The Pennsylvania State University the Graduate School The

Home , Botryococcus, Lycopane

The Pennsylvania State University

The Graduate School

The Department of Chemistry

NEW ANALYTICAL APPROACHES TO UNDERSTAND BIOLOGICAL

SYSTEMS WITH SECONDARY ION MASS SPECTROMETRY

A Dissertation in

Chemistry

Lauren Marie Jackson

 2014 Lauren Marie Jackson

Submitted in Partial Fulfillment of the Requirements for the Degree of

Doctor of Philosophy

August 2014

The dissertation of Lauren Marie Jackson was reviewed and approved* by the following:

Nicholas Winograd Evan Pugh Professor of Chemistry Dissertation Advisor Chair of Committee

Wayne Curtis Professor of Chemical Engineering

Thomas Mallouk Evan Pugh Professor of Materials Chemistry and Physics Associate Head of the Chemistry Department

Barbara Garrison Shapiro Professor of Chemistry Head of the Chemistry Department

*Signatures are on file in the Graduate School

iii

ABSTRACT

The work of this thesis has focused on investigating biological systems with the most current secondary ion mass spectrometry instrumentation and novel sample preparation techniques. The microalga biofuel candidate, Botryococcus braunii, has been thoroughly investigated by three-dimensional chemical imaging to answer several questions regarding its metabolism and chemical composition. These findings provide a breadth of information beneficial and crucial to the successful advancement of B. braunii as a biofuel. The enhancement effects and cellular vitrification capabilities of trehalose have been explored and confirmed trehalose entry into cells has been found. This work is evidence of the great contributions to the biological realm to which SIMS can contribute.

TABLE OF CONTENTS

List of Figures ...... vii

List of Tables ...... xi

Acknowledgements ...... xii

Chapter 1 Introduction ...... 1

1.1. Background of Secondary Ion Mass Spectrometry Imaging and Instrumentation ...... 1 1.2. Research Overview ...... 4 1.3. References ...... 7

Chapter 2 The Use of Botryococcus braunii as a Simple Biological Model Sample for Diverse Secondary Ion Mass Spectrometry Analysis ...... 20

2.1. Introduction ...... 20 2.2. Experimental ...... 21 2.2.1. Algal Culture Conditions ...... 21 2.2.2. Algal Sample Preparation for SIMS Analyses ...... 21 + 2.2.3. SIMS and SEM Algal Analyses with C60 Primary Ion Beam ...... 22 2.2.4. SIMS and SEM Algal Analyses with Argon Cluster Primary Ion Beam ...... 22 2.3. Results and Discussion ...... 23 2.3.1. The Tolerance of B. braunii Cells to the Vacuum Environment ...... 23 2.3.2. Three-Dimensional Analysis without the Effects of Charging ...... 26 2.3.3. The Variety of Chemical and Morphological Features in the B. braunii Colony ...... 28 3.3.4. The Chemical Diversity of B. braunii observable by SIMS ...... 30 2.4. Conclusions...... 35 2.5. References ...... 37

Chapter 3 Understanding the Behavior of the Triterpene Hydrocarbons of Botryococcus braunii during SIMS Depth Profiling Experiments ...... 46

3.1. Introduction ...... 46 3.2. Experimental ...... 48 3.1.1. Algal Culture Conditions ...... 48 3.1.2. Algal Sample Preparation for TOF-SIMS Analyses ...... 49 3.1.3. Squalene standard sample preparation for depth profiling by SIMS analysis ...... 49

3.1.3. SIMS depth profiling conditions for squalene standard and B. braunii analysis ...... 50 3.3. Results and Discussion ...... 50 3.4. Conclusions...... 56 3.5. References ...... 56

Chapter 4 Elucidation of the Metabolic Processes of Three Dimensional Chemical Imaging of the Algal Biofuel Candidate, Botryococcus braunii ...... 60

4.1. Introduction ...... 60 4.2. Experimental ...... 62 4.2.1. Algal Culture Conditions ...... 62 4.2.2. Algal Sample Preparation for TOF-SIMS Analyses ...... 63 4.3.3. Instrument Parameters for SEM and TOF-SIMS Algal Analysis ...... 63 4.3. Results and Discussion ...... 64 4.4. Conclusion ...... 77 4.5. References ...... 78

Chapter 5 Trehalose as a biologically-relevant matrix for secondary ion enhancement and quantitative detection of purines for SIMS analysis ...... 82

5.1 Introduction ...... 82 5.1.1. The de novo Purine Biosynthetic Pathway and Associated Metabolites .... 84 5.2. Semi-Quantitative Detection of Purines ...... 86 5.2.1. Experimental ...... 86 5.2.2. Results and Discussion ...... 89 5.2.3. Conclusions ...... 94 5.4. Determining the Enhancement Effect of Trehalose for Purine Detection ..... 96 5.4.1. Experimental ...... 96 5.4.2. Results and Discussion ...... 98 6.4.3. Conclusions ...... 104 5.5. New Trehalose Vitrification Technique for Three-Dimensional Single Cell Studies ...... 105 5.5.1. Experimental ...... 105 5.5.2. Results and Discussion ...... 107 5.5.3. Conclusions ...... 116 5.6. References ...... 116

Chapter 6 Conclusions and Future Directions ...... 121

6.1. Ongoing and Future Research ...... 121 6.1.1. The Potential Symbiotic Role of the Flavonoids of B. braunii ...... 121 6.1.2. Wax Monoester Expression and the Maintenance of an Axenic Line of B. braunii Race B ...... 122 6.1.3. Physiological Engineering of the Oil Bodies of B. braunii ...... 122

6.1.4. Time-Course Chemical Imaging of Mammalian Cells ...... 123 6.1.5. Guiding Principles for the Development of Biological Analyses with SIMS ...... 124 6.2. Conclusory Remarks ...... 125 6.3. References ...... 125

vii

LIST OF FIGURES

Figure 2-1: A comparison of Race A UTEX #2441 and #572 out- and in-vacuum environment of the SIMS instrument. Optical microscopy images of B. braunii race A shows that the cells of (A) UTEX #2441 form colonies while cells of (D) #572 do not. Secondary emission micrographs taken inside SIMS instrument show that in (E) and (F) the intracellular contents of UTEX #572 have spilled outside of the ruptured cells whereas the cells of UTEX #2441 in (B) and (C) have not ruptured. The scale bars of (A) and (D) are 50 μm and 25 μm; (B) and (E) are 25 μm; (C) and (F) are 12 μm...... 24

Figure 2-2: The absence of charging effects with depth during a B. braunii imaging experiment. The total ion signal has been divided by four to directly compare the signals...... 27

+ Figure 2-3: Chemical images from B. braunii race A by C60 primary ion beam. (A) The total ion image from approximately 1 micron inside the colony reveals features unique to this depth. The circular features 1 – 3 microns in diameter contain (B) hydrocarbon signals (m/z 410 – 430) and are localized to intracellular oil bodies which are about 1-2 microns underneath the surface of the sample. (C) The surface of the colony is covered in wax monoesters (m/z 530, 558, 586). The scale bars are 12 μm...... 29

Figure 2-4: Secondary electron micrographs from B. braunii taken with Ar4000 primary ion beam. The colonies give off fewer secondary electrons than the substrate and appear darker than the signal from the silicon substrate. The scale bars in (A) and (B) are 12 μm and 25 μm, respectively...... 31

+ Figure 2-5: Chemical images from B. braunii race A by Ar4000 primary ion beam. (A) The total ion of the surface of the algal cell colonies. The (B) hydrocarbon signals (m/z 410 – 430) cover the colonies, as well as (C) the wax monoesters (m/z 530, 558, 586). The scale bars are 12 μm...... 32

Figure 3-1: Mass spectrum of the squalene standard and the corresponding mass assignments compared to the exact masses of isotopes and random rearrangements. The exact masses of isotopes and rearrangements are in green text. The observed masses are in blue text...... 52

Figure 3-2: Comparison of the squalene standard spectrum versus the same m/z range in the B. braunii Race B sample. Primary ion dose for both spectra is 1 13 + 2 x 10 C60 /cm . (A) is the squalene standard and (B) is of the algae...... 53

Figure 3-3: The squalene standard’s signals as a function of depth and continuous + bombardment by the C60 ion...... 55

viii

Figure 4-1: The overall diversity of hydrocarbons increase with depth. (A) and (B) are spectra from the top surface layer of the colony obtained with a dose 13 + 2 of 1 x 10 C60 /cm . (B) and (D) are zoomed into the m/z 300 – 600 region of the spectrum in (A) and (C), respectively. (C) and (D) are spectra from the intracellular region of the cells within the colony obtained after a dose of 3 x 14 + 2 10 C60 /cm corresponding with a depth of about 1 -2 μm deep into the colony...... 65

Figure 4-2: The C34H58 distribution is different with depth. This hydrocarbon’s distribution on the surface of the colony is nearly homogenous, yet when 1-2 microns deep into the colony, at the intracellular level, it is homogeneously distributed in the oil bodies. All of the oil bodies have the same contents...... 66

Figure 4-3: The mass spectrum of B. braunii features two distinct lysolipids. Peak assignments were determined by comparing the observed mass with the accurate masses of the lysolipids and by comparing the isotopic patterns with a predictive isotopic distribution calculated by the chemical formula. Lysolipids have been reported in race B before, but this is the first time their identity and distribution have been described...... 71

Figure 4-4: The chemical structures of the two lysolipids of B. braunii race B identified by SIMS. Each lysolipid is characterized by its zwitterionic nature, hydrophobic tail, and a phosphate group...... 72

Figure 4-5: The two lysolipids have the same chemical distribution through the colony (A) in the extracellular matrix and surface of the colony and (B) at the intracellular level around the regions of oil bodies. The field-of-view of each image is 100 x 100 µm2...... 73

Figure 4-6: Proposed biochemical and physiological role for lysolipds in triterpene biosynthesis and export. (1) botryococcene bionsythesis enzymes SSL-1 (red) and SSL-3 (dark blue) associate with the zwitterionic lysolipid choline head groups and facilitate C30 botryococcocene biosnthesis. (2) Methyltransferases (green) decorate the botyrococcene; also hypothecised to be lipid-body associated. (3) Lysolipids facilitate localized destabilization of plasma membrane for oil excretion...... 76

Figure 5-1: The relative quantitation of the (a) IMP- and (b) AMP-doped trehalose spin-cast films is shown. The coefficient of determination is (a) R2 = 0.992 and (b) R2 = 0.999, respectively...... 90

Figure 5-2: The mass signature peaks for [IMP-H]- and [AMP-H]- are resolved here within a mass spectrum of a trehalose film doped with IMP and AMP, respectively...... 92

Figure 5-3: The relative quantitation of the (a) IMP- and (b) AMP-doped cell homogenate/trehalose spin-cast films is shown. The coefficient of determination is (a) R2 = 0.797 and (b) R2 = 0.975, respectively...... 93

Figure 5-4: The enhancement effect of trehalose in negative and positive mode SIMS. The spectrum of the neat solid is shown in (A) positive mode and (C) negative mode, respectively. When IMP is doped into a trehalose film, the positive ion is enhanced by 4,750 X and the negative ion is increased to a lesser extent by 125 X...... 99

Figure 5-5: The chemical interactions of IMP and trehalose investigated by H- NMR. (A) and (B) are the control spectra obtained from pure IMP and anhydrous trehalose, respectively. Spectra from (A) and (B) were overlayed with software to obtain spectrum (C), representative of a predictive spectrum which would be obtained if IMP and trehalose did not interact to form ions. The spectrum in (C) is from the sample contained a mixture of IMP and trehalose. It is nearly identical to (C), indicating no proton rearrangements between the two species...... 102

Figure 5-6: Comparison of the signal obtained in IMP-doped trehalose films using D2O versus H2O as a solvent. The green depth profile represents the sample created with D2O as the solvent, while the blue had H2O as the solvent. The error bars represent the standard deviation calculated from three trials...... 103

Figure 5-7: The RAW and BOAC cells vitrified in trehalose. The (A) RAW Cells retain a spherical shape and the nucleus is seen intact. The (B) BOAC cells are spherical and encased in a glassy trehalose film. The scale bars in (A) and (B) are 15 μm and 50 μm, respectively...... 109

Figure 5-8: Purines and other biochemical in vitrified RAW cells. The chemical images shown are the result of all the layers of the depth profile summed together in two dimensions (x-y). Chemical images (A) through (F) are purines. (A) is guanine, [M+H]+, at m/z 152; (B) is phenylalanine, [M+H]+, at m/z 120; (C) is thymine/isoleucine, [M+H]+, at m/z 84; (D) is adenine, [M+H]+, at m/z 136; (E) is arginine, [M+H]+, at m/z 175; and (F) is inosine monophosphate, [M+H]+, at m/z 349. (G) is trehalose, [M+H]+, at m/z 343. + (H) is the phosphocholine head group, C5H15NPO4 at m/z 184. (I) is Indium, [M]+ at m/z 115...... 111

Figure 5-9: Three-dimension chemical image of RAW cells vitrified in trehalose. Yellow is trehalose, fuchsia is adenine, blue is the phosphocholine headgroup. The field of view is 80 x 80 μm2...... 113

Figure 5-10: Imaging depth profile comparing cellular signal with trehalose distribution. (A) The trehalose signal is observed throughout the depth profile. (B) The trehalose and adenine signal are correlated and the trehalose signal in (A) clearly overlaps with the adenine signal in here. In (C) the trehalose signal is observed completely encircling the PC signal, implying complete coverage around the spherical shape of the cell. Each of the four chemical images in each series is representative of a dose of approximately 1 14 + 2 x 10 C60 /cm ...... 115

LIST OF TABLES

+ Table 2-1: Assorted metabolites of B. braunii observable by SIMS by C60 and + Ar4000 . Chemical formulas denoted by an asterisk have isomers. † denotes there are several other rearrangements for ion formation; the two most common are listed here. ‡ denotes the identity assignments supported by tandem MS...... 33

+ Table 2-2: Lipids discoverable by Ar2000 bombardment in the higher mass region. The lipids were identified by their accurate masses. All of the Lipid Maps Database matches for each different chemical formula is listed in the table. Chemical formulas denoted by an asterisk have isomers. Mass assignments assumed to represent molecular ions, [M]+...... 36

xii

Acknowledgements

Working towards my Ph.D. has been the most rewarding and exciting goal of my life. The pursuit of scientific knowledge is my passion whether it has related to my current research or not. It is for this reason that I truly value the people who have supported and enriched my growth as a scientist.

I would like to thank my family. From the time I was three years old, my parents recognized my interest in science and provided me with the tools and training to allow me to explore my interests. They made sure I was involved in Montessori schooling with a range of scientific and engineering activities available to me and set up a large

“laboratory” in my playroom. The tremendous sacrifices they made to send my brother and I to the best possible schools without blinking an eye is a testament to their dedication. While they are not scientists, they are both creative and hard-working people.

My mother has trained me in public speaking and other theatrical techniques; these skills have proven invaluable in communicating science. Since we could have conversations, my father would point to any random object and ask me, “How can this be made better?”

These conversations have shaped my thinking and approach to research. It is a question I ask myself several times a day. It is the reason that I have been so attracted to analytical chemistry. My brother has been my friend (and now my counselor) throughout my life, even though we are different in our approaches, I appreciate his views and consistently seek out his guidance.

The science educators I have met through my undergraduate career were instrumental in encouraging my direction towards graduate school. My chemistry

xiii professors at Marist introduced me to what chemistry was really all about. When I returned from winter break my freshman year, I approached Dr. Neil Fitzgerald about where I could start a research project and he gladly offered me my first appointment in analytical chemistry. After that, there was no turning back for me. I have learned a great deal from Dr. Jocelyn Nadeau. In her office hours, I learned how to think more universally about chemistry. Her extensive and selfless guidance in helping me learn more about graduate school is something I have thought back on often during my time here at Penn State. No matter who was my professor during the semester, all of my professors at Marist have been dedicated to supporting my progression as a chemist. I know I can still count on their guidance to this day.

I would like to thank my previous research mentors at the Smithsonian Institution and Rensselaer Polytechnic Institute. When I became a Smithsonian Research Fellow, I was assigned a project in biogeochemistry, but Dr. Gerhadt “Fritz” Riedel, gave me the freedom to start a concurrent analytical chemistry project with the most expensive instrument on the site, maybe because I filled the bass guitar position in his SI band.

