SINGLE MOLECULE STUDIES OF ENZYMES HORSERADISH PEROXIDASE AND ALKALINE PHOSPHATASE USING TOTAL INTERNAL REFLECTION

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1 SINGLE MOLECULE STUDIES OF ENZYMES HORSERADISH PEROXIDASE AND ALKALINE PHOSPHATASE USING TOTAL INTERNAL REFLECTION FLUORESCENCE MICROSCOPY AND CONFOCAL MICROSCOPY Leonora Kaldaras A Thesis Submitted to the Graduate College of Bowling Green State University in partial fulfillment of the requirements for the degree of MASTER OF SCIENCE August 2013 Committee: Peter H. Lu, Advisor Neocles Leontis George Bullerjahn

2 2013 Leonora Kaldaras All Rights Reserved

3 iii ABSTRACT Dr. Peter H. Lu, Advisor Conformational dynamics plays an important role in determining protein reactivity including affinity to substrate and product, catalytic rate constant and product release process. Whereas the influence of conformational dynamics on kinetic parameters of enzymatic reaction have been widely studied by ensemble averaging techniques, in these experiments conformational variations tend to average out and therefore ensemble averaging methods lack the level of resolution necessary to characterize time dependent behavior of individual enzyme molecules, which is essential to understanding enzymatic catalysis. This thesis consists of two parts. The first part focuses on the new development of integrated spectroscopy system, which combined the advantages of wide plane view field of objective-type TIRFM imaging with high time-resolution confocal single-molecule spectroscopy to address the challenge of probing both temporally and spatially stochastic events of the single-molecule reactions, such as fluorogenic enzymatic reactions with enzyme tethered to a solution/glass interface. Tthe calibration of this integrated apparatus is demonstrated with fluorescence microspheres and the feasibility by spectroscopy measurement of single molecule Rhodamine 6G on glass surface. Finally, the application of the apparatus is demonstrated by studying spatially and temporally randomly distributed single-molecule enzymatic reaction of fluorogenic substrate, horseradish peroxidase-catalyzed oxidation of amplex red.

4 The second part focuses on exploratory studies of enzyme Alkaline Phosphatase from E. coli and its potential to be studied on single molecule level using fluorogenic substrate 3-o-methylfluorescein phosphate (MFP) and two polarization total internal reflection fluorescence microscopy apparatus. Ensemble averaged kinetics data and several single molecule trajectories are presented here. iv

5 This thesis is dedicated to my friend, Mr. Allen Baldwin (Uncle Al). v

6 vi ACKNOWLEDGMENTS I would like to thank my advisor, Dr. Peter H. Lu, for his support and guidance throughout my work. I would like to thank Dr. Desheng Zheng, for all the time he spent teaching and explaining things to me in the lab. Without his valuable advice and help it would have been impossible for me to finish this thesis. I would like to thank Dr. Papatya Sevinc for all her help with the graduate coursework (especially Photochemistry) as well as valuable advice in lab work. I also thank Dr. Yufan He and all my lab mates for their help and support. I would like to thank Dr. Neocles Leontis for all his help, encouragement and the invaluable teaching and professional experience I have gained while working with him. I would also like to thank BGSU faculty and staff members, especially Alita Frater and Nora Cassidy for all their help and support. Special thanks go to Vassiliki Leontis for being such a wonderful, supportive and encouraging friend. I also thank my loving family, my mom Marina Kaldaras, my dad, Nikolay Kaldaras, and my little brother Konstantin Kaldaras, for believing in me no matter what.

7 vii TABLE OF CONTENTS Page INTRODUCTION... 1 The Significance of Single Molecule Experiments... 1 Timescales of Single Molecule Experiments... 3 Single Molecule Optical Imaging Techniques... 4 CHAPTER I. TOTAL INTERNAL REFLECTION FLUORESCENCE MICROSCOPY IMAGING-GUIDED CONFOCAL SINGLE MOLECYLE FLUORESCENCE SPECTROSCOPY... 6 Instrument Design... 6 Calibration and Test... 9 Conclusion Sample Preparation CHAPTER II. SINGLE MOLECULE STUDIES OF ENZYME ALKALINE PHOSPHATASE FROM E.COLI Introduction Determination of K m and V max Kinetic Parameters using Ensemble Averaging Methods Discussion of Ensemble Averaging Kinetic Data Sample Preparation for Ensemble Averaging Kinetic Measurements Single Molecule Studies: Apparatus and Sample Preparation Single Molecule Studies of Alkaline Phosphatase: Experimental Results Single Molecule Studies of Alkaline Phosphatase: Discussion and Conclusion... 41

8 viii CONCLUSION REFERENCES... 44

9 ix LIST OF FIGURES Figure Page 1 Schematic overview of basic characteristics of fluorescence confocal microscope and total internal reflection fluorescence microscope Schematic of total internal reflection fluorescence microscopy-guided confocal single molecule fluorescence spectroscopy Calibration of total internal reflection fluorescence microscopy-guided confocal single molecule spectroscopy apparatus Feasibility of total internal reflection fluorescence microscopy-guided confocal single molecule spectroscopy apparatus Schematic representation of enzyme immobilization on (3-Aminopropyl) trimethoxysilane modified coverglass Application of total internal reflection fluorescence microscopy imaging guided confocal single molecule spectroscopy apparatus to study single molecule enzymatic reaction of Horseradish Peroxidase Aggression by a Male Against a Female of His Harem General mechanism of enzymatic catalysis Crystal structure of E. coli alkaline phosphatase in complex with the natural product, phosphate General mechanism of dephosphorylating by alkaline phosphatase Dephosphorylation of 3-o-methylfluorescein phosphate by alkaline phosphatase Enzyme progress curve Michaelis-Menten curve for an enzyme... 25

10 x 13 Examples of enzyme progress curves for alkaline phosphatase at different substrate concentrations Rate of 3-o-methylfluorescein production at different substrate concentrations Michaelis-Menten curve for E.coli Alkaline Phosphatase Lineweaver-Burk plot for an enzyme Experimental Lineweaver-Burk plot for Alkaline Phosphatase Definition of Hill coefficient using Michaelis-Menten equation Determination of kinetic parameters using IgorPro software Scheme of the two channel total internal reflection fluorescence microscopy for single molecule studies of alkaline phosphatase Sample preparation for single molecule studies of alkaline phosphatase Apparatus calibration for single molecule studies of alkaline phosphatase Sample trajectories of single molecule reaction of alkaline phosphatase Analysis of single molecule trajectories for alkaline phosphatase in Excel Signal to noise analysis of single molecule trajectories in Excel... 40

