactions with other biomolecules may be the first

step to effective disease diagnoses as well as finding cures. As a good alternative to

traditional antibody-based ligands, protein ligands based on short nucleic acids have

advantages such as inexpensive production, easy labeling, good stability, and relatively

small sizes. Oligonucleotide probes can not only target many DNA-binding proteins

nonspecifically, but also bind to and sense target proteins with high selectivity thanks to

recent developments in molecular aptamers. Both nonspecific and specific

oligonucleotide probes can be easily labeled with fluorescent tags for easy and sensitive

analyses of proteins of interest. We have explored the application of those fluorescently-

labeled oligonucleotide probes for study of DNA-protein and protein-protein interactions









in homogeneous solutions, under heterogeneous separation conditions, and even in real

time in live cells.

By combining protein-binding aptamers and signal transduction mechanisms such

as fluorescence resonance energy transfer and fluorescence anisotropy, we were able to

monitor protein-protein interactions in real time without labeling either of the two

interacting proteins, posing minimum effects on the binding properties of the proteins.

Our method has been shown to be simple and effective, with the capability of providing

detailed information regarding binding sites and binding kinetics.

We have also shown that fluorophore-labeled aptamers, with their small sizes and

charged backbones, are ideal capturing and sensing agents in affinity capillary

electrophoresis (ACE) for sensitive protein detection and protein-protein interaction

analyses. The aptamer-based ACE should be valuable in many areas of protein research.

Lastly, we combined the imaging capability of confocal microscopy with

fluorescence anisotropy measurements to monitor interactions between fluorophore-

labeled DNAs and cellular proteins in live cells. As a demonstration, DNA digestion by

nucleases was studied inside cell nuclei in real time. We found that DNAs with telomere-

like sequences were more resistant to cellular nucleases than other sequences we tested.














CHAPTER 1
INTRODUCTION

Importance of Protein Detection and Protein Interaction Study

Proteins are macromolecules that consist of one or more unbranched chains of

amino acids. They are the driving force for proper function of cells and ultimately

organisms in human life. A great variety of proteins work together or with other

molecules to realize almost all important biological functions, such as catalysis of most

biochemical reactions, regulation of cell growth and apoptosis, and controlling motion

and locomotion of cells and organisms. Because of the deep involvement of proteins in

life, diseases caused by disorders in functions of cells and organisms will always result in

changes in expression level of certain proteins and disruptions of various interactions

between proteins or between proteins and other molecules. The role of analytical

chemistry in protein related studies would be the development of simple yet effective

techniques for sensitive and selective detection of important proteins as well as their

interactions with other biomolecules. These methods can then be used to greatly facilitate

early disease diagnoses, disease mechanism understanding, and eventually drug

discovery.

With the better understanding of the genetic basis of many diseases such as various

cancers, certain biomolecules may emerge as a new class of disease markers with greater

sensitivity and selectivity. Take cancer as an example, traditionally, diagnosis of cancers

have used biomarker molecules that are produced in higher than normal level either

directly by tumor cells or by the response of the human body to the presence of cancers.









Detection of the biomarkers in a patient's body fluids can serve as the first step in cancer

diagnosis and provide critical information to doctors as whether biopsy is needed. Some

classic tumor markers include a-fetoprotein (AFP), carcinoembryonic antigen (CEA), and

prostate specific antigen (PSA). They are usually not very specific to a particular cancer

as the level of one tumor marker can be elevated by more than one type of cancer.

Another problem is that presence of cancer does not necessarily cause a detectable level

of tumor markers, especially in the early stage of the cancer. Extra caution is thus needed

in some cases to avoid false negatives. In contrast, changes induced by gene mutation can

be more specific. The mutated genes in cancer cells can lead to expression of new

proteins not found in normal cells, over-expression of certain proteins that promote cell

growth and division such as growth factors and related proteins, or mutation of proteins

that inhibit tumor cell proliferation. One example of proteins capable of indicating

cancers is the human epidermal growth factor receptor 2 (HER2) usually found on cell

membranes. HER2 has been shown to be over-expressed in about 25% of all breast

cancer patients.1 Therefore, tests for HER2 protein in tissue samples are recommended

for breast cancer diagnosis. Over-expression of many other growth factors, including

insulin-like growth factor-I (IGF-I), epidermal growth factor (EGF) and platelet-derived

growth factor (PDGF), are also related with tumor progression.2-4 Sensitive detection of

these important proteins may serve as good indications of both the presence and stage of

cancers.

Since proteins often work together with many other molecules on a specific task,

detection and analyzing of such interactions may help understand the cause of the

diseases as well as find the cure. For instance, the p53 protein is a tumor suppressor that









induces cell apoptosis under stressful conditions such as DNA damage by activate

transcription of certain genes.5 Mutation or inhibition of the wild type p53 makes p53 not

able to prevent cells from growing out of control and eventually becoming tumors. It has

been found that the protein product of MDM2 gene can bind to p53 and inhibit its

transcriptional activity, thus contributing to cancer development. The revelation of this

protein-protein interaction makes targeting MDM2 protein of great therapeutic interest.6

Inhibition of the p53-MDM2 interaction will certainly help in early tumor suppression.

Analytical Techniques for Protein and Protein Interaction Study

Many analytical techniques have been used in protein research. Overall, they can be

summarized into two major categories. One of them uses molecular ligands that

selectively recognize the proteins of interest, while the other does not rely on such

ligands. The latter approach includes many separation-based methods such as those based

on electrophoresis and chromatography. These methods usually take advantage of

differences in mass, size and charge between proteins. Among them, gel electrophoresis

allows proteins to migrate at different rates on cross-linked polyacrylamide sieving

support. When the detergent sodium dodecyl sulfate (SDS) is used, the proteins are

detangled and negatively changed, and the separation is based only on size. Another

related technique known as isoelectric focusing often uses polyamino-polycarboxylic

acid ampholytes to form a solid support with a pH gradient. A protein with charges

migrates on the support until it reaches the region where the pH equals the protein's

isoeletric point (pI) at which the protein carries no charge. In this way, proteins with

different pi's can be separated and analyzed. Introduced not long ago, another form of

electrophoresis called capillary electrophoresis (CE), is conducted in open fuse-silica

capillaries.7 Unlike traditional electrophoresis, Joule heat generated during









electrophoresis in CE can be dissipated very quickly because of the small capillary

diameter. This allows ultrahigh voltage to be applied for the separation, leading to very

rapid analyses and high efficiency with theoretical plates up to one million.8 CE

separation is based on both charge and size of the molecules, making it ideal for analyses

of biopolymers such as nucleic acids and proteins.

Filled-column based chromatography is also used for protein analyses. Besides the

widely used HPLC with stationary phases based on hydrophobicity and hydrophilicity,

one commonly used method is ion-exchange chromatography, in which resins with

charged groups are present in the stationary phase to retain molecules with opposite

charges. Molecules of the same charge as the resins come out first. Another type is the

size-exclusion chromatography, where beads with pores of certain sizes are packed in the

column. Molecules smaller than the pore size will get trapped more easily in the pores,

and thus migrate more slowly than molecules of bigger sizes.

One thing especially noteworthy is that recent advances in proteomic research have

resulted in a powerful approach for large-scale protein analyses. The development of

two-dimensional gel electrophoresis (2-D gel), combined with breakthroughs in mass

spectrometry (MS), especially soft ionization techniques such as electrospray ionization

(ESI)9 and matrix-assisted laser desorption ionization (MALDI),10 have made high-

throughput protein analyses possible.11

The separation-based techniques for protein analyses require no protein

modifications and can often detect multiple targets. However, a few factors limit their

application in some areas of protein study. First, most of the time, good separation may

require specifically optimized buffer conditions, which may be very different from those









of cellular environments. Proteins may likely behave very differently than in their native

system, which makes it undesirable to study protein interactions. Furthermore, detection

of proteins is done after the separation is completed. Thus it may take more time than

protein assays that are capable of detection in real time. When it comes to large-scale

protein analyses, the time difference may be even more significant. More importantly,

detection in a non-real-time fashion makes it near impossible to study kinetics of protein

functions or interactions.

A very different approach for protein study than the separation techniques is the use

of selective binding ligands for targeting proteins. The ligands can be proteins, nucleic

acids and many other molecules. However, the most widely used protein ligands are a

special category of proteins called antibodies. Antibodies are produced by the immune

systems of human or animals upon the invasion of foreign molecules (antigens) such as

proteins, carbohydrate polymers, and nucleic acids.12 They usually contain two heavy

polypeptide chains and two light chains, and have molecular weights ranging from 150

kDa to about 950 kDa. Antibodies are able to selectively bind to targets with very high

affinity constants, between 105 and 1010 M-1. As a result, they have been the preferred

recognition agents for protein detection for the last decades.

One of the commonly used antibody-based protein assays is the enzyme-linked

immunosorbent assay (ELISA).13 Although in many different forms, this technique is

generally based on antibody-antigen recognition. In a simplified form of the ELISA

assay, an antibody is immobilized on a solid surface as a capture agent for the antigen.

Samples containing the target protein are incubated with the surface-bound antibodies.

The targets will be captured by the antibody while other molecules can be washed away.









The following step usually involves the use of another antibody that also binds to the

antigen at a different site. Thus the two antibodies and the antigen form a sandwich-like

complex on the surface. The second antibody has been linked to an enzyme previously,

so that when solution containing substrate for the enzyme is added to the sandwich

complex, the enzymatic reaction can take place. The enzyme and substrate pair is

carefully chosen in a way that the colorless substrate will become colored after the

enzymatic reaction. ELISA assay is very sensitive because the enzymatic reaction is

actually a signal amplification process. The other major advantage is that sample does not

need to undergo stringent purification before the test, which may save sample preparation

time. ELISA has been routinely used to detect proteins in serum samples. Despite the

popularity of this technique, one problem with ELISA is that the antibodies that can be

used are limited. The assay may take hours for the enzymatic reaction to complete, so

apparently it is not a real-time detection method.

Fluorescence Techniques for Signal Transduction

Fluorescence has been one of the most important tools for molecular sensing in

many fields of research. Upon absorbance of the energy of the incident light, electrons of

the fluorescent molecules can be excited to the excited singlet states. Return of the

electrons to the ground state is accompanied by emission of photons, resulting in

fluorescence. The process of fluorescence is illustrated by the Jablonski diagram in

Figure 1-1.

So, S1, and S2 stand for the singlet ground, first, and second electronic states

respectively, while T stands for triplet state. As one can see, there is less emitted energy

than absorbed energy, thus fluorescence always occurs at a longer wavelength than that

of the incident light. The difference between a fluorescent molecule's wavelength of









maximum absorption and its wavelength of maximum emission is termed Stokes' shift.

Figure 1-2 shows typical absorption and emission spectra of a commonly used

fluorophore known as fluorescein.


S2

Si



Absorptio


>n


hv hvj


rsion

-- Intersystem Crossing

T1


Fluorescence
L hv


Phosphorescence


Figure 1-1. A typical Jablonski diagrm.


350 400 450 500 550 600 650
Wavelength (nm)



Figure 1-2. Absorption and fluorescence emission spectra of fluorescein in pH 9.0
buffer.14


\ \ I i


i Internal Conve
,t









Because fluorescent molecules have their distinct absorption and emission profiles,

they can be selectively detected using related instruments. That is, by measuring

fluorescent emission from an unknown sample, it is possible to tell if a certain

fluorophore is present. Another major advantage of fluorescence techniques is its very

high sensitivity. Since fluorescence is a photon-producing process, use of ultra-sensitive

detectors capable of single-photon counting results in very sensitive fluorescence

measurements, and even detection of single molecules is possible.15

Even though there are many ways to utilize fluorescence for molecular recognition,

the most basic approach is to tag fluorophores to the analytes that are not fluorescent by

themselves for detection purpose. Once the fluorescent molecule is linked to the non-

fluorescent target, it is possible to sense the target by measuring fluorescence from the

fluorophore. There are two ways to link fluorophores to target molecules. First, in some

applications, the fluorophore can be directly adsorbed onto the target through non-

covalent forces. One example is the dye ethidium bromide for probing double-stranded

DNA (dsDNA). When mixed with target sample, it can intercalate into the double strand

of the double-helical DNA and give strong fluorescence.16 The second approach to tag a

target with a dye is more indirect. The fluorophore is first attached to a probe molecule,

usually via covalent bonding. The probe should have good affinity selectively for the

target molecule. So when the target and dye-labeled probe are mixed together, the target

can be fluorescently tagged and later detected. For protein targets, antibodies are the most

widely used probes. Fluorescent labels have replaced radioactive labels in antibody-based

immunoassays for highly sensitive antigen detection.17









One thing that needs to be addressed when conducting fluorescence-based analyte

detection is how to separate the fluorescence signal of the fluorophore linked to the

analyte from that of the fluorophore in its free form. A simple way to do this is to use

ELISA-like format to capture the target on a solid surface and then stain it with

fluorophore. The unbound fluorophore in solution can be easily washed away later. In

this way, any fluorescence coming from the solid support should be originated from

target-bound fluorophore. Another way to isolate useful fluorescence signal is to combine

dye-labeled protein probe with separation techniques. For example, after the analyte and

the dye-labeled antibody are mixed together, separation can be carried out using CE or

HPLC. The antibody-analyte complex is isolated from the free dye-labeled antibody due

to their size and charge differences. Under fluorescence detection, two peaks representing

each of them will be present, indicating the presence of the analyte. Any other species

will not be recognized because of the lack of fluorescence.

Both approaches can take advantage of the high sensitivity and selectivity of

fluorescence measurements. However, the detection does not proceed until after the

binding event has happened, leading to difficulties in kinetics study. As a result, a

detection scheme is needed to produce signal change during the target recognition

process. Fortunately, fluorescence based detection can adopt various designs to achieve

such a goal. Two major approaches capable of real time target detection have been

employed in our work.

Fluorescence Quenching and Fluorescence Energy Transfer

There are a variety of ways to decrease fluorescence emission of a fluorescent

molecule. The process of decreasing fluorescence is termed quenching. s Quenching

happens through two major mechanisms. One of them is collisional quenching. When a









fluorophore is excited by an incident light, it leaps onto the excited state and will stay

there for a very short period of time, usually in the nanosecond range, before coming

back to the ground state. The time it stays on the excited state is called fluorescence

lifetime. During its lifetime, the excited fluorophore could collide with other molecules in

the solution. The collision may cause energy loss of the fluorophore, which means that

the absorbed energy is dissipated in a way other than fluorescence emission resulting in

fluorescence quenching. The decrease in fluorescence intensity can be described using

the Stem-Volmer equation:

Fo/F = +K [Q] = 1+ kqo[Q]

where K is the Stern-Volmer quenching constant, kq is the bimolecular quenching

constant, To is the unquenched lifetime, and [Q] is the quencher concentration.18 Many

molecules can be collisional quencher, including oxygen, halogens, amines and

acrylamide.

The other type of quenching is called static quenching, where the quencher can

form non-fluorescent complex with the fluorophore. This quenching happens even before

the fluorophore is excited to the excited state.