There, I was able to learn what kind of researcher I am. Denise Butera helped me learn how to keep organized in lab and, most importantly, how to stand my ground as a scientist. It was my pleasure to have the opportunity to work with Dr. Linda McGown at

Rensselaer Polytechnic Institute. Her outlook was that as a professor, she was able to be the artist of her own research; this approach continues to strike a chord with me.

xiv

Throughout my years in graduate school, I have had the opportunity to share and discuss science and friendship with the members of my group. I feel that when they succeed, we all succeed and I take pride in the people with whom I work.

I thank my committee members Dr. Barbara Garrison, Dr. Thomas Mallouk, and

Dr. Wayne Curtis for helping me move towards the completion of my goals here at Penn

State.

I want to particularly thank Dr. Wayne Curtis for welcoming me as an unofficial group member of his lab. Our collaboration has been scientifically exciting and enriching since the first day of its inception. We have been able to enter completely uncharted territory in our research multiple times. I have had the pleasure of attending his group’s scientific discussions, which have helped me interface with how chemical engineers are approaching biofuel research. The members of his group have been eager to work with me and contribute to these projects

Ultimately, I want to thank Dr. Nicholas Winograd. When I first met Nick, he was a few hours off a long flight from England, excitedly explained to me all about SIMS and their goals to chemically image a cell in three dimensions, gave me a tour of the lab, and decided that I should come to Penn State. The science is his lab was unique and I knew this was where I needed to develop into an analytical chemist. From the start, he offered me a unique blend of freedom to develop my own research and guidance to bring those projects to fruition. I appreciate him pushing me to become a better and more informed instrumentalist. He helped me to think about my experiments more comprehensively. The opportunities I have had in his lab, I would not have been able to

xv receive anywhere else. He has worked hard to create this environment and, in appreciation of that, my goal has been to make more than the most of it. I admire his research goals to take SIMS into the biological arena and my thesis work has been dedicated to putting that into action.

Chapter 1

Introduction

1.1. Background of Secondary Ion Mass Spectrometry Imaging and Instrumentation

Imaging Mass Spectrometry (IMS) is a spatially-resolved method where mass spectra are collected from an array of positions across a sample surface. Intensities of selected signals from the mass spectra are plotted to create an ion image, often ranging from 104 to 107 pixels.1 Two common techniques used for IMS are matrix-assisted laser desorption/ionization (MALDI) mass spectrometry and secondary-ion mass spectrometry

(SIMS). Secondary-ion mass spectrometry is an ionization technique where a focused continuous or pulsed beam of energetic primary ions is used to sputter a sample surface.

This bombardment initiates the ejection of secondary ions, which can be separated according to their mass-to-charge ratio, the recording of which yields a mass spectrum.

Primary ion beams for SIMS may be grouped by two categories: atomic ion

+2 +3 + 1,4-12 beams (such as, Ga and Cs ) and cluster ion beams ( C60 and Arn). Research has

+ found that the use of Buckminister fullerene ions (C60 ) as a primary ion source has several advantages over non-cluster sources. In general, the bombardment process causes a great deal of surface damage which is a problem particularly when analyzing organic samples.6 The extent of surface damage is influenced variedly by the differing ion beam sources; for instance, differences in the interaction of an atomic surface with

+ + 13 C60 versus Ga can be clearly observed by molecular dynamic simulations. A single

2 Ga+ ion penetrates deeply into the sample and causes mixing of the many of the layers beneath the surface.10 The energy of the atomic primary ion mostly penetrates deep down into the subsurface; therefore, there is little desorption of secondary ions from the

+ surface. However, the C60 cluster ion shatters upon impact and creates an energy effect similar to that of 60 individual carbon atoms simultaneously bombarding the surface.

This action allows for a greater energy distribution closer to the surface14 which, in turn, results in greater surface disruption and ejection of secondary ions. Since fewer layers below the surface are disrupted and mixed, this allows for the opportunity for depth profiling; whereas, with single-ion bombardment, the intrinsic order of sub-surface layers is lost.

Secondary-ion mass spectrometry was first developed as a technique for studying the reactions and characteristics of inorganic surfaces but has also developed into a valuable method of analysis for organic and biological systems.15,16 Inorganic studies by

SIMS have ranged from surface studies of metal single crystals17,18,19,19,20 to surface oxidation21,22,23,24,25,26,27,28 and similar chemical reactions of metals.29,30 SIMS has been applied to a wide range of biological studies. It has been used to obtain tissue images revealing lipids,31,23,31-40 metabolites,41,42 and small proteins.43-45,46,47,48-50,51,44 Even macrophages41,52,41,53 have been successfully imaged by SIMS. These achievements alone show the range of bio-analytes which can be investigated with SIMS.

The emergence of C60 ion sources and recent advances in SIMS instrumentation allow for a wide range of biological applications. SIMS instrumentation requires high vacuum to perform optimally to avoid undesirable scattering of the primary ion beam and the secondary ions, as well as to prevent the adsorption of non-native gases to the sample

3 surface.54 Since high vacuum is required in most SIMS instrumentation, this presents a problem when attempting to analyze biological samples because the native biochemical distribution must be maintained. Water has a high vapor pressure and because cells largely contain water, an unmodified cell will immediately rupture in high vacuum conditions.55 As a consequence, the native biochemical distribution of the cell is disrupted and the vacuum pressure required for analysis may be increased unfavorably.55,56 Frozen-hydrated cellular analysis has so far yielded the most promising results for cellular analysis.57-59 In this methodology, living cells are flash-frozen to cryogenic temperatures and quickly introduced into the instrument. This allows the cells to be analyzed more closely to their native state. Unfortunately, this sample preparation is subject to problems such as excessive ice build-up on the surface and the building up charge on the sample surface during SIMS analysis.

Time-of-flight (TOF) mass analyzers are widely used in SIMS instrumentation.

They allow for simultaneous detection of all secondary ions. Prior knowledge about analytes is not necessary, thereby allowing TOF-SIMS instrumentation to be more of a discovery tool by widening the detectable mass range. The incorporation of TOF to

SIMS instrumentation has made the method highly applicable to the high mass heterogeneity and complexity of cells.60 The incorporation of large gas cluster primary ion beams has extended the achievable mass range of SIMS up to over m/z 12,000. As many SIMS facilities are incorporating gas cluster ion beams onto their existing instrumentation, the utility of the TOF mass analyzer is more crucial than ever.

The J105 Chemical Imager by Ionoptika is a relatively new instrument, which has been developed for 2D and 3D chemical imaging of biological samples. It utilizes a

+ continuous 40 keV C60 ion beam capable of rastering across a sample, allowing for submicron lateral resolution.61 The high vacuum in the J105’s stage area decreases scattering of the beam by unwanted gas particles, thus, the beam can be focused to a 250 nm diameter.61 The stage may be cooled to temperatures as low as 100 K This feature makes it well-suited to biological samples which have been flash frozen to maintain their biomolecular integrity.61 The TOF is in an axial orientation, which allows for higher sensitivity but lower mass resolution.61,62 The instrument is capable of tandem MS: a valuable tool in confirming the identity of biomolecules by their fragmentation patterns.

61,62

1.2. Research Overview

The green microalga, Botryococcus braunii, has the unique ability to consistently synthesize and excrete C25+ hydrocarbons. This inherent characteristic makes it an excellent candidate for a renewable source of biofuel. Its cells congregate in colonies constructed with a rigid, hydrophobic extracellular matrix with peculiar properties. The presence of the extracellular matrix protects B. braunii from rupture in the typical vacuum environment of a SIMS instrument. Intracellular organelles, known as ‘oil bodies,’ remain intact during a SIMS three-dimensional analysis. The long-chain hydrocarbons, similar in structure to squalene, have a vapor pressure in excess of 1 x 10-8 torr, allowing them to remain in their endogenous configurations. The proton-rich chemistry of the overall cellular colonies makes it resistant to the effects of negative

+ charging existent in the J105-Chemical Imager. C60 and large argon gas cluster primary

5 ion beams were used to identify biomolecules ranging from m/z 90 to over m/z 1600. In

Chapter 2, the properties of this cellular system are shown to make B. braunii and ideal candidate for a range of SIMS analyses.

In order to probe the three-dimensional distribution of the several unique hydrocarbons of B. braunii, it had to be established that the hydrocarbons could withstand the high-dose, continuous bombardment required of that analysis. In Chapter

3, squalene, a triterpene synthesized by B. braunii in parallel to botryococcenes, was used

+ as a model system to determine its behavior as a function of depth during continuous C60 bombardment.63,64 The [M] + peak of squalene was found to remain relatively constant in a depth profile. This behavior was predictive of the target botryococcenes of B. braunii, therefore, hydrocarbon signals during the three-dimensional cellular imaging was deemed to be a reliable and true indication of their distributions.

The hydrocarbon secretion mechanisms of B. braunii are largely unknown. The cells consistently produce and store the hydrocarbons in membrane-bound oil bodies.65

Because the oil bodies have a delicate structure, it was previously unknown which hydrocarbons they contained.66,67 The cells consistently produce and store the hydrocarbons in the oil bodies.68,69 The location of the botryococcene methylation pathway was undetermined and assumed to be somewhere between the hydrocarbons’ passage from intracellular to extracellular.63,64 In Chapter 4, SIMS has been used to answer several biological questions surrounding the secretion mechanism and the chemical composition of the oil bodies. The methylation of botryococcenes was determined to occur intracellularly by the membrane-tethered methyltransferases. The end-product of the methylation pathway was found to have two unique distributions, in

6 the intracellular oil bodies and in the extracellular matrix. The oil bodies were found to all contain the same, homogenous distribution of the end-product, C34H58. In an exciting discovery, specific lysolipids were discovered and their two distinctive distributions were observed. The lysolipids are incorporated into the membranes of the oil bodies, thus, contribution to the secretion mechanism and exported extracellularly into the extracellular matrix along with the hydrocarbons.

Trehalose is a disaccharide produced by several species of plants and animals to protect their cellular structures’ integrity in times of prolonged desiccation.70-73 It can be combined with water to create an amorphous, glassy crystal.53,73,74 The use of trehalose in SIMS analysis has ranged from being used as a model system for molecular depth profiling to the preservation of single cells for chemical imaging. 75,76-79,80-86,35,41,53,87 An in-depth analysis of the use of trehalose as a biologically-relevant matrix for SIMS analyses is described in Chapter 5.

In the first set of experiments, trehalose was determined as a biological matrix suitable for reproducible relative quantitation of purines.88 Subsequently, the previously proposed reasons for the enhancement effect of analytes in a trehalose film are investigated.79,82 The enhancement effect was found to field more than 4,000 times more signal than neat analyte preparation. H-NMR results determined that trehalose and purines do not interact in solution. But, rather the enhancement is due to the liberation of protons by the bombardment process, thus, creating a proton source for [M+H]+ formation.

A new method to vitrify mammalian cells in trehalose was developed. Spin- casting cells incubated in trehalose allows for a thinner glassy encapsulation surrounding

7 the cell. Samples remain preserved at ambient, low-humidity condition for several weeks. This feature enabled analysis across instruments and non-consecutive days.

Chemical imaging the vitrified cells revealed several purines localized to specific intracellular organelles. A three-dimensional chemical image was constructed to reveal morphological information corresponding to the chemical distributions.

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20 Chapter 2

The Use of Botryococcus braunii as a Simple Biological Model Sample for Diverse Secondary Ion Mass Spectrometry Analysis

This chapter is adapted from L.M.Jackson, H. Tian, N. Winograd, “Botryococcus braunii

as a Biological Model for Diverse Secondary Ion Mass Spectrometry,” Rapid

Communications in Mass Spectrometry Protocols. To be submitted. 2014

2.1. Introduction

Time-of-Flight Secondary Ion Mass Spectrometry (ToF SIMS) has enabled the collection of a diverse amount of data since it has been utilized as a tool in biological analyses. Yet, this success has not come without great amounts of time and attention paid to the details of sample preparation and cryogenic analysis. Biological systems often require the SIMS researcher to consider each organism’s highly complex and sensitive nature.

As the instrumental choices in SIMS advance and grow in diversity, there is a desire to balance the investigation of spatially-resolved intercellular features with extending the observable mass range. Therefore, many SIMS instrument arrangements require the use of multiple primary ion sources to achieve the goals of a single-cell and tissue analyses.1,2,3,4 A complete analysis may require comparison between several instruments or primary ion sources in a facility.5,6 However, due to the sensitive and complex nature of biological samples and the instrumental characteristics (operating

21 pressure, availability of cryogenic analysis, primary ion source configuration, etc.), it is difficult to use the same sample preparation for each experiment and instrument.7,8

Botryococcus braunii is a unique algal species with the capability of constantly producing lipids to the point that up to 85% of its dried biomass is long-chain hydrocarbons. This curious feature has made B. braunii a promising candidate as a renewable source for biofuel. Here, B. braunii is suggested as a model biological sample with a minimal sample preparation, and morphological features from sub-micron to dozens of microns with observable biomolecules over 1,000 Da. Thereby, making this sample easily amendable a range of SIMS instruments and capabilities.

2.2. Experimental

2.2.1. Algal Culture Conditions

B. braunii A race UTEX #2441 and UTEX #572, respectively, were obtained from The Culture Collection of Algae, The University of Texas at Austin. Each strain was maintained identically. The culture conditions for UTEX 2441 were previously described elsewhere, but summarized here. The strains were maintained with Bold 3N

Medium (UTEX, The Culture Collection of Algae) at 20 oC, 115 μEm light intensity on a

16:8-hour light-dark cycle under gentle agitation.

2.2.2. Algal Sample Preparation for SIMS Analyses

The sample preparation was slightly modified from a previously described

22 method and summarized here. UTEX #2441 and UTEX #572 algal samples were prepared identically. About 500 mg of algae was washed 3 x with 1-mL 0.15 M ammonium formate (Alfa Aesar, 97%) per wash to remove residual media salts. 2 μL of the cellular pellet was placed on each cleaned Si-shard. Each sample was dried for 8 minutes in a dessicator before mass spectral analyses.

+ 2.2.3. SIMS and SEM Algal Analyses with C60 Primary Ion Beam

Algal cells were analyzed at room temperature with the J105 – 3D Chemical

53 + Imager (Ionoptika, Ltd.) with a 40 keV C60 primary ion beam focused to a 300 nm diameter and sample current of 0.5 pA. Each SIMS image was collected over a 100 x

100 μm2 field-of-view with a 2.87 x 1013 ions/cm2 ion fluence at 256 x 256 pixels. SEM images were acquired with the aforementioned ion beam conditions over 512 x 512 pixels.

2.2.4. SIMS and SEM Algal Analyses with Argon Cluster Primary Ion Beam

The following analyses were performed on the vacuum dried samples at room temperature using the J105 3D Chemical Imager. The system is equipped with a 40 keV

+ C60 and a 20 keV gas cluster ion source (GCIB). This GCIB is fed with UHP Ar and

+ generates Arn (n = 1~6000) at 20 bar of backpressure. The cluster size distribution was checked by pulsing the gun and measuring the flight time spectrum of the projectile ions between the pulser and the sample surface via the ion induced secondary electron

23 emission signal. The resulting flight time spectrum can be converted into a cluster mass distribution by means of the known flight path (42.9 cm) and kinetic energy (20 keV) of

+ + the projectiles. The Ar2000 and Ar4000 were focused to ~10 µm to detect the high mass species from the UTEX #2441 algal strain. The pure Ar cluster beams were rastered across 1000×1000 µm2 /500×500 µm2 with 128×128 pixels, the total ion dose was

3×1012 ions/cm2 /1.2×1013 ions/cm2. Charge compensation was performed using 20 eV

Helium source emitting at 10 mA.

2.3. Results and Discussion

2.3.1. The Tolerance of B. braunii Cells to the Vacuum Environment

The colony-forming strain of B. braunii race A has the peculiar ability to withstand the vacuum conditions of a typical SIMS instrument at room temperature, a practically unheard of feature of a cell. In a typical cell experiment, if cells are introduced into an instrument under vacuum, they immediately rupture and the intracellular contents are spilled around the cell body.10,11 Thus, cellular preparation for SIMS requires extensive dehydration techniques or cryogenic preservation. In Figure 2-1 B and C,

UTEX #2441 cells were introduced into the instrument and secondary electron micrographs revealed that the cells remained spherical in shape and the intracellular content was not observed around the cells. The same phenomenon has been observed before with the Showa race B strain, an alga also growing in colonies with an extracellular matrix.