11 xi LIST OF TABLES Table Page 1 Kinetic parameters for Alkaline Phosphatase determined from ensemble averaging measurements... 29

12 1 INTRODUCTION The significance of single-molecule experiments When teaching chemistry to students, we generally start by describing the basic principles of molecular interactions and chemical reactions using single molecules as examples. However, our understanding of molecular interactions and chemical dynamics has come almost exclusively from experiments on ensembles of molecules. The capability of studying the behavior of single molecules has been developed quite recently and is now a well-established, rapidly developing field of fundamental science. Recent advances in optical spectroscopy and microscopy have made it possible not only to detect and image single molecules, but to conduct spectroscopic measurements and monitor dynamic processes. This made optical spectroscopy and microscopy a powerful method for studying the behavior of biomolecules such as proteins and their interactions with the environment because optical measurements provide the time resolution required to monitor dynamic processes such as enzymatic catalysis. Even though single-molecule measurements represent a technological breakthrough with practical applications in biological sciences, it has been debated whether any fundamentally new information can be obtained by using single molecule techniques. This comes from the fundamental postulate of statistical mechanics, which states that the time-average of a physical quantity along the trajectory of a member of the ensemble is equivalent to the average of that quantity at a given time over the ensemble. Why, then, study a single molecule for a period of time instead of obtaining the same information by sampling the ensemble of molecules simultaneously? In order for the fundamental postulate to be applicable, the ensemble must consist of entirely equivalent members, i.e. it must be homogeneous 1. However, biological

13 2 systems such as enzymes are inhomogeneous. The inhomogeneity can be static (different enzyme molecules can show different properties), or the measurement time can be shorter than the relevant fluctuations. Whatever the origin, in an inhomogeneous system, the trajectory average varies among the members of the ensemble and is no longer equivalent to the ensemble average. The advantage of single-molecule experiments over ensemble-averaged experiments lies in two major reasons 1. First, ensemble-averaged measurements can be used to determine the mean value of a quantity but cannot generally be used to determine the distribution of the quantity. Without the distribution, the behavior of individual members of the ensemble cannot be deduced. Second, single-molecule trajectories contain detailed dynamical and statistical information and therefore analyses of trajectories are more informative than are ensembleaveraged results. Both of these statements are supported by recent single molecule studies. For example, when looking at enzyme kinetics, it has been demonstrated by Brian English and coworkers 2 that enzymes exhibit catalytic rate fluctuations of large amplitude and broad timescales at the single-molecule level. This behavior was uncovered by analyses of long turnover time trajectories and it supports the fact that an enzyme molecule is an ever-fluctuating dynamic entity during catalysis. This phenomenon cannot be observed in ensemble averaged experiments for two reasons. First, in pre steady-state ensemble measurements, data often lack the dynamic range necessary to identify long tails in multiexponential kinetics. Second, in steady-state ensemble measurements, data are masked by the fact that the Michaelis-Menten equation holds for both single conformer and multiple slowly interconverting conformers with different enzymatic activities. The effects of enzymatic fluctuations would be less significant for a system comprising many enzyme molecules. However, if a system contains a small number of

14 3 enzyme molecules, (like a living cell), the enzymatic fluctuations could be biologically important. Therefore, studying protein dynamics on single molecule level can provide important additional information on the forces that drive enzymatic reactions therefore allowing for better understanding of this fundamental biological process. Timescales of single molecule experiments Single molecule fluorescence measurements can be obtained on the millisecond to tenthsecond timescale, which include enzymatic turnover trajectories. Single molecule techniques don t have the capability of recording trajectories on the timescale of molecular dynamics simulations (femtosecond to nanosecond) due to signal-to-noise considerations and repetitive excitation, each cycle of which occurs maximally on the nanosecond timescale. Fluorescence lifetimes of single molecules can nevertheless be measured with picosecond resolution using time-correlated photon counting. The measured lifetime is averaged over the period of milliseconds or longer. Such measurements allow protein motions, photoinduced electron transfer, and fluorescence resonant energy transfer to be probed on single molecules. The major limitation to recording and interpreting time trajectories on single molecule level is that repetitive excitation may create photoinduced artifacts associated with photophysical and photochemical processes not related to the process under study. Understanding the photophysics and photochemistry of the system under study is therefore crucial in interpreting the obtained data correctly. The most common of these interfering processes is the irreversible photochemical bleaching of the chromophore, which terminates the single molecule trajectory.

15 4 Single-molecule optical imaging techniques Fluorescence confocal microscopy (FCM) has been widely applied in the single-molecule imaging 3-8 with diffraction limited spatial resolution near half of the excitation light wavelength nm, as illustrated in Figure 1(a). Its capability of decreasing detection volume has shown a significant advantage in extraction of weak signals from background, through confining the excitation and detection in the conjugated diffraction limited focal volumes 3-5, 9, 10. Combined with fast-gated single photon counting techniques and ultrafast pulse laser excitation, confocal configuration is well used for single-molecule time-resolved fluorescence measurements and fluorescence correlation spectroscopy 9-12, which can provide the information about single molecule dynamics, associated with molecular interactions and biochemical reactions in biological and chemical systems. Compared to fluorescence confocal microscopy, total internal reflection fluorescence microscopy (TIRFM) takes the advantages of evanescent light field with exponential intensity decay at the interface to simultaneously probe a wide plane area, typically 104 times larger than the size of a confocal focus spot, as illustrated in Figure 1(b). Fluorescence confocal microscope is often equipped with avalanche photodiodes or single photon avalanche diodes (SPAD), which are suitable for high time resolution measurements and time-resolved dynamics analyses. However, the single pinpoint detection constrains the efficiency of detecting spatially and temporally randomly distributed single molecule fluorogenic events, such as fluorogenic product turnovers of single-molecule enzymatic reactions. TIRFM cannot provide high time resolution due to two-dimensional imaging.

