APPLIED AND ENVIRONMENTAL MICROBIOLOGY, July 2008, p. 4264–4270 Vol. 74, No. 14
Copyright © 2008, American Society for Microbiology. All Rights Reserved.
Potential Prebiotic Properties of Almond (Amygdalus communis L.) Seeds
* C. Nueno-Palop,
M. S. J. Wickham,
and A. Narbad
The Model Gut Platform, Institute of Food Research, Norwich Research Park, Colney, Norwich NR4 7UA, United Kingdom
Commensal and Microﬂora Programme, Institute of Food Research, Norwich Research Park, Colney, Norwich NR4 7UA,
; and Department of Pharmacobiology, University of Messina, Viale Annunziata, 98100 Messina, Italy
Received 28 March 2008/Accepted 15 May 2008
Almonds are known to have a number of nutritional beneﬁts, including cholesterol-lowering effects and
protection against diabetes. They are also a good source of minerals and vitamin E, associated with
promoting health and reducing the risk for chronic disease. For this study we investigated the potential
prebiotic effect of almond seeds in vitro by using mixed fecal bacterial cultures. Two almond products,
ﬁnely ground almonds (FG) and defatted ﬁnely ground almonds (DG), were subjected to a combined
model of the gastrointestinal tract which included in vitro gastric and duodenal digestion, and the
resulting fractions were subsequently used as substrates for the colonic model to assess their inﬂuence on
the composition and metabolic activity of gut bacteria populations. FG signiﬁcantly increased the popu-
lations of biﬁdobacteria and Eubacterium rectale, resulting in a higher prebiotic index (4.43) than was
found for the commercial prebiotic fructooligosaccharides (4.08) at 24 h of incubation. No signiﬁcant
differences in the proportions of gut bacteria groups were detected in response to DG. The increase in the
numbers of Eubacterium rectale during fermentation of FG correlated with increased butyrate production.
In conclusion, we have shown that the addition of FG altered the composition of gut bacteria by
stimulating the growth of biﬁdobacteria and Eubacterium rectale.
Functional foods are known as dietary components that may
cause physiological effects on the consumer, leading to justiﬁ-
able claims of health beneﬁts (36). According to the European
consensus document “Scientiﬁc concepts of functional foods,”
a food ingredient may be regarded as functional if it beneﬁ-
cially affects one or more target functions in the body, beyond
its nutritional effects, in order to improve the state of health
and/or reduce the risk of disease (9). A prebiotic is deﬁned as
“a nondigestible food ingredient which beneﬁcially affects the
host by selectively stimulating the growth and/or activity of one
or a limited number of bacteria in the colon and thus improv-
ing host health” (15). Prebiotics of proven efﬁcacy are able to
modulate the gut microbiota by stimulating indigenous bene-
ﬁcial ﬂora while inhibiting the growth of pathogenic bacteria,
such as proteolytic bacteroides and clostridia (43). Biﬁdobac-
teria and lactobacilli are able to inhibit the growth of clostridia
and pathogenic Enterobacteriaceae by the production of short-
chain fatty acids and antimicrobial compounds, as well as by
competition for growth substrate and adhesion sites (16, 19,
23). As such, these beneﬁcial gut bacteria, together with mu-
cins and antimicrobial peptides, represent part of the host’s
front line of defense against harmful microorganisms (24, 40).
In order to be effective as a prebiotic, an ingredient must
neither be hydrolyzed nor absorbed in the upper part of the
gastrointestinal tract (GIT). Although any dietary material that
enters the large intestine can be considered as potentially pre-
biotic, currently the best-known prebiotics are nondigestible
oligosaccharides (17). Different oligosaccharides with prebiotic
properties, such as inulin, fructooligosaccharides (FOS), galac-
tooligosaccharides, and lactulose, are commercially available,
but currently there is increasing interest in the identiﬁcation
and development of new prebiotic compounds, perhaps with
added functionality (26, 29, 33, 42).
The almond nut (Amygdalus communis L.) is a species of
Prunus belonging to the family Rosaceae. The global pro-
duction of almonds is around 1.7 million metric tons, with
California producing 80% of the world’s almonds. Lipid, the
main storage component in almond seeds, constituting over
50% of the total weight of the seeds, is located as intracel-
lular oil bodies (35). Proteins comprise about 22 to 25% of
the seeds, while 11 to 12% is represented by dietary ﬁber.
