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2
Laboratoire de Chimie Biologique, INRA, Institut National Agronomique Paris-Grignon, 78850 Thiverval-Grignon, France;
*
Unité d’Ecologie et de Physiologie du Système Digestif, INRA, 78352 Jouy-en-Josas cedex, France; and
Laboratoire des Maladies Métaboliques et Micronutriments, INRA, 63122 Saint-Genès-Champanelle, France
2To whom correspondence should be addressed.
 |
ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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KEY WORDS: • proanthocyanidins • flavonoids • bioavailability • biodegradation • human colonic microflora
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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,
Shahidi and Naczk 1995
, Swain 1962
). PA
differ from all other natural polyphenols by their polymeric nature.
They are made of flavan-3-ol units and their average degree of
polymerization generally varies between 3 and 11 (Fig. 1
) (Santos-Buelga and Scalbert 2000
). Polymerization
degree may reach values as high as 17 as was shown in an apple cider
extract by liquid chromatographic-mass spectrometric (LC-MS)
analysis (Guyot et al. 1997
). Two main types of PA can
be distinguished according to the substitution pattern of their
B-ring, i.e., procyanidins have two vicinal hydroxy groups and
prodelphinidins three. The phenolic and polymeric nature of PA makes
them good complexants of proteins and explains the astringent character
of foods and beverages rich in PA (e.g., grape skin and seeds, unripe
fruits, wine or cider) (Haslam 1998
). The daily intake
of PA has been estimated at 0.1–0.5 g (Kühnau 1976
, Santos-Buelga and Scalbert 2000
,
Scalbert and Williamson, 2000
). However, the complexity
of their chemical structure and the lack of a suitable method for their
estimation make these figures uncertain.
|
]. These effects suggest that some PA are
absorbed through the gut barrier. Their absorption depends on their
degree of polymerization. In some in vitro experiments, only PA dimers
and trimers, but not polymers with an average polymerization degree of
7, were absorbed through an intestinal epithelium cell monolayer
(Déprez 1999
). PA polymers were not better
absorbed in vivo in chicken and sheep (Jimenez-Ramsey et al. 1994
, Terrill et al. 1994
).
These PA polymers could be degraded by the colonic microflora into
low-molecular-weight compounds, which would be subsequently absorbed.
This has been demonstrated for other dietary polyphenols such as
isoflavones by comparing normal and germfree animals or humans before
and after antibiotic administration (Axelson and Setchell 1981
, Axelson et al. 1982
, Setchell et al. 1981
). A procyanidin dimer (dimer B3) (Groenewoud and Hundt 1986
) and (+)-catechin (Groenewoud and Hundt 1984
, Scheline 1991
) were shown to be degraded
into phenolic acids and nonphenolic aromatic metabolites. Some of these
metabolites were also found in the urine of rats fed a grape extract
containing a mixture of catechin monomers and PA oligomers of low
polymerization degree (Harmand and Blanquet 1978
). No
study of this kind has been carried out to date with human microflora
and high polymerization degree PA, the most abundant in the human diet.
We report here the in vitro degradation of
14C-labeled PA polymers by human microflora.
Labeled PA polymers were obtained by incorporation of a radiolabeled
precursor to willow tree shoots. They were carefully purified and were
free of any PA dimer and trimer or any other phenolic or nonphenolic
contaminants (Déprez et al. 1999
). PA in willow
tree leaves are procyanidins, typical of the most common PA in food
(Santos-Buelga and Scalbert 2000
). The radiolabeled PA
had an average polymerization degree of 6, close to that of other PA
found in apples (Guyot et al. 1998
) or grape seeds
(Prieur et al. 1994
).
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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5-(m-Hydroxyphenyl)valeric and 5-(m,p-dihydroxyphenyl)valeric acids were kindly provided by Ronald Scheline (University of Bergen, Norway). 3-(m-Hydroxyphenyl)propionic acid was a gift of Gerhard Groenewoud (Farmovs Research Center, Bloemfontein, South Africa). Benzoic acid was purchased from Prolabo (Nogent-sur-Marne, France) and all other aromatic acids from Sigma-Aldrich (Saint-Quentin Fallavier, France). Total radioactivity of samples was determined by liquid scintillation counting of aliquots (10 µL). Liquid scintillation radioactivity counting was controlled with a Packard Tri-Carb 1500 analyzer (Meriden, CT) (liquid scintillation cocktail: Ecolite, ICN Pharmaceuticals France, Orsay, France); counting efficiency was 95%.
