DEVELOPMENT OF AN ANALYTICAL METHOD FOR β … · proteinogenic basic amino acid,...
Transcript of DEVELOPMENT OF AN ANALYTICAL METHOD FOR β … · proteinogenic basic amino acid,...
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
IN
CHEMISTRY
MAY 2014
by
Yoshiaki Miyasaka
Thesis Committee:
Thomas K. Hemscheidt, Chairperson
Phillip Williams
Joseph T. Jarrett
ii
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 .
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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.
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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
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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
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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
ix
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 ( ), 15
N 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
xi
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
[α]T
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
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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
2
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.
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.
H3CNH
NH2
OH
O
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.
8
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.
9
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.
11
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
12
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
Hydrolysate
Crude amino acid mixture
Tagged amino acid mixture
0 - 140 mL fractions 140 - 160 mL fraction
19.9 - 20.45 min fraction0 - 19.9 min fraction
Si fraction
Isolated FMOC-DAPA
6 N HCl @ 100C, O.N.
Dowex-50 cation exchange resin
FMOC Derivatization
100 cm LH-20 Column
HPLC C18 Column
Si Column, 3 times
LH-20 Column
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.
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.
OH
O O
HO
NH2 NH2
2,6-meso-Diaminopimelic acid
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.
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
NO2
NO2
F
+HN
NH2
COOHN
NH
COOH
DNB
DNB
Tagged BMAA
[M+H]+=451.1
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.
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.
4 x10
0 0.25 0.5
0.75 1
1.25 1.5
1.75 2
2.25 2.5
2.75 3
3.25 3.5
3.75 4
4.25 4.5
4.75 + Product Ion (13.147 min) (451.0 -> **) BMAA Prod Ion Scan 10.d
134.1 198.0
210.1
236.1
254.1
268.1
451.1
Counts vs. Mass-to-Charge (m/z) 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
NH
HOOC
N
H3C
O2N
NO2O2N
NO2
210
268
254
18
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
451.1>210.1
451.1>254.1
451.1>268.1
451.1>210.1
451.1>268.0
451.1>164.0
451.1>254.1
451.1>268.2
451.1>134.1
451.1>196.0
451.1>268.1
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
H3CN
HN
OH
O
DNB
DNB
N
HN OH
ODNB
BND
N
OHNH
O
BND
BNDHN
HN
OH
O
DNB
BND
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.
20
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.
21
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.
23
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)
24
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
synthesized by using methylamine-d3 according to the method of Ziffer et al,22
due to
unavailability of methylamine-d3 in Hawaii, BMAA-d3 was synthesized through a different
synthetic scheme as follows. (Scheme 1.2)
25
EtO Cl
O
+NH2 N
H
O
EtO
NH
O
EtOTHF, reflux, 3 hours
NH
D
DD
NH
D
DD
COOMeNH
O
+60 oC, 48 hours MeOOC
HN
O
N
D3C
MeOOC
HN
O
N
D3C
Pearlman's catalyst
H2 (g), RT, 24 hours
MeOOC
HN
O
NH
D3C
MeOOC
HN
O
NH
D3C
3 M HCl
110 oC, 4 hours HOOC
H2N
NH
D3C
2 HCl
1) Dowex-50 Clean-up
2) 1 equiv. HCl
HOOC NH2
NH
CD3
BMAA-d3 monohydrochloride
HOOC
H2N
NH
D3C
2 HCl HCl
LiAlD4
Scheme 1.2 Synthetic scheme of BMAA-d3
26
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.
27
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)
4 x10
0 0.2 0.4 0.6 0.8
1 1.2 1.4 1.6 1.8
2 2.2 2.4 2.6 2.8
3 + Product Ion (13.1 min) (454.00 -> **) D3BMAA Trial#1 prod10.d
213.10
454.20 254.10
271.10 134.10 201.00 236.00 168.10
Counts vs. Mass-to-Charge (m/z) 80 100 120 140 160 180 200 220 240 260 280 300 320 340 360 380 400 420 440 460 480 500
NH
HOOC
N
D3C
O2N
NO2O2N
NO2
213
271
254
28
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
(ng/mL)
Mass (ng) Concentration
(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
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.
