Synthesis and Characterisation of α-Glycosyloxyamides Derived from Cyanogenic Glycosides

Article abstract of DOI:10.1002/pca.1237

Introduction - After exposure to oxidative stress, the leaves of some cyanogenic plants contain primary α-glycosyloxyamides with structures corresponding to their original cyanogenic glycosides.Objectives - The aim of this study was to prepare such amides

Full text of DOI:10.1002/pca.1237

Research Article
Received: 2 November 2009;
Revised: 12 April 2010;
Accepted: 15 April 2010
Published online in Wiley Online Library: 5 August 2010
(wileyonlinelibrary.com) DOI 10.1002/pca.1237
Synthesis and Characterisation of
a-Glycosyloxyamides Derived from
Cyanogenic Glycosides
Jandirk Sendker* and Adolf Nahrstedt
ABSTRACT:
Introduction – After exposure to oxidative stress, the leaves of some cyanogenic plants contain primary a-glycosyloxyamides
with structures corresponding to their original cyanogenic glycosides.
Objectives – The aim of this study was to prepare such amides from their nitrile precursors and to characterise the new
substances in order to facilitate their early identification in forthcoming studies.
Methods – A simple but highly specific method is described for the in-vitro synthesis of the amides from their nitrile glycoside
precursors using the Radziszewski reaction with hydrogen peroxide and a single-step purification of the reaction product. A
TLC method is presented for the preliminary and fast identification of the a-glycosyloxyamides.
Results – Following this procedure, seven representative a-glycosyloxyamides, five of them new, were obtained and analyti-
cally characterised by means of 1H, 13C NMR and ATR-IR spectroscopy, highlighting the differences from their respective nitrile
glycoside precursors.
Conclusion – Thus, a-glycosyloxyamides can be obtained in sufficient amounts and purity to serve as references for further
studies on the catabolism of cyanogenic glycosides and the detoxification of cyanogenic foodplants using the new aspect of
nitrile hydrolysis with (endogenous) hydrogen peroxide. Copyright © 2010 John Wiley & Sons, Ltd.
Keywords: cyanogenic glycosides; glycosyloxyamides; Radziszewski reaction; synthesis; purification; NMR; IR; TLC
fruit ripening (Hadfield and Bennett, 1997; del Rio et al., 1998),
Introduction
primary a-glycosyloxyamides corresponding to cyanogenic gly-
The capability of an organism to release hydrogen cyanide upon
cosides are likely to be found in other cyanogenic plants includ-
injury is referred to as cyanogenesis. Hydrogen cyanide release is
predominantly attributed to a-glycosyloxynitriles (cyanogenic
glycosides), which are found in numerous structural variations
throughout the plant kingdom (Lechtenberg and Nahrstedt,
1999). After drying, some cyanogenic plants contain primary
a-glycosyloxyamides with structures corresponding to their
respective cyanogenic glycosides (Nahrstedt and Rockenbach,
1993; Takeda et al., 1997; Jaroszewski et al., 2002; Backheet et al.,
2003; Hungeling et al., 2009). These amides, though first being
assumed to be artefacts resulting from the isolation procedure,
have proven to be generated from cyanogenic glycosides during
the drying of fresh cyanogenic leaf material (Olafsdottir et al.,
1991; Adersen et al., 1993; Sendker and Nahrstedt, 2009). It was
shown for the leaves of Prunus laurocerasus L. and other plant
ing important cyanogenic food plants like sorghum, cassava, flax,
black beans or almonds (Jones, 1998). Further, when investigat-
ing the cyanogenic potential of plant material, the formation of
a-glycosyloxyamides will, even during careful drying, also be
responsible for the loss of cyanogenic glycosides without libera-
tion of hydrogen cyanide.