In addition to fluorescence quenching, another related important fluorescence

process is the fluorescence resonance energy transfer (FRET). This process involves an

energy donor which should be fluorescent, and an energy acceptor. FRET happens when

the emission spectrum of the donor overlaps with the absorption spectrum of the

acceptor. Because of the overlapping, the photon energy may be able to transfer between

the donor-acceptor pair. One important thing to note about FRET is that the donor does

not emit photons for the energy transfer to occur. The donor and acceptor have to be









coupled by a dipole-dipole interaction.18 For this interaction to take place, the two

molecules need to be very close, typically less than 10 nm. In fact, the efficiency of the

energy transfer is highly dependent on the distance between the donor and acceptor. The

relationship can be described using the following equation:

E=Ro6 / (R6 + r6)

where E is the energy transfer efficiency, r is the distance between the donor and

acceptor, and Ro is the Firster distance of the donor-acceptor pair. The Firster distance is

defined as the distance between the specific donor-acceptor pair at which the FRET

efficiency is 50%. The Firster distance is in the range of 3-6 nm, within the size of many

macromolecules.

The acceptors do not have to be fluorescent. When a non-fluorescent acceptor is

used, the result of the energy transfer is quenching of the donor fluorescence. Some non-

fluorescent acceptors can act as "dark quenchers", meaning they can effectively quench a

broad range of fluorophores whose emission spectra overlap with their absorption

spectra. Examples of dark quenchers include dabcyl, Black Hole QuenchersTM (BHQTM-1

and BHQTM-2, Biosearch Technologies, Inc., Novato, CA).

Since FRET is highly dependent on distance between the donor and acceptor, if

there is a way to design a probe such that with and without target molecules, there would

be a change in the donor and acceptor distance, then the fluorescence change can directly

report the presence of the target. One important example of utilizing this idea is the

development of molecular beacons (VMBs) for detection of nucleic acids in real time.19

Figure 1-3 illustrates the principle of how a MB recognizes its target nucleic acid:














S-C



MB Target Hybridization

Figure 1-3. Principle of nucleic acid detection using MB.

The MB is a synthetic oligonucleotide, generally ranging from 25-50 bases in

length. The molecule contains a stem and loop structure (Figure 1-3). The single-

stranded loop region consists of a probe sequence complementary to the intended target

nucleic acid. The sequences directly flanking the loop region are complementary to each

other but unrelated to the target. Thus, these flanking sequences anneal to form the MB

stem. Signal transduction in MBs is accomplished by FRET. A fluorophore is covalently

attached to one end of MB, and a quencher, often dabcyl, is covalently coupled to the

other end. The fluorescent dye acts as the energy donor, and the quencher acts as a non-

fluorescent acceptor. When the stem sequences are hybridized with each other, these two

moieties are kept in close proximity to each other, causing the fluorescence of the donor

to be quenched by energy transfer. In the presence of target DNA (tDNA), however, the

loop region forms a hybrid that is longer and more stable than that of the stem. This

forces the MB to undergo a spontaneous conformational change that forces the stem

apart. With the quencher no longer positioned near the fluorophore, fluorescence is

restored, thus signaling the binding of the MB to its target. Since signal is generated only

in the presence of the target DNA, there is no need to separate the hybrid and the MB

itself. As a result, continuous monitoring of the fluorescence signal can be used for real-

time tDNA detection as well as for hybridization kinetics study.









Another report used a second fluorophore as the acceptor instead of a quencher.20

In such a design, the energy transferred from the donor will excite the acceptor which

then emits photons. If we assign ID and IA as emission intensity of the donor and acceptor

respectively, the result of FRET between a closely located donor and acceptor is the

decrease ofID and increase ofIA. In fact, for the MB labeled with two dyes, the ratio of

A/ ID is used to sense the presence of tDNA, which provides better sensitivity than

simply measuring IA or ID.

Besides being widely employed for real-time nucleic acid detection, FRET can also

be suitable for analyzing proteins. One example is the detection of a DNA-binding

protein, catabolite activator protein (CAP), using DNA as the probe.21 CAP binds to a

sequence-specific short piece of double-stranded DNA (dsDNA). To construct the probe

for CAP, the specific dsDNA was broken into halves from the middle of the sequence.

The two pieces had short complementary overhangs labeled with a fluorophore and

quencher respectively. The overhangs could help the two pieces come back and anneal to

form the protein-binding dsDNA. However, the length of the overhang was designed to

be short enough that in the absence of CAP, the two fragments would not be able to

anneal. Only when CAP was present, could the annealing be stabilized because the

protein binding favored the formation of the dsDNA. As the two half probes came close

to each other, quenching of fluorescence was induced by FRET between the fluorophore

and quencher. In this way, rapid protein detection was successfully conducted in

homogeneous solutions.

Fluorescence Anisotropy

Like FRET, fluorescence anisotropy (FA) is another choice for real-time analyte

detection. FA is related to the phenomenon that upon excitation with polarized light,









fluorescent molecules often give depolarized emission. Anisotropy can be considered to

describe the extent of such depolarization. Figure 1-4 shows an illustration of the

principles of ansitropy.

'VH
Vertically polarized w
excitation IV
STime (ns) Emission






Time (ns) Emission




S- Excited fluorophore Unexcited fluorophore


Figure 1-4. Illustration of the principle of fluorescence anisotropy.

The subscripts V and H in Figure 1-4 refer to the orientation (vertical or horizontal)

of the polarization for the intensity measurements, with the first subscript indicating the

direction of the polarization of the excitation and the second for the polarization of the

fluorescence emission. When excited by a polarized light, the fluorescent molecules that

have absorption transition moments oriented along the electric vector of the incident light

are preferentially excited.18 During the lifetime on the excited state, usually nanoseconds,

those molecules may still rotate to other directions before returning from the excited state

to the ground state and emitting light. As a result, the emission contains not only

components with the same polarization state as the excitation source, but also light of

other polarization directions. To define this depolarization quantitatively, the concept of

anisotropy (r) is introduced and described using the following equation:

r = (Ivv-IvH) / (Ivv+2lvH)









Apparently, the extent of the depolarization is related to the fluorescence lifetime of

the fluorophore and how fast the fluorophore can rotate in its microenvironments. Also

illustrated in Figure 1-4, larger fluorescent molecules rotate more slowly on the excited

state, resulting in a smaller IVH component in the emission and consequently a higher

anisotropy value. There are other factors that can influence rotation of molecules and

their anisotropy as well, e.g., the viscosity of the solution.

Measurement of anisotropy is usually carried out with two polarizers. One of them

is placed in front of the light source to generate vertically polarized excitation. The other

polarizer is located somewhere between the sample and the detector. By rotating the

emission polarizer to the direction parallel or perpendicular to that of the excitation

polarizer, the two polarization components of the emission Ivv and IVH can be obtained.

Figure 1-5 shows a simplified setup for anisotropy measurements.

The Ivv and IVH values obtained are then applied to an equation for anisotropy

calculation,

r = (Ivv-GIvH) / (Ivv+2G-IvH)

Compared to the previous equation for anisotropy, one may notice the addition of a

G parameter, which is called G-factor. The introduction of G-factor is because the optical

components and the detector in the instrument may have different response to different

polarization states. For example, even though the sample generates same amount of vv

and IVH, after all the optical path and detector readout, the final result may be that the two

values become very different. In such cases, the G-factor is needed to calibrate and make

up for the instrument caused error. Determination of G-factor is usually done by

measuring IHH and IHV, and calculating using G = IHV / IHH.











Polarizer Iv Sample

Light
source ------
Sample
emission

Polarizer





Detector


Polarizer Iv Sample

-k~----t---P

Sample
emission

Polarizer

IVH



Detector


Figure 1-5. Measuring fluorescence anisotropy.

The connection between molecular weight of the fluorescent molecule and

fluorescence anisotropy makes anisotropy an ideal method for detection of

macromolecules and biomolecular interactions. A probe molecule can be labeled with a

dye and used to interact with its protein target. The change in the size of the probe caused by

the interaction can lead to a detectable anisotropy change, thus reporting the target molecule.

Dye-labeled DNA molecules have been employed to study interactions with proteins under

different conditions.22 Alternatively, protein can also be labeled with fluorophores to study

protein-DNA and protein-protein interactions.23;24 In theory, as long as the biological

interaction induces changes in the rotation of the fluorescently labeled probe, anisotropy

should be applicable for real-time analyzing of such interactions. On the other hand, other

commonly used fluorescence techniques including FRET often require precise

conformational change of the probe molecule upon target binding to correctly report the

interaction. Other advantages of fluorescence anisotropy include: 1) FA is able to use only

one dye label to report target molecules in real time and in homogeneous solutions; 2) FA is a









ratiometric approach, thus less affected by sample fluorescence fluctuation and photo-

bleaching.

One important thing needs to be taken into account when analyzing anisotropy data

is that measured anisotropy from a solution sample is the sum of the contribution from all

fluorescent species in the solution. For example, a probe P is labeled with a fluorophore F

to give PF. In the given solution, PF has an anisotropy value of rpF. When analyte A

containing sample is added to the solution of the PF, the probe will bind to the analyte to

form a complex A-PF. Anisotropy of the complex, rA-PF is higher that rPF. However, the

anisotropy measured for this mixed solution is determined by the following equation,

r = (l-) rpF + X' rA-PF

where X is the fraction of the total PF that has been converted to the complex. It is clear

that the more A is added to form more A-PF, the higher the measured anisotropy will be.

When A is in excess and all PF is bound to the analyte, the highest anisotropy is

achieved, which equals to rA-PF. This constructs the basis for quantitative detection of

target using fluorescence anisotropy. However, a disadvantage of fluorescence anisotropy

is also revealed here, which is the limited dynamic range of detection, in this case,

between rPF and rA-PF.

Oligonucleotides as Probes for Protein Interactions

As discussed before, antibodies have been the preferred protein-binding ligands for

decades. However, some of their intrinsic properties may limit their applications in

certain protein research fields. First, the production of antibodies involves using animal

hosts and is a rather time-consuming process. Second, like any other proteins, antibodies

are also sensitive to their surrounding environments and may be denatured by changes in

pH and salt concentrations. Nonspecific adsorption of antibodies to many solid surfaces









can sometimes destabilize them, which limited their application on a solid supports.

Moreover, the larger molecular weight of the antibodies, over 150 kD, is not practical for

many assays such as the fluorescence anisotropy where a significant size difference

between the probe and the target is desired. In an addition, antibodies lack the ability to

have a built-in signal transduction mechanism. Consequently, the detection of target

proteins requires either fluorescent labeling of the targets or a second signaling probe.

Recent development in antibody fragments uses only the antigen-binding fragments of

the antibody for target capturing.25 Antibody fragments improve upon antibodies in the

areas that they can be produced inexpensively in large scale and are much smaller. But

the stable Fab fragment is about 50 kD, still comparable to or larger than many common

proteins.

On the other hand, nucleic acid based protein-binding ligands may be excellent

candidate for studying protein-DNA and protein-protein interactions for a couple of

reasons. First, nucleic acid probes can be small and flexible, and they are routinely

synthesized using DNA synthesizers. Second, there are many ways to attach functional

groups and fluorescent dyes to any position of the sequence. Third, nucleic acids,

especially DNAs, can withstand harsher environments than proteins and are less sensitive

to many buffer conditions. Immobilization of DNAs onto solid surfaces is also relatively

easy, with multiple methods to choose from. DNA ligands can be used to study proteins

that have non-specific affinity for DNA, such as nucleases. Proteins like transcription

factors bind to specific DNAs thus can be studied using the DNAs with the right

sequences. Recent advances in selecting protein-specific DNA ligands make it possible to

analyze any protein of interest with nucleic acid probes.









A few approaches have been adopted to analyze proteins in real time and in

homogeneous solution, such as FRET and anisotropy based assays.

Molecular Beacons for Nonspecific Protein Detection

Although molecular beacons were originally designed for binding and recognition

of specific nucleic acids, these probes can also lead to increased fluorescence upon

binding of certain proteins. The protein recognition ability of MBs was first

demonstrated with a single-stranded DNA binding protein (SSB).26 SSB is a 75.6 kDa

tetrameric protein that acts as an accessory protein in DNA replication, recombination,

and repair. A tetramethylrhodamine (TAMRA)/dabcyl molecular beacon was used, and

SSB concentrations as low as 20 nM could be detected using a conventional fluorescence

spectrophotometer. Monitoring fluorescence over time shows that the SSB-MB

interaction is rapid, reaching equilibrium within 10 s. In fact, the MB binds with the SSB

much more quickly than with its complementary DNA. The MB-based SSB assay is,

however, not particularly specific. SSB leads to a fluorescence enhancement nearly equal

to that of the complementary DNA, but other proteins can also bind with the MB and

cause a fluorescence increase. For example, histone and RecA proteins have both been

demonstrated to bind with MB and augment fluorescence.

The selectivity of MBs for protein detection was further examined through detailed

binding studies of the ssDNA-binding enzyme lactate dehydrogenase (LDH).27 LDH

occurs as five distinct isoenzymes, and MBs were used to elucidate how minor structural

changes in the protein affect its ability to bind ssDNA. Two LDH isoenzymes from three

different species were assayed using MBs, and the resulting fluorescence varied by as

much as 80% between the various samples.









Together, these results demonstrate that while MBs are sensitive and somewhat

selective to DNA-binding proteins, they are not specific enough to be capable of

distinguishing a particular protein. However, the utility of MBs for studying non-specific

DNA-binding proteins in enzyme cleavage assays has already been noted.

Aptamers for Specific Protein Detection Based on FRET

In order to overcome the limitation of non-specific DNA probes for proteins and be

competitive with antibodies, nucleic acids ligands that selectively recognize target

proteins need to be developed. As a result of great efforts in this direction, aptamers have

emerged and gained great attention from researchers. Aptamers are nucleic acids that

have high affinity and selectivity for their target molecules. By using the systematic

evolution of ligands by exponential enrichment (SELEX) process,28;29 oligonucleotide

sequences can be isolated to recognize virtually any class of molecules.30 The SELEX

process begins with a library of synthesized oligonucleotides usually containing 1014 to

1015 random sequences. This library is then incubated with the target molecule of interest

under certain conditions. The sequences that interact with and bind to the target

molecules are isolated for the next round of incubation. This process is repeated until a

sequence that binds to the target with the highest affinity and selectivity is determined.

Compared to antibodies, aptamers also have high affinity and selectivity for proteins.31;32

Advantages of aptamers over antibodies roots from the easy production, easy labeling,

and chemical stability of nucleic acid molecules in comparison to protein molecules.

In order to employ aptamers for real-time target detection, FRET has been chosen

by many researchers as the signal transduction mechanism. In the FRET assay, the

aptamer is labeled with a fluorophore and a quencher similar to an MB assay. When the

aptamer specifically binds to its target, a consequent conformational change of the









aptamer may take place. This change usually results in a change in the distance between

the fluorophore and the quencher, leading to changes in the quenching efficiency as well

as in the measured fluorescence. By carefully designing the positions of the two labels, a

large difference in fluorescence intensity before and after aptamer-target binding can be

achieved. Therefore, very small amounts of target molecules are detected using highly

sensitive fluorescence measurements.

The first MB-like aptamer probe was reported for a small molecule. Stojanovic et

al. developed such a sensor based on a previously reported aptamer for cocaine

detection.33 The cocaine-binding aptamer sequence was labeled at the two ends with a 6-

FAM dye and a dabcyl respectively. The presence of cocaine would bring the two termini

together to form a stable secondary structure. The fluorophore and quencher were also

brought close so that the quenched fluorescence could be used to report cocaine

molecules.