y = 0.0145x - 0.508
R² = 0.9994 CurveA
y = 0.0137x - 0.5247
R² = 0.9988 CurveB
y = 0.012x - 0.2427
R² = 0.9998 CurveC
-10
0
10
20
30
40
50
60
70
80
0 1000 2000 3000 4000 5000 6000
STD Curve A
STD Curve B
STD Curve C
Linear (STD Curve A)
Linear (STD Curve B)
Linear (STD Curve C)
Concentration
(ng/mL)
STD/ISTD area ratio
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 rangey = 0.0113x - 0.0021
R2 = 0.9999
-1
-0.5
0
0.5
1
1.5
2
0 10 20 30 40 50 60
Concentration (ng/mL)
STD
/IST
D a
rea
rati
o
Curve A Middle range
y = 0.0118x - 0.0139
R2 = 0.9998
-1
0
1
2
3
4
5
6
7
8
0 100 200 300 400 500 600
Concentration (ng/mL)
STD
/IST
D a
rea
rati
o
31
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).
Curve A High rangey = 0.0148x - 1.7109
R2 = 0.9999
0
10
20
30
40
50
60
70
80
0 1000 2000 3000 4000 5000 6000
Concentration (ng/mL)
STD
/IST
D a
rea
rati
o
32
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.
34
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.
35
Table 1.5 Summary of Matrix-adapted curves
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
y = 0.0145x - 0.508
R² = 0.9994 STD
y = 0.0138x - 0.6053
R² = 0.9982 TOM
y = 0.0149x - 0.1865
R² = 0.999 FISH
y = 0.0144x - 0.4716
R² = 0.9991 SP
-10
0
10
20
30
40
50
60
70
80
0 1000 2000 3000 4000 5000 6000
STD Curve A
STD Tomato
STD Spirulina
STD Fish
Linear (STD Curve A)
Linear (STD Tomato)
Linear (STD Spirulina)
Linear (STD Fish)Concentration (ng/mL)
STD/ISTD area ratio
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%
39
1.11 Sample and LC/MS sample preparation protocol
In summary, the overall protocol is shown in Scheme 1.3.
Scheme 1.3 Summary of sample preparation
~20 mg Biological
samples
80% EtOH extract
- 2 mL 80% EtOH
- Stir overnight at room temp.
- Internal Standard spike (50 ng)
- Centrifuged
- Evaporation (Speedvac, 35 °C)
- Re-suspend in 2 mL DI water
- Centrifuged
Supernatant
Cell disruption Vessel
(Cyanobacterial samples only)
- 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
(Speedvac, 35 °C)
- Derivatization
- Derivatization
Hydrolyzable BMAA Free BMAA
~20 mg Biological samples
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.
41
Scheme 1.4 Overall flow of the method
Small Scale Extraction
LCMS/MS Analysis
Large Scale Extraction
SPE Cartridge
- Sample Preparation
- Sample Preparation
The result is ambiguous
LCMS/MS Analysis
Successful Quantificatio Successful Quantification
42
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.
43
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.
44
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
A) and CH3CN with 0.1% formic acid (mobile phase B); the flow rate was 3.0 ml min-1
. 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.
Isolated FMOC-2,6-meso-DAPA: 1H 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);
13C 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
Standard FMOC-2,6-DAPA: 1H 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);
13C 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
47
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.
d3-Methylbenzylamine (3a): 1H 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
48
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.
BMAA-d3 mono-hydrochloride: 1H 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:
UPLC/FD Method A was performed using and AccQ-TAGTM
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.
UPLC/TUV Method B was performed using and ACCQ-TAGTM
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.
52
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
56
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.
58
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.
Figure 2.1 The control extraction protocol #1
Freeze-dried cycad leaves
Remaining pellet
Supernatant
80% EtOH, O.N.
No Hydrolysis
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.
61
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.
Aqueous extract
Top layer
Bottom layer
10,000 Da exclusion
millipore membrane
62
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.
Supernatant (80% EtOH)
EtOAc fraction
1-BuOH fraction
Aqueous extract
Ethyl Acetate wash
1-Butanol wash
63
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.
64
Scheme 2.1 Isolation diagram of Compound 1
freeze-dried leaves
Crude extract
Aqueous extract
fraction 1 - 8 fraction 9, 10 and 11
BMAA conjugate
80% EtOH, RT, O.N.
EtOAc wash, followed by 1-BuOH
Si Column, 3 times
HPLC Primesep100 Column
65
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.