Nonlabeled and 14C-labeled proanthocyanidin polymers.
PA polymers were purified from the leaves of an adult willow tree
(Salix caprea L.) by chromatography on a Sephadex LH
20 (Pharmacia, Guyan-Court, France) (Déprez et al. 1999
). They were free of any catechin monomers or PA dimer and
trimer. 13C nuclear magnetic resonance spectra indicated a
procyanidin-type polymer, with a predominance of
2,3-trans (catechin) units. Polymers were analyzed by
thiolysis (Fig. 1
(Déprez 1999
, Matthews et al. 1997
)
and gave two main thioadducts, 3,4-cis- and
3,4-trans-benzylthiocatechins, formed from inner catechin
units (yield 45.0%), low amounts of
3,4-trans-benzylthioepicatechin derived from inner
epicatechin units (yield 1.4%) and (+)-catechin derived from terminal
units (yield 7.7%). The ratio of thioethers and (+)-catechin allowed
the calculation of an average polymerization degree of 7.
14C-PA polymers were prepared by administration
of [1-14C]-acetate to willow shoots and similar
purification by chromatography on Sephadex LH 20 (Déprez et al. 1999
). Specific activity was 7.0 MBq/g. The yield of
thioadducts formed by thiolysis was 42.2%. Thioadducts
3,4-cis- and 3,4-trans-benzylenzylthiocatechins
represented 95% of the thioethers. An average polymerization degree of
6 was calculated. All other characteristics were similar to those given
above for nonlabeled PA.
Degradation of proanthocyanidin polymers by the microflora.
The brain heart infusion (BHI) medium was made of calf brain and beef heart infusion (37 g/L; Difco, Becton Dickinson France SA, Le Pont de Claix, France), hemine (5 mg/L; Sigma-Aldrich) and yeast extract (5 g/L; Difco). The pH was adjusted to 7.4 with NaOH and the medium sterilized (120°C, 20 min). It was placed in anaerobic conditions (5% CO2, 10% H2, 85% N2) 48 h before fermentation experiments. Fecal suspensions were prepared by mixing fecal samples (1 g) (freshly collected from a human subject habitually consuming a Western type diet) in BHI medium (100 mL).
Aqueous solutions of nonlabeled PA polymers (100 mmol/L expressed as catechin unit equivalents, i.e., 29 g/L) were sterilized by filtration on Millex-GS sterile units (25 mm, 0.22 µm, Millipore, Saint-Quentin en Yvelines, France); 150 µL was added to the fecal suspensions (2.85 mL; final PA concentration, 5 mmol/L) under anaerobic conditions (Freter glove box) at 37°C without shaking. Aliquots (500 µL) were sampled at 0, 6, 12, 24 and 48 h. All experiments were carried out in duplicate.
Similar experiments were carried out 1 mo later with aqueous solutions of radiolabeled PA polymers (50 mmol/L expressed as catechin unit equivalents). These solutions (200 µL) were added to fecal suspensions (1.80 mL; final concentration of polymers, 5 mmol/L; 18 kBq/flask). Samples (300 µL) were taken at 0, 6, 12, 24, 36 and 48 h. Four controls were performed, i.e., polymers incubated in water or in BHI medium without flora; flora in BHI medium without PA; flora heat-inactivated (120°C, 20 min) and in BHI medium with PA. All experiments were conducted in duplicate.
Samples were stored at -20°C before analysis. The raw extracts were
centrifuged for 5 min at room temperature (15,000 x g) before analysis by cellulose TLC. Aliquots
(40 µL) were also acidified with HCl (6 mol/L)
saturated with NaCl (final pH 
1), extracted three times by ethyl
acetate and concentrated under vacuum before silica TLC and gas
chromatography coupled to mass spectrometry (GC-MS) analysis. Due to
the identification of an arylsulfotransferase enzyme in a bacterial
strain isolated from human feces (Koizumi et al. 1990
and 1991
), some fermentation samples (48 h) were treated with a
sulfatase of Patella vulgata (Sigma-Aldrich; 5 h at
37°C, 0.5 mg; 10 U/mg) before extraction by ethyl acetate
(EtOAc); however, no change in GC chromatograms was observed.
Analysis of PA by thiolysis.