A similar approach was reported for the detection of the Tat protein of HIV-1.34

The RNA aptamer was also split into two subunits. One of them was labeled with both

fluorophore and quencher in a MB-like structure, so that fluorescence was quenched in

the absence of the target. The presence of the Tat protein would stabilize the

hybridization between the two subunits and open the MB-like structure of the first

subunit. A fluorescence enhancement as high as 14-fold could be obtained for sensitive

detection of the Tat protein.

Two groups developed aptamer beacons for human a-thrombin in about the same

time period.35;36 Thrombin is a serine protease that works with fibrinogen and Factor 13

to help stop bleeding. The first reported aptamer for thrombin contains a 15-nucleotide









consensus sequence, 5'-GGTTGGTGTGGTTGG-3'. NMR and X-ray diffraction studies

have demonstrated that this sequence can adopt a compact unimolecular conformation

termed a quadruplex containing two G-quartet structures.37-39 When bound to thrombin,

the aptamer exists primarily in its quadruplex form, while in free solution, it can adopt a

more relaxed linear conformation, depending in part upon the ionic strength and

temperature (Figure 1-6). Low salt concentrations favor the linear state of the aptamer at

room temperature.


F

F



10
Thrombin 1
-------------------------------6

2 14---- -
\ ^ '3---- --4.
2,
13 12
3 4



Figure 1-6. Conformation change of thrombin-binding aptamer induced by thrombin.
When bound to thrombin, the 15-base aptamer forms a quadruplex
conformation with the protein binding primarily in the base 4-12 region.
Fluorescence is quenched upon thrombin binding as the fluorophore and
quencher moieties are pulled close together.

Based on this conformational change, Li et al. developed an MB-like aptamer

probe.36 When the aptamer is labeled with a fluorophore-quencher pair, the distance

between the two labels changes as the aptamer binds to thrombin and takes the compact

quadruplex form. To maximize this distance change, and thus to obtain the largest

fluorescence decrease upon thrombin binding, a 6-FAM dye and a dabcyl quencher were









linked to the two ends of the aptamer. The fluorescence of 6-FAM was seen to decrease

instantly when excess thrombin was added to the aptamer probe. Furthermore, a titration

of the probe by thrombin revealed a detection limit of 370 pM (S/N>3). The selectivity of

the aptamer-based probe was well demonstrated by mixing the aptamer with different

proteins of interest. The results clearly showed that other proteins, including the closely-

related y-thrombin, did not display much quenching of 6-FAM. In order to obtain even

better sensitivity, the fluorophore-quencher pair was replaced by a two-fluorophore pair,

coumarin and 6-FAM. The absorption spectrum of 6-FAM overlaps with emission

spectrum of coumarin, making FRET between the two dyes possible. Binding of the two-

fluorophore aptamer probe to thrombin resulted in decreased coumarin fluorescence and

increased 6-FAM intensity. When the ratio of the two intensities was used to build a

calibration curve, a lower detection limit of thrombin was achieved at -112 pM.

The approach Hamaguchi et al. took to detect thrombin was to add a few bases to

one end of the thrombin aptamer that were complementary to the sequence at the other

end.35 With a fluorophore-quencher pair labeled at the two ends, this sensor gave low

fluorescence in the absence of thrombin because of the double helix stem formed

between the two complementary termini. Upon binding with thrombin, a more compact

quadruplex structure of the aptamer would be favored and the two stem ends were

separated. Thus, an enhanced fluorescence was the indication of the presence of the

target.

FRET-based aptamer probe was later used for real-time detection of cancer-related

proteins. One of the often targeted oncoproteins is platelet-derived growth factor (PDGF),

a dimeric protein composed of a combination of subunit A and B. Out of the three









isoforms of the protein (PDGF-AA, PDGF-AB and PDGF-BB), the BB form, in

particular, has been implicated in tumor growth and progression. While generally

undetectable in normal cells, PDGF is overexpressed in a variety of human tumors

including gliboblastomas and sarcomas.

Aptamers that are selective to the PDGF-B chain have been previously isolated.40

When bound to the protein, the consensus secondary structure motif of the PDGF

aptamers is a three-way helix junction with a conserved single-stranded loop at the

branch point. However, without PDGF, only two of the helixes are stable in physiological

conditions while the third helix containing the two ends of the aptamer is separated into

two strands (Figure 1-7). This conformational difference forms the basis for the

construction of a FRET-based PDGF probe. By labeling the two ends of the aptamer with

a fluorophore-quencher pair, the fluorescence of the fluorophore is expected to decrease

when the aptamer is bound to PDGF.

Such a PDGF probe was designed by labeling the 5'-end of the aptamer with

fluorescein and the 3'-end with a dabcyl quencher.41 To test the capability of this probe

for real-time quantitation of PDGF, a series of titration experiments were conducted in a

physiological buffer at 370C. The fluorescence of fluorescein was found to decrease as

the PDGF concentration increased. The lowest PDGF concentration that could be

detected was determined at 0.11 nM. The dissociation constant, Kd, for PDGF-aptamer

complex was calculated to be 3 nM. The very high affinity of aptamers for their targets

ensures the probe's capability to quantitate trace amounts of target proteins in

homogeneous solutions. This aptamer-based assay showed a good ability to differentiate

between PDGF-BB and other proteins. The addition of a 5-fold excess of unrelated









proteins including lysozyme, BSA, hemoglobin, and myoglobin had no observable effect

on the fluorescence. Unrelated peptide growth factors such as epidermal growth factor

and insulin-like growth factor 1 also did not appear to bind with the aptamer probe.


,A\ \ C,
S- T
a G A, /

G COG




Physiological buffer PDGF present


Figure 1-7. Conformation change of PDGF aptamer induced by PDGF. Binding of PDGF
causes the aptamer to change from a relatively unconstrained structure to a
tightly packed one containing a 3-way helical junction with a conserved
single-stranded loop.

In the same report, the authors continued their work and applied this FRET-based

assay to detect PDGF in real cellular samples. For a cell line that was known to secret

PDGF, Serial dilutions of protein preparations from each cell line were incubated with a

fixed amount of the aptamer probe in a 96-well microtiter plate to obtain dose response

curves. Presence of PDGF-BB gave a dose response curve with sharp slope. In
comparison, PDGF-AB and AA gave less steep slopes.

All the above mentioned examples have demonstrated that by combining the FRET

signal transduction mechanism, and the excellent affinity and selectivity of the aptamers

for their target proteins, easy yet sensitive assays can be built to analyze proteins of

interest in real time without need for any separation, even in relatively complex
biological samples.
biological samples.









Aptamers for Protein Detection Using Fluorescence Anisotropy

The primary advantage of using aptamers as anisotropy probes over many other

molecules, especially antibodies, is their relatively small sizes. They are usually not only

much smaller than monoclonal antibodies, but also smaller than their target proteins in

most cases. This makes aptamer-based anisotropy probes ideal for protein detection. With

the aptamer conveniently labeled with a fluorophore, the anisotropy probe will report

binding with a target protein via the increase in the anisotropy of the fluorophore. Unlike

the FRET-based assays, where the conformational change of the aptamer is essential for

target detection, the aptamer anisotropy assay is not as heavily dependent on the structure

of the aptamer probe, and thus may be highly useful for applications where the

understanding of the aptamer structure is limited.

In an effort to explore the application of an aptamer anisotropy probe for protein

analysis, the PDGF aptamer was labeled with a single fluorescein dye at the 5'-end.42 PDGF

in the nM range, when added to the aptamer solution, caused a significant anisotropy increase

due to the overall larger molecular weight of the complex. Controls were done using pure

fluorescein dye mixed with PDGF and no perceptible change in anisotropy was observed.

The detection limit of this assay was determined to be 2 nM.

In conclusion, the selectivity of the anisotropy probe was tested to be as good as the

FRET-based assays. Among a variety of control proteins, not only did unrelated proteins

produce no observable changes in the anisotropy of the aptamer, but also the three

isoforms of PDGF could be differentiated by the anisotropy assay, with highest

selectivity for PDGF-BB. The fluorescence anisotropy increases measured with the

PDGF-AB and PDGF-AA isoforms were only -40% of that resulting from PDGF-BB

binding.






27


Overall, the nucleic acid based protein probes may be able to replace antibodies in

many fields of protein research, especially with current trend of proteomics aimed at

large-scale and high-throughput analyses of whole cell proteins. While much of the

research with nucleic acid probes is simply focused on protein detection, we have applied

those probes for protein function study by investigating protein-DNA and protein-protein

interactions.














CHAPTER 2
MOLECULAR APTAMERS FOR REAL TIME PROTEIN-PROTEIN INTERACTION
MONITORING

The functions of living cells are mostly executed and regulated by proteins. The

important roles of proteins are often realized through interactions between two or more

proteins. As an example, growth factor proteins interact with their receptors on the cell

membrane to regulate the proliferation of the cells. In another example, serine protease

thrombin works with fibrinogen and Factor 13 to help stop bleeding. In order to

understand how cells fulfill their functions and how they react to changes in the

environments, it is necessary to gain insight into how proteins interact with each other

under different conditions. The function of proteins is regulated by their structure, which

is necessary for correct interaction with other molecules small (e.g., the substrate for an

enzyme protein) or large (e.g., the chromosomal DNA for DNA and RNA polymerases).

The less the proteins are perturbed, the truer information one can obtain about protein-

protein interactions.

It is well known that proteins fold into certain tertiary structures through a variety

of bonds and interactions such as hydrogen bonds and hydrophobic effects. The functions

of proteins in biological systems are highly dependent on their tertiary structures. As a

result, chemical modifications to proteins such as dye labeling may cause a reduction or

even a loss of protein activities by either directly blocking the active binding sites or

affecting the three-dimensional folding of the proteins. Therefore, it is highly desirable to

avoid any modifications of proteins when monitoring protein-protein interactions in order









to obtain the most "true-to-life" information. However, it remains challenging to

accomplish label-free protein interaction monitoring in real time. Bioanalytical

techniques based on molecular separation such as gel electrophoresis and capillary

electrophoresis (CE) can provide label-free protein-protein binding detection in a

relatively complex system,43 but they lack the ability of real-time analysis in

homogeneous solutions. Surface plasmon resonance (SPR) is another technique used to

probe two interacting proteins. It requires no protein labeling but usually needs one of the

proteins to be immobilized on a sensor silica surface.

More recent development in protein-protein interactions is the yeast two-hybrid

system that was first reported in 1989.44 In this system, two proteins are fused to the

DNA-binding domain (DNA-BD) and the transcription activation domain (TA) of a yeast

transcription factor respectively in the yeast nucleus. The interaction between the two

proteins will bring DNA-BD and TA in close proximity which results in the expression of

the reporter gene. This method has been widely used to study protein-protein interactions

and recently it has been adapted to map protein interactions on a proteome-wide

scale.45;46 However, several factors may limit the application of the two-hybrid system in

certain areas. First, it consists of rather time-consuming and labor-intensive procedures

compared to some other techniques when used on a small number of proteins. Second,

the interactions have to take place in the yeast nuclei, which may be problematic for

proteins that do not function well in the nuclei such as the membrane proteins. Moreover,

transcription factors and some other proteins can activate the reporter gene expression by

themselves, leading to false positives. Another commonly used technique for protein-

protein interactions is based on fluorescence resonance energy transfer (FRET), where









two dye-labeled proteins interact with each other to trigger a fluorescence signal change

of the fluorophores due to the energy transfer. It is simple and can be easily adapted for

most proteins. Still, some proteins may lose their biological activities after they are

modified or labeled with dye molecules.

Here we describe a convenient and versatile method for real-time protein-protein

interactions based on a competitive assay using protein-binding aptamers. Aptamers

possess affinity and selectivity, comparable to those of antibodies, for their intended

protein targets. At the same time, as discussed previously, aptamers have many inherent

advantages over antibodies in the field of real-time protein analyses.

Despite being excellent molecular probes for proteins, aptamers have not been used

extensively to study the interactions between their target proteins and other proteins. It

was reported that aptamers were used for protein-binding small molecule screening and

radioactive isotope labeling was used for detection.47 More recently, an assay was

constructed for protein interactions based on protein-dependent ribozymes combined with

aptamers.48 Here we have developed a new competitive assay using protein-binding

aptamers directly for protein-protein interactions. Based on the highly stable and flexible

structures of aptamers, two signal transduction strategies were established to detect the

binding events between the aptamer-binding protein-"bait protein", and a second

protein- "prey protein". As illustrated in Figure 2-1, in one approach, the aptamer was

labeled with a fluorophore and a quencher to form internal FRET. Binding of the aptamer

to the bait protein caused a quenched fluorescence while the binding of the prey protein

to the bait protein may displace the aptamer and result in a restoration of fluorescence. In

the other approach, the aptamer was labeled with only one fluorophore and the









fluorescence anisotropy of the aptamer was monitored in real time. Binding of the

aptamer to the much larger bait protein molecules resulted in an increased anisotropy.

Further change in the fluorescence anisotropy of the aptamer could be triggered by the

interaction between the bait and prey proteins.


CIZD2+


+ N^, ,
Q


F

+ \ 1Th


:L~J~~'" L~ +


Figure 2-1. Dye-labeled protein-binding aptamers reporting protein-protein interactions.
(A) Aptamer is dual-labeled with a fluorophore and a quencher. The folded
form of the aptamer when it binds to the bait protein results in a quenched
fluorescence. The bait-prey protein interaction causes release of aptamer from
the bait protein, leading to a restored fluorescence; (B) Aptamer is labeled
with only one dye. When bound to the much larger bait protein, the aptamer
displays slow rotational diffusion. The interaction between bait and prey
proteins displaces the aptamer. The unbound aptamer has much faster
rotational diffusion. The change in the rotation rate is reported by fluorescence
anisotropy of the dye molecule.

Neither approach requires labeling of the interacting proteins, allowing the real

interaction between the two proteins to be revealed based on their unaffected biological

activities. Both methods enabled monitoring of protein-protein interactions in real-time









and homogeneous solutions with great ease and effectiveness. While they excel in

different aspects of protein interaction study, we found that the combination of these two

methods capable of providing detailed and solid knowledge about the kinetics of the

protein-protein binding as well as mechanism and binding site information of the

interactions, which is not possible or easily obtainable with many other techniques.

Experimental Section

Materials. Dye-labeled aptamers were obtained from Integrated DNA

Technologies, Inc. (Coralville, IA). The sequences of the 15mer and 27mer thrombin-

aptamer are 5'-GGT TGG TGT GGT TGG-3', and 5'-ACC CGT GGT AGG GTA GGA

TGG GGT GGT-3' respectively. For FRET-based assays, both aptamers were dual-

labeled with 6-FAM at the 5' end and Dabcyl at the 3' end to form FQ-15Ap and FQ-

27Ap respectively. For fluorescence anisotropy assays, both aptamer sequences were

labeled with only TAMRA at the 3' end to make T-15Ap and T-27Ap respectively. A

control 15mer aptamer was labeled with only 6-FAM at the 3' end (F-15Ap). All

aptamers were purified with HPLC.

Human a-thrombin (M.W.-36.7 kDa), human antithrombin III (AT3) (M.W.-58

kDa) and a monoclonal antibody anti-human thrombin (AHT) (M.W.-150 kDa) were

obtained from Haematologic Technologies Inc. (Essex Junction, VT). Bovine serum

albumin (BSA) (M.W.-67 kDa) and a sulfated fragment 54-65 of protein hirudin, Gly-

Asp-Phe-Glu-Glu-Ile-Pro-Glu-Glu-Tyr(SO3H)-Leu-Gln (HirF) (M.W.-1.5 kDa), were

from Sigma-Aldrich, Inc. (St. Louis, MO).