66
HOOCNH2
N
H3C
HOOCNH
N
H3C
HOOCNH
NH
H3C
HOOCNH
N
H3C
HOOCNH
NH
HOOCNH2
N
-formyl BMAA
-formyl BMAA
i) ii)
iii)
ii) i)
iv)
OH O
H
OO
H3C
OO
HO
CH3
1b
3a
(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 1
H 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 13
C gHMBC, 15
N gHMBC, 13
C 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 15
N gHMBC data to
support the assignment of the two three-line patterns to individual rotamers. Amides are
expected to show 15
N 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 15
N 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 15
N gHMBC spectrum.
To complete the assignment of all resonances to the isomers, 13
C gHMBC data were
acquired, which corroborated the interpretation of the 15
N 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 13
C 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 15
N gHMBC data and confirm that the two most
intense signals in the formyl and N-methyl chemical shift range in both the 13
C and 1H
69
domain belong to the Z and E isomers of 1a. 13
C 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 13
C 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. 13
C gHSQC was also used to see the correlation between the
barely detectable N-methyl carbon in the 13
C NMR spectrum resonating at 34.0 ppm and the
N-methyl proton resonance at 2.76 ppm. 13
C 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 13
C 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 13
C 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
Figure 2.4 Summary of selected nOe ( ), 15
N 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 (13
C) (1H)
HMBC (13
C) (1H)
HMBC (13
C) (1H)
HMBC
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.
N
O
NH2HOOC
CH3
HN
H
NH2HOOC
CH3
O NH
HNHOOC
CH2
O
H
H
(Z)-1a (E)-1a 1b
71
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.
72
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.
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.
N
NHHOOC
HO
O2N
NO2
CH3
H
N
NHHOOC
NO2
NO2
N
NHHOOC
NO2
NO2
CH3
H
O
O
H
CH3(Z)-2a (E)-2a 2b
74
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
75
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.
76
Figure 2.8 HPLC/UV chromatogram of Marfey-tagged DL-BMAA (Top),
L-BMAA (Middle), and partially racemized natural BMAA conjugate (Bottom)
77
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.
Figure 2.9 HPLC Chromatogram of aqueous extract from C. thouarsii.(Top) and
extracted mass spectrum (bottom)
5 x10
0 0.2 0.4 0.6 0.8
1 1.2 1.4 1.6 1.8
2 2.2 2.4 2.6 2.8
3 + Scan (16.2 min) LO Frac 80%.d Subtract
130.10
147.10
119.10
471.30 101.10 220.10 309.10 347.10
234.10
Counts vs. Mass-to-Charge (m/z) 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
[M+H]+ - 1a/1b
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.
79
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).
Thouarsiine 1: colorless oil: [α]24
D = -11.5 (c 1.0, MeOH): 1H 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)
13C 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.
81
Intermediate 3a (-benzyl BMAA): m.p. 199-203 C; 1H 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).
Synthetic BMAA-conjugate Z/E-1a/1b: 1H 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)
13C 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.
83
mono-DNB formamides E/Z-2a: 1H 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
mono-DNB formamide 2b: 1H 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.
85
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
Alzheimer’s disease. Acta. Neurolog. Scan. 120:216-225
4. Lobner, D., Piana, P. M., Salous, A. K., Peoples, R. W. (2007). β-N-methylamino-L-
alanine enhances neurotoxicity through multiple mechanisms. Neurobiology of Disease
25(2):360-366
5, Rao S.D., Banack S.A., Cox P.A., Weiss J.H. (2006). BMAA selectively injures motor
neurons via APA/Kainate receptor activation. Exp. Neurol. 201:244-252
6, Liu X., Rush T., Zapata J., Lobner, D. (2009). beta-N-methylamino-l-alanine
induces oxidative stress and glutamate release through action on
system Xc. Exp. Neurol. 217:429-433.
7. Abe, N., Fujusaki, F., Sumoto, K. (1998). Synthesis of -(sec-Amino)alanines. Chem.
Pharm. Bull. 46(1):142-144
8. Benkovic, S. J., Bullard, W. P., Benkovic, P. A. (1972). Models for tetrahydrofolic
acid. III. Hydrolytic interconversions of the tetrahydroquinoxaline analogs at the formate
level of oxidation. J. Am. Chem. Soc. 94:7542-7549.
86
9. Levy, G. C., Lichter, R. L. (1979). Nitrogen-15 Nuclear Magnetic Resonance
Spectroscopy. Wiley-Interscience, NY
10. Nunn, P. B., O’Brien, P. (1989). The interaction of beta-N-methylamino-L-alanine
with bicarbonate: an 1H-NMR study. FEBS Lett. 251:31-35.