The in-vitro conversion of cyanogenic glycosides into their cor-
responding amides using concentrated ammonia to hydrolyse
the nitrile moiety has been previously reported (Turczan et al.,
1978; Takeda et al., 1997; Jaroszewski et al., 2004). However, with
this method, the corresponding acids are formed as a major
by-product besides the low amount of the primary amides
(Turczan et al., 1978). Moreover, under these strong alkaline con-
ditions, cyanogenic glycosides with asymmetric benzyl carbons
undergo a partial stereoinversion of this carbon (Fig. 2; Nahrstedt,
species which contain the cyanogenic glucoside prunasin that
the corresponding a-glycosyloxyamide, prunasinamide,1 is gen-
1975) forming epimeric mixtures of the respective primary
amides and carboxylic acids during hydrolysis of the nitrile
erated via a facilitated reaction of the nitrile moiety of prunasin
with hydrogen peroxide evolving during the drying process
moiety (Turczan et al., 1978). In contrast, the in-vitro reaction of
(Sendker and Nahrstedt, 2009).The reaction mechanism is known
cyanogenic glycosides with hydrogen peroxide (Radziszewski
from organic chemistry as the Radziszewski reaction (Fig. 1). As
reaction) allowed a complete conversion of prunasin into its
excessive production of hydrogen peroxide in plant tissues can
generally be expected during desiccation and other pathological
processes (Smirnoff, 1993), but also during senescence including
* Correspondence to: J. Sendker, Institute of Pharmaceutical Biology
and Phytochemistry, Hittorfstr. 56, 48149 Münster, Germany. E-mail:
jandirk.sendker@uni-muenster.de
1
Although formally incorrect, we follow the established practice of naming
the amides by appending the suffix ‘amide’to the trivial name of the corre-
sponding cyanogenic glycoside.
Institute of Pharmaceutical Biology and Phytochemistry, Hittorfstr. 56, 48149
Münster, Germany
Phytochem. Anal. 2010, 21, 575–581
Copyright © 2010 John Wiley & Sons, Ltd.

J. Sendker and A. Nahrstedt
Figure 1. The Radziszewski reaction of cyanogenic glycosides with hydrogen peroxide yields the corresponding primary amides starting with the
facilitated cycloaddition of a hydroperoxide anion (Schaefer, 1970).
Figure 2. The benzylic a-carbon of aromatic cyanogenic glycosides undergoes a stereoinversion when treated under alkaline conditions (Nahrstedt,
1975) as used for the nitrile hydrolysis described by Turczan et al. (1978), Takeda et al. (1997) and Jaroszewski et al. (2004).
Figure 3. Primary amide glycosides obtained from Radziszewski reaction of cyanogenic glycosides.
corresponding a-glucosyloxyamide prunasinamide with con-
servation of its stereochemistry (Sendker and Nahrstedt,
2009).
As described by Jaroszewski et al. (2002), a-glycosyloxyamides
may give positive results in the cyanide-specific sandwich-picrate
TLC detection when using Helix pomatia enzyme (Brimer et al.,
1983). Also, other simple detection methods which provide
evidence of the sugar moiety will not distinguish between
cyanogenic glycosides and a-glycosyloxyamides; thus the
unambiguous differentiation of a putative cyanogenic glycoside
from a primary amide may be precarious and retarded until a
suitable 13C-NMR experiment reveals the lack of a nitrile carbon
resonance (Jaroszewski et al., 2002). The aim of this study there-
fore was to prepare a-glycosyloxyamides (Fig. 3) from cyano-
genic glycosides using the cheap and simple Radziszewski
reaction and to characterise the substances in order to facilitate
their early identification in forthcoming studies. Dhurrinamide,
holocalinamide, linamarinamide, neolinustatinamide and osma-
roninepoxideamide were hitherto unknown and are described
here for the first time.
