identification of novel lignans in the whole grain rye bran by non-targeted lc–ms metabolite...
TRANSCRIPT
ORIGINAL ARTICLE
Identification of novel lignans in the whole grain rye branby non-targeted LC–MS metabolite profiling
Kati Hanhineva • Ilana Rogachev • Anna-Marja Aura •
Asaph Aharoni • Kaisa Poutanen • Hannu Mykkanen
Received: 28 March 2011 / Accepted: 24 May 2011 / Published online: 4 June 2011
� Springer Science+Business Media, LLC 2011
Abstract Rye (Secale cereale) is among the richest die-
tary sources of lignan phytochemicals. Lignans are one of
the suggested metabolite groups to contribute to the ben-
eficial health effects of whole grain products evidenced in
epidemiological studies. So far, the complete repertoire of
lignan derivatives in rye, especially in the bran, has not
been fully described. In this study, ten novel oligomeric
sesqui- and dilignans were identified in rye bran by the use
of high resolution LC–MS analysis (i.e., UPLC-qTOF-MS/
MS). Putative identification of lignan components in the
bran was performed by combining: (i) detailed inspection
of the fragmentation behavior of available standard com-
pounds belonging to different lignan types, (ii) interpreta-
tion of MS/MS data obtained from unknown metabolites in
the samples. This combined analysis, particularly detailed
MS/MS characterization, is most valuable for non-targeted
assays in metabolite-rich matrices such as plant extracts, in
which the verification of identity with authentic standards
for each detected metabolite is normally not possible.
Metabolomics analysis will increasingly aid in deciphering
the active compounds in dietary products as part of studies
aiming at elucidating the link between human health and
nutrition.
Keywords Rye � Secale cereale � Whole grain � Bran �Lignan � Phytochemicals � Metabolite profiling �Metabolomics � LC–MS
1 Introduction
Lignans are a widely occurring group of natural products.
The core structure of lignans contains two phenylpropanoid
units (C6–C3) oxidized via a carbon–carbon single bond
(C8–C80). The core dimer can be further linked to addi-
tional phenylpropanoid units to form trimeric and tetra-
meric oligolignans commonly referred to as sesquilignans
and dilignans, respectively. In addition, other linkages than
C8–C80 have been observed, and such structures are termed
neolignans (Pan et al. 2009; Willfor et al. 2006). The dif-
ferent combinations of linkage formation allow enormous
structural diversity, and several hundred different lignan
structures have been characterized form various natural
sources mainly from cereals and conifers (Morreel et al.
2004; Pan et al. 2009). Lignans are found both as free
aglycones, sugar decorated and also esterified to the cereal
matrix (Liggins et al. 2000; Milder et al. 2004; Popova
et al. 2009; Willfor et al. 2006). The role of lignans in
plants is likely defensive, similarly as for other polyphe-
nols, and they may act as allelochemicals as well (Cutillo
et al. 2003; Harmatha and Dinan 2003; Macias et al., 2004;
Willfor et al., 2006). The composition of different lignan
metabolites varies largely not only between different spe-
cies but also within species depending on growth region
and season (Smeds et al. 2009).
The analysis of lignans is usually carried out by either
LC–MS (Eklund et al. 2008; Milder et al. 2004; Morreel
et al. 2004; Morreel et al. 2010; Willfor et al. 2006) or
GC–MS (Mazur et al. 1996; Penalvo et al. 2005a).
K. Hanhineva (&) � K. Poutanen � H. Mykkanen
Institute of Public Health and Clinical Nutrition, Food and
Health Research Centre, University of Eastern Finland,
P.O. Box 1627, 70211 Kuopio, Finland
e-mail: [email protected]
I. Rogachev � A. Aharoni
Weizmann Institute of Science, Department of Plant Sciences,
P.O. Box 26, 76100 Rehovot, Israel
A.-M. Aura � K. Poutanen
VTT Technical Research Centre of Finland,
P.O. Box 1000, 02044 VTT, Finland
123
Metabolomics (2012) 8:399–409
DOI 10.1007/s11306-011-0325-0
Lignan aglycones have several free hydroxyl groups which
are easily glucosylated, and thus the quantification of
lignans typically involves a treatment with e.g., strong acid
in order to hydrolyse the sugar conjugates (Liggins et al.
2000; Smeds et al. 2007).
Estimates on daily intake vary and have been reported
for example 430 lg/day in Finland (Valsta et al. 2003),
670 lg/day in Italy (Pellegrini et al. 2010) and 980 lg/day in
Netherlands (Milder et al. 2005a, b). Among the richest die-
tary sources of lignans are cereal brans, legumes and several
vegetables. The highest dietary source of lignans is flaxseed
(300 mg/100 g) (Milder et al. 2005a, b), also instant powder
coffee contains substantial concentration of lignans having
900 lg lignans in 100 g of powder (Kuhnle et al. 2008). In the
Japanese diet legumes like dropwort and asparagus contribute
to dietary intake of lignans both having over 1000 lg lignans
in 100 g fresh weight (Penalvo et al. 2008).
Whole grain rye is the richest cereal source of lignans;
5000–7000 lg/100 g in bran (Smeds et al. 2007), and
1900 lg/100 g in the whole kernel (Penalvo et al. 2005a).
The qualitative repertoire of lignans present in whole grain
rye has been studied earlier, and lignans belonging to dif-
ferent structural classes have been identified, including:
furofurano (syringa- and pinoresinol), furano (lariciresi-
nol), dibenzylbutyrolactone (matairesinol), and dibezylbu-
tanediol (secoisolariciresinol). Quantitatively the most
abundant lignans in rye have been reported to be syringa-
and pinoresinol, followed by lariciresinol, hydroxyma-
tairesinol, medioresinol, matairesinol, oksomatairesinol
and secoisolariciresinol (Smeds et al. 2007). Additionally,
minor lignans detected in rye include anhydro-secoisolar-
iciresinol, a-conidendrin, todolactol A and iso-hydrox-
ymatairesinol (Smeds et al. 2007).