Recent results have shown that the encapsulation of intra-
cellular lipids by the cell walls restricts their digestion in the
stomach and small intestine (13, 27). Furthermore, if undi-
gested lipid from almond tissue reaches the large intestine,
it could be used by resident microbiota, and evidence of
bacterial fermentation was previously shown (13). Almond
cell wall material contains pectic substances that are rich in
arabinose, and the observation of their partial degradation
by the gut microbiota in the fecal samples can be explained
by the erosion of the middle lamella (10, 13). The results of
our recent study have demonstrated that the bioaccessibility
of nutrients and phytochemicals from almond seeds is im-
proved by increased residence time in the gut and is regu-
lated by almond cell walls. The results of these in vitro and
ileostomy digestibility studies have shown high amounts of
lipid and protein remaining in the almond tissue after duo-
denal digestion and therefore available for fermentation in
the colon by the gut microbiota (27).
Here we describe the investigation of the potential prebiotic
effect of almond seeds by using a full model of GIT digestion,
including gastric and small intestinal environments and a co-
* Corresponding author. Mailing address: Institute of Food Re-
search, Norwich Research Park, Colney Lane, Norwich NR4 7UA,
United Kingdom. Phone: 44 1603 251405. Fax: 44 1603 507723. E-mail:
Published ahead of print on 23 May 2008.
lonic model consisting of in vitro fermentation systems with
representative human gut bacteria.
MATERIALS AND METHODS
Almond products. Blanched ﬁnely diced and powdered almonds (Amygdalus
communis L; variety Nonpareil) were kindly provided by the Almond Board of
California. Commercial FG did not contain skin coat and had a mean particle
size of 200 m. DG was prepared by extracting 25 g of FG three times with 400
ml of n-hexane for2honeach occasion, as previously described (27).
Chemicals and enzymes. Egg
L-␣-phosphatidylcholine (PC; lecithin grade 1,
99% purity) was purchased from Lipid Products (South Nutﬁeld, Surrey, United
Kingdom). Porcine gastric mucosa pepsin (activity of 3,300 U/mg of protein
calculated by using hemoglobin as substrate), bovine ␣-chymotrypsin (activity of
40 U/mg of protein using benzoyl-L-tyrosine ethyl ester as substrate), porcine
trypsin (activity of 13,800 U/mg of protein using benzoyl-
L-arginine ethyl ester as
substrate), porcine pancreatic lipase (activity of 25,600 U/mg protein), porcine
colipase, sodium taurocholate, and sodium glycodeoxycholate were obtained
from Sigma (Poole, Dorset, United Kingdom). The lipase for the simulated
gastric phase of digestion was a gastric lipase analogue from Rhizopus oryzae
(F-AP15; activity of ⱖ150 U/mg) obtained from Amano Enzyme (Nagoya,
Japan). All other chemicals were of Analar quality.
In vitro digestion studies. The protocol previously developed to study almond
digestion under gastric and duodenal conditions (27) was used to simulate gas-
trointestinal processing for both FG and DG. Each simulated digestion was
performed at least four times, and the solid material recovered for analysis.
Control digestions of the two almond products (FG and DG) were performed in
saline solution (150 mM NaCl, pH 2.5 or 6.5 for gastric and duodenal digestion,
respectively) without enzyme additions.
In vitro gastric digestion. Phospholipid vesicles were prepared as previously
described (27). Brieﬂy, solvent was removed from 0.94 ml of PC stock solution
(63.5 mM), and the thin ﬁlm of phospholipids was then suspended in 12.2 ml of
warmed saline (150 mM NaCl, pH 2.5, at 37°C). The suspension was then
sonicated at 5°C in a coolant-jacketed vessel, using a sonication probe (Status US
200; Avestin) with a pulsed cycle of 30% full power on for 0.9 s and off for 0.1 s.
The single-shelled liposome suspension was ﬁltered through a 0.2-m nylon
syringe ﬁlter (Nalgene, United Kingdom) and equilibrated in an orbital shaking
incubator (170 rpm) at 37°C. Each almond product (1.5 g) was suspended in 12.4
ml acidic saline (150 mM NaCl, pH 2.5) in the presence of the PC vesicle
suspension, pepsin, and the gastric lipase analogue at concentrations of 2.4 mM,
146 U/ml, and 0.56 mg/ml, respectively. In vitro gastric digestion was performed
In vitro duodenal digestion. Following gastric digestion, the pH was immedi-
ately raised to 6.5 in order to simulate the duodenal conditions. Bile salt solution
(4 mM sodium taurocholate and 4 mM sodium glycodeoxycholate), CaCl
mM), and bis-Tris buffer, pH 6.5 (0.73 mM) were also added. Duodenal diges-
tions were initiated by the addition of ␣-chymotrypsin (5.9 U/ml), trypsin (104
U/ml), colipase (3.2 g/ml), and pancreatic lipase (54 U/ml) and performed in a
shaking incubator (170 rpm) at 37°C for 1 h.