Thiolysis was performed on the raw fermentation samples according to a
procedure established on plant extracts (Matthews et al. 1997
) except that the amount of added acetic acid (AcOH) was
increased to take into account the buffering capacity of the growth
medium. Toluene-
-thiol (90 µL), AcOH (300
µL) and the fermentation sample (200
µL) were added to ethanol/water 5:1 (2.5 mL) and the
mixture acidified to pH 3.8 by further addition of AcOH, purged with
nitrogen. The tubes were tightly closed (Teflon-lined screw cap), and
the thiolysis reaction was conducted at 105°C in an oil bath over a
period of 24 h under stirring. Samples were cooled and centrifuged
(2 min, 20°C, 11,300 x g) and the supernatant
dried under reduced pressure. Thiolysis products were solubilized in
methanol (200 µL) and the solution filtered on
Millex-GV units (Millipore, 4 mm, 0.22 µm) before
HPLC analysis (Matthews et al. 1997
). The
chromatographic conditions were slightly modified to avoid an overlap
between one of the thioadducts and a compound present in the samples
but not derived from the PA polymers. Analyses were carried out on a
Lichrospher 100 RP-18 column (5 µm, 25 cm x 4 mm i.d.; Merck,
Darmstadt, Germany) with the following elution conditions: solvent A,
H3PO4 0.1% in water; solvent B, methanol;
linear gradient: 20–70% B in 30 min; flow rate, 1 mL/min; detection
was performed at 280 nm. Yields are expressed as the percentage of the
products obtained by thiolysis of the nonfermented PA polymer.
TLC analysis.
Raw extracts were analyzed on high performance cellulose thin layer
plates (aluminum foils, thin layer thickness 0.1 mm, Merck) with
AcOH/water 6:94 (v/v) as eluent. EtOAc extracts were spotted on silica
plates (aluminum foil, 5 x 10 cm, 0.25 mm, Merck) and eluted
with CHCl3/AcOH/water 4:1:1 (v/v/v) on a length of 5–8 cm
(Griffiths and Smith 1972
). Electronic autoradiographies
of thin layer chromatograms were obtained with a Storm 860 (Molecular
Dynamics, Amersham Pharmacia Biotech, Orsay, France) electronic system.
Dried cellulose plates were imaged for 3–24 h. Calibration was
performed with a standard solution of
[U-14C]-phenylalanine.
GC-MS analysis.
EtOAc extracts were dried over Na2SO4 and
aliquots (10 µL) silylated with 50 µL
bis-trimethylsilyltrifluoroacetamide and 5
µL pyridine before GC-MS analysis. Silylated
compounds were analyzed with a Varian Star 3400 chromatograph (Les
Ulis, France), on a SPB5 polydimethylsiloxane capillary column (30 m
x 0.2 mm i.d., 0.25 µm, Supelco, Sigma-Aldrich)
with helium as carrier gas (0.5 bar inlet pressure). The column
temperature was raised by 30°C/min from 40 to 110°C, then by
2.5°C/min from 110 to 280°C. MS of trimethylsilylated derivatives
(70 eV electronic impact, 50–650 m/z range, positive
mode) was performed with a Saturn 2000 ion trap instrument (Varian).
Compounds were identified by comparison of their MS spectra to those of
reference compounds. Standards of phenolic acids hydroxylated in the
ortho, meta or para
position or nonhydroxylated were well separated by GC; the elution
order was as follows: nonhydroxylated < ortho
< meta < para as previously
reported (Groenewoud and Hundt 1986
). Positional isomers
all gave different mass spectra by GC-MS. The relative abundance of
the phenolic acids was estimated from the relative surface area of
their trimethylsilylated derivatives on the total ion chromatogram.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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(Matthews et al. 1997
).
A major drop of thiolysis yield was observed after 48 h of
incubation (Fig. 2
). This drop was not observed in the presence of a heat-inactivated
flora or in the absence of flora.
|
). These products represented 9–22% of the initial PA radioactivity
(according to the sample and duration of incubation). They were not
observed in killed microflora controls. The total radioactivity
measured in the liquid medium after incubation was close to the initial
radioactivity introduced into the medium. The formation of gaseous
metabolites such as CO2 and
CH4 was therefore negligible.
