SDSU Template, Version 11.1

NONLINEAR LASER WAVE-MIXING DETECTION FOR CAPILLARY

ELECTROPHORESIS AND MULTI-CHANNEL ARRAYS FOR

BIOMEDICAL AND ENVIRONMENTAL APPLICATIONS

______

A Thesis

Presented to the

Faculty of

San Diego State University

______

In Partial Fulfillment

of the Requirements for the Degree

Master of Science

Chemistry

______

Eric J. Maxwell

Spring 2015

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Eric J. Maxwell

DEDICATION

To my wife, Selena, for supporting me through all of the long nights and weekends of work that this program required. I cannot wait to start the next chapter in this journey.

ABSTRACT OF THE THESIS

Nonlinear Laser Wave-Mixing Detection for Capillary Electrophoresis and Multi-Channel Arrays for Biomedical and Environmental Applications by Eric J. Maxwell Master of Science in Chemistry San Diego State University, 2015

Degenerate Four-Wave Mixing is demonstrated as a highly sensitive nonlinear spectroscopic detection method for biomedical and environmental targets. This is achieved through refractive index change within an absorbing liquid medium, which produces a laser- like signal beam. This signal has high spatial resolution, and may be collected with high efficiency against a nearly 100% dark background. The cubic dependence on laser power and square dependence on analyte concentration allow for high signal intensity in trace analysis applications. In this work, the Degenerate Four-Wave Mixing technique is coupled with capillary electrophoresis, immunoprecipitation or color-forming reactions to provide specificity. The veterinary drugs malachite green and crystal violet are shown to be detectable at concentrations as low as 6.9 x 10-10 M (2.5 x 10-19 mol) and 8.3 x 10-11 M (3.0 x 10-20 mol) respectively (S/N = 2). Capillary electrophoresis is used in conjunction with a 2- laser Degenerate Four-Wave Mixing detector to allow simultaneous identification of both analytes. For another small molecule target, ammonium nitrate, sample preparation with diphenylamine is used to produce a colored compound capable of absorbing 635 nm light. This explosive component is of significant forensics interest due to its use in the manufacture of improvised explosive devices. The limit of detection for ammonium nitrate in 1 mm ID capillary cells is determined to be 1.5 x 10-9 M (5.2 x 10-18 mol). The detection limit in a 1.5 mm thin-film cell is found to be 3.2 x 10-7 M (2.0 x 10-15 mol) for S/N = 2. In addition to small molecule environmental targets, a detector for the cancer biomarker carcinoembryonic antigen is demonstrated with a combination of magnetic immunoprecipitation, multi-channel capillary arrays and Degenerate Four-Wave Mixing. Isolated protein samples are reacted with bicinchoninic acid (BCA) to yield a colored product, and absorbance of a 532nm laser produces a wave-mixing signal. Multiple samples may be processed rapidly though computer-controlled positioning of the multi-channel capillary array. The limit of detection for carcinoembryonic antigen is determined to be 3.3 x 10-12 M (0.59 ng/mL), with a corresponding mass detection limit of 1.2 x 10-21 mol (0.22 fg) for S/N = 2.

TABLE OF CONTENTS

PAGE

ABSTRACT ...... v

LIST OF TABLES ...... ix

LIST OF FIGURES ...... x

ACKNOWLEDGEMENTS ...... xiv

CHAPTER

1 INTRODUCTION ...... 1 1.1. Detection of Biomedical and Environmental Targets ...... 1 1.1.1. Motivation ...... 1 1.1.2. Spectroscopic Detection Methods...... 2 1.1.3. Lasers ...... 4 1.1.4. Laser Spectroscopic Methods ...... 4 1.2. Nonlinear Wave-Mixing ...... 5 1.3. History of Nonlinear Wave Mixing ...... 6 1.4. Thesis Outline ...... 8 2 FOUR-WAVE MIXING SPECTROSCOPY ...... 9 2.1. Overview ...... 9 2.1.1. Backward-Scattering Degenerate Four-Wave Mixing...... 9 2.1.2. Forward-Scattering Degenerate Four-Wave Mixing ...... 10 2.2. Interference Patterns ...... 10 2.3. Laser-Induced Dynamic Gratings ...... 12 2.4. Wave-Mixing Signal ...... 14 2.4.1. Signal Intensity ...... 16

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2.4.2. Signal Collection ...... 17 3 DETECTION OF MALACHITE GREEN AND CRYSTAL VIOLET BY DEGENERATE FOUR-WAVE MIXING AND CAPILLARY ELECTROPHORESIS ...... 19 3.1. Abstract ...... 19 3.2. Introduction ...... 20 3.3. Experimental ...... 23 3.3.1. Chemicals ...... 23 3.3.2. Degenerate Four-Wave Mixing Apparatus ...... 23 3.3.3. Capillary Electrophoresis System ...... 25 3.3.4. Capillary Treatment ...... 27 3.4. Results and Discussion ...... 29 3.4.1. Continuous Flow Detection ...... 29 3.4.2. Background Electrolytes ...... 30 3.4.3. Field Amplified Sample Stacking ...... 36 3.4.4. Bare Capillary Separation ...... 39 3.4.5. Effects of Capillary Dynamic Coating ...... 41 3.4.6. Detection Limits and Linearity ...... 43 3.5. Conclusions ...... 45 4 AMMONIUM NITRATE EXPLOSIVE SCREENING USING DEGENERATE FOUR-WAVE MIXING SPECTROSCOPY ...... 48 4.1. Abstract ...... 48 4.2. Introduction ...... 48 4.3. Experimental ...... 50 4.3.1. Chemicals ...... 50 4.3.2. Degenerate Four-Wave Mixing Optical Setup ...... 50 4.3.3. Capillary Sample Cells ...... 52 4.3.4. Thin-Film Sample Cells ...... 52 4.3.5. Nitrate Derivatization...... 55 4.4. Results and Discussion ...... 58 4.4.1. Nitrate Derivatization Kinetics ...... 58 4.4.2. Detection in Capillary Cells ...... 60 4.4.3. Detection in Thin-Film Cells ...... 66

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4.4.4. Linearity of Response ...... 69 4.5. Conclusions ...... 71 5 SENSITIVE DETECTION OF CARCINOEMBRYONIC ANTIGEN BY DEGENERATE FOUR-WAVE MIXING SPECTROSCOPY ...... 74 5.1. Abstract ...... 74 5.2. Introduction ...... 74 5.2.1. Degenerate Four-Wave Mixing ...... 76 5.2.2. Bradford Protein Assay ...... 77 5.2.3. BCA Protein Assay ...... 77 5.3. Experimental ...... 79 5.3.1. Chemicals ...... 79 5.3.2. Degenerate Four-Wave Mixing Optical Setup ...... 81 5.3.3. Multichannel Capillary Array ...... 81 5.3.4. Linear Actuator Control ...... 83 5.3.5. Magnetic Immunoprecipitation Protocol ...... 85 5.3.6. HPLC Protocol ...... 85 5.3.7. Bradford Protein Assay Protocol ...... 88 5.3.8. BCA Protein Assay Protocol ...... 88 5.4. Results and Discussion ...... 89 5.4.1. Bradford and BCA Assays ...... 89 5.4.2. Magnetic Immunoprecipitation ...... 92 5.4.3. Multi-Channel Array Scanning Modes ...... 96 5.4.4. Linearity of Response ...... 100 5.4.5. Limit of Detection ...... 102 5.5. Conclusions ...... 104 REFERENCES ...... 106

LIST OF TABLES

PAGE

Table 3.1. Detection Limits for Crystal Violet Analysis Methods...... 47 Table 3.2. Detection Limits for Malachite Green Analysis Methods...... 47 Table 4.1. Detection Limits for Ammonium Nitrate in Varied Sample Cells...... 73 Table 4.2. Detection Limits for Various Ammonium or Nitrate Ion Analysis Methods...... 73 Table 5.1. Detection Limits for CEA Analysis Methods...... 105

LIST OF FIGURES

PAGE

Figure 1.1. Electron energy state changes in spectroscopy...... 3 Figure 2.1. Backward-Scattering Degenerate Four-Wave Mixing (A) and Forward- Scattering Degenerate Four-Wave Mixing (B)...... 11

Figure 2.2. Interference grating formed by two light waves of intensity I1 and I2, with wave vectors k1 and k2 intersecting to form an interference pattern...... 13

Figure 2.3. Intersection of pump beams (E1 & E2) within an absorbing medium to generate a wave mixing signal E4 with wave vector k4...... 15 Figure 2.4. Spatial orientation of Forward Degenerate Four-Wave Mixing pump/probe beams (E1\E3, E2\E3) and signal beams (E4)...... 18 Figure 3.1. Structures of malachite green and crystal violet...... 21 Figure 3.2. UV-visible spectra of malachite green (1.0 µM) and crystal violet (1.0 µM)...... 24 Figure 3.3. Experimental setup for wave-mixing detection of malachite green and crystal violet. 532 nm and 635 nm lasers are used for simultaneous detection. S1, S2: Beam splitters; B1, B2: Beam blockers; L1, L2: Lenses...... 26 Figure 3.4. Wave-Mixing Detection for Capillary Electrophoresis...... 28 Figure 3.5. 5.0 x 10-7 M malachite green in acetonitrile. 360 μm OD/75 μm ID capillary. A: pump beams blocked; B: pump beams unblocked; C: non- modulated beam blocked...... 31 Figure 3.6. 1.0 x 10-7 M crystal violet in methanol. 360 μm OD/75 μm ID capillary. A: pump beams blocked; B: pump beams unblocked; C: non-modulated beam blocked...... 32 Figure 3.7. 2.5 x 10-5 M crystal violet (CV) in 50 mM ammonium acetate. Capillary electrophoresis parameters: +20 kV, 360 μm OD/75 μm ID x 40 cm capillary (20 cm effective length), 30 s electrokinetic injection...... 33 Figure 3.8. 1 x 10-4 M crystal violet (CV) in 50 mM citric acid (A). 1 x 10-4 M CV in 50 mM citric acid diluted with acetonitrile (1:2) (B). Capillary electrophoresis parameters: +20 kV, 360 μm OD/75 μm ID x 40 cm capillary (20 cm effective length); 30 s (A) and 5 s (B) electrokinetic injections...... 35

Figure 3.9. Field Amplified Sample Stacking (FASS). E: Electric field...... 37 Figure 3.10. Field Amplified Sample Stacking. 10 μM CV in 50 mM aqueous citric acid/acetonitrile (1:2) (Left). 10 μM CV in 0.5 mM aqueous citric acid / acetonitrile (1:2) stacking buffer (Right). Capillary electrophoresis parameters: +20 kV, 360 μm OD/75 μm ID x 40 cm capillary (20 cm effective length); 5 s electrokinetic injection...... 38 Figure 3.11. (A) Electropherogram of a mixture of 10 μM MG and 10 μM CV in 0.5 mM aqueous citric acid/acetonitrile (1:2) stacking buffer. (B) Migration times for repeated injections of the same sample mixture in (A); MG (○), CV (◊). Capillary electrophoresis parameters: +20 kV, 360 μm OD/100 μm ID x 35 cm capillary (17 cm effective length), 10 s electrokinetic injection...... 40 Figure 3.12. Electropherograms of a mixture of 10 μM MG and 10 μM CV in 0.5 mM aqueous citric acid/acetonitrile (1:2) stacking buffer. (Top) Bare fused silica capillary (Bottom) Ultratrol LN dynamic capillary coating. Capillary electrophoresis parameters: +20 kV, 360 μm OD/100 μm ID x 35 cm capillary (17 cm effective length), 3 s electrokinetic injection...... 42 Figure 3.13. Electropherograms of 2.0 x 10-10 M MG and 2.0 x 10-9 CV in 0.5 mM aqueous citric acid/acetonitrile (1:2) stacking buffer. Capillary electrophoresis parameters: +20 kV, 360 μm OD/100 μm ID x 35 cm capillary (17 cm effective length); 15 s electrokinetic injection...... 44 Figure 3.14. Signal vs. concentration of CV (A). log(signal) vs. log(concentration) of CV (B). Signal vs. concentration of MG (C). log(signal) vs. log(concentration) of MG (D). 360 μm OD/75 μm ID x 40 cm capillary...... 46 Figure 4.1. UV-visible spectra of 0.1 M ammonium nitrate (A) and the product of the reaction of ammonium nitrate (1.0 x 10-5 M) and 4.5 mM diphenylamine in 13.8 M sulfuric acid (B)...... 51 Figure 4.2. Experimental setup for wave-mixing detection of ammonium nitrate. S1: Beam splitter; B1, B2: Beam blockers; L1, L2: Lenses...... 53 Figure 4.3. Ammonium nitrate capillary sample cell...... 54 Figure 4.4. Ammonium nitrate thin-film sample cells...... 56 Figure 4.5. Diphenylbenzidine derivative produced through reaction of diphenylamine with ammonium nitrate and sulfuric acid...... 57 Figure 4.6. (A) Absorbance at 620 nm for the product of ammonium nitrate (2.5 x 10- 4 M) reacted with 4.5 mM diphenylamine in 13.8M sulfuric acid at 25 °C (1:10 dilution). (B) Absorbance of ammonium nitrate (2.5 x 10-4 M) reacted with 4.5 mM diphenylamine in 13.8 M sulfuric acid at 25 °C, 45 °C and 60 °C for 1 minute (1:10 dilution)...... 59 Figure 4.7. Ammonium nitrate (2.5 x 10-5 M) in diphenylamine reagent. 75 µm ID/360 µm OD capillary. A: pump beams blocked; B: pump beams unblocked; C: non-modulated beam blocked...... 61