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a-Glycosyloxyamides Derived from Cyanogenic Glycosides
Preparation of a-glycosyloxyamides
Experimental
Aliquots of 30 mg of each cyanogenic glycoside were dissolved in 1 mL of
10% hydrogen peroxide which had been adjusted to pH 6.8 using a
250 mM solution of sodium monohydrogenphosphate. After keeping the
reaction batches at 40°C for 1 day, each solution was subjected to a
column containing 1 g of Extrelut™, which had been packed in a 2 mL
Luer™ syringe. The Extrelut™ was eluted with 10–100 mL of ethylacetate
saturated with water (monoglucosides) or a mixture of 2–3 parts ethylac-
etate and 1 part 1-butanol saturated with water (diglycosides, e.g. gen-
tiobiosides) to remove residual cyanogenic glycosides and hydrogen
peroxide; the column was subsequently eluted with 10–100 mL of a
mixture of 3 parts ethylacetate and 1 part 1-butanol saturated with water
(monoglucosides) or a mixture of 1–2 parts ethylacetate and 1 part
1-butanol saturated with water (diglycosides, e.g. gentiobiosides) to elute
the a-glycosyloxyamides in high purity (yields between 30% and 80%).
Both reaction and separation were monitored using TLC.
Materials
All organic solvents, buffers, reagents, disposables and Extrelut™
were purchased from VWR International GmbH, Darmstadt; 4-
hydroxyphenylacetonitrile was from Sigma. Prunasin was isolated as
described previously (Sendker and Nahrstedt, 2009). Linamarin, linusta-
tin, dhurrin, holocalin, amygdalin and osmaroninexpoxide were obtained
from the collection of A.N.
Epimerisation
Aliquots of 10 mg of prunasin and prunasinamide were separately dis-
solved in 10.0 mL 50 mM phosphate buffer pH 9.0 and kept at 60°C for
30 min. The ratio of epimers was estimated using HPLC: 10 mL of each
solution was chromatographed over a 4 ¥ 150 mm ProSep C18 5 mm
column with a mixture of trifluoroacetic acid:methanol:water (1:150:849)
at 1.0 mL/min; detection was at 202 nm. The HPLC system consisted of a
Waters 600 + Waters 2690 seperations module, a Waters 717plus
autosampler and a Waters 996 PDA.
Spectroscopic analysis
Figure 4. TLC analysis shows strict correlation of the Rf-values of pri-
mary amides and their corresponding nitriles. Rf(amide) = 1.08 · Rf(nitrile)2
- 0.34 · Rf(nitrile) + 0.13 (R2 = 0.972). 4-Hydroxyphenylacetonitrile and its
corresponding amide (8) were included to confirm the predicitive power
of this model at higher Rf-values (numbers here refer to the correspond-
ing pair nitrile–amide).
1H-NMR (400 MHz) and 13C-NMR (100 MHz) spectra were recorded on a
Varian Mercury 400 plus FT-NMR spectrometer with reference to TMS. All
spectra were recorded in perdeuterated dimethyl sulfoxide at 23°C. IR
spectra were recorded with about 20 mg of pure solid substance on a
Nicolet 4700 FTIR-FMIR ATR spectrometer using a diamond as ATR crystal.
Optical rotation was measured with an Autopol™ Automatic Polarimeter.