All tests were performed in a 20 mM Tris-HCl buffer with a pH of 7.6 that

contained 50 mM NaCl and 5% (V/V) glycerol. All reagents for the buffer were obtained

from Fisher Scientific Company L.L.C. (Pittsburgh, PA).









Fluorescence measurements. Fluorescence measurements were done on a

Fluorolog-3 spectrofluorometer (Jobin Yvon Inc., Edison, NJ). For FRET-based assays,

fluorescence of 6-FAM was monitored with excitation wavelength of 490 nm and

emission wavelength of 520 nm. For anisotropy-based experiments, Fluorescence of

TAMRA was monitored with 555 nm as excitation and 580 nm as emission wavelength.

Slit widths were varied to yield best signals. All measurements were carried out in a 100

[L cuvette. In the aptamer/thrombin binding experiments, very small volume of a-

thrombin with high concentration was added to an aptamer solution in the cuvvette to

make molar ratio of aptamer and thrombin 1:1, and the fluorescence signals were

recorded before and after the addition. For protein-protein binding reaction,

aptamer/thrombin mixture at 1:1 molar ratio was placed in the cuvette, very small volume

of the second protein solution with high concentration was added to the mixture to make

a desired prey protein concentration. All dilution effects caused by addition of samples to

the original solutions were corrected during data analysis.

Anisotropy measurements. Anisotropy measurements were based on the

following equation:

Iw G IVm
Anisotropy r = -Gv
Iw + 2G IVH

where the subscripts V and H refer to the orientation (vertical or horizontal) of the

polarizers for the intensity measurements, with the first subscript indicating the position

of the excitation polarizer and the second for the emission polarizer. G is the G-factor of

the spectrofluorometer, which is calculated as G = IHV/HH by the instrument. G-factor is

dependent on the emission wavelength. For a certain dye, the G-factor would be

measured and used throughout the experiments that used the same dye. Then the









spectrofluorometer would keep the excitation polarizer vertical and rotate the emission

polarizer from vertical to horizontal position to measure the intensities for anisotropy

calculation. For TAMRA, all intensities were measured at emission wavelength of 580

nm with excitation wavelength of 555 nm. Time based measurements were carried out by

continuously monitoring the anisotropy readout. With an integration time of 0.5 second,

each anisotropy measurement would take about 4.1 seconds.

Kinetic Studies. Experiments were done in a 100uL cuvette in the

spectrofluorometer. While the detection system was running, the reaction samples were

quickly mixed together. Data were recorded from the point of mixing to when the signal

reached plateau and stabilized. The reaction was regarded completed when the signal was

at the plateau. Detections were either done using steady state anisotropy measurements

for AT3 and AHT study with T-15Ap, or steady state fluorescence measurements for

HirF study with FQ-15Ap. Study with AT3 was conducted at room temperature while

AHT and HirF experiments were done at 5 OC to enable monitoring of the otherwise too

fast reactions by our instrument. The temperatures of reactions were maintained using a

RTE-111 water bath/circulator (Neslab Instruments, Inc., Newington, NH).

Gel electrophoresis. Gel electrophoresis was performed on a Mini-Protean 3

precast gel system (Bio-Rad Laboratories, Inc., Hercules, CA). Samples loaded on a 7.5%

resolving Tris-HCl native gel (Bio-Rad Laboratories, Inc., Hercules, CA) were run at 150

V for 150 minutes. The gel was then taken out, rinsed with ultra-pure water and stained

with Coomassie blue stain reagent (Fisher Scientific Company L.L.C., Pittsburgh, PA)

for 1 hour. A digital camera was used to image the stained gel.









Results and Discussion

Human a-thrombin (a-thrombin) and its aptamers were used as a model system to

demonstrate the capability of aptamers to probe protein-protein interactions. Human a-

thrombin is a serine proteinase which has two positive-charged sites termed Exosite I and

II on the opposite sides of the protein.49 Exosite I was found to bind to fibrinogen50 and

hirudin51 while Exosite II binds to heparin. Two different aptamers have been identified

that have high affinity and selectivity for a-thrombin. The first one is a 15mer single-

stranded DNA aptamer which was reported to bind to the fibrinogen-binding site of a-

thrombin (Figure 2-2),52 namely Exosite I. The other DNA aptamer, with a 27mer

backbone length, was determined to bind to the Exosite II of a-thrombin.53 Both aptamers

were found to adopt a G-quadruplex structure when bound to a-thrombin. A 15mer

Exosite I binding aptamer (15Ap, Table 2-1) and a 27mer Exosite II binding aptamer

(27Ap, Table 2-1) with similar thrombin-binding affinity were chosen to study the

interactions of a-thrombin with other proteins.








''










Figure 2-2. 3-dimentional structure of human a-thrombin in complex with 15Ap.54 15 Ap
is shown in purple and blue at the bottom.









Table 2-1. Sequences of the fluorophore-labeled aptamers used in this work.
Oligo name Oligo sequence
FQ-15Ap 5'-(6-FAM)-GGT TGG TGT GGT TGG-(Dabcyl)-3'

T-15Ap 5'-GGT TGG TGT GGT TGG-(TAMRA)-3'

FQ-27Ap 5'-(6-FAM)-ACC CGT GGT AGG GTA GGA TGG GGT
GGT-(Dabcyl)-3'
T-27Ap 5'-ACC CGT GGT AGG GTA GGA TGG GGT GGT-
(TAMRA)-3'


FRET-Based Signaling Aptamer for Protein Binding.

We previously reported a molecular beacon aptamer for a-thrombin detection

based on the 15Ap.36 Here a slightly modified aptamer (FQ-15Ap, Table 2-1) has been

used that incorporates a 6-carboxyfluorescein (6-FAM) at the 5' end of the DNA as the

donor and a Dabcyl at the 3' end as the quencher. The quenching of 6-FAM emission is

caused by energy transfer between 6-FAM and Dabcyl in the protein-binding induced G-

quartet structure where the two labels are in close proximity. When excess a-thrombin

was added to an FQ-15Ap solution at room temperature, the fluorescence of 6-FAM

dropped about 55 percent (Figure 2-3). It is known that high metal ion concentrations,

especially the presence of K+, can promote the formation of G-quartet,55;56 which will

result in a much lower fluorescence signal change upon aptamer/a-thrombin binding.

However, using a buffer without any metal ions was found to inhibit protein-protein

interactions. By keeping a 50 mM NaCl concentration in the buffer, we were able to

sustain the protein activities and get relatively high fluorescence quenching induced by

protein binding to the aptamer. When a-thrombin was added into a control 15mer

aptamer that was labeled only with 6-FAM, no significant fluorescence change was

observed (Figure 2-3), indicating that the fluorescence decrease in the FQ-15Ap-

thrombin binding experiment was due to the binding-induced conformational change of









the aptamer rather than a direct quenching of the dye 6-FAM by a-thrombin. We did not

observe the quenching (i) under conditions where thrombin would not bind the aptamer

beacon, and (ii) with a scrambled aptamer beacon to which thrombin does not bind. This

result was consistent with reported molecular beacon aptamer study.36



1.2


1.0




S0.6


C 0.4



0.2


0.0
FQ-15Ap F-15Ap


Figure 2-3. Human a-thrombin binding induced relative fluorescence change of dual-
labeled 15mer aptamer. On the left, 6-FAM florescence intensity of 100 nM
FQ-15Ap aptamer in physiological buffer before (1, blue column) and after
(white column) the addition of 500 nM a-thrombin. One the right, a control
15mer aptamer labeled with only 6-FAM (F-15Ap) underwent the same
experiment and the relative fluorescence before (1, blue column) and after
(white column) the addition of 100 nM a-thrombin was measured.

Dual-labeled aptamer for thrombin-protein binding study.

The 1:1 molar ratio FQ-15Ap/a-thrombin solution (bait solution) was used to

identify interactions of a-thrombin with other proteins. When a second protein (prey

protein) binds to the same site of a-thrombin as the FQ-15Ap, the aptamer is expected to

be displaced and the freed aptamer will shift back to a more relaxed conformation,










resulting in restored 6-FAM fluorescence. A sulfated fragment of hirudin that contained

the C-terminal 13-residue51 (HirF) instead of hirudin was used for binding a-thrombin.

The addition of HirF to the FQ-15Ap bait solution caused a sharp fluorescence increase

(Figure 2-4), which was expected since both HirF and FQ-15Ap bound to the same site of

a-thrombin. Control experiments showed that there was no fluorescence change when

HirF was added to a FQ-15Ap in the absence of thrombin, indicating that there was no

direct interaction between the aptamer and HirF. The time course results showed that this

competitive binding reaction was fast as the aptamer departed within seconds after HirF

was added to the aptamer-thrombin complex solution.


I 0I
0 500


1000 1500
Time (s)


2000
2000


2500 3000
2500 3000


Figure 2-4. Thrombin bound to FQ-15Ap interacts with other proteins. In a solution of
mixed 100 nM FQ-15Ap and 100 nM a-thrombin, 200 nM AT3 (0), 500 nM
HirF (0) or 300 nM AHT ( .) was added at 0 sec. and fluorescence of 6-FAM
was continuously monitored.


170-

165-

160-

155-
-

150-

145-

140-

135-

130-

125-









Several other proteins were also investigated for interactions with a-thrombin using

the FQ-15Ap bait solution. The addition of an antibody, anti-human thrombin (AHT),

caused no significant change in the fluorescence of 6-FAM (Figure 2-4). While this result

indicates that AHT does not compete with the aptamer for the Exosite I of a-thrombin,

we can not exclude the possibility that AHT still binds to a-thrombin but at a different

site of a-thrombin. More experiments were done to address this issue (results are

presented later in this paper). A serine protease inhibitor antithrombin III (AT3) was also

tested in the bait solution. A slow-signal increasing trend was observed for AT3 (Figure

2-4). Addition of excess AT3 further increased the 6-FAM fluorescence, but the

fluorescence intensity never exceeded that of the FQ-15Ap solution in the absence of a-

thrombin. This result could be explained in that the binding of AT3 to a-thrombin may

have caused a conformational change in a-thrombin that rendered the binding with the

aptamer at Exosite I unstable.7 The slow reaction rate of AT3 was probably due to the

fact that its interaction with the active site of serine proteinases is a multi-step, covalent-

bond-forming process.5

Bovine serum albumin (BSA) was used as a control protein for interaction with a-

thrombin. No fluorescence change was observed for BSA. Another set of control

experiments were conducted by adding the prey proteins to be tested to an FQ-15Ap

buffer solution without a-thrombin. None of the proteins affected fluorescence of the

aptamer, meaning they did not interact with either the aptamer or the fluorophore.

It is also possible to quantify the amount of prey protein that is interacting with

thrombin using different level of signal change. We found that at higher thrombin to

aptamer ratio such as 2:1, it took more prey protein to cause similar quantity of signal







40


change, thus diminishing the sensitivity of this assay. For that reason, 1:1 ratio of

thrombin and aptamer was used in all our experiments.

FRET-based 27mer aptamer for thrombin-protein binding.

The sequence of the Exosite II-binding 27mer aptamer was adopted from a

previous report.53


150-

140-

130-

120-

110-

100-

90-

80-

70-


I I
1000 1500
Time (s)


2000 2500 3000


Figure 2-5. Dual-labeled 27mer aptamer for a-thrombin/protein interactions. In a solution
of mixed 100 nM FQ-27Ap and 100 nM a-thrombin, 300 nM AT3 (0), 500
nM HirF (0) or 300 nM AHT ( ) was added at 0 sec. and fluorescence of 6-
FAM was continuously monitored.

This aptamer was labeled with 6-FAM and Dabcyl similar to FQ-15Ap. With the

addition of a-thrombin, FQ-27Ap also displayed decreased 6-FAM fluorescence because

6-FAM and Dabcyl at the two ends of the aptamer were brought closer in the quadruplex

structure. The relative fluorescence decrease was found to be a little larger than that in the


0 500
0 500









FQ-15Ap experiments (Figure 2-3). Compared to the noise level, the absolute

fluorescence difference between the bound and the unbound FQ-27Ap provided adequate

sensitivity for our thrombin-protein interaction study.

Different proteins were investigated in a FQ-27Ap/a-thrombin bait solution in a

similar way as in the FQ-15Ap based assay. The results for HirF and AHT showed

slightly decreased signals (Figure 2-5), indicating no displacement of FQ-27Ap took

place. The fluorescence reduction could be caused by interactions of thrombin with those

two molecules. In contrast, antithrombin III still displayed a gradual increase in 6-FAM

fluorescence, meaning that, contrary to a previous report,5 binding between thrombin

and the serpin antithrombin III can also destabilize binding at Exosite II. Again, the slow

interaction between AT3 and thrombin caused rather gradual displacement of FQ-27Ap.

Fluorescence Anisotropy (FA) Based Aptamer Probes for Protein Interactions.

To address some of the unresolved problems in FRET experiments such as how

AT3 really binds to a-thrombin and what happens between AHT and a-thrombin, we

developed a complementary strategy based on fluorescence anisotropy. Fluorescence

anisotropy is widely used for studying the interactions of biomolecules due to its

capability of sensing changes in molecular size or molecular weight. We labeled the

thrombin aptamers with only one TAMRA dye at the 3' end to create anisotropy aptamer

probes, the 15mer T-15Ap and the 27mer T-27Ap (Table 2-1).

The T-15Ap was first investigated for its ability to probe protein interactions. When

T-15Ap/a-thrombin (1:1) solutions were mixed together, the anisotropy of T-15Ap

increased more than 30%. This bait solution was then tested with different prey proteins

(Figure 2-6). The anisotropy dropped within seconds upon addition of HirF to the bait

solution and remained almost constant after that. This result correlates well with the










result from the FRET-based experiment and may be explained as a quick displacement of

the aptamer by HirF at the Exosite I binding site of a-thrombin. The anisotropy decreased

as a result of the increased concentration of unbound aptamer which had a much lower

molecular weight than that of the aptamer-protein complex. The reaction was rapid,

indicating a simple binding between HirF and a-thrombin through non-covalent bonds.



0.19-

0.18-

0.17-

0.16 -
0
", 0.15-
o

< 0.14-

0.13- "

0.12-

0.11 I I
0 500 1000 1500 2000 2500
Time (s)



Figure 2-6. TAMRA-labeled 15Ap for a-thrombin/protein interactions based on
fluorescence anisotropy. In a solution of mixed 100 nM T-15Ap and 100 nM
a-thrombin, 200 nM AT3 (0), 500 nM HirF (i) or 300 nM AHT ( ) was
added at 0 sec. and anisotropy of TAMRA was recorded in real time.