Lignans contribute to the polyphenol intake in our diet
remarkably. Lignan-rich dietary items like whole grain rye
may have important health implications, since experimen-
tal studies have suggested that lignans possess antioxidant,
anticarcinogenic and antimicrobial, anti-inflammatory and
immunosuppressive properties (Adlercreutz and Mazur
1997; Saleem et al. 2005). Lignans have been detected
postprandially in plasma after lignan-rich meal, which
indicates that they can be absorbed from the small intestine
(Nurmi et al. 2003; Penalvo et al. 2005b). However, sev-
eral lignans (including matairesinol, secoisolariciresinol,
pinoresinol and lariciresinol) are known to proceed to
proximal colon, where they are converted by the colonic
microbiota to so called enterolignans (or mammalian
lignans), enterolactone and enterodiol, that are detectable
in urine (Adlercreutz et al. 1982; Adlercreutz et al. 1995),
plasma (Kuijsten et al. 2006) or serum (Knust et al. 2006).
Enterolignans are metabolites belonging to the phytoes-
trogen group, they have been shown to interact with
estrogen receptors. They are therefore suggested to be
among the main contributors of the biological effects of
lignan rich food like whole grain rye bread (Adlercreutz
and Mazur 1997; Clavel et al. 2006).
A prerequisite for investigating the bioavailability and
biochemical effect of any dietary phytochemical is to know
the qualitative and quantitative composition as well as
occurrence of the metabolite group in the taken plant
species or the food product made of it. Advanced meta-
bolomics technologies offer the possibility for metabolite
characterization in extensive detail, and it is thus likely that
knowledge regarding the phytochemical diversity in die-
tary plant-based food products will increase dramatically in
the near future. Here, we performed a detailed qualitative
analysis of one phytochemical group of whole grain rye
bran. We evaluated lignan composition by carrying out a
non-targeted analysis of all likely lignan derivatives. Using
this approach we have discovered a group of sesqui- and
dilignans that have not been characterized in rye before.
This work therefore adds a new set of compound to the
known repertoire in rye and further demonstrates the power
of non-targeted assays by metabolomics technologies.
2 Materials and methods
2.1 Plant material and extraction
Bran samples of rye (Secale cereale) from Finnish origin
were used for the study. The preparation of the samples
was done as described earlier (Hanhineva et al. 2011). In
brief, the water extractable fraction of bran was enriched
for phenolics in the SPE-chromatography; the residue was
dried and redissolved for UPLC-qTOF-MS (Waters)
metabolite profiling in ESI(-). LC–MS analysis, data
processing and identification were performed as described
previously (Hanhineva et al. 2011) with MassLynx and
MarkerLynx software in the data-analysis. The lignan
standards used in the LC–MS analysis were secoisolaric-
iresinol (CAS: 29388-59-8), lariciresinol (CAS: 27003-73-
2), pinoresinol (CAS: 487-36-5), and matairesinol (CAS:
580-72-3), all purchased from Arbonova, Turku, Finland.
3 Results
The qualitative analysis of the lignan metabolites in rye
bran was carried out as part of a larger study aiming at
resolving the phytochemical composition of whole grain
rye in detail. A set of four lignan compounds from struc-
turally different classes were included to serve as guide for
the lignan chromatographic behavior and MS/MS structural
fragmentation. The detailed inspection of the fragmenta-
tion of the lignan standards in ESI(-) MS/MS was
400 K. Hanhineva et al.
123
followed by search for similar compounds in the raw data
from the rye bran analysis. The examination of the
metabolite signals in the rye bran allowed the tentative
detection of ten sesqui- and dilignans that have not been
previously characterized in rye, namely stereoisomeric
pairs of buddlenol C, buddlenol D, buddlenol E, hedyotisol
A, and methoxyhedyotisol A.
3.1 LC–MS/MS analysis of the standard compounds
3.1.1 Lariciresinol (furano lignan)
One of the most characteristic fragmentations in ESI analysis
of lignans containing hydroxymethyl groups is the neutral
loss of formaldehyde from the aliphatic alcohol part of the
molecule (Table 1) (Eklund et al. 2008; Morreel et al. 2010).
The lariciresinol standard exhibits such a loss already in MS
total ion chromatogram, as the m/z 329.136 fragment is the
base peak in the spectrum (Fig. 1). In addition to this main
signal, fragments resulting from the fragmentation of the
furano ring are visible as signals of m/z 175.077 and 160.053
(Fig. 1; Table 2), in accordance with earlier publication
(Popova et al. 2009). Additionally, a smaller signal of m/z
178.064 is visible and this has been reported earlier by
Eklund et al. (2008). In the rye bran fraction a relatively small
signal could be observed in the chromatogram at the same
retention time (RT) as that of the lariciresinol standard,
having similar fragmentation with the standard.
3.1.2 Matairesinol (dibenzylbutyrolactone lignan)
Typical ESI fragmentation for methoxy containing (phe-
nolic) metabolites is the neutral loss of methyl radical ion
Table 1 Observed neutral losses in the ESI(-) MS/MS analysis of the
lignan compounds
Moiety Neutral
loss
1 Methyl radical H3C• 15.0235
2 H2O 18.0106
3 Formaldehyde H2C=O 30.0106
4 O=C=O 43.9898
5 CH2CHOH 44.0262
6 Water ? formaldehyde 48.0212
7 Guaiacylglycerol 196.0736
8 Syringoylglycerol 226.0841
Fig. 1 LC–MS analysis of the lignan standard compounds. The fragmentation is in-source fragmentation at collision energy of 25 eV
Rye lignan MS-analysis 401
123
(Table 1) (Hanhineva et al. 2008). Such cleavage is visible
for the matairesinol standard as fragment ion m/z 342.110
(Fig. 1). A reported characteristic cleavage of dib-
enzylbutyrolactones occurs on the lactone ring structure
resulting in loss of CO2. For the standard matairesinol, a
fragment ion at m/z 313.143 corresponding with the loss of
43.9896 amu was detected. The accurate mass measure-
ment enables distinguishing this loss from the cleavage of
the CH2CHOH-fragment that has the same nominal mass
(44) (Table 1). The cleavage of the benzyl group results in
the fragment of m/z 221.081, as has been reported also
earlier (Guo et al. 2007). Additional fragments in the MS/
MS analysis include m/z 298.120, 161.060, 137.060,
122.037, which also correspond to the previously reported
fragmentation of matairesinol (Guo et al. 2007). In the rye
bran sample there is a small signal observable in the
chromatogram at the same RT as that of the matairesinol
standard, matching with the fragmentation of the standard.