Table 1. 1H-NMR (400 MHz) and 13C-NMR (100 MHz) chemical shifts of amides 1 and 2
Position
CONH2
Prunasinamide (1)
1H-NMRa
Amygdalinamide (2)
1H-NMRa
13C-NMRa
172.26
13C-NMRa
172.16
7.57 (1H, brs)
7.57 (1H, brs)
7.32 (1H, m)b
5.14 (1H, s)
—
7.32 (1H, m)d
5.06 (1H, s)
—
2
77.60f
136.95
128.10
128.49c
128.49c
128.49c
128.10
98.76
78.01
137.03
128.18
128.36e
128.36e
128.36e
128.18
99.14
1*
2*
3*
4*
5*
6*
1’
7.42 (1H, dd, 1.7 Hz, 8.0 Hz)
7.48 (1H, dd, 1.7 Hz, 7.8 Hz)
7.28–7.40 (3H, m)b
7.29–7.40 (3H, m)d
7.42 (1H, dd, 1.7 Hz, 8.0 Hz)
3.83 (1H, d, 7.8 Hz)
7.48 (1H, dd, 1.7 Hz, 7.8 Hz)
3.89 (1H, d, 7.8 Hz)
6’
3.41 (1H, m)
3.64 (1H, dd, 6.5 Hz, 11.8 Hz)
—
—
61.34
3.56 (1H, dd, 6.5 Hz, 11.5 Hz)
3.96 (1H, dd, 1.4 Hz, 11.0 Hz)
4.28 (1H, d, 7.8 Hz)
3.44 (1H, dd, 5.1 Hz, 12.5 Hz)
3.67 (1H, brd, 11.8 Hz)
2.96–3.21 (8H, m)
68.42
1”
6”
—
—
103.63
61.25
2’–5’
2”–5”
2.81–3.14 (4H, m)
70.41
73.74
76.10
77.45f
70.29, 70.32
73.55, 73.86
76.08, 76.33
76.96, 77.10
a Spectra recorded in DMSO-d6, values shown in ppm with reference to TMS.
b–e Signals bearing the same symbol overlap.
f Interchangeable.
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J. Sendker and A. Nahrstedt
sulphuric acid and subsequent heating to 120°C for 15 min. LOD for pure
a-glycosyloxyamides was about 0.2 nmol.
Mass spectrometric analysis
High-resolution ESI-MS spectra were recorded on a Bruker micrOTOF II in
positive mode using lithium formiate for mass calibration.
Prunasinamide (1). C14H19NO7; amorphous powder; HR-ESI-MS (posi-
tive ion mode) m/z 314.1232 [M + H]+ (calculated for C14H20NO7 m/z
314.1234); αD20 = −141.0° (CH3OH, c 0.173); IR nmax cm-1: 3332 (–OH, st),
2926 (–CH), 2890 (–CH, st), 1671 (C=O, st), 1596 (N–H), 1502 (C=C), 1459
(C=C), 1420, 1023 (st).
TLC analysis
2 mL aliquots of sample solutions were applied as a spot to a 10 ¥ 10 cm
precoated Silica gel 60 F254 aluminium plate (Merck) and developed over
a distance of 7 cm in a non-equilibrated 10 ¥ 10 cm double trough
chamber. The mobile phase was a mixture of ethylacetate:methanol:ac-
etone:dichloromethane:water (20:5:15:6:4). Glycosidic compounds were
detected by spraying with a solution of 0.5% thymol in 5% ethanolic
Amygdalinamide (2). C20H29NO12; amorphous powder; HR-ESI-MS
(positive ion mode) m/z 476.1762 [M + H]+ (calculated for C20H30NO12 m/z
Table 2. 1H-NMR (400 MHz) and 13C-NMR (100 MHz) chemical shifts of amides 3 and 4
Position
CONH2
Dhurrinamide (3)
1H-NMRa
Holocalinamide (4)
1H-NMRa
13C-NMRa
173.29
13C-NMRa
172.23
7.48 (1H, brs)
7.29 (1H, brs)
7.54 (1H, brs)
4.95 (1H, s)
—
7.18 (2H, d, 8.6 Hz)
6.70 (2H, d, 8.7 Hz)
—
6.70 (2H, d, 8.7 Hz)
7.18 (2H, d, 8.6 Hz)
4.25 (1H, d, 7.4 Hz)
3.43 (1H, m)
7.54 (1H, brs)
5.03 (1H, s)
—
6.81 (1H, s)
—
6.71 (1H, dd, 1.4 Hz, 8.2 Hz)
7.12 (1H, t, 7.8 Hz)
6.84 (1H, d, 7.9 Hz)
3.86 (1H, d, 7.5 Hz)
3.42 (1H, m)
2
79.38
128.60
128.73
114.91
157.26
114.91
128.73
101.93
61.23
77.95
138.17
115.01
157.48
115.42
129.32
118.86
98.55
1*
2*
3*
4*
5*
6*
1’
6’
61.31
3.66 (1H, dd, 5.2 Hz, 11.5 Hz)
2.99–3.21 (4H, m)
3.63 (1H, d, 11.3 Hz)
2.99–3.21 (4H, m)
2’–5’
70.20
73.78
76.21
77.39
70.42
73.73
76.06
77.39
a Spectra recorded in DMSO-d6, values shown in ppm with reference to TMS.
