In Cycas Thouarsii Has Been Identified As a Mixture of Regioisomeric Formamides

DEVELOPMENT OF AN ANALYTICAL METHOD FOR β-METHYLAMINO-L-ALANINE, A CYANOBACTERIAL METABOLITE AND POTENTIAL ENVIRONMENTAL TOXIN

AND

SELECTIVE EXTRACTION PROTOCOL AND STRUCTURE OF FORMAMIDES OF β-METHYLAMINO-L-ALANINE (BMAA) FROM CYCAS THOUARSII

A THESIS SUBMITTED TO THE GRADUATE DIVISION OF THE UNIVERSITY OF HAWAI'I AT M NOA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF

MASTER OF SCIENCE

CHEMISTRY

MAY 2014

by Yoshiaki Miyasaka

Thesis Committee:

Thomas K. Hemscheidt, Chairperson Phillip Williams Joseph T. Jarrett

We certify that we have read this thesis and that, in our opinion, it is satisfactory in scope and quality as a thesis for the degree of Master of Science in Chemistry

THESIS COMMITTEE

______

iii

Acknowledgement

I would like to thank the members of my thesis committee for their time, helpful comments and suggestions. First, I would like to show my greatest appreciation to my advisor,

Professor Hemscheidt, for his continuous and generous support during my years in a graduate program. Additionally, I would like to express my gratitude to Professor Williams and

Professor Jarrett for their support when needed.

I have been also supported by all faculty members and graduate students at the chemistry department. I would like to show my deep gratitude to them wholeheartedly. In addition, I would like to thank my collaborators in the Bidigare laboratory. (Professor

Bidigare, Stephanie Christensen and Daniel Elsey). My work could not have been accomplished without their continuous support.

I would also like to thank the University of Hawaii for financial support in the form of teaching assistantship. Being a TA was great learning experience along with my research experience.

Finally, I would like to express my gratitude to my family and friends for their moral support and warm encouragements.

Abstract

Part I:

The development of a qualitative/quantitative analysis of β-Methylamino-L-alanine

(BMAA), including a sample preparation protocol and SPE protocol is described. A common interfering metabolite was isolated as the FMOC-derivative. Its structure was determined by spectroscopic methods and confirmed by chemical synthesis.

Part II:

The qualitative analysis of the BMAA content in samples from cycad leaf and oyster are described. Three different extraction protocols have been developed to categorize the form of BMAA present in either cycad leaf or oyster muscle. The low molecular mass conjugate of β-Methylamino-L-alanine (BMAA) in Cycas thouarsii has been identified as a mixture of regioisomeric formamides. The structures were elucidated on the basis of spectroscopic data and confirmed by chemical synthesis.

Table of Contents

Acknowledgements...... iii

Abstract...... iv

List of Tables...... viii

List of Figures...... ix

List of Schemes...... xi

List of Abbreviations...... xii

Part I: Development of an Analytical Method for β-Methylamino-L-alanine, a

Cyanobacterial Metabolite and Potential Environmental Toxin

1.1 Introduction...... 2

1.2 Approaches to BMAA analysis, pros/cons...... 5

1.2.1 Direct method...... 5

1.2.2 Indirect method...... 6

1.3 Detections for BMAA...... 8

1.3.1 UV/FD detection...... 8

1.3.2 MS/MS detection...... 9

1.4 False positive detection of BMAA due to co-elution of similar compounds...... 11

1.5 The isolation and structure elucidation of the contaminant peak...... 12

1.6 Development of a DNFB based pre-column derivatizing method...... 16

1.6.1 MRM method development...... 16

1.6.2 Isomers of BMAA: Possible interferences...... 18

1.7 Qualitative Analysis of BMAA...... 20

1.8 Quantitative Analysis...... 24 vi

1.8.1 Internal standard...... 24

1.8.2 Calibration Standards...... 28

1.8.3 Calibration Curves...... 28

1.8.4 Detection limit/Sensitivity...... 32

1.9 Matrix effect...... 34

1.10 SPE cartridge cleaning/concentration, Dowex-50 strong cation resin...... 36

1.11 Sample and LC/MS sample preparation protocol...... 39

1.11.1 Large Scale Extraction...... 40

1.11 Summary...... 42

1.12 Experimental section...... 43

1.13 Reference...... 52

vii

Part II: Selective Extraction Protocol and Structure of Formamides of β-Methylamino-

L-alanine (BMAA) from Cycas thouarsii

2.1 Introduction...... 56

2.2 Free BMAA vs. Protein associated BMAA in Cycad leaf...... 58

2.3 Isolation of the formamide of BMAA from Cycas thouarsii...... 63

2.4 Elucidation of the formamide of BMAA...... 65

2.5 mono-DNB deivatives of the BMAA formamides...... 71

2.6 The absolute configuration...... 75

2.7 Several notes...... 77

2.8 Conclusion...... 78

2.9 Experimental section...... 79

2.10 References...... 85

viii

List of Tables

Table Page

1.1 List of isobaric molecules of BMAA and observed fragmentations in

order of decreasing ion intensity...... 18

1.2 Summary of calibration standards...... 28

1.3 Summary of three calibration curves, A, B and C...... 29

1.4 Summary of curve A with separate ranges...... 30

1.5 Summary of Matrix-adapted curves...... 35

1.6 Recovery from SPE protocol...... 38

1.7 Recovery after entire sample preparation...... 40

2.1 500 MHz (1H) and 125 MHz (13C) NMR data for the two observable

rotamers of 1a and for 1b in CD3OD...... 70

List of Figures

Table Page

1.1 Structure of BMAA...... 3

1.2 Section of the UPLC/FD chromatogram of hydrolyzed Spirulina.

(UPLC/FD Method A)...... 9

1.3 Mass spectrum of AQC-tagged co-eluting peak with fragmentation at 10

eV...... 12

1.4 Structure of the compound co-eluting with BMAA...... 15

1.5 Derivatizing reaction of BMAA and DNFB...... 16

1.6 Fragmentation of bis-DNB-BMAA at 10 eV...... 17

1.7 Chromatogram of standard isomer mix (Top) and hydrolyzed Spirulina

(Bottom) by LCMS/MS Method A...... 21

1.8 LC Chromatogram of isomer mix (Top) and hydrolyzed Spirulina

(Bottom) by LCMS/MS Method B...... 23

1.9 Fragmentation of bis-DNB- BMAA-d3...... 27

1.10 451>210 extracted ion chromatogram of BMAA (top) and BMAA-d3

(bottom) ...... 27

1.11 Calibration curve by the Agilent quantitative program...... 32

2.1 Selective extraction protocol #1...... 59

2.2 Selective Extraction Protocol #2...... 61

2.3 Selective Extraction Protocol #3...... 62

2.4 Summary of selected nOe ( ), 15N HMBC ( ), HMBC ( ) and

HSQC ( ) data for the region-isomeric formamides 1a and 1b...... 70 x

2.5 HPLC/UV(340 nm) chromatogram of mono-DNB-BMAA conjugate

derivatives...... 72

2.6 Summary of selected nOe ( ), for the regio-isomeric formamides 2a

and 2b...... 73

2.7 Fragmentations of α-formyl-β-DNB derivative (Top) and β-formyl-α-

DNB derivative of BMAA (Bottom) at 10 eV collision energy...... 74

2.8 HPLC/UV chromatogram of Marfey-tagged DL-BMAA (Top), L-

BMAA (Middle), and partially racemized natural BMAA conjugate

(Bottom)...... 76

2.9 HPLC Chromatogram of aqueous extract from C. thouarsii.(Top) and

extracted mass spectrum (bottom)...... 77

List of Scheme

Table Page

1.1 Isolation scheme of the FMOC derivative of the unknown...... 14

1.2 Synthetic scheme of BMAA-d3...... 25

1.3 Summary of sample preparation...... 39

1.4 Overall flow of the method...... 41

2.1 Isolation diagram of Compound 1...... 64

2.2 Synthetic scheme of α-formyl and β-formyl BMAA...... 66

xii

List of Abbreviations

T [α] D Specific optical rotation at 589 nm and temperature T in °C

AccQ-Tag AccQ-Tag Ultra Derivatization Kit

ACN acetonitrile

AEG N-(2-aminoethyl)glycine

ALS amyotrophic lateral sclerosis

ALS-PDC Amyotrophic Lateral Sclerosis-Parkinson/Dementia complex amu atomic mass unit aq. aqueous

AQC 6-aminoquinolyl-N-hydroxysuccinimidyl carbamate br broad atm atmospheric pressure

BAMA β-amino-methylalanine

BMAA -methylamino-L-alanine

BOC tert-butyloxycarbonyl group c concentration in g/100 mL calcd calculated

Cbz carbobenzoxy group

C degrees Celsius

13C carbon-13 isotope d doublet

D deuterium, 2H d3 trideuterated xiii

Da Dalton

DAB 2,4-diaminobutyric acid

DAPA 2,6-diaminopimelic acid dd doublet of doublets

DI deionized

DNB 2,6-Dinitrobenzene group

DNFB 2,6-Dinitrofluorobenzene

1D-nOe one dimensional nuclear Overhauser effect spectroscopy dq doublet of quartets

ELSD evaporative light scattering detector em emission

ESI electrospray ionization ex excitation eV electron volt

FD fluorescence detector

FLT fluorescence lifetime

FMOC 9-fluorenylmethyloxycarbonyl group

GC gas chromatography

1H proton isotope

HILIC hydrophobic interaction chromatography

HMBC heteronuclear multiple-bond correlation spectroscopy

HPLC high performance liquid chromatography

HRMS high-resolution mass spectrometry xiv

HSQC heteronuclear single-quantum correlation spectroscopy

ISTD internal standard

J coupling constant (in Hz)

L liter

LC liquid chromatography

LiAlD4 lithium aluminum deuteride

LOD limit of detection

LOQ limit of quantification m multiplet

M molarity

[M ]+ molecular ion

MRM multiple reaction monitoring

MS mass spectrometry m/z mass to charge ratio

15N nitrogen isotope

N normality

NMR nuclear magnetic resonance

O.N. overnight ppm parts per million psi pound per square inch q quartet

R2 coefficient of determination

RT room temperature xv s singlet

S standard sat. saturated

SCX strong cation exchange

SIM single ion monitoring

S/N signal-to-noise ratio sp. species

SPE solid phase extraction t triplet

TLC thin layer chromatography

TOF time of flight

TUV tunable ultraviolet

UPLC ultra performance liquid chromatography

UV ultraviolet

V volt

 chemical shift (in ppm)

 wavelength

Part I:

Development of an Analytical Method for -Methylamino-L-Alanine,

a Cyanobacterial Metabolite and Potential Environmental Toxin

1.1 Introduction

It has been said that our diet strongly affects our health. In the 1950’s a peculiar neurodegenerative illness was reported on the island of Guam. A significant number of native people showed symptoms such as paralysis, shaking and dementia at 50-100 times the incidence of amyotrophic lateral sclerosis (ALS)1, 2 compared to other populations. This illness was known as lytico-bodig among native people in Guam and later received the clinical name Amyotrophic Lateral Sclerosis-Parkinson/Dementia complex (ALS-PDC).

A significant body of research suggested that consumption of the traditional diet of native people in Guam was the only variable significantly associated with disease incidence.3

Specifically, seeds of cycad plants (Cycas Micronesia or Cycas circinalis) were consumed by the native people in Guam as a source of tortilla flour. From the seeds of the cycad a non- proteinogenic basic amino acid, -methylamino-L-alanine (BMAA) (Figure 1.1) was isolated and suspected to be a neurotoxin causing such disease.1, 2 In subsequent research in the

1980’s the neurotoxicity of BMAA in vitro and vivo was demonstrated in mice and nonhuman primates.3, 4, 5 However, these reports were criticized because the results were obtained by feeding a high dose of BMAA to the animals and these far exceeded the doses that humans could possibly be exposed to. As a result, interest in BMAA as a causative agent of ALS-PDC waned.

In recent years, the ethnobotanist Paul Cox has revived this hypothesis by invoking biomagnification of BMAA6 from a primary producer through a food chain ending with humans as the penultimate consumer. Using HPLC-FD techniques, Cox’s group also tested brains of ALS-PDC patients and of Canadians who had died of Alzheimer disease for BMAA, which was detected in all brains tested. A careful analysis of the data available in the form of figures in this publication reveals that other amino acids one would expect to show up in 3 these samples were not detected. In view of the reports by others who could not confirm the presence of BMAA in brain tissue, it appears possible that what was analyzed as BMAA really was some other compound. This is not entirely unlikely as HPLC-FD is a problematic technique for analysis, particularly if the fluorescence originates from a tag and is not intrinsic to the analyte of interest, unless the tag is highly specific for the analyte. A less specific tag may react with a variety of compounds and the resulting adduct may elute with the same retention time as that of the analyte of interest resulting in a false-positive result.

O H C 3 N OH H NH2 Figure 1.1 Structure of BMAA

The food chain postulated by Cox envisions the primary producer to be symbiotic cyanobacteria of the genus Nostoc, known to be present in the root nodules of cycads. The

BMAA produced by these organisms was reported to be translocated to the plant, to be accumulated in the fruit of the cycads, which in turn would be eaten by "flying foxes", bat- like animals considered to be delicacies among the Chamorro. Cox's analytical work suggested a 10,000-fold enrichment of BMAA along this food chain (cyanobacteria to flying foxes).6 In further support of this hypothesis, Cox reported to have found BMAA in representative genera of all five groups of cyanobacteria, in amounts up to 6 mg/g dry weight.

This raised the concern that, rather than just a narrowly defined ethnic group consuming a traditional diet, a much larger fraction of the human population might be exposed to BMAA through, say, drinking water contaminated by BMAA. Precedents for such exposure to cyanobacterial toxins by way of contaminated drinking water exists, e.g. 4 microcystins, anatoxin-a and cylindrospermopsin.7 In addition, as interest in BMAA has grown, this amino acid has been identified in various samples of marine animals such as oysters, mussels, and scallops, suggesting that humans may be exposed to BMAA through consumption of BMAA-containing food.8

Due to the public health implications of potential widespread BMAA, many studies on BMAA detection and quantification have been reported with quite contradictory conclusions, of which studies using UV- or fluorescence- based detection of BMAA are the most controversial. Inherently more specific methods can detect BMAA, if at all, only at much lower concentrations than those reported using HPLC-FD methodology.

As more high-tech detection technology, such as LC/MSMS, is applied to the problem of BMAA analysis, a consensus is emerging about BMAA levels in natural samples and the extent of its distribution. However, even with advanced methodology in hand, such as

LCMS/MS, a false positive result may still be observed if the analytical method is not carefully developed. Of greatest concern is the presence in natural samples of BMAA isomers that may generate isobaric ions. Initially we set out to modify existing methodology based on

AccQ-Tag® technology. In the course of this work we identified a common bacterial metabolite whose presence leads to false positive results when the commercial AccQ-Tag®

AQC-based methodology is used. The isolation and identification of the compound co-eluting and causing a false positive result will be described.

In working with the AccQ-Tag® chemistry we found the method had a few significant weaknesses. We therefore developed a new qualitative/quantitative method for BMAA analysis using 2,4-dinitrofluorobenzene (DNFB) as a pre-column derivatizing reagent. The method is based on LC/MSMS using a MRM scan of the bis-DNB derivative of BMAA. The stepwise method development including matrix effects, a solid phase extraction (SPE) 5 protocol, and sample preparation will be described.

1.2 Approaches to BMAA analysis, pros/cons

In order to be able to explain some of the decisions we made initially or during the course of the research, some general remarks on direct and indirect methods will be presented.

Both approaches have pros/cons when a method for a specific analyte is developed.

Appreciating these will help us to explore the most suitable method for BMAA analysis.