16 5 Figure 1. Schematic overview of basic characteristics of fluorescence confocal microscope and total internal reflection fluorescence microscope. (a) Fluorescence confocal microscope (FCM) with excitation beam waist nm. (b) Total internal reflected fluorescence microscope (TIRFM) with wide excitation area, which is typically 104 times larger than confocal spot in objective-type configuration, and with vertical resolution 100 nm. However, the much larger simultaneously sampling view fields dramatically improve the possibility of identifying the tethered enzymes only by the temporally random fluorogenic signal. Therefore, it is desirable to design an integrated spectroscopy combining the advantage of wide field imaging of TIRFM with the advantage of high time resolution of confocal fluorescence spectroscopy, through (1) detecting reaction active sites and recording their coordinates by TIRFM and (2) relaying on the recorded coordinates to guide the confocal single-molecule spectroscopy measurements for individual pinpoints of interest. Combining the advantages of spatial resolution of confocal microscopy and total internal reflection microscopy had been well demonstrated by the integrated apparatus in previously reported studies 19-27, which deployed vertical (z-axial) resolution of total internal reflection configuration and lateral resolution of confocal configuration to improve the imaging spatial resolution.

17 CHAPTER I. TOTAL INTERNAL REFLECTION FLUORESCENCE MICROSCOPY IMAGING- GUIDED CONFOCAL SINGLE MOLECYLE FLUORESCENCE SPECTROSCOPY 6 Instrument design This objective-type TIRFM imaging-guided confocal single-molecule fluorescence spectroscopy system (Figure 2) is based on an inverted microscope (Axiovert 200M, Carl Zeiss) in an epiillumination configuration to combine TIRFM mode with confocal mode, facilitated with a home developed software for recording imaging from TIRFM mode and shifting the pinpoints of interest from TIRFM imaging to confocal single-molecule spectroscopy measurements. In the TIRFM mode, a cw laser (GCL-050-L, Crysta- Laser) beam is expanded by the telescope of lens 1 and lens 2, and aligned to the side port of the microscope stand by a dichroic mirror beam splitter (DM1) with axis of rotation located in the intermediate image plane. The beam from the side port is then reflected by the side port prism (P1), passed through the empty filter cube, and focused by the tube lens (TL) on the back focal plane of the objective (Plan-Fluar, 1.45NA, 100, Carl Zeiss). After passing through the objective, the beam is tightly collimated and incident on the cover glass. By fine adjusting the rotation angle of the DM1 slightly from 45 o relative to the incident direction of the laser beam, the incident angle at the cover glass/solution interface can be adjusted to exceed the critical angle of total internal reflection. An evanescent electromagnetic field with exponential intensity decay in the normal direction is generated at the cover glass-solution interface. The fluorescence molecules located in the evanescent field range are selectively excited. The fluorescence emission is collected by the same microscope objective, passing P1, reflected by M2 to the top port of the microscope stand. Before entering the multiplication charge coupled device (EMCCD) camera (Photomax 512B, Princeton

18 7 Instruments), the emission signal is purified by dichroic mirror beam splitter (DM2) and emission filter (F1) with deflection and blocking the excitation laser light, respectively. In our current experiments, the refractive index of solution is 1.33 and that of the cover glass is 1.515; here, 0.68o deviation from the 45o incident angle of the DM1 provides total internal reflection with evanescent field depth about 100 nm. Figure 2. Schematic of TIRFM imaging-guided confocal single-molecule fluorescence spectroscopy. Laser 1: cw laser source; Laser 2: pulse laser source; L1 L5: lenses; M1: reflection mirror; DM1 DM3: dichroic mirror beam splitters; P1: side port prism for left/vis obs; TL: tube lens; Obj: Objective; XYSS: x-y scanning stage; M2: beam path switch for visual obs; F1 F2: emission filters; PH: excitation pin hole; P2: side port prism for left. TIRFM mode beam path: Laser 1-L1-L2-M1-DM1-P1-TLObj-TL-P1-M2-DM2-F1-EMCCD; confocal mode beam path: Laser 2-L3-PH-L4-DM3-Obj-DM3-TL-P2-DM1-L5-F2-SPAD. In the confocal mode, a femtosecond pulse laser (Laser 2, Ti:Sapphire Mira 900F/P, Coherent Inc.) is used. After an optical parametric oscillator (OPO BASIC, Coherent Inc.) and frequency doubling by a β-barium borate crystal (BBO), the linear-polarized pulse laser is aligned through a lens-pinholelens system which includes lens (L3), pinhole (PH, 10 μm), and

19 8 lens (L4) to expand the beam diameter. Then the extended parallel beam, passing through the back port of the microscope stand, is reflected by the dichroic mirror beam splitter (DM3) in the filter cube, and focused on cover glass-solution interface by the objective (Obj). The cover glass is fixed on the piezoelectric scanning stage (Nano-H100, MCL) with a positioning resolution of 0.2 nm for fine-tune. Fluorescence emission from the excitation focal volume is collected by the same objective (Obj). After transmitting DM3, the signal is reflected by the side port prism (P2), transmitted DM1, and then refocused by the re-magnification lens (L5) to the SPAD (PDM50ct,MPD) entrance. Here the sensor target diameter of SPAD is 50 μm, which also serves as a pinhole to reject stray and ambient light noise. The signal from the detector is processed by a single photon counting module (SPC-830, Becker & Hickl GmbH) to record both chronic arrival time and delay time between the pulse excitation and molecule excited state emission with high time resolution for each single photon event. The common part of the optical paths for the two modes lies between DM1 and the cover glass. Switching between the two modes is achieved by toggling the filter cube turret and the side port prism. To guide the confocal spectroscopic measurements of the spots of interest in the TIRFM frame, the coordinates of the spots of interest in sample scanning stage frame is determined by matrix transformation. The sample does not move relative to the stage, which makes the system stable under experimental conditions. The transformation relationship is as follows: ( ) ( ) ( ) [ ] [ ] (1)

20 9 Here (x ccd, y ccd ) is any spot of interest in TIRFM frame, and (x sss, y sss ) is the coordinate of this spot in sample scanning stage frame. T is the transformation matrix that can be determined from any three non-collinear patterned spots coordinates in TIRFM frame and the sample scanning stage frame. Calibration and Test The calibration experiment was performed using 100 nm fluorescence microspheres (F-8800, Invitrogen) as imaging probes to construct the transformation matrix. Excitation and emission maxima of the fluorescence microspheres are 540 nm and 560 nm, respectively. The sample was prepared by dropping diluted solution of microspheres on a clean cover glass (Gold seal, 3419). When the microspheres were dry, the cover glass was placed on the sample scanning stage. Then, about 30 μl of water was slowly dropped near the laser position on cover glass, and the microspheres were still attached to the cover glass. Figures 3(a) and 3(b) show the fluorescence images from EMCCD in the TIRFM imaging mode and from SPAD through scanning the sample scanning stage in the confocal imaging mode, respectively. From the distribution pattern of the microspheres, it is identifiable that the microsphere a in Figure 3(a) is the same one as the microsphere a in Figure 3(b). Similarly, the microsphere b and microsphere c in Figure 3(a) are the same ones as the microsphere b and microsphere c in Figure 3(b). Substituting the corresponding coordinates of a, b, and c in Figures 3(a) and 3(b) to Eq. (1), the transformation matrix is then determined, and this transformation relationship is used to guide registration of any spot of interest in EMCCD view field to the confocal excitation spot.