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Figure 4.8. Ammonium nitrate (5.0 x 10-6 M) in diphenylamine reagent. 360 µm ID/1000 µm OD capillary. A: pump beams blocked; B: pump beams unblocked; C: non- modulated beam blocked...... 64 Figure 4.9. Ammonium nitrate (1.0 x 10-8 M) in diphenylamine reagent. 1 mm ID/1.5 mm OD capillary. A: pump beams blocked; B: pump beams unblocked; C: non- modulated beam blocked...... 65 Figure 4.10. Ammonium nitrate (1.0 x 10-5 M) in diphenylamine reagent (left) and diphenylamine reagent blank (right). 6.5 µm thin-film cell. A: pump beams blocked; B: pump beams unblocked; C: non-modulated beam blocked...... 68 Figure 4.11. Ammonium nitrate (1.0 x 10-5 M) in diphenylamine reagent (left) and diphenylamine reagent blank (right). 1500 µm thin-film cell. A: pump beams blocked; B: pump beams unblocked; C: non-modulated beam blocked...... 70 Figure 4.12. Logarithmic plot of ammonium nitrate concentration (12.5 to 100 µM) versus relative wave-mixing signal...... 72 Figure 5.1. Bradford protein assay reaction. Nitrogen deprotonation results in a color change from red to blue...... 78 Figure 5.2. Two-step BCA protein assay reaction. Reduction of Cu2+ to Cu+ via the Biuret Reaction allows formation of the BCA-Cu+ chelate...... 80 Figure 5.3. Experimental setup for wave-mixing detection of carcinoembryonic antigen. 532 nm Nd:YAG or 635 nm diode laser. S1: Beam splitter; B1, B2: Beam blockers; L1, L2: Lenses...... 82 Figure 5.4. Capillary array sample cell and actuator...... 84 Figure 5.5. JavaScript for delayed four-position movement of Zaber TLA-28 linear actuator used for the capillary array...... 86 Figure 5.6. Magnetic immunoprecipitation procedure for isolation of CEA from a heterogeneous protein solution...... 87 Figure 5.7. UV-visible spectra of Bradford reagent alone and with BSA (100 µg/mL)...... 90 Figure 5.8. UV-visible spectra of BCA reagent alone and with 2 to 10 µg/mL BSA...... 91 Figure 5.9. UV-visible spectra of BCA reagent alone and with 10 µg/mL CEA...... 93 Figure. 5.10. Size exclusion (SEC)-HPLC chromatograms for molecular weight standards (465, 310, 155, 67 and 29 kDa) (A), CEA standard (B), magnetic-IP purified CEA (C) and magnetic-IP control (D)...... 95 Figure 5.11. Continuous vertical scan of capillary array containing 270 µg/mL BSA- BCA. 75 µm ID/360 µm OD fused silica capillaries. 25 µm/s scan rate...... 97 Figure 5.12. Three-position scan of capillary array containing 200 µg/mL BSA-BCA. 75 µm ID/360 µm OD fused silica capillaries...... 99 Figure 5.13. Signal vs. concentration of BSA (left). log(signal) vs. log(concentration) of BSA (right). 360 μm OD/75 μm ID x 40 cm capillary array...... 101

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Figure 5.14. Wave-mixing signal response for 20 ng/mL CEA reacted with the BCA Reagent. 150 µm ID/360 µm OD fused silica capillary. A: pump beams blocked; B: pump beams unblocked; C: non-modulated beam blocked...... 103

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ACKNOWLEDGEMENTS

Thank you to Dr. Tong and the entire San Diego State University Chemistry

Department. I had high expectations coming into this program, and I can honestly say I ended up getting more out of it than I thought I would. I greatly appreciate the knowledge and skills that I have picked up along the way, as well as the flexibility the program allows for working professionals. I know many other chemistry graduate programs would not have allowed me to work full-time while being pursuing this degree.

Manna and Marcel, thank you both for your friendship and all of the troubleshooting advice over the past few years. It helped get me started early on, and it helped get some projects get through rough patches. As the three of us graduate, I hope the work we produced will make a positive impact on the future direction of the group.

Finally, I want to thank my managers, Adriana and Emily, at Alere San Diego.

Completing this program would not have been possible without you two allowing me to work modified schedules to plan around classes and research. At times it was tough to balance it all, but I know the end result will be extremely rewarding.

CHAPTER 1

INTRODUCTION

1.1. DETECTION OF BIOMEDICAL AND ENVIRONMENTAL TARGETS

1.1.1. Motivation

The field of analytical spectroscopy is critical to research performed in numerous sciences, including chemistry, biology and physics. A wealth of information can be obtained by utilizing spectroscopic techniques, and they have historically been used for a vast array of inorganic and organic samples. These techniques often offer relatively simple sample preparation, along with the capability to generate significant qualitative and quantitative information. Modern medicine relies heavily on diagnostic assays to diagnose, treat and monitor diseases and other health conditions. Other applications can have an environmental focus such as analysis of toxic or explosive compounds. At the heart of almost any diagnostic assay is a spectroscopic detection method. This can range from simple colormetric techniques to advanced mass spectrometry. These methods are designed to take what cannot be seen and turn it into something tangible, and the information that can be obtained gives an understanding of our surroundings that would normally be beyond the capability of human beings. Spectroscopic detection methods allow us to see more than what is possible through physiology alone.

1.1.2. Spectroscopic Detection Methods

Many spectroscopic methods exist for the detection of single atoms, small molecules and proteins (Figure 1.1). A large number of these methods make use of atomic absorption, which is the excitation of an atom or molecule through absorption of photons. This absorption causes a temporary transition to an excited energy state, followed by relaxation with the release of energy. Absorption of specific wavelengths of light can determine the identity and concentration of a compound through change in light intensity, temperature or refractive index (1).

Fluorescence is relaxation that releases energy in the form of radiation. Incident radiation is used to stimulate emission from a sample capable of fluorescence. The emitted light is typically of lower energy than the input radiation, which is referred to as a Stokes emission. But the opposite can also be observed under certain circumstances; this is called anti-Stokes emission. In either case, photons can be collected to generate a signal with intensity relative to concentration. Fluorescence detection methods can also provide good specificity due to the limited number of fluorophores found in nature (2).

Atomic emission is also a form of relaxation that releases energy in the form of radiation. However, atomic emission differs from fluorescence in that excitation is brought on by thermal or electrical stimulation rather than by radiation. Energy is introduced by an atomizer to excite atoms, which then emit photons at a characteristic wavelength as they return to their ground state. For elemental analysis, the specific wavelength of the emitted light can be used to identify atoms present in a sample. This type of analysis is particularly useful for detection of metal ions such as sodium or calcium (1).

Ionization can also be useful for analysis of atomic, small molecule and protein samples. Energy is introduced into the system by a high power-density light source

(photoionization) or electrical/thermal sources (collisional ionization). The charged ions that are released in the process can be monitored by electrodes within a sample cell.

Figure 1.1. Electron energy state changes in spectroscopy.

Changes in current are directly proportional to concentration. Ionization also plays a large role in mass spectrometry, which requires charged species. A variety of detector setups are used to measure the mass-to-charge (m/z) ratio, which can provide both quantitative and qualitative data (3).

1.1.3. Lasers

Lasers are a unique light source. Since the first laser was developed in 1960 (4), they have become a powerful tool in the area of analytical spectroscopy. Lasers possess many beneficial properties that conventional lamp-based light sources do not, which opens doors to new possibilities. Tunability of laser wavelength using a variety of nonlinear optics allows for the production of radiation from ultraviolet to infrared (0.2 nm to 40 µm). Because of this wide range of available laser sources, they can be applied to many applications that are not possible with conventional lamps. Lasers can also possess intensity that is orders of magnitude greater than other sources. This can be used to greatly improve sensitivity of detection methods, while also allowing for techniques such as ionization, optical saturation and excitation of forbidden transitions. Most conventional light sources are not capable of delivering the power density required to produce these effects. The spectral linewidth of a laser is also narrower than that of atomic or molecular emissions, which allows for efficient and selective excitation of targets. The beam produced has high spatial coherence, and may be focused as small as a few micrometers. This also allows the beam to be reduced, expanded or collimated with only minor loss of photons in the form of stray light (5).

1.1.4. Laser Spectroscopic Methods

Lasers can be used as a light source for many different analytical spectroscopic techniques. Some of the more common methods are described in this section. Conventional

5 absorption methods may utilize lasers in place of conventional light sources (e.g. lamps) to take advantage of the temporal and spatial coherence properties. The same can be true for methods that rely on stimulated emission. Laser-Excited Atomic Fluorescence (LEAF)

Spectroscopy uses lasers tuned to a resonance frequency of atomized samples to generate fluorescence. Laser Induced Fluorescence (LIF) utilizes laser light in a similar manner, except atomization is not required. LIF is commonly used with liquid samples, including proteins labeled with small molecule fluorophores. Lasers also make a number of ionization techniques possible. Laser Desorption Mass Spectrometry and Matrix Assisted Laser

Desorption Ionization (MALDI) Mass Spectrometry use thermal energy generated by lasers to cause desorption and ionization of samples (5).

1.2. NONLINEAR WAVE-MIXING

The spectroscopic methods described thus far are commonly referred to as “linear”.

This is due to the fact that they involve the interaction of matter with a single electromagnetic field. In contrast, methods that involve two or more electromagnetic fields are referred to as

“nonlinear”. In nonlinear spectroscopy, the relationship between analyte and signal response can be more complex. By definition, a response curve will follow a nonlinear trend. While this can make interpretation of results less clear, nonlinear response is a significant advantage. When compared to signal response of linear methods, nonlinear spectroscopy generally offers superior sensitivity for trace detection. Within this category of spectroscopy, there are a large number of different techniques. Nonlinear Wave Mixing includes three- wave and four-wave mixing arrangements, but Degenerate Four-Wave Mixing (DFWM) is one of the most sensitive and versatile variations. DFWM can use relatively low-power

(mW) laser sources for the detection of trace levels of ions, small molecules and proteins in

6 their native form (6). This subject is covered in greater detail in Chapter 2, which focuses on the theory and application of Four-Wave Mixing.

1.3. HISTORY OF NONLINEAR WAVE MIXING

In 1972, Boris Zeldovich et al. (7, 8) demonstrated that Stimulated Brillouin

Scattering (SBS) with a ruby laser could be used to create an amplified, phase conjugated wave in a nonlinear medium. Acoustic phonons were used to create refractive index changes within the medium, which allowed a portion of the laser to be reflected at a slightly lower frequency. Remarkably, when the input beam was intentionally distorted prior to reaching the nonlinear medium, the reflected wave was found to be nearly undistorted. This is referred to as a phase conjugate mirror, which allows for a time-reversed wave to be created from an input wave.

In 1976, Amnon Yariv (9) proposed a three-wave mixing technique that was designed to overcome the problem of frequency change encountered by Zeldovich. This method did not use SBS, and instead used parametric mixing of light waves in an acentric crystal to induce phase conjugation. While this new methodology succeeded in avoiding frequency change, it had significant limitations in terms of beam-acceptance angles and phase-matching requirements. A year later, Yariv and David Pepper proposed Degenerate Four-Wave Mixing

(DFWM) (10). For DFWM, two counter-propagating pump waves were allowed to meet within a non-resonant optical medium. A third wave of the same frequency was directed into the medium at an arbitrary angle, with the result being a phase conjugated wave reflected back along the same path. At the same time, Robert Hellwarth developed a similar DFWM method with a single laser providing both of the pump waves, as well as the probe wave (11).

Hellwarth also demonstrated that phase matching was ensured for all angles, which made the process attractive for holography applications requiring wide fields of view.

Up to this point, optical phase conjugation techniques all used input waves with frequencies that did not correspond to any resonant electronic transition within the nonlinear media. That changed in 1978 when Abrams and Lind published a letter extending the analysis to absorbing nonlinear media (12, 13). With an absorbing system, the counter- propagating beam created by phase conjugation was found to contain quantitative information about the media and the pump intensity. Additionally, absorption served to amplify the intensity of the wave greatly. This proved to be a major breakthrough since the technique could not only provide qualitative information for holography, but it could also be further developed as a spectroscopic tool. In the following years, the technique was extended to kinetic studies (14), and the impact of laser and absorbing media changes on wave mixing intensity was explored (15, 16).

In 1987, the first instances of DWFM as a quantitative detection method for analytical spectroscopy were published (17, 18). In these works, a very low detection limit was achieved when DFWM was used as a detector for elemental analysis of sodium in an analytical flame. These initial works were followed by additional DWFM analysis using an analytical flame (19-21), as well as others using a hollow cathode discharge (22-25), a continuously flowing liquid cell (6, 26, 27) and liquid chromatography (28).

While numerous applications for DWFM have been devised using the “backward- scattering” experimental geometry described by Yariv and Hellwarth, other alignments are capable of creating a wave-mixing signal. In 1993, Zhiqiang Wu and William Tong introduced a “Forward-Scattering” Degenerate Four-Wave Mixing (F-DWFM) geometry as a

8 modification of the original spectroscopic technique (29). While the Backward-Scattering arrangement (B-DFWM) requires two counter-propagating pump beams and a separate probe beam, F-DFWM only uses two beams. Alignment is simplified significantly by the reduction in the number of beams, and increases in wave-mixing efficiency allow for lower power requirements than that of B-DFWM. Works using F-DWFM detection have successfully paired it with continuously flowing liquid cells (30) and capillary electrophoresis (31) with high sensitivity.

1.4. THESIS OUTLINE

The following chapters demonstrate DFWM as a spectroscopic detection method for use with biological and environmental small molecule and protein targets. Chapter 2 details the theoretical aspects of DFWM and explains the nature of wave-mixing signal diffraction.

Chapter 3 presents a method for detection of the small molecule veterinary drugs, malachite green and crystal violet, through the coupling of F-DFWM with capillary zone electrophoresis. The inherent optical absorbance of these compounds allows for direct measurement without the use of any molecular labels. Chapter 4 describes a means of detecting trace quantities of nitrate-based explosives using F-DFWM with both capillary liquid cells and thin-film cells. A color-forming agent is used to induce absorption of visible light from naturally colorless samples. Chapter 5 presents an F-DFWM multi-channel capillary array system for analysis of protein biomarkers. The cancer biomarker carcinoembyronic antigen (CEA) is detected through the use of the copper-based bicinchoninic acid (BCA) protein assay and magnetic immunoprecipitation.

CHAPTER 2

FOUR-WAVE MIXING SPECTROSCOPY

2.1. OVERVIEW

Four-wave mixing spectroscopy is a broad subject that includes a large number of nonlinear experimental configurations all of which are centered on the concept of three light waves interacting to produce a fourth wave. Several Raman spectroscopic techniques are included under this umbrella, including Coherent Anti-Stokes Raman Spectroscopy (CARS).

CARS is an example of a four-wave mixing technique that uses light sources of two different frequencies (ω1 and ω2) to produce a signal. Two photons of ω1 interact with one photon of

ω2, where ω1 is greater than ω2, to generate a signal of ωs = 2ω1 – ω2 (32). In contrast,

Degenerate Four-Wave Mixing techniques use three light waves of the same frequency to form a fourth wave that shares the same frequency as the original three (ω1 = ω2 = ω3 = ω4).