1.2.1 Direct method

Direct analytical methods involve the direct injection of a sample containing the target without any pre-column derivatization. Ease of sample preparation is the major advantage of this approach. Owing to the absence of a significant chromophor in BMAA, a direct method for its analysis would have to use an ELSD or similar general detector in an LC approach or the use of a mass spectrometer as a detector. However, owing to the low molecular mass of

BMAA (BMAA has a mass of 118 amu), any mass spectrometric method would require analysis in the low mass region of the spectrum. This is not preferred for MS/MS analysis since analysis in the low mass region suffers from high background noise and ion suppression from the sample matrix. This could cause false positive and false negative results, respectively, in the detection of BMAA. Since BMAA is not only small, but also highly polar, it is not readily amenable to reversed phase chromatography. Instead any direct method of analysis involving HPLC likely will require the use of hydrophobic interaction chromatography (HILIC) with its associated problems of longer column equilibration times between runs, the need for a more complex buffer solvent, and poor separation of amino acids.9 Therefore, it seemed to us that the cons overwhelm the pros for the direct injection method of BMAA analysis. 6

1.2.2 Indirect method

Some of those problems seen for the direct method discussed above can be overcome by using an indirect analytical approach. In an indirect method, the sample is subjected to chemical manipulation that modifies the target, and likely other components of the sample matrix, prior to injection into the analytical apparatus, in order to increase sensitivity or to modify chromatographic behavior. This inevitably requires more time for sample preparation in comparison with a direct method, but with proper choice of the derivatization reagent and the chromatographic conditions, a remarkable increase in sensitivity over a direct method can be achieved.

Derivatization of BMAA could not only make it chromatographically more tractable in standard reversed phase chromatography. Another advantage would be that derivatization increases the molecular weight of the target, thereby moving the molecular ion from the crowded low-mass range to one less dominated by sample matrix ions. As a result one may expect better selectivity and sensitivity.

Various reagents have been used for pre-column derivatization of amino acids prior to analysis by either GC and GC/MS or LC and LC/MS, respectively. Two of these, have also been used for BMAA analysis, namely ethyl chloroformate10 and (6-aminoquinolyl-N- hydroxysuccinimidyl carbamate (AQC)). The latter is based on aminoquinoline, which yields highly fluorescent tagged amino acids.11 Both reagents are available as commercial kits at rather significant cost. The former is often used for GC/MS analysis since in the tagging process relatively little mass is added to the analyte and the derivative is still sufficiently small to be amenable to gas chromatography. AQC is the most commonly used tag for LC analysis of amino acids. The advantage of this tag is ease of use, safety, the availability of already well-established protocols for derivatization and analysis, the commercial availability 7 in the form of convenient kits as well as support from the vendor in the form of databases. If standards are available and the amino acid to be analyzed is relatively abundant relative to other amino acids, this is a very robust method of analysis. The method is more problematic if the analyte of interest is minor in comparison to other species that also react with the tag.

The more abundant reactive species may outcompete a minor component in the derivatization reaction, resulting in the underestimation of the analyte of interest. Additionally, a substantial mass is added to the analyte with each tag and this may lead to precipitation of the derivative and other sample components before injection onto the LC column. Since injection volumes are typically small, with an analyte of low relative abundance in the matrix it may be difficult to inject a sufficiently large fraction of the sample onto the column such that a minor component is still detectable in the presence of tagged background compounds.

1.3 Detections for BMAA

The choice of detectors also plays an important role as well. Two types of detectors have often been employed in BMAA studies. Specifically, UV, FD, and ELSD type of detectors were the most typical detectors used for BMAA detection in the 90’s and early

2000’s, while in more recent work detection is accomplished by mass spectrometric methods.

1.3.1 UV/FD detection

The UV/FD detection of BMAA tagged as the ACQ derivative was the early method of choice for BMAA analysis and is still used by some analysts. Whether detection of the tagged analyte occurs by UV absorbance or by fluorescence, in these analytical methods the retention time is the only parameter for compound identification. Any co-eluting species can lead to false positive results qualitatively and lead to an overestimation of analyte abundance during quantitative analysis. Some of the early published papers on BMAA detection that have come under criticism in later work used this methodology.12 For example, there were strong indications from the Bidigare laboratory at UH, that use of the standard gradient recommended by the manufacturer of the AccQ-Tag® kit, when applied in the analysis of

BMAA, results in co-elution, or near-co-elution, of a compound that is ubiquitous in cyanobacteria. As shown in Figure 1.2, this compound results in a shoulder peak almost at the same retention time as that of BMAA, the typical retention time difference being 0.2 min or less. If one does not spike the sample to be analyzed with a BMAA reference standard, the chromatogram in Figure 1.2 could easily be interpreted as BMAA positive. Therefore, many of the results from LC analysis of BMAA by UV/FD might need to be re-examined.

Figure 1.2 Section of the UPLC/FD chromatogram of hydrolyzed Spirulina. (UPLC/FD

Method A)

1.3.2 MS/MS detection

The problem of co-eluting tagged contaminants can be overcome by using LCMS/MS where the mass spectrometer selects out the parent/precursor ion of the analyte of interest and thereby suppresses other tagged moieties of different molecular masses. Upon fragmentation through collisional activation of the selected parent ion, daughter/product ions are created that can be used for quantification. Therefore MS-based detection in the absence of isobaric ions enables us to achieve highly selective analysis.

This approach cannot, however, protect the analysis from interference by isomers of the analyte, which might co-elute with the target molecule and generate isobaric ions upon ionization. Isobaric ions possess the same elemental composition as the parent ion of the target analyte. Occurrence of such isobaric ions may lead to false positive results unless 10

MS/MS techniques are used and the fragmentation path of the contaminant is different from the one for the target analyte. Typically, the AQC tagged compounds yield a major product ion of m/z = 171 as the most abundant product ion.12 This fragment ion only incorporates atoms from the tag and none originating from the analyte. Thus, the major ion, which should be important for quantification due to the associated high signal-to-noise ratio, is not characteristic of the analyte of interest and hence is non-specific. However, this will inevitably decrease the selectivity due to the fact that the most abundant daughter ion from

AQC-tagged amino acids comes from the tag itself. As a consequence, some published papers have attempted to modify the standard AQC method to avoid the use of the m/z = 171 peak.13

Therefore, we wished to develop an alternative indirect method that would eliminate interference from isobaric ions, was inexpensive and had at least comparable sensitivity to that of the AQC method.

1.4 False positive detection of BMAA due to co-elution of similar compounds

As discussed earlier, there are some contradictions between past published papers on

BMAA detection. Of most immediate interest to us was the report by Cox that 95% of the genera of cyanobacteria produce BMAA.10 Therefore, when we started our method development we selected Spirulina-based nutritional supplements as starting material. This material is cheap and available in bulk without the long lead times required for cyanobacterial cell mass derived from in-house fermentations. At the time we were using AQC as derivatizing reagent and LC/UVFD for detection. In our analysis of Spirulina cell mass samples obtained after acid hydrolysis, we observed a peak that eluted at the retention time of a BMAA standard or close to it. Careful spiking of the Spirulina extract with BMAA led to broadening or doubling of the peak, depending on the chromatographic run. This suggested that while this unknown peak behaved chromatographically and chemically much like

BMAA, it was not BMAA. Hence, this material was cause for a false positive result as shown in Figure 1.2 unless a careful BMAA spike was used. In order to investigate the metabolite giving rise to the AQC derivative co-eluting with BMAA, the compound was isolated from

Spirulina as a 9-fluorenylmethyloxycarbonyl (FMOC) derivative.14 Further details will be discussed in a later section.

Likewise, Nostoc sp. CMMED001, a cyanobacterial isolate originating from the

Hawaiian islands, was reported by Cox in 2005 as containing 1,243 μg/g free BMAA content.8 However, our recent screening using our LCMS/MS method showed no trace of

BMAA content in CMMED001. Thus, false positive results due to co-elution of species similar to BMAA might have contributed to the contradictory results among BMAA analyses in past published papers, especially those based on UV/FD detection

1.5 The isolation and structure elucidation of the contaminant peak

The molecular mass of the peak co-eluting with the bis-AQC derivative of BMAA in the extract of Spirulina was first determined based on MS fragment analysis (Figure 1.3).

Figure 1.3 Mass spectrum of AQC-tagged co-eluting peak with fragmentation at 10 eV

The presence of two amino sites in this unknown was deduced based on MS analysis.

In two consecutive fragmentations AQC tags are lost from the parent ion (m/z = 531>361) and then from the resulting daughter ion (m/z = 361>171) (Figure 1.3). This analysis suggested that the compound of interest likely was a diamino acid having a mass of 190 Da.

We chose to isolate this compound from Spirulina in order to characterize it chemically. In a series of scouting experiments it was observed that the compound giving rise to the m/z = 531 peak after derivatization with AQC, bound to Dowex-50 resin and could be eluted with aqueous ammonia, much like typical monobasic amino acids. We then chose to derivatize the components of the amino acid fraction eluted from the Dowex resin with an

FMOC group. This was attractive as the resulting derivatives are well-behaved in standard 13 reversed phase chromatography16 and have a significant UV chromophor so that the purity of any component is readily assessed. The latter part is the reason why another standard carbamate derivative commonly used in amino acid chemistry, such as BOC or Cbz, was not chosen. A last consideration for the choice of an FMOC tag was that the addition of two

FMOC groups to the analyte, as suggested by the observation of a bis-AQC derivative by MS, results in a significant mass increase for a compound that was a minor component of the mixture. This increased the likelihood that we would be observing sufficient increases in mass recovery over an approach trying to isolate the underivatized unknown. This would allow us to obtain an amount of the pure, derivatized unknown that could be handled easily and would allow us to determine the structure.

Using the purification scheme shown in Scheme 1.1, the minor “BMAA-like” compound from Spirulina was isolated by use of a combination of reverse phase (C18), size exclusion (LH-20) and normal phase chromatography (silica gel). The tagged unknown was tracked in the fractions by using LCMS targeting a m/z = 634 ion (= 2 × FMOC + 190 – 2H).

The purification scheme is shown in Scheme 1.1. 14

Powdered Spirulina

6 N HCl @ 100C, O.N.

Hydrolysate

Dowex-50 cation exchange resin

Crude amino acid mixture

FMOC Derivatization

Tagged amino acid mixture

100 cm LH-20 Column

0 - 140 mL fractions 140 - 160 mL fraction

HPLC C18 Column

0 - 19.9 min fraction 19.9 - 20.45 min fraction

Si Column, 3 times

Si fraction

LH-20 Column

Isolated FMOC-DAPA

Scheme 1.1 Isolation scheme of the FMOC derivative of the unknown

The co-eluting peak was identified as 2,6-meso-diaminopimelic acid (2,6-meso-

DAPA) on the basis of 1H NMR, 13C NMR and mass spectral analysis. This result of analysis was confirmed by comparison of the spectroscopic data of the bis-FMOC derivative of the unknown compound to those of a sample of chemically synthesized FMOC-DAPA prepared 15 from a commercial mixture 2,6-meso-DAPA and D/L-DAPA.17 The comparison of 1H NMR,

13C NMR and mass spectra is shown in the Appendix.

O O

HO OH

NH2 NH2

2,6-meso-Diaminopimelic acid Figure 1.4 Structure of the compound co-eluting with BMAA

2,6-meso-DAPA is contained in a bacterial cell wall16, 17 and is essential for bacteria to grow normally. Since 2,6-meso-DAPA is ubiquitous in cyanobacteria, its presence may lead to false positive results during BMAA analysis using the AQC method unless precautions are taken that result in an improved separation of bis-AQC DAPA from bis-AQC BMAA during

LC analysis. This is critical in an LC method with UV- or fluorescence-based detection.

Fortunately, 2,6-DAPA is not one of the isobaric isomers of BMAA so that it would not be problematic as long as an MS/MS based method of detection is used. However, the observation of co-elution of bis-AQC DAPA and bis-AQC BMAA suggests that when using a

MS/MS-based detection system, close attention needs to be paid to co-elution of compounds generating isobaric ions. Therefore, testing possible isomeric molecules seemed essential for the development of a robust LCMS/MS method.

Even employing ultra-high resolution techniques such as UPLC/FD analysis for the determination of BMAA using the AQC tag, we observed a potential co-elution problem that might result in false positive qualitative analyses or quantitative overestimation. Therefore, we explored an alternative derivatizing reagent. 16

1.6 Develoment of a DNFB based pre-column derivatizing method

2,6-Dinitrofluorobenzene (DNFB), known as Sanger’s reagent, is commonly used for amino acid analysis.18 The major drawback of this compound is its toxicity. The primary and/or secondary amino group of amino acids is tagged by the nucleophilic aromatic substitution reaction as shown in figure 1.5. The reaction time is about 15 min in the heat block at 60 °C. This is almost the same reaction time as required for the AQC method protocol. DNFB is sold for about $90 for 10 mL, and in our protocol more than 30,000 assays are possible with this volume of reagent. This compares favorably with the AccQ-Tag® kit which allows 200 analyses for $450. In the final version of our method, the DNB method showed sensitivity comparable to that of the AQC method and, significantly, even better selectivity was achieved. Therefore, the DNB tag seems an excellent alternative to AQC and we developed a qualitative/quantitative method for the analysis of BMAA with this pre- column derivatizing reagent.

NO2 DNB

NH2 DNB NH H + N N COOH COOH NO2 Tagged BMAA F [M+H]+=451.1 Figure 1.5 Derivatizing reaction of BMAA and DNFB

1.6.1 MRM method development

An MRM method for bis-DNB BMAA was established by monitoring a product ion resulting from fragmentation of the molecular ion of the doubly tagged bis-DNB-BMAA

([M+H]+ = 451.1) at 10 eV collision energy as shown in figure 1.6. Three transitions were monitored as follows: The most abundant 451>210 transition was chosen for the quantifier 17 ion and the 451>254 and 451>268 transitions for the qualifier ions. Significantly, unlike the bis-AQC derivative of BMAA, the bis-DNB derivative of BMAA did not generate any daughter ion that contained only atoms from the tag itself. Thus, increased selectivity will be observed since the fragment ions contain structural elements of the analyte and therefore reflect its structure.

x104 + Product Ion (13.147 min) (451.0 -> **) BMAA Prod Ion Scan 10.d 4.75 210.1 4 4.5 25 4.25 4 NO2 3.75 H C 3.5 3 3.25 O2N NH N NO2 3 2.75 2.5 HOOC O2N 2.25 2 6 2 8 1.75 210 451.1 1.5 254.1 1.25 1 268.1 0.75 198.0 0.5 134.1 236.1 0.25 0 80 90 100 110 120 130 140 150 160 170 180 190 200 210 220 230 240 250 260 270 280 290 300 310 320 330 340 350 360 370 380 390 400 410 420 430 440 450 460 470 480 490 500 Counts vs. Mass-to-Charge (m/z)

Figure 1.6 Fragmentation of bis-DNB-BMAA at 10 eV

The peak area ratio of the most abundant m/z = 210 (Quantifier ion) to the m/z = 254 and m/z = 268 (Qualifier ions) was also calculated. The 254/210 ratio was 1:5 and 268/210 ratio was 1:10, averaged over 10 calibration standards. The consistency of the peak area ratio should be maintained within ±10% to achieve a high selectivity of the method. This condition was fulfilled as well. On the basis of these results the fundamental method of BMAA qualification/quantification had been established. The next problem to be addressed is that of potential isomers of BMAA acting as confounders.

1.6.2 Isomers of BMAA: Possible interferences

The same product ion scan analysis was also conducted for various possible isomers of BMAA, which had been proposed by Jiang et al.13 These authors found that there are 260 theoretical structural isomers of BMAA on the basis of a database search (Scifinder, PubMed).