The AT3 curve showed a different decreasing trend with time. It was rather slow

and gradual, similar to the FRET-based result. However, in the FRET assay, while it

clearly illustrated that the aptamer was displaced, it did not provide much information

about how this displacement took place. There could be several pathways that the AT3/a-

thrombin interaction might have taken. One of them is that the AT3 molecules would









quickly bind to the active site of a-thrombin, and a slow conformational change of a-

thrombin induced by AT3 binding then caused FQ-15Ap to leave Exosite I. In another

pathway, AT3 would slowly attack thrombin and such interaction would force the

aptamer to leave thrombin. The FRET-based method could not differentiate between

these two mechanisms. On the other hand, using fluorescence anisotropy, if the AT3/a-

thrombin interaction underwent the first pathway, the increased molecular weight through

the binding of AT3 to a-thrombin/aptamer complex in the first step would introduce an

initial anisotropy increase. Then, the anisotropy would slowly decrease from that point on

as the T-15Ap slowly became unbound. However, the real time anisotropy detection of

the AT3/a-thrombin interaction (Figure 2-6) demonstrated no such initial anisotropy

jump. Combined with the result from FQ-27Ap, the anisotropy experiments seemed to

better support the second pathway as the mechanism for this protein-protein interaction.

The anisotropy approach is shown here to be able to provide insight into the kinetics and

mechanisms of the targeted interactions, which will be highly useful in understanding

proteins' functions. It is our belief that site-directed aptamers enable real-time, sensitive

studies on protein-protein interaction.

It is interesting to observe that AHT caused an immediate anisotropy increase of T-

15Ap when added to the aptamer/a-thrombin bait solution (Figure 2-6). While the lack of

a decreased anisotropy correlated with the FRET-based result that showed AHT had no

effect on binding between the 15mer aptamer and a-thrombin, the anisotropy increase

suggested the presence of a binding between AHT and a-thrombin. Furthermore, this

binding happened at a different site than Exosite I, which added extra weight to the

aptamer/a-thrombin complex. The binding of AHT and a-thrombin was further









confirmed using gel electrophoresis (Figure 2-7). One advantage of the anisotropy-based

method over the FRET-based method and many other techniques might be that it can

differentiate interactions at different binding sites.







:iil



















Figure 2-7. Binding between a-thrombin and anti-human thrombin (AHT) confirmed by
gel electrophoresis on a 7.5% native Tris-HCl gel. Left lane contained 50
pmole a-thrombin. Middle lane had 32 pmole AHT. Right lane had mixture of
32 pmole AHT and 50 pmole a-thrombin.

Bait solutions containing T-27Ap and a-thrombin were also used to probe protein-

protein interactions at the Exosite II of a-thrombin (Figure 2-8). HirF caused a slightly

lower anisotropy change even though it binds to Exosite I. Considering HirF is a rather

small molecule (M.W. =~1.5 KDa), the small anisotropy decrease was likely caused by

HirF displacing T-27Ap. However, this displacement was much smaller compared to that

of T-15Ap. AT3 displayed a gradually decreasing anisotropy as it slowly displaced T-










27Ap. In contrast, AHT induced an instant anisotropy increase similar to what was found

with T-15Ap, suggesting that AHT does not affect binding at Exosite II and probably

binds to a third site of a-thrombin other than Exosite I and II.


0.21 -

0.20-

0.19-

0.18-

0.17-

0.16-

0.15-

0.14-

0.13-


2500 3000 3500


Figure 2-8. TAMRA-labeled 27Ap for a-thrombin/protein interactions based on
fluorescence anisotropy. (A) In a solution of mixed 100 nM T-27Ap and 100
nM a-thrombin, 200 nM AT3 (0), 500 nM HirF (U) or 300 nM AHT ( ) was
added at 0 sec. and anisotropy of TAMRA was recorded in real time.

Quick Evaluation of Binding Constants of Protein-Protein Interactions.

In many research areas and biological applications, it is important to not only

identify a protein-protein interaction, but also determine how strong the affinity is

between the two proteins. The binding affinity of a protein-protein interaction can be

represented by the dissociation constant (Kd) of the binding reaction. Typically, for a

newly found interaction, no matter what detection technique is used, it is often necessary


-El'.~+, L-b


0 500 1000 1500 2000
Time (s)









to build a calibration curve using various analyte concentrations. Kd of the interaction can

then be derived for the calibration curve. In a competitive assay, such as described in this

work, the interaction of aptamer and its target protein is a known system. Addition of the

prey protein may shift the equilibrium of the aptamer/bait protein binding reaction and

cause a new signal from the aptamer when a new equilibrium is reached. We found that,

based on the known aptamer/a-thrombin interaction and equilibrium conditions, it was

possible to theoretically calculate the Kd of a-thrombin/prey protein binding reaction

using the signal change occurred when the prey protein was added to the aptamer/a-

thrombin complex solution.

Assume CA molar of T-15Ap aptamer and CT molar of a-thrombin are mixed

together. When Cp molar of prey protein is added to the mixture, it will displace T-15Ap

and result in a decreased anisotropy value of rnew. rnew can be represented using the

following equation:

rA X + rAT (1 X) = ew

where rA and rAT are anisotropies of the two fluorescent species of the solution, T-15Ap

and T-15Ap/a-thrombin complex respectively, and x is fraction of the unbound T-15Ap

aptamer. Since TA and rAT are known properties of the aptamer/a-thrombin system and

rnew is the measured new anisotropy, it is easy to find out that:

rnew rAT
x=--
rA rAT
Then the concentrations of unbound and bound T-15Ap are:

[T15Ap] = CA x [T-15Ap/a-thrombin] = CA (1-x)

Because the dissociation constant of aptamer/a-thrombin reaction (Kd/AT) is already

known, then:









S Kd AT* [T -15Ap/a thrombin]
[a-thrombin] =
[T-15Ap]

Since C = [a-thrombin] + [T-15Ap/a-thrombin] + [prey/a-thrombin],

[prey/a-thrombin] = Cr [a-thrombin] [T-15Ap/a-thrombin]

Similarly, Cp = [prey/a-thrombin] + [prey protein], so

[prey protein] = Cp [prey/a-thrombin]

Finally, the dissociation constant of a-thrombin/prey protein binding reaction

(Kd/Tp) is given by the following equation:

Kd = [prey protein] [a thrombin]
Kd/TP -
[prey/a thrombin]

It is clear that theoretically, the aptamer signal change induced by competitive

binding can be easily applied to elucidate affinity of the protein-protein binding

interaction. There is no requirement to take multiple measurements using different prey

protein concentrations since aptamer/a-thrombin serves as a good reference system.

Using a simple computer program, it is possible to routinely calculate protein-protein

binding affinity using data obtained from the aptamer-based competitive assay for

protein-protein interactions. We demonstrated this capability by calculating Kd Of a-

thrombin/HirF binding reaction to be -190 nM using a reported 15mer aptamer-thrombin

Kd/A of 75 nM.59 This Kd is close to what other people have observed (150 nM).53

Despite the quick and easy evaluation of Kd using the above described method, it is

important to note that errors in sample handling and fluorescence signal measurements

might lead to a considerable amount of uncertainty in the calculated dissociation

constants. In this case, multiple measurements may be required. We also notice that there

is no information about the stoichiometry of binding and the potential of"co-operativity"









in any of the binding reactions in our estimation. This simple method for evaluating

binding constant can at least be used as a quick estimation in protein-protein interaction

studies.

Kinetics of Protein-Protein Interactions in Competitive Assays.

While the thermodynamic properties of the protein-protein interactions will

probably not be affected by the competitive binding of the aptamer, the reaction rates are

most likely still dependent on the kinetics of aptamer-protein binding. The detection of

protein-protein interactions where the aptamer is displaced consists of two major steps,

the dissociation of the aptamer and thrombin, and the association of thrombin and the

prey protein.

k._
Apt-Thr -> Apt+ Thr
ki

k2
Thr +P > Thr-P
k_2

where Apt, Thr and P are designated to aptamer, thrombin and the prey protein,

respectively. The affinities of the aptamer and the prey protein for thrombin can be

represented by their binding constants:

KApt-Thr = ki/k.1

KThr-P= k2/k-2

One situation that should be considered in the aptamer-based competitive assay is

that even though the two binding constants could be very close, there could still be large

differences between k.1 and k-2, and kl and k2. In the cases where k.<< k2, namely the

"off" rate of aptamer is vastly smaller than that of the prey protein, it may take an










enormously long time to detect a signal change even though thermodynamically the

protein should be able to displace the aptamer from thrombin.




1.0 _-___


0.8-
o

S0.6
4--
0
o 0.4

E
o 0.2


0.0 -

0 20 40 60 80 100
Time (s)



Figure 2-9. Effect of the order of incubation with thrombin on thrombin-protein
interaction. 500 nM HirF was first incubated with 100 nM thrombin (0) and
then 100 nM FQ-15Ap was added at time 0 to replace HirF. In another case
(A), 100 nM FQ-15Ap was incubated with 100 nM thrombin first and 500
nM HirF was added later at time 0. Completion of reactions was monitored
using fluorescence changes of 6-FAM.

In order to evaluate the possibilities of such false negatives in our assays, it is

necessary to compare the reaction rates of aptamer and prey proteins with thrombin. One

direct way to do the comparison is to change the order aptamer and the prey protein are

incubated with thrombin. In one experiment, the prey protein was incubated with

thrombin first and then the aptamer was used to displace the prey protein from the

thrombin/protein complex. In another experiment, the order of adding aptamer and prey

protein to thrombin was reversed. By comparing kinetic profiles of these two









experiments, it is possible to find out if aptamer binding makes interaction between

thrombin and prey protein difficult to take place. HirF was tested along with FQ-15Ap in

this way because they compete for the same Excsite I on thrombin. It was found that

aptamer replacing HirF was even slower than HirF replacing aptamer (Figure 2-9),

meaning "off' rate of aptamer would not be so slow as to affect thrombin/HirF

interaction.

Another indirect method was also used to study the effects of aptamer on

thrombin/protein interactions. If aptamer binding to and dissociation from thrombin was a

much slower process than thrombin/protein interaction, one would expect that changing

prey protein concentration would not change the observed rates of thrombin/protein

binding in the aptamer-based assay since the prey protein was not in the rate-limiting step

of the two steps mentioned earlier. On the other hand, changing aptamer concentration

should greatly affect the observed rates since aptamer was in the rate-limiting step.

Experiments were conducted to study the thrombin/AT3 interaction. Different

concentrations of AT3 were added to thrombin/T-15Ap incubation solution and the

anisotropy of the aptamer was monitored as AT3 would displace T-15Ap. The results

show a clear dependence of thrombin/AT3 kinetics on AT3 concentration (Figure 2-

10A), which contradicts with the assumption that aptamer binding was the rate limiting

step. In another study with AHT, T-15Ap concentration was varied to see if the aptamer

had any effects on the observed rates of thrombin/AHT reaction even though T-15Ap and

AHT were found to bind to different parts of thrombin (Figure 2-10B). The results show

no noticeable change in the kinetics, indicating aptamer had no effects on thrombin/AHT

binding either.















A 1.1-

1.0-

0.9-

0.8-

O 0.7-

2 0.6-

S0.5-

o 0.4-

0.3-
E
o 0.2-
0
0.1-

0.0-

-0.1-





B
1.2-


1.0-
c-
0
O 0.8-
C5

0.6-
O
0
0.4-
0.
E
0 0.2-


0.0


0 500 1000 1500 2000

Time (s)


2500 3000 3500


.. 0. o .
.0 o .



* *.
* i


100 200

Time (s)


400 500


Figure 2-10. Rate-limiting step in aptamer-thrombin-protein interactions. (A) 100nM T-
15Ap and 100 nM thrombin were first incubated. Various amounts of AT3
were added at time 0: (0) 100 nM; (0) 200 nM; ( ) 300 nM. (B) Various
concentrations of T-15Ap were incubated with 100 nM thrombin: (I) 50 nM;
(0) 100 nM; ( ) 200 nM. Then 300 nM of AHT was added at time 0.
Completion of reactions was monitored using anisotropy changes.


/. ,.,












The protein-protein interactions we studied here have shown to be not affected by

the aptamer binding to its target. Compared to two interacting proteins, aptamers are

usually much smaller than their target proteins and tend to bind to the targets only

through non-covalent forces. They are also less likely to cause induced conformational

changes of the target proteins than in protein-protein interactions. Thus it is not surprising

that aptamers interacting with their targets would not kinetically interfere with protein-

protein detection in our competitive assay.

Conclusions

Aptamers have great potential in molecular recognition due to their excellent

structural stability and exceptional flexibility with various intra-molecular modifications.

While most previous work has been focused on using aptamers as probes for direct

detection of their target molecules, this work has opened novel applications for aptamers

in areas where understanding of the interactions between known proteins and other

molecules bears great significance. Many aptamers can be easily labeled with a

fluorophore and a quencher to form intra-molecular FRET. Folded conformations of

many aptamers have shown to be stabilized by binding to their target molecules.33;39;40;53

This makes possible a fluorescence signal change of the fluorophore induced by FRET

when the aptamer binds to its target. In some cases, target-binding induced FRET can

cause up to 90% fluorescence quenching, making FRET-based detection very sensitive.

In an alternative approach, FRET can be formed within an aptamer even if the aptamer

lacks the necessary conformational changes accompanying the binding to the target

molecules.35

The other detection strategy used in this work, the fluorescence anisotropy, relies

on the relatively smaller sizes of aptamers compared to proteins. Aptamers have shown to









be suitable for fluorescence anisotropy based protein studies and detections.42;60 It is

demonstrated here that aptamer based anisotropy probes can provide sufficient signal

change for protein-protein interaction study.

Both above-mentioned methods allow real time monitoring of protein-protein

interactions without any modifications to the interacting proteins. In a recent report,

aptamers were used for a similar purpose, RNA ribozymes were required to produce a

fluorescence signal change.48 Aptamers completed with another molecule may have

affected binding affinity toward their targets. As a result, more target protein is necessary

to cause enough initial signal change for protein-protein binding study. In fact, a

thrombin concentration of 20 times of the ribozyme/aptamer complex was used,48

compared to the 1:1 molar ratio of a-thrombin and aptamer used in this work. With

excess bait protein in a competitive assay, a considerable amount of prey protein would

be necessary to significantly affect the signal of the aptamer, which may easily lead to

false negatives. Using assays directly based on aptamers could preserve aptamers'

affinity to the proteins and monitor protein-protein interactions with high sensitivity.

Our results have shown that two detection methods complement each other. The

FRET-based assay relies on direct measurements of sample fluorescence, which makes it

highly sensitive and selective. It can also be easily adapted for binding site-specified high

throughput protein interaction screening in an array format. On the other hand,

fluorescence anisotropy has shown to offer a large amount of information about protein-

protein binding that is not readily available using many other techniques including FRET.

The aptamer-based competitive assay should be useful for finding protein-binding

targets with comparable affinities in a large array of compounds. When detection of









weaker protein-protein binding is desired, it is possible to lower the aptamer's affinity

towards the target protein by adding, removing or changing bases of the aptamer that are

not directly involved in the aptamer/protein binding. This flexibility or tenability makes

aptamers more appealing for competitive assays than antibodies.

The aptamer competitive assay should also hold the potential for studying

interactions between proteins and other molecules such as small organic molecules,

DNAs and RNAs. With aptamers being rapidly developed for a growing number of

proteins, it is possible to build a large array of aptamers in protein-drug candidate

interactions for large scale drug discovery, or in whole cell protein-protein interactions

for disease diagnosis and functional proteomics.














CHAPTER 3
MOLECULAR APTAMERS-BASED AFFINITY CAPILLARY ELECTROPHORESIS
FOR PROTEIN-PROTEIN INTERACTIONS

We have demonstrated the capability of oligonucleotide aptamers serving as protein

probes for real-time protein interaction study. However, except its competence in

homogeneous solution systems, aptamer couple with fluorescence detection can also

excel in protein analyses using separation-based techniques, particularly capillary

electrophoresis (CE).