This type of Four-Wave Mixing is further broken down into two methods: Backward-

Scattering Degenerate Four-Wave Mixing (B-DFWM) and Forward-Scattering Degenerate

Four-Wave Mixing (F-DFWM). This work will focus on F-DFWM; however, the underlying principles of signal formation are the same.

2.1.1. Backward-Scattering Degenerate Four-Wave Mixing

Backward-Scattering Degenerate Four-Wave Mixing (B-DFWM) operates on the basis of optical phase conjugation, which is the reversal of the temporal and spatial

10 development of a wave (32). The experimental configuration consists of two phase-matched, counter-propagating waves directed to meet head-on within an absorbing medium, while a third wave (a probe beam) is directed through the intersection point at a small angle. When the probe beam reaches the intersection point, a fourth wave is created and reflected back in the opposite direction (6, 19, 32, 33). This setup is depicted in Figure 2.1.

2.1.2. Forward-Scattering Degenerate Four-Wave Mixing

In an F-DFWM experimental setup, a single wave is split into two pump waves and directed through a focusing lens. The absorbing medium is placed at the focal point of the lens to allow convergence at the sample. Interaction of the waves within the medium results in the formation of two additional waves at angles equivalent to the angle of separation between the pump waves. Unlike B-DFWM, there is not a separate third wave source.

Photons from either pump wave also play the role of the third beam, which results in a fourth wave being generated (34-37). Figure 2.1 shows the optical arrangement of an F-DFWM experiment.

2.2. INTERFERENCE PATTERNS

When two light waves intersect, a pattern of constructive and destructive interference is created. The grating vector K associated with this interference pattern may be described as

(2.1) K = ± (k1 – k2) where k1 and k2 are the propagation vectors of the intersecting waves. The period Λ of this spatially modulated light field may be described as

2휋 (2.2) 훬 = 퐾

Figure 2.1. Backward-Scattering Degenerate Four-Wave Mixing (A) and Forward- Scattering Degenerate Four-Wave Mixing (B).

The equation for the grating vector period may also be expressed in terms of the pump beam wavelength λ and intersection angle θ.

휆 훬 = (2.3) 2sin(휃/2)

For small intersection angles (θ < 1), Equation 2.3 may be simplified further as (33, 38)

휆 훬 ≈ (2.4) 휃

Figure 2.2 illustrates the arrangement of the interacting light waves and the interference pattern that is formed. From these equations, it is evident that the period of the interference pattern is directly proportional to the wavelength of the waves and the angle between them.

2.3. LASER-INDUCED DYNAMIC GRATINGS

The formation of laser-induced gratings can be initiated by a number of mechanisms depending on the type of medium. In liquid samples, thermal gratings, as a result of Dufour and Soret effects, contribute most significantly (34, 39). When an interference pattern is created within an absorbing liquid medium, absorption of photons leads to excitation of the analyte, which initially forms a population density grating. This population of molecules in the excited state can return to their ground state through non-radiative decay. This type of decay transfers energy in the form of heat to the surrounding medium. The heat transfer causes thermal expansion of the sample matrix, which results in changes in the refractive index, which in turn form a thermal grating. This newly formed thermal grating has a period

Λ and propagation vector K that are determined by the period of the interference pattern formed by the pump waves. The thermal grating that forms as a result of refractive index change within the interference pattern may then act as a stationary phase grating for wave- mixing (33-35).

Figure 2.2. Interference grating formed by two light waves of intensity I1 and I2, with wave vectors k1 and k2 intersecting to form an interference pattern.

2.4. WAVE-MIXING SIGNAL

In a Forward-Scattering Degenerate Four-Wave Mixing configuration, two pump beams intersect and interact within an absorbing medium to create a thermal grating. Once the grating has been established, a portion of the photons from each pump beam will diffract off of the grating at an angle defined by rearrangement of the Bragg equation (35, 40):

휆 휃 = 푠푖푛−1 ( ) = 휃 (2.5) 푠푖푔푛푎푙 2훬

Accordingly, Figure 2.3 shows the angle of the signal beams is the same as the angle of incidence between the pump beams (E1 & E2). Given that either pump beam may act as a probe, the signal beams that are formed are mirror images of one another within the same plane. Under Bragg conditions, both energy and momentum input into the system by the pump beams must be conserved, which requires that

(2.6) ω4 = ω1 + ω3 – ω2 where ω4 is the frequency of either signal beam, and ω1, ω2 and ω3 are that of the pump and probe beams. Since all input waves are of the same frequency, the resulting signal wave must also have this frequency to satisfy the conservation of energy requirement. Similarly, the phase vectors associated with each signal beam are defined by

(2.7) k4 = k1 + k3 – k2 where k4 denotes the most effective propagation vector for either signal with respect to the pump and probe inputs (k1, k2 & k3). Conservation of momentum is satisfied in this regard; however, minor differences in the phases of the interacting waves may result in some deviation from the ideal propagation vector (Figure 2.3).

Figure 2.3. Intersection of pump beams (E1 & E2) within an absorbing medium to generate a wave mixing signal E4 with wave vector k4.

This phase mismatch Δk may be expressed as

(2.8) Δk = k1 + k3 – k2 – k4

This difference in phase can be influenced by the interaction length within the nonlinear medium. The wave-mixing signal is maximized when Δk = 0; however, this is difficult to achieve (34, 41).

2.4.1. Signal Intensity

The primary advantage nonlinear wave-mixing has over linear spectroscopic methods is the strength of the signal that may be formed in the presence of an absorbing medium. The

F-DFWM signal has a square dependence on analyte absorptivity and a cubic dependence on laser power, as shown by Equation 2.9 (34-37).

푏 2 휆 푑푛 2 훼2 퐼 = ( ) 퐼2퐼 푒 ( ) (2.9) 푠 8휋 2 1 sin4(휃⁄2) 푑푡 휅2

Signal intensity is represented by Is, and the pump beam intensities are assigned the identifiers I1 and I2. b is the optical path length of the sample cell, λe is the excitation wavelength, θ is the angle between the pump beams, and dn/dt is the change in the refractive index with respect to temperature (due to the thermal grating). α is the absorptivity/extinction coefficient of the analyte, and κ is the solvent thermal conductivity. For weakly absorbing analytes, signal intensity is most easily enhanced by increases in laser power. In the opposite case, with strongly absorbing analytes, a very strong wave-mixing signal may be generated even with a relatively low-power laser source. This tunable nature of the technique and strong nonlinear response allow for a great deal of versatility in terms of analytes that can be detected.

2.4.2. Signal Collection

Isolation of an F-DFWM signal from the pump beams and any stray light created by diffraction of the laser beams off of an analyte sample cell can be accomplished through the use of specialized instrumentation. The signal itself can be given an induced frequency through the use of an optical chopper interfaced with a lock-in amplifier. The optical chopper physically blocks and unblocks a pump beam at a specified frequency, while the lock-in amplifier is used to separate the signal from background noise that does not have that frequency. Additionally, the optical chopper is ideally placed in the path of the pump/probe beam with the least power. If used to create frequency modulation in the stronger beam, a larger amount of stray light with the same frequency as the signal would be observed. The final step taken to reduce background noise is the use of spatial filters. Since the angle between all output beams is identical, the trajectory of a signal beam and position relative to each other beam can be predicted (Figure 2.4). This allows for optical alignment even with low-concentration analytes that do not produce a visible wave-mixing signal. Proper placement of spatial filters can prevent pump beams from reaching the signal collection point

(a photodetector) while allowing the signal to propagate unhindered.

Figure 2.4. Spatial orientation of Forward Degenerate Four-Wave Mixing pump/probe beams (E1\E3, E2\E3) and signal beams (E4).

CHAPTER 3

DETECTION OF MALACHITE GREEN AND CRYSTAL VIOLET BY

DEGENERATE FOUR-WAVE MIXING AND CAPILLARY

ELECTROPHORESIS

3.1. ABSTRACT

An ultrasensitive, label-free, antibody-free detection method for traces of malachite green (MG) and crystal violet (CV) using degenerate four-wave mixing laser spectroscopy

(DFWM) and capillary zone electrophoresis (CZE) is presented in this chapter. Wave-mixing spectroscopy provides a sensitive, absorption-based detection method for trace analytes. This is accomplished by creation of a dynamic grating within a sample cell, which diffracts light to create a coherent, laser-like signal beam with high optical efficiency and high signal-to- noise ratio. A cubic dependence on laser power and square dependence on analyte concentration make DFWM sensitive enough to detect molecules in their native form without the use of fluorescent labels for signal enhancement. A 200 mW 532 nm laser and a 100 mW

635 nm laser were used for malachite green and crystal violet sample excitation. The use of two lasers of different wavelengths allows the method to simultaneously detect both analytes.

Specificity is obtained through the capillary zone electrophoresis separation, which results in characteristic migration times. Measurement in capillary zone electrophoresis resulted in a limit of detection (LOD) of 6.9 x 10-10 M (2.5 x 10-19 mol) for crystal violet and 8.3 x 10-11 M

(3.0 x 10-20 mol) for malachite green at S/N of 2.

3.2. INTRODUCTION

Malachite green (Figure 3.1) is a triphenylmethane dye that may be used as a fungicide and paraciticide in aquaculture (42). Despite this beneficial application, it is not approved for use as an aquaculture veterinary drug in the United States, Canada or the

European Union (43, 44). Malachite green also has uses as a food coloring agent, as a medical disinfectant and as a dye for textiles (45). Although a wide range of uses exists, it has also been reported to have toxic effects that increase with concentration, exposure time and temperature. This toxicity may present itself in the form of organ damage and mutagenic, carcinogenic, and developmental abnormalities in mammals. Due to reports of this nature, the U.S. Food and Drug Administration (FDA) nominated malachite green as a priority chemical for carcinogenicity testing in 1993 (46).

Crystal violet (Figure 3.1) is another triphenylmethane dye used as an anti-microbial and anti-fungal agent for treatment of infections. It has a long history of medical use under the name Gentian Violet, but, like malachite green, crystal violet can be highly toxic and has been shown to enhance tumor growth (47, 48). Crystal violet also sees use as a DNA stain in gel electrophoresis (49), which may speak to the carcinogenicity of the compound. As is the case with malachite green, the FDA does not recognize crystal violet as safe for use as a veterinary drug (21CFR500.29 and 21CFR500.30) (47).

Due to these health risks and the potential for highly dilute samples, a very sensitive analytical detection method is needed to identify trace quantities in an aquaculture setting.

Both malachite green and crystal violet are readily absorbed into fish tissues, which may then be ingested by consumers.

Figure 3.1. Structures of malachite green and crystal violet.

Current detection methods often involve expensive, complicated techniques such as mass spectrometry to obtain low detection limits (50-56), while many of the simpler methods are not capable of measuring below nanomolar concentrations (57-60). This leaves a need for simple, yet sensitive, detection methods that can be field-deployed to confirm the presence of these hazards in water supplies or animal tissues. The method of detection described in this work utilizes absorptivity, molecular weight and charge to identify these drugs at sub- nanomolar concentrations by Degenerate Four-Wave Mixing coupled to capillary electrophoresis (CE).

As detailed in Chapter 2, Degenerate Four-Wave Mixing is a powerful spectroscopic method that produces a signal through the formation of a thermal diffraction grating at the intersection point of two input laser beams (pump and pump/probe) passing through an absorbing analyte. The angle of the input beams determines the angle of the two signal beams, which allows a predictable location for collection by a photodetector. Because the input beams are derived from one laser source split by a 70:30 beam splitter, the signal beam produced from the pump/probe beam is stronger than the one produced by the pump beam.

This signal has a square dependence on analyte absorptivity and a cubic dependence on laser power (29, 36, 61). These properties allow the method to take advantage of high power- density laser sources to excite molecules even if they do not have a large extinction coefficient. This allows a very dilute sample to produce a strong signal that is proportional to concentration.

Given that malachite green (MG) and crystal violet (CV) both yield strong optical absorption in the visible range (Figure 3.2), visible laser wavelengths closest to the peak absorption wavelengths will yield the strongest wave-mixing signals. For crystal violet, peak

23 absorption was determined to be at 580 nm with an extinction coefficient of approximately

160,000 cm-1 M-1. Malachite green peak absorption was determined to be at 618 nm with an extinction coefficient of approximately 140,000 cm-1 M-1. Using 532 nm and 635 nm lasers, the 532 nm excitation wavelength falls at 61% of crystal violet λmax and the 635 nm excitation wavelength at 68% of malachite green λmax . The broad absorption peak for malachite green actually allows a small amount of absorption from the 532 nm Nd:YAG laser, as well as the 635 nm diode laser used in this work. However, the same is not true for crystal violet, which does not show significant absorption near 635 nm.

3.3. EXPERIMENTAL

3.3.1. Chemicals

Malachite green chloride and crystal violet were purchased from Sigma Aldrich.

Ultratrol LN was purchased from Target Discoveries. All other chemicals were of analytical grade. Deionized water was distilled prior to use in capillary electrophoresis buffer solutions.

Buffers were filtered through 0.2 µm syringe filters prior to use as a sample diluent or in the capillary electrophoresis system.

3.3.2. Degenerate Four-Wave Mixing Apparatus

A Degenerate Four-Wave Mixing configuration described in our previous work (62) was utilized for analyte detection. A 200 mW 532 nm Nd-YAG laser (Changchun New

Industries Optoelectronics Tech Co. Ltd., MGL-FN-523-200mW) and a 100 mW 635nm diode laser (Laserglow Technologies, LRD-0635-TSR-00100-10) were positioned so that both beams converged at a 90-degree angle on a single 70T/30R (70% transmitted/30% reflected) beam splitter (Figure 3.3). This allowed approximately 70% of the 635 nm beam and 30% of the 532 nm beam to pass through into the optical setup.

Figure 3.2. UV-visible spectra of malachite green (1.0 µM) and crystal violet (1.0 µM).