Lipid content determination. Total lipid and vitamin E extraction of FG and
of FG after in vitro gastric and gastric plus duodenal digestion was performed as
previously described (27).
Total protein assays. Original almond materials (FG and DG) and solid
residues recovered after in vitro gastric and duodenal digestion were analyzed for
total nitrogen by using the micro-Kjeldahl method, as previously reported (27).
Cell wall analysis. Cell wall material was prepared from FG and FG after
gastric plus duodenal digestion by using the method previously described (27).
The alditol acetates were quantiﬁed by gas-liquid chromatography, and total
uronic acids determined colorimetrically at 580 nm (2, 3).
Fecal batch culture fermentations. Water-jacketed fermenter vessels (300
ml) were ﬁlled with 135 ml of presterilized basal growth medium (2 g/liter
peptone water, 2 g/liter yeast extract, 0.1 g/liter NaCl, 0.04 g/liter K
0.04 g/liter KH
, 0.01 g/liter MgSO
O, 0.01 g/liter CaCl
, 2 ml Tween 80, 0.02 g/liter hemin, 10 l vitamin K
g/liter cysteine HCl, 0.5 g/liter bile salts, pH 7.0) and inoculated with 15 ml of
fecal slurry. Before the addition of the fecal slurry, prepared by homogenizing
10% (wt/vol) freshly voided fecal material from one healthy donor in 0.1 M
phosphate-buffered saline (PBS), pH 7.0, the almond extract (FG or DG after
gastric and duodenal digestion) or FOS was added to give a ﬁnal concentra-
tion of 1% (wt/vol). Each vessel was magnetically stirred, the pH automati-
cally controlled and maintained at pH 6.8, and the temperature set at 37°C.
Anaerobic conditions were maintained by sparging the vessels with oxygen-
free nitrogen gas at 15 ml/min. Samples (5 ml) were removed over 24 h for the
enumeration of bacteria and short-chain fatty acid analysis. Fermentations
were run on three separate occasions.
Enumeration of bacteria. Bacteria were counted by using ﬂuorescent in situ
hybridization (FISH) (37). Duplicate fermentation samples were diluted four
times in 4% (wt/vol) ﬁltered paraformaldehyde and ﬁxed overnight at 4°C.
Samples were then washed twice with ﬁltered PBS (0.1 M, pH 7.0) and stored at
⫺20°C in PBS-ethanol (1:1, vol/vol) until further analysis. Hybridization was
performed at an appropriate temperature by using genus-speciﬁc 16S rRNA-
targeted oligonucleotide probes labeled with the ﬂuorescent dye Cy3 for the
different bacterial groups or with 4⬘,6-diamidino-2-phenylindole (DAPI) for total
cell counts. The probes used were Bif164, speciﬁc for Biﬁdobacterium (22);
Bac303, speciﬁc for bacteroides (28); Lab158, speciﬁc for Lactobacillus/Entero-
coccus spp. (18); His150, speciﬁc for most species of the Clostridium histolyticum
group (Clostridium clusters I and II) (14); and EREC482, speciﬁc for most of the
Clostridium coccoides-Eubacterium rectale group (Clostridium clusters XIVa and
XIVb) (5). The hybridized mixture was then vacuum ﬁltered using a 0.2-m
membrane ﬁlter (Millipore, Watford, United Kingdom), and the ﬁlter was
mounted on a microscope slide. At least 15 random ﬁelds were counted on each
slide by using a Nikon Microphot ﬂuorescent microscope.
Short-chain fatty acid analysis. One-milliliter samples removed from the
batch culture fermenter were centrifuged at 15,000 ⫻ g for 5 min, and 20 lof
the supernatant was injected into a high-pressure liquid chromatography system
equipped with a refractive index detector. We used an ion exclusion Aminex
HPX-87H column (7.8 by 300 mm; Bio-Rad, Watford, United Kingdom), main-
tained at 50
as eluent at a ﬂow rate of 0.6 ml/min. Quan
tiﬁcation of the organic acids was carried out by using calibration curves of acetic,
propionic, butyric, and lactic acids in concentrations between 0.5 and 100 mM,
and the results expressed in mmol/liter (37).
Statistical analysis. Differences between bacterial numbers at 0, 8, and 24 h of
fermentation for each batch culture were checked for signiﬁcance by paired t test,
assuming normal distribution and equal variances and considering both sides of
the distribution. The differences were considered signiﬁcant when the P value
Almond product characterization after in vitro digestion.