21 10 Figure 3. Calibration of TIRFM imaging-guided confocal single-molecule fluorescence spectroscopy. (a) Fluorescence image of the microspheres in EMCCD coordinates by TIRFM mode. (b) Fluorescence image of the microspheres in scanning translation stage coordinates by confocal mode. According to the special pattern, the microspheres a, b, and c in (a) are the same ones a, b, and c in (b). Considering that the accuracy of localization of the selected microspheres in different coordinates influences the accuracy of transformation matrix T, we calculated the center of each microsphere of interest by using the two-dimensional Gaussian fit of the spatial distribution of fluorescence intensity. The uncertainty of the center localization is determined by 29 ( ) (2) where the index i refers to x axis or y axis, and other parameters are the number of collected photons (N), the standard deviation of Gaussian distribution (s), the pixel size of imaging detector (a), and the background noise (b). The number of collected photons is crucial in determining the uncertainty (σ i ). In this experiments, using the high brightness fluorescence

22 11 microspheres and the long exposure time, the uncertainty of the center can be controlled in nm scale. We have demonstrated the feasibility of the TIRFM imaging-guided confocal singlemolecule fluorescence spectroscopy with single-molecule spectroscopic measurement of Rhodamine 6G molecules. After placing cover glass on the sample scanning stage, 30 μl water was dropped on the cover glass, which provided a solution/cover-glass interface. Figure 4(a) shows a total internal reflection fluorescence microscopy imaging result of Rhodamine 6G molecules. According to transformation relationship, the emission of any pinpoint spot of interest in Figure 4(a) can be shifted to the focal spot of the femtosecond laser by coordinating the computer controlled x-y scanning stage at nanosclae precision. Figure 4(b) shows the time resolved fluorescence intensity decay of single Rhodamine 6G molecule from the selected spot a of Figure 4(a) imaged by TIRFM. The lifetime of the confined single Rhodamine 6G was determined as 3.1 ± 0.3 ns by a single exponential fit. Figure 4. Feasibility of TIRFM imaging-guided confocal single-molecule fluorescence spectroscopy. (a) Image of Rhodamine 6G molecules in TIRFM mode. (b) The corresponding confocal single-molecule spectroscopy measurement on molecule a in (a). The fluorescence intensity decay from single-molecule photon stamping recording is fitted with exponential decay with fit residual; the fluorescence lifetime of the single Rhodamine 6G molecule is 3.1 ± 0.3 ns. Inset: The intensity trajectory of the single molecule.

23 12 Finally, the application of TIRFM imaging-guided confocal single-molecule fluorescence spectroscopy was demonstrated through detecting spatially and temporally randomly distributed single-molecule enzymatic reaction of fluorogenic substrate, horseradish peroxidase-catalyzed oxidation of amplex red. The spatially and temporally randomly distributed turnover events of fluorescent products of the enzymatic reaction were recorded by EMCCD in TIRFM mode (figure 5). Figure 5. Schematic representation of the enzyme immobilization on (3-Aminopropyl) trimethoxysilane modified coverglass. A fluorogenic substrate amplex red in PBS buffer is converted to fluorescent resorufin product by the single enzyme HRP under the initiation of hydrogen peroxide.

24 13 The single-molecule photon time-stamping data contains more detailed information than a conventional time-correlated single photon counting delay time histogram, adding the chronic arrival time trajectory, i.e., the consecutive arrival time of each detected photon. Figure 6(a) shows the single molecule photon time-stamping recording of an individual turnover event of fluorescent product from one of the active tethered enzyme. Each dot in Figure 6(a) corresponds to a detected photon plotted by its chronic arrival time and delay time. From the photon density along the x axis, the background and the signal associated with recording a product turnover before product releasing can be clearly discriminated. Shown here is only the lifetime analysis of the enzymatic product resorufin confined by a single enzyme before it was released. The fluorescence intensity decay is shown as dots (which were binned with 0.25 ns in histogram) in Figure 6(b). The lifetime of the detected enzymatic product resorufin molecule is determined as 3.9 ± 0.2 ns by a single exponential fit, and the corresponding fitting residual is also shown.

25 14 Figure 6. Application of TIRFM imaging-guided confocal single-molecule fluorescence spectroscopy to studying spatially and temporally randomly distributed single-molecule enzymatic reaction, horseradish peroxidase-catalyzed oxidation of amplex red. (a) The singlemolecule photon time-stamping raw data of an individual turnover event of fluorescent product from one of the active tethered enzyme. Each dot corresponds to a photon stamped with the chronic arrival time (t) and the single-photon delay time (Δt) between the photon emission and femtosecond laser pulse excitation. (b) Fluorescence intensity decay of a newly formed single resorufin molecule confined in the enzyme. The fluorescence intensity decay from singlemolecule photon time stamping recording is fitted with exponential, with fit residual. The fluorescence lifetime of the confined single resorufin molecule is 3.9 ± 0.2 ns.

26 15 Conclusion In conclusion, demonstrated here is a new integrated single-molecule spectroscopy approach that uses total internal reflection fluorescence microscopy imaging to guide confocal single molecule fluorescence spectroscopic measurements. With the localization by two-dimensional Gaussian fit and coordinates transformation, any single-molecule fluorescence spot of interest in the imaging field of TIRFM imaging mode is accurately registered and shifted to the excitation focus of confocal mode for single-molecule fluorescence spectroscopic measurements through a computer-controlled close loop piezoelectric x-y scanning stage. This integrated system has a number of significant advantages including the low requirement for laser power of the femtosecond pulse laser, absence of photon bleaching by wide field excitation and high signal collection efficiency for the confocal single-molecule fluorescence spectroscopic measurements. The system s capability to simultaneously carry out multiple molecules sampling and in situ fluorescence time-resolved analyses for pinpoint individual molecules is demonstrated here. This apparatus can be used to conduct the measurements of the spatially and temporally stochastic single-molecule on interfaces. Sample Preparation The cover glass was cleaned by corrosion of fresh chromic acid solution to eliminate the possible fluorescent spots. After washing with water and drying with nitrogen gas, the clean cover glass was first spin-coated (3000 rpm, Spincoat G3-8) with 20 μl Rhodamine 6G aqueous solution ( M), then 30 μl PMMA/toluene (1 wt. %) solution was spin-coated on the cover glass to confine Rhodamine 6G molecules from diffusion. The thickness of PMMA film is about 26 nm, and the optical index of the PMMA film is similar to that of the cover glass (both are close to 1.52).