Taking into account additional losses due to mirrors used to align the beams and spatial filters, the final effective laser powers were 65 mW and 40 mW respectively. After the initial power reduction, the two beams were directed to a second beam splitter (70R/30T) to create two pairs of overlapping beams, with the weak-side beams passing through an optical chopper (Stanford Research Systems SR540). These beams were then redirected by three mirrors to allow parallel paths into a 10 cm focusing lens. A polyimide coated silica capillary

(Polymicro Technologies) with 1 cm of its coating removed was placed at the focal point to allow the two pairs of beams to converge at the detection window. A spatial filter was placed behind the capillary to isolate only the strong-side signal beams from the remaining three pairs of beams. These overlapping signal beams were then reflected off of a mirror and through another 10cm focusing lens. A photodetector (Thorlabs PDA36A Si Amplified

Detector, 400-1100nm) was used to collect the signal, which was processed by a lock-in amplifier (Stanford Research Systems, SR810 DSP) and then digitized using AIDA data acquisition software.

3.3.3. Capillary Electrophoresis System

The custom-built capillary electrophoresis system (Figure 3.4) utilized a Glassman

PS/MJ30P0400-11 30 kV power supply with a custom-built control system to apply current across the system. Polyimide coated fused silica capillaries, 360 µm OD/100 µm ID x 35 cm

(17 cm effective) and 360 µm OD/75 µm ID x 40 cm (20 cm effective), were used for various experiments. Platinum electrodes were placed in glass sample vials at both end of a capillary, while system voltage and current were monitored by a multi-meter (Cen-tech

P35017) and AIDA data acquisition software. 50 mM ammonium acetate buffer (pH 5.1) was prepared in distilled deionized water.

Figure 3.3. Experimental setup for wave-mixing detection of malachite green and crystal violet. 532 nm and 635 nm lasers are used for simultaneous detection. S1, S2: Beam splitters; B1, B2: Beam blockers; L1, L2: Lenses.

Buffer solutions of 50 mM (pH 2.2) or 0.5 mM citric acid (pH 3.5) buffer were prepared in distilled deionized water and then diluted 2:1 with acetonitrile. These buffers were inspired by separate publications by Tsai, et al. and Sun, et al. (57, 60), which used similar background electrolytes in analysis of MG and CV. MG and CV samples were prepared in

0.5 mM citric acid and acetonitrile (1:2), 50 mM citric acid and acetonitrile (1:2) or 50 mM ammonium acetate . Electrokinetic sample injection (+20 kV) was performed at the anode.

3.3.4. Capillary Treatment

Prior to use, capillaries were treated with 0.1N NaOH for 10 minutes at 5 μL/min using a peristaltic pump (Rainin Dynamax RP-1). This was followed by priming with run buffer for 1 minute at the same flow rate. For capillary coating, Ultratrol LN capillary dynamic coating polymer (Target Discoveries) was flowed into the capillary at 5 μL/min between the standard 0.1N NaOH and run buffer treatment steps. Between sample runs, capillaries were flushed with 0.1N NaOH and run buffer for 1 minute at 5 μL/min.

Figure 3.4. Wave-Mixing Detection for Capillary Electrophoresis.

3.4. RESULTS AND DISCUSSION

3.4.1. Continuous Flow Detection

The simplest form of wave-mixing detection is direct analysis in a fixed sample cell.

For liquid samples, this may be accomplished through the use of any fused silica or quartz cell capable of holding liquids. To conduct direct analysis of MG and CV samples, a fused silica capillary (75 µm ID/360 µm OD), with 1 cm of its polyimide coating removed, was placed in the capillary electrophoresis apparatus shown in Figure 3.4. Individual samples were then pulled into the capillary by a peristaltic pump, and were allowed to flow at a slow rate (5 µL/minute). No electrical current was applied in this case, as no separation was required for this type of analysis. With either the 532nm or 635nm laser online, wave-mixing signal was generated by either sample type. Figure 3.5 shows raw signal obtained for 5.0 x

10-7 M MG in acetonitrile. The overall baseline noise is represented between 0 and 10 seconds, which was captured with both pump beams blocked (Figure 3.5A). At this stage, virtually no light entered the photodetector, which resulted in a baseline that was almost exactly zero. From 10 to 20 seconds, both pump beams were unblocked, which allowed for generation of the wave-mixing signal (Figure 3.5B). The relative signal recorded had a value of approximately 3. Lastly, between 20 and 30 seconds, the pump beam without frequency modulation (via an optical chopper) was blocked (Figure 3.5C). Since the remaining beam had frequency modulation, stray light that was not blocked by spatial filters entered the photodetector and was amplified by the lock-in amplifier tuned to the same frequency. This resulted in a relative signal that was somewhat elevated from the baseline depending on the position of each mirror and spatial filter in the wave-mixing apparatus. The net wave-mixing signal measured for this sample is the difference between the average of all data points

30 recorded with both pump beams unblocked and those recorded with the non-modulated beam blocked.

These same three levels of relative signal can also be seen for a sample of 1.0 x 10-7

M CV in acetonitrile (Figure 3.6). As with the MG sample, signal and noise levels were obtained through repeated blocking and unblocking of the pump beams. This type of assessment allowed for a fundamental understanding of the intensity of the signals generated by samples of high and low concentration, which was then used to establish appropriate detector and lock-in amplifier settings. Additionally, daily alignment of the optics in the wave-mixing apparatus was performed in a continuous-flow-mode using high concentration

MG and CV samples (50 – 100 mM).

3.4.2. Background Electrolytes

Initial attempts to detect MG and CV in capillary electrophoresis mode were made using an aqueous 50 mM ammonium acetate buffer (pH 5.1). This buffer system was chosen due to its acidic pH and the successful application of a similar solution in a publication by

Sun, et al. (60). However, when used as the sample solvent and background electrolyte in the capillary electrophoresis system, the current was found to be excessively high (> 240 µA) at

20 kV applied voltage. Through multiple sample injections with this buffer, only two successful runs were recorded without loss of system conductivity. Figure 3.7 shows an electropherogram for one of these runs. A 25 µM CV sample was injected electrokinetically for 30 seconds at +20 kV in a 360 μm OD/75 μm ID x 40 cm capillary with a 20 cm effective length. Despite the long injection duration, signal response was found to be relatively poor in the 50 mM ammonium acetate system. This weak signal, combined with the frequent boiling of the buffer due to high current, ultimately led to a change in CE run buffer.

Figure 3.5. 5.0 x 10-7 M malachite green in acetonitrile. 360 μm OD/75 μm ID capillary. A: pump beams blocked; B: pump beams unblocked; C: non-modulated beam blocked.

Figure 3.6. 1.0 x 10-7 M crystal violet in methanol. 360 μm OD/75 μm ID capillary. A: pump beams blocked; B: pump beams unblocked; C: non-modulated beam blocked.

0.5

0.4

0.3 Relative Signal Relative

0.2

0.1 150 200 250 300 Time (s)

Figure 3.7. 2.5 x 10-5 M crystal violet (CV) in 50 mM ammonium acetate. Capillary electrophoresis parameters: +20 kV, 360 μm OD/75 μm ID x 40 cm capillary (20 cm effective length), 30 s electrokinetic injection.

While a reduction in ammonium acetate concentration would have reduced system current to more manageable levels, the inadequate signal generation suggested the potential for low MG and CV detection limits was very limited. For this reason, another acidic run buffer was formulated using 50 mM citric acid. This system was chosen for its reported success as a capillary electrophoresis buffer in a 2007 work by Tsai, et al. (57). Initial testing of this 50 mM citric acid solution in the CE system showed a significant reduction in current to an average of 70 µA with a constant +20 kV applied voltage. This system current decrease versus ammonium acetate is due to a change in conductivity resulting from the higher molecular weight of citric acid. Although a more stable CE system was possible with this buffer, the wave-mixing signal response was found to be even lower than with the ammonium acetate buffer. The electropherogram peaks were also very broad and non-

Gaussian in shape (Figure 3.8A). This observation suggested poor analyte solubility in a purely aqueous citric acid solution, which may have resulted in capillary wall interaction. In light of this result, the decision to add a significant organic component to the run buffer was made. The 50 mM aqueous citric acid solution was then diluted 1:2 with acetonitrile to produce a more suitable run buffer. When this new buffer was used in a CE separation run, the reduction in net citric acid concentration and the change in the dielectric constant were found to further reduce the average system current to 10 µA at +20 kV. This allowed for highly consistent electropherograms and minimal instances of total current loss. The change also resulted in nearly a 50-fold increase in relative wave-mixing signal (Figure 3.8B) when compared to that of 50 mM citric acid alone (Figure 3.8A). This dramatic increase is believed to be due to a combination of effects. First, CV and MG both have greater absorptivity in acetonitrile than in water.

Figure 3.8. 1 x 10-4 M crystal violet (CV) in 50 mM citric acid (A). 1 x 10-4 M CV in 50 mM citric acid diluted with acetonitrile (1:2) (B). Capillary electrophoresis parameters: +20 kV, 360 μm OD/75 μm ID x 40 cm capillary (20 cm effective length); 30 s (A) and 5 s (B) electrokinetic injections.

Second, solubility of both compounds is increased in acetonitrile, which may have reduced capillary wall interactions and improved electrokinetic injections. Given the observations made with each of these capillary electrophoresis run buffers, the 50 mM citric acid: acetonitrile (1:2) solution was the clear choice in terms of signal response and system stability. This run buffer was used for all further capillary electrophoresis runs.

3.4.3. Field Amplified Sample Stacking

Sample preparation can have a major impact on separation and detection of analytes in capillary electrophoresis. Sample stacking is a preparation technique that aims at pre- concentration of analytes prior to separation to increase efficiency. There are numerous types of sample stacking procedures; however, many of these methods require complex control over the capillary system. Field Amplified Sample Stacking (FASS) is a simple stacking method that can achieve large increases in signal-to-noise ratio (S/N) with only minor system modifications (63). By injecting samples in a low-conductivity solvent, analytes experience a strong electric field, and therefore migrate at an increased rate relative to those injected in higher conductivity solvents. Analytes in a low-conductivity zone slow once the interface with a high-conductivity run buffer is reached, which causes a stacking effect (Figure 3.9).

Due to the basic nature of our capillary electrophoresis system, this technique was chosen for separation of MG and CV. Figure 3.10 shows electropherograms for 10 μM CV prepared in both 50 mM citric acid/acetonitrile (1:2) and 0.5 mM citric acid/acetonitrile (1:2). 50 mM citric acid/acetonitrile (1:2) run buffer was used in both trials. The peaks obtained showed a dramatic difference in S/N, with the sample prepared in low-conductivity stacking buffer yielding nearly a 5-fold increase over the sample prepared in run buffer.

Figure 3.9. Field Amplified Sample Stacking (FASS). E: Electric field.

Figure 3.10. Field Amplified Sample Stacking. 10 μM CV in 50 mM aqueous citric acid/acetonitrile (1:2) (Left). 10 μM CV in 0.5 mM aqueous citric acid / acetonitrile (1:2) stacking buffer (Right). Capillary electrophoresis parameters: +20 kV, 360 μm OD/75 μm ID x 40 cm capillary (20 cm effective length); 5 s electrokinetic injection.

Given this significant improvement in detection sensitivity, all further samples were prepared in 0.5 mM citric acid/acetonitrile (1:2) stacking buffer.

3.4.4. Bare Capillary Separation

Figure 3.11A shows a typical electropherogram for a mixture of 10 μM MG and 10

μM CV using capillary zone electrophoresis (CZE). Eleven successive injections of the sample mixture were performed to observe variation in run-to-run migration times. Migration times for each compound showed good repeatability at 57-65 seconds (RSD = 5.2%) for MG and 60-71 seconds (RSD = 5.6%) for CV (Figure 3.11B), despite a lack of temperature control or automated sample injection. For all trials, both MG and CV peaks were resolved from one another; however, it was observed that after 3 to 4 runs, migration times for both compounds increased. This is most evident with the migration times for runs 3 and 8, which show a significant increase in relation to the previous run. Additional treatment of the capillary with 0.1N NaOH was performed for 1 minute after each of these runs, and migration times were found to stabilize. This change in migration times is believed to be the result of re-protonation of capillary silanol groups due to the use of an acidic run buffer.

Decreases in the net negative charge on the capillary surface reduce electroosmotic flow

(EOF), while treatment with base serves to restore the net negative charge and EOF. Further reduction in run-to-run RSD may be obtained through flushing the capillary with 0.1N NaOH between each run.

Another observation from this experiment is the difference in analyte wave-mixing signal strengths. The signal for MG is significantly stronger than that of CV due to two factors.

Figure 3.11. (A) Electropherogram of a mixture of 10 μM MG and 10 μM CV in 0.5 mM aqueous citric acid/acetonitrile (1:2) stacking buffer. (B) Migration times for repeated injections of the same sample mixture in (A); MG (○), CV (◊). Capillary electrophoresis parameters: +20 kV, 360 μm OD/100 μm ID x 35 cm capillary (17 cm effective length), 10 s electrokinetic injection.

First, the 635 nm laser has a greater effective power (65 mW) than that of the 532 nm laser

(40 mW). Since wave-mixing signals have a cubic dependence on laser power, a higher intensity signal is generated for MG. Additionally, MG also has a small molar absorptivity at

532 nm, which allows signal formation from both input lasers. This amounts to total MG signal strength that is approximately 5 times greater than that generated by CV. The peaks shown in Figure 3.11A reflect this difference.

3.4.5. Effects of Capillary Dynamic Coating

Although separation of MG and CV was successfully obtained using a bare fused- silica capillary, a degree of system control is required to prevent shifts in migration time from causing peak overlap between runs. To this end, modification of the CZE system could potentially allow for improved separation. One means of creating greater separation between analytes is through reduction of electroosmotic flow. Dynamic capillary coatings offer a simple means of EOF reduction through non-permanent alteration of the capillary surface.

Figure 3.12 shows CE separation of MG and CV in a 100 μm ID capillary with and without

Ultratrol LN dynamic coating. In the uncoated capillary, MG and CV were detected at 66.3 and 72.3 seconds. However, the addition of Ultratrol LN shifted these migration times to 90.6 and 98.4 respectively. This is a 30% improvement in migration time differential; however, there was no net gain in overall peak resolution with this modification. In both cases, band broadening negatively affected peak shape, which negated improvement in migration time change. For this reason, bare capillaries were used for further measurement of samples.

Figure 3.12. Electropherograms of a mixture of 10 μM MG and 10 μM CV in 0.5 mM aqueous citric acid/acetonitrile (1:2) stacking buffer. (Top) Bare fused silica capillary (Bottom) Ultratrol LN dynamic capillary coating. Capillary electrophoresis parameters: +20 kV, 360 μm OD/100 μm ID x 35 cm capillary (17 cm effective length), 3 s electrokinetic injection.

