Planta

, 226:639

Occurrence of “mammalian” lignans in plant and water sources

Authors

    • Department of Biochemistry and PharmacyÅbo Akademi University
  • Stefan M. Willför
    • Process Chemistry Centre, Laboratory of Wood and Paper ChemistryÅbo Akademi University
  • Suvi P. Pietarinen
    • Process Chemistry Centre, Laboratory of Wood and Paper ChemistryÅbo Akademi University
  • Pirjo Peltonen-Sainio
    • Plant Production ResearchMTT Agrifood Research Finland
  • Markku H. T. Reunanen
    • Process Chemistry Centre, Laboratory of Wood and Paper ChemistryÅbo Akademi University
Original Article

DOI: 10.1007/s00425-007-0512-4

Cite this article as:
Smeds, A.I., Willför, S.M., Pietarinen, S.P. et al. Planta (2007) 226: 639. doi:10.1007/s00425-007-0512-4

Abstract

Enterolignans, also called “mammalian” lignans because they are formed in the intestine of mammals after ingestion of plant lignans, were identified for the first time in extracts of four tree species, i.e., in knot heartwood of the hardwood species Fagus sylvatica and in knot or stem heartwood of the softwood species Araucaria angustifolia, Picea smithiana, and Abies cilicia. They were also identified for the first time in grain extracts of cultivated plants, i.e., in 15 cereal species, in 3 nut species, and in sesame and linseeds. Furthermore, some plant lignans and enterolignans were identified in extracts of water from different sources, i.e., in sewage treatment plant influent and effluent and in humic water, and for the first time also in tap and seawater. They were present also in water processed through a water purification system (ultrapure water). As enterolignans seem to be abundant in the aquatic environment, the occurrence of enterolignans in plant sources is most likely due to uptake by the roots from the surrounding water. This uptake was also shown experimentally by treating wheat (Triticum aestivum ssp. vulgare) seeds with purified lignan-free water spiked with enterolactone (EL) during germination and growth. Both the remaining seeds and seedlings contained high EL levels, especially the roots. They also contained metabolites of EL, i.e., 7-hydroxy-EL and 7-oxo-EL.

Keywords

AbiesAraucariaCerealsEnterolignansFagusPicea smithiana

Abbreviations

ASE

Accelerated solvent extraction

CLar

Cyclolariciresinol

DHEL

4,4′-Dihydroxyenterolactone

ED

Enterodiol

EL

Enterolactone

GC-MS

Gas chromatography-mass spectrometry

HEL

7-Hydroxyenterolactone

HMR

7-Hydroxymatairesinol

HPLC-MS/MS

High-performance liquid chromatography-tandem mass spectrometry

Lar

Lariciresinol

MDMR

Monodemethylated matairesinol

MR

Matairesinol

OEL

7-Oxoenterolactone

RP

Reversed-phase

Sec

Secoisolariciresinol

STP

Sewage treatment plant

TOC

Total organic carbon

Introduction

Plant lignans are secondary plant metabolites which structurally are dimers of phenylpropane units linked together by a β-β bond. Plant lignans occur in tree species, especially in wood knots (the branch bases encased in the tree stem), which have recently been shown to contain exceptionally large amounts of lignans (Willför et al. 2003; Holmbom et al. 2003). Plant lignans are also present in our food, i.e., in vegetables such as legumes, fruits, berries, cereals, nuts, and oilseeds (Mazur and Adlercreutz 1998; Milder et al. 2005; Peñalvo et al. 2005; Smeds et al. 2007), with flaxseed and sesame seeds (Milder et al. 2005) as the richest food sources.