3.1.3 Secoisolariciresinol (dibenzylbutanediol lignan)
The fragmentation of the secoisolariciresinol standard
showed the loss of methyl radical from the molecular ion
visible as fragment of m/z 346.141, as reported earlier for
butanediol lignans (Eklund et al., 2008). In our study,
however, the loss of 48 amu mentioned earlier was hardly
observed, but the fragments resulting from the cleavage of
the C8–C80-carbons, namely m/z 179.071 and 165.055
were clearly visible in the MS/MS spectrum (Eklund et al.
2008). In addition to such reported fragmentation, also the
breakage between the carbons 7 and 8 on one of the phe-
nolic arms was visible as neutral loss of 138.0675, which is
very close to the monoisotopic mass calculated for the
detached fragment of C8H10O2 (138.0681). After such a
loss, the remaining deprotonated part of the molecule is
visible as m/z 223.098, which corresponds to the calculated
value for C12H16O4 (ES(-) 223.0970). Further fragmen-
tation of the carbon side chain may be detected as loss of
44.0268, which corresponds to the detachment of
CH2CHOH moiety (44.0262) as breakage in the bond 8–80
carbons resulting in the fragment m/z 179.071. It should be
notified that this fragmentation has the same nominal mass
as the neutral loss of CO2, (44), but accurate mass mea-
surement allows distinguishing these two fragmentations
from each other (Table 1). Alternatively, the m/z 179.071
fragment may be due to beta cleavage of the syringaresinol
molecule as has been reported (Eklund et al. 2008), but
MS3 analysis would be required to resolve which of the
Table 2 MS/MS fragmentation in ESI(-) of the putatively identified lignans in rye bran samples
Ret.
time
ESI(-)
m/zFormula ID ESI(-) MS/MS m/z
Standards
13.9 361.165 C20H26O6 Secoisolariciresinol 346.141, 315.123, 223.098, 179.071, 165.055, 147.045
14.6 359.151 C20H24O6 Lariciresinol 329.136, 192.080, 178.064, 175.077, 160.053, 116.928
17.7 357.133 C20H22O6 Pinoresinol 342.110, 311.127, 175.075, 151.040, 136.016
19.7 357.134 C20H22O6 Matairesinol 342.110, 313.143, 298.120, 221.081, 161.060, 147.046, 137.060, 122.0367
Metabolites in samples
13.9 433.150 C22H26O9 Hydroxysyringaresinol 418.127, 403.132, 385.130, 373.128, 358.104, 181.049, 166.026, 138.031
14.1 403.138 C21H24O8 Hydroxymedioresinol 388.116, 373.104, 343.114, 221.084, 181.051, 166.028, 151.004
15.5 373.129 C20H22O7 Hydroxymatairesinol 355.116, 340.093, 329.139, 311.130, 299.128, 296.105, 284.104, 178.063, 160.053,
161.062, 148.052
17.2 417.156 C22H27O8 Syringaresinol 402.132, 387.111, 372.112, 355.083, 205.052, 181.051, 166.026, 151.003
19.2 643.239 C33H40O13 Buddlenol D 595.218, 565.207, 417.154, 387.143, 357.137, 255.077, 195.066, 180.043, 165.016
19.5 613.228 C32H38O12 Buddlenol C 565.207, 535.196, 417.152, 387.143, 221.077, 195.066, 165.055, 150.031
19.8 583.218 C31H36O11 Buddlenol E 535.193, 505.170, 387.136, 372.121, 357.118, 329.116, 195.074, 165.056, 150.030
20.1 643.239 C33H40O13 Buddlenol D 595.218, 565.215, 417.154, 387.135, 357.137, 225.074, 195.066, 180.042, 165.021,
137.021
20.5 613.229 C32H38O12 Buddlenol C 565.210, 417.154, 387.143, 225.081, 195.066, 150.032
20.8 839.304 C43H52O17 Methoxyhedyotisol A 821.297, 791.290, 643.238, 613.229, 595.206, 417.148, 225.076, 195.063, 165.057
20.9 583.218 C31H36O11 Buddlenol E 387.125, 357.090, 195.061, 165.054, 150.029
21.1 809.318 C42H50O16 Hedyotisol A 743.276, 713.255, 667.221, 613.222, 565.212, 535.209, 417.155, 387.154, 195.067,
165.057, 150.032
21.6 839.304 C43H52O17 Methoxyhedyotisol A 821.298, 791.288, 645.449, 643.230, 613.223, 376.449, 225.075, 195.068
22.0 809.318 C42H50O16 Hedyotisol A 743.260, 713.259, 613.238, 595.220, 565.204, 535.204, 417.158, 387.139, 195.065,
165.056, 150.032
402 K. Hanhineva et al.
123
fragmentation patterns is in question in our case. With
comparison to the authentic standard, secoisolariciresinol
was not detected in the rye bran samples. It has been
reported that secoisolariciresinol occurs frequently as a
double glucosylated metabolite of m/z 685 (Popova et al.
2009), however, no such signal was found, neither single
sugar bearing derivative was observed in our analysis.
3.1.4 Pinoresinol (furofurano lignan)
Furofurano lignans have been shown in negative ESI to
cleave inside the tetrahydrofuran ring to give product ion of
m/z 151 in case of single methoxy-substituted phenol ring
(guaiacyl) or m/z 181 for di-methoxy substituted (syringyl)
derivatives (Eklund et al. 2008; Morreel et al. 2004;
Ye et al. 2005). The resulting fragment can be further
observed to lose a methyl radical (15 amu) (Morreel et al.
2010; Eklund et al. 2008; Ye et al. 2005), with product ions
of m/z 136 or 166, respectively. Alternative fragmentation
for furofurano lignans has been demonstrated also to occur
by breakage of bonds from the carbon 7 to carbons 8 and 9
releasing a unit of m/z 136 or 166 amu (Morreel et al.