25 The cells of the UTEX #2441 strain are organized into a colony, bound together by a complex extracellular matrix, as seen in Figure 2-1A. Since the extracellular matrix has been known to survive extreme desiccation and fossilization, it was feasible to assume its presence was preventing rupture in vacuum.12,13,14,15 To prove this assumption, a non- colony forming race A strain, UTEX #572, which does not possess an extracellular matrix, seen in Figure 2-1D, was introduced into vacuum. UTEX #572 ruptured in the vacuum environment of the instrument as seen in Figure 2-1E and F. The intracellular contents of the cells are spilled around the single cells. Furthermore, the cell shape is puckered and shrunken to 2-5 μm diameter instead of its native spherical shape, indicating the intracellular fluid that supported the shape is absent. In Figure 2-1F, circular pools of intracellular fluid are adjacent to each ruptured cell. The pools are approximately 10 μm in diameter, which is approximately what would be expected from an 8 μm diameter cell with a 268 μm3 volume. Therefore, it is clear that the presence of the extracellular matrix prevents the cells from rupturing and losing their inherent shape.

The chemical composition of the extracellular matrix and the oil composition contribute to its protective nature. This matrix is composed of a cross-linked hydrocarbon network formed by the polymerization of the hydrocarbons and lipids, generally referred to as algaenan.16,17,18 Surrounding the cells are rigid, polysaccharidic cell walls. This structural and chemical arrangement has been linked to the preservation of labile macromolecules from decomposition and in shale deposits.19,20,21,22 In addition, after extreme decomposition and fossilization, the algaenan and cell wall structure remains.21,23 It follows that these properties contribute to the colony-forming cells not rupturing in the SIMS instrument. Furthermore, the long-chain hydrocarbons in the algae

26 can comprise of over 50% of its biomass.24 A typical SIMS instrument has a vacuum of

1x10-7 torr. The vapor pressure of squalene, a hydrocarbon similar in chain-length and structure to those of B. braunii, is 1x10-8 torr. Therefore, the hydrocarbons do not go into the gas phase in the SIMS instrument, thereby, contributing to the observation.

Furthermore, the chemical imaging experiment reveals that intracellular organelles remain intact, as will be discussed later.

2.3.2. Three-Dimensional Analysis without the Effects of Charging

When performing high spatially-resolved analysis under high vacuum with a dc beam, preventing the occurrence of charging is a great consideration. Often during single cell analysis, cells must be analyzed in a frozen-hydrated state to remain in its native structure.25,26,9 Yet, the cryogenic temperatures prevent charge build-up to dissipate and the effects of charging are amplified.27,28,29 In particular, the effects of charging can limit the detection of high mass species.29,30

The colony-forming B. braunii does not suffer the effects of charging. In Figure

2-2, a depth profile of the total ion count and a biologically-ubiquitous fragment, C7H7, was constructed from the algal region of a three-dimensional imaging experiment. The signals remain relatively stable during the depth profile, an uncharacteristic feature of room temperature analyses.31,29 There is a slight increase followed by a decrease in both signals in the middle of the experiment. This coincides with an increase in signal as the

1.0x104 total ion C H 7 7

8.0x103

6.0x103 cts 4.0x103

2.0x103

0.0 5.0x1013 1.0x1014 1.5x1014 2.0x1014 2.5x1014 3.0x1014 C + Dose (C +/cm ) 60 60 2

Figure 2-2: The absence of charging effects with depth during a B. braunii imaging experiment. The total ion signal has been divided by four to directly compare the signals.

28 hydrocarbon-rich intracellular organelles are reached and then etched away. The chemical composition of the sample is the most probable reason for the prevention of charging. Negative charge builds up on the sample in this particular instrument. The primary ion bombardment can release protons from the rich hydrocarbon environment of the sample and compensate for the negative charging. Also, because there is no need to perform cryogenic analysis, the effects of excessive charging are not incurred.

The primary ion beam technology in SIMS is evolving as the move towards biological research advances. Most SIMS research facilities require a primary ion source to provide nanometer or submicron spatial resolution.32,33,34 Yet, large cluster beams are gaining momentum to increase the mass range.35,36,37 Both of these capabilities are important to probe biological systems. B. braunii has chemical and morphological features that are particularly amendable to observation by current SIMS technology.

2.3.3. The Variety of Chemical and Morphological Features in the B. braunii Colony

There are a variety of observable features and chemicals in the B. braunii cells and colonies. Better suited to submicron primary ion beams are the intracellular organelles known as oil bodies, as chemically imaged in Figure 2-3B. They are spherical, membrane-encased organelles which store long chain hydrocarbons

38,39 intracellularly. In Race A, they contain aliphatic alkenes ranging from C25 to C31 corresponding to a mass range of m/z 320 to 460.40,41 The cells of UTEX #2441 range in size from 8 to 15 microns in length and contain 0-4 oil bodies per cell. In the total ion image in Figure

30 2-3A, the apexes are observable; they are the widest features of the cells with widths ranging from 10- 15 microns. The surface of the colony is covered in wax monoesters,

Figure 2-3C, which are in the mass range of m/z 530 to 586, completely separate from the hydrocarbons and other molecular species. The apexes of the cells covered in wax monoester would be readily observable with larger beam diameters.

Chemical imaging cells with large cluster beams is difficult because of the spatial resolution limitations; however, the algal cell colonies can be imaged well with Argon cluster beams. Figure 2-4 shows secondary electron micrographs taken with Ar4000 to exhibit the algal cells with a 5 micron beam diameter. Even at 100 x 100 μm2 field-of- view, the outlines of the cells from the substrate can be observed in Figure 2-4A. At a larger, 500 x 500 μm2 field-of-view with the same focus, images of the entire colony distribution may be observed in Figure 2-5A. The hydrocarbons at the surface of the colony are chemically mapped in Figure 2-5B. The wax monoesters can be observed clearly on the surface of the colony, as seen in Figure 2-5C. This allows the researcher to chemically map features of biological cells with large cluster beams (Arn) in conjunction

+ with beams able to achieve a sub-micron focus (C60 ).

3.3.4. The Chemical Diversity of B. braunii observable by SIMS

There are several biomolecules observable in the mass range accessible by atomic and gas cluster primary ion beams. The chemical diversity of the molecules and their structure allow for different types of analyses by SIMS. All of the biomolecules in Table

Observed Chemical m/z Formula Ion Name category + 151.06 C5H10O5 [M+H] arabinose polysaccharide

+ 181.07 C6H12O6 [M+H] galactose polysaccharide

+ 152.06 C5H5N5O ‡ [M+H] guanine nucleotide

+ 145.10 C40H39O2 [M-C29H39O2+H] lutein and beta-carotene carotenoid

+ 238.23 C16H34O2 ‡ [M+H] oleic acid fatty acid

+ + 320 - 460 CnH2n † ‡ [M] , [M+H] , … alkenes hydrocarbon

+ 530.56 C36H66O2 ‡ [M] oleyl linoleate * wax monoester

+ 558.59 C38H70O2 ‡ [M] arachidyl linolenate * wax monoester

+ 586.63 C40H74O2 ‡ [M] behenyl linolenate * wax monoester

+ 743.56 C44H80O6 [M+K] TG(12:0/12:0/17:2(9Z,12Z))* triacylglycerol

+ Table 2-1: Assorted metabolites of B. braunii race A observable by SIMS by C60 and + Ar4000 . Chemical formulas denoted by an asterisk have isomers. † denotes there are several other rearrangements for ion formation; the two most common are listed here. ‡ denotes the identity assignments supported by tandem MS.

+ + + 2-1 are observable with C60 , Ar2000 , and Ar4000 . The identities were made by searching for the accurate mass in Scripps Metabolite Database and/or Lipid Maps Database and comparing the experimental isotopic distributions with those obtained by an isotopic distribution calculator. Furthermore, several of the species assignments have been supported by tandem MS experiments on a Sciex QStar XL mass spectrometer equipped

+ with a C60 primary ion source (data not shown).

Primary ion beams near nanometer spatial resolution often have a limited mass range up to m/z 300. Polysaccharides, like arabinose and galactose are in high concentrations in the B. braunii colonies and cells walls and in the mid m/z 100s, as seen in Table 2-1. The species’ DNA is rich in G-C base pairs and guanine ionizes well with

SIMS. Carotenoids are pigments abundant in chloroplasts and yield a large common fragment, C29H39O2H, common to lutein and beta-carotene. The C18 fatty acid, oleic acid, is the starting materials for the long chain hydrocarbons of B. braunii. Because of its single double bond and aliphatic nature, it is currently being used as part of part of a model system to research the effects of aromaticity in laser post-ionization studies with

SIMS. The hydrocarbon region of the spectrum is wide from m/z 320 up to 460 representative of the multiple hydrocarbon identities and their rearrangements. There are over 15 reported hydrocarbon structures in race A algae which could be effectively resolved using tandem MS experiments.40,42 There are three major wax monoesters present in relatively high amounts on the surface of the colony in the mid m/z 500s.

These molecules have spurred exciting results as they are the largest molecules to be ionized by laser point ionization with SIMS up to this point in time. Furthermore, the

35 wax monoester and hydrocarbon distribution has already served as a model system to develop chemical image fusion technique.

Gas cluster ion beams have extended the mass range of SIMS experiments and are

+ a popular choice with dual beam analysis. Ar2000 was used to detect the presence and identity of a range of lipid classes and sub-classes, as shown in Table 2-2. The assignments were made as previously described by using the Lipid Maps Database and by comparing isotopic distributions. Diacylglycerophosphocholines (DAGs) and triacylglycerols (TAGs) were observed in the m/z 800 – 1000 region. These lipids have been reported before as involved in the biochemical process to synthesize the hydrocarbons of B. braunii race A.43,44,45 In addition, these classes of lipids are relevant to mammalian organisms, as well. DAGs and TAGs are highly-present in mammalian cell and tissue systems, especially brain tissue.46,47 B. braunii would prove a suitable system to tune an instrument for these molecules without using animal tissue and sacrifice. Multiple flavonoids were found near and above m/z 1000; these molecules are pigmentation molecules. Since the mass resolution in the high mass region is known to be lower at M/ΔM = 3,000 (versus 6,000 at lower masses), tandem MS analysis is planned to resolve the specific structures using the tandem MS capability is being developed on the instrument equipped with an Argon gas cluster source.

2.4. Conclusions

The inherent characteristics of B. braunii make it an excellent cellular system for a range of SIMS analyses. The colony-forming strains are able to withstand the vacuum

Observed Chemical Main Class Sub-Class m/z Formula

873.724 C50H100NO8P * Glycerophosphocholines Diacylglycerophosphocholines

901.429 C52H104NO8P * Glycerophosphocholines Diacylglycerophosphocholines

- C50H96NO10P * Glycerophosphoserines Diacylglycerophosphoserines

- C41H41O23 Flavonoids Anthocyanidins

1663.08 C80H146N2O33 * Neutral glycosphingolipids GalNAcβ1-3Galα1-4Galβ1-4Glc-

1078.2 C53H90O22 * Sterols Furostanols and derivatives

- C52H86O23 Sterols Spirostanols and derivatives

976.715 C65H100O6 * Triacylglycerols Triacylglycerols

918.265 C39H50O25 * Flavonoids Flavones and Flavonols

- C42H46O23* Flavonoids Flavones and Flavonols

1287.65 C58H63O33 Flavonoids Anthocyanidins

- C59H67O32 Flavonoids Anthocyanidins

Table 2-2: Lipids discoverable by Ar2000 bombardment in the higher mass region. The lipids were identified by their accurate masses. All of the Lipid Maps Database matches for each different chemical formula is listed in the table. Chemical formulas denoted by an asterisk have isomers. Mass assignments assumed to represent molecular ions, [M]+.

37 environment of a SIMS instrument at room temperature without the extensive sample preparation that is required for typical cellular analyses. Its inherent highly reduced chemical composition makes the sample resistant to the effects of charging, thus, allowing intensity-steady, three-dimensional chemical imaging. There is a variety of

+ biochemicals over 1,000 Da and morphological features observable by C60 and Argon gas cluster beams, including metabolites, nucleotides, carotenoids, and multiple classes of lipids. The B. braunii race A colony-forming strain that produces hydrocarbons from fatty acids, cellular system presents the opportunity for a range of experiments with current SIMS instrumentation capabilities and biological research objectives.

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46 Chapter 3

Understanding the Behavior of the Triterpene Hydrocarbons of Botryococcus braunii during SIMS Depth Profiling Experiments

This chapter was adapted from L. M. Jackson, J. Yoo, W. Curtis, N. Winograd.

“Three Dimensional Chemical Imaging of the Hydrocarbon Distribution of Bortyococcus

braunii.” Nature. To be submitted. 2014.

3.1. Introduction

Alternative energy research has become increasingly popular lately as fears continue to grow concerning the eventual depletion of fossil fuels available to the Earth.

Biofuels offer a means to capture energy from sunlight into chemical bonds. They are an increasingly popular alternative to fossil fuels as they are renewable, biodegradable, and a non-toxic fuel source.54 Microalgae have been suggested to be the best candidates for efficient fuel production because of their higher biomass production, high photosynthetic efficiency, and faster growth compared to other suggested biodiesel-source crops.54,55,56

Certain microalga species offer a much higher amount of energy resources in the form of hydrocarbons and other lipid molecules than other oilseed crop without having to compromise the production of foods and feed.57 Botryococcus braunii is a freshwater green microalgae capable of continuously producing an excess of long-chain hydrocarbons with some strains reaching a hydrocarbon content of 85% of its dried biomass.58 Because of this unique characteristic, B. braunii has been proposed as a promising species for the production of renewable biofuels.59

47 Botryococcus braunii has the evident ability to synthesize and accumulate a diversity of hydrocarbon oils. B. braunii strains are classified by the structures of their lipids which separate into three chemical races, A, B, and L. Algae of race A typically produce odd-carbon-numbered n-alkadiene and triene hydrocarbons from C23 to C33 derived from fatty acids;60 race L produces tetraterpenoid hydrocarbons;61,62 strains of race B produce triterpenoid hydrocarbons, C31-C24 methylated squalenes, and C30-C37 botryococcenes.60,63 Race B has stood apart from the others as the most advantageous biofuel candidate. It continuously produces the greatest quantity of hydrocarbons compared to the other races.64 B. braunii race B is an ancient organism known to date back to at least 500 MYA; its hydrocarbons already account for approximately 1.4 % of total hydrocarbon in oil shale.65-68 The chemical structures of the botryococcenes and methylated squalenes make them ideal starting materials for a hydrocracking and distillation approach to create biofuels and renewable petrochemical products at a near

97% yield.64,69 The major limitation to using B. braunii race B as a robust material for biorefinery is its relatively slow reproduction rate.70-72 Yet, the overall value in using B. braunii remains because it is able to capture more energy in its hydrocarbons than faster growing algae which produce fatty acids. In addition, the presence of the relatively thick and chemically complex extracellular matrix presents challenges for methods of hydrocarbon harvest, most of which involve sacrifice of the culture, resulting in a decreased hydrocarbon yield.73,74 There has been great interest in genetically engineering the species to have faster reproduction rates and increase the ease of oil harvest.75-77

However, to responsibly and efficiently fine tune the species, the mechanism of hydrocarbon production and secretion must be understood.

48 Our research seeks to use SIMS as a unique investigative tool to gain more information to help answer some of these questions. Here, we show that it is possible with current instrumental and sample preparation capabilities to depth profile through a hydrocarbon-rich sample where neat squalene is used as a model system, to create 2D images of a depth profile of B. braunii race B to show the distribution of specific hydrocarbons as a function of their depth, and provide insight into the process of the methylation of the hydrocarbons associated to their distribution in a multi-cellular colony.

3.2. Experimental

3.1.1. Algal Culture Conditions

The well-studied algae species Botryococcus braunii race B strain Berkeley

(Showa) was maintained using a high-density culturing strategy as previously described and briefly summarized here. The strain was cultivated in WFAM (Wayne’s Freshwater

Algae Medium) which was formulated for the maintenance of high density freshwater algae species. Cultures were grown up on a gyratory shaker at 125 rpm at room temperature and exposed to light intensities between 130 – 250 μE/m2/s. Lighting was set to a 16/8 hr light/dark cycle to simulate a 24-hr diurnal light cycle. Cultures were aerated with a humidified gas mixture of 5% (v/v) CO2 in air. In order to maintain an algal monoculture, 0.2 μm Gelman Acro gas vent filters and sterile cotton plugs were placed upstream and downstream, respectively, of each culture flask.