3.4.6. Detection Limits and Linearity

To determine the lowest detectable concentration of both MG and CV, stock solutions were prepared at high concentration (1.0 x 10-3 M) in 0.5 mM aqueous citric acid/acetonitrile

(1:2) stacking buffer. Serial dilutions were then made ranging from 1.0 x 10-4 M to 1.0 x 10-12

M and tested individually. It should be noted that after initial optical alignment with high concentration samples, no changes were made to the positions of mirrors, spatial filters or the sample cell. The only changes made during this process were to detector gain and signal amplification scaling. Figure 3.13 shows the lowest concentration MG and CV samples (2.0 x 10-10 M MG and 2.0 x 10-9 M CV) that generated a detectable wave-mixing signal. From these electropherograms, the limits of detection (LOD) were calculated using peak height

(signal) and the standard deviation of the baseline (noise). LOD was determined to be 8.3 x

10-11 M for MG and 6.9 x 10-10 M for CV (S/N=2). Using an estimated probe volume of 360 pL, these values also correspond to mass detection limits of 3.0 x 10-20 mol and 2.5 x 10-19 mol respectively. These limits are very small since Degenerate Four-Wave Mixing only requires a short optical path length and the focused beam diameter is less than 70 μm.

Investigation of the technique as a quantitative tool led to generation of standard curves using a series of MG and CV calibrators. For each set of calibrators, instrument response versus analyte concentration was characterized. Instead of electrokinetic injection, each sample was drawn into a 360 μm OD/75 μm ID x 40 cm capillary using a peristaltic pump at a continuous flow rate of 5 μL/min. Wave-mixing signal produced by each sample was monitored over a 30 second period (10 data points/second). This methodology allowed for direct evaluation of wave-mixing signal linearity without variation introduced by the capillary electrophoresis system.

Figure 3.13. Electropherograms of 2.0 x 10-10 M MG and 2.0 x 10-9 CV in 0.5 mM aqueous citric acid/acetonitrile (1:2) stacking buffer. Capillary electrophoresis parameters: +20 kV, 360 μm OD/100 μm ID x 35 cm capillary (17 cm effective length); 15 s electrokinetic injection.

The mean signal values obtained for 1.0 – 50 μM CV samples are shown in Figure 3.14A.

Given the square dependence of wave-mixing signal on absorptivity, a second-order polynomial response was observed. When converted to a logarithmic scale, the curve became linear with a slope of approximately 2 (Figure 3.14B). Linear fit over this range was found to be good, with the relationship defined as y = 1.784x + 8.045 (R2 = 0.993). Similarly, Figures

3.14C and 3.14D illustrate mean wave-mixing signals for 6.25 - 50 µM MG calibrator samples on both absolute and logarithmic scales. Linear line fit for these samples also resulted in a high correlation coefficient (y = 1.736x + 7.615, R2 = 0.996).

3.5. CONCLUSIONS

Degenerate Four Wave-Mixing coupled with capillary electrophoresis provides a highly sensitive detection method for malachite green and crystal violet. The method does not require any derivatization or labeling of the target analytes, and it is capable of producing reproducible results in less than two minutes. This offers picomolar detection limits in a system that can be scaled down to field-deployable designs, while other published techniques could not achieve one or the other (Table 3.1 and Table 3.2). Further development of this method using commercial capillary electrophoresis equipment may offer even greater sensitivity and reproducibility through more advanced sample stacking techniques and enhanced system controls.

Figure 3.14. Signal vs. concentration of CV (A). log(signal) vs. log(concentration) of CV (B). Signal vs. concentration of MG (C). log(signal) vs. log(concentration) of MG (D). 360 μm OD/75 μm ID x 40 cm capillary.

Table 3.1. Detection Limits for Crystal Violet Analysis Methods. Method Detection Limit Reference

LC–QqQLIT-MS/MS 3.29 x 10-11 M (12.2 ppt) ( 52 )

LC-ESI-MS/MS 4.1 x 10-10 M (152 ppt) ( 53 )

CE-D4WM 6.9 x 10-10 M (256 ppt) This Work

ELISA 1.01 x 10-9 M (374 ppt) ( 55 )

LC–MS/MS 2.2 x 10-9 M (815 ppt) ( 51 )

SPE-UV/Vis 1.23 x 10-9 M (456 ppt) ( 48 )

CE - UV/Vis 9.8 x 10-8 M (36.3 ppb) ( 58 )

Table 3.2. Detection Limits for Malachite Green Analysis Methods. Method Detection Limit Reference

TiO SERS Sensor -12 2 1 x 10 M (0.3 ppt) ( 54 )

-11 LC–QqQLIT-MS/MS 3.29 x 10 M (10.8 ppt) ( 52 )

-11 CE-D4WM 8.3 x 10 M (27.3 ppt) This Work

-10 LC-DAD 1.08 x 10 M (35.6 ppt) ( 59 )

-10 LC-ESI-MS/MS 4.1 x 10 M (135 ppt) ( 53 )

-10 ELISA 6.33 x 10 M (209 ppt) ( 56 )

-10 LC-IT-MS 6.85 x 10 M (226 ppt) ( 50 )

CHAPTER 4

AMMONIUM NITRATE EXPLOSIVE SCREENING USING

DEGENERATE FOUR-WAVE MIXING SPECTROSCOPY

4.1. ABSTRACT

Degenerate Four-Wave Mixing (DFWM) is demonstrated as a sensitive detection method for ammonium nitrate explosives. This method can serve as an alternative to Ion

Mobility Spectrometers (IMS), which utilize conductivity or mass spectrometric detectors and are currently used by the Transportation Security Administration (TSA) for airport explosives screening. To produce a sample suitable for wave-mixing analysis with a 635 nm diode laser, an aqueous ammonium nitrate sample is converted to an optically absorbent compound through reaction with diphenylamine in sulfuric acid. Samples can be analyzed in capillary or thin-film cells depending on specific application. The limit of detection for 1 mm

ID capillary cells was determined to be 1.5 x 10-9 M (5.2 x 10-18 mol). The detection limit for a 1.5 mm thin-film cell was found to be 3.2 x 10-7 M (2.0 x 10-15 mol). Detection limits were calculated for a signal-to-noise ratio of 2.

4.2. INTRODUCTION

Ammonium nitrate is an ionic compound commonly used in the manufacture of explosives due to the strong oxidizing character of nitrates. Other nitrate-based oxidizers such as sodium nitrate and urea nitrate may also be used in a similar manner. A mixture of

94% fertilizer-grade ammonium nitrate and 6% fuel oil (ANFO) makes up one of the most common nitrate explosives. Additionally, many slurry explosives have been formulated using a blend of ammonium nitrate and other explosives including trinitrotoluene (64-67). While these types of explosives typically serve an industrial purpose, they may also be exploited in the form of improvised explosive devices (IEDs) and other bombs. In 2012, ammonium nitrate explosives were ranked as the most common type of devices used by Taliban insurgents in Afghanistan due to the presence of fertilizer plants operating in Pakistan (68).

Such devices also pose a threat to homeland security, and, for this reason, nitrates are part of explosives screenings performed by the Transportation Security Administration (TSA). The current chemical-based TSA method for trace explosive detection is ion mobility spectrometry (IMS), and, while this method has had success, there are significant chemical specificity and sensitivity limitations (69, 70). Pairing of IMS with mass spectrometry (MS) has been investigated as a means of increasing sensitivity, but false positives as a result of poor selectivity remain an issue (71, 72). In some cases, methods using simple UV-visible spectrophotometers for detection of ammonium and nitrate ions outperform both IMS and

MS in terms of detection sensitivity (73-75).

In this work, an optical detection method for trace amounts of ammonium nitrate using Degenerate Four-Wave Mixing is demonstrated. Degenerate Four-Wave Mixing is a powerful spectroscopic method that produces a signal through the formation of a thermal diffraction grating at the intersection point of two input laser beams (pump and pump/probe) passing through an absorbing analyte. The angle of the input beams determines the angle of the two signal beams, which allows a predictable location for collection by photodetector.

Because the input beams are derived from one laser source split by a 70:30 beam splitter, the

50 signal beam produced from the pump/probe beam is stronger than the one produced by the pump beam. This signal has a square dependence on analyte absorptivity and a cubic dependence on laser power (29, 36, 61). These properties allow the method to take advantage of high power-density laser sources to excite molecules even if they do not have a large extinction coefficient. This allows a very dilute sample to produce a strong wave-mixing signal that is proportional to concentration.

Given the dependence of a wave-mixing signal on sample optical absorbance, steps must be taken to convert a weakly absorbing sample into one with greater absorptivity.

Ammonium nitrate alone has very poor optical absorption, which necessitates indirect detection. Using a method adapted from Grebber et al. and Roberts (76, 77), nitrate ions were reacted with diphenylamine in sulfuric acid to give a blue-violet diphenylbenzidine derivative product. This colored product can be excited in a wave-mixing setup with a laser wavelength near 600 nm (Figure 4.1). Products of this reaction are suitable for continuous- flow detection through a capillary and on surfaces using glass slides/plates.

4.3. EXPERIMENTAL

4.3.1. Chemicals

Ammonium nitrate and diphenylamine were purchased from Sigma Aldrich. All other chemicals were of reagent grade.

4.3.2. Degenerate Four-Wave Mixing Optical Setup

For analyte detection, a Forward-Scattering Degenerate Four-Wave Mixing configuration was used (Figure 4.2). A 100 mW 635nm diode laser (Laserglow Technologies,

LRD-0635-TSR-00100-10) was directed to a 70R/30T (70% reflected/30% transmitted) beam splitter to create two input beams.

Figure 4.1. UV-visible spectra of 0.1 M ammonium nitrate (A) and the product of the reaction of ammonium nitrate (1.0 x 10-5 M) and 4.5 mM diphenylamine in 13.8 M sulfuric acid (B).

The weak-side beam was passed through an optical chopper (Stanford Research Systems

SR540). The two input beams were then redirected by three mirrors to allow parallel paths into a 10 cm focusing lens. A fused silica capillary or a thin-film sample cell was positioned at the focal point to allow the two beams to converge at the detection window. A spatial filter

(beam blocker) was placed behind either sample cell to isolate the strong-side signal beam from the remaining three beams. This signal beam was then reflected off of a mirror and through another 10cm focusing lens. A photodetector (Thorlabs PDA36A Si Amplified

Detector, 400-1100nm) was used to collect the signal, which was processed by a lock-in amplifier (Stanford Research Systems, SR810 DSP) and then digitized using AIDA data acquisition software.

4.3.3. Capillary Sample Cells

For continuous-flow detection of ammonium nitrate, three sizes of fused silica capillaries were used: 75 μm ID/360 μm OD (Polymicro Technologies), 360 μm ID/1000 μm

OD (VWR) and 1000 μm ID/1500 μm OD (Fisherbrand). Each capillary was fixed in a slotted metal block with adjustable clamps and spacers to prevent damage and prevent movement. The sample holder was attached to an XYZ translation stage for position adjustment in all three directions. Tubing was connected to one or both ends of the capillary, with one side routed through a peristaltic pump for sample injection, and the other side placed in sample vials (Figure 4.3).

4.3.4. Thin-Film Sample Cells

For surface-mode detection of ammonium nitrate, analytes were deposited onto quartz windows. A second quartz window was then placed on top, which dispersed the liquid across the surface to create a thin film with an approximate thickness of 6.5 μm.

Figure 4.2. Experimental setup for wave-mixing detection of ammonium nitrate. S1: Beam splitter; B1, B2: Beam blockers; L1, L2: Lenses.

Figure 4.3. Ammonium nitrate capillary sample cell.

This value was determined using the volume of the liquid added and the diameter of the thin- film spot as shown in Figure 4.4A. A second type of thin-film sample cell was prepared in the same manner, except a Teflon-coated rubber o-ring with a thickness of 1.5 mm was placed between the quartz slides to act as a spacer. For this type of sample cell, a larger volume of liquid could be added, which also allowed for a larger volume to be probed and detected by wave mixing (Figure 4.4B).

4.3.5. Nitrate Derivatization

To produce a colored compound from ammonium nitrate, a mixture of 4.5 mM diphenylamine and 13.8 M sulfuric acid was used to treat ammonium nitrate samples. While the reactants were colorless, the blue-violet diphenylbenzidine derivative product of this reaction absorbs red light and can be detected by wave-mixing using a red laser (76, 77). The structure of the colored product is shown in Figure 4.5. Color formation in this reaction is rapid, with initial change observable within minutes at room temperature. Prior to analysis, all samples were incubated at room temperature for 30 minutes to allow for complete color change. Due to instability of the colored diphenylbenzidine derivative in solutions of neutral or basic pH, samples were not transferred to another solution before use in the wave-mixing detection system.

Figure 4.4. Ammonium nitrate thin-film sample cells.

Figure 4.5. Diphenylbenzidine derivative produced through reaction of diphenylamine with ammonium nitrate and sulfuric acid.

4.4. RESULTS AND DISCUSSION

4.4.1. Nitrate Derivatization Kinetics

To assess the reaction kinetics involved in the formation of the diphenylbenzidine derivative from ammonium nitrate, a sample of 2.5 x 10-4 M ammonium nitrate was prepared with 4.5 mM diphenylamine in 13.8M sulfuric acid. The sample was left at room temperature

(25 °C), and absorbance measurements were taken over the course of 15 minutes at 620 nm using a UV-visible spectrophotometer (Agilent Technologies, 8453). Color change was observed to be rapid, with significant change within 1 minute of sample preparation. After approximately 8 minutes, the maximum absorbance at 620 nm was reached (Figure 4.6A).

For all measurements, 1 to 10 dilutions were prepared using 4.5 mM diphenylamine in

13.8M sulfuric acid as a diluent. Based on these observations, an additional three samples of

2.5 x 10-4 M ammonium nitrate were prepared in the same manner as previously described.