2004), which have the same nominal mass as in the case of
loss of methyl radical in the first mentioned fragmentation
pattern. In our study, the accurate mass measurement in the
MS/MS analysis of the pinoresinol standard showed, that
the resulting fragment of m/z 136.016 is due to loss of
methyl radical having elemental composition C7H5O3,
which matches the calculated monoisotopic mass 136.0160
in ES(-) (Fig. 1). In case of the above mentioned alter-
native fragmentation, this fragment is expected to be
C8H9O2, with m/z of 136.0524 and a difference that would
have been detected.
In the extractable rye bran fraction there is a small signal
observable in the chromatogram at the same RT as that of
the pinoresinol standard. A closely related metabolite to
pinoresinol is syringaresinol, which has a methoxy group in
both the ortho and meta positions of both of the phenyl
rings (Fig. 2). A candidate metabolite for syringaresinol
was observed eluting at 17.2 min, based on the accurate
mass/elemental composition, and the fragmentation in MS/
MS, being identical to the earlier reported ones (Fig. 2,
Table 1) (Eklund et al. 2008; Morreel et al. 2004; Ye et al.
2005).
3.2 Identification of new lignan structures in rye bran
Several larger molecules in the rye bran samples elute in the
same chromatographic region and show similar fragments in
the MS/MS analysis as the lignan standards. Two such
metabolites with identical m/z value 613.228 elute at 19.5
and 20.5 min (Fig. 3). The suggested elemental formula for
these molecules is C32H38O12 exhibiting three suggestions in
the Dictionary of Natural Products (DNP), two lignans and
one antibiotic. The suggested lignans are sesquilignan
metabolites containing a syringaresinol backbone with an
additional guaiacylglycerol moiety linked via ether bond to
the free hydroxyl group of one of the phenyl moieties
(Fig. 4a). The MS/MS analysis indicated a loss of 196.0773,
which can be attributed to the fragmentation of the additional
guaiacylglycerol moiety (Fig. 4a) (Morreel et al. 2004; Yang
et al. 2007). The breakage of this ether bridge occurs easily in
the ionization of lignans, as has been demonstrated with the
sesquilignan compounds found in ‘‘butterfly bush’’ Buddleja
davidii (Houghton 1985). The resulting fragment of m/z
417.154 represents the syringaresinol backbone of the fur-
ofuran molecule, and the compound is thus assigned as
buddlenol C (Houghton 1985). The two metabolites closely
eluting with the m/z 613.228 (Fig. 3) have generally the same
fragments with slightly different proportional fragmentation,
which implies that these are two stereoisomeric forms of the
same metabolite, and can thus be distinguished from bud-
dlenol F, which has the same nominal mass, but contains a
medioresinol backbone with a syringoylglycerol side chain
and in such case the 417 mass fragment would not be visible
(Houghton 1985). These two diastereomers of buddlenol C,
holding the guaiacylglycerol side chain in two different
projections (erythro and threo) could not be distinguished
without authentic standards. All other fragments observed in
the MS/MS analysis were observed also in the fragmentation
Fig. 2 MS/MS fragmentation
of the syringaresinol lignan
detected in the rye bran. The
numbering for the neutral losses
refers to Table 1
Rye lignan MS-analysis 403
123
of the standard lignan compounds (Table 2), including
565.207 (loss of 48.0214), which correspond to the con-
comitant loss of formaldehyde and water (calc. monoiso-
topic mass 48.0212) and several losses of formaldehyde
30.0106 (Fig. 4a). The smallest fragments visible in the
spectrum resulted from the breakage of the syringaresinol
backbone and from the guaiacylglycerol moiety retaining the
charge (Fig. 4a).
Analogous fragmentation occurs also on a peak pair
with m/z 643.239 eluting close by the m/z 613.228 peaks
(Fig. 3). The molecular weight suggests a compound hav-
ing an additional methoxy group (30 amu) as compared to
buddlenol C. The elemental composition suggested for m/z
643.239 was C33H40O13, and it corresponded to five
metabolites listed in DNP, one of those, buddlenol D,
posses a syringaresinol backbone that is decorated with a
syringoylglycerol moiety. The MS/MS fragmentation
analysis confirmed the structure, as the detachment of
syringoylglycerol moiety is visible as neutral loss of
226.0838 (C11H14O5 calc. 226.0841) and the syringoyl-
glycerol moiety displays a mass signal at m/z 225.077
(Fig. 4b). All the other neutral losses were detected as in
the case of buddlenol C (Fig. 4a–b; Table 2).
Slightly after the buddlenol C and D peaks, we detected
a peak pair of m/z 809.304, both 196 amu larger than
buddlenol C (Fig. 3). A dilignan metabolite matching such
structure has been described in literature, namely hedyo-
tisol A (Matsuda et al. 1984; Yang et al. 2007). The sug-
gested elemental composition for the observed metabolite,
C42H50O16 matched the reported compound. This lignan
has the guaiacylglycerol substitution in both of the phe-
nolic moieties of the syringaresinol backbone. The MS/MS
fragmentation analysis confirmed such a dilignan structure,
as all the observed fragments matched the calculated ones,
and the smaller fragments were the same as for the bud-
dlenol C and D compounds (Fig. 5a, Table 2).
In the same chromatographic region there is an addi-
tional peak pair, 30 amu larger than the m/z 809.304
metabolite, (m/z 839.316; see Fig. 3). The increase in the
mass indicated an additional methoxy unit in the molecule.
Such a metabolite, in which one of the phenolic groups of
syringaresinol contains guaiacylglycerol, and the other
retains syringoylglycerol, has been reported previously,
namely methoxyhedyotisol A (Yang et al. 2007). All the
observable fragments in the MS/MS analysis could be
explained by such a structure (Fig. 5b), and thus the ten-
tative identification was assigned as methoxyhedytisol A.