49 3.1.2. Algal Sample Preparation for TOF-SIMS Analyses

A 5-mL aliquot of algal suspension was deposited onto a 11 μm nylon filter paper

(Millipore) held over vacuum at approximately 1 torr. The algae were gently washed three times with 0.15 ammonium formate (Alfa Aesar, 97%) solution to remove excess media materials. The vacuum was turned off and 0.5 ml of deionized water was deposited immediately onto the algae on the filter and mixed gently. A silicon-shard (Ted Pella,

Inc.) cleaned by sonication 3x with methanol and water solutions, respectively. 10 μL of the algae/water was deposited on the shard and dried in a desiccator for 10 minutes before introduction into the mass spectrometer for analysis.

3.1.3. Squalene standard sample preparation for depth profiling by SIMS analysis

Approximately 300-μL of Squalene (Alfa Aesar, 98%) was deposited onto a 5 x 5 mm silicon-shard (Ted Pella, Inc.). The sample was plunge frozen in liquid ethane (-89 o o C) for 10 seconds and subsequently plunge frozen in a 500-mL vessel of LN2 (-210 C).

This freezing process allows for a more gradual and less violent freezing method, necessary for the liquid-state of the squalene at standard conditions. The vessel containing LN2 and the sample were transferred into the nitrogen-filled glove box connected to the vacuum chamber entrance of the J105 Chemical Imager mass spectrometer (Ionoptika, Ltd.). The squalene sample was transferred from the LN2 with long forceps and dropped into the instrument’s sample block cooled to LN2 temperature, facing squalene droplet-side up.

50 3.1.3. SIMS depth profiling conditions for squalene standard and B. braunii analysis

The SIMS analysis was done on the J105 Chemical Imager at liquid nitrogen

+ temperatures. A 0.5 pA C60 primary ion current with a 75% duty cycle on a 100 x 100

μ2 area at 32 x 32 pixels2 was used to depth profile through the frozen squalene sample at

13 + 2 a total dose of 2.35 x 10 C60 /cm .

3.3. Results and Discussion

While the ultimate goal of this study is to garner three-dimensional information concerning the distribution of hydrocarbons, it is crucial to ensure that these specific hydrocarbons can withstand the dc bombardment of the C60 primary ion. In a sample with a diversity of hydrocarbons with minimal differences in molecular structure, a controlled model system was created to observe the behavior of the hydrocarbons of B. braunii during a SIMS depth profile.

Squalene is a ubiquitously produced in plants and animals for the synthesis of sterols and steroids. In B. braunii race B, it is produced in parallel with botryococenes from the same FPS precursor. Squalene and botryococcenes are both isoprenoid triterpenes with the former having 30 carbons and the latter containing a C30 backbone with subsequent methylations from +2 to +7, therefore, squalene was the ideal molecule for the controlled model SIMS experiment.

+ Frozen squalene was subjected to a constant bombardment of C60 primary ions under the same conditions as those for the depth profiling imaging of the B. braunii. The

51 mass fragmentation pattern of squalene is shown in the spectrum in Figure 3-1.

Hydrocarbons with an alkane-nature tend to undergo random rearrangements and the carbon skeletal arrangement can become scrambled at the point of ionization. The mass assignments have been compared with the exact masses of isotopes and the most probable rearrangements of the squalene molecule, and peak identities were made by comparing the accurate mass with the exact masses of rearrangements and isotopes of squalene. These identities were made with less than 1% error. Since the mass resolution of the instrument is approximately M/Δ M = 5000, the isotopes of certain ions are isobaric with other quasi-molecular ions. Therefore, the fragmentation pattern of squalene in the SIMS instrument offers a straightforward spectrum.

The spectrum of the squalene sample was compared with the spectrum from an imaging depth profile from the algae to compare the squalene control to the actual sample in Figure 3-2. Before we could draw conclusions from the algae depth profiling data, we were interested to see if the fragmentation pattern of squalene was not convoluted by the complex mixture. Since a similar ionization pattern is observed in the B. braunii spectrum, we can assume comparable behavior for the standard and the algal sample.

Chain branching can cause a decrease in the M+ molecule; therefore, the molecular ion is more probable in hydrocarbons with reduced branching. Consistent with this prediction, the molecular ion, [M]+ , and two pseudo-molecular ions, [M+H]+ and [M-H]+, are observed in both the algae sample and standard spectrum. This validates our ability to utilize predictions of pure compound ionization patterns as basis of interpreting additional hydrocarbon signatures.

53 A.

Figure 3-2: Comparison of the squalene standard spectrum versus the same m/z range in 13 + 2 the B. braunii Race B sample. Primary ion dose for both spectra is 1 x 10 C60 /cm . (A) is the squalene standard and (B) is of the algae.

54 It was crucial to determine the squalene’s signals as a function of depth during a depth profile by ToF-SIMS. Excessive fragmentation of these long chain hydrocarbon species could lead to especially misleading chemical distributions in the z-direction. The signal of the molecular ion and two pseudo-molecular ions of squalene were plotted over increasing primary ion fluence in Figure 3-3 to observe their stability in a depth profile.

As expected in depth profiles performed at liquid nitrogen temperatures, there is a slight enhancement of signal at the initiation of the depth profile. This is due to the presence of a thin layer of ice, which has formed due to water adsorbing to the surface mostly during introduction of the sample into the instrument. The signals reach the steady state that

+ indicates that the secondary ion yield remains stable under continuous sputtering by C60 bombardment. During the introduction of the sample into the instrument, a dry nitrogen environment in the instrument’s glove box was used to prevent copious adsorption of water and, thus, ice accumulation at the sample surface. However, not all ice formation at the surface can be prevented. Due to the nature of the freezing process, the frozen squalene droplet had an amorphous surface causing the slight variation seen in the depth profile.

Figure 3-3: The squalene standard’s signals as a function of depth and continuous + bombardment by the C60 ion.

56 3.4. Conclusions

+ The results of these experiments the damage accumulation induced by the C60 bombardment is negligible and a high-quality depth profile of the squalene molecule can be achieved. The behavior of the squalene signal as a function of depth is predictive of the behavior of the multiple methylated C30+ hydrocarbons of B. braunii. Therefore, it can be said that the hydrocarbons of B. braunii will also be able to remain stable during three-dimensional imaging experiments that require the same depth profiling procedure.

3.5. References

1 Miao, X. & Wu, Q. Biodiesel production from heterotrophic microalgal oil.

Bioresour. Technol. 97, 841-846 (2006).

2 Banerjee, A., Sharma, R., Chisti, Y. & Banerjee, U. C. Botryococcus braunii: A

renewable source of hydrocarbons and other chemicals. Critical Reviews in

Biotechnology 22, 245-279, doi:10.1080/07388550290789513 (2002).

3 Li, X., Xu, H. & Wu, Q. Large-scale biodiesel production from microalga

Chlorella protothecoides through heterotropic Cultivation in bioreactors.

Biotechnol. Bioeng. 98, 764-771 (2007).

4 Chisti, Y. Biodiesel from microalgae. Biotechnology Advances 25, 294-306,

doi:10.1016/j.biotechadv.2007.02.001 (2007).

5 Metzger, P. L., C. in Chemicals from Microalgae (ed Z. Cohen) (CRC Press,

1999).

57 6 Tran, H., Hong, S. & Lee, C. Evaluation of extraction methods for recovery of

fatty acids from Botryococcus braunii LB 572 and Synechocystis sp PCC 6803.

Biotechnol. Bioprocess Eng. 14, 187-192 (2009).

7 Metzger, P., Berkaloff, C., Casadevall, E. & Coute, A. ALKADIENE-

PRODUCING AND BOTRYOCOCCENE-PRODUCING RACES OF WILD

STRAINS OF BOTRYOCOCCUS-BRAUNII. Phytochemistry 24, 2305-2312,

doi:10.1016/s0031-9422(00)83032-0 (1985).

8 Metzger, P. & Casadevall, E. LYCOPADIENE, A TETRATERPENOID

HYDROCARBON FROM NEW STRAINS OF THE GREEN-ALGA

BOTRYOCOCCUS-BRAUNII. Tetrahedron Letters 28, 3931-3934,

doi:10.1016/s0040-4039(00)96423-2 (1987).

9 Metzger, P. & Largeau, C. Botryococcus braunii: a rich source for hydrocarbons

and related ether lipids. Applied Microbiology and Biotechnology 66, 486-496,

doi:10.1007/s00253-004-1779-z (2005).

10 Huang, Z. & Poulter, C. D. STEREOCHEMICAL STUDIES OF

BOTRYOCOCCENE BIOSYNTHESIS - ANALOGIES BETWEEN 1'-1 AND

1'-3 CONDENSATIONS IN THE ISOPRENOID PATHWAY. J. Am. Chem. Soc.

111, 2713-2715 (1989).

11 Hillen, L. W., Pollard, G., Wake, L. V. & White, N. HYDROCRACKING OF

THE OILS OF BOTRYOCOCCUS-BRAUNII TO TRANSPORT FUELS.

Biotechnology and Bioengineering 24, 193-205, doi:10.1002/bit.260240116

(1982).

58 12 Largeau, C. et al. OCCURRENCE AND ORIGIN OF ULTRALAMINAR

STRUCTURES IN AMORPHOUS KEROGENS OF VARIOUS SOURCE

ROCKS AND OIL SHALES. Organic Geochemistry 16, 889-895,

doi:10.1016/0146-6380(90)90125-j (1990).

13 Derenne, S., Largeau, C., Casadevall, E., Berkaloff, C. & Rousseau, B.

CHEMICAL EVIDENCE OF KEROGEN FORMATION IN SOURCE ROCKS

AND OIL SHALES VIA SELECTIVE PRESERVATION OF THIN

RESISTANT OUTER WALLS OF MICROALGAE - ORIGIN OF

ULTRALAMINAE. Geochimica Et Cosmochimica Acta 55, 1041-1050,

doi:10.1016/0016-7037(91)90162-x (1991).

14 Summons, R. E., Metzger, P., Largeau, C., Murray, A. P. & Hope, J. M.

Polymethylsqualanes from Botryococcus braunii in lacustrine sediments and

crude oils. Organic Geochemistry 33, 99-109, doi:10.1016/s0146-6380(01)00147-

4 (2002).

15 Moldowan, J. M. & Seifert, W. K. First discovery of botryococcane in petroleum.

Journal of the Chemical Society, Chemical Communications, 912-914 (1980).

16 Tran, N. H. et al. Catalytic upgrading of biorefinery oil from micro-algae. Fuel

89, 265-274, doi:10.1016/j.fuel.2009.08.015 (2010).

17 Li, Y. & Qin, J. G. Comparison of growth and lipid content in three Botryococcus

braunii strains. Journal of Applied Phycology 17, 551-556, doi:10.1007/s10811-

005-9005-7 (2005).

59 18 Volova, T. G., Kalacheva, G. S., Zhilo, N. O. & Plotnikov, V. F. Physiological

and biochemical properties of the alga Botryococcus braunii. Russian Journal of

Plant Physiology 45, 775-779 (1998).

19 Tsukahara, K. & Sawayama, S. Liquid fuel production using microalgae. Journal

of the Japan Petroleum Institute 48, 251-259, doi:10.1627/jpi.48.251 (2005).

20 Lee, S. J. et al. Effects of harvesting method and growth stage on the flocculation

of the green alga Botryococcus braunii. Letters in Applied Microbiology 27, 14-18

(1998).

21 Moheimani, N. R., Cord-Ruwisch, R., Raes, E. & Borowitzka, M. A. Non-

destructive oil extraction from Botryococcus braunii (Chlorophyta). Journal of

Applied Phycology 25, 1653-1661, doi:10.1007/s10811-013-0012-9 (2013).

22 Niehaus, T. D. et al. Functional Identification of Triterpene Methyltransferases

from Botryococcus braunii Race B. Journal of Biological Chemistry 287, 8163-

8173, doi:10.1074/jbc.M111.316059 (2012).

23 Ioki, M., Baba, M., Nakajima, N., Shiraiwa, Y. & Watanabe, M. M. Codon usage

of Botryococcus braunii (Trebouxiophyceae, Chlorophyta): implications for

genetic engineering applications. Phycologia 52, 352-356, doi:10.2216/12-041.1

(2013).

24 Niehaus, T. D. et al. Identification of unique mechanisms for triterpene

biosynthesis in Botryococcus braunii. Proceedings of the National Academy of

Sciences of the United States of America 108, 12260-12265,

doi:10.1073/pnas.1106222108 (2011).

60 Chapter 4

Elucidation of the Metabolic Processes of Three Dimensional Chemical Imaging of the Algal Biofuel Candidate, Botryococcus braunii

This chapter was adapted from L. M. Jackson, J. Yoo, W. Curtis, N. Winograd.

“Three Dimensional Chemical Imaging of the Hydrocarbon Distribution of Bortyococcus

braunii.” Nature. Submitted. 2014.

4.1. Introduction

The green microalga, Botryococcus braunii, has garnered substantial attention as a biofuel candidate for its unique ability to synthesize and accumulate long-chain hydrocarbon oils. Despite its slow growth, it achieves carbon fluxes and productivities that can exceed those of faster growing algae. The algal colonies contain hydrocarbons in excess of 50% of its dried biomass; there are very few precedents for such a large fraction carbon flux constitutively metabolized into such a highly reduced metabolic pool of oil.

Even more amazing is the oil is not for energy storage and subsequently secreted from the cells. The metabolic pathway for hydrocarbon synthesis and secretion is a tantalizing target for heterologous introduction into fuel producing organisms; however, recapitulation of this efficient pathway appears to require more than metabolic engineering. This will require transplanting more of the biomolecular and subcellular biomolecular physiology that enables this massive commitment of carbon towards the production of what has been called bio-crude oil. This study relates to connecting the biochemistry to cellular morphology through three-dimensional chemical images towards

61 developing strategies for “physiological engineering” organisms for hydrocarbon biofuels production.

The critical biosynthetic step from farnacyl pyrophosphate (FPP) to botryococcene in B. braunii race B has been determined recently by a collaborator, Joe

Chappell. It was discovered that the biosynthetic enzymes appear to have evolved from the enzyme squalene synthase, which produces C30 triterpene sterols. The biosynthetic pathway to the extracellular oils has evolved from two homologs which act sequentially

(SSL-1 and SSL-3) where there is a third homolog that can divert carbon flux back to squalene (SSL-2) from the pre-squalene diphosphate (PSPP) intermediate.1

Botryococcene can be subsequently methylated up to 4 times to add additional branching, these methylated molecules are collectively termed ‘botryococenes.2 A recent examination of productivity in continuous culture, suggests that the in vitro enzyme kinetics are insufficient to explain the massive carbon flux.3 This suggests there is likely subcellular association of enzymes and substrates in a manner that facilitates this very hydrophobic chemistry. There is much to be learned from understanding the physiological basis of this unique biochemical capability.

In parallel with genetic and biochemical advances, there has been intensive efforts to unravel these secrets from physiochemical, microscopic and histological examination of B. braunii. Attempts to determine the location of the specific hydrocarbon species of

B. braunii have given conflicting results. Sequential extractions of the hydrocarbons indicated the majority of extracellular triterpenes are methylated, suggesting the methylation may take place during oil excretion.4 Microscopy studies have provided insight into the formation of oil bodies, including their synchronization with cellular

62 division.5,6,7, However, it is extremely difficult to make the connection between subcellular organization and biochemical analytics. This disconnect can be resolved using chemical imaging by time-of-flight secondary ion mass spectrometry (TOF-SIMS).

TOF-SIMS is capable of three-dimensional chemical imaging of several biomolecules with sub-micron spatial resolution with minimal sample preparation. The ability to combine precision rastered etching while obtaining mass spectra, presents an opportunity to conjoin biomolecular chemistry and microscopic images with spatially resolved three- dimensional chemical images.

The aggregate of the biochemical and microscopy work has led to two basic question which can be readily answered by SIMS analysis: (1) are there chemical gradients inside and outside the cells that may offer insight into the location of methyltransferases and the secretion mechanism? (2) are there characteristics of the membrane-bound vesicles that could confer localization of enzymes and enhancements in biosynthetic rates for the isoprene hydrocarbons?