|
). After 48 h of incubation, these EtOAc-soluble metabolites
represented 2.7% of the initial radioactivity. Their chemical
structures were determined by GC-MS analysis. Six phenolic acids
[2(p-hydroxyphenyl)acetic acid (pHPAc),
2-(p-hydroxyphenyl)- propionic acid (pHPP) and
their m-hydroxy isomers 2-(m-hydroxyphenyl)acetic
acid (mHPAc) and 2-(m-hydroxyphenyl)propionic
acid (mHPP), 5-(m-hydroxyphenyl)valeric acid
(mHPV) and phenylpropionic acid (PP)] were specifically
observed after incubation of PA polymers with the living flora and were
absent in all controls (Figs. 5
and 6
). No dihydroxylated phenolic acid could be detected. The nonphenolic
aromatic compounds phenylacetic acid (PAc) and benzoic acid (BA) were
also identified. However, they were not specific for PA and could also
derive from the metabolism of other compounds such as protein aromatic
acids present in the brain heart growth medium or could have been
present initially in the fecal sample. Co-chromatography with
authentic samples allowed us to identify the metabolites M2 and M3 as
mHPP and mHPV, respectively. Radioactive
compounds eluted in the solvent front (M4) are likely nonhydroxylated
aromatic acids as suggested by the similar chromatographic behavior of
BA, PAc and PP.
|
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|
). pHPAc and pHPP acids detected after 6
h-incubations disappeared progressively after a longer period of
incubation. The proportions of their m-isomers and of PP
increased in the first 10 h of incubation and then remained
stable. mPP was the most abundant aromatic acid in all
samples whatever the time of incubation. Some differences in the
kinetics of accumulation of mHPP were observed in different
experiments. In the first experiment, mHPP (Fig. 4
,
metabolite M2) disappeared after 36 h of incubation, whereas its
concentration was found to be stable when the experiment was repeated 1
mo later. These differences may be explained by variations in the
composition and activity of the microflora induced by diet
modifications (Cummings et al. 1989
).
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|
DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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), can be
degraded by the colonic microflora into low-molecular-weight aromatic
acids, which differ according to their hydroxylation profile and the
length of the aliphatic side chain. Three types of hydroxylation
pattern were observed for the phenolic acids. Phenolic acids with a
m-hydroxyphenyl group and the nonhydroxylated aromatic acids
both reached stable values after 24 h of incubation. In contrast,
phenolic acids with a p-hydroxyphenyl group rose up to a
maximum after 6 h of incubation to decrease to near-zero
values after 24–48 h.
p-Hydroxy–substituted phenolic acids are likely produced by
dehydroxylation in the 3-position. They may be further rearranged into
their m-isomers by migration of either the hydroxyl group or
the aliphatic side chain or dehydroxylated into their nonphenolic
analogs as suggested by microflora degradation experiments of
deuterated pHPP (Curtius et al. 1976
).
However, the major route leading to the
m-hydroxy–substituted phenolic acids is likely to occur
through the selective dehydroxylation in the 4-position of
3,4-dihydroxylated phenolic acid intermediates as previously shown in
catechin metabolism studies by human and rat microflora (Meselhy et al. 1997
). Nonhydroxylated phenolic acids are formed by
dehydroxylation of the monohydroxylated phenolic acids. Only PP could
be identified as a metabolite of PA polymers. PAc and BA are also
likely formed because they are known metabolites of some aromatic acids
identified here (Curtius et al. 1976
, Drasar and Hill 1974
, Scheline 1991
). No definitive
evidence of their formation could be obtained because of the
simultaneous presence of other precursors in the growth medium.
Metabolites also differ by the length of the aliphatic side chain.
Phenylvaleric acids are transformed into propionic acids and benzoic
acids by a progressive loss of carbon atoms through ß-oxidation
(Das and Griffiths 1968
, Harmand and Blanquet 1978
, Meselhy et al. 1997
, Scheline 1991
). -Oxidation, which would explain the formation of
mHPAc from mHPP (Curtius et al. 1976
), may also occur.
All aromatic acids presently identified as metabolites of PA polymers
are similar to those produced by colonic microflora metabolism of
either (+)-catechin (Meselhy et al. 1997
) or procyanidin
dimer B3 (Groenewoud and Hundt 1986
). In agreement with
the present work, m-HPP was the main metabolite produced
from tea catechins by human or rat microflora (Meselhy et al. 1997
) and also the main phenolic acid found in urine of
volunteers who had ingested (+)-catechin (Das 1971
).