After putatively identifying the buddlenol- and hedyo-
tisol-type syringaresinol derivatives, we screened the raw
data for other, related metabolites reported with the bud-
dlenol and hedyotisol lignans. The signal m/z 583.218
eluting in the same chromatographic region as the bud-
dlenol- and hedyotisol derivatives was identified as such
a candidate (Fig. 3). The elemental composition of
C31H36O11 suggested a metabolite with a similar structure
as the buddlenol C and D metabolites, and is called bud-
dlenol E (Houghton 1985). The m/z 583.218 metabolite
displayed a lignan-like fragmentation in the MS/MS anal-
ysis, as the loss of 196 indicated fragmentation of a gua-
iacylglycerol moiety resulting in a m/z 387.136 fragment
(Table 2). This fragment corresponded to a medioresinol
type backbone holding two methoxy groups in one of the
phenolic end groups of the furofuran structure, and one
methoxy group in the other. The fragmentation pattern of
the m/z 387.136 ion after the loss of guaiacylglycerol is
similar to the other buddlenol metabolites (Table 2). Based
on the fragmentation evidence and elemental composition
the m/z 583.218 molecule was tentatively assigned as
buddlenol E. In the same chromatographic region, elution
of other metabolites with identical mass was detected
(Fig. 3), but as they did not display a lignan-type frag-
mentation in the MS/MS analysis, they were not identified.
In addition to these metabolites, several other spectral
signals due to in-source fragmentation indicated the pres-
ence of additional lignan structures, but they had too low
signals for MS/MS analysis.
Fig. 3 Extracted ion chromatograms for the buddlenol and hedyo-
tisol lignans tentatively identified in rye bran
404 K. Hanhineva et al.
123
In addition to the large sesqui- and dilignans, also
smaller compounds resembling a lignan structure in the
MS/MS analysis were detected. Two such metabolites had
similar fragments in the MS/MS analysis as the other sy-
ringaresinol derivatives, the fragments resulting from the
cleavage of the furano ring (m/z 181.051, 166.03, 151.00)
(Fig. 6). Based on the elemental composition and frag-
mentation pattern, these two were tentatively assigned as
hydroxysyringaresinol m/z 433.150 and hydroxymediores-
inol m/z 403.138. (Table 2), both characterized earlier e.g.,
from bark of Chinese oak (Fraxinus mandschurica) (Tsu-
kamoto et al. 1984). Hydroxymedioresinol contains one
monomethoxylated phenol and one dimethoxylated phenol,
thus the residual fragments after the breakage of the fur-
ano-ring are identical with those upon MS/MS of syring-
aresinol. The additional hydroxyl group in the furano ring
results in m/z fragment 221.084, which has also been
reported earlier for hydroxymedioresinol (Tsukamoto et al.
1984). For both hydroxysyringaresinol and hydroxymedi-
oresinol the hydroxyl-group can be located to either
carbons 8 (or 80) or 9 (or 90), however, with our method the
precise location of the hydroxyl-group could not be
determined.
Finally, an additional putative lignan metabolite with m/
z 373.129 was detected. The base peak in its MS/MS
spectrum corresponded with a loss of -44 amu (Fig. 6).
Such a fragmentation was previously reported to be typical
for matairesinol in ESI MS/MS analysis, and was also
observed for the matairesinol standard (Table 2). The
smaller ions visible in the MS/MS analysis differed from
the ones observed for the pinoresinol standard and the
syringaresinol derivatives, thus it is more likely that the
observed metabolite is hydroxymatairesinol and not
hydroxypinoresinol which has the same mass and ele-
mental composition. The presence of hydroxymatairesinol
has also been reported in rye before (Smeds et al. 2007),
and the MS/MS fragments observed for hydroxymataires-
inol including m/z 355.116, 340.093, 311.130, 296.105 and
160.053 are the same as reported by others (Eklund et al.
2004).
Fig. 4 MS/MS fragmentation in ESI(-) at collision energy of 20 eV
for the m/z 613.228 metabolite putatively identified as Buddlenol C
(a); and m/z 643.239 metabolite putatively identified as Buddlenol D
(b). The numbering for the neutral losses refers to Table 1. The m/z181.051 fragment from the syringaresinol backbone is visible also in
the lower spectrum with low intensity
Rye lignan MS-analysis 405
123
4 Discussion
4.1 Occurrence of buddlenol- and hedyotisol-type
lignans
Novel buddlenol and hedyotisol-type sesqui- and dilignans
were characterized in the water extractable fraction of rye
bran. These metabolites all originate from the syringaresinol
backbone with additional decorations with the guaiacyl-
glycerol and syringoylglyserol moieties. The guaiacylglyc-
erol and syringylglycerol fragments are the major units in the
lignin polymer that are derived from coniferyl alcohol
(originating from ferulic acid) and sinapyl alcohol (origi-
nating from sinapic acid) (Morreel et al. 2004).
The buddlenol-type sesquilignans containing the guaia-
cylglycerol domain attached to the furofuran lignan were
first characterized in the plant ‘‘butterfly bush’’ (Buddleja
davidii) that is used widespreadly in folk medicine in China
(Houghton 1985). Since, such metabolites have been
detected in several other plants, e.g., sunflower (Helianthus
annuus) (Macias et al. 2004), Cestrum parqui, a shrub
indigenous to South America and widely distributed in the
Mediterranean area (Fiorentino et al. 2007), and willow
(Salix spp.) (Huvenne et al. 2008). Also the hedyotisol-type
dilignans have been known for long in traditional Asian
medicine, as they were first characterized from the leaves of
Hedyotis lawsoniae (Matsuda et al. 1984) as part of studies
focusing on the bioactive phytochemicals present in plants
used in Asian folk medicine. The other dilignan observed in
our study was methoxyhedytisol A, which has been found in
plant Tarenna attenuata together with several other lignans
and neolignans (Yang et al. 2007). Both Hedyotis lawsoniae
and Tarenna attenuata belong to the family Rubiaceae,
whereas rye is one of the grasses of the Poaceae family, and
to our knowledge this is the first report of these metabolites in
this family well known for its cereal members.