4.2. Experimental

4.2.1. Algal Culture Conditions

The algal culture conditions have been described previously in section 2.1.1. of this thesis.

63 4.2.2. Algal Sample Preparation for TOF-SIMS Analyses

The algal sample preparation has been described previously in section 2.1.2. of this thesis.

4.3.3. Instrument Parameters for SEM and TOF-SIMS Algal Analysis

Room temperature analysis of algal cells was also done with the J105 – 3D

Chemical Imager, an instrument previously described in section 3.1.3. of this thesis. For

+ SEM and ToF-SIMS analysis, a primary ion beam of 40 keV C60 was focused down to an approximately a 300 nm beam diameter, as measured by line scan of an SEM image of a 300-mesh copper grid over a hole. A sample current of 0.5 pA at a 50% duty cycle was used for all SEM and SIMS analyses. The SEM images were acquired at 512 x 512

+ pixels. SEM imaging was done with the focused C60 primary ion beam at the same conditions used for TOF-SIMS imaging. Subsequent images were obtained with the aforementioned conditions over the identical sample area, thus, allowing for the depth profiling conditions required for three-dimensional analysis of the algal sample. Each image was acquired with an ion fluence of 4.55 x 1012 ions/cm2, therefore, the top image was acquired with 4.55 x 1012 ions/cm2 and the image at the intracellular level was

14 + 2 2 obtained with 3 x 10 C60 /cm . Each SIMS analyses were acquired on 100 x 100 μm area at 256 x 256 pixels2. These conditions for the SEM and SIMS analyses, thus, featured a pixel size of 290 and 590 nm/pixel, respectively. All SIMS analyses were obtained with a +10 V stage bias.

64 Based on a comparison of the intracellular features revealed with depth in the chemical images with previously obtained confocal microscopy images [references of papers which show microscope images with scale bars] it has been assumed that the

14 + 2 fluence of 3 x 10 C60 /cm corresponds to reaching a depth of approximately 1 – 2 μm, dependent on the characteristics of the sampled area, by depth profiling with SIMS.

4.3. Results and Discussion

Different hydrocarbons are shown to have unique distributions in the colony most specifically with depth. Figure 4-1 compares the difference of hydrocarbon presence and diversity on the outside of the colony versus intracellularly. While there are very few hydrocarbons observed on the outside of the colony, the overall quantity and diversity of hydrocarbons observed increased significantly.

Since very few hydrocarbons are on the surface, these may represent end- products of the methylation process. The increased diversity of hydrocarbon species intracellularly suggests that within the cells, there are additionally more intermediates.

This observation would imply that the methylation process is occurring intracellularly.

However, to be certain, it is necessary to map the distribution of an end product.

In Figure 4-2, the chemical distribution of the tetramethylated botryococcene, C34H58, has been mapped in three dimensions. First, the total ion image offers insight into the meaning of its distribution. The total ion image of the surface of the colony shows a continuous coating over the bound cells underneath. This continuous coating can be

Figure 4-1: The overall diversity of hydrocarbons increase with depth. (A) and (B) are 13 + 2 spectra from the top surface layer of the colony obtained with a dose of 1 x 10 C60 /cm . (B) and (D) are zoomed into the m/z 300 – 600 region of the spectrum in (A) and (C), respectively. (C) and (D) are spectra from the intracellular region of the cells within the 14 + 2 colony obtained after a dose of 3 x 10 C60 /cm corresponding with a depth of about 1 - 2 μm deep into the colony.

67 inferred to be the ubiquitously-observed extracellular matrix, containing a cross-linked hydrocarbon network. 8 The bound cells are about 5-10 μm in diameter and appear as spherical protrusions; where each protrusion accounts for the apex of a cell underneath the extracellular matrix. After depth profiling 1 – 2 μm deep into the colony, several smaller circular features approximately 1-3 μm in diameter are shown to be clustered within the regions associated with the individual cells. These small circular features have a relatively brighter intensity than their surroundings, thus, implying a physical barrier on the outside of the features. The features correspond to the presence of oil bodies, which have been reported to localize near the apex of each cell.9

Previous studies have questioned whether the distribution of hydrocarbons is homogenous or gradational through the oil body and whether there are different contents between oil bodies within each cell.5,10 The 1-3μm circular features that show brightly in the total ion image can be assumed to correlate to the hydrocarbon contents of the oil bodies. Assuming squalene’s ionization potential is directly comparable to that of botryococcenes, as previously learned by the SIMS depth profile, and the other like molecules of race B. Therefore, since the spots are homogenous in the total ion image, we assume that they all fully contain methylated botryococcenes, an observation also mirrored by Weiss.10 However, we can go one step further to conclude that since these features in the total ion image match the same areas where C34H58 is localized, the distribution of C34H58 is homogenous throughout the oil body, a novel observation. There are several cells shown in the chemical image and they all contain these identical features. Here, the hydrocarbon content between oil bodies in the same cell and oil

68 bodies between different cells are confirmed to have the same homogenous C34H58 distribution, whereas, it was previously unknown if these distributions were different.

C34H58 is distributed homogenously on the outside of the colony and suggests the tetramethylated botryococcene has been secreted into the extracellular matrix. This distribution agrees with hydrocarbon extraction studies which report that over 99% of the

11,12 tetramethylated botryococcene, C34H58, is present in the extracellular matrix. In the

SIMS images, relatively, there is a greater amount of C34H58 outside of the cells; however, there is also a considerable distribution within the intracellular oil bodies.

From this data, it can be implied that there is much more than 1% of the total C34H58 is found intracellularly. This observation suggests that the oil bodies participate in temporary storage of C34H58 before excretion into the extracellular matrix.

Previously, there was confusion over whether the methylation process occurred intracellularly or extracellularly.2,13 The previously-held belief was that since the methyltransferases are membrane-bound and over 99% of the end product is located extracellularly, the triterpenes might be methylated during passage through the cell membrane out of the cell. However, since the chemical distribution of C34H58 indicates that this end-product is present in large amounts intracellularly, the location of the methylation process must also be intracellular and not as the triterpenes are exported extracellulary.

The observation of intracellular methyltransferases activity leads to the question as to where their specific intracellular location is. An obvious candidate would be in association with the oil bodies to provide the hydrophobic chemical environment required for botryococcene synthesis. It is reasonable to speculate that the membrane-bound

69 methyltransferases might be attached to the membrane of the oil bodies to take advantage of the hydrophobic environment. Since the substrate and end-product are both localized in the oil bodies, a co-localization with the biosynthetic enzymes would facilitate efficient bioconversion.

Botryococcus braunii has a nearly unprecedented ability to produce a high yield of oil comprising of a major part of its biomass during active growth, yet, the recent assessments of chemical kinetics are not indicative of the observed biosynthetic capacity.

The step to create botryococcene triterpenes from FPP is catalyzed by SSL-1, yet, this

1 enzyme has a rather low reported kcat at 0.027 mole product/mole enzyme/s. Previously, we have determined that this kcat would imply that there is an unreasonably high absolute value of enzyme per cell.14 There, it has been suggested that these hydrophobic substrates are channeled in a metabolon, thus, considerably less of the enzyme is necessary per cell to achieve the observed high rate of botryococcene triterpenes production.14 The formation of metabolons to facilitate growth-dissociated lipid accumulations of approximately 70% of biomass has been reported in some oleaginous species of eukaryotic micro-organisms, many of which have also been identified as potential biofuel sources.15,16 Perhaps, the only way to achieve accumulations this high is through the use of metabolons.

It is assumed the environment for the metabolon is hydrophobic in nature, however, the colonies have several unique hydrophobic features, a specific area of formation is unclear by this logic alone. However, since we have determined that the end product of the botryococcene pathway is located in the oil bodies, we propose that metabolon formation occurs in the membrane-enclosed oil body as it provides an

70 excellent environment for the membrane-bound methyltransferases to channel substrates to achieve the noticeably high degree of botryococcenes synthesis.

Since biochemical and microscopic evidence logically converge to the enclosed oil bodies, the SIMS analysis was focused on discovering the biochemical nature of the interfaces between the cytoplasm and the intracellular oil storage. The microscopic and kinetic data are suggestive of an important role for the membrane that encloses the intracellular lipid bodies. This region of the chemical imaging data explored for polar lipids which could effectively surround the oil as a monolayer.5 Two different species of lysolipids were readily observable and identified in the SIMS mass spectrum. They were identified by using the Lipid Maps Database to match the experimentally observed masses to the exact masses of the corresponding lipids within 5 ppm. To further confirm the identities, the isotopic patterns of the lysolipids, shown in Figure 4-3, were compared to the predicted spectra obtained by entering the chemical formulas into an isotope distribution calculator. The observed masses of m/z 505.362 and 523.359 were identified as the molecular ions, [M]+, of 1-(1Z,9Z-octadecadienyl)-sn-glycero-3- phosphocholine and 1-octadecanoyl-sn-glycero-3-phosphocholine , respectively. Their structures are characterized as having a phosphocholine head group and a single-chain hydrocarbon tail, as shown in Figure 4-4.

While lipids are typically associated with the cell membrane bilayer and other membranes of other intracellular organelles, the lysolipids in race B have two distinct intracellular and extracellular distributions. In Figure 4-5, the lysolipids are homogenously distributed on the surface and throughout the extracellular matrix. When the intracellular level is reached, the lysolipids have different distributions; here they

A. LPC(p18:0/0:0)

m/z 505.353

B. LPC(18:0/0:0)

m/z 523.363

Figure 4-4: The chemical structures of the two lysolipids of B. braunii race B identified by SIMS. Each lysolipid is characterized by its zwitterionic nature, hydrophobic tail, and a phosphate group.

A. B.

74 form small rings of 1 – 3 μm in diameter corresponding to the same order of size as the oil bodies. This suggests that the lysolipids are integrated within the membrane that encapsulates the oil bodies. It has been previously reported that the oil bodies have a phospholipid membrane.5 The presence of lipids in the phospholipid and lysolipid classes have been found in lipid profiling experiments of race B lipid extracts, however, information regarding the distribution and function of lysolipids in the algae have not been reported.17 Here, information concerning the function of lysolipids can by inferred by their complementary distributions with the end-product botryococcene, C34H58. Their distributions are indicative of a process that must move intracellularly to extracellularly.

This close association of the lysolipids and hydrocarbons would be consistent with a role for lysolipid involvement in the excretion of hydrocarbons from the cell.

In the determination of the connection and motions of hydrocarbon and lysolipids through the cells and colony, the cell cycle uniquely characterized by the appearance and disappearance of oil bodies must be examined. With increased maturation of the cell, the lipid bodies increase in number and size.5 When a mature cell enters into mitosis, the multitude of lipids bodies it had essentially disappear.5 During this virtual absence of observable oil bodies, droplets of oil have been seen to accumulate in the septum between and around the daughter cells.5 It is assumed that the contents of the oil bodies have been excreted from the mother cell in a mechanism that is not clear. However, the SIMS data shows that in addition to this oil being excreted, it appears that the lysolipids incorporated into the membrane are also excreted into the extracellular matrix. The process we will propose here matches with these previous observations of hydrocarbon movement with cell division.18

75 Lysolipids have the unique ability to be incorporated into membranes at nearly lytic concentrations, yet, they have the power to breakdown the integrity of lipid bilayer or make it susceptible to breakdown.19,20,21 There is chemical evidence here that the lysolipids in race B play a crucial role in hydrocarbon excretion as explained by Figure 4-

6. Because the lysolipid distribution correlates with a membrane around the oil body contents, this implies that the lysolipids are present in the membrane. As lysolipid concentration in a membrane increases, the breakability of the membrane increases; at high lysolipid concentrations, the membrane can be broken by small amounts of force or even temperature fluctuations.20 At cell division, the septum formation induces an intracellular positive pressure.22 This pressure increase can provide enough disturbance, thus, activation energy to initiate a defect to break down the oil body membrane. After the oil body membrane is broken, the lysolipids and other membrane lipids mix with the hydrocarbons. This coincides with the observation that as septum formation begins, the oil body structures effectively disappear and form one large droplet of oil.5 At this stage the lysolipids would be present as free monomers and at high lysolipid concentrations they can form micelle.21 Their aggregation into micelles through the hydrocarbon droplet is facilitated by the lysolipids’ zwitterionic nature. However, since the lysolipid concentration must have been high to enable breakability by positive pressure, their concentration would be high enough for micelle formation.21,20 The micelles can interact with the cell membrane and create a pore.23 Since there is still positive pressure inside the cell, the first pore created by the micelles allows the

77 hydrocarbon and lysolipid droplet is the pore by which together they are excreted into the extracellular matrix. The cell membrane can close after excretion. Each newly formed daughter cell is absent of the presence of oil bodies.5

4.4. Conclusion

We have used this methodology to determine novel information regarding the hydrocarbon synthesis and distribution within the cells and the colony of B. braunii. (1)

The methylation process occurs intracellulary, therefore, the membrane-bound methyltransferases are confirmed to act inside the cells and not as the triterpenes are exported extracellularly. (2) The end-product of the methylation process, C34H58, has been determined to be present in great abundance both in the extracellular matrix and inside the intracellular oil bodies. (3) The biochemical nature of the oil bodies within the cells and between cells has been explored: oil bodies within the same cell and in comparison to other cells in the colony have been found to contain the same contents, homogenously and without a gradient. (4) The oil bodies have been found to primarily contain C34H58 and offers support to the hypothesis that the enzymes involved in methylation form a metabolon to accomplish fast rates of C34H58 synthesis. (5) Specific lysolipids of race B B. braunii have been identified for the first time. Two distinct distributions within the organism have been determined. Intracellularly, they are shown to contribute to the lipid membranes encapsulating oil bodies. Yet, throughout the extracellular matrix, they are present in relatively large quantities. These findings

78 suggest that the oil body contents and its membrane are both excreted from the cells to the extracellular matrix.

4.5. References

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81 21 Needham, D. & Zhelev, D. V. Lysolipid exchange with lipid vesicle membranes.

Annals of biomedical engineering 23, 287-298 (1995).

22 Charras, G. T., Coughlin, M., Mitchison, T. J. & Mahadevan, L. Life and times of

a cellular bleb. Biophysical Journal 94, 1836-1853 (2008).

23 Siegel, D. P. Energetics of intermediates in membrane fusion: comparison of stalk

and inverted micellar intermediate mechanisms. Biophysical Journal 65, 2124-

2140 (1993).

82 Chapter 5

Trehalose as a biologically-relevant matrix for secondary ion enhancement and quantitative detection of purines for SIMS analysis

This chapter has been adapted from L.M. Jackson, J.J. Hue, N. Winograd,

“Quantitative detection of purines in biologically-relevant films with TOF-Secondary Ion

Mass Spectrometry,” Applied Surface Science (2013), 45, 237-239.

5.1 Introduction

Trehalose is a disaccharide synthesized by a range of plants and animals to protect their cellular integrity during long periods of desiccation. The anhydrous molecule has 8

–OH groups and will readily form a dihydrate structure. As a result, it promotes an active hydrogen-bonding network, making it able to interface well with water and biological structures. This characteristic allows trehalose to effectively mimic the interactions of water with cell membranes and proteins even when trehalose has formed an encapsulating, amorphous crystal. The ability of trehalose to preserve aqueous biochemistry makes it an excellent matrix for biological samples. Here, this disaccharide will be exploited for its ability to easily lend these qualities to biological analyses with

SIMS.

A method of relative purine quantitation in biologically-relevant matrices was developed to eventually make conclusions about the de novo purine biosynthetic pathway during a three-dimensional analysis of a cell. Two approaches are considered. In the first instance, inosine monophosphate (IMP) standard solutions were doped into a

83 trehalose spin-cast thin film. Trehalose has been previously shown to be a successful matrix for depth profiling of peptides. 6-7Since IMP is readily converted into adenosine monophosphate (AMP), AMP-doped trehalose spin-cast standards were created to demonstrate linearity, specificity, and simultaneous detection for purines related to the pathway. In a second instance, spin-cast films of a HeLa cell homogenate, spiked with known concentrations of IMP resuspended in trehalose were shown to yield quantitative concentration information. The results show the feasibility of simultaneously determining analyte concentration of biomolecules, a powerful approach for cellular analysis.