The list was narrowed down by imposing certain restrictions such as 1) presence of two amino groups, 2) tolerance to hydrolysis conditions and 3) no net charge at neutral pH. This set of restrictions reduced the number of potential isomers to be considered during BMAA analysis from 260 to seven candidates.13 Of these, three isomers (AEG, BAMA and DAB) were picked as being of interest as potentially interfering with BMAA analysis on the basis of possible natural occurrence. The list of BMAA isomers that were tested in our method together with their respective major fragment ions is shown in Table1.1. The actual mass spectra of product ion scans for all BMAA isomers tested are shown in the Appendix A.1-3.

Table 1.1 List of isobaric molecules of BMAA and observed fragmentations in order of decreasing ion intensity

BMAA AEG BAMA DAB O O O H H3C H BND N N OH N OH N OH BND OH BND DNB N H HN N HN DNB DNB O BND DNB 451.1>210.1 451.1>210.1 451.1>254.1 451.1>196.0 451.1>254.1 451.1>268.0 451.1>268.2 451.1>268.1 451.1>268.1 451.1>164.0 451.1>134.1 451.1>240.1

One of the isomers, β-amino-methylalanine (BAMA), was synthesized according to the method by Jiang et al.13 BAMA was identified in an oyster sample in our screening program, therefore this compound is an actual interference. DAB has been found in many prokaryotic and eukaryotic organisms, and it has been investigated widely.19 AEG was 19 recently identified in cyanobacterial samples as a possible interference by Banack.20 AEG was also detected in Spirulina by our method as the possible cause of false positive results during MS/MS detection. In the standard HPLC protocol, the its retention time is very similar to that of BMAA as discussed in more detail later. For all three potential contaminants, analysis of the product ion scans revealed that, except for AEG, the most abundant product ions are different from the most abundant fragment ions of bis-DNB BMAA (Appendix A.1-

3). It should therefore be possible to distinguish the DNB derivatives of BAMA and of DAB from BMAA on the basis of the m/z ratio of the most abundant fragment ion of their respective bis-DNB derivatives. Bis-DNB-2-aminoethylglycine, on the other hand, has a peak at m/z = 210 as the most abundant product ion, just as BMAA does. Of greatest concern for quantitative analytical work therefore was the DNB derivative of 2-aminoethylglycine, which, if it could not be separated from BMAA, would contribute strongly to the 451>210 transition and could be mistaken for a contribution from BMAA. We therefore expected that if we could achieve a baseline separation of the various isomers of BMAA by optimization of LC conditions, a false positive result due to the presence of any of the tested BMAA isomers could be avoided. Therefore, base-line separation of these isomers from our target BMAA was an essential goal to be achieved.

1.7 Qualitative Analysis of BMAA

The initial LC conditions used a 10% ACN - 100% ACN with 0.1% (v/v) formic acid to both components of the mobile phase (0.1% Formic acid adduct to ACN and Water). The linear gradient was performed in 20 min with a flow rate of 0.7 mL/min (LCMS/MS Method

A). The retention time of BMAA, AEG and DAB, respectively, is shown in figure 1.7, where

BMAA and AEG almost co-elute. It thus appeared that the two compounds that needed to be clearly separated chromatographically were eluting closest to each other. It is straightforward to qualitatively distinguish between bis-DNB AEG and bis-DNB BMAA since the 451>254 transition is missing in the AEG derivative. However for any quantitative analysis the

451>210 transition is crucial as discussed earlier. Therefore a baseline separation of bis-DNB

AEG from bis-DNB BMAA is necessary. This is all the more important since it is known that

AEG occurs in cyanobacteria and it is therefore possible that a sample contains both AEG and BMAA. In that case even the qualification becomes invalid in that a contribution to the

451>210 transition from bis-DNB AEG will skew the 210 vs. 254/268 peak area ratio which is used to qualify a fragment ion as originating from bis-DNB BMAA. As is shown in Fig.

1.7 (bottom), the AEG known to be present in cyanobacteria can be a major confounder for

BMAA if chromatographic separation cannot be achieved. Therefore, the bis-DNB AEG/bis-

DNB BMAA co-elution issue has to be resolved.

Figure 1.7 Chromatogram of standard isomer mix (Top) and hydrolyzed Spirulina (Bottom) by LCMS/MS Method A (linear gradient 10% - 100% ACN w/ 0.1% formic acid in 20 min)

After some experimentation optimized conditions were discovered that achieved the desired separation without changing eluting solvents (Solvent A: 0.1% Formic acid in water;

Solvent B: 0.1% Formic acid in acetonitrile). The only change required was a shallower linear gradient from 40% ACN - 55% ACN in 15 min. After the elution of the bis-BMAA derivative, the column needs to be washed for a better reproducibility. The total run time including column washing and re-equilibration is 25 min. Under the new conditions BMAA is separated from all its isomers that are of concern as shown in figure 1.8. In addition, not only all BMAA isomers but also lysine and 2.6-DAPA are well separated from BMAA. The 22 method enables us to do highly selective qualitative analysis of the samples along with

BMAA quantification. The chiral isomer, LL/DD-DAPA, elutes close to the BMAA peak.

However, this is not of great concern as LL/DD-DAPA has not been identified in Nature and therefore this is not a significant problem. Moreover, since DAPA has a different molecular mass, it will not influence qualification/quantification result of BMAA using the MRM method. The details of the method are described in the Experimental section.

The newly explored LCMS/MS Method B also enabled us to identify the presence of

AEG in Spirulina (Figure 1.8) and in Leptolyngbya, which confirms in our own hands that cyanobacterial samples frequently contain AEG. The concern that AEG could be a major interference in cyanobacterial sample is therefore warranted and the time and effort on optimizing the separation was well spent.

Figure 1.8 LC Chromatogram of isomer mix (Top) and hydrolyzed Spirulina (Bottom) by

LCMS/MS Method B (linear gradient 40% - 55% ACN w/ 0.1% formic acid in 25 min)

1.8 Quantitative Analysis

With a selective LC method in hand, we moved on to develop a quantitative method for the analysis of BMAA. This is usually done by constructing a calibration curve and quantification typically involves the use of a standard to account for losses during sample preparation and fluctuations in instrument sensitivity. This standard may either be analyzed as a separate sample (external standard) or the standard may be admixed with the sample

(internal standard). Our method employs an internal standard.

1.8.1 Internal standard

An internal standard is often used by analytical chemists in order to account for losses during a sample preparation protocol, or instrumental response variations and injection volume variations.21 It involves the construction of a calibration curve using the peak area ratio of the analyte vs. the internal standard. The internal standard should be chemically close to the target molecule so that ideally it behaves as much as possible the same as the target molecule, while still being distinguishable from the analyte.

For this reason isotopically labeled derivatives of the analyte of interest are often employed as an internal standard. We therefore decided to prepare BMAA-d3 with a deuterated methyl group as our standard. With this number of deuterium atoms in the molecule isotope fractionation was not likely going to be a problem and a mass difference of

3 amu would suffice to distinguish analyte from standard. While BMAA-d3 could be easily

22 synthesized by using methylamine-d3 according to the method of Ziffer et al, due to unavailability of methylamine-d3 in Hawaii, BMAA-d3 was synthesized through a different synthetic scheme as follows. (Scheme 1.2)

O O + NH2 EtO N EtO Cl H

O D D LiAlD4 EtO N D N H THF, reflux, 3 hours H

O D D O HN D N + N H N COOMe 60 oC, 48 hours H MeOOC D3C

O O

Pearlman's catalyst HN HN

N H2 (g), RT, 24 hours NH MeOOC MeOOC D3C D3C

O 3 M HCl H2N 2 HCl HN NH 110 oC, 4 hours HOOC NH D3C MeOOC D3C

HOOC NH2 H2N 1) Dowex-50 Clean-up 2 HCl HCl NH HOOC 2) 1 equiv. HCl NH D3C CD3

BMAA-d3 monohydrochloride

Scheme 1.2 Synthetic scheme of BMAA-d3

The labeled methyl portion was prepared by LiAlD4 reduction of a carbamate and the resulting amine combined with acetamidoacrylic acid methyl ester to give protected BMAA- d3. The N-benzyl group was removed by catalytic hydrogenolysis, while the acetamide and ester protecting groups were removed by acidic hydrolysis, respectively, to give unprotected

BMAA-d3 in the dihydrochloride form, which was converted to the BMAA-d3 mono- hydrochloride since our BMAA standard was purchased as a mono-hydrochloride.

BMAA-d3 was synthesized successfully in the mono-hydrochloride form according to scheme 1.2. The sample of deuterated BMAA was subjected to a product ion scan of its bis-

DNB derivative. The labeled compound behaved exactly as the unlabeled reference compound with exception of its mass. The product ion scan of BMAA-d3 as a doubly tagged

DNB derivative is shown in figure 1.9. The parent ion showed up as m/z = 454 and the most abundant product ion appeared as m/z = 213, which is exactly +3 amu from the 451>210 transition of the natural abundance material. The 454>213 transition can therefore be used as our internal standard transition. Lastly, we made sure that the synthesized BMAA-d3 showed no trace contamination by unlabeled BMAA. The MS/MS analysis after tagging was shown in figure1.10. Therefore, the BMAA-d3 internal standard will not contribute to the ion intensity of the analyte.

x10 4 + Product Ion (13.1 min) (454.00 -> **) D3BMAA Trial#1 prod10.d 3 213.10 54 2.8 2 2.6 NO2 2.4 2.2 D3C 2 O2N NH N NO2 1.8 1.6 1.4 HOOC O N 2 2 1.2 7 1 454.20 1 213 254.10 0.8

0.6 271.10 0.4 134.10 168.10 201.00 236.00 0.2 0 80 100 120 140 160 180 200 220 240 260 280 300 320 340 360 380 400 420 440 460 480 500 Counts vs. Mass-to-Charge (m/z)

Figure 1.9 Fragmentation of bis-DNB- BMAA-d3

The comparison of BMAA and BMAA-d3 fragmentation patterns suggests some structural information on the most abundant fragment. The shift of m/z = 210 to m/z =

213(+3) in BMAA-d3 is evidence that the N-methyl group and one DNB tag are present in the most abundant fragment ion.

Figure 1.10 451>210 extracted ion chromatogram of BMAA (top) and BMAA-d3 (bottom)

1.8.2 Calibration standards

A total of 10 calibration standards were prepared by spiking a fixed amount of

BMAA-d3 (50 ng). The peak area ratio of BMAA and BMAA-d3 was plotted against the concentration of BMAA to construct the calibration curve. The summary of all calibration standards is shown in Table 1.2.. From total volume of 0.5 mL, 10 L of each standard of bis- derivatives was subjected to the LCMS/MS analysis.

Table 1.2 Summary of calibration standards

Standard BMAA ISTD Mass (ng) Concentration Mass (ng) Concentration (ng/mL) (ng/mL) S1 2.5 5 50 100 S2 5.0 10 50 100 S3 12.5 25 50 100 S4 25 50 50 100 S5 50 100 50 100 S6 125 250 50 100 S7 250 500 50 100 S8 500 1000 50 100 S9 1250 2500 50 100 S10 2500 5000 50 100

1.8.3 Calibration Curves

Three calibration curves were constructed at one to three month intervals to investigate the effect of change over time, (3 month - 1 month - 1 month). All calibration standards were freshly prepared before the analysis. The linear regression analysis was performed for three calibration curves with ISTD to evaluate each curve. Calibration curve C was constructed in the first month. Then curve B and A were constructed three month and one month later, respectively. The linear regression was performed using Excel data analysis. The slope of all three curves was 0.013383 on average with a standard deviation of 0.00125. The 29 coefficient of variation was calculated as 9.3%. This result suggests that the slopes agree within 10% error even among the curves that were constructed more than a month from each other. This proves that the ISTD approach works well to account for daily/monthly variation of instrumental response. Moreover, all curves showed extremely good R square values, indicating the precision within the single curve is well maintained for all three calibration curves as well. The summary for the analysis is shown in Table 1.3.

Table 1.3 Summary of three calibration curves, A, B and C

80 STD/ISTD area ratio y = 0.0145x - 0.508 R² = 0.9994 CurveA

70 y = 0.0137x - 0.5247 R² = 0.9988 CurveB

y = 0.012x - 0.2427 60 R² = 0.9998 CurveC

50 STD Curve A

STD Curve B 40

STD Curve C

30 Linear (STD Curve A)

20 Linear (STD Curve B)

Linear (STD Curve C) 10

0 0 1000 2000 3000 4000 5000 6000 Concentration -10 (ng/mL)

Curve A Curve B Curve C R Square 0.999 0.999 1.000 Standard Error for y 0.800 0.800 0.278 # of standards 10 10 10 Equation 0.014x - 0.508 0.014x - 0.525 0.012x - 0.243

In order to obtain even more accurate BMAA quantification, curve A was divided into three regions encompassing the low, middle and high range, respectively. Standard curve A was chosen as the calibration curve for further analysis since it is the most recently constructed curve. For the low range, standards S1 - S4 (5 - 50 ng/mL) were used, standards

S4 – S7 (50 - 500 ng/mL) for the middle range, and standards S7 - S10 (500 - 5000 ng/mL) for the high range. The summary and curves are shown below in Table 1.4. 30

Table 1.4 Summary of curve A with separate ranges

Low Middle High R Square 1.000 1.000 1.000 Standard Error 0.003 0.042 0.287 # of standards 4 4 4 Equation y = 0.011x - 0.002 y = 0.012x - 0.014 y = 0.015x - 1.711 Curve A Low range 2 y = 0.0113x - 0.0021 R2 = 0.9999 1.5

0.5

STD/ISTD area ratio area STD/ISTD 0 10 20 30 40 50 60 -0.5

-1 Concentration (ng/mL)

Curve A Middle range 8 y = 0.0118x - 0.0139 7 R2 = 0.9998 6 5 4 3

2 STD/ISTD area ratio area STD/ISTD 1 0 -1 0 100 200 300 400 500 600 Concentration (ng/mL)

Curve A High range 80 y = 0.0148x - 1.7109 2 70 R = 0.9999 60 50 40 30

20 STD/ISTD area ratio area STD/ISTD 10 0 0 1000 2000 3000 4000 5000 6000 Concentration (ng/mL)

Even better R square values are observed when the three divided ranges are considered separately. It might be beneficial to use each range separately to obtain more accurate quantification in the analysis of samples from natural sources. Since to date we have not encountered a high BMAA content in marine samples or cyanobacterial samples, the low or mid-range standard is the most appropriate for these, whereas BMAA-rich samples such as

Cycad seed or leaf may fall within the range of the High Range standard.

The quantification program (Agilent MassHunter Workstation QQQ Quantitative analysis) also gave the best R square value using best fitted localisms function with an excellent R square value satisfying almost all calibration standards. Standard 9 was ignored

(shown as a light dot) for the result with the best fit (Figure 1.11).

Figure 1.11 Calibration curve by the Agilent quantitative program (S9 is shown as a light dot)

1.8.4 Detection limit/Sensitivity

The instrumental and methodological limit of detection (LOD) and the limit of quantification (LOQ) were determined experimentally using a dilution series of a standard solution. The commonly accepted value for LOD is S/N = 3, and S/N = 10 for LOQ, therefore these values were used for our definition of LOD/LOQ. The lowest standard used was 5 ng/mL (2.5 ng BMAA). It showed the least abundant qualifier transition 451>268 with a S/N

(Signal-to-noise) ratio = 3.8. Since 5 ng/ml corresponds to a 50 pg on-column amount of

BMAA with a 10 L injection and we have a method to concentrate BMAA (which will be discussed in a later section), a standard lower than 5 ng/mL has not been tested. Moreover, final volume of samples, injection volume, columns diameter or instrumental settings could be changed to enhance S/N, resulting in even lower values for LOD/LOQ if the need for even 33 higher sensitivity arose.