27 16 To immobilize horseradish peroxidase enzyme the clean cover glass was silanized overnight with mixture solution of (3-aminopropyl) trimethoxysilane (Fluka, 09324), isobutyltrimethoxysilane (Sigma, ), and dimethylsulfoxide (Sigma, D4540) with ratio at 1:30 000: (in volume). After being rinsed with methanol and water, the silanized cover glass was incubated in 10 nm dimethyl suberimidate (Thermo Scientific, 20700) in 50 mm PBS buffer (ph 8.0) for 4 h. The phosphate buffer (PBS) was prepared with potassium phosphate monobasic solution (Sigma Aldrich, P8709) and potassium phosphate dibasic solution (Sigma Aldrich, P8584). At this step, one amine-reactive imidoester group of dimethyl suberimidate is linked to the amino group on the modified cover glass surface. After additional washing, the cover glass was incubated with 1 nm horseradish peroxidase (HRP) (Sigma Aldrich, P8375) in 50 mm PBS buffer (ph 8.0) for 4 h followed by rinsing with water and PBS buffer. Then, another amine-reactive imidoester group of dimethyl suberimidate reacted with the amino group in HRP and linked the HRP to the cover glass. After placing a modified cover glass on the sample scanning stage, 30 μl reaction solution containing amplex red (200 nm), H 2 O 2 (2 mm) in PBS buffer solution (ph 7.3) was dropped on the cover glass.

28 CHAPTER II. SINGLE MOLECULE STUDIES OF ENZYME ALKALINE PHOSPHATASE FROM E.COLI 17 Introduction Enzymes fulfill their function as biocatalysts by lowering the activation energy required for a chemical reaction to occur. The catalytic reaction consists of multiple steps each occurring at different rates starting from substrate binding proceeding to product formation and product release as shown in figure 7. Figure 7. General mechanism of enzymatic catalysis Each step involves complex molecular interactions, inhomogeneous conformational changes, and the fluctuations of local environment within the enzyme s active site. Recently, fluctuating enzyme paradigm has been developed, according to which the properties of enzymes fluctuate over time. Specifically, enzymes exhibit static and dynamic disorder The former is supported by the fact that different enzyme molecules can show different properties and the latter one by the fact that properties of individual enzyme can fluctuate over time. In the case of the enzyme these properties are related to the three dimensional structure (if enzyme is a monomer) or to the quaternary structure (if enzyme consists of multiple subunits). The folding of the enzyme determines its properties and therefore is essential for enzymatic activity 31. The requirement for a specific configuration is especially necessary for enzymes because most of them rely on a certain, very precise spatial coordination of specific amino acid residues to form

29 18 an active site. This specific configuration must be maintained in the presence of thermal motion, which constantly induces fluctuations in the amino acid chain 33. The experimental observation that each individual enzyme s activity is not constant in time 34, is easily explained by these fluctuations in the three dimensional structure of the enzyme: the enzyme can appear in different conformations, each with its own specific activity. In addition to the observation that conformations may differ in terms of catalytic activity, it is now well established that conformational changes are essential in many enzymatic reactions. They can even be the ratelimiting step in the catalytic turnover cycle Single molecule fluorescence microscopy is a powerful approach in characterizing dynamic conformations and understanding enzymatic reactions mechanisms 39. The photophysical properties of fluorophores such as such as fluorescence band intensities, fluorescence quantum yield, and fluorescence decay rate constants are sensitive to the local environment and therefore can be used to study enzyme conformational dynamics. In particular, fluorescence quantum yields increase dramatically with increase in the solvent polarity 40. Fluorescence anisotropy provides information about the motion of the protein fragment to which the fluorescence probe is attached. Anisotropy decay yields information on the protein structure and conformational changes as well as on the protein flexibility 41. Fluctuations of single molecule fluorescence lifetime are induced by the electrostatic interaction of the probe molecule with its local, inhomogeneous environment 42. This intrinsic ability of the probe molecule to sense its surrounding makes the single-molecule spectroscopy a useful tool for studying conformational dynamics of enzymes. In contrast to classical ensemble measurements, which average over the whole population, single molecule experiments are able to detect conformational heterogeneities,

30 to identify transient or rare conformations, to follow the time series of conformational changes and to reveal parallel reaction pathways. 19 There are generally three categories of reporter systems used for studying single molecule conformational dynamics of enzymes. They include the systems that take advantage of the intrinsic fluorescence from tryptophan or tyrosine residues of enzymes, systems that have undergone site-directed fluorescent labeling (like those used for FRET studies), and systems that use artificially derived fluorogenic molecules as substrates for enzymatic reaction. When available, the fluorogenic substrate reporter system is the best choice. The reason is that fluorescent product molecules are produced constantly as a result of the catalytic reaction and therefore eliminate the photo bleaching problem. As a result, long time trajectory can be recorded for the enzymatic reaction of a single enzyme for dynamic disorder analysis. These trajectories are sufficiently long to represent a statistically relevant sampling of all possible conformational states and therefore provide invaluable information about the mechanism of enzymatic reaction. Alkaline phosphatase from E. coli is a metalloenzyme system that requires magnesium and zinc ions for full activity In E. coli this enzyme is produced under the conditions of phosphate starvation. Phosphate is needed for metabolism and is especially important in signal transduction pathway. Since the main function of alkaline phosphatase is to dephosphorylate a wide variety of phosphoester substrates, it is produced in the bacterial cell when free phosphate is no longer available for uptake directly from the environment. In that case, alkaline phosphatase dephosphorylates various phosphate containing substrates present in the cell 46.