Trehalose spin‐cast films as a matrix and trehalose encasements for cells have been shown to enhance secondary ion yields of biomolecules.1,2,3 It has been suggested that the molecular ionization efficiency is attributed to the availability of protons from the

+ water molecules throughout spin-cast trehalose films by C60 bombardment during molecular depth profiling.3 It is unclear whether there are preformed ions in solution or in the trehalose film.4 Furthermore, the bombardment process has been suspected to induce the liberation of protons from water molecules.5,3 The enhancement effect of trehalose will be investigated in a multi-instrument analysis to discern its basis.

Here, a method of trehalose vitrification by spin-casting is developed for SIMS to enable three-dimensional imaging of mammalian single cells. Purines and other biomolecules were imaged intracellularly with sub-micron spatial resolution. Trehalose vitrification is a technique first developed for clinical purposes to enable cellular preservation without the use of cryogenic techniques.6,7 As a result, the method established here allows cells to be preserved in their native state for several months at

84 ambient conditions. Insight into trehalose’s entry into the cell was observable through imaging depth-profiling experiments.

5.1.1. The de novo Purine Biosynthetic Pathway and Associated Metabolites

In purine-depleted environments, the de novo purine biosynthetic pathway is catalyzed by six enzymes in ten biochemical steps to ultimately produce inosine monophosphate (IMP), a ribonucleotide of hypoxanthine which can serve as a precursor for other purines.8 While it has been hypothesized that these enzymes interact to form a

“cluster” to facilitate substrate channeling and regulate the flux of metabolites, the

Benkovic group has clearly observed that all six enzymes colocalize to create cluster formations in the cytoplasm of the cell in purine-depleted environments presumably to synthesize a purine, IMP, via the de novo purine biosynthetic pathway.8 They have termed this cluster the “purinosome.”8 They have used fluorescence microscopy facilitated by fluorescently labeling the six enzymes to witness the aforementioned observation.8 In addition, with this methodology they have witnessed the association and dissociation of the purinosome by dynamically altering the purine levels of the media to provide evidence for the formation of a multi-enzyme complex, the purinosome.8

Abnormal regulation of the de novo purine biosynthesis is related to various human diseases, including immunodeficiency, myopathies, and others.9,10 Therefore, the purinosome may be considered as a pharmacological target for therapeutic treatment.11,9

Extensive chemical information concerning the purinosome is sparse and details about which subcellular compartment contains these enzymes is unknown.11,9 The most

85 direct way to prove and trace dynamic action of the purinosome, chemically, is to determine the associative distributions of IMP, the sole product of the pathway.

Ultimately, the distribution of IMP could be conveyed in a three-dimensional chemical representation to provide evidence of purinosome function. However, it is clearly a challenge to develop analytical methods to measure micromolar concentrations of IMP at the single cell level.

While IMP is the only purine whose presence after a purine-depleted environment is the only molecular evidence of the de novo purine biosynthetic pathway,8 knowledge of the distribution of other purines could serve to benefit and strengthen the IMP distributions obtained. For example, an environment nearly free of all purines could be ensured by the absence or near absence of their signals, not just that of IMP. In another instance, in an environment, which would affect purinosome disassembly, the distribution of other purines and their relative concentrations to that of IMP can be determined.

Chemical information of other purines could be detected easily their mass signature would likely be picked up during the analysis of IMP. Many biologically common purines can be identified during a TOF-MS analysis, including adenine, thymine, isoleucine, hypoxanthine, and guanine.

86 5.2. Semi-Quantitative Detection of Purines

5.2.1. Experimental

5.2.1.1. Instrumental Parameters

The experiments to create a model system for the quantitation of purine were performed on a TOF-SIMS instrument described previously elsewhere.12 Briefly, the

+ instrument is equipped with a C60 primary ion source by Ionoptika Ltd. (Southampton,

13 + U.K.). For depth profiling, a dc 100 pA beam of 40 keV C60 was utilized to sputter through the spin-cast films with a raster area of 500 µm x 500 µm in 4 s intervals.

Between these erosion cycles, negative ion mode TOF-SIMS mass spectra were obtained

10 + -2 at an ion fluence of 3 x 10 C60 ions cm into a raster area of 200 µm x 200 µm within the eroded area. The primary ion beam was pulsed for 100 ns at 3000 Hz.

For the instance concerning the purines doped in trehalose, the magnitude of the purine signal was determined by integrating the peak area at half-width and half- maximum at the corresponding mass value for each mass spectrum acquired during the complete depth profile – from the surface of the film to the Si interface. Because the spin-casting preparations are reproducible in terms of thickness and surface homogeneity, this allowed the comparison of the integrated purine intensities of the depth profiles for the different concentration standards, respectively.14 For the more biologically-relevant instance concerning the analyses with cell homogenate, the value of “total counts” was calculated by taking the integrated peak area at half-width and half-maximum of each

87 sample, respectively, during a surface spectrum from a 150 x 150 µm2 area with a

12 + -2 primary ion dosage of 1.0 x 10 C60 cm .

5.2.1.2. Cell Homogenate with Purine-Doped Trehalose Sample Preparations

HeLa cells, an immortal cervical cancer cell line, were grown in a T-75 flask using Dulbecco’s Modification of Eagle’s Medium (DMEM) (Cellgro), supplemented with 10% Fetal Bovine Serum (Cellgro), and a penicillin-streptomycin solution (10,000

I.U. penicillin, 10,000 μg/mL streptomycin) (Cellgro). Cells remained in an incubator maintained at 5% CO2 and 37 °C, until 100% confluence.

The homogenization protocol was adapted from Core G Subcellular Fractionation of RAW264.7 Protocol PP00004301 to suit the HeLa cell line. To detach cells from the flask, 1X Trypsin-EDTA (Cellgro) was added. After centrifugation, the growth media was removed and cells were resuspended in isolation media, [250 mM sucrose (Sigma, ≥

99.5%), 10 mM HEPES (Cellgro), 1 mM EDTA (EMD, 99.0-101.0%), nanopure water, at pH of 7.4)], and centrifuged at 1000 rpm for 5 minutes. Isolation media were removed, cells were resuspended in hypotonic media (100 mM sucrose (Sigma, ≥ 99.5%), 10 mM

HEPES (Cellgro), 1 mM EDTA (EMD, 99.0-101.0%), at pH of 7.4), and centrifuged at

1000 rpm for 5 minutes. The supernatant was discarded. The cellular pellet and 2 mL supernatant was homogenized, centrifuged at 1000 rpm for 10 minutes.

Following this step, two different routes of sample preparation were performed.

First, for the IMP-doped cell homogenate/sucrose solutions, the pellet was resuspended in

88 the hypotonic media, 5 µL drops of this solution were dried on Si shards, and dried under a stream of nitrogen.

5.2.1.3. Purine-Doped Spin-Cast Film Preparations

Spin-cast films were prepared with 0.125 M aqueous D-(+)-trehalose dehydrate

(Sigma, ≥ 99%), solution. Pre-sliced 5 mm x 5 mm Si wafers (Ted Pella Inc.) were used as substrates for all films. All water for the preparations was purified by a Milli-Q

System (Millipore, U.S.A.) with a resistivity of 18.2 MΩ cm and an organic content less than 5 ppm. The Si wafers were prepared by a procedure described elsewhere.14

The respective IMP- and AMP-doped trehalose solutions at varying concentrations were prepared via serial dilutions. Solid IMP (Sigma, ≥ 99.5%) and AMP

(Spectrum, ≥ 99%), respectively, were dissolved in the trehalose solution to make a stock solution. Different aliquots of this solution were diluted with trehalose solution to make the concentration standards.

For the IMP-doped cell homogenate/trehalose spin-cast films, the hypotonic media was removed and the homogenate pellet was re-suspended in 8 mL of trehalose solution. The resulting solution was separated into two aliquots. Solid IMP was dissolved in the first to create a stock solution. This stock solution was diluted serially with the remaining aliquot to create the concentration standards.

Concentration standards of IMP and AMP were confirmed with a NanoDrop 1000 spectrophotometer (R2 = 0.999) at 223 and 227 nm, respectively. All of the spin-cast films were created with 250 µL of the respective solutions. They were spin-cast onto the

89 prepared wafers at 5000 rpm. First, approximately a 10 µL drop of the solution was dropped onto the stationary shard, spinning was then initiated, and the process repeated.

5.2.2. Results and Discussion

Spin-cast films of trehalose doped with varying concentrations of IMP were prepared and analyzed as a biologically-relevant model to determine if differences in IMP concentration could be detected in a defined volume. The concentration of IMP in a

HeLa cell is approximately 100 µM (6 x 1016 IMP molecules/cm3).1 As demonstrated in

Figure 5-1a, using the IMP-doped spin-cast film standards, IMP can be precisely determined well into the low nanomolar regime. This experiment has been reproduced successfully in triplicate with a coefficient of determination, R2, greater than 0.99 at each instance. This result implies that trehalose is an excellent matrix for reliable quantitation between spin-cast films containing nanomolar concentrations of IMP and, potentially, other purines. Similar to related work with biomolecules6-7, it appears trehalose offers enhanced stability to the purines during the primary ion bombardment. The combination of these features lends to the belief that IMP and other purines could be detected within a cell if matrix effects, which currently prevent this type of analysis, are overcome.15,16,17

While IMP is the only purine whose presence after a stage of purine depletion would be the only molecular evidence of the de novo purine biosynthetic pathway, it is valuable to monitor the distributions of other purines during the process.11 As demonstrated in Figure 1b, AMP was examined using the same protocol as for IMP in the same nanomolar concentration range. With this capability, it should be feasible to

Figure 5-1: The relative quantitation of the (a) IMP- and (b) AMP-doped trehalose spin- cast films is shown. The coefficient of determination is (a) R2 = 0.992 and (b) R2 = 0.999, respectively.

91 monitor the rate and distribution of the conversion from IMP to AMP during purine replenishment.

As shown in Figure 5-2, the mass signature peaks for [IMP-H]- and [AMP-H]-, respectively, can be resolved within a mass spectrum of a trehalose spin-cast film doped with identical concentrations of IMP and AMP, despite an insignificant amount of isotopic overlap. When observing this spectrum, it should be noted that the trehalose peak, [trehalose-H]-, is present at m/z 341, closely within the mass range of IMP and

AMP. It has a similar intensity to the purine samples at nanomolar concentrations. This peak provides a convenient reference and internal standard. Tracking the intensity of this peak between depth profiles of different spin-cast films serves as an indicator of film homogeneity between samples and possible primary ion beam current fluctuations.

While trehalose is an accepted biologically-relevant model matrix, nanomolar concentration standards of IMP were created in a cell homogenate solution to serve as a matrix most similar to the cellular environment. As noticed from Figure 5-3A, the correlation reflects greater variability between concentrations compared to pure trehalose.

This result is most probably a result of the characteristic surface inhomogeneity of dried cellular homogenate. In addition, there is chemical noise associated with the homogenate that can generate isobaric interferences. While qualitative data may be obtained, there is excessive variability making obtaining a reproducible result difficult. However, when the cellular homogenate is resuspended in trehalose solution and spin-cast, as seen in Figure

5-3B, the results are much improved. The slight variability is suspected to be introduced by matrix effects often caused by excessive salts in cellular growth media.18 Similar to previous spin-cast trehalose films, the homogenate films have a glassy appearance, thus,

Figure 5-2: The mass signature peaks for [IMP-H]- and [AMP-H]- are resolved here within a mass spectrum of a trehalose film doped with IMP and AMP, respectively.

94 lending to a more uniform surface, which was key to the correlation and repeatability of the experiment. IMP-doped cell homogenate/trehalose spin-cast films could provide a standard technique to determine the concentration of a certain biomolecule in a cell or any other defined biological volume.

When trehalose is used as a matrix the number of intact purine molecules is enhanced relative to when neat IMP is bombarded directly, an observation which will be probed later in the chapter. The trehalose matrix appears to be linked to the prevention of purine molecule fragmentation. Thus, both purine quasi-molecular ions are formed, presumably by the loss of a proton from the parent molecule. The purine molecules were completely consumed in the depth profile with little fragmentation indicated by the major fragment of IMP and AMP, hypoxanthine as [M-H]- present at m/z 135. The purine quasi-molecular ions appear to be protected from fragmentation largely by the trehalose matrix; perhaps, similar in theory as to how MALDI matrices prevent excessive fragmentation of high-mass molecules. In addition, despite damage accumulation characteristically incurred by depth profiling, the integrated purine signal is reproducible and gives a linear response to varying concentration.

5.2.3. Conclusions

These concentration studies provide promising evidence that IMP concentration can be seen at biological levels relevant to the dynamic actions of the de novo purine biosynthetic pathway. Biologically relevant concentration standards can be analyzed quickly on the day of cellular analysis to obtain relative concentration data. Trehalose

95 proves to be a matrix essential to observe quasi-molecular purine ions linearly in the presence of homogenate, in regards to concentration. These results suggest the possibility of temporal studies for the distribution and concentration of purines in a biological sample.

96 5.4. Determining the Enhancement Effect of Trehalose for Purine Detection

5.4.1. Experimental

5.4.1.1. SIMS Instrumental Parameters

Positive and negative SIMS analyses were acquired with the Bio-ToF mass

+ spectrometer equipped with a 40 keV C60 primary ion source by Ionoptika Ltd.

(Southampton, U.K.) at a 40o angle relative to sample normal, previously described elsewhere.12

+ 2 A 100 pA beam of 40 keV C60 was rastered across a 2000 x 2000 µm area with

100 ns pulses at 3000 Hz and 300,000 shots were collected per mass spectra. All spectra

-9 + 2 were taken with an ion dose of 1.6 x 10 C60 /cm , within the static limit.

+ For depth profiling experiments, a dc 100 pA beam of 40 keV C60 was utilized to sputter through the spin-cast films with a raster area of 500 x 500 µm2 in 4 s intervals.

Between these erosion cycles, mass spectra were obtained at an ion fluence of 3 x 1010

+ -2 C60 ions cm into a raster area of 200 µm x 200 µm within the eroded area. The primary ion beam was pulsed for 100 ns at 3000 Hz. Three trials were obtained per sample.

5.4.1.2. NMR Instrumental Parameters

All 1-dimension H-NMR spectra were collected on a Bruker DPX 300 MHz spectrometer.

97 5.4.1.3. Sample Preparation for SIMS Analyses

The neat IMP sample was prepared by gently pressing 800 mg of solid IMP

(Sigma, ≥ 99.5%) with a metal spatula into 0.1 mm thick Indium foil (99.99%, Sigma) until the foil was nearly covered with sample.

To observe the behavior of IMP for the enhancement study, the spin-cast films were prepared with 0.125 M aqueous D-(+)-trehalose dehydrate (Sigma, ≥ 99%) solution doped 0.1% volume IMP. Pre-sliced 5 mm x 5 mm Si wafers (Ted Pella Inc.) were used as substrates for all films. The Si wafers were prepared by a procedure described elsewhere.14 All water for the preparations was purified by a Milli-Q System (Millipore,

U.S.A.) with a resistivity of 18.2 MΩ cm and an organic content less than 5 ppm. The anhydrous trehalose film doped with 0.1% IMP and prepared with anhydrous trehalose

(Sigma, ≥ 99%) and D2O (99.8%, Cambridge Isotope Laboratories, Inc.).

All of the spin-cast films were created with 250 µL of the respective solutions as previously described in section 5.2.1.3.

5.4.1.4. Sample Preparation for NMR Analyses

Each sample was ran in a 5-mL NMR tube. The IMP and anhydrous trehalose standards, respectively, were prepared by dissolving 500 mg of analyte to 1 mL of D2O

(99.8%, Cambridge Isotope Laboratories, Inc.) before pouring the solution into the tube.

The IMP and anyhydrous trehalose combined sample included 500 mg of each dissolved into 1 mL of D2O.

98 5.4.2. Results and Discussion

Through an approach utilizing ToF-SIMS supplemented by H-NMR, the aim of the following set of experiments was to determine if the ionization enhancement of IMP is due to preformed ions in a aqueous trehalose spin-casting solution or the

+ mixing of the film’s components by the C60 cluster ion/solid interactions.

5.4.2.1. Ionization of Inosine Monophosphate Solid versus Doped into a Trehalose Film

The ionization efficiency of IMP appears to be directly linked to its presence in a spin-cast trehalose film. When neat IMP solid is analyzed by SIMS, a nearly negligible

[M+H]+ is obtained, as seen in Figure 5-4A. The signal-to-noise ratio of the resulting peak is well below 10; the signal is considered to be at the detection limit, as seen in

Figure 5-4A. IMP is more likely to lose a proton to achieve a negative quasi-molecular ion than it is to gain an ion. This tendency is observable in the positive ion spectrum in

Figure 5-4A where the [M+H]+ is just over its detection limit, whereas, in Figure 5-4C the [M-H]+ ion is more readily achieved.