The least abundant qualifier ion 451>268 (S/N>3) was used for LOD/LOQ in our method. When the least abundant 451>268 transition shows a signal-to-noise greater than 3, the other transitions 451>254 and 451>210 show one greater than 10. For the purposes that the present method was developed for, a sensitivity of 50 pg on-column for LOD/LOQ seems satisfactory.

1.9 Matrix effect

In some analyses the effect of the sample matrix on analytical performance of a method is sometimes drastic, especially if one expects to analyze samples with complex matrices such as marine/cyanobacterial samples. Any interference might invalidate the method as was seen with the AEG co-elution issue. Matrices also decrease sensitivity due to more complicated and messy background noise or via ionization suppression. Therefore, it is important to investigate the effect of the matrix deemed to be present in the samples to be analyzed.

Three types of samples were selected to examine the possible interferences and effects from samples we expected to analyze. We chose extracts from tomato, fish (Cephalopholis argus), and Spirulina to construct matrix-adapted calibration curves. Tomato represents plant supply, fish represents marine samples and Spirulina represents cyanobacterial samples.

Spirulina was treated initially with 80% aq. ethanol and was then broken up by way of a cell disruption vessel to achieve complete cell lysis. The other two matrices were extracted with

80% aq. ethanol overnight at room temperature. The resulting extracts were filtered to remove solids and matrix-adapted calibration curves were constructed from each in order to investigate ion suppression effects and the presence of any interfering peaks. A total of 10 standards (S1 - S10) of matrix-adapted standards were prepared for each matrix. The summary of the result and curves were listed in Table 1.5.

Table 1.5 Summary of Matrix-adapted curves

STD/ISTD area ratio 80 y = 0.0145x - 0.508 R² = 0.9994 STD

70 y = 0.0138x - 0.6053 R² = 0.9982 TOM

60 y = 0.0144x - 0.4716 R² = 0.9991 SP

50 y = 0.0149x - 0.1865 R² = 0.999 FISH 40 STD Curve A

30 STD Tomato

STD Spirulina 20 STD Fish

10 Linear (STD Curve A)

Linear (STD Tomato) 0 Linear (STD Spirulina) 0 1000 2000 3000 4000 5000 6000 Linear (STD Fish) -10 Concentration (ng/mL)

Curve A Tomato STD Fish STD Spirulina STD Retention time (min) 14.04 ± 0.05 13.83 ± 0.02 13.83 ± 0.05 13.99 ± 0.03 451>254 ratio 18.83 ± 1.11 22.48 ± 0.89 22.34 ± 1.01 18.29 ± 1.41 451>268 ratio 9.87 ± 0.74 11.19 ± 0.53 11.20 ± 0.74 9.94 ± 0.42

The calibration curves showed almost no ion suppression effect and the retention time also was consistent over several runs with samples from each matrix. The slopes of the three matrix-adapted curves were compared to standard curve A to evaluate any ion suppression.

Based on student’s t test, the slopes for the three curves have no difference between the slope for curve A and the slopes for matrix-adapted curves at the 99.9% confidence level.23 It clearly shows that there is no ion suppression/ion enhancement observed from either of the three tested matrices, nor was a significant retention time shift or a change in the peak area ratio of product ions observed for either of them as shown in table 1.5.

Assuming that the matrices we tested are similar to a variety of other potential matrices, the matrix analysis has shown that our method can be applied to plant, marine and cyanobacterial samples without any significant interference. Since especially with marine samples it was difficult to find a completely BMAA negative sample for use as a model 36 matrix, only one species was tested for each type of sample, i.e. plant, marine and cyanobacterial, respectively.

1.10 SPE cartridge cleaning/concentration using Dowex-50 strong cation resin

As discussed in a previous section, we might encounter a sample containing BMAA, albeit below our detection limit. This can be overcome by adjusting instrumental settings, injection volume, or column diameter. However, these adjustments will likely not drastically increase sensitivity. In order to achieve a substantially increased sensitivity, a pre- concentration of BMAA relative to other sample components appears to have the greatest potential. As a dibasic amino acid, BMAA will be strongly retained on a cation exchange resin. This has the added advantage that the contamination by neutral amino acids would be significantly reduced since these can be eluted under milder conditions than the basic amino acids.

Solid phase extraction procedures for cleanup of BMAA-containing samples have already been explored by several research groups.11 A cleaner sample usually results in an enhanced S/N ratio due to the reduction of matrix effects (less background noise) and ion suppression. In attempts to reproduce these reports we observed that not all strong cation resins performed equally well. Especially resins that had mixed-mode characteristics appeared to be inferior. We selected Extract CleanTM SCX solid phase extraction cartridges purchased from Alltech for use in our SPE protocol.

In the low pH range the two amino sites are charged in the BMAA molecule. It therefore binds to the strong cation resin more strongly than neutral/acidic amino acids. Thus, a SPE protocol that removes these less strongly retained sample components selectively will concentrate the BMAA content in the sample, relative to other components. Furthermore, an 37

SPE protocol could result in the concentration of BMAA and consequently limits to

LOD/LOQ could be overcome, even for samples having BMAA content below LOD/LOQ of the standard method. If this is the case, the use of SPE cartridge during sample preparation becomes critical when quantification of low BMAA samples needs to be accomplished. To observe such a concentration, the volume necessary for elution of BMAA from the resin must be smaller than the sample volume and the recovery of the analyte from the SPE medium must be good. It should be noted that the question concerning recovery can be addressed by inclusion of an internal standard.

Several trials were made to optimize the nature of the eluent and the order in which it was applied to the resin. These experiments resulted in the following optimized protocol. The

SCX cartridge (500 mg resin) is equilibrated by washing with 0.1 N HCl and H2O respectively, until the eluate is almost neutral at which point the sample dissolved in 0.1 N

HCl is loaded onto the SCX cartridge. The resin bed is washed first with approximately 6 bed volumes each (6 mL) of 0.1 N HCl, 0.5 N HCl, 1.0 N HCl, and DI water, respectively.

BMAA is eluted by 6 mL of 2.0 N NH4OH. In order to evaluate the protocol, UPLC/TUV analysis was used to examine how the other amino acids behave under these conditions. For each fraction collected, amino acid analysis by UPLC/TUV using the AQC tag was performed. The chromatograms of these fractions are shown in the Appendix A.4.

While neutral and acidic amino acids are eluted in the 0.1 N HCl - H2O fractions, none of the BMAA applied to the column was eluted under these conditions. BMAA eluted, as expected, together with the basic amino acids such as lysine and arginine in the ammonia fraction. Unexpectedly, phenylalanine was also identified in the basic fraction, probably as a result of the interaction of the aromatic ring with the styrene backbone of the Dowex resin resulting in retention by a mechanism different from ion exchange. 38

Qualitatively, these results show that excellent selectivity during this SPE protocol is observed and that the background from neutral and acidic amino acids can be almost completely removed. The SPE protocol helps not only to clean the sample but also to concentrate the BMAA content effectively.

In order to establish the effect of SPE protocol on the qualitative results, the recovery for selected BMAA concentrations was investigated using our quantitative method of BMAA analysis using the DNFB reagent as described in earlier sections. Three calibration standards were selected (S2, S5 and S8) and ISTD was spiked to the basic fraction after the SPE protocol. Ten μL samples of 10, 100 and 500 ng/mL STD were passed through an SPE cartridge with one replicate each. Recoveries of between 75 and 90% were observed. Since the addition of an internal standard during quantitative analysis of a sample accounts for the loss from the SPE procedure (if the ISTD is spiked BEFORE the SPE protocol), a recovery of

75 - 90% seems reasonable enough. The summarized results were shown in Table 1.6.

Table 1.6 Recovery from SPE protocol

Recovery Trial #1 and 2 (%) Average S2 93% 89% 91% S5 76% 74% 75% S8 87% 83% 85%

1.11 Sample and LC/MS sample preparation protocol

In summary, the overall protocol is shown in Scheme 1.3.

~20~20 mg Biological mg Biological samples samples - 2 mL 80% EtOH - Stir overnight at room temp. - Internal Standard spike (50 ng)

Cell disruption Vessel 80% EtOH extract (Cyanobacterial samples only)

- Centrifuged - Evaporation (Speedvac, 35 °C) - Re-suspend in 2 mL DI water - Centrifuged

Supernatant

- Extraction with 2 mL EtOAc - Extraction with 2 mL 1-BuOH (×2 for cyanobacterial samples) - Evaporation (Speedvac, 35 °C)

Aqueous extract

- Hydrolysis (6 N HCl, 100 °C, O.N.) - Evaporation - Derivatization (Speedvac, 35 °C) - Derivatization

Hydrolyzable BMAA Free BMAA

Scheme 1.3 Summary of sample preparation 40

The overall recovery after the sample preparation shown in Scheme 1.3 using

BMAA-negative fish sample is summarized in table 1.7.

Table 1.7 Recovery after entire sample preparation

Trial #1 Trial #2 Average S2 48% 47% 48% S5 42% 42% 42% S8 53% 61% 57%

The overall recovery seems low which is probably due to multiple extraction steps.

Especially the n-BuOH wash is somewhat problematic as even water-saturated n-BuOH absorbs some of the aqueous layer (73 g/L at 25 C) containing BMAA. However, inclusion of an internal standard can correct for the losses incurred during sample preparation.

1.11.1 Large Scale Extraction

In those cases where the result of the BMAA analysis suggested that the concentration is below the LOD/LOQ, but where there were nonetheless some indications for the presence of BMAA, (e.g. tiny bump, off product ion ratio, DNFB tag inefficiency, etc) a large scale extraction could be carried out. The large scale extraction protocol involves the use of a SPE

SCX cartridge in order to concentrate BMAA in the sample. The extraction steps used are the same as in the small scale sample preparation except that the sample amount is 300 mg, the extraction solvent volume is 12 mL 80% EtOH and the washes consist of two 10 mL of

EtOAc/n-BuOH washes. The aqueous extract after the EtOAc/n-BuOH wash (hydrolyzed or non-hydrolyzed) is applied to an SPE cartridge. The 2.0 N NH4OH eluate fraction is derivatized and analysed by LCMS/MS. The scheme of the overall protocol of this variant of our method is shown below in Scheme 1.4.

Small Scale Extraction

- Sample Preparation

LCMS/MS Analysis

The result is ambiguous

Large Scale Extraction SuccessfulSuccessful QuantificationQuantificatio

- Sample Preparation

SPE Cartridge LCMS/MS Analysis

Scheme 1.4 Overall flow of the method

1.12 Summary

A qualitative/quantitative method for BMAA analysis has been established successfully by means of LC/MSMS MRM scanning. DNFB was selected as our derivatizing reagent. The detection of possible isobaric molecules is integrated into the method and false positive results owing to these contaminants are not expected. The detection and quantification limits (LOD/LOQ) were determined as 50 pg on-column amount. Three representative natural samples were selected and examined for matrix effects. Our experiments showed no evidence of undesired effects from components of the selected species. A detailed protocol for the sample preparation was also established for samples of high or low BMAA content. Application of this method to the screening of real samples has revealed the presence of BMAA in oysters, mussels, scallops and sample of the cyanobacterium Leptolyngbya. The concentrations observed are in the single to low double digit μg/g range.

1.13 Experimental section

General Instrumentation:

UPLC analysis was carried out using a Waters Acquity-UHPLCsystem with a Binary

Solvent Manager, Sample Manager and TUV/FLT Detector. LCMS/MS analysis was performed on an Agilent Technologies 1200 series LC system (a Quaternary Solvent Manager,

Sample Manager and UV Detector) with a 6410 Triple Quad LC/MS. All high-resolution mass spectral data were obtained on an Agilent 6100 TOFMS instrument interfaced with an

Agilent 1100 series LC (a Quaternary Solvent Manager, Sample Manager and Diode Array

Detector). All NMR spectral data were recorded either on a Varian Unity Inova 500

Spectrometer or a Mercury 300 Spectrometer.

Derivatization procedure (AQC and DNFB):

The derivatizing reagent (AQC or DNFB) was dissolved in dry ACN at 10 mg/mL solution. To 10 L of sample solution, 110 L of 0.2 N borate buffer (pH 9) and 30 L of the derivatizing solution were added. This was heated in a heat block at 60 C for 15 min and cooled to room temperature. The total volume was adjusted to 0.5 mL by adding 350 L of

1:1/ACN:H2O with 0.1% Formic acid.

Spirulina hydrolysate preparation for UPLC/FD analysis (1.3.1):

To 20 mg of Spirulina powder, 0.5 mL of 6 N HCl was added and heated in a heat block at 120 °C overnight for complete hydrolysis. The sample was dried, derivatized as described in the derivatization procedure with AQC and analyzed using UPLC/FD Method A.

Isolation of 2,6-meso-DAPA from Spirulina and standard 2,6-DAPA (1.5):

To 2.5 g of green powdered Spirulina in a sealed round neck flask, 70 mL of 6 N HCl was added and heated at 80 °C for over 24 hours with constant stirring. After evaporation to dryness in vacuo, the hydrolysate was re-suspended in 80 mL of deionized-water and filtered through a Buchner funnel. The residue was washed two times with deionized-water, giving a total volume of about 100 mL. The resulting filtrate was a clear deep green solution. This was again dried in vacuo and adjusted to 15 mL with DI water. About 100 mL of Dowex 50W-X8,

H+ 20 - 50 mesh (J.T. Baker Chemical Co , Phillipsburg, N.J.) was used to isolate basic amino acids in the hydrolysate. Ion-exchange resin (100 mL) was packed into a 35/20 glass column and washed with 50 mL of 2 N HCl, then washed with deionized-water until the eluate’s pH was between 5 or 6. The hydrolysate was then applied to the column and washed with deonized-water until the pH reaches almost neutral. The resin was then washed with 2 N

NH4OH solution. After pH of the eluate reached pH 10, 200 mL of NH4OH solution was collected.The NH4OH fraction was concentrated to about 20 mL (pH 7), gave a clear deep green-orange solution. This was applied to a C18 SPE Cartridge (2 g, Alltech) and washed with deionized-water for further purification/decolorization. The resulting eluate was ~ 300 mg in weight. 5 mL of deionized water was added and gave a clear yellow solution (Crude

Amino Acid Mixture).

To 3 mL of the Crude Amino Acid mixture, 30 mL of 0.2 N borate buffer (pH 9) was added. The derivatization was then performed by adding 10 mL of 100 mg/mL FMOC-Cl

(ACROS ORGANICS ,N.J., USA) in acetonitrile was added to the mixture and stirred for 15 minutes in an ice bath. The mixture was then acidified to pH 2 with a concentrated HCl and concentrated in vacuo. The mixture was then extracted three times with dichloromethane and concentrated to give a white yellowish solid. Lipophilic Sephadex LH-20 (SIGMA Chemical 45

Co., St Louis, MO) was used to prepare a 70 cm LH-20 column. An appropriate amount of

LH-20 gel was equilibrated in MeOH overnight and added to a 100 cm column until the packed height reached to 70 cm and the flow rate was adjusted to 0.625 mL per minute. To the dried derivatized sample, 5 mL of MeOH was added. About half of this MeOH solution was applied to the column and 5 mL fractions were collected by use of a fraction collector to remove all mono-derivatized amino acids. The operation was carried out at room temperature.