11, Bhushan, R, Brückner, H. (2004). Marfey's reagent for chiral amino acid analysis: a
review. Amino Acids. 27(3-4):231-47.
12. John, C., Sheehan, J. C., Yang, D. H. (1958). The use of N-Formylamino Acids in
Peptide Synthesis J. Am. Chem. Soc. 80(5):1154-1158.
Appendix:
Selected Fragmentations, UPLC Chromatograms and NMR Spectra
88
A.1 Fragmentation of AEG at 10 eV
A.2 Fragmentation of DAB at 10 eV
A.3 Fragmentation of BAMA at 10 eV
89
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)
AMQ - 2.978
NH3 - 4.140
6.286
Ser - 6.698
Gly - 7.126
7.224
7.583
Asp - 7.709
8.013
Glu - 8.338
8.834
8.946
9.123
Ala - 9.493
9.772
Pro - 10.879
Deriv peak - 12.867
AU
0.00
0.02
0.04
0.06
0.08
0.10
0.12
0.14
0.16
Minutes
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
AMQ - 3.000
NH3 - 4.111
4.644
6.290
Ser - 6.697
Gly - 7.119
7.504
7.580
Asp - 7.706
8.005
Glu - 8.339
8.565
Thr - 8.836
9.133
9.247
9.423
Ala - 9.492
9.775
Pro - 10.883
12.614
Deriv peak - 12.870
Cys - 13.146
Lys - 13.249
13.304
Val - 14.095
nor-Val - 14.207
14.329
14.444
14.506
14.572
14.636
14.689
14.766
14.815
Leu - 15.653A
U
0.000
0.010
0.020
0.030
0.040
0.050
0.060
0.070
0.080
0.090
0.100
0.110
0.120
0.130
0.140
Minutes
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
AMQ - 2.992
NH3 - 4.122
Asp - 7.578
9.141
Pro - 10.886
Deriv peak - 12.870
Tyr - 13.698
Val - 14.095
nor-Val - 14.313
14.534
14.837
14.930
15.486
Leu - 15.652
Phe - 15.918A
U
0.00
0.02
0.04
0.06
0.08
0.10
0.12
0.14
0.16
Minutes
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
AMQ - 2.967
4.151
Asp - 7.581
9.146
Deriv peak - 12.870
AU
0.00
0.02
0.04
0.06
0.08
0.10
0.12
0.14
0.16
0.18
Minutes
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
AMQ - 3.046
3.588
NH3 - 4.074
His - 5.213
5.662
Ser - 6.627
Arg - 6.842
Gly - 7.017
7.124
Asp - 7.583
7.706
Glu - 8.294
8.849
9.150
Ala - 9.507
Pro - 10.859
BMAA - 11.705
11.859
12.539
Deriv peak - 12.824
12.873
13.077
13.111
Cys - 13.184
Lys - 13.258
13.309
13.417
13.514
Tyr - 13.698
Met - 13.820
13.898
13.963
Val - 14.031
14.081
14.202
nor-Val - 14.300
14.533
14.694
14.831
15.004
15.092
15.284
Ile - 15.314
Leu - 15.652
Phe - 15.918
AU
-0.02
0.00
0.02
0.04
0.06
0.08
0.10
0.12
0.14
0.16
0.18
0.20
0.22
0.24
0.26
Minutes
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
90
1H NMR Spectrum of isolated FMOC-2,6-meso-DAPA (500 MHz, CD3OD)
1H NMR Spectrum of synthesized FMOC-2,6-DAPA (500 MHz, CD3OD)
91
13C 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)
92
1H NMR Spectrum of d3-methylbenzylamine (300 MHz, CD3Cl)
1H NMR Spectrum of BMAA-d3 mono-hydrochloride (300 MHz, D2O)
93
1H NMR Spectrum of isolated Compound 1 (500 MHz, CD3OD)
13
C NMR Spectrum of isolated Compound 1 (75 MHz, CD3OD)
94
1H NMR Spectrum of synthetic mixture of (E/Z)-1a/1b (500 MHz, CD3OD)
13C NMR Spectrum of synthetic mixture of (E/Z)-1a/1b (75 MHz, CD3OD)
95
1H NMR Spectrum of intermediate 3a (300 MHz, D2O)
96
1H NMR Spectrum of synthetic (E/Z)-2a (500 MHz, CD3OD)
1H NMR Spectrum of synthetic 1b (500 MHz, CD3OD)