In the intestine, plant lignans are known to be converted by intestinal microbiota to the enterolignans enterodiol (ED) and enterolactone (EL) (Axelson and Setchell 1980). These compounds were first discovered in urine from humans and some other mammalian species (Setchell et al. 1980), and were thus called “mammalian” lignans. They have been shown to possess health-promoting properties such as antioxidant and antitumour effects (Kitts et al. 1999; Prasad 2000; Saarinen et al. 2002a). Epidemiological studies show that a high serum EL concentration is associated with a reduced risk for breast cancer (Adlercreutz et al. 1982; Ingram et al. 1997; Pietinen et al. 2001) and coronary heart disease (Vanharanta et al. 1999; 2002). More recently, also other enterolignans were identified as plant lignan metabolites, i.e., 7-hydroxy-EL (HEL) (Heinonen et al. 2001; Smeds et al. 2004), 4,4′-dihydroxy-EL (DHEL), monodemethylated matairesinol (MDMR), and 7-oxo-EL (OEL) (Smeds et al. 2005).

It has been assumed that enterolignans are produced only in the intestine of mammals and are therefore likely to be found only in body fluids of mammals. Recently, however, Lee et al. (2004) detected ED and EL at low concentrations in herbs of several plant families. Nevertheless, the uniqueness of this discovery was not discussed at all in the work. In our HPLC-MS/MS analyses, we discovered that the HPLC eluent water contained EL although it had been produced by processing deionised tap water through a water purification system (Saarinen et al. 2002b; Smeds and Hakala 2003). Our findings on the presence of EL in water were very recently confirmed in a study by Kang et al. (2006). They detected ED and EL in wastewater treatment plant influent and in creek water.

In this work, plant lignans and/or enterolignans were analysed in 22 species of cultivated plants, in knot- and stemwood of more than 50 tree species, and in water from various sources. Furthermore, in order to find a possible link between the occurrence of enterolignans in plants and in water (which could be up taken by the roots), wheat seeds were treated with purified, lignan-free water spiked with EL during germination and growth, after which enterolignans were analysed in the extracts of roots, coleoptiles, and remaining seeds.

Materials and methods

Chemicals

All solvents were of analytical grade and purchased from commercial sources. The chemical structures of the analysed compounds are shown in Fig. 1. The reference substances were prepared in our laboratory. (-)-7-Hydroxymatairesinol (HMR), (−)-secoisolariciresinol (Sec) and (+)-lariciresinol (Lar) were isolated from knots of Picea abies, Araucaria angustifolia, and Abies balsamea, respectively (Freudenberg and Knof 1957; Anderegg and Rowe 1974; Smeds et al. 2004). (-)-Matairesinol (MR) was prepared from (-)-HMR (Freudenberg and Knof 1957). (±)-HEL (Mäkelä et al. 2001), (±)-OEL, and (±)-EL (for the germination experiment and for preparation of EL-d6) were prepared using total synthesis. MR-d6, DHEL, and MDMR, were prepared from (-)-MR (Adlercreutz et al. 1995a; Mäkelä et al. 2000), and EL-d6 from (±)-EL (Wähälä et al. 1986). (+)-Cyclolariciresinol (CLar) was prepared by treating (+)-Lar with concentrated formic acid. HMR and HEL were mixtures of two isomers differing in stereochemistry at C-7. ED and EL for analytical purposes were purchased from Fluka Chemie (Buchs, Switzerland). The purities of the prepared lignans as determined by GC-MS (as silylated compounds) were the following (%): (-)-HMR 98.3, (-)-MR 99.8, (-)-Sec 98.2, (+)-Lar 97.0, (+)-CLar 92.3, HEL 80.8, DHEL 94.0, OEL 93.2, MDMR 99.0, EL-d6 92.6, MR-d6 98.2.
https://static-content.springer.com/image/art%3A10.1007%2Fs00425-007-0512-4/MediaObjects/425_2007_512_Fig1_HTML.gif
Fig. 1

Chemical structures of the analysed lignans

β-Glucuronidase/sulfatase (type H-1, from Helix pomatia) (Sigma-Aldrich Co., St Louis, MO, USA) containing 492 units/mg of β-glucuronidase and 10 units/mg of sulphatase was used for the enzymatic hydrolysis.