FMOC-DAPA appeared in fractions from 140 to 160 mL. These fractions were combined and concentrated to 5 mL. The preparative HPLC system consisted of two LC-10AS HPLC pumps (SHIMADZU) and a Spectra 100, variable wavelength detector (THERMO

SEPARATION PRODUCT). Mobile phases were H2O with 0.1% formic acid (mobile phase

-1 A) and CH3CN with 0.1% formic acid (mobile phase B); the flow rate was 3.0 ml min . The detection wavelength was 263 nm. All separations were performed on a Phenomenex Luna 5

μm C18(2), 250 × 10 mm at room temperature. Gradient conditions: 3.0 mL/min, initial =

90 % A, 20 min = 0 % A, followed by a wash with 100 % B for 10 min and re-equilibration for 10 min at 90 % A. Injection volume was 150 μL each time. FMOC-DAPA appeared at about 20.10 min and the fraction from 19.9 min – 20.45 min from multiple injections was pooled and dried yielding about 10 mg of a slightly yellow solid. The solid was chromatographed initially on 2.0 g of silica gel, then on 500 mg of silica gel and last on 200 mg of silica gel eluting with ethyl acetate:1-propanol = 1:2 with 0.1% formic acid added.

After evaporation in a speedvac a white solid was obtained which was further purified by chromatography on 2.0 g of LH-20 gel packed in a 10.0 × 300 mm column. The chromatographed product was applied to the column and 1.8 ml fractions were collected.

FMOC-DAPA appeared from 12 to 14 mL. The fraction was dried and recrystallized from

H2O:MeOH = 9:1 solvent to give approximately 1 mg of pure product (0.04% recovery). 46

The standard FMOC-2,6 DAPA was synthesized by the procedure by Wang et al.16

The HREIMS, 1H NMR, and 13CNMR were measured compared to the isolated FMOC-2,6-

DAPA.

1 Isolated FMOC-2,6-meso-DAPA: H NMR (500 MHz, CD3OD): δ 7.74 (t, J = 8.1 Hz, 4H),

7.62 (dd, J = 13.3, 6.1 Hz, 4H), 7.34 (dd, J = 15.2, 7.6 Hz, 4H), 7.27 (dt, J = 14.2, 5.5 Hz,

4H), 4.28 (m, 2H), 4.16 (m, 2H), 1.89-1.72 (m, 4H), 1.52 (m, 2H);

13 C NMR (500 MHz, CD3OD): δ 176.5, 158.6, 145.2, 142.5, 128.7, 128.1, 126.3, 120.8, 67.9,

55.6, 32.5, 23.4

+ + HREIMS m/z exact mass calcd for [C37H35N2O8 + H] 635.2388 found [M+H] 635.2383

1 Standard FMOC-2,6-DAPA: H NMR (500 MHz, CD3OD): δ 7.74 (t, J = 8.5 Hz, 4H), 7.62

(dd, J = 13.5, 7.1 Hz, 4H), 7.34 (m, 4H), 7.27 (dt, 14.2, 5.6 Hz, 4H), 4.28 (m, 2H), 4.16 (m,

2H), 1.89-1.72 (m, 4H), 1.52(m, 2H);

13 * * C NMR (500 MHz, CD3OD): δ 176.5, 158.7/158.6 , 145.3/145.2 , 142.5, 128.7, 128.1,

126.3, 120.8, 67.9, 55.6, 32.5, 23.5/23.4

+ + HREIMS m/z exact mass calcd for [C37H35N2O8 + H] 635.2388 found [M+H] 635.2393

* The commercial 2,6-DAPA standard contained both LL/DD and meso isomers

Qualitative analysis of BMAA isomers (1.6.2):

Isomer mix solution was prepared by mixing 1.0 mg/mL of each standard solution of

AEG, BMAA, BAMA, DAB, DAPA and lysine. From the standard mix solution, 10 μL was derivatized by using the DNFB derivatizing procedure and 10 μL was injected to LCMS/MS.

LCMS/MS Method A was used for product ion scanning mode.

Synthesis of BMAA-d3 (1.8.1):

To 3.997 g, (37.3 mmol, 2.7 ml) of benzylamine in 5 mL ice-cold water, 40 mmol

(4.34 g, 3.8 ml) of ethyl chloroformate and 1.64 g (41 mmol) of NaOH in 5 mL of water were added dropwise simultaneously. The mixture was stirred for 1 hour. The mixture was extracted by DCM and dried. The ethyl carbamate of benzylamine was recovered as white crystals 7.0 g (99 % yield). The carbamate, 6.0 g (33.5 mmol), in 10 ml of THF was added to

2.8 g (2 equiv.) of LiAlD4 in 20 mL THF dropwise at 0 °C and the mixture was refluxed for 4 hours. After cooling down to 0 °C, 15 mL of diethyl ether was added to the mixture. The reaction was quenched by 3 mL H2O, 3.5 ml of 15% NaOH and 4 mL of water, sequentially.

After quenching, the solution was filtered and dried. It gave 3.0 g (72% yield) of oily d3- methylbenzylamine.

1 d3-Methylbenzylamine (3a): H NMR (300 MHz, CDCl3): δ 7.30-7.26 (m, 5H), 3.73 (s, 2H),

+ 1.76 (br s, 1H) HREIMS m/z extract mass calcd for [C4H10D3N2O2 + H] 122.1049, found

[M+H]+ 122.1040

To 500 μL (0.468 g, 3.77 mmol) of benzylamine-d3, 0.250 g (1.75 mmol) of methyl 2- acetamidoacrylate in 1 mL 1,4-dioxane was added and stirred for 2 days at 60 °C. The presence of the intermediate was verified by LC MS and TLC. The intermediate was dried and directly hydrolyzed by 6 N HCl at 80 °C overnight. The hydrolysate was dried and re- suspended in 1 mL 1,4-dioxane. This solution was transferred to a flask containing 50 mg

Pearlman’s catalyst and the hydrogenation reaction was conducted under H2 gas in a balloon for overnight. The mixture was dried in a Speedvac and cleaned by chromatography over

Dowex-50 strong cation exchange resin. The product, an oily solid, was then mixed with one equivalent of HCl and the mono-hydrochloride was recrystalized from EtOH. It gave about

100 mg (22.4% recovery) of BMAA-d3 mono-hydrochloride.

1 BMAA-d3 mono-hydrochloride: H NMR (300 MHz, D2O): δ 3.81 (dd, J = 8.6, 6.0 Hz, 1H),

3.29 (dd, J = 12.8, 8.6 Hz, 1H), 3.21 (dd, J = 12.8, 6.0 Hz, 1H)

+ + HREIMS m/z extract mass calcd for [C4H10D3N2O2 + H] 122.1049, found [M+H] 122.1040

Preparation of matrix-adapted curves (1.9):

A freeze-dried and well-ground 100 mg of sample (Tomato, Fish and Spirulina) was extracted with 80% EtOH overnight. The Spirulina sample was then further treated in a cell disruption vessel (N2 gas under 2000 psi for 30 min with constant stirring at RT, then abrupt pressure release). All extracts were filtered and dried. The extracts were then suspended in water and extracted with EtOAc and n-BuOH. The aqueous phases were then filtered again and dried. The extracts were suspended in 20 mM HCl so that the concentration of the extracts was 10 mg/mL. To each calibration standard, 10 L (0.1 mg) of extract was added, equivalent to 2 mg of dried weight of the matrix samples, and the final volume was adjusted 49 to 0.5 mL. The calibration standards with matrix spike were prepared by following the calibration STD preparation mentioned earlier. All standards were analyzed by LCMS/MS

Method B overnight sequentially.

SPE cartridge (Alltech SCX) protocol (1.10):

The SPE cartridge was first washed with 500 μL of 0.1N HCl and then washed with

DI water until the pH settles down to almost neutral. Spirulina hydrolysate (200 μL ) with 30 ng of standard BMAA added was applied to the pre-equilibrated Alltech SCX SPE cartridge.

The following fractions are collected.

- 6 mL of 0.1 N HCl

- 6 mL of 0.5 N HCl

- 6 mL of 1.0 N HCl

- 6 mL of DI water

- 6 mL of 2.0 N NH4OH

Each fraction was dried in a Speedvac and the residue was derivatized using the AQC method as in the standard procedure. The derivative was injected to UPLC/TUV using

UPLC/TUV Method B to determine the distribution of all amino acids in each fraction.

UPLC and LC conditions:

TM UPLC/FD Method A was performed using and AccQ-TAG Ultra C18 Column (100

× 2.1 mm, 1.7 μm particle size, Waters, Needham, MA) at 55 °C with a binary mobile phase

(solvent A, 1:20 dilution of AccQTag Ultra Eluent A; Solvent B, AccQTag Ultra Eluent B) delivered at a flow rate of 0.7 mL/min. Separation was achieved using the linear gradient program as follows: 0.0 min, 0.1% B; 0.54 min, 0.1% B, curve 6; 5.74 min, 9.1% B, curve 7; 50

7.74 min, 21.2% B, curve 6; 8.04 min, 59.6% B, curve 6; 8.64 min, 59.6% B, curve 6; 8.73 min, 0.1% B, curve 6; 9.5 min, 0.1% B, curve 6. The FD detector was set at λex=266 and

λem=473.

TM UPLC/TUV Method B was performed using and ACCQ-TAG Ultra C18 Column

(100 × 2.1 mm, 1.7 m particle size, waters) at 45 °C with a binary mobile phase (solvent A,

1:20 dilution of AccQTag Ultra Eluent A; Solvent B, AccQTag Ultra Eluent B) delivered at a flow rate of 0.4 mL/min. Separation was achieved using the linear gradient program as follows: 0.0 min, 0.1% B; 1.08 min, 0.1% B, curve 6; 8.0 min, 5.8% B, curve 7; 10.0 min,

5.8% B, curve 6; 11.48 min, 9.1% B, curve 7; 15.48 min, 21.2% B, curve 6; 16.08 min,

59.6% B, curve 6; 17.28 min, 59.6% B, curve 6; 17.46 min, 0.1% B, curve 6; 20.0 min, 0.1%

B, curve 6. The TUV detector was set at 260 nm.

LCMS/MS Method A was carried out with a Waters Xbridge column (100 × 2.1 mm,

3.5 m particle size) at room temperature and a binary mobile phase (Solvent A: 0.1%

Formic acid in water; Solvent B: 0.1% Formic acid in acetonitrile) delivered at fixed flow rate of 0.7 mL/min. The linear gradient elution program used was as follows: 0.0 min, 10%

B; 20 min, 100% B; 25 min, 10% B. The UV detector was set at 360 nm.

LCMS/MS Method B was carried out with the Waters Xbridge column (100 ×

2.1mm, 3.5 m particle size) at room temperature and a binary mobile phase (Solvent A:

0.1% Formic acid in water; Solvent B: 0.1% Formic acid in acetonitrile) The linear gradient elution and flow rate program used was as follows: 0.0 min, 40% B, 0.7 mL/min; 15 min,

55% B, 0.7 mL/min; 20 min 5% B, 1.0 mL/min; 22 min 100% B, 1.2 ml/min; 24 min, 40% B, 51

1.2 mL/min. After 15 min program was used to wash off all unnecessary compounds from the column. A procedure without the 15min wash program causes poor reproducibility of retention times. The UV detector was set at 360 nm.

6410 Triple Quad MS/MS setting:

The Triple Quad MS was operated in positive ion detection mode using MRM scan with electrospray ionization (ESI). Three transitions for DNB derivatives of BMAA

(451>210/254/268) and two transitions for BMAA isomers/basic amino acids were monitored as follows. (AEG: 451>210/268; BAMA: 451>254/268; DAB: 451>196/268; DAPA:

523>294; LYS: 479>267) for qualifying purpose. For the quantification program, three transitions for the bis-DNB BMAA derivative and the 454>213 transition for internal standard (BMAA-d3) were monitored at a collision energy of 10 eV. The acquisition/ion source parameters were optimized as follow: fragmentor (135 V), dwell (200), collision energy (10 eV), delta EMV(+) (400), delta EMV(-) (0). Capillary voltage (3500 V), gas temperature (325 °C), gas flow (10 L/min), nebulizer (30 psi). Nitrogen was used for the gas.

1.13 References

1. Kurland, L. T., Mulder, D. W. (1954). Epidemiologic investigations of amyotrophic lateral sclerosis: 1. Preliminary report of geographic distribution, with special reference to the

Mariana Islands, including clinical and pathologic observations. Neurology 4:355-378.

2. Mulder, D. W., Kurkland, L. T. (1987). Motor neuron disease: epidemiologic studies.

Adv. Exp. Med. Biol. 209:325-332

3. Whiting, M. G. (1988). Toxicity of cycads: implications for neurodegenerative diseases and cancer. Transcripts of Four Cycad Conferences. New York: Third World

Medical Research Foundation.

4. Spencer, P. S., Nunn, P. B., Hugon, J., Ludolph, A., Roy, D. N. (1986). Motorneuron disease on Guam: possible role of a food neurotoxin [letter]. Lancet 1:965-966

5. Spencer, P. S. (1987). Guam ALS/Parkinsonism-dementia: a long latency neurotoxin disorder caused by ‘’slow toxin(s)’’ in food? Can. J. Neurol. Sci. 14:347-357

6. Cox, P., Banack, S., Murch, S. (2003). Biomagnification of cyanobacterial neurotoxins and neurodegenerative disease among the Chammoro people of Guam. Proc.

Natl. Acad. Sci. USA 100:13380-13383

7. United States Environmental Protection Agency. (2012). Cyanobacteria and

Cyanotoxins: Information for Drinking Water Systems. Available online at: http://water.epa.gov/scitech/swguidance/standards/criteria/nutrients/upload/cyanobacteria_fac tsheet.pdf

8. Cox, P. A., Banack, S. A., Murch, S. J., Rasmussen, U., Tien, G., Bidigare, R. R.,

Metcalf, J. S., Morrison, L. F., Codd, G. A., Bergman, B. (2005). Diverse taxa of cyanobacteria produce beta-N-methylamino-L-alanine, a neurotoxic amino acid. Proc. Natl. 53

Acad. Sci. USA 102:5074-5078

9. Li, A., Fan, H., Ma, F., McCarron, P., Thomas, K., Tang, X., Quilliam, M. A. (2012).

Elucidation of matrix effects and performance of solid-phase extraction for LC-MS/MS analysis of β-N-methylamino-L-alanine (BMAA) and 2,4-diaminobutyric acid (DAB) neurotoxins in cyanobacteria. Analyst 137:1210-1219

10. Guo, T., Geis, S., Hedman, C., Arndt, M., Krick, W. (2007). Characterization of ethyl chloroformate derivative of beta-methylamino-L-alanine. J. Am. Soc. Mass. Spectrom.

18:817-82511. Bosch, L., Alegría, A., Farré, R. (2006). Application of the 6-aminoquinolyl-

N-hydroxysccinimidyl carbamate (AQC) reagent to the RP-HPLC determination of amino acids in infant foods. J. Chromatogr. B. Analyt. Technol. Biomed. Life Sci. 831:176-183.

12. Banack, S. A., Metcalf, J. S., Spáčil, Z., Downing, T. G., Downing, S., Long, A.,

Nunn P. B., Cox, P. A. (2011). Distinguishing the cyanobacterial neurotoxin β-N- methylamino-L-alanine (BMAA) in the presence of other diamino acids. Toxicon 57:730-738.

13. Jiang, L., Aigret, B., De Borggraeve, W. M., Spacil, Z., Ilag, L. L. (2012). Selective

LC-MS/MS method for the identification of BMAA from its isomers in biological samples.

Anal. Bioanal. Chem. 403:1719-1728

14. Melucci, D., Xie, M., Reschiglian, P., Torsi, G. (1999). FMOC-Cl as Derivatizing

Agent for the Analysis of Amino Acids and Dipeptides by the Absolute Analysis Method.

Chromatographia. 49:317-320

15. Kok, M. W., Scanlon, D. B., Karas, J. A., Miles, L. A., Tew, D. J., Parker, M. W.,

Barnham, K. J., Hutton, C. A. (2009). Solid-phase synthesis of homodimeric peptides: preparation of covalently-linked dimers of amyloid beta peptide. Chem. Comm. 41(41):

6228-6230

16. Van Heijenoort, J., Elbaz, L., Dezelee, P., Petit, J. F., Bricas, E., Ghuysen, J. M. 54

(1969). Structure of the meso-diaminopimelic acid containing peptidoglycans in Escherichia coli B and Bacillus megaterium KM. Biochemistry 8:207-213.