Electrophoresis has been an established method for protein analyses. Traditionally,

the mobility shift gel electrophoresis has been used to study DNA-protein interactions,

until recent, development of the capillary electrophoresis mobility shift assay and affinity

capillary electrophoresis (ACE).61-63 ACE refers to a collection of techniques in which

high-affinity binding probe is used in conjunction with capillary electrophoresis (CE)

separation to determine analytes. When coupled with fluorescent labels and laser-induced

fluorescence detection (LIF), this immunoassay technique has shown advantages such as

high mass sensitivity, rapid separations, simultaneous determination of multiple analytes,

and compatibility with automation.64-66 Antibody-antigen (Ab-Ag) interaction is widely

employed in ACE. Both competitive and non-competitive immunoassays canbe used to

detect the antigen.67-69 In a competitive immunoassay, antigen (Ag) is mixed with a

fluorophore-labeled antigen (Ag*) and a limiting concentration of antibody (Ab). CE-LIF

analysis yields two zones that correspond to the Ag* and the Ab-Ag* complex. The

relative intensities of the Ag* and Ab-Ag* allow the quantification of the original









concentration of Ag. In the non-competitive assay, a dye-labeled Ab (Ab*) is mixed with

Ag. The Ag can be quantified by determining Ag-Ab* after CE separation. Though

simple in design, antibody-based ACE has a few limitations. First, it is not always easy to

uniformly label either Ab or Ag with fluorophores. Second, for protein analytes,

antibodies may not be the ideal ligand in electrophoresis. Compared to many proteins,

antibodies are much larger in size, making it difficult to separate Ab* from Ab*-Ag

complex due to a small electrophoretic mobility difference between Ab* and Ab*-Ag.

Furthermore, charge density on the antibody molecule may very well be similar to that on

the protein antigen, which does not help the separation in CE.

One way to overcome these disadvantages of the antibody-based ACE is to develop

alternative ligands that not only possess the high binding strength to the analyte, but also

are easy to label and separate. Aptamers have been successfully used in capillary

electrochromatography and affinity chromatography.70;71 Aptamers, especially DNA

aptamers possess several advantages in ACE assays.72-74 Size of aptamers is considerably

smaller than antibodies, and often smaller than the protein analyte. The small size ensures

a better separation between the aptamer and the aptamer-protein complex in CE. In

addition, aptamers have predictable behavior in electrophoresis as a result of their

uniform charge-to-size ratios. Aptamers hold negative charges in a wide range of pH

owing to the negative phosphate backbone of nucleic acids. This makes it possible to fine

tune the pH of the running buffer to render the protein target neutral or positive for even

better separation of the aptamer and complex. The advantages of DNA aptamers that

apply to the homogeneous assays are also applicable here in the ACE, such as

inexpensive syntheses, easy labeling, and chemical stability for long-time storage.









Kennedy and coauthors have successfully applied DNA aptamers to the

quantitative analysis of two proteins (IgE and thrombin) in CE.72 In their experiments, the

fluorophore labeled aptamer and protein mixtures were injected and separated by CE.

The peak areas of free and protein-bound aptamer were used for the quantification of the

proteins. In this way, the detection limits of IgE and thrombin were 46 pM and 40 nM,

respectively. The authors pointed out that the binding constant between aptamer and

thrombin was significantly weaker than that of aptamer and IgE, resulting in the higher

detection limit of thrombin relative to that obtained for IgE. It was presumably believed

that lower binding affinity caused a significant loss of the complex of protein-aptamer by

dissociation during the electrophoresis process. Unstable aptamer-protein complexes may

completely dissociate during the separation, leading to a very broad peak or no peak

corresponding to the complex, which in turn makes it difficult to use aptamers for the

quantitative analyses of proteins. In a more recent work, they further investigated the

electrophoresis conditions required to successfully detect aptamer-ligand complexes.74 In

their report, the tris(hydroxyamino)-methane-glycine-potassium (TGK) buffer at pH 8.4,

minimal column length and a high electric field were required for the successful detection

of aptamer-protein complexes. They concluded that these results showed potential for

aptamer based ACE. In another study, Krylov and coauthors proposed a new method that

allows for the use of low-affinity aptamers as affinity probes in the quantitative analyses

of proteins.73 The method is based on the nonequilibrium capillary electrophoresis of the

equilibrium mixture (NECEEM), which allows for the accurate quantitative analysis of

proteins even when the aptamer-protein complexes may completely decay during the

separation.75









In this work, we have investigated the effects of an electrophoresis buffer

containing different metal ions on the conformation of the 15mer thrombin-binding

aptamer. Furthermore, we found that adding an appropriate concentration of

poly(ethylene glycol) (PEG) to the aptamer and thrombin mixture might stabilizing the

complex of aptamer and thrombin. Finally, we further studied the protein-protein

interactions of thrombin with a few anti-thrombin proteins using the fluorophore-labeled

aptamers. These studies should provide useful information on using molecular aptamers

for protein-protein interactions in ACE.

Experimental Section

Chemicals and Buffers

The 6-carboxyfluorescein (6-FAM) was labeled at the 5' end of aptamer (5'-(6-

FAM)-GGT TGG TGT GGT TGG-3') obtained from Integrated DNA Technologies, Inc.

(Coralville, IA). Thrombin (M.W. 36.7 kDa), human anti-thrombin III (AT III, M.W.

58 kDa) and a monoclonal antibody anti-human thrombin (AHT, M.W.-150 kDa) were

obtained from Haematologic Technologies Inc. (Essex Junction, VT). A sulfated hirudin

fragment 54-65, Gly-Asp-Phe-Glu-Glu-Ile-Pro-Glu-Glu-Tyr(SO3H)-Leu-Gln (HirF,

M.W. 1.5 kDa), fluorescein, PEG (M.W. 8 kDa) and poly(N-vinyl-2-pyrrolidone) (PVP,

1.3 MDa) were from Sigma-Aldrich, Inc. (St. Louis, MO). The electrophoresis buffer

consisted of 25 mM tris(hydroxy-amino)methane (Tris), 192 mM glycine and 0-10 mM

KC1, LiC1, MgC12, and BaCl2 at pH 8.4 was used for separating aptamer. The

electrophoresis buffer consisted of 10 mM Tris-HCl pH 8.4 and 15 mM KC1 was for

quantifying thrombin and studying protein-protein interactions. All solutions were made

in the electrophoresis buffer including the stock solutions of protein, aptamer, PEG and








fluorescein (internal standard). All reagents for the buffers were obtained from Fisher
Scientific Company L.L.C. (Pittsburgh, PA).
Apparatus
The basic design of the separation system has been previously described (see
Figure 3-1).76


PMT


Pinhole -T-


Objective



Filter c


Laser Focus lens


( 0


High voltage
power supply


Buffer reservoir

Figure 3-1. Schematics of the capillary electrophoresis setup.









Briefly, a high-voltage power supply (Gamma High Voltage Research Inc.,

Ormond Beach, FL) was used to drive electrophoresis. The entire detection system was

enclosed in a black box with an HV interlock. The high-voltage end of the separation

system was put in a laboratory-made plexiglass box for safety. A 2.5-mW Ar ion laser

with 488 nm output (Spectra Physics, Mountain View, CA) was used for excitation. The

emission was collected with a 20 X objective (numeric aperture = 0.25). One 520-nm

interference filter was used to block scattered light before the emitted light reached the

photomultiplier tube (Hamamatsu R928, Hamamatsu Photonics K.K., Hamamatsu,

Japan). The amplified currents were transferred directly through a 50-kQ resistor to a 24-

bit A/D interface at 10 Hz (AT-MIO-16, National Instruments, Austin, TX) and stored in

a personal computer. Capillaries (Polymicro Technologies, Phoenix, AZ) 50 |tm i.d. and

365 |tm o.d. were used for electrophoresis separations with or without PVP coating.

Separation of Aptamer

Sample prepared in a separation buffer and consisted of 200 nM aptamer and 10

nM fluorescein (internal standard). Samples were injected into the capillary (total length,

35 cm; effective length, 10 cm) hydrodynamically (Ah = 5 cm) for 10 s. Separation buffer

were constituted of 25 mM Tris, 192 mM glycine and 0 -10 mM KC1 at pH 8.4. The

electrophoresis separation was carried out with an electric field of 285 V/cm.

Aptamer-Based ACE

In thrombin quantification, the aptamer was mixed with thrombin and fluorescein

(internal standard) in the electrophoresis separation buffer (10 mM Tris-HCl pH 8.4 and

15 mM KC1) and incubated for 60 min at room temperature. The final concentrations of

aptamer, fluorescein and thrombin were 200 nM, 10 nM and 0-1.0 lM, respectively. The









resulting samples injected into the capillary (total length, 50 cm; effective length, 25 cm)

hydrodynamically (Ah = 10 cm) for 10 s. The electrophoresis separation was carried out

with an electric field of 350 V/cm. For the quantification of anti-thrombin proteins (AT

III, HirF and AHT), aptamers were mixed with thrombin in the electrophoresis separation

buffer (10 mM Tris-HCl pH 8.4 and 15 mM KC1) and incubated for 60 min at room

temperature. The desired concentrations of anti-thrombin proteins were mixed with

aptamer-thrombin complex solutions and incubated for another 60 min. Fluorescein was

added to the resulting samples as an internal standard to 10 nM. The final concentrations

of aptamer, thrombin and anti-thrombin proteins were 200 nM, 200 nM and 0-10.0 aM,

respectively. The samples were injected into the capillary (total length, 40 cm; effective

length, 25 cm) hydrodynamically (Ah = 10 cm) for 10 s. The electrophoresis separation

was carried out with an electric field of 500 V/cm. At the end of each run, the capillary

was rinsed with 0.5 N NaOH for 10 min to remove the protein adsorbed on the capillary.

PEG-Assisted Aptamer ACE

To study the effect of PEG, aptamers were mixed with thrombin in 10 mM Tris-

HCl pH 8.4, 15 mM KC1 and 0-10% PEG and incubated for 60 min at room temperature.

The final concentrations of aptamer and thrombin were 200 nM. The electrophoresis

buffer consisted of 10 mM Tris-HCl pH 8.4 and 15 mM KC1. The resulting samples were

injected into the PVP coated capillary (total length, 15 cm; effective length, 5 cm)

hydrodynamically (Ah = 1.5 cm) for 20 s. The electrophoresis separation was carried out

with an electric field of 666 V/cm. Similarly, to quantify thrombin and study the

interactions of thrombin and anti-thrombin proteins (AT III, AHT and HirF), the desired

concentrations of thrombin and anti-thrombin proteins were mixed with aptamer and









aptamer-thrombin complex solutions, respectively. The resulting samples were injected

and separated in a PVP coated capillary. The electrophoresis buffer consisted of 10 mM

Tris-HCl pH 8.4, 15 mM KC1 and the separation was carried out with an electric field of

666 V/cm. At the end of each run, the capillary was rinsed with 5% PVP for 10 min.

Results and Discussions

Conformation of aptamer

It has been reported that a 15-mer thrombin binding aptamer adopts an

intramolecular G-quadruplex structure (Figure 1-6) in the presence of K+.7780 Its affinity

for thrombin has been associated with the inhibition of thrombin-catalyzed fibrin clot

formation.81;82 Studies with circular dichroism, temperature-dependent UV spectroscopy,

differential scanning calorimetry, isothermal titration calorimetry, capillary

electrophoresis, NMR, and mass spectrometry have revealed intramolecular G-

quadruplex structures of the 15-mer aptamer in the presence of various metal ions.77-80;83-

85 The G-quadruplex aptamer (G-Apt) is stable both kinetically and thermodynamically

because of the tight association of cations with quinine residues.86-88

The stability of the G-Apt is a very important factor in the detection of thrombin in

ACE because thrombin will only bind to the G- quadruplex form of the aptamer.37;82 Here,

we evaluated the stability of the G-Apt in the presence of several metal ions (K+, Li+,

Ba2+ and Mg2+). Figure 3-2 shows the electropherograms for the aptamer with increasing

concentrations of KC1.

The aptamer was separated into two peaks when we first used a K+-containing

running buffer. That was when we started to think that the two conformations of the

aptamer might be able to be separated in CE even though a few related reports did not

mention that.72;74






63


A. KC 0 mM 8D. 1.0 mM



4



0 0 '
50 100 150 200 50 100 150 200

8 8
r B. 0.25 mM E. 5.0 mM






8D 8



C. 0.5 mM F. 10.0 mM
04 4
0 0 _






50 100 150 200 50 100 150 200
S8
C. 0.5 mM F. 10.0 mM



4 i4



S..----.... ----- .
50 100 150 200 50 100 150 200

Time (sec)

Figure 3-2. Capillary electrophoresis traces of thrombin binding aptamer in the presence
of 25 mM Tris, 192 mM glycine and various concentrations of KC1.
Separation buffer was composed of 25 mM Tris, 192 mM glycine and 0 mM
(A), 0.25 mM (B), 0.5 mM (C), 1.0 mM (D), 5.0 mM (E), 10.0 mM (F) KC1 at
pH 8.4. Samples were prepared in a separation buffer and contained a final
concentration of 200 nM aptamer and 10 nM fluorescein (internal standard).
The samples were injected into the capillary (total length, 35 cm; effective
length, 10 cm) hydrodynamically (Ah = 5 cm) for 10 s and an electric field of
285 V/cm was applied to drive the separation.

To confirm this hypothesis, running buffers without K+ or Na+ were tested. It can

be seen in Figure 3-2 that the aptamer came out a single peak at the time where the first









peak showed up in the K -containing buffers. Since G-quadruplex structure is not stable

without metal ions, the single peak was believed to be primarily the linear form of the

aptamer. We suspected that there might have been an unstable G-Apt based on the slight

tailing of the single aptamer peak. The first and second peak except the internal standard

peak, in Figure 3-2 B to F, corresponded to the linear aptamer (L-Apt) and the G-Apt,

respectively. The peak area ratio of the G-Apt to L-Apt form increased with the increased

concentration of K+. This result agreed with other reports that the stability of the G-Apt

was dependent upon the concentration of K+.78;79;83 Furthermore, the increase in the K

concentration was found to improve the resolution between the L-Apt and G-Apt peak.

Figure 3-3 shows that other metal ions, such as Li+, Mg2 and Ba2 were not

capable of stabilizing the G-Apt or effectively separating the G-Apt from the L-Apt. It

was reported that cations with an ionic radius in the range of 1.3-1.5 A fit well within the

two G-quartets of the complex.79 Our results were in agreement with the reports that K+,

Rb+, NH+, Sr2+, and Ba2+ are able to form stable cation-Apt complexes, while Li+, Na+,

Cs+, Mg2+, and Ca2+ only form weak complexes. Among the four metal ions (K+, Li+,

Mg2+ and Ba2+) chosen in our experiments, the aptamer displayed a stable G-Apt peak

with K+ or Ba2+ present. As shown in Figure 3-3, as little as 2.5 mM Ba2+ could

effectively stabilize the peak of the G-Apt. However, similar to Mg2+, Ba2+ associates

with high affinity to the phosphate backbone of oligonucleotides and decreases the

mobility of the aptamer. This was demonstrated by the change in the aptamer peak

position relative to the fluorescein peak. On the other hand, the decrease of the mobility

of fluorescein indicated a severely reduced electroosmotic flow (EOF), which led to peak

broadening, increased separation time and decreased resolution. For this reason, Ba2+ was









certainly not suitable for the study of protein-DNA and protein-protein interactions in

ACE. Instead, KC1 was chosen to be added to sample matrix and electrophoresis buffer in

further experiments.