These samples were then incubated at room temperature (25 °C), 45 °C and 60 °C for 1 minute. Each sample was diluted 1 to 10 with 4.5 mM diphenylamine in 13.8M sulfuric acid and measured by the UV-visible spectrophotometer. Absorbance spectra for these samples are shown in Figure 4.6B. Absorbance at 620nm for the 25 °C sample was found to be consistent with the results of the initial study of absorbance change over time. However, significantly greater absorbance around 620 nm was observed for the 45 °C and 60 °C samples. After 1 minute at 45 °C, color development had progressed to yield absorbance at

620 nm that was 87% of the maximum value recorded upon completion of the reaction. After

1 minute at 60 °C, 100% of maximum absorbance was reached. These results demonstrate that continuous heating results in complete conversion of diphenylamine to the diphenylbenzidine derivative in a shorter amount of time than when performed at 25 °C.

Figure 4.6. (A) Absorbance at 620 nm for the product of ammonium nitrate (2.5 x 10-4 M) reacted with 4.5 mM diphenylamine in 13.8M sulfuric acid at 25 °C (1:10 dilution). (B) Absorbance of ammonium nitrate (2.5 x 10-4 M) reacted with 4.5 mM diphenylamine in 13.8 M sulfuric acid at 25 °C, 45 °C and 60 °C for 1 minute (1:10 dilution).

However, it was also observed that heating samples at 60 °C in excess of 30 minutes would result in initial formation of blue-violet color, followed by reversion to a colorless solution.

This may be the result of a secondary acid-catalyzed reaction that proceeds at a much slower rate, since colored samples that were stored at room temperature overnight were also found to lose their color. For the purposes of this work, samples used for further wave-mixing analysis were made fresh daily and incubated at room temperature. However, adaptation of this technique for an explosives detection system could utilize a short heating period for rapid analysis.

4.4.2. Detection in Capillary Cells

To assess wave-mixing signals generated from an ammonium nitrate samples prepared with the diphenylamine reagent described in Section 4.3.5, three separate capillary systems were tested with samples in continuous-flow mode. The first capillary tested was a fused silica capillary (75 µm ID/360 µm OD) with 1 cm of its polyimide coating removed, which was placed in the path of the 635 nm laser using the capillary holder depicted in

Figure 4.3. Through the use of a peristaltic pump, samples were drawn into the capillary for direct analysis at 5 μL/min. Samples of varying concentration were tested individually. For each sample that demonstrated a result that could be distinguished from random baseline noise, signal-to-noise ratio and limit of detection were calculated.

Figure 4.7 shows wave-mixing signal obtained for 2.5 x 10-5 M ammonium nitrate in diphenylamine reagent. The data points recorded for the first 10 seconds represent the overall baseline noise picked up by the photodetector (Figure 4.7A). This was captured by blocking of both pump beams. With the input laser beams blocked, only a negligible amount of light may enter the photodetector, which results in a baseline that is nearly zero.

Figure 4.7. Ammonium nitrate (2.5 x 10-5 M) in diphenylamine reagent. 75 µm ID/360 µm OD capillary. A: pump beams blocked; B: pump beams unblocked; C: non- modulated beam blocked.

For the next 10 seconds, both pump beams were unblocked, which allowed generation of the wave-mixing signal (Figure 4.7B).The relative signal recorded here had a value close to 3.

Lastly, for 10 additional seconds, the pump beam that was not modulated by an optical chopper was blocked (Figure 4.7C). Since the remaining beam was modulated, stray light that was not blocked by spatial filters could enter the photodetector. This generates a relative signal that is somewhat elevated from the baseline depending on the position of each mirror and spatial filter in the wave-mixing optical setup. The net wave-mixing signal measured for this sample is the difference between the average of all data points recorded with both pump beams unblocked and those recorded with the non- modulated beam blocked. By comparing this net signal to the standard deviation of the noise observed with the non-modulated pump beam blocked, the limit of detection was calculated to be 1.1 x 10-6 M (S/N = 2). This value was converted to a mass detection limit of 3.1 x 10-16 mol using an estimated probe volume of 270 pL. While the mass detection limit was already low, significant improvement of the concentration detection limit was expected with minor changes to the detection system as described below.

To improve the concentration detection limit for ammonium nitrate samples, the path length of the sample cell was increased in order to enhance the wave-mixing signal. To accomplish this, the 75 μm ID/360 μm OD capillary was replaced with an uncoated 360 μm

ID/1000 μm OD capillary. This change increased the probe volume to an estimated 1.3 nL, which was nearly a 5-fold increase over the smaller capillary. As with previous continuous- flow mode testing, ammonium nitrate samples were prepared at multiple concentrations in the diphenylamine reagent. The signal recorded for a 5.0 x 10-6 M ammonium nitrate sample is shown in Figure 4.8.

As with the previous result, the pump beams were blocked and unblocked in 10 second intervals to observe relative signal with and without actual wave-mixing signal being generated from the sample. The net wave-mixing signal is the difference between the average of all data points recorded with both pump beams unblocked and those recorded with the non- modulated beam blocked. The concentration limit of detection for this result was determined to be 6.4 x 10-7 M (S/N = 2), which represented a modest improvement over that calculated for the 75 μm capillary. However, the corresponding mass detection limit was found to increase to 8.3 x 10-16 mol. This was expected given the use of a larger capillary, which increased the probe volume.

To further assess the impact of probe volume and sample cell path length on the signal- to-noise ratio, the 360 μm ID/1000 μm OD capillary was replaced with an uncoated 1000 μm

ID/1500 μm OD fused-silica capillary. This change resulted in a 3-fold increase in both path length and probe volume (3.6 nL) when compared to that of the previous capillary (360 μm).

Once more, varied concentrations of ammonium nitrate samples reacted with the diphenylamine reagent were prepared and analyzed. The overall wave-mixing signal intensity observed with this sample cell was found to be significantly greater than that captured when using the 75 μm and 360 μm capillaries. This allowed for detection of samples of much lower concentration. The lowest concentration sample measured was an ammonium nitrate solution of 1.0 x 10-8 M (Figure 4.9). As with prior capillary samples, the pump beams were blocked and unblocked to determine raw relative signal and background noise levels. Comparison of the net signal with the standard deviation of the background noise level yielded a good signal-to-noise ratio despite the low sample concentration.

Figure 4.8. Ammonium nitrate (5.0 x 10-6 M) in diphenylamine reagent. 360 µm ID/1000 µm OD capillary. A: pump beams blocked; B: pump beams unblocked; C: non- modulated beam blocked.

Figure 4.9. Ammonium nitrate (1.0 x 10-8 M) in diphenylamine reagent. 1 mm ID/1.5 mm OD capillary. A: pump beams blocked; B: pump beams unblocked; C: non- modulated beam blocked.

The concentration detection limit was calculated as 1.5 x 10-9 M (S/N = 2), with a corresponding to a mass detection limit of 5.2 x 10-18 mol. This represents a significant gain in terms of both the concentration detection limit and the mass detection limit for ammonium nitrate. This is interesting in that the mass detection limit was shown to be superior to that of smaller diameter capillaries, even though probe volume was increased. This is due to increases in wave-mixing signal strength, which provided greater overall sensitivity to small quantities of ammonium nitrate.

4.4.3. Detection in Thin-Film Cells

In addition to detection in capillary cells, it is also possible to detect nitrate explosives on surfaces. To this end, sample cells were designed to produce a thin film of analyte solution that could behave in a manner that would be similar to if it were on an exposed surface. The two types of thin-film sample cells described in Section 4.3.4 were used to demonstrate this capability. For both thin-film cells, small volumes of aqueous ammonium nitrate were mixed with the diphenylamine reagent and then allowed to incubate at room temperature in excess of 30 minutes. A portion of this mixture was placed between the quartz plates of the sample cells, which were then situated at the focal point in the wave-mixing optical setup. Additionally, measurements were taken with a blank sample containing no ammonium nitrate. Figure 4.10 shows wave-mixing signal obtained for 1.0 x 10-5 M ammonium nitrate in diphenylamine reagent in the thin-film cell without a spacer o-ring. As with the results for capillary detection, baseline noise was compared to relative signal. From time point 0 to 10 seconds, both pump beams were blocked to record the overall baseline noise picked up by the photodetector (Figure 4.10A). In this state, the amount of light that can be picked up by the photodetector is very small, which results in a near-zero baseline.

After 10 seconds, both pump beams were unblocked to record the wave-mixing signal generated from the ammonium nitrate sample (Figure 4.10B). For this sample, the relative signal recorded had a value close to 4, while the corresponding measurements for the blank sample were also elevated to a slightly lower level. Upon blocking the pump beam without modulation, only a minor reduction in relative signal was observed from the ammonium nitrate sample, while the signal from the blank sample actually showed a minor increase

(Figure 4.10C). This suggests that the amount of stray light created by the laser passing through the two quartz plates was significantly greater than that observed with capillary cells, and that the wave-mixing signal itself was also weaker.

The net wave-mixing signal for the ammonium nitrate sample is calculated as the difference between the average of all data points recorded with both pump beams unblocked and those recorded with the non-modulated beam blocked. The background noise level recorded in the latter case is also equivalent to the relative signal value associated with the blank sample. This demonstrates that the relative signal value recorded corresponds to the presence or absence of ammonium nitrate. For this system, the limit of detection was calculated to be 1.6 x 10-6 M (S/N = 2). This value was converted to a mass detection limit of

3.6 x 10-17 mol using an estimated probe volume of 20 pL. This small probe volume corresponds to a path length of only 6.5 µm, which partly explains the lower wave-mixing signals generated from this thin-film cell. In comparison, the capillary cells used with the same ammonium nitrate samples yielded lower limits of detection due to greater path length

(75 – 1000 µm) and smaller surface area. This resulted in a larger wave-mixing zone within the sample, as well as a reduction in stray light background levels from the sample cell itself.

Figure 4.10. Ammonium nitrate (1.0 x 10-5 M) in diphenylamine reagent (left) and diphenylamine reagent blank (right). 6.5 µm thin-film cell. A: pump beams blocked; B: pump beams unblocked; C: non-modulated beam blocked.

Based on these observations, the original thin-film sample cell was modified using a

Teflon-coated rubber o-ring spacer between the two quartz plates. This increased the path length to approximately 1.5 mm and the probe volume to 6.1 nL. As with capillary cells, a large increase in path length with thin-film cells also resulted in signal-to-noise ratio improvement. Wave-mixing signal obtained for 1.0 x 10-5 M ammonium nitrate in diphenylamine reagent in a thin-film cell with a 1.5 mm spacer o-ring is shown in Figure

4.11. As with the previous example, the pump beams were blocked and unblocked in a sequence designed to show signal versus background noise. A comparison of the net signal with the standard deviation of the background noise level (or blank signal) yielded a concentration detection limit of 3.2 x 10-7 M (S/N = 2), which corresponds to a mass detection limit of 2.0 x 10-15 mol using an estimated probe volume of 6.1 nL. The gains for this thin-film cell are significant in terms of detection of low concentrations of ammonium nitrate, as the increased path length resulted in an improvement of nearly one order of magnitude. However, the mass detection limit is reduced when compared to the thin-film cell with a 6.5 µm path length. This is expected due to the change in probe volume associated with modification with an o-ring spacer.

4.4.4. Linearity of Response

Through collection of results for varied concentrations of ammonium nitrate, the relationship between signal and concentration can be observed as being distinctly nonlinear.

This is as expected given the square dependence on analyte concentration given by the

DFWM signal intensity equation. This relationship was confirmed through the use of a series of ammonium nitrate calibrators, where instrument response could be compared with analyte concentration.

Figure 4.11. Ammonium nitrate (1.0 x 10-5 M) in diphenylamine reagent (left) and diphenylamine reagent blank (right). 1500 µm thin-film cell. A: pump beams blocked; B: pump beams unblocked; C: non-modulated beam blocked.

Samples were drawn into a 75 μm ID/360 μm OD capillary cell using a peristaltic pump at a continuous flow rate of 5 μL/min. Wave-mixing signal produced by each sample was monitored over a 30 second period (10 data points/second). The logarithm of mean signal values obtained for 12.5 to 100 μM ammonium nitrate in diphenylamine reagent are shown plotted against the logarithm of concentration in Figure 4.12. A line plotted through these points has a slope of 1.5, or approximately 2, which is not ideal but consistent with the expected quadratic relationship. Linear fit over this range was found to be good, with the relationship defined as y = 1.5154x - 2.4834 (R2 = 0.988). Deviation from a slope of exactly

2.0 can be explained by low levels of stray light collected by the photodetector.

4.5. CONCLUSIONS

Degenerate Four-Wave Mixing offers sensitive detection of trace quantities of nitrate- based explosives in different sample types as described in this chapter. The technique is viable with liquids in both capillary cells and thin-film analytes, which may be applied to both airport explosives screening and open-environment samples. Rapid identification of nitrate compounds is accomplished without separation equipment due to the use of a color- forming reaction unique to the strong oxidizing characteristics of the nitrate ion. The lowest detection limits are achieved through the use of large-diameter capillaries or thin-film cells with a spacer (Table 4.1). Further optimization may determine the ideal path length to optimize both concentration and mass detection limits. These results demonstrate superior detection limits and specificity when compared to existing published spectroscopic methods

(Table 4.2). Given this, the method detailed in this chapter could therefore prove useful in field screening applications.

1 y = 1.515x - 2.483 R² = 0.988

0.5

0 log(Relative Signal) log(Relative -0.5

-1 1 1.2 1.4 1.6 1.8 2 log [Concentration (µM)]

Figure 4.12. Logarithmic plot of ammonium nitrate concentration (12.5 to 100 µM) versus relative wave-mixing signal.

Table 4.1. Detection Limits for Ammonium Nitrate in Varied Sample Cells. Cell Type Path Length (µm) LOD (M) LOD (mol)

Flow 75 1.1 x 10-6 3.1 x 10-16

Flow 360 6.4 x 10-7 8.3 x 10-16

Flow 1000 1.5 x 10-9 5.2 x 10-18

Thin-Film 6.5 1.6 x 10-6 3.6 x 10-17

Thin-Film 1500 3.2 x 10-7 2.0 x 10-15

Table 4.2. Detection Limits for Various Ammonium or Nitrate Ion Analysis Methods. Detection Method LOD (M) Reference

Laser Wave Mixing 1.5 x 10-9 This Work

UV/Visible Spectrophotometer 1.8 x 10-8 ( 12 )

Ion Mobility Spectrometer 2 x 10-7 ( 7 )

UV-visible Spectrophotometer 1.8 x 10-5 ( 10 )

UV-visible Spectrophotometer 5.6 x 10-5 ( 11 )

Electrospray Ionization-MS/MS 3.7 x 10-4 ( 9 )

CHAPTER 5

SENSITIVE DETECTION OF CARCINOEMBRYONIC ANTIGEN BY

DEGENERATE FOUR-WAVE MIXING SPECTROSCOPY

5.1. ABSTRACT

Degenerate Four-Wave Mixing (DFWM) is demonstrated as an ultrasensitive detection method for proteins including carcinoembryonic antigen (CEA). Isolation of CEA was achieved through magnetic immunoprecipitation using anti-CEA (CD66E). Purified samples were reacted with Bradford Protein Reagent or BCA Protein Reagent to produce colored products with strong optical absorption near laser lines of 532 nm or 635 nm.