17. Work, E., Dewey, D. L. (1953). The distribution of α,ε-diaminopimelic acid among various micro-organisms. J. Gen. Microbiol. 9:394-409.

18. Wang, F., Chen, X., Chen, Q., Qin, X., Li, Z. (2000). Determination of neurotoxin 3-

N-oxalyl-2,3-diaminopropionic acid and non-protein amino acids in Lathyrus sativus by precolumn derivatization with 1-fluoro-2,4-dinitrobenzene. J. Chrom. A. 883:113-118

19. Rosen, J., Hellenaes, K. (2008). Determination of the neurotoxin BMAA (ß-N- methylamino-L-alanine) in cycad seed and cyanobacteria by LC-MS/MS. Analyst. 133(12):

1785-1789

20. Banack, S. A., Metcalf, J. S., Jiang, L., Craighead, D., Ilag, L. L., Cox, P. A. (2012).

Cyanobacteria Produce N-(2-Aminoethyl)Glycine, a Backbone for Peptide Nucleic Acids

Which May Have Been the First Genetic Molecules for Life on Earth. PLoS ONE 7(11): e49043.

21. Skoog, D. A., West, D. M., Holler, F. J., Crouch, S. R. (2004). Fundamentals of

Analytical Chemistry (8th ed.). Belmont, CA, Thomson-Brooks/Cole. 210-211

22. Ziffer, H., Hu, Y. (1990). Synthesis and optical resolution of the neurotoxin BMAA. J.

Labelled Compd. Radiopharm. 28:581-586

23. Skoog, D. A., West, D. M., Holler, F. J., Crouch, S. R. (2004). Fundamentals of

Analytical Chemistry (8th ed.). Belmont, CA, Thomson-Brooks/Cole. 142-169

Part II:

Selective Extraction Protocol and Structure of

Formamides of -Methylamino-L-alanine (BMAA) from Cycas thouarsii

2.1 Introduction

In past published papers, the BMAA content in various samples is often categorized in two forms, free BMAA and “protein-associated BMAA”.1 Free BMAA refers to the BMAA that is detectable without any acid hydrolysis, whereas the protein-associated BMAA is described as the BMAA that is detectable only upon acid hydrolysis. Most of the published papers on BMAA detection only focus on the total BMAA content as they conduct acid hydrolysis before the analysis. However, in the report of Cox and coworkers it was proposed that BMAA was present in protein from human brains of Canadian Alzheimer's patients. The authors also suggested that this protein-associated BMAA acted as a reservoir for BMAA and that it could cause neurodegenerative symptoms.2,3. Although questions regarding how

BMAA leads to neurodegenerative symptoms still remain,4, 5, 6 establishing the protocol that clearly categorizes the form of BMAA in addition to a quantification analysis seems important as well.

In Part II of this thesis, the qualitative analysis of BMAA content in samples from cycad leaf and oyster are covered. Mainly three extraction protocols are developed to categorize the form of BMAA in both cycad leaf and oyster muscle. From our initial BMAA screening after total acid hydrolysis both are known as BMAA positive samples.

Furthermore, in our own work on aspects of the BMAA problem, we had found that a local specimen of cycad, Cycas thouarsii R. Br. ex Gaudich, contained in fresh leaves up to 2,000 g BMAA /g FW. Since this analysis did not include an acid hydrolysis step, we conclude that the BMAA must have been present in its "free" (unconjugated form). However, in senescent leaves of the same plant the BMAA was not present in the form of the free amino acid, but rather in a conjugated form from which BMAA is released upon acid hydrolysis. This conjugate possesses an apparent MW of below 10,000 Da as indicated by 57 membrane filtration. We therefore set out to isolate the BMAA conjugate from C. thouarsii and determine its structure. The synthetic proof of the structure of this BMAA conjugate is also described in this chapter.

2.2 Free BMAA vs. Protein associated BMAA in Cycad leaf

We developed mainly three selective extraction protocols in order to classify/characterize the form of BMAA present in the samples. We therefore categorize the form of BMAA into three possible forms as follows: free BMAA, protein-associated

BMAA and low mass conjugated BMAA. These terms are defined as follows: "protein- associated BMAA" indicates the form of BMAA that is released upon hydrolysis of a protein fraction having more than 10 kDa MW. In this context the term “protein” is defined loosely and would include, for instance, BMAA homopolymers. The "low mass BMAA conjugate" form, on the other hand, indicates the form of BMAA that is released upon hydrolysis of a fraction containing molecules of less than 10 kDa. This could be an oligomeric form of

BMAA or a low mass conjugate of the amino acid. The total BMAA content of a specific sample determined after hydrolysis should therefore be the sum of the contributions of these three individual forms.

The first extraction was conducted in order to investigate whether BMAA exists in free form or in protein-associated/low mass conjugated form (Figure 2.1). Finely ground cycad leaf, which was already known to be BMAA-rich from our initial screening, was extracted with 80% EtOH and the extract was analyzed without any hydrolysis. In the case of cyanobacterial samples, extraction with 80% EtOH for cyanobacterial samples causes only a partial disruption of the cells, as observed by examination of the cells by microscopy. While a more complete extraction can be achieved following an acid hydrolysis protocol, this is only useful if the amount of “total BMAA” is of interest as opposed to the speciation of the

BMAA in a sample. We therefore chose to use a high-pressure cell to disrupt the cell wall in samples derived from cyanobacterial source material. This method is mechanical rather than chemical and is commonly used during the isolation of DNA and proteins from cyanobacteria. 59

If BMAA exists as “free BMAA”, BMAA will be detectable right after the initial 80% EtOH extraction or cell disruption.

As shown in Fig. 2.1, when senescent cycad leaf is extracted with 80% ethanol and the resulting extract is analyzed for BMAA content, no free BMAA is detected as shown in the middle panel. Upon acid hydrolysis of the extract, BMAA is released and can be detected as shown in the top panel. The pellet remaining after extraction was washed, subsequently hydrolyzed and analyzed for BMAA. Almost no BMAA was detected under these conditions

(bottom panel). The small amount detected (approx. <1% of total BMAA) could be due to either BMAA present in a high-molecular mass form if the latter is not efficiently extracted with 80% ethanol or could be remaining low-mass conjugated form of BMAA that had not been removed by washing of the extraction residue.

Freeze-dried cycad leaves

80% EtOH, O.N.

Supernatant

No Hydrolysis

Remaining pellet

Figure 2.1 The control extraction protocol #1 60

The next protocol was developed to examine whether BMAA is present in a high- molecular mass form, either as a homopolymer or integrated into the protein chain during protein synthesis by mis-incorporation in lieu of another amino acid. The 80% EtOH extract of cycad leaf was dried under N2 gas flow and re-suspended in water resulting in a cloudy green solution. It was centrifuged to remove green pigment and some hydrophobic proteins.

This pellet was also hydrolyzed and analyzed for BMAA, however, no trace of BMAA was detected. The aqueous supernatant was then passed through a size exclusion membrane with a

10,000 Da cutoff. It was expected that a compound having a molecular mass greater than 10 kDa will remain in the compartment on top of the membrane and all compounds with a molecular mass below 10 kDa will pass through the membrane. In order to avoid any mechanical trapping of BMAA or its conjugate in the top compartment, the membrane was washed repeatedly with water. What remained on top of the membrane and the filtrate were hydrolyzed separately. The hydrolysates were analyzed for BMAA content. As shown in

Figure 2.2, most of the BMAA content of the sample was accounted for by the material found in the filtrate (bottom panel). The small amount of BMAA detected in the hydrolysate from the top compartment is likely due to still incomplete washing of the membrane and not due to

BMAA conjugated into a high-molecular mass form. We conclude that less than 1%, if any, of the total BMAA contained in the leaf sample of C. thouarsii was present in a protein- associated or high-molecular mass form.

Aqueous extract

10,000 Da exclusion millipore membrane

Top layer

Bottom layer

Figure 2.2 Control Extraction Protocol #2

In order to characterize the low-mass conjugate of BMAA further, the aqueous phase that had been subjected to membrane filtration was washed with EtOAc and 1-butanol respectively. As shown in Fig. 2.3, the majority of BMAA remained in the aqueous extract.

While a trace amount of BMAA was detected in the n-butanol layer, this is probably due to the high solubility of water in n-butanol (73 g/L at 25 C). An aqueous solution containing a significant amount of BMAA, when extracted with 1-butanol, may therefore yield a positive test for BMAA in the n-butanol layer even if the 1-butanol used is water saturated. The results from protocol #3 further characterize the BMAA conjugate as a compound of low mass and high polarity.

Supernatant (80% EtOH)

Ethyl Acetate wash

EtOAc fraction

1-Butanol wash

1-BuOH fraction

Aqueous extract

Figure 2.3 Control Extraction Protocol #3

The same sequence of extraction protocols was also used for the analysis of a BMAA- positive sample of oyster muscle and exactly the same result was obtained: the total BMAA content is accounted for a polar low-molecular mass conjugate of BMAA.

In conclusion, in view of the Cox hypothesis invoking the incorporation of BMAA into protein in humans, we found it noteworthy that another eukaryote, Cycas sp. and oyster muscle, would contain significant amount of BMAA yet none of it was present in either the

"protein-associated" or the free amino acid form. We therefore set out to isolate the BMAA conjugate from C. thouarsii and determine its structure.

2.3 Isolation of the formamide of BMAA from Cycas thouarsii

The 80% aq. ethanol extract of freeze-dried, ground senescent leaves of C. thouarsii was dried and partitioned first between water and EtOAc and then 1-butanol. The aqueous layer from the partition was concentrated and applied to a silica flash column eluted with

EtOAc/ACN/MeOH/H2O (60/20/20/20). The sample was initially adsorbed on the silica gel resin and loaded on the column due to a solubility of the aqueous layer residue in the solvent we use. Fractions containing the BMAA conjugate were pooled and concentrated. The resulting brownish oil was subjected to repeated HPLC purification using aqueous acetonitrile (ACN/H2O = 60/20) isocratically with formic acid (0.5%) as a mobile phase additive. The only stationary phases on which any retention of the BMAA conjugate was observed were a Microsolv Diamond Hydride column, a SIELC Primesep 100 and a pentafluorophenyl column. Use of a plethora of different C18, phenyl, or HILIC columns did either not result in any retention of the compound (C18, Phenyl) or did not result in measurable separation from contaminants (HILIC). The isolation scheme is summarized in

Scheme 2.1.

freeze-dried leaves

80% EtOH, RT, O.N.

Crude extract

EtOAc wash, followed by 1-BuOH

Aqueous extract Si Column, 3 times

fraction 1 - 8 fraction 9, 10 and 11

HPLC Primesep100 Column

BMAA conjugate

Scheme 2.1 Isolation diagram of Compound 1

2.4 Elucidation of the formamide of BMAA

Compound 1 was isolated as optically active oil (approx. 3 mg) in approximately

0.002% yield. LC-HRESITOF analysis indicated a mass of 146.0683 suggesting a molecular formula of C5H10N2O3. This observed mass is 28 mass units higher than that of BMAA itself and suggests that this conjugate is a formamide derivative of BMAA as the alternative, an additional ethyl group (28 mass units higher as well), is biosynthetically unlikely. In an attempt to determine which nitrogen, alpha or beta, in the BMAA molecule is formylated, the isolated BMAA conjugate was tagged with DNB. The resulting derivative was analyzed by

LCMS using a fragmentation voltage of 10 eV. The parent ion has [M+H]+ = 313 and a major fragment ion at m/z = 210 was observed. Analysis of possible fragmentation pathways of the pseudomolecular ion suggests that the fragment ion m/z = 210 contains the β DNB moiety, the N-methyl group, the β nitrogen, and the C-3 methylene of bis-DNB-BMAA. This interpretation is supported by the appearance of a m/z= 213 fragment in the mass spectrum of the bis-DNB derivative of BMAA-d3. Since the m/z = 210 fragment does not contain the formyl group, it must be attached to the α nitrogen.

However, upon NMR analysis, it was found that three forms of the analyte appeared to be present as suggested by the presence of three formyl-H resonances in the ratio of 1:6:3 resonating at 8.15, 8.08 and 8.02 ppm, respectively. The same ratio was observed for the resonances of the N-methyl protons appearing in the ratio of 6:3:1 resonating at 3.06, 2.92 and 2.73 ppm, respectively. While one may readily explain the presence of two rotamers due to hindered rotation around the C-N bond of the amide, the presence of the third set of signals was puzzling. We therefore prepared the two regioisomeric formamides by synthesis as shown in Scheme 2.1.

O O H H HOOC HOOC HOOC NH2 NH i) ii) NH

N N NH H3C H3C H3C 3a -formyl BMAA 1b iii)

O O O HOOC HOOC O NH2 NH HOOC ii) NH i) N CH3 N NH iv) O H3C H H3C -formyl BMAA (E/Z)-1a

Scheme 2.2 Synthetic scheme of α-formyl and β-formyl BMAA. i) Acetic-formic anhydride, dioxane; ii) H2/Pearlman’s cat., 1,4-dioxane; iii) BOC anhydride, triethylamine,

O.N.; iv) TFA/CH2Cl2

For the preparation of α-formyl and β-formyl BMAA, the β-benzyl derivative of

BMAA 3a is a starting material. This material was prepared by a slight modification of the

Mannich reaction procedure described by Abe.7 For the α-formyl BMAA derivative, 3a was treated with excess formic acetic anhydride in 1,4-dioxane solution to give the  formamide.

Subsequently the benzyl group was removed by catalytic hydrogenation. The β-formyl

BMAA 3b was obtained after BOC protection of the -amino group followed by debenzylation and reaction of the amine with formic-acetic anhydride. The resulting BOC- protected formamide was deprotected using TFA/CH2Cl2 followed by evaporation in vacuo.

Both synthetic samples showed the same two three-line patterns for the formyl-H and 67 the N-methyl resonances, respectively, as the material isolated from C. thouarsii. This observation suggests that compound 1 is an equilibrium mixture between E/Z-1a and 1b, a situation similar to that observed in formyl derivatives of tetrahydrofolic acid and related model compounds.8 It appeared most likely that geometrical isomerism around the amide C-

N bond of 1a would result in the appearance of doubled signals in the methyl region of the 1H

NMR spectrum. Isomer 1b may also exist as an E/Z mixture, but this likely does not result in two N-methyl resonances for the two rotamers of 1b as the distance of the methyl to the stereogenic bond seems too large, i.e. the N-methyl is too far from the difference-causing portion. It is more likely that a second rotamer of 1b would be evident in the formyl-H region of the 1H NMR spectrum. However, inspection of the region of the spectrum where the formyl protons resonate does not indicate the existence of a second rotamer of 1b. This might be because the presence of hydrogen bonding between formamide and carboxylic acid site hinders the rotation. Furthermore, if the distribution between the putative rotamers in this regioisomer 1b is heavily skewed to a major one, a minor second conformer may not be detectable, being the minor conformer of a minor regioisomer. Lastly, it is also possible that the formyl-H resonance for the putative second rotamer of 1b overlaps with one of the resonances due to the formyl-H resonance of one of the rotamers of 1a.

Confirmation for the assignment of the resonances to the α- and β-formamide, respectively, was obtained through 13C gHMBC, 15N gHMBC, 13C gHSQC and nOe experiments. Thus, the most intense N-methyl resonance at 3.06 ppm showed a nOe interaction to the most intense formyl proton resonance at 8.08 ppm, suggesting that this pair of signals represents the Z isomer of 1a around the s-cis bond of the amide (Figure 2.4). The two other methyl resonances did not show any interaction with the other two formyl-H resonances, nor was there any evidence for exchange of magnetization between the three 68 methyl resonances. Thus, interconversion between the E and Z isomers is apparently sufficiently slow at room temperature that exchange is not evident using a 500 MHz instrument.