Equipment

Deionised tap water samples were purified using a Simplicity 185 water purification system (Millipore Corp.) to produce ultrapure water and then by eluting through a glass column packed with approx. 50 g of Bondesil 40 μm RP-18 material (Varian Inc., Harbor City, CA, USA). This lignan-free water was used in the HPLC eluent, as blank water, and in the germination experiment. The water samples were solid-phase extracted using 1 cc Oasis cartridges (Waters Corp., Milliford, MA, USA).

The wheat seeds and seedlings for the germination experiment were freeze-dried using a Christ Gamma 1–20 freeze-drier (Martin Christ Gefriertrocknungsanlagen GmbH, Osterode am Harz, Germany). Nuts and oilseeds were milled using a Polymix analytical mill A 10 (Kinematica AG, Lucerne, Switzerland). The bran was separated from the cereal grains using a Faribon 300 grain mill (Gensaco Inc., New York, NY, USA) and the bran was further milled using an IKA MF 10 mill with cutting grinding head (IKA Group). Samples of wood and cultivated plants were extracted using an Accelerated Solvent Extractor (ASE) apparatus (Dionex Corp., Sunnyvale, CA, USA). Quartz sand (granular 1–2 mm) used in the ASE extraction was purchased from J.T. Baker. A Christ Beta 1–8 freeze drier was used for evaporation of water in the ASE extracts of wheat seeds and seedlings and in enzyme solution of a methyl tert-butyl extracted tap water sample.

Lignans in water extracts and in extracts of Fagus sylvatica (beech) and cultivated plants were quantified by HPLC-electrospray ionisation-MS/MS using a Micromass Quattro Micro instrument (Micromass, Manchester, UK) in the multiple reaction monitoring and negative mode as described previously (Smeds et al. 2004, 2005). Lignans in extracts of softwood species were quantified on a 25 m × 0.20 mm i.d., 0.11 μm HP-1 capillary column (Agilent Technologies, Palo Alto, CA, USA) using a Perkin Elmer AutoSystem XL GC (Perkin Elmer, Boston, MA, USA). The identity of individual lignans in extracts of softwood, a tap water sample, and a reference sample was confirmed by GC-MS analysis with an HP 6890-5973 GC-quadrupole-MSD instrument using a similar column as above.

Sample collection, pretreatment, and quantification procedures

Water samples

Ultrapure water was produced from deionised tap water by using a Millipore water purification system. The analysed tap water samples were Turku municipal drinking water. The drinking water treatment plant in Turku utilises stream water with a total organic carbon (TOC) content around 10 mg/l as a raw water source. Seawater was collected from the archipelago in southwestern Finland and humic water from Lake Pilvilampi in Western Finland. Sewage treatment plant (STP) influent and effluent flow was collected at the Turku STP, which utilises activated sludge treatment with simultaneous addition of ferric salt, and sedimentation and biological treatment of the sludge.

The water samples were prepared as follows: To 5 ml of the samples, 1 mg of enzyme preparation dissolved in 0.5 ml of 0.01 M acetate buffer (pH 5.0) was added and the samples were incubated at 37°C overnight. The internal standards were added (250 ng of MR-d6 and 110 ng of EL-d6) and the samples were solid-phase extracted. The lignans were eluted with 1 ml of acetone. The acetone was evaporated to dryness under nitrogen gas and the samples were redissolved in 0.3 ml of methanol/0.1% acetic acid 20/80, v/v.

In another laboratory, a 0.5 l tap water sample was extracted three times with 300 ml of methyl tert-butyl ether, the combined organic phase was evaporated to dryness, and enzymatic hydrolysis was performed as described above. The solution was freeze-dried overnight before qualitative GC-MS analysis. This analysis was done in order to check that EL detected in water samples was not due to contamination in the laboratory.

Both HMR and HEL were quantified as a sum of their two stereoisomers. The quantification was performed using calibration curves of standard solutions. The standard solutions were not hydrolysed; otherwise they were treated as the other samples. ED and EL were quantified against EL-d6, the other lignans against MR-d6. The standard solutions consisted of blank water spiked with HMR, MR, Sec, Lar, CLar, HEL, ED, and EL at six different concentration levels.