Table 3. 1H-NMR (400 MHz) and 13C-NMR (100 MHz) chemical shifts of amides 5–7
Position
CONH2
Linamarinamide (5)
Neolinustatinamide (6)
Osmaroninepoxideamide (7)
1H-NMRa
13C-NMRa
177.21
1H-NMRa
13C-NMRa
1H-NMRa
13C-NMRa
168.71
7.14 (1H, brs)
7.72 (1H, brs)
7.07 (1H, brs)
7.52 (1H, brs)
—
1.28 (3H, s)
1.64 (2H, m)
176.49
7.40 (1H, brs)
7.45 (1H, brs)
3.28 (1H, s)
2
3
4
—
79.59
23.21
—
82.41
24.20
29.22
59.33
60.91
69.03
1.29 (3H, s)
—
—
3.54 (1H, d, 11.7 Hz)
3.81 (1H, d, 11.7 Hz)
1.38 (3H, s)
4.13 (1H, d, 7.8 Hz)
3.41 (1H, dd, 5.4 Hz, 11.8 Hz)
3.65 (1H, d, 11.4 Hz)
—
5
1’
6’
1.35 (3H, s)
4.31 (1H, d, 7.9 Hz)
3.39 (1H, m)
3.61 (1H, dd, 3.2 Hz, 12.0 Hz)
27.83
98.42
61.20
0.81 (3H, t, 6.9 Hz)
4.32 (1H, d, 7.6 Hz)
3.48 (1H, m)
8.20
97.84
68.99
19.71
103.30
61.28
3.94 (1H, d, 10.8 Hz)
4.18 (1H, d, 7.6 Hz)
3.40 (1H, m)
1”
6”
—
—
—
—
103.40
61.23
—
—
—
3.63 (1H, d, 11.7 Hz)
2.87–3.56 (8H, m)
2’–5’
2”–5”
2.97–3.20 (4H, m)
70.31
73.80
76.95
76.98
70.25, 70.29 2.92-3.14 (4H, m)
73.76, 73.81
75.78, 76.67
70.22
73.64
76.93
77.04
76.91, 77.11
a Spectra recorded in DMSO-d6, values shown in ppm with reference to TMS.
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a-Glycosyloxyamides Derived from Cyanogenic Glycosides
476.1763); αD20 = −83.4° (CH3OH, c 0.263); IR nmax cm-1: 3312 (–OH, st),
2926 (–CH), 2885 (–CH), 1679 (C=O, st), 1593 (N–H), 1502 (C=C), 1458
(C=C), 1413, 1021 (st).
Osmaroninepoxideamide (7). C11H19NO8; amorphous powder;
HR-ESI-MS (positive ion mode) m/z 294.1183 [M + H]+ (calculated for
C11H20NO8 m/z 294.1183); αD20 = −17.7° (CH3OH, c 0.143); IR nmax cm-1:
3323 (–OH, st), 2940 (–CH), 2897 (–CH), 1674 (C=O, st), 1607 (N–H), 1436,
1377, 1032 (st).
Dhurrinamide (3). C14H19NO8; amorphous powder; HR-ESI-MS (posi-
tive ion mode) m/z 330.1180 [M + H]+ (calculated for C14H20NO8 m/z
330.1183); αD20 = +29.0° (CH3OH, c 0.102); IR nmax cm-1: 3291 (–OH, st),
2924 (–CH), 2888 (–CH), 1671 (C=O, st), 1617, 1600 (N–H), 1518 (C=C), 1451
(C=C), 1368, 1262 (phenolic), 1235 (phenolic), 1021 (st).