31 20 Alkaline phosphatase is a dimer of identical subunits, each consisting of 429 amino acids, two zinc atoms and two magnesium atoms. The crystal structure of alkaline phosphatase from E. coli is shown in figure 7. The zinc ions are essential for enzymatic activity, whereas magnesium Figure 8. Crystal structure of E. coli alkaline phosphatase in complex with natural product phosphate 44. ion does not directly participate in the enzymatic reaction, but is believed to play a role in subunit communication that leads to a phenomenon called half-site reactivity 45. It is also sometimes referred to as negative cooperatively in phosphate binding and the result of this process is that phosphate binds to one of the sites with higher affinity than to the other. It is also believed that magnesium ion is involved in allosteric regulation and might play a role in establishing the half-site reactivity phenomenon of alkaline phosphatase From X-ray crystallographic structure and ensemble averaged experiments is has been established that the mechanism for ester dephosphorylation proceeds according to the mechanism in figure :

32 21 Figure 9. Mechanism of dephosphorylation by enzyme E. coli alkaline phosphatase. E is the alkaline phosphatase enzyme, ROP is the phosphoester substrate, E*ROP is the enzyme substrate complex, E-P is reaction intermediate formed via covalent bond between phosphate and Ser 102 residue in the active site of the enzyme and E*P is the non-covalent complex of enzyme with phosphate. The rate of phosphorylation and dephosphorylation is strongly dependent upon the ph of the environment as was showed by the NMR experiments 46. The experimental evidence shows that phosphorylation of serine residue is a rate-limiting step at acidic ph (5.5 and below) 46, whereas at alkaline ph (8-9), where the enzyme is most active, the rate limiting step is the release of phosphate group 46. Magnesium ion is believed to be involved in facilitating conformational change that promotes product release. This conformational change is thought to be responsible for generating the subunit with higher and lower affinity for the substrate However, the role of Mg ion in this process as well as the structural changes involved in subunit communication still remains poorly understood and the evidence for site homogeneity also exists 47. Alkaline phosphatase has been studied by means of single molecule techniques Both studies were performed by professor Dovichi and coworkers, but the evidence for catalytic heterogeneity is quite controversial. In the first study the kinetic information was obtained by performing replicate incubations on individual molecules in capillaries and measuring the

33 22 amount of product generated during incubation period from fluorogenic substrate 48.In this experiment it was found that the reaction rate had a very broad distribution centering around 60, 160 and 290 s -1 the most abundant being around s -1. This evidence could serve as a proof for multiple conformations of the enzyme present during the observation time. In the other study, however, highly purified alkaline phosphatase showed no heterogeneity in observed reaction rates 49. Both studies were performed using endpoint measurement technique and did not provide dynamic information on the degree of conformational change involved in allosteric regulation of product release by alkaline phosphatase. In order to resolve detailed conformational changes it is necessary to employ single molecule methods with single fluorophore sensitivity. This approach will allow for continuously monitoring single turnover reaction and accompanying conformational changes. In particular, enzyme substrates that are converted into fluorescent dye molecules in combination with timeresolved techniques provide a powerful tool for studying conformational dynamics of proteins. Since the rate of alkaline phosphatase reaction is on the order of s -1, time resolution measurement on this scale should be sufficient to visualize processes involved in reaction turnover. This degree of time resolution can be achieved using continuous wave (cw) laser in combination with CCD detection system and fluorescent microscope capable of detecting two polarizations from the emission signal. The following paragraphs describe ensemble average kinetic studies of enzyme alkaline phosphatase using fluorogenic substrate 3-o-methylfluorescein phosphate (MFP), single molecule sample preparation techniques, microscope apparatus description and preliminary single molecule enzymatic reaction imaging and trajectory examples that could be used by future scientists who intend to continue single molecule studies on alkaline phosphatase.

34 23 Determination of K m and V max Kinetic Parameters for Alkaline Phosphatase Using Ensemble Averaging measurements Prior to performing single molecule experiments, it is useful to perform ensemble averaged kinetics studies of the chosen system to ensure that the chemical reaction works, the auto oxidation of fluorogenic substrate is not very significant to interfere with single molecule measurements and to obtain kinetic parameters for the system under study. In this section ensemble averaged data for alkaline phosphatase reaction with 3-o-methylfluorescein phosphate (MFP) fluorogenic substrate is presented. The reaction is illustrated in figure 10. Figure 10. Dephosphorylation of MFP substrate by alkaline phosphatase yielding fluorescent 3-o-methylfluorescein product that has excitation maximum at 488 nm and emission maximum at 520 nm. The kinetic analysis was performed using standard Michaelis-Menten (MM) approach, where the fluorescence product generation was monitored on a fluorimeter at the emission maximum of 520 nm and using different increasing concentrations of substrate keeping the enzyme concentration constant.

35 24 An example progress curve for an enzyme assay is shown in figure 11. Phase 1 of the enzyme progress curve is usually very fast (first few milliseconds) and is called pre-steady-state phase. Since pre-steady-state kinetics is therefore concerned with the formation and consumption of enzyme substrate intermediates (such as ES or E*) until their steady-state concentrations are reached, this phase is not taken into consideration in Michaelis-Menten kinetic analysis which assumes steady-state conditions 52. Figure 11. Enzyme progress curve During phase two (steady state phase) of the reaction the enzyme produces product at an initial rate that is approximately linear for a short period after the start of the reaction. As the reaction proceeds and substrate is consumed, the rate continuously slows down as shown in phase 3 of the progress curve. To measure the initial (and maximal) rate, enzyme assays are typically carried out while less than 5 percent of available substrate is consumed 50, 51. Once the progress curve is measured for a given concentration of the substrate, the slope can be computed which gives the maximum velocity at this concentration of substrate. The same experiment is

36 25 carried out then for other substrate concentrations and Michaelis-Menten curve can then be constructed from which K m and V max are determined as shown in figure 12. Figure 12. Michaelis-Menten curve for an enzyme The data for alkaline phosphatase kinetics at different concentrations of MFP substrate is shown in figure 13. The reaction was monitored for 600 and 900 seconds. The best fit from 600 seconds curve was used to construct the substrate-velocity curve. The control sample contained all the chemicals except for the enzyme and as the data suggests, does not exhibit significant rate of auto hydrolysis. The overall plot for the catalytic reaction for different substrate concentrations is shown in figure 14. The slope value of every curve in figure 14 (velocity) was plotted against 3-o-methylfluorescein product concentration as shown in figure 15. As you can recognize, this is the standard curve for determining Michaelis-Menten kinetic parameters such as K m and V max (like the theoretical curve shown in figure 12).