Wood samples

More than 50 tree species (including both soft- and hardwood) available at our laboratories were screened for enterolignans in the knot- and stemwood. Knot heartwood of A. angustifolia and of five Picea smithiana trees were sampled from the Brazilian rain forest and from Pakistan (Shawar forest, District Swat, Northwest Frontier Province), respectively. Stem heartwood of Abies cilicia was sampled from a full-grown tree in Northern Turkey and knot heartwood of F. sylvatica from a forest in Slovenia. The wood samples were splintered, freeze-dried, and ground. The beech sample was treated as the samples of cultivated plants (see below). The other wood samples, except the P. smithiana sample, which was Soxhlet extracted with acetone for 8 h, were sequentially extracted in an ASE apparatus according to Willför et al. (2003). The hydrophilic extracts were stored at −18°C until GC-MS analysis, which was done according to a previously described method (Ekman and Holmbom 1989).

Samples of cultivated plants

Grains of rye, wheat, oats, barley, spelt wheat, buckwheat, millet, quinoa, amaranth, brown rice, wild rice, red rice, sesame seeds, linseeds, peanuts, almonds, cashew nuts, and walnuts were purchased from local stores. Dhurra grains and Japanese rice bran were kindly provided by Dr. Takeshi Deyama, Yomeishu Seizo Co., Ltd., Nagano, Japan. Corncobs were purchased from a local store; the grains were gathered, frozen, and freeze-dried. Grains of triticale, var. Prego were obtained from Boreal Kasvinjalostus Oy, Jokioinen, Finland.

Approximately 1 g of the cereal bran samples and the beech sample was weighed, thoroughly mixed with quartz sand, and poured into ASE extraction tubes. The ASE extraction was performed as described previously (Willför et al. 2003) with the following modification: the material was extracted first with n-hexane, then with acetone, and finally with acetone–water (70:30, v/v). The three fractions were collected into separate glass tubes. The acetone and acetone–water fraction were combined and the solvent was evaporated to dryness using a rotary evaporator. The extract was redissolved in 7–10 ml of acetone–water (70:30, v/v). An aliquot of 0.5 ml was transferred into a test tube and the solvent was evaporated using a stream of nitrogen gas. Enzymatic hydrolysis was performed as with the water samples. One hundred and ten nanograms of EL-d6 was added and the solutions were liquid–liquid extracted with 2 × 0.75 ml of ethyl acetate. The ethyl acetate phase was evaporated to dryness under nitrogen gas and 0.5 ml of methanol/acetonitrile/0.1% acetic acid (15:15:70, by vol.) was added. The samples were placed in an ultrasonic bath for 5 min and then centrifuged for 10 min at 3,200 g. Quantification was performed as with the water samples, however, the standard solutions consisted of EL, ED, HEL, OEL, MDMR, and DHEL at six concentration levels dissolved in 0.01 M acetate buffer pH 5.0 and extracted with ethyl acetate as the real samples.

Germination procedure

The experiment included two treatments, control and EL application. For the control treatment, five groups containing 100 grains each of spring wheat cultivar Amaretto with 99% germination capacity were evenly placed on blotting paper that was thoroughly moistened with purified, lignan-free water. Five replicates of control treatment blotting papers were all loosely rolled up and placed into individual polythene bags, which were closed. To enable gas exchange, one corner of each bag was cut. All bags were placed in an upright position and kept at 5°C for 4 days and thereafter at room temperature (about 22°C) for 6 days. EL treatment was carried out similarly to the control treatment, except that the blotting paper was moistened with EL solution, which was prepared by adding 50 mg EL to 500 ml of lignan-free water, stirring the solution for 4 h and filtering the solution prior to its use for moistening. EL treatment was started and terminated 1 day after the control treatment.