However, when less than 0.1% IMP was doped into a trehalose film, the [M+H]+ was enhanced by a factor of 4,750 and the [M-H]- by 125, as observed in Figure 5-4B and

D, respectively. The [M+/-H]+/- enhancement was calculated as follows

100

where I[M+/-H]+/- is the intensity as peak area for IMP from the analysis of the film or the solid, [nIMP,film /ntrehalose,film] is the molar ratio of IMP to trehalose in the spin-cast film.

Linear signal response is assumed due to previous research.19

Since the enhancement in negative ion mode is less pronounced than that in positive ion mode, it leads to the assumption that the trehalose crystal matrix provides a source of protons to enhance [M+H]+ formation. The minor enhancement in negative mode could be attributed to the trapped water molecules being excellent acceptors of protons; whereas, in the neat sample, the proton acceptors are not as strong. The enhancement is more prominent in positive mode as the bombardment of trehalose releases protons, thus, creating a proton source for [M+H]+ formation. However, it is unclear whether the enhancement is due to protons released from the trehalose matrix or from the water-trapped molecules within the amorphous trehalose crystal matrix.

5.4.2.2. The Chemical Interaction of IMP and Trehalose as Examined by H-NMR

The goal of the H-NMR experiments was to determine if the IMP and trehalose interact with each other in a solution virtually free of water molecules. With this preparation, we can determine if proton exchanges occur between the two molecules leading to pre-formed ions in solution responsible for the enhancement effect.

101

Solutions of D2O with IMP, trehalose, and a mixture of the two compounds, respectively, were analyzed by H-NMR. In Figure 5-5 A and B served as controls to determine the H-NMR signature of IMP and trehalose, respectively. These spectra were combined by computer software to create a predicted spectrum, Figure 5-5 C, representing IMP and trehalose not interacting in solution. Expected interactions were protonation of the IMP molecule and a rearrangements of the –OH groups of trehalose.20

When a true mixture of IMP and trehalose was analyzed, the resulting spectrum, Figure

2D, matched the predicted spectrum in Figure 5-5C. Thereby, indicating that in solution, trehalose and IMP do not interact to form preformed ions. This result implies that water may have a role in the enhancement effect, or the bombardment process of the trehalose/water amorphous crystal structure contributes to the enhancement. The following experiment was performed to determine the role of water and the bombardment-induced liberation of protons.

5.4.2.3. Bombardment-Induced Liberation of Protons and the Role of Water

When IMP is doped into a spin-cast trehalose film, there is a ~4,800 X enhancement in the [M+H]+, supporting the assumption that the bombardment process induces the liberation of protons from water molecules within the film. To isolate the influence of water, IMP-doped trehalose films with water and D2O as a solvent,

-14 + 2 respectively, were prepared and analyzed under a total ion dose of 1.6 x 10 C60 /cm .

As seen in Figure 5-6, the [M+H]+ signal for IMP is depressed in the film primarily

+ composed of D2O, while the [M+H] in the film with H2O as the solvent is

102

A. B.

C. D.

Figure 5-5: The chemical interactions of IMP and trehalose investigated by H-NMR. (A) and (B) are the control spectra obtained from pure IMP and anhydrous trehalose, respectively. Spectra from (A) and (B) were overlayed with software to obtain spectrum (C), representative of a predictive spectrum which would be obtained if IMP and trehalose did not interact to form ions. The spectrum in (C) is from the sample contained a mixture of IMP and trehalose. It is nearly identical to (C), indicating no proton rearrangements between the two species.

103

104 enhanced. This indicates that the protons of water provide a proton source for the

[M+H]+ ionization of the analyte. The differences between the two films are outside the standard deviation. Thereby, supporting the belief that water molecules act as a proton

+ source in C60 cluster/surface interactions, thus, contributing to the enhanced ionization of IMP.

6.4.3. Conclusions

Spin-cast films of trehalose provide an environment for a substantial enhancement of IMP ionization during the cluster ion bombardment event. With supporting H-NMR information, trehalose was found to not alter or interact in the proton transfer with IMP.

These results imply the cluster ion/surface interactions are necessary for ionization. In addition, when isolating water as a variable for enhancement during bombardment, the amount of water within a film was found to contribute to the ionization of a biomolecule.

105 5.5. New Trehalose Vitrification Technique for Three-Dimensional Single Cell

Studies

5.5.1. Experimental

5.5.1.1. Instrumental Parameters

Secondary ion mass spectrometry analyses were performed on the J105 – 3D

21 + Chemical Imager, an instrument previously described. A 0.5 pA 40 keV C60 primary ion beam run at a 100% duty cycle was focused to a 750 nm beam diameter. Each image was acquired with an ion fluence of 4.0 x 1012 ions/cm2 and the total dose for the depth

14 + 2 profile was 4 x 10 C60 /cm . The Mouse leukaemic monocyte macrophage (RAW) cell

SIMS images were acquired at 80 x 80 μm2 area at 128 x 128 pixels2. The Bovine Aortic

Endothelial cell (BAOEC) SIMS images were acquired at 100 x 100 μm2 area at 256 x

256 pixels2. All SIMS analyses were obtained with a +10 V stage bias at room temperature.

5.5.1.2. Culture Maintenance of RAW-264 and Bovine Aortic Endothelial Cells

RAW 246 cells, a mouse leukemic macrophage cell line, were grown in a T-75 flask using Dulbecco’s Modification of Eagle’s Medium (DMEM) (Cellgro). Bovine

Aortic Endothelial Cells (BAOEC) were grown with Bovine EC Growth Medium (Cell

106 Applications, Inc.). Both cell lines were supplemented with 10% Fetal Bovine Serum

(Cellgro), and a penicillin-streptomycin solution (10,000 I.U. penicillin, 10,000 μg/mL streptomycin) (Cellgro). Each cell line was maintained in separate incubators both at 5%

CO2 and 37 °C, until 100% confluence.

5.5.1.3. Vitrification of RAW-264 and Bovine Aortic Endothelial Cells in Trehalose

The RAW and BOAC cells were treated with 4 mL of trypsin solution (Sigma-

Aldrich), respectively, to release the cells from the flasks. The trypsin/cell solution was centrifuged at 3,000 rpm for 5 minutes and the supernatant was discarded. The cellular pellet was washed with 0.15 M ammonium formate, centrifuged for 5 minutes at 3,000 rpm, and the supernatant discarded; this washing process was repeated three times. The cellular pellet was resuspended in 0.2 M aqueous trehalose solution and incubated at room temperature for 1 hour. The resulting cell solution was used for spin-casting.

The cell/trehalose solution was spin-cast onto hydrophilic 5 x 5 mm2 Indium Tin

Oxide (ITO) - coated glass shards. The shards were made hydrophilic by exposure to ozone for 3 hours. 10 μL of the cell/trehalose solution was dropped incrementally onto the ITO-coated glass shards and spun at 3500 rpm for 60s between drops. Each spin-cast sample was made with a total of 100 μL cell/trehalose solution.

107 5.5.1.4. Data Analysis for Three-Dimensional Constructions

Using custom software, a stack of 2D images was converted into a 3D representation by creating semi-transparent, colored voxels.

5.5.2. Results and Discussion

5.5.2.1. Spin-Cast Trehalose Vitrification of Cells

Trehalose vitrification of cells by spin-casting is a sample preparation technique for SIMS analyses which offers a balance between the benefits of freeze-dried and frozen hydrated preparations while avoiding some of their pitfalls. For typical cellular analysis, the cells require several days to adhere and proliferate on a poly-lysine coated substrate.22

Whereas, with the spin-casting method, cells can be harvested from a flask required to maintain continuous cultures. The coverage of cells on the substrate can be determined by how much trehalose/cell solution is added to the spin-cast sample. This enables multiple concentrations of cells on substrates to be prepared on the same occasion. For the typical cellular analysis, some substrates inserted into the cellular colonies can produce a toxic environment to cells and they will grow slowly onto the substrate or not at all. Yet, the creation of a cell-coated spin-casting sample is independent of the toxicity of the substrate, provided it has a hydrophilic surface for even film formation.

Furthermore, the method is well-suited to cell lines which do not adhere to any surface

108 during growth. As seen in Figure 5-7, cells are not misshapen from the spin-cast procedure. They retain a spherical shape: the same shape they assume during suspension.

In Figure 5-7A, the RAW cells’ nuclei are observable while vitrified in the trehalose film.

The film maintains a glassy appearance similar to that seen in spin-cast trehalose films.14

The trehalose appears to completely encircle the cells. The centrifugal force inherent to the spin-casting procedure reduces the entrapment of air and water pockets in the film, as observed in Figure 5-7B, and results in a thinner glass coating, less prone to incurring charging.23,6

Trehalose vitrification by spin-casting and other techniques have been previously investigated as a method of preservation alternative to cryogenics.7,24 As a result, vitrified spin-cast RAW cells samples can maintain their state for several months when kept in a low-humidity environment. 25,24,7 This allows for multiple days of SIMS analysis and even analysis between instruments with the same sample, allowing for a variety of experiments. Whereas, with a frozen-hydrated sample, the cells rupture while approaching room temperatures in vacuum.

5.5.2.2. Intracellular Detection of Purines

The overall goals of the purine quantitation and enhancement studies were in preparation and planning for the capabilities of a single cell experiment, those goals are realized ultimately in this study. In Figure 5-8, several purines and other biomolecules are observable in the trehalose vitrified RAW cells. Nucleotides, Adenine and Guanine,

109

A. B.

110 are observed in Fig. 5-8 A and B, respectively. The majority of the adenine signal is localized to the nucleus, where the majority of genetic material is stored. Yet, adenine and guanine are also observed in lesser concentrations inside the cells, this observation correlates to the fact that nucleotides are synthesized and free-floating in the cytoplasm until they are used for construction of genetic material or salvaged. Both of these distributions of nucleotides are well-resolved in this subcellular chemical image.

Aminoacids, the building blocks of proteins, are detected in their expected distributions within the cell in Figures 5-8 B, C, and E. The phosphocholine head group fragment is indicative of the cell membrane. The “holes” in their otherwise round distribution coincide with the dense areas of the adenine signal, therefore, the “holes” show the area of the nucleus. Furthmore, the adenine and PC-headgroup distribution show that the nucleus, nuclear envelope, and cell membrane have maintained their inherent distribution. Therefore, the other chemical distributions are indicative of a preserved cell.

The purine of interest in the end-product of the de novo purine biosynthetic pathway, inosine monophosphate, was observed in the spectrum as a [M+H]+ and adjacent isotope peak, however, its distribution is low intracellularly. The reason could lie in that the cells were well-maintained at the time of vitrification and would not need to activate this “starvation” pathway, thus, the IMP concentration would not be amplified.11

Two components of the sample preparation can be observed in chemical images: trehalose and indium. The trehalose distribution in Figure 5-8 G shows the trehalose surrounding the cells and the area around them. The areas of higher intensity imply that the amount of trehalose around the cells was higher than the surrounding areas, which indicates that the trehalose covered the sides of the cells, as well.

111

Figure 5-8: Purines and other biochemical in vitrified RAW cells. The chemical images shown are the result of all the layers of the depth profile summed together in two dimensions (x-y). Chemical images (A) through (F) are purines. (A) is guanine, [M+H]+, at m/z 152; (B) is phenylalanine, [M+H]+, at m/z 120; (C) is thymine/isoleucine, [M+H]+, at m/z 84; (D) is adenine, [M+H]+, at m/z 136; (E) is arginine, [M+H]+, at m/z 175; and (F) is inosine monophosphate, [M+H]+, at m/z 349. (G) is trehalose, [M+H]+, at m/z + + 343. (H) is the phosphocholine head group, C5H15NPO4 at m/z 184. (I) is Indium, [M] at m/z 115.

112 5.5.2.3. Three-Dimensional Chemical Imaging of Cells

Three-dimension chemical imaging creates the most informative depiction of a cell by SIMS. Whereas in Figure 5-8 it was obvious that the cells had a circular shape, in the three-dimensional image in Figure 5-9, morphology is introduced into the data and the shapes of the cells are spherical. This observation further confirms that the spin-cast trehalose vitrification method preserves the inherent morphology and ultrastructure of the cells. In cryopreservation research, spin-casting cells in trehalose has shown that 90% of cellular membranes retain their integrity in the process.6 Trehalose is clearly surrounding the cell. This is indicated by the green areas on the image; this color is created by the yellow trehalose signal and the blue PC-headgroup signal being adjacent. Where in

Figure 5-8, it was assumed that the brighter trehalose regions around the cell were due to the height of trehalose surrounding the cell to be higher, the coating and valleys of the trehalose glass in between cells is apparent by the morphological information. The top

SIMS images, corresponding to approximately 20 nm, are removed from the 3-D depiction to clearly observe the intracellular contents, most specifically, the nucleus in fuschia.

5.5.2.4. Evidence of Trehalose Entry into the Cytoplasm and Nucleus of the Cell

Research concerning trehalose vitrification as a means of cellular preservation was first created as a method for long-term storage of cells and cell-based products as

113

Figure 5-9: Three-dimension chemical image of RAW cells vitrified in trehalose. Yellow is trehalose, fuchsia is adenine, blue is the phosphocholine headgroup. The field of view is 80 x 80 μm2.

114 clinical therapies involving cells and cell-based research is on the rise. While cryopreservation is the leading long-term storage solution, the difficulty transporting cryogenically frozen cells and the high cost and space requirement poses serious limitations for clinical research trials and medical treatment centers. Trehalose vitrification has shown success as an alternative method of cellular preservation and

However, viability of cells after trehalose dessication is still low at 10 – 50% viability after reanimation.26,6,27 This raises the question as to whether or not trehalose is able to enter the cell so it is able to preserve the aqueous chemistry via the hydrogen-bonding network it promotes with its multitude of OH groups.28,29,30

The intracellular trehalose distribution was evident in the BAOC cells, as shown in Figure 5-10. In Figure 5-10 A, the trehalose signal is plotted in a series of chemical images. The intracellular signal inside the cell is significantly more intense than the static shown outside the cellular areas. This result indicates that the trehalose enters the cytoplasm and the nucleus of the cell. As seen in Figure 5-10 B, the trehalose distribution corresponds to the intracellular area occupied by adenine. The trehalose signal is thicker outside of the cell membrane and forms a ring. When trehalose is plotted with the PC-headgroup signal in Figure 5-10 C, this trehalose ring formation surrounds the cell membrane.

To ensure that residual trehalose fragments from the top coating were not left behind with depth, the [M+H] + of trehalose was plotted. The relative intracellular intensity remains constant throughout the depth profile of the cell. This result indicates

+ that during this room temperature depth profile, the C60 ion beam is able to provide

115

116 secondary ions while also removing damage incurred from the bombardment, resulting in depth resolution appropriate for cellular studies.

5.5.3. Conclusions

Spin-cast trehalose vitrification of cells enables SIMS analyses of cell in nearly natively-preserved state. The sample preparation allows for a more tunable cellular analysis: different cell coverage, substrates, and instruments may all be used in the same day or across several weeks. Several purines and other biomolecules may be chemically mapped in three-dimensions from one complete imaging depth profile. Furthermore, the analyses have shown that trehalose does enter the cytoplasm and the nucleus in spin-cast trehalose vitrified cells.

5.6. References

1 Parry, S. A. et al. Imaging macrophages in trehalose with SIMS. Applied Surface

Science 255, 929-933, doi:10.1016/j.apsusc.2008.05.251 (2008).

2 Parry, S. & Winograd, N. High-resolution TOF-SIMS imaging of eukaryotic cells

preserved in a trehalose matrix. Analytical Chemistry 77, 7950-7957,

doi:10.1021/ac051263k (2005).

117 3 Lu, C., Wucher, A. & Winograd, N. Ionization effects in molecular depth

profiling of trehalose films using buckminsterfullerene (C-60) cluster ions.

Surface and Interface Analysis 43, 99-102, doi:10.1002/sia.3449 (2011).

4 Lerach, J. O. & Winograd, N. Evidence for the formation of dynamically created

pre-formed ions at the interface of isotopically enriched thin films. Surface and

Interface Analysis 45, 54-56, doi:10.1002/sia.5102 (2013).