Other known metabolites of (+)-catechin and dimer B3 such as
phenylvalerolactones (Groenewoud and Hundt 1986
,
Meselhy et al. 1997
, Scheline 1991
) and
diarylpropan-2-ol metabolites (Groenewoud and Hundt 1984
and 1986
, Meselhy et al. 1997
, Scheline 1991
) could not be detected here, possibly because the duration
of exposure to the microflora was too long. (+)-Catechin was not
identified as an intermediate in contrast to previous results obtained
with a rat microflora and the dimer B3 (Groenewoud and Hundt 1986
). This contradiction may arise from the difference in the
nature of the microflora in both rats and humans as suggested by the
incapacity of the rat microflora to hydrolyze galloyl esters in tea
catechins (Meselhy et al. 1997
). It could also be
explained by diet-mediated changes in the microflora.
The results obtained by in vitro anaerobic degradation by an isolated
colonic microflora can be compared with animal and human in vivo data.
Aromatic acid metabolites are formed exclusively by the colonic
microflora and not by animal tissues as indicated by the comparison of
the metabolites formed after oral and intraperitoneal administration of
(+)-catechin and the suppression of their formation after
administration of antibiotics (Das and Griffiths 1968
,
Griffiths 1964
). The total yield of such aromatic acids
recovered in 24-h urine from (+)-catechin or a mixture of (+)-catechin
and PA oligomers fed to either rats or humans varies between 2 and 4%
(Das and Griffiths 1969
, Das 1971
,
Harmand and Blanquet 1978
). These figures are close to
those obtained after a 48-h in vitro degradation by the microflora (see
above the yields of ethylacetate-soluble metabolites). However, the
similarity of these figures may be fortuitous because the amounts of
aromatic acids excreted in urine depend not only on the catabolism of
polyphenols in the colon but also on the amounts of substrate reaching
the colon (which decrease with its absorption in the small intestine)
and on the absorption of the products through the colon barrier. The
absorption of aromatic acids may be quite high after oral
administration. BA (Bridges et al. 1970
) or PAc
(James and Smith 1973
) are absorbed very efficiently
when ingested by humans, with 100% of the dose recovered in urine. The
absorption of hydroxylated aromatic acids may be lower. When
mHPP was administered orally to guinea pigs, 5% of the dose
was recovered in urine as unchanged mHPP, mHBA or
m-hydroxyhippuric acid (Das and Griffiths 1968
). However, these data do not give any indication on their
specific absorption at the colon level.
These results also suggest the formation of metabolites different from the aromatic acids described above. Indeed, radioactivity of EtOAc-soluble metabolites formed from PA polymers accounted for only 2.7% of the initial activity of PA. Their formation cannot compensate for the quasi-total disappearance of PA-derived thiolysis products. Some of the metabolites may still have a polymeric structure but may be modified in a way that would explain their loss of reactivity toward the thiolysis reagents. Some of the phenolic compounds may have been converted into microbial biomass and would therefore have escaped detection. A more detailed scrutiny of the distribution of the radioactivity in different soluble and insoluble fractions of the raw incubation media should give more insight into the nature of these metabolites.
PA polymers, which were usually considered to be relatively inert in
the digestive tract and recovered unchanged in the feces, appear to be
as easily degraded as other flavonoid monomers. More interest has been
paid to their effects on microorganisms rather than to the effects of
the microorganisms on PA polymers (Scalbert 1991
). The
only previous report clearly showing a chemical biodegradation of PA
polymers was published more than 25 years ago and concerned fungi
(Grant 1976
). To understand the nutritional properties
of dietary PA, it will be necessary to study not only their biological
properties as such, but also those of their degradation products.
Anticancer properties of aromatic acids such as PAc are well documented
(Samid et al. 1997
, Thibout et al. 1999
).
Such degradation products may be equally important as has been realized
for fibers and their fatty acid metabolites.
|
FOOTNOTES |
|---|
3 Abbreviations used: AcOH, acetic acid; BA, benzoic acid; BHI, brain heart infusion; EtOAc, ethyl acetate; GC-MS, gas chromatography coupled to mass spectrometry; mHPAc, 2-(m-hydroxyphenyl)acetic acid; mHPP, 2-(m-hydroxyphenyl)propionic acid; mHPV, 5-(m-hydroxyphenyl)valeric acid; PA, proanthocyanidin; PAc, phenylacetic acid; pHPAc, 2-(p-hydroxyphenyl)acetic acid, pHPP, 2-(p-hydroxyphenyl)propionic acid; PP, phenylpropionic acid.
Manuscript received January 19, 2000. Initial review completed March 13, 2000. Revision accepted June 8, 2000.
|
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