As this situation was not entirely satisfactory, we acquired 15N gHMBC data to support the assignment of the two three-line patterns to individual rotamers. Amides are expected to show 15N resonances between -220 and -270 ppm, while secondary amines resonate between -280 and -360 ppm in theory.9 In the event, the N-methyl resonance at 3.08 ppm and 2.92 ppm, respectively, showed a correlation to a 15N atom resonating at -268 ppm.

This indicates a contribution from an isomer in which the formyl group is attached to the methyl-bearing nitrogen atom. This pair of signals must therefore represent the rotamers of β- isomer 1a, which amounts to 84% of the total. The intensity of the N-methyl signal at 2.73 ppm for the minor isomer 1b and of all three formyl-H resonances was insufficient to observe correlations in the 15N gHMBC spectrum.

To complete the assignment of all resonances to the isomers, 13C gHMBC data were acquired, which corroborated the interpretation of the 15N gHMBC data, and allowed information to be collected also on 1b. Specifically, the most intense formyl proton resonance at 8.08 ppm correlated to the most intense N-methyl carbon resonance at 36.2 ppm and the most intense N-methyl peak at 3.06 ppm showed correlation to the most intense of the two observable formamide carbons at 167.1 ppm in the 13C NMR spectrum. The intermediate strength formyl proton resonating at 8.02 ppm correlated with the intermediate intensity N- methyl carbon at 30.4 ppm, and the intermediate intensity N-methyl proton at 2.92 ppm showed correlation to the intermediate intensity formyl carbon at 165.6 ppm. These data points corroborate the interpretation of the 15N gHMBC data and confirm that the two most intense signals in the formyl and N-methyl chemical shift range in both the 13C and 1H 69 domain belong to the Z and E isomers of 1a. 13C gHMBC also allowed us to assign the methylene carbons for 1a-(Z) and (E). The most intense formyl-H and N-methyl at 8.08 and

3.06 ppm, respectively, showed correlations to the carbon resonating at 47.0 ppm. Likewise, the intermediate intensity formyl-H and N-methyl resonating at 8.02 and 2.923 ppm, respectively, correlate with the carbon resonating at 51.6 ppm. This assigns the two methylene carbons for 1a-(Z) and 1a-(E). Conversely, the least intense N-methyl signal at

2.72 ppm did not show any correlation with a formyl 13C resonance in the gHMBC spectrum and instead correlated to methylene carbon resonance, C-3, at 52.3 ppm. The least intense formyl-H resonance showed a correlation with a methine carbon, C-2, resonating at 53.1 ppm.

These latter data points conclusively prove that the least abundant formyl derivative of

BMAA is the α-formamide 1b. 13C gHSQC was also used to see the correlation between the barely detectable N-methyl carbon in the 13C NMR spectrum resonating at 34.0 ppm and the

N-methyl proton resonance at 2.76 ppm. 13C gHSQC also showed clear correlations between the N-methyl carbon atoms and protons in 1a. However, these data are not shown in Figure

2.4. for clarity. Additionally, the 13C gHSQC experiment revealed that the methine H-2 and the H-3 methylene protons in (E/Z)-1a and 1b are resonating at 3.82 - 3.76 ppm as a multiplet.

This was proven by analysis of the phases of the cross peaks in the 13C gHSQC spectrum, i.e. two different phase colors were observed under the multiplet at 3.82 - 3.76 ppm.

The rest of the carbon atoms (carbonyl carbons in carboxylic acid, C-1) and the two methine carbons in Z/E-1a and 1b (C-2) were also assigned based on the intensity relationship of the isomers. The apparent greater stability of the β-formamide 1a parallels that of the analogous carbamic acid derivative of BMAA under physiological conditions.10 70

HOOC NH2 H HOOC NH2 HOOC N H O H

O N H N O NH

CH3 CH CH3 2 H (Z)-1a (E)-1a 1b

Figure 2.4 Summary of selected nOe ( ), 15N HMBC ( ), HMBC ( ) and HSQC

( ) data for the region-isomeric formamides 1a and 1b.

Table 2.1 500 MHz (1H) and 125 MHz (13C) NMR data for the two observable rotamers of 1a and for 1b in CD3OD.

Z-1a (major) E-1a (minor) 1b Position   HMBC   HMBC   HMBC (13C) (1H) (13C) (1H) (13C) (1H) 1 C.A. 177.0 ------172.0 ------169.9 ------2 Methine 55.1 3.82 53.9 3.78 53.1 3.76 3 Methylene 47.0 3.82 H-4,5 51.6 3.78 H-4,5 52.3 3.76 H-4 4 N-Me 36.2 3.06 H-3,5 30.4 2.92 H-3,5 34.0 2.73 H-3 5 H(C=O) 167.1 8.08 H-3,4 165.6 8.02 H-3,4 164.4 8.15

The NMR-based analysis successfully elucidated the structural information of the α- and β-formamides. However, the observation of the intense m/z = 210 fragment in the mass spectrum of the formamide mixture still needed to be explained.

2.5 mono-DNB derivatives of the BMAA formamides

As mentioned above, an ESIMS spectrum of the mono-DNB derivative of a mixture of Z/E-1a and 1b shows a strong fragment ion peak at m/z = 210, which initially we interpreted as evidence for the presence of the α-formamide 1a over the β-formamide 1b in the sample. This interpretation could not be maintained in view of the NMR evidence discussed above. We therefore prepared samples of the individual isomers of the mono-DNB derivatives of the formyl regioisomers by use of HPLC separation. A chemically synthesized equilibrium mixture of formyl BMAA (mixture of α- and β-formamides) was derivatized with

DNFB under standard condition and subjected to HPLC. After several trials, a C6-phenyl column with 20 - 100% ACN w/ 0.1% formic acid additive in 30 min using a linear gradient elution showed baseline separation of two peaks with mass m/z = 313 at the retention times of

12.1 and 13.5 - 14.1 min, respectively. The former appears as a sharp peak while the latter peak seems broadened (Figure 2.5). Mass spectra recorded in segments across the width of the entire late-eluting peak were identical, suggesting that the compound was chemically pure.

We interpreted this observation as indicating that there was some separation of the rotamers of 2a. However, this was not pursued further. Preparative separation of the mono-DNB derivative of α- and β-formamides followed by 1H NMR analysis allowed an unambiguous assignment of the structures.

Figure 2.5 HPLC/UV(340 nm) chromatogram of derivatives of the regioisomeric mono-DNB-BMAA conjugate derivatives

The nOe experiment was conducted to verify the location of the DNB and formyl moieties. The 1H NMR spectrum of the first-eluting α-formyl-β-DNB derivative 2b showed a characteristic nOe between the aromatic proton of the DNB moiety and the N-methyl protons when the N-methyl protons were irradiated. The rotamers of 2a showed distinct, different and characteristic nOe's. Specifically, the (Z)-isomer of the mixture of rotamers of 2a was characterized by the previously discussed nOe between the N-methyl resonance at 3.06 ppm and the formyl-H proton at 8.08 ppm, while irradiation of the N-methyl resonance at 2.92 ppm in the E rotamer of 2a led to enhancements of the signals for H-3a,b at 3.92 ppm and for

H-2 at 4.61 ppm. This allowed a clear assignment to be made for these two samples: the early eluting isomer is the mono-DNB derivative 2b, while the late eluting peak is due to a mixture of the rotamers of the mono-DNB derivative 2a.

The 1H NMR spectrum of E/Z-2a suggests that it is present as a 1:1 mixture of rotamers. In the case of α-formamide derivative 2b there is evidence of doubling of all signals in the 1H NMR spectrum with a ratio of approximately 5:1. The largest chemical shift difference between the two components of the mixture is observed for H-2 and the formyl-H with a Δδ of 0.45 and 0.1 ppm, respectively. We propose that this indicates that a mixture of rotamers of the α-formyl-β-mono DNB derivative is present, but we were not able to make a 73 rigorous assignment with respect to identity and configuration due to lack of sample.

O H NO2 NO2

HOOC NH

CH NO2 NO2 3 N HOOC NH HOOC NH O H O2N H

N H N O

(Z)-2a CH3 (E)-2a CH3 2b NO2

Figure 2.6 Summary of selected nOe ( ), for the regio-isomeric formamides 2a and 2b.

The fragment ion analysis on ESIMS was also conducted (Figure 2.7). The strong m/z

= 210 signal is only observed in the α-formyl-β-DNB derivative since it arises from fragmentation between C-2 and C-3 and includes a DNB group, the β-nitrogen atom, the β- methyl and C-3 methylene (Figure 1.6. and 1.9.). While our initial consideration that the intense m/z =210 fragment constitutes evidence for the presence of the -formamide 2b was correct qualitatively, it was false insofar as it arises from the minor isomer present. The major isomer in the mixture, 2a, fragments entirely differently and this fragmentation path was not understood when the first mass-spectrometry based assignment was made. With the differences in fragmentation patterns of the mono-DNB derivatives of BMAA now recognized and understood, the qualitative and quantitative differences between the NMR- based and mass-spectrometry-based results are resolved.

Figure 2.7 Fragmentations of α-formyl-β-DNB derivative (Top) and β-formyl-α-DNB derivative of BMAA (Bottom) at 10 eV collision energy

2.6 The absolute configuration

The absolute configuration of the BMAA backbone of 1a and 1b was determined by

Marfey's method after hydrolysis of the formamide11. This has to be done carefully since extended heating in 6 N HCl leads to complete racemization of BMAA. However, hydrolysis in 1 N HCl in methanolic solution for 24 hours at room temperature liberates BMAA that is only partially racemized.12

Derivatization with Marfey's reagent under the standard conditions yields the bis- dinitrophenyl alanine amide derivative, which upon LCMS analysis shows two peaks of mass m/z = 623.1 with retention times of 22.5 min and 23.2 min, respectively (Figure 2.8). The former coincides with retention time of the Marfey derivative of authentic commercial L-

BMAA, while the Marfey derivative of DL-BMAA gives two peaks in a 1:1 ratio at 22.5 and

23.2 min. The sample obtained by acid hydrolysis of the natural formamides, partially racemized during hydrolysis, was derivatized with Marfey's reagent under the standard conditions. It yields two peaks but in approximately 3(22.5 min):1(23.2 min) ratio, suggesting that N-formyl BMAA is present in the L-form. BMAA isolated without hydrolysis from young fresh C. thouarsii leaves with a BMAA content of ~2,000 g/g of free BMAA were also subjected to Marfey's analysis and showed only the peak at 22.5 min. Therefore, we conclude that the BMAA backbone of the formamides isolated from C. thouarsii and free

BMAA from young fresh C. thouarsii have L- or (S) configuration.

Figure 2.8 HPLC/UV chromatogram of Marfey-tagged DL-BMAA (Top),

L-BMAA (Middle), and partially racemized natural BMAA conjugate (Bottom)

2.7 Several notes

An isolation scheme that features the use of formic acid as a mobile phase additive and results in the isolation of a formamide needs to be critically evaluated to ensure that the formamide is not an artifact. We therefore extracted C. thouarsii leaf tissue with aq. ethanol and subjected the resulting extract to water/n-butanol partition.

Compound 1 ([M+H]+ = 147) is readily detected in the crude aqueous phase from this partition upon direct analysis by Triple Quad LCMS without using formic acid in the mobile phase as shown in Figure 2.9. Thus, the detection of the formamide of BMAA is not an artifact and thouarsiine, named after Cycas thouarsii, is a genuine natural product of C. thouarsii.

x10 5 + Scan (16.2 min) LO Frac 80%.d Subtract 3 2.8 2.6 130.10 2.4 2.2 2 + 147.10 [M+H] - 1a/1b 1.8 1.6 1.4 1.2 1 119.10 0.8 0.6 471.30 0.4 101.10 220.10 309.10 347.10 0.2 234.10 0 100 120 140 160 180 200 220 240 260 280 300 320 340 360 380 400 420 440 460 480 500 520 540 560 580 Counts vs. Mass-to-Charge (m/z)

Figure 2.9 HPLC Chromatogram of aqueous extract from C. thouarsii.(Top) and extracted mass spectrum (bottom) 78

2.8. Conclusion

In conclusion, we have determined the structure of a hydrolysable form of a low-mass conjugate of BMAA as a mixture of regioisomers of formyl- BMAA. This constitutes a first report of the determination of a structure of a hydrolysable form of BMAA and, significantly, a low molecular mass form of a BMAA conjugate. The fact that such a hydrolysable, low molecular mass form of BMAA exists should caution any interpretation that if a hydrolysable form of BMAA is found, that it is a “protein-associated” form of BMAA, whatever the precise meaning of this term may be. In our screening effort we have identified several other sources containing BMAA-conjugates. In all cases tested thus far, these conjugates are in the low-mass form. Even in the plant that contains 2 mg/g FW free BMAA, to date no evidence for the existence for a high-mass conjugated form has been found.

2.9 Experimental section

Instrumentation:

LCMS/MS analysis was performed by using an Agilent Technologies 1200 series LC system (a Quaternary Solvent Manager, Sample Manager and UV Detector) with a 6410

Triple Quadrupole LC/MS. All high-resolution mass spectral data were obtained on an

Agilent 6100 TOFMS instrument interfaced with an Agilent 1100 series LC (a Quaternary

Solvent Manager, Sample Manager and Diode Array Detector).All NMR spectral data were recorded either on a Varian Unity Inova 500 Spectrometer or a Mercury 300 Spectrometer.

Optical activity was measured on a Jasco DP-100 optical polarimeter using a 1 mL cell with

10 cm pathlength.

Isolation of BMAA conjugate (2.3):

The cycad leaf was collect from Lyon Arboretum in Manoa Valley, Honolulu, Hawaii.

The leaves were cut and flash-frozen using liquid N2, crushed and then freeze-dried.

To 150 g of freeze-dried leaves, 500 mL of 80% EtOH was added and stirred overnight at room temperature. The crude extract was dried in vacuo, re-suspended in water and then filtered through a glass fiber filter (Whatman GF1). The resulting extract was washed with an equal volume of EtOAc, and two times with an equal volume of n-BuOH to remove green pigments in the extract. The washed aqueous extract was dried in vacuo yielding about 30 g of oily black material. The oily extract was first chromatographed on a Silica gel column [100 mL Silica gel, 6:2:2:2 EtOAc:ACN:MeOH:H2O]. Sixteen fractions, each containing half a bed volume, were collected and tested for the presence of BMAA by the LCMS/MS assay described in Part I of this thesis. Fraction #9, 10, and 11 had the highest BMAA content and 80 were combined. This silica gel chromatography procedure was repeated twice. The concentration of the combined fractions was adjusted to 100 mg/mL in 20% ACN and directly separated by HPLC isocratically [Primesep100 (100 × 2.1 mm, 5 m particle size),

60% aq. ACN w/ 0.5% formic acid]. The peak of interest appeared from 10 min until 13 min and was collected. About 5 mg of isolated compound 1 was obtained (0.003% recovery).