Both buddlenol type sesquilignans and hedyotisol-type
dilignans have been described as part of the oligolignol
Fig. 5 MS/MS fragmentation in ESI(-) at collision energy of 35 eV for the m/z 809.318 metabolite putatively identified as hedyotisol A (a) and
m/z 839.304 putatively identified as methoxyhedyotisol A (b). The numbering for the neutral losses refers to Table 1
406 K. Hanhineva et al.
123
pool of poplar (Populus spp.) xylem (Morreel et al. 2004).
Xylem tissue is known for extensive lignifications, and
most likely these sesqui- and dilignans are used as building
blocks for the lignan synthesis. It may be that the occur-
rence of the oligolignans such as the ones detected in this
study, is much more common than thought up to now as
only relatively few reports regarding lignans exist. It is,
therefore, likely that they will be further characterized in
other plant species, especially as part of studying lignin
formation. Even larger oligolignols such as rhyncoside,
which contains an additional guaiacylglycerol moiety
connected via an ether bond in the dilignan hedyotisol A,
are known to exist. Such structure has been described
previously in the Chinese mangrove plant (Bao et al. 2007).
4.2 Lignan content in rye
Smeds et al. (2007) reported that the most abundant lignan in
rye bran is syringaresinol (3540 lg/100 g bran) followed by
pinoresinol (1547 lg/100 g bran) (Smeds et al. 2007). In our
analysis, all the novel lignan metabolites are of furofurano-
type, mostly syringaresinol derivatives. The syringaresinol
and pinoresinol core metabolites are observable in our rye
bran LC–MS analysis, but by far more intensive signals
result from the novel buddlenol-derivatives. It will be
interesting to quantify these novel metabolites in rye bran, to
see whether the intense signals observed here are because of
higher ionization tendency than the pino- or syringaresinol
backbones, or if the sesqui- and dilignan derivatives detected
in this study are truly at higher concentration than the clas-
sical lignans. The fact that buddlenols have not been char-
acterized in rye before may be due to the fact that earlier
analyses have usually been carried out in a targeted manner
using standard compounds. One reason for the fact that the
buddlenol and hedyotisol lignans were not part of the earlier
analyses is that potentially the ether bridges connecting the
syringaresinol backbone with the additional phenylpropane
units are sensitive to the harsh extraction methods that typ-
ically have been used in the quantitative analysis of lignans
(Liggins et al. 2000; Mazur et al. 1996; Milder et al. 2004;
Smeds et al. 2007). It is known that the yields of different
lignan metabolites vary depending on the extraction method
and a combination of alkaline and acid extraction provides
the highest yield for pinoresinol and syringaresinol releasing
the esterified forms (Smeds et al. 2007). The structures
reported here may well be the precursors for the pinoresinol
or syringaresinol detected after such extractions.
In our analysis not all of the previously reported rye
lignans were detected, even when authentic standards were
included in the analysis. For example, secoisolariciresinol
and its diglucoside could not be detected. It has been
reported that the amounts of secoisolariciresinol in rye are
relatively low (400 lg/100 g versus the most abundant
lignan, syringaresinol: 3500 lg/100 g (Smeds et al. 2007)).
Fig. 6 MS/MS fragmentation in ESI(-) at collision energy of
20 eV for the metabolites putatively identified as hydroxymediores-
inol (upper panel), hydroxysyringaresinol (middle panel) and
hydroxymatairesinol (lower panel). The numbering for the neutral
losses refers to Table 1. The insert is a close-up of the spectral region
m/z 120–190 showing the fragmentation of the lignan core
Rye lignan MS-analysis 407
123
It may be that since in our study the extraction of metab-
olites was not especially optimized for concentrating
lignans, the content of secoisolariciresinol was too low to
be detected. Variation between rye varieties may also
explain the difference, as our analysis was carried on a
single Finnish rye cultivar. Several other minor lignans as
reported by Smeds et al. (2007) were searched for in the
course of this study. Candidates based on accurate mass
and elemental composition were indeed observed in the
chromatographic region where the lignans reported here
elute. However, the chromatographic signals for most of
them were too small to be used for a detailed structural
analysis by ESI–MS/MS.
4.3 Colonic formation of enterolignans from rye
lignans
Here we report the presence of eight novel oligomeric
lignan derivatives in the whole grain rye. Several plant
lignans are converted by the colonic microbiota to en-
terolignans that are absorbed from diet. It was long thought
that secoisolariciresinol and matairesinol are the only
precursors for enterolignan formation until novel precur-
sors such as pinoresinol and lariciresinol were proven to
contribute to enterolignan metabolism, whereas syring-
aresinol was suggested to be converted via an alternative
route (Heinonen et al. 2001). Furthermore, medioresinol
was suggested to be an enterolignan precursor (Penalvo
et al. 2005a) however, this has not been verified by in vitro
colonic conversion experiments. In addition, it has been
postulated that lignans released from the lignin matrix in
the acidic conditions of the digestive system could serve as
precursors for enterolignan formation in the colon by
microbiota (Begum et al. 2004). The oligomeric lignans
characterized in this study may as well be precursors and
substrates for microbiota and contribute to enterolignan
formation.
5 Concluding remarks
Our study shows how non-targeted metabolite profiling can
be utilized in order to widen the knowledge regarding the
chemical composition of edible plants and dietary products.
The targeted quantitative measurements are typically based
on earlier knowledge of the chemical composition, and thus
ignore all the rest of the molecules that, for reason or
another, have not been characterized in the plant species or
food product earlier. On the contrary, using a non-targeted
LC–MS profiling method and detailed interpretation of the
MS/MS fragmentation the assignment of metabolites with
high probability can be achieved when authentic standards
are not available for all the detected metabolites. By this
approach, we have identified and added a set of sesqui- and
dimeric derivatives to the repertoire of lignans present in
rye.
Acknowledgments This work is funded by the Nordforsk Nordic
Centre of Excellence project ‘‘HELGA—whole grains and health’’
(KH, HM). Funding from Academy of Finland is gratefully
acknowledged (KP). AA is the incumbent of the Adolpho and Evelyn
Blum Career Development Chair. The work in the Aharoni lab was
supported by the European Research Council (ERC) project SAMIT
(FP7 program) and the Benoziyo Institute. We are grateful to Arye
Tishbee for operating the LC–MS instrument and to Sagit Meir for
assistance in LC–MS sample preparation. Michael Bailey and Olavi
Myllymaki are acknowledged for technological expertise in prepa-
ration of rye fractions.