Results and Discussion
The preparation of the glycosyloxyamides prunasinamide (1),
amygdalinamide (2), dhurrinamide (3), holocalinamide (4),
linamarinamide (5), neolinustatinamide (6) and osmaroninep-
oxideamide (7) was achieved by simply dissolving the respective
cyanogenic glycoside in diluted hydrogen peroxide at pH 6.8 and
keeping the solution at a temperature of 40°C. Using prunasin, it
was shown that an almost complete conversion to the corre-
sponding amide was achieved after 1 week reaction time;
notably, >95% of the employed prunasin could be analytically
recovered as 1, while no by-products were detected (Sendker
and Nahrstedt, 2009). However, in order to shorten experimental
time during the present experiments we decided to isolate the
amides from the reaction mixture already after 1 day with
30–80% of the theoretical yield. The amide glycosides could
easily be obtained in high purity (no major impurities were
Holocalinamide (4). C14H19NO8; amorphous powder; HR-ESI-MS
(positive ion mode) m/z 330.1177 [M + H]+ (calculated for C14H20NO8 m/z
330.1183); αD20 = −96.5° (CH3OH, c 0.223); IR nmax cm-1: 3294 (–OH, st),
2927 (–CH), 2879 (–CH), 1673 (C=O, st), 1594 (N–H), 1489 (C=C), 1461
(C=C), 1279 (phenolic), 1265 (phenolic), 1022 (st).
Linamarinamide (5). C10H19NO7; amorphous powder; HR-ESI-MS
(positive ion mode) m/z 288.1060 [M + H]+ (calculated for C10H19NO7Na
m/z 288.1054); αD20 = −15.5° (CH3OH, c 0.076); IR nmax cm-1: 3341 (–OH, st),
2993 (–CH), 2924 (–CH), 2887 (–CH), 1665 (C=O, st), 1602 (N–H), 1403,
1366, 1038 (st).
Neolinustatinamide (6). C17H31NO12; amorphous powder; HR-ESI-MS
(positive ion mode) m/z 442.1912 [M + H+] (calculated for C17H32NO12 m/z
442.1919); αD20 = −18.3° (CH3OH, c 0.154).
Figure 5. The reduced acidity of the benzylic hydrogen of 1 is demonstrated by the failure of 1 to epimerize at pH 9.0 (lower panel). However, when
treated at pH 9.0 the corresponding nitrile prunasin [9(2R)] epimerizes to a ca. 40:60% mixture of prunasin and sambunigrin [9(2S)] (upper panel); in this
mixture we also observed a partial hydrolysis of 9(2R)/9(2S) to a mixture of c. 40% 1(2R) and 60% of its epimeric sambunigrinamide [1(2S)], thus
demonstrating that epimerisation occurs much faster than hydrolysis of the nitrile group under alkaline conditions. Nahrstedt (1975) measured a similar
epimeric ratio for alkali-treated aromatic cyanogenic glycosides using GC and trimethylsilyation. However, he could not observe the products of nitrile
hydrolysis as the corresponding amides were obviously not volatile enough due to insufficient silylation for GC analysis.
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J. Sendker and A. Nahrstedt
detected by TLC and NMR spectroscopy) by liquid–liquid chro-
matography of the reaction mixture using Extrelut™.
58.9 ppm; (ii) to the a-hydrogen resonance (if present) which is
shifted upfield by 0.49 ppm for 7 and by 0.89 to 1.15 ppm for the
aromatic compounds 1–4, representing a reduced acidity of their
benzylic proton (Fig. 5); (iii) to the resonance of the a-carbon
which is shifted downfield by 6 to 13.5 ppm; and (iv) to the reso-
nance of the adjacent sugar’s anomeric proton which is shifted
upfield by 0.21–0.63 ppm (reference data for nitriles are from
Möhrle and Fangerau, 1980; Seigler and Brinker, 1993; Lechten-
berg et al., 1994; Nakajima and Ubukata, 1998; Backheet et al.,
2003). An HMBC spectrum of 4 is shown in Fig. 6.