37 26 3-o- Mthyl Fluorescein generation for 5000nM MFP+1e- 11 M AP in Glycine ph 9.08 Fluorescein concentration, nm Time, sec y = 0.094x R² = nM MFP+AP (900sec) 500nM MFP+AP (600sec) 5000nM MFP Linear (500nM MFP+AP (600sec)) Fluorescein, nm o-Methyl Fluorescein generation for 6000nM MFP+1e- 11 M AP in Glycine ph 9.08 y = x R² = Time, sec 6000nM MFP+AP (900sec) 6000nM MFP+AP (600sec) 6000nM MFP Linear (6000nM MFP+AP (600sec)) Figure 13. Examples of enzyme progress curves for alkaline phosphatase at different concentrations of MFP substrate.

38 27 3-o-Methyl Fluorescein production for 1e-11M AP in Glycine ph 9.08 at different concentrations of MFP substrate nM MFP 2000nM MFP 3000nM MFP 4000nM MFP 5000nM MFP 6000nM MFP 7000nM MFP 1000nM MFP 2500nM MFP 3500nM MFP 4500nM MFP 5500nM MFP 6500nM MFP 15000nM MFP R² = R² = Fluorescein concentration, nm Time, sec R² = 0.98 R² = R² = R² = R² = R² = R² = R² = R² = y = x R² = y = x R² = Figure 14. Rate of 3-o-methylfluorescein production at different MFP substrate concentrations by enzyme alkaline phosphatase. The equation of straight line is shown for two bottom graphs. The slope for every straight line from this figure is plotted against product concentration in figure 15 in order to find K m and V max for the reaction.

39 28 Velosity, nm/sec Michaelis-Menten curve for AP enzyme with MFP substrate MFP, nm Figure 15. Michaelis-Menten curve for enzyme alkaline phosphatase from E. coli. The K m and V max parameters can be determined using the data in figure 15. The standard approach is to take a reciprocal of both the velocity and the substrate concentration and to construct the Lineweaver-Burk plot 51 like the one shown in figure 16. Figure 16. Lineweaver-Burk plot for determination of kinetic parameters

40 29 The experimental data is shown in figure 17. The kinetic parameters calculated from Lineweaver-Burk plot are shown in table 1. Lineweaver-Burk plot for AP+MFP reaction 60 y = 23136x R² = /velocity(nM/sec) /[MFP, nm] Figure 17. Experimental data for Alkaline Phosphatase Lineweaver-Burk plot. Vmax(nM 3-o-m-fluorescein/sec) Vmax(microM 3-o-m-fluorescein/sec) 1.69E-03 Km (nm) Km(microM) Kcat=Vmax/[active sites], sec^ Kcat, msec^ msec/3-o-m-fluorescein molecule sec/3-o-m-fluorescein molecule Table 1. Kinetic parameters of alkaline phosphatase using 3-o-methylfluorescein substrate

41 30 The same parameters were also calculated using computational software IgorPro and the built-in function for calculating kinetic parameters for enzymatic reactions by fitting Michaelis- Menten equation for enzyme progress curve. This method allows to determine the cooperatively coefficient for multi-subunit enzymes. As shown in figure 18, the cooperativity (Hill) coefficient h is equal to 1 when no cooperativity occurs. Values greater than one indicate positive cooperativity in ligand binding (once one ligand molecule is bound to the enzyme, its affinity for other ligand molecules increases). Maximum value of Hill coefficient can t exceed the number of active sites in the enzyme (two for the case of alkaline phosphatase) 51. Experimental results are shown in figure 19. Figure 18. Definition of Hill coefficient using fitting of Michaelis-Menten equation. The sigmoidal shape of enzymatic velocity-substrate curve changes as the degree of cooperativity changes.

42 31 Velocity, nm fluorescein/sec Coefficient values ± one standard deviation base = ± max = ± same as Vmax rate = ± degree of cooperativity xhalf = ± 467 Same as Km MFP, nm x10 3 Figure 19. Determination of kinetic parameters for alkaline phosphatase using enzyme progress curve fitting in IgorPro software. Units for V max are[nm product/min] and units for K m are nm product.

43 32 Discussion of Ensemble-Averaging Kinetic Data for Alkaline Phosphatase The kinetic analysis of alkaline phosphatase from E. coli using MFP substrate has never been performed before. This substrate, however, has been widely used in several studies to determine the level of alkaline phosphatase activity in seawater as well as to study activity levels of alkaline phosphatase in different aquatic microbial species. Even though it is difficult to compare the values of kinetic parameters obtained in this experiment to those reported in the studies because of significant differences in methodologies, sample preparation and sample purity, the results for K m and V max obtained in this study were in the range of those reported for alkaline activity in different microbial populations. Single molecule studies of calf intestinal alkaline phosphatase by Dovichi group report a broad non-gaussian distribution of catalytic rate constant K cat with 10-fold range of activity that tends to cluster around 60, 160 and 290s -1. The mean activity for the combined data set is 111 s 1, standard deviation of distribution is 83 s -1. For the bulk assay the average activity is 380+/-108s 1. In the current studies, the K cat has been found to have the value of s 1 which is significantly lower than the one reported for calf intestinal alkaline phosphatase. This may be due to the fact that alkaline phosphatase from E.coli exhibits different kinetics from calf intestinal alkaline phosphatase or because at concentrations used in the current study (1e-11M) the dimer of alkaline phosphatase dissociates and therefore the number of active alkaline phosphatase molecules is smaller than the apparent concentration. It could therefore be necessary to repeat the kinetic experiment to ensure the accurate value for the Kcat parameter. Furthermore, the two methods for calculating kinetic parameters, the traditional Lineweaver- Burk method and a more recent method of directly fitting the Michaelis-Menten equation using