24 1 Thouarsiine 1: colorless oil: [α] D = -11.5 (c 1.0, MeOH): H NMR (500 MHz CD3OD): δ

8.15 (s, 1H), 8.08 (s, 1H), 8.03 (s, 1H), 3.82-3.76 (m, 3H/3H/3H), 3.06 (s, 1H), 2.92 (s, 1H),

2.73 (s, 1H)

13 C NMR (500 MHz CD3OD): δ 177.0, 172.0, 169.9, 167.1, 165.6, 164.4, 55.0, 53.9, 53.0,

52.3, 51.6, 47.0, 36.2, 34.0, 30.4

+ + HREIMS calcd for [C5H10N2O3 + H] 147.0764, found [M+H] = 147.0772

Synthesis of α-formyl BMAA (2.4):

The precursor compound 3a was prepared by a Mannich reaction following a slightly modified procedure of Abe.7 Five grams (24.6 mmol) of diethyl formamidomalonate was added to a mixture of 2.98 g (24.6 mmol) N-methylbenzylamine and 2.03 g (25.0 mmol) 37% solution of formaldehyde at room temperature. The mixture was stirred for 2 hours after which 50 mL of concentrated HCl was added to the mixture. The mixture was stirred for 5 days at room temperature and then heated for 1 hour at 90 C. After concentration, the oily residue was passed through a column packed with strong anion resin (Dowex 1X8, hydroxide form) and the flow-through was collected and dried. -benzyl BMAA 3a, 1.3 g (25% recovery) was obtained as white powdery crystals after recrystallization from MeOH.

1 Intermediate 3a (-benzyl BMAA): m.p. 199-203 C; H NMR (D2O, 300 MHz): δ 7.27 (m,

5H) 3.63 (q, J = 13.0 Hz, 1H), 3.38 (s, 2H), 2.74 (m, 2H), 2.22 (s, 3H);

HREIMS: Calcd for C11H16N2O2 208.1201, found 208.1212

To 100 mg (0.48 mmol) of 3a 3 mL (34 mmol) of acetic formic anhydride prepared by mixing 1.5 mL 96% formic acid and 2.5 mL acetic anhydride, was added and stirred for 3 hours at 60 C. The resulting mixture was stirred overnight and dried by evaporation. The resulting residue (0.1 g, 0.42 mmol) was mixed with Pearlman’s catalyst (50 mg) in 3 mL of

1,4,-dioxane and debenzylated under H2 filled in a latex balloon (1.05 atm) overnight. This gave 58 mg (0.397 mmol, 83 % recovery) of an oily product (Equilibrium mixture of Z/E-1a and 1b).

Synthesis of β-formyl BMAA (2.4):

To 100 mg (0.48 mmol) of -benzyl BMAA (3a) in 8 mL of 1:1: solution of 1,4- dioxane:water, a few drops of triethylamine and 110 mg of BOC anhydride were added. The mixture was stirred overnight at RT and the BOC protected intermediate was extracted with ether. The ether layer was dried and evaporated to give 98 mg (0.32 mmol, 67% yield) of a clear oily residue. The residue was then dissolved in a 1:1 mixture of 1,4-dioxane:water (5 mL) followed by the addition of 50 mg Pearlman's catalyst and a few drops of acetic acid.

The hydrogenation reaction was conducted under balloon pressure overnight with constant stirring. After the reaction, the mixture was filtered through a glass fiber filter and dried in a speedvac. To the resulting oily residue, 3 mL (34 mmol) of formic acetic anhydride was added. The mixture was stirred overnight at RT and then evaporated. The reaction mixture was then dissolved in TFA/CH2Cl2 (1:1, 3 mL each) and stirred for 2 hours. It gave about 38 82 mg (0.26 mmol, 54% yield) of an oily product (Equilibrium mixture of Z/E-1a and 1b).

1 Synthetic BMAA-conjugate Z/E-1a/1b: H NMR (500 MHz CD3OD): δ 8.15 (s, 1H), 8.08

(s, 1H), 8.03 (s, 1H), 3.82-3.76 (m, 3H/3H/3H), 3.06 (s, 1H), 2.92 (s, 1H), 2.73 (s, 1H)

13 C NMR (500 MHz CD3OD): δ 177.0, 172.0, 169.9, 167.1, 165.6, 164.4, 55.0, 53.9, 53.0,

52.3, 51.6, 47.0, 36.2, 34.0, 30.4

+ + HREIMS calcd for [C5H10N2O3 + H] 147.0764, found [M+H] = 147.0772

Isolation of α-formyl-β-DNB and β-formyl-α-DNB derivatives (2.5.):

To 50 mg (0.34 mmol) product of α-/-formyl BMAA mixture, 100 mg (0.54 mmol) of

DNFB and 0.5 mL 1:1 ACN:0.2 M borate buffer (pH 9) were added. The mixture was heated in the heat block at 60 °C for 30 min. After the derivatization, the mixture was acidified by adding 1 N HCl and extracted with ether. The solvent was dried in vacuo and yellow oil was obtained. The product was then dissolved in MeOH and injected to HPLC [C6-Phenyl column

(150 × 2.1 mm, 5 m particle size). The HPLC was carried out using a binary mobile phase

(Solvent A: 0.1% Formic acid in water; Solvent B: 0.1% Formic acid in acetonitrile). The linear gradient elution and flow rate program used was as follows: 0 min, 20% B; 30 min,

100% B; 35 min, 20% B at fixed flow rate of 1.0 mL/min. The peaks of interest appeared at

12.1 min (sharp) and the other at 13.5 - 14.1 min (sharp). Two peaks were collected separately during repeat injections. After evaporation to dryness of these two fractions, chemically pure mono-DNB formamides E/Z-2a and mono-DNB formamide 2b, respectively, were obtained.

1 mono-DNB formamides E/Z-2a: H NMR (CD3OD, 500 MHz): δ 9.06 (d, J = 2.7 Hz, 1H) and 9.04 (d, J = 2.7 Hz, 1H), 8.30 (dd, J = 9.6, 2.8 Hz, 1H) and 8.28 (dd, J = 9.6, 2.8 Hz, 1H),

8.06 (s, 1H) and 7.95 (s, 1H), 7.21 (d, J = 9.6 Hz, 1H) and 7.14 (d, J = 9.6 Hz, 1H), 4.63 (m,

1H/1H), 3.92 (m, 2H/2H), 3.09 (s, 3H) and 2.93 (s, 3H)

+ + HREIMS m/z exact mass calcd for [C11H12N4O7 + H] 313.0779, found [M+H] = 313.0765

1 mono-DNB formamide 2b: H NMR (CD3OD, 500 MHz): δ 8.60 (d, J = 2.8 Hz, 1H), 8.22

(dd, J = 9.5, 2.8 Hz, 1H), 7.97 (s, 1H, formyl-H), 7.40 (d, J = 9.5 Hz, 1H), 4.78 (dd, J = 8.3,

4.9 Hz, 1H), 3.96 (dd, J = 14.6, 4.8 Hz, 1H), 3.65 (dd, J = 14.6, 8.5 Hz, 1H), 3.024 (s, 3H).

+ + HREIMS m/z exact mass calcd for [C11H12N4O7 + H] 313.0779, found [M+H] = 313.0765

The absolute configuration (2.6):

A 10 L sample of a 10 mg/mL solution of thouarsiine 1 in MeOH, obtained by isolation from Cycas thouarsii, was mixed with 100 L 1 N HCl in methanolic solution and stirred for 24 hours at room temperature. The solution containing de-formylated 1 was neutralized with sat. NaHCO3 and dried in a Speedvac. To this dried residue, 30 L of 10 mg/mL 1-fluoro-2-4-dinitrophenyl-5-L-alanine amide (FDAA, Marfey’s reagent) in acetonitrile and 120 L 0.2 N borate buffer (pH 9) were added. The mixture was heated in a heat block at 60 C for 30 min. The volume was adjusted to 400 L by adding 250 L of

1:1/ACN:H2O solution and 10 L of the resulting mixture was subjected to LC/MSMS. The

LC/MSMS was carried out using a Waters Xbridge column (100 × 2.1 mm, 3.5 m particle size) and a binary mobile phase (Solvent A: 0.5% Formic acid in water; Solvent B: 0.5%

Formic acid in acetonitrile) The linear gradient elution and flow rate program used was as 84 follows: 0.0 min, 10% B; 20 min, 40% B; 30 min, 100% B; 35 min, 10% B at fixed flow rate of 1.0 mL/min. The UV detector was set at 340 nm.

The Triple Quad MS was operated in positive ion detection mode using SIM scan with electrospray ionization (ESI). SIM scan was established by monitoring m/z = 623.1. The acquisition/ion source parameters were optimized as follow: fragmentor (135 V), dwell (200), collision energy (10 eV), delta EMV(+) (400), delta EMV(-) (0). Capillary voltage (3500 V), gas temperature (325 °C), gas flow (10 L/min), nebulizer (30 psi). Nitrogen was used for the gas.

2.10 References

1. Cox, P. A., Banack, S. A., Murch, S. (2003). Biomagnification of cyanobacterial neurotoxins and neurodegenerative disease among the Chammoro people of Guam. Proc.

Natl. Acad. Sci. USA 100:13380-13383

2. Murch, S. J., Cox, P. A., Banack, S. A. (2004). A mechanism for slow release of biomagnified cyanobacterial neurotoxins and neurodegenerative disease in Guam. Proc. Natl.

Acad. Sci. USA 101:12228-122231

3. Pablo, J., Banack, S. A., Cox, P. A., Johnson, T. E., Papapetropoulos, S., Bradley,

W. G., Buck, A., Mash, D. C. (2009). Cyanobacterial neurotoxin BMAA in ALS and

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Appendix:

Selected Fragmentations, UPLC Chromatograms and NMR Spectra

A.1 Fragmentation of AEG at 10 eV

A.2 Fragmentation of DAB at 10 eV

A.3 Fragmentation of BAMA at 10 eV

A.4 UPLC/FD chromatograms of eluted fractions from SPE protocol. (0.1 N, 0.5 N, 1.0 N

HCl, DI water and 2.0 N NH4OH fraction, respectively) 7 6 8 . 2 8 1 7 9 0 - . 4 2 1 k . a - 4 e 0.16 p Q - M v 0.14 A 3 i H r N e 0.12 D

0.10

U 8 A 0.08 6 9 3 8 2 0 3 3 9 9 1 7 . 9 7 0.06 6 . . 8 4 8 . 7 7 . . 6 - 9 0 1 0.04 - - - 3 u - 6 4 3 46 3 2 - y p 1 l 8 r 2 8 34 2 7 l s 0 G a 0.02 2 e 2 5 89 1 7 o G A . l . S ...... r 8 A 6 7 7 88 9 9 P 0.00 0 7 1 9 0 8 3.00 4.001 5.00 6.00 7.001 8.00 9.00 10.00 11.00 12.00 13.00 14.00 15.00 16.00 0 . 1 1 0 Minutes 2 . . . 1 4 7 3

- 0.140 - - -

k 0.130 3 y 6 Q 2 a H l 0 M 9 9 e 0.120 N G 7 A 3 4 p . . 0.110 3 7 7 . 9 v

0.100 9 8 i - 6 - r

0.090 . - e p 7 6 a D 0 0.080 s u 6 l - A 3 A 52 l 3 U 0.070 8 9. A G 8 9 r . 6 04 3 8 4 0.060 e 0 8 4 . 1 5 . 2 S 8 1 4 6 0 . 0.050 5 - . 1- . 1 33 . 5 5 0.040 3 11 7 r 7 - l 4 1 5 3 - 9 62965 5 h 7 4 4 a 4 6 0.030 0 6 1 - - 2 07861 4 4 0 T 7 3 . 1 0 l V 4 3 - 9 5 . o 3 55678 4 0 0 4 2 9 6 3 a- . 6 0.020 2 . 9 r ss ...... 6 5 . 2 4 . . Vr 4 . u . 8 P 0 yy 4 44444 . . 8 . . 2 3 o 1 4 e 0.010 7 CL 1 11111 4 6 7 9 9 1 1 n 1 L 8 0.000 . 2 2 2 93.00 4.002 5.00 6.00 7.00 8.00 9.00 10.00 11.00 12.00 13.00 14.00 15.00 16.00 1 9 1 Minutes . . - 2 4

0.16 k - - a

e Q 3 8 0.14 p M H 7

A N 5 . v 0.12 7 i r 2 e - 5 D 0.10 6 p 3 . s 1 5 6 A 8 3 1 U 0.08 1 8 9 . A 4 8 5 8 6 4 - 1 . 9 1 . 1 0.06 . 0 0 9 3 u 9 1 . . 1 - e 4 5 L - 1 1 0.04 - l 6 70 a 4 8 o - 33 - r V 3 4 r 89 y - 5 . 0.02 P l . . e T r . 5 a 44 h o 4 1 0 V 11 P 7 n 1 0.00 8 . 7 3.00 4.00 5.00 6.00 7.00 8.00 9.00 10.00 11.00 12.00 213.00 14.00 15.00 16.00 6 1 9 Minutes . - 2 k 0.18 - a e Q p 0.16 M A v i 0.14 r e 0.12 D

0.10 U 1 A 5 0.08 1 1 . 8 4 5 0.06 . 7 0.04 - 6 4 0.02 p 1 s . A 9 0.00 6 4 3.004 4.007 5.00 6.00 7.00 8.00 9.00 10.00 11.00 12.00 13.00 14.00 15.00 16.00 0 0 Minutes . . 3 4 5

0.26 0 - - 7

0.24 . Q 3 1 H 1 0.22 M A N - 0.20

A 4 0.18 A 2 M 8 0.16 B . 0 2 0 0.14 1 3 8 1 U 8 . A 0.12 2 - 48 0 1 4 4 2 9 9 9 3 4 3 85 2 3 1 1 5 . 7 7 4 7 5 3 9 6 0.10 1 8 8 k 12 8 0 3 6 5 2 1 9 0 8 7 0 . 2 . 5 a .. . . - . . 1 6 0 2 5 . 8 3 3 0.08 . 6 . e 33 3 4 5 5 . . . . 0 . . 1 5 7 p 11 1 1 l 1 1 - 6 7 8 9 1 2 3 0.06 - 9 71 8 a 9 1 174 - 312 3 4 1 42 4 - - 5 v 71-- -9 - V - - e 8 2 - - 4 6 - 9 0 - - 3 11 680 3 9 3 09 8 0.04 g 8 i 01 8 - h 8 6 2 0 4 5 5 45 r 902 5 6 8 00 2 s r p . r ..ss t. l r e u P 5 6 r y1 7 u 8 1 a o . . . y ...... 0.02 i A s 1 e 33yy e3 a o l e . . e l . . l . . l r 2 33 T 344 4 4 4 55 5 H A 1 D 11CL M1 V n I L 3 5 S G7 7 G 8 9 A P 1 11 111 1 1 1 11 1 0.00

-0.02 3.00 4.00 5.00 6.00 7.00 8.00 9.00 10.00 11.00 12.00 13.00 14.00 15.00 16.00 Minutes

1 H NMR Spectrum of isolated FMOC-2,6-meso-DAPA (500 MHz, CD3OD)

1 H NMR Spectrum of synthesized FMOC-2,6-DAPA (500 MHz, CD3OD)

13 C NMR Spectrum of isolated FMOC-2,6- meso-DAPA (75 MHz, CD3OD)

13 C NMR Spectrum of isolated FMOC-2,6-DAPA (75 MHz, CD3OD)

1 H NMR Spectrum of d3-methylbenzylamine (300 MHz, CD3Cl)

1 H NMR Spectrum of BMAA-d3 mono-hydrochloride (300 MHz, D2O)

1 H NMR Spectrum of isolated Compound 1 (500 MHz, CD3OD)

13 C NMR Spectrum of isolated Compound 1 (75 MHz, CD3OD)

1 H NMR Spectrum of synthetic mixture of (E/Z)-1a/1b (500 MHz, CD3OD)

13 C NMR Spectrum of synthetic mixture of (E/Z)-1a/1b (75 MHz, CD3OD)

1 H NMR Spectrum of intermediate 3a (300 MHz, D2O)

1 H NMR Spectrum of synthetic (E/Z)-2a (500 MHz, CD3OD)

1 H NMR Spectrum of synthetic 1b (500 MHz, CD3OD)