For chromatographic identification of the amide glycosides
along with their corresponding cyanogenic glycosides, thin-layer
chromatography was the method of choice because all com-
pounds investigated here were consistently detectable with high
sensitivity due to their sugar moieties using thymol–sulphuric
acid reagent. Separation was achieved by chromatography over
silica gel coated plates with a mobile phase widely used for
cyanogenic glycosides (Brimer et al., 1983). When using this
system, we observed a strict correlation (see Fig. 4) between the
Rf-values of the cyanogenic glycosides (always at higher Rf) and
their corresponding amides (always at lower Rf, in the case of the
amide monoglucosides almost identical to the nitrile digluco-
sides). This behaviour may support the preliminary identification
of hitherto unknown amides corresponding to known cyano-
genic glycosides or other nitriles in plant extracts.
Although these differences overcome usual solvent-
caused resonance variation and were also observed for
the pairs of acalyphin/acalyphinamide (Hungeling et al.,
2009),
lucumin/lucuminamide
(Takeda
et al.,
1997),
gynocardin/gynocardinamide (Jaroszewski et al., 2004), volkenin/
volkeninamide (Jaroszewski et al., 1987) and tetraphyllin-B/
tetraphyllin-B-amide (Jaroszewski at al., 1987), they do not
allow an unambiguous proof of the primary amide moiety.
This is especially due to the possible presence of the correspond-
ing carboxylic acids which have been repeatedly found in cyano-
genic plants (Kitajima and Tanaka, 1993; Takeda et al., 1997;
D’Abrosca et al., 2001; Fukuda et al., 2003) and are hardly
distinguishable from the corresponding amides by the NMR
characteristics mentioned above.Thus, direct spectroscopy of the
amide protons in aprotic solvents is required, e.g. in acetone-d6,
acetonitrile-d3 or dimethylsulfoxide-d6, which was also used in
this study.
The 1H- and 13C-NMR data of compounds 1–7 were recorded in
DMSO-d6 and were assigned to the respective nuclei as shown in
Tables 1–3. The NMR properties of the primary amides investi-
gated here strongly resemble those of their corresponding
nitriles. Spectral differentiation of an amide glycoside from its
corresponding nitrile is hampered by the fact that the molecular
constitution is homologue and the amide protons will not
1
appear in H-NMR spectra recorded in protic NMR-solvents like
methanol-d4. The clearest differences observed here pertain to
nuclei closely related to the carbonic acid derivative site namely:
(i) to the amide carbon resonance which, in comparison to the
respective a-glycosyloxynitriles, is shifted downfield by 51.4–
In each case, amide protons appeared between 6.7 and
7.8 ppm as two distinct broad singlets. The protons are not
Figure 6. HMBC spectrum of 4 in DMSO-d6 (400/100 MHz).
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a-Glycosyloxyamides Derived from Cyanogenic Glycosides
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Conclusion
The occurrence of a-glycosyloxyamides alone or together with
their corresponding cyanogenic glycosides is most likely in air
dried cyanogenic plant material (Sendker and Nahrstedt, 2009).
The TLC correlation presented here hints as to the presence of
a-glycosyloxyamides in an early stage of future investigations.
Hereafter the NMR and IR data sets presented will facilitate the
identification of this class of compounds even for hitherto
unknown a-glycosyloxyamides. Finally, our protocol to produce
and purify a-glycosyloxyamides from corresponding cyanogenic
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Acknowledgements
The Official Medicines Control Laboratory (OMCL) of the Institute
for Public Health Nordrhein-Westfalen is gratefully acknowl-
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