References
Adlercreutz, H., Fotsis, T., Heikkinen, R., Dwyer, J. T., Woods, M.,
Goldin, B. R., et al. (1982). Excretion of the lignans enterolac-
tone and enterodiol and of equol in omnivorous and vegetarian
postmenopausal women and in women with breast cancer.
Lancet, 2, 1295–1299.
Adlercreutz, H., & Mazur, W. (1997). Phyto-oestrogens and western
diseases. Annals of Medicine, 29, 95–120.
Adlercreutz, H., van der Wildt, J., Kinzel, J., Attalla, H., Wahala, K.,
Makela, T., et al. (1995). Lignan and isoflavonoid conjugates in
human urine. The Journal of Steroid Biochemistry and Molec-ular Biology, 52, 97–103.
Bao, S., Ding, Y., Deng, Z., Proksch, P., & Lin, W. (2007).
Rhyncosides A-F, phenolic constituents from the Chinese
mangrove plant Bruguiera sexangula var. rhynchopetala. Chem-ical and Pharmaceutical Bulletin, 55, 1175–1180.
Begum, A. N., Nicolle, C., Mila, I., Lapierre, C., Nagano, K.,
Fukushima, K., et al. (2004). Dietary lignins are precursors of
mammalian lignans in rats. The Journal of Nutrition, 134,
120–127.
Clavel, T., Dore, J., & Blaut, M. (2006). Bioavailability of lignans in
human subjects. Nutrition Research Reviews, 19, 187–196.
Cutillo, F., D’Abrosca, B., DellaGreca, M., Fiorentino, A., & Zarrelli,
A. (2003). Lignans and neolignans from Brassica fruticulosa:
Effects on seed germination and plant growth. Journal ofAgricultural and Food Chemistry, 51, 6165–6172.
Eklund, P. C., Backman, M. J., Kronberg, L. A., Smeds, A. I., &
Sjoholm, R. E. (2008). Identification of lignans by liquid
chromatography-electrospray ionization ion-trap mass spectrom-
etry. Journal of Mass Spectrometry: JMS, 43, 97–107.
Eklund, P. C., Sundell, F. J., Smeds, A. I., & Sjoholm, R. E. (2004).
Reactions of the natural lignan hydroxymatairesinol in basic and
acidic nucleophilic media: Formation and reactivity of a quinone
methide intermediate. Organic & Biomolecular Chemistry, 2,
2229–2235.
Fiorentino, A., DellaGreca, M., D’Abrosca, B., Oriano, P., Golino, A.,
Izzo, A., et al. (2007). Lignans, neolignans and sesquilignans
from cestrum parqui l’her. Biochemical Systematics and Ecology,35, 392–396.
Guo, H., Liu, A. H., Ye, M., Yang, M., & Guo, D. A. (2007).
Characterization of phenolic compounds in the fruits of forsythia
suspensa by high-performance liquid chromatography coupled
with electrospray ionization tandem mass spectrometry. RapidCommunications in Mass Spectrometry: RCM, 21, 715–729.
Hanhineva, K., Rogachev, I., Aura, A. M., Aharoni, A., Poutanen, K.,
& Mykkanen, H. (2011). Qualitative characterization of
408 K. Hanhineva et al.
123
benzoxazinoid derivatives in whole grain rye and wheat by LC–
MS metabolite profiling. Journal of Agricultural and FoodChemistry, 59, 921–927.
Hanhineva, K., Rogachev, I., Kokko, H., Mintz-Oron, S., Venger, I.,
Karenlampi, S., et al. (2008). Non-targeted analysis of spatial
metabolite composition in strawberry (fragariaxananassa) flow-
ers. Phytochemistry, 69, 2463–2481.
Harmatha, J., & Dinan, L. (2003). Biological activities of lignans and
stilbenoids associated with plant-insect chemical interactions.
Phytochemistry Reviews, 2, 321–330.
Heinonen, S., Nurmi, T., Liukkonen, K., Poutanen, K., Wahala, K.,
Deyama, T., et al. (2001). In vitro metabolism of plant lignans:
New precursors of mammalian lignans enterolactone and entero-
diol. Journal of Agricultural and Food Chemistry, 49, 3178–3186.
Houghton, J. P. (1985). Lignans and neolignans from Buddlejadavidii. Phytochemistry, 24, 819–826.
Huvenne, H., Goeminne, G., Maes, M., & Messens, E. (2008).
Identification of quorum sensing signal molecules and oligolig-
nols associated with watermark disease in willow (salix sp.).
Journal of Chromatography B, 872, 83–89.
Knust, U., Spiegelhalder, B., Strowitzki, T., & Owen, R. W. (2006).
Contribution of linseed intake to urine and serum enterolignan
levels in german females: A randomised controlled intervention
trial. Food and Chemical Toxicology: An International JournalPublished for the British Industrial Biological Research Asso-ciation, 44, 1057–1064.
Kuhnle, G. G., Dell’Aquila, C., Aspinall, S. M., Runswick, S. A.,
Mulligan, A. A., & Bingham, S. A. (2008). Phytoestrogen
content of beverages, nuts, seeds, and oils. Journal of Agricul-tural and Food Chemistry, 56, 7311–7315.
Kuijsten, A., Arts, I. C., Hollman, P. C., van’t Veer, P., & Kampman,
E. (2006). Plasma enterolignans are associated with lower
colorectal adenoma risk. Cancer Epidemiology, Biomarkers &Prevention: A Publication of the American Association forCancer Research, cosponsored by the American Society ofPreventive Oncology, 15, 1132–1136.
Liggins, J., Grimwood, R., & Bingham, S. A. (2000). Extraction and
quantification of lignan phytoestrogens in food and human
samples. Analytical Biochemistry, 287, 102–109.
Macias, F. A., Lopez, A., Varela, R. M., Torres, A., & Molinillo, J. M.
(2004). Bioactive lignans from a cultivar of Helianthus annuus.
Journal of Agricultural and Food Chemistry, 52, 6443–6447.