100-
100


0
100


200 300 400 500 600


200 300 400 500 600


Time (sec)

Figure 3-3. Capillary electrophoresis traces of thrombin binding aptamer in the presence
of 25 mM Tris, 192 mM glycine and various other metal ions. Separation
buffer were composed of 25 mM Tris, 192 mM glycine, and 10 mM LiCl (A),
2.5 mM MgC12 (B), 2.5 mM BaC12 (C) at pH 8.4. Samples prepared in a
separation buffer and contained a final concentration of 200 nM aptamer and
10 nM fluorescein (internal standard). Other experimental conditions same as
in Figure 3-2.









Quantification of Thrombin

As mentioned previously, literature reports had pointed out that the unstable

aptamer-protein complex would undergo dissociation in the electrophoresis process.7274

This caused broadening or disappearance of the complex peak unless hydrodynamic flow

was used to force the complex through the cappillary.72 Although the flow-assisted

method allows quantification of the aptamer-protein complex, it is inconvenient and may

eliminate some of the advantages of CE. A relatively long capillary (effective length 25

cm) was used in our experiments. The advantage of a long capillary was that there was

less interference with the peak of the G-Apt by the aptamer-thrombin peak (Apt*Thrmb)

because the complex of aptamer-thrombin would almost completely decay in a long

capillary. Therefore, we were able to focus on the peak of G-Apt for quantification.

Figure 3-4A and B compare the electropherograms obtained for 200 nM aptamer

without and with 200 nM thrombin in the sample. Fluorescein was used as an internal

standard (IS) to correct variations in the injection volume. The electrophoresis buffer was

further optimized from before (25 mM Tris, 192 mM glycine and 10 mM KC1 at pH 8.4,

Figure 3-2F) to 10 mM Tris-HCl added 15 mM KC1 at pH 8.4. In Figure 3-4A, the peak

area ratio (G-Apt/L-Apt) was calculated to be 2.97, which was significantly higher than

that in Figure 3-2F (1.09). The electropherogram with thrombin (Figure 3-4B) shows a

large decrease in the free G-Apt peak which can be attributed to the binding of the G-Apt

to the thrombin. A calibration curve was constructed (Figure 3-4C) based on the peak

area of the free G-Apt as a function of thrombin concentration. The curve is linear up to

200 nM thrombin and a 1:1 binding ratio between the aptamer and thrombin is clearly

demonstrated. The limit of detection (LOD), which was calculated as the concentration

yielding a signal change 3 times the peak to peak noise, was 9.8 nM, and was lower than






67


previous reported 40 nM.72 We suspect that the main reason is that the quantification in

the literature did not differentiate between G-Apt and L-Apt, while we focused on the

peak area change of the G-Apt, which might have resulted in a higher sensitivity.

8A.






0.
S 0 100 200 300 400 500
8



0
4




0 1() 200 300 400 500
Time (sec)
8
C.



4 4


200 400 6(0 800
Thrombin i' nM)


1000 1200


Figure 3-4. Detection of thrombin using aptamer-based ACE. Electropherograms
obtained for 200 nM aptamer with 0 (A) and 200 nM thrombin (B).
Calibration curve constructed using samples containing 200 nM aptamer with
various concentrations of thrombin (C). The electrophoresis separation buffer
was 10 mM Tris-HCl and 15 mM KC1 at pH 8.4. Samples prepared in
separation buffer and consisted of a final concentration of 200 nM aptamer, 10
nM fluorescein (internal standard), and 0-1.0 [tM thrombin. The samples were
injected into the capillary (total length, 50 cm; effective length, 25 cm)
hydrodynamically (Ah = 10 cm) for 10 s and an electric field of 350 V/cm was
applied to drive the separation. Peak areas were corrected for variations in
injection volume by dividing by the area of the internal standard peak.









Determination of Dissociation Constant (Kd)

The value of Kd was determined under the conditions of the experiments. The

equilibrium concentration of free G-Apt is proportional to the area of the G-Apt peak

(Age).73

[G-Apt]eq = cAgeq (1)

where c is a constant. The equilibrium concentration of the complex (G-

Apt*Thrmb) is equal to:

[G-Apt*Thrmb]eq = [G-Apt]o [G-Apt]eq = c(Ag Ageq) (2)

where Ag is the peak area G-Apt without thrombin. The ratio R of the two

equilibrium fractions:

R = [G-Apt]eq / [G-Apt*Thrmb]eq = Ageq / (Ag Ageq) (3)

The knowledge of ratio R is sufficient for the determination of Kd:

Kd = {[thrombin]o(1+R)-[G-Apt]o} / (1 + 1/R) (4)

where [thrombin]o and [G-Apt]o are the initial concentrations of thrombin and G-

Apt, respectively.

The fluorescence intensities of aptamer were found to be unchanged on a

fluorometer under different concentrations of KC1, indicating the conformational change

of the aptamer did not affect the fluorescence intensity of the labeled 6-

carboxyfluorescein. As a result, the [G-Apt]o could be calculated from Figure 3A by the

areas of peak L-Apt (Ato) and G-Apt (Ag)

[G-Apt]o = [Apt]o Ago / (At + Ag) (5)

Based on six experiments with different concentrations of thrombin and aptamer,

the value ofKd (G-Apt*Thrmb) was calculated using Equations 4 and 5 to be 20 nM.

This value is smaller than those obtained by Kennedy et al. (450 nM) and Krylov et al.









(240 nM). This can be explained by the fact that the binding affinity measured here is

between the G-Apt and thrombin, which is supposed to be stronger than that between

thrombin and the overall aptamer (G-Apt + L-Apt).

Competitive Assay

Thrombin has two positive-charged sites termed Exosite I and II on the opposite

sides of the protein.89 Exosite I was found to bind to fibrinogen50 and hirudin90-92 while

Exosite II binds to heparin and a monoclonal antibody93. Two different aptamers have

been identified that have high affinity and selectivity for thrombin. The first one is a 15-

mer single-stranded DNA aptamer (in this work) which was reported to bind to the

fibrinogen-binding site of a-thrombin, namely Exosite I.52 The other DNA aptamer, with

a 27-mer backbone length, was determined to bind to the Exosite II of a-thrombin.53 Both

aptamers were found to adopt a G-quartet structure when bound to a-thrombin.53;94 In the

previous chapter, we used the complex of aptamer-thrombin to probe thrombin-protein

interactions in a competitive assay where the binding of the aptamer to thrombin was

altered by a second protein that interacts with thrombin. Two signal transduction

strategies, fluorescence energy transfer and fluorescence anisotropy, have been designed

to study the interactions of thrombin with different proteins using two aptamers specific

for two binding sites on a-thrombin.95 Here, we have further demonstrated the results by

aptamer based ACE. As shown in Figure 3-5, the increasing concentration of AT III

caused an increase in the G-Apt peak area as expected. This result agreed with another

report that the binding of AT III to thrombin may cause a conformational change in

thrombin that rendered the binding with the aptamer at Exosite I unstable.57 In Figure 3-6,

the reaction between AT III and thrombin was monitored in a close-to-real-time fashion.

The reaction of thrombin and AT III was completed within 10 min, and the result agreed







70


with our previous work.95 A calibration curve was made based on the peak area of free G-

Apt vs. AT III concentration (Figure 3-7). The curve is almost linear up until 200 nM AT

III and a 1:1 binding ratio between the thrombin and AT III was displayed. Based on

these data, the LOD was calculated to be 2.1 nM.


A. AT Il 0 nM


4


0
0 150 3(









8 10 nM
C.


4
_,,AA


50 nM


4


0
30 0
Sr-


)0 o 0


"0 150 300 -0
8 20 nM 8
D. H.


4 4


0 L_
0 150 300 0

Time (sec)


150


100 nM


-L J
150 300
200 nM






150 300
400 nM






150 300


Figure 3-5. Analyses of AT III-thrombin interaction using aptamer-based ACE.
Electropherograms obtained for 200 nM aptamer with 200 nM thrombin and
varies concentrations of AT III. In electropherograms A-H, AT III
concentrations were 0, 5, 10, 20, 50, 100, 200 and 400 nM respectively.
Aptamers were mixed with thrombin and incubated for 60 min at room
temperature. The desired concentrations of AT III were mixed with aptamer-
thrombin complex solutions. The resulting samples were added fluorescein as
an internal standard to 10 nM and incubation another 60 min. Separation was
carried out at a constant electric field of 500 V/cm. Other conditions as in
Figure 3-4.






























Time (minn)


Figure 3-6. Using thrombin binding aptamer to monitor thrombin/AT III interaction. In a
solution of mixed aptamer and thrombin incubated in electrophoresis buffer
for 60 min. Then AT III were mixed with the aptamer/thrombin complex
solution and incubated for another 60 min. The final concentration of aptamer,
thrombin and AT III were 200 nM. After the resulting sample incubation for
different time, rapid inject into capillary and the separation were carried out at
a constant electric field of 500 V/cm. Other conditions the same as in Figure
3-4.

8.0


- 4,0
Ca
*L
u
0
uJ


LI'3 ----i i
0 200 400 600 800 1000 1200
AT III (nM)


Figure 3-7. Quantification of AT III-thrombin interaction. Calibration curve constructed
using samples containing 200 nM aptamer, 200 nM thrombin and various
concentrations of AT III (0-1.0 pM). Peak area of G-Apt was corrected for
variations in injection volume by dividing by the area of the internal standard
peak. Other conditions as in Figure 3-4 and Figure 3-5.









The addition of another antibody, 400 nM AHT (2 times concentration of

thrombin), caused no significant change in the peak of the G-Apt (data not shown). While

this result indicates that AHT does not compete with the aptamer for the Exosite I of

thrombin, it does not mean that AHT does not bind to thrombin at a different site. More

experiments were done to demonstrate this point, and the results are presented later in

this paper. A sulfated fragment of hirudin that contained the C-terminal 13-residue (HirF)

instead of hirudin was used for studying binding with thrombin. Although the Kd of HirF-

thrombin (150 nM) at pH 7.4 is similar to the reported Kd of aptamer-thrombin (200 nM)

and both HirF and aptamer bind to the same site (Exosite I) of thrombin, HirF caused no

significant change in the peak of the G-Apt even when the added concentration was as

high as 10 [tM (50 times concentration of thrombin). It has been reported that binding of

thrombin to hirudin (65 amino acids) and some derived hirudin fragments strongly

depends on pH.96 The optimum pH for the interaction between hirudin and thrombin was

found to be between pH 7.5 and pH 8.0. The Kd value increased at higher pH values, and

the plot of -log Kd against pH displayed an asymptotic slope of -2 in the alkaline pH

range. As a result, the sample and electrophoresis buffer at pH 8.4 in our experiments

may also cause a much higher Kd between HirF and thrombin than reported in literatures.

On the other hand, the relative lower salt concentration of buffer used in this experiment

may also have an impact on the binding affinity of HirF.

Effect of PEG on Aptamer-Thrombin Complex in CE

Unlike AT III, which can displace G-Apt from a G-Apt*Thrmb complex and cause

changes in the area of the G-Apt peak, the thrombin antibody AHT would not affect G-

Apt/thrombin binding. It is necessary to observe the peak of the G-Apt*Thrmb complex









and analyze its mobility shift to study the interaction between AHT and thrombin. To

resolve the problem of complex dissociation during CE, a short PVP coated capillary

(effective length 5 cm) was used to shorten the analysis time and prevent the protein

adsorption on the capillary. However, with a total separation time less than 50 sec, peak

broadening was still not improved (Figure 3-8A). There have been reports suggesting that

linear polymers can promote the cage effect which stabilizes protein complexes during

electrophoresis.97-99 In addition, polymers in electrophoresis buffers may aid the

separation by interacting with solutes and capillary walls to prevent adsorption.

Moreover, polymer in the region where the complexes have dissociated, may hinder

further separation of the two components, and lead to an enhanced probability for a re-

association. The dissociation step may also be slowed down by the polymer, if the

dissociation requires complex-complex interactions. Finally, polymers may cause macro-

crowding effect that increases local concentrations of analytes that cause lower level of

dissociation of complex.

Given all the background information, we tried adding PEG to the sample matrix.

As shown in Figure 3-8 B-F, addition of PEG clearly revealed the peak of the G-

Apt*Thrmb complex. The L-Apt and G-Apt can not be separated due to the fact that the

experiments were done in a very short time. However, it did not effect the quantification

of thrombin and AT III by the complex peak of G-Apt*Thrmb. The optimum

concentration of PEG was found to be 2.0%. A higher concentration of PEG would cause

peak broadening, probably because that the collisions of aptamer and protein molecules

into the polymer network may have induced distribution of the migration rates of those

molecules.










A. PEG 0%





c-



25 50 75 1f


0 25 50 75 100


0 25 50 75 100 0 25 50 75 100


Time (sec)

Figure 3-8. Effect of PEG on the stability of G-Apt*Thrmb complex. Aptamers were
mixed with thrombin in 10 mM Tris-HC1 at pH 8.4, 15 mM KC1 and 0% (A),
0.5% (B), 1.0% (C), 2.0% (D), 5.0% (E) and 10% (F) PEG, incubated for 60
min at room temperature. The final concentrations of aptamer and thrombin
were 200 nM. The electrophoresis buffer was 10 mM Tris-HC1, 15 mM KC1
at pH 8.4. The resulting samples were injected into the PVP coated capillary
(total length, 15 cm; effective length, 5 cm) hydrodynamically (Ah = 1.5 cm)
for 20 s. The electrophoresis separation was carried out with an electric field
of 666 V/cm.













A.


5
-'C
0

0 20 40 60 80
10
B.
F-
5


0
0 20 40 60 80

10 C.



5


0
0 20 40 60 h.
Time

8
D.



4



0


0 400 800
Thrombin (nM)


u E.



5


0
0 20 40 60 S,
10 F



5


0
0 20 40 60 80

10 G.



5


0
0 20 40 60 80
[sec)

8
H.



4



0


1200 0


400 800
AT III (nM)


1200


Figure 3-9. Analyses of thrombin and thrombin-AT III interaction using PEG-containing
sample matrix. Each sample contained a final concentration of 200 nM
aptamer and 0 (A), 50 (B), 200 nM (C) thrombin. In figure E-G, each sample
contained a final concentration of 200 nM aptamer, 200 nM thrombin, and 50
nM (E), 100 nM (F), 200 nM (G) AT III. Figure D and H are the calibration
curves constructed with various concentrations of thrombin and AT III
respectively. The sample matrix consisted of 10 mM Tris-HC1, 15 mM KC1
and 2% PEG at pH 8.4. The electrophoresis buffer was 10 mM Tris-HC1, 15
mM KC1 at pH 8.4. Other conditions as in Figure 3-5 and Figure 3-8.









Figures 3-8 A-C show the increased G-Apt*Thrmb to free aptamer peak area ratio

with increasing thrombin. The calibration curve in Figure 3-9 D has a linear range up to

200 nM thrombin. The LOD of thrombin is 10.9 nM. In a similar way, AT III was

quantified using the G-Apt*Thrmb based competitive assay (Figure 3-9 E-H). The

electropherograms with increasing AT III have shown a decreased peak of G-Apt*Thrmb

and an increased peak of free aptamer. These results once again confirmed the

displacement of G-Apt by AT III in CE. The LOD of AT III was estimated at 21.2 nM.