5 Lu, C., Wucher, A. & Winograd, N. Investigations of molecular depth profiling

with dual beam sputtering. Surface and Interface Analysis 45, 175-177,

doi:10.1002/sia.4838 (2013).

6 Chakraborty, N. et al. A spin-drying technique for lyopreservation of mammalian

cells. Annals of biomedical engineering 39, 1582-1591 (2011).

7 Lee, Y. A. et al. Cryopreservation of porcine spermatogonial stem cells by slow-

freezing testis tissue in trehalose. Journal of Animal Science 92, 984-995,

doi:10.2527/jas2013-6843 (2014).

8 An, S. G., Kumar, R., Sheets, E. D. & Benkovic, S. J. Reversible

compartmentalization of de novo purine biosynthetic complexes in living cells.

Science 320, 103-106, doi:10.1126/science.1152241 (2008).

9 Zhang, T., Zhang, J., Derreumaux, P. & Mu, Y. G. Molecular Mechanism of the

Inhibition of EGCG on the Alzheimer A beta(1-42) Dimer. Journal of Physical

Chemistry B 117, 3993-4002, doi:10.1021/jp312573y (2013).

10 Smith, G. K., Mueller, W. T., Wasserman, G. F., Taylor, W. D. & Benkovic, S. J.

Characterization of the enzyme complex involving the folate-requiring enzymes

of de novo purine biosynthesis. Biochemistry 19, 4313-4321 (1980).

118 11 An, S., Kumar, R., Sheets, E. D. & Benkovic, S. J. Reversible

compartmentalization of de novo purine biosynthetic complexes in living cells.

Science 320, 103-106 (2008).

12 Braun, R. M. et al. Performance characteristics of a chemical imaging time-of-

flight mass spectrometer. Rapid Communications in Mass Spectrometry 12, 1246-

+, doi:10.1002/(sici)1097-0231(19980930)12:18<1246::aid-rcm316>3.0.co;2-c

(1998).

13 Weibel, D. et al. A C60 primary ion beam system for time of flight secondary ion

mass spectrometry: its development and secondary ion yield characteristics.

Analytical Chemistry 75, 1754-1764 (2003).

14 Cheng, J. & Winograd, N. Depth profiling of peptide films with TOF-SIMS and a

C-60 probe. Analytical Chemistry 77, 3651-3659, doi:10.1021/ac048131w (2005).

15 Fletcher, J. S., Rabbani, S., Henderson, A., Lockyer, N. P. & Vickerman, J. C.

Three-dimensional mass spectral imaging of HeLa-M cells - sample preparation,

data interpretation and visualisation. Rapid Communications in Mass

Spectrometry 25, 925-932, doi:10.1002/rcm.4944 (2011).

16 Altelaar, A. M., Luxembourg, S. L., McDonnell, L. A., Piersma, S. R. & Heeren,

R. M. Imaging mass spectrometry at cellular length scales. Nature protocols 2,

1185-1196 (2007).

17 Fletcher, J. S. Cellular imaging with secondary ion mass spectrometry. Analyst

134, 2204-2215 (2009).

18 Fletcher, J. S. et al. A new dynamic in mass spectral imaging of single biological

cells. Analytical Chemistry 80, 9058-9064 (2008).

119 19 Jackson, L. M., Hue, J. J. & Winograd, N. Quantitative detection of purines in

biologically relevant films with TOF-Secondary Ion Mass Spectrometry. Surface

and Interface Analysis 45, 237-239, doi:10.1002/sia.5098 (2013).

20 Wucher, A., Cheng, J. & Winograd, N. Molecular Depth Profiling Using a C-60

Cluster Beam: The Role of Impact Energy. Journal of Physical Chemistry C 112,

16550-16555, doi:10.1021/jp8049763 (2008).

21 Hill, R., Blenkinsopp, P., Thompson, S., Vickerman, J. & Fletcher, J. S. A new

time‐of‐flight SIMS instrument for 3D imaging and analysis. Surface and

Interface Analysis 43, 506-509 (2011).

22 Orive, G., Tam, S. K., Pedraz, J. L. & Hallé, J.-P. Biocompatibility of alginate–

poly-l-lysine microcapsules for cell therapy. Biomaterials 27, 3691-3700 (2006).

23 Lechene, C. P., Lee, G. Y., Poczatek, J. C., Toner, M. & Biggers, J. D. 3D multi-

isotope imaging mass spectrometry reveals penetration of 18O-trehalose in mouse

sperm nucleus. Plos One 7, e42267 (2012).

24 Qu, B. et al. Trehalose Maintains Vitality of Mouse Epididymal Epithelial Cells

and Mediates Gene Transfer. Plos One 9,

doi:e9248310.1371/journal.pone.0092483 (2014).

25 Cheng, J., Wucher, A. & Winograd, N. Molecular depth profiling with cluster ion

beams. Journal of Physical Chemistry B 110, 8329-8336, doi:10.1021/jp0573341

(2006).

26 Chakraborty, N. et al. Cryopreservation of spin-dried mammalian cells. Plos One

6, e24916 (2011).

120 27 Chesné, C. et al. Viability and function in primary culture of adult hepatocytes

from various animal species and human beings after cryopreservation. Hepatology

18, 406-414 (1993).

28 Crowe, J. H., Crowe, L. M. & Chapman, D. Preservation of membranes in

anhydrobiotic organisms: the role of trehalose. Science 223, 701-703 (1984).

29 Leslie, S. B., Israeli, E., Lighthart, B., Crowe, J. H. & Crowe, L. M. Trehalose

and sucrose protect both membranes and proteins in intact bacteria during drying.

Applied and Environmental Microbiology 61, 3592-3597 (1995).

30 Wolkers, W. F., Walker, N. J., Tablin, F. & Crowe, J. H. Human platelets loaded

with trehalose survive freeze-drying. Cryobiology 42, 79-87 (2001).

121 Chapter 6

Conclusions and Future Directions

6.1. Ongoing and Future Research

6.1.1. The Potential Symbiotic Role of the Flavonoids of B. braunii

Using an Argon gas cluster source, flavonoids were determined to be present in the UTEX #2441 Race A strain of B. braunii. The molecules are produced by plants for a range of functions, including protection from excessive UV light, antimicrobial protection, and pollinator attractants.1-6 Currently the most interesting function is that they have been linked to enabling their organisms to have symbiotic relationships, most specifically, a nitrogen-fixing symbiosis. This type of action could indicate an interesting evolutionary advancement.1 If B. braunii is capable of forming such a symbiotic relationship, this characteristic could potentially be genetically expressed in other organisms or strains of B. braunii. Genetic screening for bacterial sequences that code for nitrogen-fixation are currently underway by collaboration with Justin Yoo and Wayne

Curtis. In addition, work to identify the bacteria that cohabitate with colony-forming B. braunii race A UTEX #2441 are currently ensuing. The continuation of this research spurred by this finding of flavonoids with SIMS has the potential to yield important results regarding symbiotic relationships in B. braunii and the plant kingdom as a whole.

122 6.1.2. Wax Monoester Expression and the Maintenance of an Axenic Line of B. braunii Race B

An axenic line of B. braunii is necessary for genetic engineering. If bacteria or other organisms are present within the culture or colony, selective genetic engineering of

B. braunii cannot be accomplished. It is difficult to obtain and maintain an axenic line of

B. braunii, but more so race B than race A. For the purposes of biofuel research, race B is preferable because its isoprene-derived hydrocarbons are excellent substrates for hydrocracking; yet, it also grows slowly and is more difficult to maintain. Using SIMS, race A was determined to have wax monoesters covering the outside of the colony, whereas, race B did not possess these biomolecules. This class of lipids is known to provide protection from desiccation and bacterial infections.7-10 Therefore, the major reason why race A is more readily axenic than race B could be the presence of wax monoesters with inherent antibacterial protective qualities. The genetic sequence that allows race A to naturally produce this antimicrobial coating could be genetically expressed in race B. Thereby, an inherent resistance in addition to traditional antimicrobial measures could be taken to create a more hardy and axenic strain of race B.

6.1.3. Physiological Engineering of the Oil Bodies of B. braunii

With three-dimensional SIMS analysis of B. braunii race B, a more informed mechanism for hydrocarbon synthesis, storage, and secretion has been elucidated. The current transgenic research to express the enzymes responsible for botryococcene methylation in other organisms has fallen short in that their rates of hydrocarbons

123 production is degrees of magnitude less than B. braunii achieves with the same enzymes.11-13 In this thesis work, the formation of metabolon associated with the hydrophobic biochemistry of the oil body has been determined to be an important contributor for the high synthesis rates of B. braunii. Therefore, to have transgenic mutants exhibit the high rates of production, genetic engineering is not sufficient. This opens the door to a new approach called ‘physiological engineering,’ where the physiological feature, the oil body, and its associated enzymes must be expressed into transgenic mutants. Then the high rate of synthesis can be achieved. The biological information revealed here with SIMS has spurred this new approach of biofuels and hydrophobic biochemical production using physiological engineering.

6.1.4. Time-Course Chemical Imaging of Mammalian Cells

The spin-cast trehalose vitrification method enables a more easily obtained time- course experiment with mammalian cells. Because this method takes less than 1 hour from cell flask to a preserved cellular sample, time points can be created in increments from every few hours to every few days. Once the time-course is complete, one control and 2 or 3 time-point samples could be run consecutively in 24-48 hours of instrument time. This would allow for maximum comparability between sample by reducing variation in factors like ion beam current and diameter, pressure fluctuations, and typical issues of instrumental drift. This type of experimental approach would allow SIMS chemical imaging to directly compete with analyses like time-course fluorescent microscopy. However, SIMS would excel over fluorescence techniques. With SIMS, a

124 large range of biomolecules could be detected simultaneously without the use of tags with sub-micron spatial resolution. With the use of spin-cast trehalose vitrification, the sample preparation that currently prevents these types of analyses can be overcome.

6.1.5. Guiding Principles for the Development of Biological Analyses with SIMS

Here, specific principles were taken into consideration when designing sample preparation and analysis techniques for biological samples. (1) When possible, methods were integrated to enhance the secondary ion yield. (2) Sample preparation or instrumental parameters known to decrease the secondary ion yield of biomolecules were avoided, especially to decrease the effects of charging. (3) The number of steps in the technique was reduced or minimal, thereby, reducing the accumulation of error and promoting reproducibility of the process. (4) Techniques were designed to be quickly learned and mastered, another feature that also contributes to the overall reproducibility of the process. Biological analyses tend to involve the detection of subtle chemical or morphological changes differences and there is a certain degree of variation is inherent to these samples because they are from living organisms. When approaching a biological analysis, these principles exemplified in this thesis work should be taken into consideration.

125 6.2. Conclusory Remarks

SIMS has come a long way in the past few years in terms of instrument advancement and innovation. New primary ion beams allow for the investigation of features of a few hundred nanometers or extend the mass range by several thousands of

Daltons.14-16,17-19 Instruments more well-suited to examining a large mass range and resolving peaks of small differences in mass have been established.20-23 Improved sample preparation techniques have enabled biological cells and tissues to be analyzed with

SIMS instrumentation.24-29 Currently, the field is poised to enter the biological arena and engage in problem solving by offering a unique, spatial chemical imaging technology.

There is a wealth of biological questions that could be solved with SIMS.

However, there is a disconnect between what biological questions SIMS researchers are aware of and what biological researchers are aware SIMS can do. In this thesis work, a concerted effort was made to determine where these fields could meet. The progression of this work exemplifies what is possible by engaging in the biological discussion.

6.3. References

1 Koes, R. E., Quattrocchio, F. & Mol, J. N. The flavonoid biosynthetic pathway in

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2 Cook, N. & Samman, S. Flavonoids—chemistry, metabolism, cardioprotective

effects, and dietary sources. The Journal of nutritional biochemistry 7, 66-76

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126 3 Petreska, J. et al. Phenolic Compounds of Mountain Tea from the Balkans:

LC/DAD/ESI/MSn Profile and Content. Natural Product Communications 6, 21-

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4 Wojakowska, A. et al. Structural analysis and profiling of phenolic secondary

metabolites of Mexican lupine species using LC-MS techniques. Phytochemistry

92, 71-86, doi:10.1016/j.phytochem.2013.04.006 (2013).

5 Calani, L. et al. Ultra-HPLC MSn (Poly)phenolic Profiling and Chemometric

Analysis of Juices from Ancient Punica granatum L. Cultivars: A Nontargeted

Approach. Journal of Agricultural and Food Chemistry 61, 5600-5609,

doi:10.1021/jf400387c (2013).

6 Wojakowska, A., Perkowski, J., Goral, T. & Stobiecki, M. Structural

characterization of flavonoid glycosides from leaves of wheat (Triticum aestivum

L.) using LC/MS/MS profiling of the target compounds. Journal of Mass

Spectrometry 48, 329-339, doi:10.1002/jms.3160 (2013).

7 Vioque, J. & Kolattukudy, P. E. Resolution and purification of an aldehyde-

generating and an alcohol-generating fatty acyl-CoA reductase from pea leaves

(Pisum sativum L). Archives of Biochemistry and Biophysics 340, 64-72,

doi:10.1006/abbi.1997.9932 (1997).

8 Negri, G., Marcucci, M. C., Salatino, A. & Salatino, M. L. F. Hydrocarbons and

monoesters of propolis waxes from Brazil. Apidologie 29, 305-314,

doi:10.1051/apido:19980401 (1998).

127 9 Perera, M. et al. Biological origins of normal-chain hydrocarbons: a pathway

model based on cuticular wax analyses of maize silks. Plant Journal 64, 618-632,

doi:10.1111/j.1365-313X.2010.04355.x (2010).

10 Walker, M. et al. Forensically Robust Detection of the Presence of Morpholine in

Apples-Proof of Principle. Food Analytical Methods 5, 874-880,

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11 Niehaus, T. D. et al. Identification of unique mechanisms for triterpene

biosynthesis in Botryococcus braunii. Proceedings of the National Academy of

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from Botryococcus braunii Race B. Journal of Biological Chemistry 287, 8163-

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14 Kraft, M. L. et al. Quantitative analysis of supported membrane composition

using the NanoSIMS. Applied Surface Science 252, 6950-6956,

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15 Kraft, M. L. et al. Chemical imaging of lipid distribution within cell membranes

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128 16 Kraft, M. L. Identifying the mechanisms for non-random sphingolipid

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17 Fujii, M. et al. Study on the detection limits of a new argon gas cluster ion beam

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18 Moritani, K. et al. Secondary ion emission from insulin film bombarded with

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20 Hill, R., Blenkinsopp, P., Thompson, S., Vickerman, J. & Fletcher, J. S. A new

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Interface Analysis 43, 506-509 (2011).

21 Fletcher, J. S. et al. A new dynamic in mass spectral imaging of single biological

cells. Analytical Chemistry 80, 9058-9064 (2008).

22 Smith, D. F. et al. High mass accuracy and high mass resolving power FT-ICR

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1 VITA

Lauren Marie Jackson

Lauren Marie Jackson was born in northern New Jersey to Marta Caridad Abascal

Jackson, a talented stage actor and television newscaster native to Santa Clara, Cuba, and

Bruce Marshall Jackson, an internationally recognized designer of men’s footwear.

Lauren was raised with her brother, Ian Joseph Jackson, in Washington Township, NJ.

Her elementary and middle school education were completed at St. Mary’s School,

Hackettstown, NJ, and Saints’ Phillip and James School, Phillipsburg, NJ. In 2004, she graduated from Immaculata High School with honors earned from academic performance, involvement, and volunteerism. She entered college determined to perform research and has held multiple research fellow appointments at Marist College, The

Smithsonian Environmental Research Center, and Rensselaer Polytechnic Institute throughout her undergraduate career. Her eagerness for research earned her the Dr.

Andrew A. Molloy Memorial Scholarship, American Chemical Society Scholar and

Ventures Scholar appointments, the Boehringer-Ingelheim Award for Overall Scientific

Research, along with other departmental research awards and grants. After graduation from her Bachelor of Science degree in Chemistry and Biochemistry cum laude, she joined the laboratory of Professor Nicholas Winograd immediately upon arrival at The

Pennsylvania State University to pursue a Ph.D. in Chemistry. Her research has been focused on developing creative ways to use SIMS to answer current biological questions, while obtaining a thorough experience in instrumentation, mass spectrometry, and biological systems.