Coupling of DFWM with a capillary array allowed for high-throughput analysis for sampling plans with large numbers of replicates, while standard curves may be generated with inexpensive protein samples instead of pure antigen targets. The limit of detection using a

150 µm ID capillary array was determined to be 3.3 x 10-12 M (0.59 ng/mL). The corresponding mass detection limit was 1.2 x 10-21 mol (0.22 fg). Detection limits were calculated for a signal-to-noise (S/N) ratio of 2.

5.2. INTRODUCTION

Carcinoembryonic antigen (CEA) is a glycoprotein that is present in high levels in the fetal colon, and typically appears to a lesser degree in adults. Elevated levels in adults can be indicative of tumor growth in the colorectal region, as well as the stomach, small intestine,

75 pancreas, liver, cervix or lungs (78, 79). CEA levels may be measured in either plasma or pleural effusion patient samples, and concentrations in excess of 2.5 to 5.0 ng/mL are commonly accepted as indicators of malignant tumors (78, 80, 81). Therefore, it has the potential for use as a biomarker for cancer screening, as well as monitoring for reoccurrences in patients in remission (82, 83). The CEA protein is heavily glycosylated, with about 60% of its 180 kDa molecular weight comprised of carbohydrates. The peptide sequence is coded by a gene on human chromosome 19, which yields a 70 kDa protein prior to glycosylation.

The structure is similar to immunoglobulins and contains three highly conserved repeats of

178 amino acids that form loops through disulfide linkages (82-84). CEA normally presents itself on the surface of epithelial cells in the colon, but may appear in a more disordered manner in tumors cells. The function of CEA is not completely known, but it may serve as a recognition protein, as is the case with other immunoglobulins. It is believed that CEA is involved in maintaining the integrity of the luminal epithelium in the adult colon through spatial orientation (83, 85).

Numerous methods have been reported for the detection of CEA, with the majority of them showing capability of measurement below the 2.5 ng/mL physiological threshold (86-

94). In each of these cases, CEA is detected indirectly via an α-CEA antibody that has been modified with a label such as horseradish peroxidase, colloidal gold or quantum dots. These are used to form an immunosensor that can capture the CEA protein and return a signal based on quantity. Unlike these methods, our technique decouples the antibody capture process from the detection step, and directly measures the CEA antigen through its interaction with a color-forming reagent. This decoupling allows for better process control, since signal generation does not depend on the degree of successful labeling of α-CEA. Furthermore,

76 while the existing immunosensor methods report concentration detection limits in the low picomolar to high femtomolar range, the corresponding mass detection limits are poor. This is due to the relatively large sample volumes used to capture enough protein for detection.

Additionally, the coupling of a multichannel capillary array with the DFWM detector allows for high-throughput analysis of large numbers of sample replicates, with very little sample consumption. Immunosensor techniques require regeneration of the sensor after every individual sample is tested, which makes testing of multiple samples challenging. The amount of patient sample required to perform multiple analyses is also prohibitive with these methods.

5.2.1. Degenerate Four-Wave Mixing

In this work, we demonstrate an optical detection method for low levels of CEA and other proteins using degenerate four-wave mixing. As described in Chapter 2, Degenerate

Four-Wave Mixing (DFWM) is a powerful spectroscopic method that produces a signal through the formation of a thermal diffraction grating at the intersection point of two input laser beams (pump and pump/probe) passing through an absorbing analyte. The angle of the input beams determines the angle of the two signal beams, which allows a predictable location for collection by photodetector. Because the input beams are derived from one laser split by a 70:30 beam splitter, the signal beam produced from the pump/probe beam is stronger than the one produced by the pump beam. The wave-mixing signal has a square dependence on analyte absorptivity and a cubic dependence on laser power (29, 36, 61).

These nonlinear properties allow the method to excite weakly absorbing analytes to produce a strong signal that is proportional to concentration. Because the wave-mixing signal is

77 coherent laser-like beam, it can be collected with high efficiency against a nearly 100% dark background. This greatly enhances the signal-to-noise ratio.

5.2.2. Bradford Protein Assay

The Bradford Protein Assay (95) is a total protein assay that functions through non- covalent binding of Coomassie Brilliant Blue G-250 (Figure 5.1). Under acidic conditions, the dye carries a positive charge and appears red in color. Upon interaction with protein, overall charge becomes negative due to the loss of protons. While bound to the protein, the dye has a blue color that is stable for about one hour. This blue product has a maximum absorbance at 595 nm, while the red form absorbs strongly at 465 nm. This wavelength shift allows for analysis of small quantities of protein against a concentrated background of unbound dye. Standard curves may be generated with inexpensive proteins due to relatively similar binding response regardless of structure. For proteins that show significant deviation from typical response, a correction factor may be used to appropriately scale calculated values (95). This assay is compatible with wave-mixing detection due to the large extinction coefficient associated with the protein-dye complex. However, the red form of the dye can produce a wave-mixing signal on its own due to absorbance across the ultra-violet and visible range.

5.2.3. BCA Protein Assay

Since its introduction in 1985 (96), the BCA Protein Assay has seen widespread use for quantitative measurement of total protein. This protein assay uses a two-step reaction to produce a colored compound in a relatively short period of time.

Figure 5.1. Bradford protein assay reaction. Nitrogen deprotonation results in a color change from red to blue.

Initially, Cu2+ interacts with nitrogen atoms in a protein peptide backbone to form a faint blue-violet chelate (λmax at 540 nm). This is known as the Biuret Reaction. The byproduct of this reaction is the reduction of Cu2+ to Cu+.The second stage of the process involves the formation of a bicinchonic acid (BCA)-Cu+ chelate (Figure 5.2). This BCA-Cu+ chelate has a

2+ deep purple color (λmax at 562 nm) with a large extinction coefficient. While the protein-Cu chelate alone has some absorption in the visible wavelength range, it has a small extinction coefficient, and is not suitable for sensitive detection of protein (97). Because four-wave mixing is an absorption-based detection method, the intense color of the BCA-Cu+ chelate allows for generation of a strong signal. Additionally, in the absence of protein, the BCA working solution does not show significant optical absorption of its own, which results in a high signal-to-noise ratio when measuring protein samples. This is true for conventional absorbance-based detection, as well as wave-mixing given the dependence on optical absorption of analytes.

5.3. EXPERIMENTAL

5.3.1. Chemicals

Carcinoembryonic antigen full length protein (Cat. ab742) was purchased from

Abcam, Inc. Anti-CEA CD66E (Cat. MS613PABX), Pierce BCA Protein Assay Kit (Cat.

23227) and Thermo Scientific Pierce Classic Magnetic IP/Co-IP Kit (Cat. PI88804) were purchased from Fisher Scientific. Bradford Protein Assay 1X Dye Reagent (Cat. B6916) was purchase from Sigma Aldrich. Ultratrol LN was purchased from Target Discoveries. All other chemicals were of reagent grade.

Figure 5.2. Two-step BCA protein assay reaction. Reduction of Cu2+ to Cu+ via the Biuret Reaction allows formation of the BCA-Cu+ chelate.

5.3.2. Degenerate Four-Wave Mixing Optical Setup

For protein detection, a Forward-Scattering Degenerate Four-Wave mixing (F-

DFWM) configuration was used (Figure 5.3). A 200 mW 532nm Nd-YAG laser (Changchun

New Industries Optoelectronics Tech Co. Ltd., MGL-FN-523-200mW) or a 100 mW 635nm diode laser (Laserglow Technologies, LRD-0635-TSR-00100-10) was passed through a

70R/30T (70% reflected/30% transmitted) beam splitter to create two input beams. Total laser power was reduced to 40 mW and 65 mW respectively prior to splitting. The weak-side beam was passed through an optical chopper (Stanford Research Systems SR540). These beams were then redirected by three mirrors to allow parallel paths into a 10 cm focusing lens. An array of fused silica capillaries was positioned at the focal point to allow the two beams to converge at the detection window. A spatial filter (beam blocker) was placed behind the capillaries to isolate only the strong-side signal beam from the remaining three beams. This signal beam was then reflected off of a mirror and through another 10cm focusing lens. The wave-mixing signal was collected by a photodetector (Thorlabs, PDA36A

Si Amplified Detector, 400-1100nm), processed by a lock-in amplifier (Stanford Research

Systems, SR810 DSP), and then digitized using AIDA data acquisition software.

5.3.3. Multichannel Capillary Array

To construct a capillary array for continuous-flow mode detection of carcinoembryonic antigen, three 75 μm ID/360 μm OD fused silica capillaries (Polymicro

Technologies) of equal length were selected. For each capillary, a 1 cm section of the polyimide coating was removed by flame. An additional array was constructed in the same manner using three 150 μm ID/360 μm OD fused silica capillaries (Polymicro Technologies).

Figure 5.3. Experimental setup for wave-mixing detection of carcinoembryonic antigen. 532 nm Nd:YAG or 635 nm diode laser. S1: Beam splitter; B1, B2: Beam blockers; L1, L2: Lenses.

The capillaries were then bonded together with epoxy and fixed into a slotted metal block with adjustable clamps and spacers to prevent damage. All three capillaries were situated to lay flat within the slot. The sample holder was attached to an XYZ translation stage for position adjustment in all three directions. The X and Z-axes were controlled by manual adjustment knobs, while the Y-axis was controlled by a linear actuator (Zaber TLA-28). A tube was connected to one end of the capillaries and routed through a peristaltic pump for sample injection. The open ends of the capillaries were placed directly into sample vials

(Figure 5.4). Prior to injection with protein samples, the capillaries were treated with 0.1N

NaOH for 10 minutes at 5 μL/min using a peristaltic pump (Rainin, Dynamax RP-1). To minimize protein interaction with the capillary walls, Ultratrol LN capillary dynamic coating polymer (Target Discoveries) was flowed through at 5 μL/min for 3 minutes after the 0.1N

NaOH treatment step. After 3 sample runs, capillaries were retreated with 0.1N NaOH and

Ultratrol LN.

5.3.4. Linear Actuator Control

To control vertical movement of the capillary array, the Zaber TLA-28 linear actuator was connected to a computer running Zaber Console software (v1.2.29.788), which allowed for precise alignment and movement of the capillary arrays in and out of the path of the laser. For analysis of samples within a single capillary, the actuator was fixed at a single position throughout collection of wave-mixing signal. For separate measurement of multiple analytes, two different methods were used. The first method commanded the actuator to move in a single direction at a pre-defined speed of 25 µm/second.

Figure 5.4. Capillary array sample cell and actuator.

The second method used instructions written in JavaScript to move the actuator to four separate capillary positions in rapid succession. This script also allowed for control over the amount of time spent at a single position and the number of times the actions were repeated

(Figure 5.5).

5.3.5. Magnetic Immunoprecipitation Protocol

Preparation of immunoprecipitated CEA samples was carried out using the instruction provided with the Thermo Scientific Pierce Classic Magnetic IP and Co-IP Kit (Figure 5.6).

For each sample, 125 µL of Pierce Protein A/G Magnetic Beads were added to microcentrifuge tubes, and then washed with the provided IP Lysis/Wash Buffer. 50 µg of anti-CEA CD66E was combined with the magnetic particles, and then allowed to incubate for 1 hour. Following a wash with the IP Lysis/Wash Buffer, a mixture containing 25 µg

CEA and 25 µg BSA protein was added to one tube, while no additional protein was added to another to serve as a control. After one hour of incubation, each tube was washed with the

IP Lysis/Wash Buffer and distilled deionized water. Elution of the target CEA was performed by addition of 100 µL of low-pH Elution Buffer, followed by 10 minutes of incubation. Samples were neutralized with 10 µL of Neutralization Buffer.

5.3.6. HPLC Protocol

High-Performance Liquid Chromatography (HPLC) protein sample analysis used an

Agilent 1100 series instrument with a Zorbax GF-250 size-exclusion column and a 200 mM ammonium sulfate/50 mM potassium phosphate/10 mM boric acid (pH 7) buffer. Injection volume was 50 µL with a flow rate of 1 mL/min. Column temperature was regulated at 30

°C. Sample run time was set at 17 minutes to allow complete elution of all proteins.

Figure 5.5. JavaScript for delayed four-position movement of Zaber TLA-28 linear actuator used for the capillary array.

Figure 5.6. Magnetic immunoprecipitation procedure for isolation of CEA from a heterogeneous protein solution.

5.3.7. Bradford Protein Assay Protocol

For the Bradford Protein Assay, the instructions provided with the kit purchased from

Bio-Rad were used to develop a protocol. Protein samples were diluted 1:10 with 1X

Bradford Dye Reagent (Coomassie Brilliant Blue G-250 in methanol and phosphoric acid).

These samples were incubated for 5 minutes at room temperature to allow development of blue color. Samples were stored at room temperature for up to 1 hour before they were discarded, as the protein-dye complex is not stable beyond this amount of time (95).

5.3.8. BCA Protein Assay Protocol

Preparation of reagents for the BCA Protein Assay was carried out using a protocol based on the instructions provided with the kit purchased from Fisher Scientific. A BCA

Reagent working solution was formulated through 1:50 dilution of BCA Reagent B (4% cupric sulfate) with BCA Reagent A (sodium carbonate, sodium bicarbonate, bicinchoninic acid and sodium tartrate in 0.1 M sodium hydroxide). This resulted in a light green colored solution. Protein samples were diluted 1:10 or 1:100 with this working solution, and then incubated for 60 minutes at 60 °C in a water bath. Upon completion of the incubation, samples were cooled to room temperature. Samples were stored at 2 - 8 °C for up to 2 months without noticeable reduction in purple color.