Matsuda, S., Kadota, S., Tai, T., & Kikuchi, T. (1984). Isolation and
structures of hedyotisol-A, -B, and C novel dilignans from
hedyotis lawsoniae. Chemical and Pharmaceutical Bulletin, 32,
5066–5069.
Mazur, W., Fotsis, T., Wahala, K., Ojala, S., Salakka, A., &
Adlercreutz, H. (1996). Isotope dilution gas chromatographic-
mass spectrometric method for the determination of isoflavo-
noids, coumestrol, and lignans in food samples. AnalyticalBiochemistry, 233, 169–180.
Milder, I. E., Arts, I. C., van de Putte, B., Venema, D. P., & Hollman,
P. C. (2005a). Lignan contents of Dutch plant foods: A database
including lariciresinol, pinoresinol, secoisolariciresinol and ma-
tairesinol. The British Journal of Nutrition, 93, 393–402.
Milder, I. E., Arts, I. C., Venema, D. P., Lasaroms, J. J., Wahala, K.,
& Hollman, P. C. (2004). Optimization of a liquid chromatog-
raphy-tandem mass spectrometry method for quantification of
the plant lignans secoisolariciresinol, matairesinol, lariciresinol,
and pinoresinol in foods. Journal of Agricultural and FoodChemistry, 52, 4643–4651.
Milder, I. E., Feskens, E. J., Arts, I. C., Bueno de Mesquita, H. B.,
Hollman, P. C., & Kromhout, D. (2005b). Intake of the plant
lignans secoisolariciresinol, matairesinol, lariciresinol, and
pinoresinol in dutch men and women. The Journal of Nutrition,135, 1202–1207.
Morreel, K., Kim, H., Lu, F., Dima, O., Akiyama, T., Vanholme, R.,
et al. (2010). Mass spectrometry-based fragmentation as an
identification tool in lignomics. Analytical Chemistry, 82,
8095–8105.
Morreel, K., Ralph, J., Kim, H., Lu, F., Goeminne, G., Ralph, S., et al.
(2004). Profiling of oligolignols reveals monolignol coupling
conditions in lignifying poplar xylem. Plant Physiology, 136,
3537–3549.Nurmi, T., Voutilainen, S., Nyyssonen, K., Adlercreutz, H., &
Salonen, J. T. (2003). Liquid chromatography method for plant
and mammalian lignans in human urine. Journal of Chroma-tography B, Analytical Technologies in the Biomedical and LifeSciences, 798, 101–110.
Pan, J. Y., Chen, S. L., Yang, M. H., Wu, J., Sinkkonen, J., & Zou, K.
(2009). An update on lignans: Natural products and synthesis.
Natural Product Reports, 26, 1251–1292.
Pellegrini, N., Valtuena, S., Ardigo, D., Brighenti, F., Franzini, L.,
Del Rio, D., et al. (2010). Intake of the plant lignans
matairesinol, secoisolariciresinol, pinoresinol, and lariciresinol
in relation to vascular inflammation and endothelial dysfunction
in middle age-elderly men and post-menopausal women living in
northern Italy. Nutrition, Metabolism, and CardiovascularDiseases: NMCD, 20, 64–71.
Penalvo, J. L., Adlercreutz, H., Uehara, M., Ristimaki, A., &
Watanabe, S. (2008). Lignan content of selected foods from
japan. Journal of Agricultural and Food Chemistry, 56,
401–409.
Penalvo, J. L., Haajanen, K. M., Botting, N., & Adlercreutz, H.
(2005a). Quantification of lignans in food using isotope dilution
gas chromatography/mass spectrometry. Journal of Agriculturaland Food Chemistry, 53, 9342–9347.
Penalvo, J. L., Heinonen, S. M., Aura, A. M., & Adlercreutz, H.
(2005b). Dietary sesamin is converted to enterolactone in
humans. The Journal of Nutrition, 135, 1056–1062.
Popova, I. E., Hall, C., & Kubatova, A. (2009). Determination of
lignans in flaxseed using liquid chromatography with time-of-
flight mass spectrometry. Journal of Chromatography A, 1216,
217–229.
Saleem, M., Kim, H. J., Ali, M. S., & Lee, Y. S. (2005). An update on
bioactive plant lignans. Natural Product Reports, 22, 696–716.
Smeds, A. I., Eklund, P. C., Sjoholm, R. E., Willfor, S. M., Nishibe,
S., Deyama, T., et al. (2007). Quantification of a broad spectrum
of lignans in cereals, oilseeds, and nuts. Journal of Agriculturaland Food Chemistry, 55, 1337–1346.
Smeds, A. I., Jauhiainen, L., Tuomola, E., & Peltonen-Sainio, P.
(2009). Characterization of variation in the lignan content and
composition of winter rye, spring wheat, and spring oat. Journalof Agricultural and Food Chemistry, 57, 5837–5842.
Tsukamoto, H., Hisada, S., & Nishibe, S. (1984). Lignans from bark
of fraximus mandshurica var japonica and F. japonica. Chemical& Pharmaceutical Bulliten (Tokyo), 32, 4482–4489.
Valsta, L. M., Kilkkinen, A., Mazur, W., Nurmi, T., Lampi, A. M.,
Ovaskainen, M. L., et al. (2003). Phyto-oestrogen database of
foods and average intake in Finland. The British Journal ofNutrition, 89(Suppl 1), S31–S38.
Willfor, S. M., Smeds, A. I., & Holmbom, B. R. (2006). Chromato-
graphic analysis of lignans. Journal of Chromatography A, 1112,
64–77.
Yang, X. W., Zhao, P. J., Ma, Y. L., Xiao, H. T., Zuo, Y. Q., He, H.
P., et al. (2007). Mixed lignan-neolignans from Tarennaattenuata. Journal of Natural Products, 70, 521–525.
Ye, M., Yan, Y., & Guo, D. A. (2005). Characterization of phenolic
compounds in the Chinese herbal drug tu-si-zi by liquid
chromatography coupled to electrospray ionization mass spec-
trometry. Rapid Communications in Mass Spectrometry: RCM,19, 1469–1484.
Rye lignan MS-analysis 409
123