Aptamer-Based Mobility Shift Assay for Thrombin-AHT Interaction

The substrate specificity of thrombin is regulated by binding of macromolecular

substrates and effectors to Exosites I and II.92 Exosites I and II have been reported to be

linked allosterically, such that binding of a ligand to one exosite results in nearly total loss

of affinity for ligands at the alternative exosite, whereas other studies support the

independence of the interactions.92 Previous results in bare capillaries revealed that AHT

had no effect on binding between the aptamer and thrombin.

In order to study the interaction between thrombin and AHT in ACE, it might be

helpful to analyze the mobility change of G-Apt*Thrmb complex with the addition of

AHT. Using the PEG-assisted ACE, electropherograms were obtained (Figure 3-10 A-D)

and clearly showed changes in the migration time of the G-Apt*Thrmb complex as AHT

concentration was varied. We further optimize the concentration of PEG in this

experiment, and found 2% PEG was again the best. As shown in Figure 3-10 B-D, this

mobility shift can be attributed to the increased overall molecular mass of the G-

Apt*Thrmb*AHT binding complex. Control experiments revealed that no mobility shift

was observed when 200 nM aptamer was mixed with only 400 nM AHT.






77


A. AHT 0 nM


4



0
0 25 50 75 100

B8
B. 50 nM






0 25 50 75 100

0 C. 100 nM


a4




0 25 50 75 100

D. H 200 nM

41



O0-
0 25 50 75 100

Time (sec)

Figure 3-10. Binding between G-Apt*Thrmb and anti-human thrombin (AHT) confirmed
by capillary electrophoresis. In a solution of mixed aptamer and thrombin
incubated in 10 mM Tris-HCi, 15 mM KC1 and 2% PEG at pH 8.4 for 60 min.
Then AHT were mixed with the aptamer/thrombin complex solution and
incubated for another 60 min. Each sample contained a final concentration of
200 nM aptamer, 200 nM thrombin and 0 nM (A), 50 nM (B), 100 nM (C)
and 200 nM (D) AHT. Other conditions as in Figure 3-8.









Figure 3-10B (aptamer 200 nM, thrombin 200 nM and AHT 50 nM) displays an

unresolved small peak (arrow point and the migration time of that peak is equal to that of

the second peak in Figure 3-10D (aptamer 200 nM, Thrmb 200 nM and AHT 200 nM).

This result reveals that only a small fraction of G-Apt*Thrmb binding with AHT when

the concentration of AHT is lower than G-Apt*Thrmb. The highly broadened peaks in

Figure 3-10 B-C indicate the shift from the G-Apt*Thrmb complex to the G-

Apt*Thrmb*AHT complex. In addition, the dissociation constant of AHT-thrombin is not

very low at 14 nM,100 which attributed to the dissociation of G-Apt*Thrmb*AHT

complex at low AHT concentrations. Even though quantification of AHT is difficult, the

interaction between thrombin and AHT, and the fact that AHT and G-Apt bind to

different sites of thrombin have been clearly revealed using this mobility shift assay in

ACE.

Conclusions

In this work, we have demonstrated that the 15-mer thrombin-binding DNA

aptamer adopts two different forms in the presence of K or Ba2+ and only the G-

quadruplex form can bind thrombin to form a complex. Binding between aptamer and

proteins is thus highly dependent on the conformation of the molecular aptamers. The

presence of thrombin and Antithrombin III only affected the G-Aptamer peak in affinity

capillary electrophoresis. The G-Aptamer based CE analysis showed a higher binding

affinity between G-Aptamer and thrombin. As a result, a better detection limit of

thrombin could be achieved. The aptamer-based competitive affinity capillary

electrophoresis assay has been also applied to quantify Thrombin/Antithrombin III

interaction and to monitor this reaction in real time. We have also shown that a mobility

shift based affinity capillary electrophoresis assay, using poly(ethylene glycol) in the






79


sample matrix, can be used to study the interactions between thrombin and proteins that

do not displace G-Aptamer binding property with Exosite I site of the thrombin. We

believe that oligonucleotide aptamers possess advantages over other protein ligands in

affinity capillary electrophoresis, and the aptamer-based ACE assay can be an effective

alternative approach for studying protein-protein interactions and for analyzing binding

site information and binding constants.














CHAPTER 4
NUCLEASE-RESISTANCE OF TELOMERE-LIKE SINGLE-STRANDED
OLIGONUCLEOTIDES MONITORED IN LIVE CELLS BY FLUORESCENCE
ANISOTROPY IMAGING

Introduction

Fluorescence Techniques for Monitoring Intracellular Biointeractions

One of the major challenges of life science is to understand roles of the vast amount

of biomolecules in cells. An ideal way to study cell functions would be monitoring

interactions between biomolecules in live cells. Most cell imaging techniques are based

on detection of fluorescence signals generated from fluorescent tags that are linked to the

molecules of interesting. The development of fluorescent proteins, such as green

fluorescent protein (GFP) has made it easy to fluorescently tag proteins for intracellular

imaging.101-103 GFP has been used to monitor cellular gene expression102 and protein

trafficking and localization. 104;105 To detect intracellular interactions between proteins,

variants of GFP that have different fluorescence spectra are used to form a fluorophore

pair for fluorescence resonance energy transfer (FRET).106 The interaction between

proteins causes changes in efficiency of energy transfer and thus in the ratio of acceptor

to donor fluorescence signals.

Although FRET-based techniques are powerful tools in imaging of cellular bio-

interactions, they have certain limitations. Compared to assays using a fluorophore and

quencher pair, FRET based on GFPs tends to have higher fluorescence background, thus

makes small signal changes caused by low level of target less appreciable. To overcome

this problem, careful selection or design of the GFP pair needs to be done to obtain low









acceptor background. In addition, photo-bleaching of the two fluorophore at different

rates may cause false change in the ratio of acceptor to donor emissions. Lastly, for

protein-protein interaction study, often the two interacting proteins need to be fused with

GFP and both host protein and GFP should remain functional. Thus, GFP-based FRET

assay should serve better for biological systems where two interacting proteins are

molecularly well characterized.107

As a complementary technique to GFP-based FRET assays, we think that

fluorescence imaging based on fluorescence anisotropy (FA) may hold great potential for

intracellular bio-interaction study. The fluorescence anisotropy of a fluorophore reflects

the molecule's ability to rotate in its micro-environments. FA is dependent on things that

affect the rotational diffusional movements of the fluorophore such as the size and mass

of the molecule the fluorophore is attached to, viscosity of the solution and the

fluorescence lifetime of the fluorophore. Since most biological events inside cells induce

changes in molecular weight of involved molecules, e.g. binding of two proteins results

in a heavier complex, it is theoretically possible to label one molecule with a fluorophore

and monitor its interaction using anisotropy. Traditionally, measurements of anisotropy

have been done mainly in homogeneous solutions where no localized anisotropy

information is available. By combining florescence imaging and anisotropy

measurements, bio-interactions in heterogeneous samples, such as live cells, can be

monitored. Fluorescence anisotropy imaging has been reported in study of cells108 and

single molecules109 based on conventional fluorescence microscope. Here we describe an

anisotropy imaging system built upon on a confocal microscope. Besides commonly

known advantages of confocal microscopes over conventional ones, such as better









resolution and 3-D imaging capability, the multi-channel image acquisition capability of

a confocal microscope enables simultaneous acquisition of two images representing two

perpendicular polarization states of the fluorescence emission. In addition, the pixel-to-

pixel scanning scheme of confocal makes alignment of the two polarization images much

less of an issue for future anisotropy image calculation. During the development of our

setup, a few groups also chose confocal microscopes to build their anisotropy imaging

systems on,110-112 probably due to the same reason.

We used our anisotropy imaging setup to monitor digestion of single-stranded

DNAs by nucleases inside live cells. Our interest lies in finding out how compositions

and structures of DNAs affect their ability of resisting cellular nuclease degradation.

Telomere and Its Presence in Live Cells

One type of DNA sequence that belongs to something called telomeres is

particularly interesting in our study of nuclease-DNA interaction. Telomeres are the end

of eukaryotic chromosomes. The length of the telomeres varies greatly between different

species, from 20bp to 150kbp per telomere.113 They are composed of simple and highly

repetitive DNA sequences, with one of the two telomeric DNA strands a GT-rich

sequence. The GT-rich repeats are not the same for different species, for example, human

and mouse have a TTAGGG repeating unit, while it is TTGGGG for Tetrahymena.114

Telomeres have been found to carry important biological functions. One of them is

believed to be the protection of chromosomes. During DNA replication, short RNA

primers are employed to initiate DNA synthesis. Removal of the terminal primers at the

end of the replication always leaves a small region of single-stranded DNA (ssDNA) that

is not replicated.115 It has been shown that length of telomeric DNA in human fibroblasts

shortened during cell aging in vitro.116 Had it not been the repetitive telomere sequence,









the genome DNA would have been damaged from the very first round of replication. On

the other hand, normal cells can only replicate certain times until the telomere end is

completely lost. In this case, the telomeric sequence ensures that cells can not be

immortal. Compared to normal cells, cancer cells can replicate indefinitely, thus grow out

of control. The reason behind this is believed to be an unusual DNA polymerase called

telomerase.117 In one report, activity of the telomerase enzyme was detected in most

human tumor samples and in none of the normal tissues.118 In another report, introduction

of telomerase-encoding genes into telomerase-negative cells evidently extended their

normal life-span.119 This clear relationship between telomere, telomerase, and cancers has

inspired great interest in this line of research.

One of the early telomere papers showed that the G-strand of the telomeric DNA

was longer than the C-strand in Oxytricha, creating a G-rich 3'-end overhang.120 The size

of the overhang was found to be about 50-100 nucleotides in mouse and human

telomeres. To understand why the single G-strand can be maintained without inducing

DNA-damage response such as degradation by nucleases, different models have been

proposed. The classical view is that the G-strand is protected by proteins that bind tightly

to single-stranded DNA.114 A new model found that the G-rich single strand can tuck

back into the double strand region of the telomere to form a closed loop structure called

T-loop, which avoids exposure of the DNA end to cellular enzymes.121

The T-loop model explains very well why the termini of telomeres are stable in cell

nuclei. However, one may ask why the repeat unit of telomeric DNA is always G-rich

instead of any random sequence that can also form the T-loop. It is well known that G-

rich telomeric DNAs in many species form parallel four-stranded structure called G-









quadruplex by bonding guanines to each other.122;123 In this work, we tried to study

whether the special structure of the telomere-like single-stranded G-rich oligonucleotides

contributes to the stability of telomeric overhang in cell nuclei.

Fluorescence anisotropy was chosen to study the digestion process over other

approaches such as FRET. A FRET design may be able to generate a fluorescence

increase when the dual-labeled probe is cleaved by the nucleases. However, further

digestion of the cleaved pieces will not induce any more signal change, thus FRET can

only monitor the "first cut". Moreover, many non-specific bind of proteins to the FRET

probe might also cause a conformational change of the DNA, leading to false positives.

In contrast, digestion of a DNA molecule can be monitored by anisotropy until the

smallest fragments. These is also likely no other process that can decrease the anisotropy

of the dye-labeled DNA. Lastly, anisotropy is a ratiometric measurement that is less

affected by photo-bleaching and system variations.

Experimental Section

Fluorescence Anisotropy Imaging (FAI) System

The FAI system was built on an Olympus FV500-IX81 confocal microscope

(Olympus America Inc., Melville, NY) (See Figure 4-1). The emission of fluorescent

samples would go through a few dichroic mirrors before entering a detection channel. A

flat polarization beam splitter (PBS) (MOXTEK, Inc., Orem, UT) replaced a dichroic

mirror in a way that emission of one polarization state would be reflected in one detection

channel while the emission of the other perpendicular polarization state would go though

and be reflected by a mirror into the next channel (Figure 4-1). Two bandpass filters for

emission of TAMRA dye were placed in the two channels right before the PMTs. One of

them was a BP560-600 filter that came with the microscope, and the other was a








580DF30 from Omega Optical, Inc. (Brattleboro, VT). The two filters have similar

transmission profiles.


Sample


Laser


Pinhole

\
I
t





t


PMT 1


PMT 2


Objective

Excitation
beam splitter





Polarization
beam splitter




Mirror


Anisotroov imaae


Figure 4-1. Schematics of the FAI system.


Imgi


Imgll









A 5 mW 543 nM He-Ne laser was the excitation for TAMRA throughout the

experiments. Usually only 20-30% of the laser power was used. The objective used for all

the experiments was a PLAPO60XO3PH 60x oil immersion objective with a numerical

aperture of 1.40 from Olympus (Melville, NY).

Scanning of the samples was controlled by the Olympus FluoView program.

Images of 512x512 pixels were taken simultaneously in the two polarization channels.

The scanning rate was chosen at slow level (-2 s per image) to obtain lower noise level.

Noise was further reduced by taking four images for each measurement and recording the

average of the four. This could be programmed on the computer. The dynamic range of

the confocal signal was 0 to 4095. The signal level could be controlled by a few

parameters such as voltage of the PMT, PMT gain (multiplying signal by a number), and

laser power. We avoided using the latter two parameters because the gain would increase

noise and high laser power could cause photo-bleaching of the dye. The two voltages

applied on the two PMT's were always identical. During the monitoring of dye-labeled

DNA inside live cells, when there was a drop in the intensity level, the both PMT

voltages were increased to keep the signal level in the range of 2000 to 3500. There

would be slight increase of background but that could be easily deducted during image

processing.

When to acquire images during monitoring of the cell was manually controlled.

Time between two acquisitions could range from 2 to 30 minutes. For different cells, the

time of the data points were different.

In an effort to determine the G-factor of the FAI system in calculation of

anisotropy, a pure TAMRA solution was used. Since TAMRA is a small organic









molecule (M.W.=430.45), its anisotropy in water should be very close to zero. Two

polarization images of the TAMRA solution was taken on the FAI system and the

intensities were averaged separately and designated Ivy and IVH. Considering the equation

for anisotropy calculation:

r = (Ivv-G-IvH) / (Ivv+2G-IvH)

Apply r = 0, and the Ivy and IVH values to the equation, we obtained a G-factor

value of 0.61 for our FAI setup. This G-factor was used for all the experiments.

The Image Processing Program

All the raw images obtained from the Olympus software were then processed in the

ImageJ program (http://rsb.info.nih.gov/ij) to calculate anisotropy images using a

homemade software plug-in. What the plug-in did was to read the two polarization

images, find the background and noise levels, and subtract the images with the

background value. Then pixel by pixel, the plug-in would try to calculate an anisotropy

value for each pixel using data from the two raw images. Anisotropy would be calculated

only when the intensity from both images at that pixel was at least 3 times higher than the

noise level, otherwise, an anisotropy of zero was given to that pixel. After doing this for

the whole image, an anisotropy image was produced and recorded. With this plug-in we

programmed, the whole process of calculating anisotropy images could be easily done at

a click of a button.

Cell Culturing

The breast cancer cell line MDA-MB-231 (ATCC, Manassas, VA) was cultured

according the provider's instructions. Cells for injection experiments were cultured in the

glass bottom culture dishes from MatTek Corporation (Ashland, MA). The thin glass

bottom would not interfere with fluorescence measurements as plastic would.