5.4. RESULTS AND DISCUSSION

5.4.1. Bradford and BCA Assays

In preparation of samples for analysis by DFWM, both the Bradford and BCA Protein

Assays were used to generate colored analytes. While both methods resulted in non-covalent binding of protein with dye or copper to yield a color change, the Bradford 1X Dye Reagent showed significant optical absorption near the 635 nm laser line while in its initial red state.

Figure 5.7 shows the absorption spectra for Bradford 1X Dye Reagent alone and after reacting with 100 µg/mL BSA protein. The Agilent 8453 UV-visible spectrophotometer was blanked with ethanol, which is the primary solvent in the 1X Dye Reagent. The expected shift from 465 nm to 595 nm was observed in the presence of protein, but approximately one third of the absorbance in the red region of the visible spectrum was due to unbound

Coomassie Brilliant Blue G-250 from the 1X Dye Reagent. When injected into the capillary array, this elevated background resulted in significant background signal generation in the absence of protein. Initial testing using this reagent showed difficulty achieving detection of trace protein quantities due to high blank signal. In contrast, the optical absorption profile for the BCA Reagent alone was found to have less than 0.1 AU near the 532 nm laser line, which is five-fold lower than that recorded for the Bradford Reagent at 635 nm. The absorption spectra for BCA samples with 0, 2, 5 and 10 µg/mL BSA protein are shown in

Figure 5.8. The UV-visible spectrophotometer was blanked with deionized water prior to these measurements. With the BCA Assay, background absorbance levels were found to be minimal, with a distinct increase near the wavelength of the 532 nm Nd:YAG laser used in the DWFM setup. For this reason, further sample preparation was performed exclusively with the BCA Reagent.

Figure 5.7. UV-visible spectra of Bradford reagent alone and with BSA (100 µg/mL).

Figure 5.8. UV-visible spectra of BCA reagent alone and with 2 to 10 µg/mL BSA.

To ensure that the BCA Assay was compatible with CEA, a 10 µg/mL sample was prepared with the BCA Reagent with the same dilution and incubation parameters used for

BSA samples. Figure 5.9 shows the absorption spectra for this sample, as well as a blank sample of the BCA Reagent alone. As with prior experiments, the UV-visible spectrophotometer was blanked with deionized water. For the 10 µg/mL CEA sample, the expected peak centered at 562 nm was observed. However, the amount of purple color formation was also observed to be somewhat less than that of a BSA sample at the same concentration. Absorbance at the 532 nm laser line was determined to be 2.8 times greater for BSA than for CEA. This may be due to the large number of carbohydrates on the surface of CEA, which could have prevented some sections of the protein from reacting with the

BCA Reagent. Quantitative determination of CEA is possible with a BSA standard curve through the use of a scaling factor based on this relationship. This procedure is part of typical BCA protocols, which account for differences in protein structure in this same manner (96, 97).

5.4.2. Magnetic Immunoprecipitation

To demonstrate the CEA purification capability of magnetic immunoprecipitation

(IP), a 25 µg sample of CEA was mixed with 25 µg of BSA. This sample was then re- purified by magnetic-IP with anti-CEA CD66E as a capture antibody. A control was also prepared with CD66E alone (no CEA or BSA protein). If magnetic-IP elution conditions cause a portion of the capture antibody to co-elute with the target antigen, an antibody control allows for a total protein correction factor.

Figure 5.9. UV-visible spectra of BCA reagent alone and with 10 µg/mL CEA.

Each sample was then analyzed by size-exclusion high-performance liquid chromatography (SEC-HPLC), as described in Section 5.3.6. Figure 5.10 shows the results of this analysis, along with two standards: a molecular weight standard containing 1 mg/mL bovine gamma globulin (155 kDa), BSA (67 kDa) and carbonic anhydrase (29 kDa), as well as a CEA standard containing 250 µg/mL CEA. When compared with the standards, the magnetic-IP purified CEA sample (Figure 5.10C) shows the presence of CEA dimer (8.036 min.), CEA monomer/CD66E dimer (8.801 min.) and CD66E monomer (9.548 min.).

Reference peaks for CEA and CD66E are shown in Figure 5.10B and Figure 5.10D. There is no observed peak at 10.0 minutes that would correspond to the 67 kDa BSA protein added to the sample prior to purification. For reference, BSA protein was present in the molecular weight standard chromatogram at 10.029 minutes (Figure 5.10A). This demonstrates 100%

CEA sample purity with respect to the BSA contaminant introduced intentionally. However, a portion of anti-CEA CD66E monomer and dimer was visible in the magnetic-IP sample.

By peak area, 51-66% of the eluted sample was made up of CD66E. This was expected given the non-covalent binding of CD66E to the magnetic particle protein A/G coating.

Upon elution of CEA under low-pH conditions, CD66E could co-elute. For this reason, the magnetic-IP control sample with CD66E alone was prepared. Figure 5.10D shows the amount of CD66E antibody expected to co-elute with the CEA target. For the wave-mixing analysis, the relative signal associated with a CD66E magnetic-IP control can be subtracted from that of a CEA/CD66E mixture to quantify CEA alone.

For further method development, two possible modifications could be made to eliminate the need for a CD66E control sample. First, the pH or ionic strength of the magnetic-IP elution buffer could be adjusted to favor CEA over CD66E.

Figure. 5.10. Size exclusion (SEC)-HPLC chromatograms for molecular weight standards (465, 310, 155, 67 and 29 kDa) (A), CEA standard (B), magnetic-IP purified CEA (C) and magnetic-IP control (D).

Fine tuning of these conditions could yield a higher purity CEA sample for wave-mixing analyses. A second approach to increasing the purity of CEA samples involves covalent bonding of the CD66E capture antibody to the protein A/G magnetic particle coating. The reagents required to accomplish this are commercially available in some immunoprecipitation kits. Adaptation of the immunoprecipitation methods used in this work would be relatively simple.

5.4.3. Multi-Channel Array Scanning Modes

To analyze multiple samples within the capillary array, small, incremental movements were required by the linear actuator used to control the vertical position of the array. With continuously flowing analytes passing through each capillary, automated movement from one capillary to another allowed for simultaneous analysis without the need for additional sample injections. To accomplish this, two different approaches were devised.

The first method used a simple movement command built into the Zaber Console software. This command instructs the actuator to move at a constant speed in one direction until fully extended or retracted. To test wave-mixing response with this method, BSA protein solutions that had been treated with the BCA Reagent were injected into all three capillaries making up the array. The actuator was manually positioned so the array would start above the path of the laser, and then commanded to move downward at a constant rate.

Multiple movement speeds were tested between 10 and 1000 µm/second, and the 25

µm/second rate was chosen as the ideal speed given relative signal height observed. Figure

5.11 shows the wave-mixing signals obtained for 270 µg/mL BSA with a scanning rate of 25

µm/second. In this case, capillaries 1 and 3 returned similar response, while the signal from capillary 2 was significantly lower.

2 Relative Signal Relative

0 200 400 600 800 1000 1200 Position (µm)

Figure 5.11. Continuous vertical scan of capillary array containing 270 µg/mL BSA- BCA. 75 µm ID/360 µm OD fused silica capillaries. 25 µm/s scan rate.

This difference is due to optical characteristics unique to each section of fused-silica capillary. This can typically be minimized through cleaning of the capillary windows with a flame, as well as changes in horizontal positioning. Individual capillary responses may also be standardized against one of the other capillaries.

The second scanning mode tested used the script described in Section 5.3.4 (Figure

5.5). This script allowed rapid movement to probe individual capillaries, followed by a set delay, and then movement to a position below the capillary array. As with testing of the first scanning mode, BSA-BCA solutions were injected into all three capillaries. The wave- mixing signals recorded for a 200 µg/mL BSA sample are shown in Figure 5.12. Because of the instructions to move to specific positions, analysis of the samples within all three capillaries was possible in a relatively short time (12 seconds). In this example, the measurements were repeated five times to determine repeatability of peak heights. This variability was determined to be 11.1% RSD for capillary 1, 1.9% for capillary 2 and 1.4%

RSD for capillary 3. Overall, this shows high repeatability for measurements in this scanning mode. The elevated variability observed with capillary 1 may be due to imperfections in the capillary itself, which could be overcome by a change in horizontal positioning in the path of the laser.

In comparing these two methods for scanning of the capillary array, the primary difference lies in the time required to complete one scan. The scripted mode took approximately five times less time than the 25 µm/second scan mode. This allowed for greater improvement in the sample measurement rate, which lends itself to higher accuracy through the capability to analyze high numbers of sample replicates.

2 Relative Signal Relative

0 0 20 40 60 80 Time (s)

Figure 5.12. Three-position scan of capillary array containing 200 µg/mL BSA-BCA. 75 µm ID/360 µm OD fused silica capillaries.

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Another observation from this testing was the occurrence of noise peaks between the signal peaks when scanning at low speed (Figure 5.11). These artifacts were the result of the laser diffracting off of the walls of the capillaries when positioned between them. Some of this light could be captured by the photodetector, which resulted in the recording of false signal peaks. At higher concentrations of analyte, this is of little concern; however at low concentrations, these noise peaks can make isolation of the true signal peaks difficult. For these reasons, the scripted scanning mode appears to be the more favorable method of measurement for capillary arrays paired with a DFWM detector.

5.4.4. Linearity of Response

To demonstrate viability of the BCA Protein Assay coupled with DFWM as a quantitative method for detection of protein samples, instrument response was measured relative to analyte concentration. To do so, a series of BSA-BCA samples were prepared with final concentrations of 50 - 200 µg/mL. These samples were drawn into one channel of a 75 μm ID/360 μm OD capillary array using a peristaltic pump at a continuous flow rate of

5 μL/min. A single channel was used to prevent response differences due to use of different capillaries. The linear actuator was positioned so the laser would only pass through the one capillary in use. The wave-mixing signal produced by each sample was monitored over a 20 second period (10 data points/second), and was then averaged to provide a response value for each BSA concentration level. The average response of a blank sample was subtracted from

BSA response to give net signal values. The net signal values obtained for obtained for 50 –

200 µg/mL BSA-BCA are shown plotted against concentration in Figure 5.13. Given the square dependence of wave-mixing signal on absorptivity, a second-order polynomial response was observed as expected.

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Figure 5.13. Signal vs. concentration of BSA (left). log(signal) vs. log(concentration) of BSA (right). 360 μm OD/75 μm ID x 40 cm capillary array.

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This was also converted to a logarithmic scale to apply a linear fit to the data points (Figure

5.13). The line plotted through these points has a slope of approximately 2, which is consistent with a square dependence on analyte concentration. Linear fit over this range was found to be very good, with the relationship defined as y = 1.881x – 3.4313 (R2 = 0.989).

Some amount of deviation from a slope of exactly 2.0 is expected due to low levels of stray light collected by the photodetector, as well as variability in the wave-mixing thermal grating.

5.4.5. Limit of Detection

To assess the trace detection capability of the DWFM capillary array for CEA protein, a series of samples were prepared with final concentrations of 20 ng/mL to 10

µg/mL CEA using the BCA Reagent. Dilution and incubation parameters were as described in Section 5.3.8. Samples were injected individually into one capillary at 5 µL/min. to minimize capillary-to-capillary variability. Figure 5.14 shows the wave-mixing signal for the lowest concentration sample measured (20 ng/mL CEA), as well as the BCA Reagent blank signal. For both samples, the pump beams were blocked and unblocked to determine raw relative signal and background noise levels. Comparison of the net CEA signal with the standard deviation of the background noise level yielded a good signal-to-noise ratio despite the low sample concentration. The limit of detection was determined to be 3.3 x 10-12 M or

0.59 ng/mL CEA (S/N = 2), with a corresponding to a mass detection limit of 1.2 x 10-21 mol or 0.22 fg. The concentration detection limit is significantly below the 2.5 ng/mL threshold commonly used in diagnosis and monitoring of cancers, and the mass detection limit demonstrates the possibility for greater sensitivity if protein concentration techniques are used prior to sample injection.

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5.5. CONCLUSIONS

The pairing of DFWM with immunoprecipitation and multichannel capillary arrays provides a highly sensitive method of detection for protein biomarkers such as CEA. The technique is capable of high-throughput analysis of multiple protein samples within a short period, and it can do so with a high degree of specificity though the use of α-CEA for sample preparation. Future work to convert to a covalent magnetic-immunoprecipitation method can be expected to yield even higher specificity. Standard curves may be designed with excellent linear line fit using pure target protein samples or with inexpensive proteins like BSA. This is an advantage over detection techniques that function through labeling of α-CEA.

Comparison of this method with the more sensitive published techniques also shows significant improvement in terms of mass detection limits due to the sub-nanoliter sample volume requirements (Table 5.1). Since the majority of the other sensitive methods are variations of colloidal gold voltammetric immunosensors, the claimed concentration detection limits are based on the built-in concentrating capability of that type of sensor. Pre- concentration of samples may be paired with our DFWM method to provide a similar effect, which would make the system compatible with even lower analyte concentrations.

Furthermore, additional sensitivity increases may be realized through the use of capillaries with an inner diameter greater than 150 µm, since wave-mixing probe volume would be increased.

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Table 5.1. Detection Limits for CEA Analysis Methods. Ref. Detection Method Volume LOD LOD LOD (L) (ng/mL) (M) (mol) ( 94 ) Gold-QD Voltammetric 4.0E-05 2.0E-6 1.1E-17 4.4E-22 Immunosensor ( 88 ) Gold Voltammetric Immunosensor 1.0E-03 1.5E-3 8.3E-15 8.3E-18

( 92 ) Gold Voltammetric Immunosensor 7.0E-04 0.01 5.6E-14 3.9E-17

( 90 ) Chemiluminescent Immunoassay 2.0E-05 0.025 1.4E-13 2.8E-18

( 87 ) Thermal Lens Microscopy 2.0E-07 0.03 1.7E-13 3.3E-20 Immunoassay ( 91 ) Chemiluminescent Immunoassay 5.0E-05 0.048 2.7E-13 1.3E-17

( 89 ) Gold Voltammetric Immunosensor N/A* 0.06 3.3E-13 N/A*

( 93 ) Gold Voltammetric Immunosensor N/A* 0.06 3.3E-13 N/A*

This DWFM Immunoassay 5.4E-10 0.59 3.3E-12 1.8E-21 Work ( 86 ) I-125 Radioimmunoassay 1.0E-02 0.5 2.8E-12 2.8E-14

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