After 10 days, each roll of replicated blotting paper was opened and the seedlings were divided with a surgeon’s knife into three fractions: coleoptiles including first leaves, roots, and remaining seeds still having some reserves. Each EL treated fraction was rinsed five times with lignan-free water to remove possible traces of EL on the plant surfaces. Seeds or seedlings contaminated with fungus species (black moulds and Fusarium sp.) were collected apart and pooled over replicates. All samples were frozen at −80°C and freeze-dried, after which they were kept in a dry place at room temperature until sample clean-up for determination of enterolignan concentration. The coleoptile, root, and seed samples were ground, weighed, and sequentially ASE extracted as the other samples of cultivated plants (see above), except that the coleoptile and root samples were not extracted with n-hexane. The solvent was evaporated to dryness using a stream of nitrogen gas at 40°C and then by freeze-drying. 5 ml of acetone was added and the samples were transferred into 6-ml test tubes. The solvent was evaporated to dryness using pressurized air at 40°C, enzyme solution was added and the samples were treated and quantified as the samples of cultivated plants. All the extracts of EL treated seeds and seedlings were diluted (root extracts 1:30 and coleoptile and seed extracts 1:5) because of the high EL concentrations.

Results

The lignan concentrations in extracts of water samples, as determined by HPLC-MS/MS, are presented in Table 1. All samples contained EL, the largest amount was found in the STP influent. The ultrapure, tap, and humic water contained equal amounts of EL. In the STP influent and effluent, the EL concentration was 25–100 times higher than in the other water sources. Also in the tap water sample analysed by GC-MS, EL could be identified (quantification was not performed), which confirmed the findings.
Table 1

Lignan concentrations (nM) in extracts of enzymatically hydrolysed water samples

Sample

HMR

MR

Sec

Lar

CLar

HEL

ED

EL

Ultrapure water

ND

0.017

ND

ND

ND

ND

ND

0.036

Tap water

0.42

0.098

ND

0.0072

0.079

ND

ND

0.044

Humic water

0.19

0.10

0.077

0.20

0.29

0.021

ND

0.041

Sea water

0.0013

ND

ND

ND

ND

0.0058

ND

0.086

STP, influent flow

0.052

ND

ND

ND

ND

0.062

0.097

4.10

STP, effluent flow

ND

ND

ND

ND

ND

0.056

0.092

2.21

ND not detected

Of the more than 50 screened tree species, EL was detected in only four species. In P. smithiana, the EL concentration was 50 μg/g of dry wood and in A. cilicia, the concentration was as high as 900 μg EL/g of dry wood (Table 2). In A. cilicia, also DHEL and MDMR were present, but in smaller amounts (100 and 400 μg/g dry wood, respectively). Furthermore, EL accounted for <0.08% of the hydrophilic extract of A. angustifolia. The predominant lignan in A. angustifolia and P. smithiana was Sec and in A. cilicia MR (4,600 μg/g dry wood). It is possible that enterolignans can be found also in other tree species although at lower levels, i.e., below the detection level of the GC-MS method. EL was indeed detected in a sample of Fagus sylvatica when HPLC-MS/MS was used, at a very low level, 0.054 μg/g of dry wood, i.e., approximately 1/1,000 of the amount found in P. smithiana and 1/17,000 of the amount in A. cilicia.
Table 2

Enterolignan content in ASE extracts of wood, cereal bran, oilseed, and nut species, μg/100 g

Plant source

EL

ED

HEL

Abies cilicia

90,000

NA

NA

Picea smithiana

5,000

NA

NA

Fagus sylvatica

5.4

ND

ND

Barley

ND

29

26

Triticale (Prego)