44 33 IgorPro software have been demonstrated to yield similar values (compare V max =0.169 nm/sec and K m =3921 nm from Lineweaver-Burk plot and V max =0.154 nm/sec and K m = 3675 nm from Michaelis-Menten equation feat using IgorPro software. Therefore, both methods are reliable when performing calculations for kinetic parameters of an enzyme. If using IgorPro software method, it is possible to get an additional parameter, the Hill coefficient, which shows the degree of cooperativity between the two subunits of alkaline phosphatase. In the current experiment, the obtained value is 1.4, which is a little higher than one and therefore shows that there is some degree of cooperativity between the two subunits of the enzyme. As one can see from figure 13, the MFP substrate exhibits almost no auto hydrolysis with time, but does contribute to background fluorescence since the the signal level of control stays at around 40 nm fluorescein when 5000nM MFP is used and rises to about 50 nm fluorescein when 6000nM MFP is used. The reason for the background is primarily contamination of the substrate sample with fluorescent end product, 3-o-methylfluorescein. Potentially this high background fluorescence might cause difficulty visualizing single molecule reaction, which will be hidden behind the background noise. One of the ways to solve this problem is to photobleach the area on the microscopic slide prior to observing the single molecule reaction. Sample Preparation for Ensemble Averaging Kinetic Measurements All the reactions were performed in 100 mm Glycine buffer (Sigma Aldrich, cat. # 50046) supplemented with 1mM MgCl 2 *6H 2 O (Sigma Aldrich, cat. # M9272) and 1mM ZnCl 2 (Sigma Aldrich, cat. # ). The buffer was prepared by dissolving g of ZnCl 2, 0.1 g of MgCl 2 *6H 2 O and g of Glycine in 50 ml of deionized water. The ph of the buffer was adjusted to 9 by 1M NaOH solution. The buffer was prepared fresh every time.

45 34 Alkaline phosphatase was purchased from Sigma Aldrich (cat. # P4252). For the experiments, the stock solution of the enzyme was prepared to taking 1μl of the enzyme suspension and dissolving it in glycine reaction buffer up to the total volume of 5ml in glycine buffer (final concentration 7e-9M). The substrate for the reaction was purchased from Sigma Aldrich (3-o-methylfluorescein phosphate cyclohexylammonium salt, cat.# M2629). The stock solution was prepared in DMSO solvent with the final concentration being 0.1e-3M. Prior to performing kinetic measurements, the fluorimeter (PTI Instruments, LPS 220B) was calibrated with different concentration of reaction product fluorescein (fluorescein has the same spectral characteristics as 3-o-methylfluorescein) to directly obtain time dependent concentration values for product generation in the kinetic measurements. The stock of fluorescein solution (1e- 6M in Glycine reaction buffer) was diluted to concentrations in the range between 0 nm to 500 nm and emission measured in calibration mode. The reaction mixture for kinetic measurements was prepared for different concentrations of MFP substrate ranging from 500 nm to nm. The AP concentration was kept constant at 1e-11M for all samples. For example, a 500nM MFP sample was prepared by combining 10μl of MFP stock with 3μl AP stock and 1990 μl of Glycine reaction buffer, 1000nM MFP sample was prepared by combining 20 μl of MFP stock with 3 μl AP stock and 1980 μl of Glycine reaction buffer and so on. These reagents are combined in a fluorimetric disposable cuvette, mixed gently with pipette and placed into the fluorimeter apparatus. Each reaction was monitored for 15 minutes in Timebased mode using emission wavelength of 512 nm. The control that had no AP added was also monitored for 15 minutes for each reaction. The intensity values are converted to concentration values for the product automatically by the software using the fluorescein

46 35 calibration curve. The resulting concentration vs. time curve is then analyzed in Excel by finding the slope of the linear phase (first 10 minutes of the reaction) and subsequently plotting the slope of each concentration vs. time curve vs. the concentration of substrate to finally obtain Michaelis-Menten curve for alkaline phosphatase shown in figure 15. Single Molecule Studies of Alkaline Phosphatase: Apparatus and Sample Preparation Single molecule experiments with alkaline phosphatase were performed using apparatus in total internal reflection configuration with two channels for detecting two light polarizations shown in figure 20. The light source is a 473 nm laser. The excitation polarizer passes s- polarized light at 473 nm and above (green). The excitation filter (Chroma Technology Z470/20X) passes excitation light at 473 nm in the range of 20 nm. Dichroic beam splitter (Chroma Technology Z470/20X) reflects polarized excitation light (green) at 473 nm and above. The microscope (objective) used in this apparatus is Zeiss axiovert 200M. The emission filter (Chroma Technology HQ500LP) passes depolarized emission light at 500 nm and above (orange). Polarizing beam splitter separates emission light in two channels: s-polarization (solid line) and p-polarization (dotted line). Finally, the light is focused onto s regions of the CCD camera by 2 mirrors and 2 focus lenses (2 channels).

47 36 Figure 20. Two channel (two polarizations) TIRF microscope for single molecule studies of alkaline phosphatase.

48 37 The sample was prepared in agarose gel as shown in figure 21. First, 1e-8 M alkaline phosphatase was spincoated on the clean cover glass. Then, 1% agarose (Sigma Aldrich, cat. #A9414, low gelling temperature agarose) in Glycine reaction buffer and 50 nm MFP was applied onto the spincoated alkaline phosphatase. Glycine buffer was applied on top of the agarose to prevent gel from drying out. The reaction started to be visible by CCD camera (ProEM CCD, Princeton Instruments) after 10 minutes. Figure 21. Sample preparation for single molecule studies of alkaline phosphatase. Alkaline Phosphatase dimer (blue) spincoated onto the clean coverglass and embedded into the agarose (pink) containing MFP substrate to be converted into fluorescent 3-o-methylfluorescein product (yellow).

49 38 Single Molecule Studies of Alkaline Phosphatase: Experimental Results Prior to performing single molecule experiments, the TIRF apparatus was calibrated with 1nM fluorescein spincoated onto the clean coverglass to ensure that the set-up can successfully detect single molecule fluorescent product. The obtained imaging results are shown in figure 22. Intensity Levels max : 8222 min : 484 total : e+008 avg : sigma : Intensity Levels : max : 1480 min : 488 total : e+008 avg : sigma : Figure 22. Apparatus calibration for single molecule studies of alkaline phosphatase. Spincoated 1 nm fluorescein dye, dry (left). Clean cover glass, dry (right).

50 39 Raw single molecule trajectories for alkaline phosphatase are shown in figure 23. Each trajectory contains 500 frames, each frame 30 milliseconds long. Each trajectory was recorded from 3X3 pixel area where the fluorescent product generation was observed. Also, for each trajectory, background signal from 9x9 pixel area was recorded. Each trajectory was further analyzed in Excel. Figure 23. Sample trajectories of single molecule reaction for alkaline phosphatase. Each trajectory contains 500 frames (x axis), each frame is 30 milliseconds long. Fluorescence signal intensity is plotted on y axis.