17

ND

0.86

Millet

0.31

5.3

0.035

Rye

ND

ND

2.6

Wild rice

1.6

ND

0.39

Quinoa

1.4

ND

0.44

Brown rice

1.4

ND

0.32

Corn

ND

ND

1.5

Japanese rice

ND

ND

1.4

Oats

0.51

0.13

0.081

Buckwheat

0.58

ND

ND

Wheat

ND

ND

0.56

Spelt wheat

0.55

ND

ND

Amaranth

0.52

ND

ND

Dhurra

0.40

ND

ND

Red rice

ND

ND

ND

Sesame seeds

5.1

ND

ND

Linseeds

0.83

ND

ND

Peanuts

638

ND

ND

Almonds

18.4

ND

ND

Cashew nuts

0.60

ND

ND

Walnuts

ND

ND

ND

ND = not detected, NA not analysed

Table 2 also shows the enterolignan levels of cultivated plants, i.e., cereal brans, oilseeds, and nuts. The plant lignan content of these plants has been presented very recently (Smeds et al. 2007). The EL levels were low compared to the levels found in softwood species, i.e., they ranged between 0.3 and 18 μg/100 g (638 μg/100 g in the peanut extract); EL could not be detected at all in six out of 22 samples. The ED level ranged between 0.13 and 29 μg/100 g (detected only in three samples). OEL, DHEL, and MDMR could not be detected in any sample.

The uptake of EL by wheat seeds and seedlings is shown in Table 3. The roots contained the highest EL (and total enterolignan) levels. Approximately 0.7% of the available EL was found in the roots. The levels of the remaining seeds and the coleoptiles were 5 and 2.5%, respectively, of the levels in the roots. EL seems to be metabolized to HEL and OEL in the plant. ED, DHEL and MDMR could not be detected in these samples.
Table 3

Enterolignan concentrations (μg/100 g) in ASE extracts of wheat coleoptiles, roots, and remaining seeds treated with purified, lignan-free water (control samples) or with lignan-free water spiked with EL during germination and growth

Sample

EL

HEL

OEL

Coleoptiles, control

ND

0.20 ± 0.02

ND

Coleoptiles, EL treated

136 ± 10.4

0.79 ± 0.10

0.06 ± 0.01

Remaining seeds, control

ND

ND

ND

Remaining seeds, EL treated

279 ± 70.6

0.76 ± 0.12

0.43 ± 0.08

Roots, control

ND

0.25 ± 0.04

ND

Roots, EL treated

5,356 ± 971

30.5 ± 2.11

19.1 ± 3.39

Five groups of 100 seedlings each, mean ± SD

ND not detected

Discussion

Plant lignans and enterolignans in water sources

Humic water contains the largest amount of plant lignans (in total) (Table 1). This is probably due to release from decaying plant material present in the water; all the plant lignans detected in the humic water are known to be present in wood (Willför et al. 2003). EL present in humic water may originate from plant lignans which are converted to EL by the action of some bacteria present in the water and/or from mammalian body excretions. EL is known to be excreted also in faeces (Adlercreutz et al. 1995b; Kurzer et al. 1995). HEL present in sea and humic water may have been formed by hydroxylation of EL, from HMR by action of bacteria present in the water, or it may originate from mammalian faeces and urine. HEL is a known metabolite of HMR (Heinonen et al. 2001; Smeds et al. 2004), and may also be a metabolite of EL.

The similar EL content of humic, tap, and ultrapure water indicates that EL is not degraded during the water treatment processes applied. Also the plant lignan content of the tap water extract was similar to the content in humic water, which may be due to the fact that humus-rich stream water is used as tap water source in Turku. The raw water consists also partly of STP effluent and of water outflow from fields, at least some of them probably fertilized with farmyard manure. Consequently, lignans seem to be largely unaffected by the chemical coagulation, activated carbon filtration, and chlorination procedures applied during the water treatment process. EL seems also unaffected by the UV irradiation in the Millipore water purification process. Previously, it has been shown that pharmaceuticals and low-molar-mass compounds are poorly removed by chemical coagulation (Vieno et al. 2006) and activated carbon filtration (Matilainen et al. 2006) during drinking water treatment.

Possible EL sources in seawater may be humic water outflow from land, production of EL from plant material present in the water, contamination with STP effluent, and/or mammalian faeces and urine.

The higher EL concentration in STP influent and effluent than in the other water sources is obvious because of the high concentrations of mammalian body excretions. The lower concentration of EL in the effluent flow than in the influent flow indicates that the compound is partly degraded in the STP, contrary to the situation in the drinking water treatment plant. This can (at least partly) be due to the usage of biological treatment in the STP.

STP influent and effluent also seem to contain ED and HEL (Table 1). Some plant lignans, e.g., Sec and Lar, which are present in foods (Milder et al. 2005), are known to be converted mainly to ED (Smeds et al. 2004). HEL has probably been metabolized from HMR, which is present in cereals, nuts, and oilseeds (Smeds et al. 2007). Very recently, the enterolignan content of aquatic environment samples was reported for the first time (Kang et al. 2006). The EL concentration was 5 ng/l (0.017 nM) in creek water and 600 ng/l (2.0 nM) in wastewater treatment plant influent; both values are about half of the concentrations we found in humic water and in STP influent, respectively (Table 1). The enterolignan concentrations in humic water are probably largely depending on the TOC content and in STP influent on dietary factors and on the number of inhabitants/total STP water flow.

Enterolignans in plant sources

The EL levels in cultivated plants (Table 2) were comparable to the amounts of some minor plant lignans like Lar sesquilignan found in the same samples (Smeds et al. 2007). Both the EL and ED levels were similar to those detected by Lee et al. (2004), who measured concentrations ranging between 1.1 and 50 μg/100 g of EL and between 0.32 and 8.0 μg/100 g of ED in Korean traditional medicinal herbs. However, the occurrence frequency of the two enterolignans was almost opposite in the study by Lee et al. compared to the present study: EL was present in 27% and ED in 45% of the samples analyzed, whereas we detected EL in 73% and ED in 14% of the analyzed samples. The EL concentration in the peanut extract was considerably higher than in the other analyzed samples of cultivated plants (Table 2). High amounts of EL in cultivated plants may be due to the use of farmyard manure as fertilizer.

There may be several possible explanations to the presence of enterolignans in plant tissues. It is possible, but not very likely, that enterolignans occur in the plants due to normal biosynthesis in these species. To the best of our knowledge, no such biosynthetic pathway has been reported for any tree or plant species (Davin and Lewis 2003; Umezawa 2003). Furthermore, no enterolignans could be detected in eight out of nine Abies species or in seven out of eight Picea species studied. Another possibility is that plants contain endophytic bacteria able to convert plant lignans into enterolignans. The most plausible explanation is, however, that enterolignans are transported from the water surrounding the roots to the wood tissue. This theory is supported by our experimental findings on EL uptake by wheat seeds and seedlings (Table 3).

The large differences in EL concentrations found in softwood species compared to those found in the cultivated plants may due to differences in the quality of the water surrounding the roots of field crops compared to the usually humus-rich water surrounding the roots of forest trees. The difference may also be due to the larger water uptake of trees, assuming that EL is accumulating in the wood tissue, or to active uptake of EL by softwood species. As EL is taken up from the water surrounding the roots, the concentration found in different plants probably varies greatly depending on the water quality or the extent of water uptake of the plant.

This study shows that different plants contain enterolignans most likely because of uptake from the water surrounding the roots. It should be noted that part of the lignans or other compounds detected in plants may not have been produced endogenously, but may have entered the plant exogenously, from the surrounding water.

Acknowledgments

Prof. Pedro Fardim at the Laboratory of Fibre and Cellulose Technology, Åbo Akademi University, Prof. Harzemşah Hafizoğlu at Bartin Faculty of Forestry, Zonguldak Karaelmas University, Turkey, and Prof. Mohammad Arfan at the Department of Chemistry, University of Peshawar, Peshawar, Pakistan are acknowledged for providing the tree samples. This work is part of the activities at the Åbo Akademi Process Chemistry Centre within the Finnish Centre of Excellence Programme by the Academy of Finland.

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© Springer-Verlag 2007