The compositions of the two almond extracts (FG and DG)
obtained after in vitro gastric plus duodenal digestion are
shown in Table 1. These fractions were subsequently used as
TABLE 1. Chemical composition of FG and DG before and after in vitro gastric plus duodenal digestion
Time of analysis
Amt of nutrient per 100 gm
Lipid (g) Vitamin E (mg) Protein (g) Dietary ﬁber (g)
Before digestion FG 54.9 ⫾ 2.5 28.2 ⫾ 1.5 25.9 ⫾ 2.6 9.8 ⫾ 1.3
DG 0 0 57.2 ⫾ 2.4 32.5 ⫾ 1.1
After digestion FG 57.4 ⫾ 3.8 34.0 ⫾ 1.8 24.7 ⫾ 1.9 13.3 ⫾ 0.9
DG 0 0 58.0 ⫾ 2.8 31.2 ⫾ 0.8
Values are the means ⫾ standard deviations of the results. Other components include carbohydrates, minerals, and vitamins.
VOL. 74, 2008 PREBIOTIC EFFECT OF ALMOND SEEDS 4265
substrates for the colonic model. As previously reported (13,
27), almond cell walls are not degraded in the upper GIT, and
therefore, the mass loss during digestion is mostly related to
loss of intracellular components, such as lipid and protein. The
simulated gastric digestion step was responsible for the major-
ity of the gravimetric losses and the highest extent of lipolysis
and proteolysis (27). FG after duodenal digestion still con-
tained 57% of the start lipid and 34 mg of vitamin E per 100 g
total almond mass, of which 96% was ␣-tocopherol, 1.3% ␤-to-
copherol, and the remainder ␥-tocopherol. The sugar compo-
sition indicated that almond cell walls are mainly composed of
arabinose-rich polysaccharides, including the pectic sub-
stances, encasing cellulose microﬁbrils. The monomeric sugar
concentrations did not change signiﬁcantly after digestion: the
sugar contents (as percentage of the total) were 39.9 and 39.5
arabinose, 12.0 and 12.6 xylose, 4.7 and 4.7 galactose, 16.7 and
16.8 glucose, and 21.1 and 20.7 galacturonic acid for FG and
for FG after gastric plus duodenal digestion, respectively. This
suggests that almond cell walls were not degraded during di-
Batch culture fermentations. Batch fermentations were used
to monitor the effects of predigested FG, DG, and FOS on the
growth of a mixed bacterial population of the human colon.
Samples were removed at intervals, and FISH was used to
quantify the levels of different bacterial groups. The results
shown in Table 2 indicate that a signiﬁcant increase in the level
of total bacteria was seen with both FG and FOS after 8 and
24 h of incubation, whereas the total bacterial number was
largely unaffected by the addition of DG. Generally, an in-
crease in the numbers of biﬁdobacteria, lactobacilli, and Eu-
bacterium rectale was observed in response to the addition of
FG and FOS at both the 8- and 24-h incubation time points.
Compared to the control vessel, the bacteroides population
with FG decreased signiﬁcantly after 24 h, whereas their num-
bers were similar to those of the control at 8 and 24 h of
incubation in the presence of DG. An increase in the number
of Eubacterium rectale was observed in the presence of both
FOS and FG, the latter showing a greater increase after 24 h of
incubation. Both fractions also stimulated the growth of bi-
ﬁdobacteria, with a 0.61 and 0.68 log increase in their numbers
at 24 h with FG and FOS, respectively. The effect of FG on
biﬁdobacteria, lactobacilli, and Eubacterium rectale numbers
was optimal after8hofincubation and did not evolve toward
the end of incubation, whereas a smaller prebiotic effect was
observed with FOS after 24 h than after 8 h. This suggests a
slower fermentation with FG, which will further the prebiotic
effect in the colon. In comparison to the results with the con-
trol, the addition of DG did not alter the bacterial numbers of
any of the groups examined. The relative changes in the num-
bers of different groups of bacteria after8hand24hof
incubation as a result of FG and FOS addition are shown in
In order to obtain a general quantitative measure of the
prebiotic effect, a prebiotic index (PI) was calculated for the
oligosaccharide fractions (31). The PI equation is described
as follows: PI ⫽ (Bif/total) ⫹ (Lac/total) ⫹ Erec/total) ⫺
(Bac/total) ⫺ (Clos/total), where Bif is biﬁdobacterial num-
bers at sample time divided by numbers at inoculation, Lac
is lactobacilli numbers at sample time divided by numbers at
inoculation, Erec is Eubacterium rectale numbers at sample
time divided by numbers at inoculation, Bac is bacteroides
numbers at sample time divided by numbers at inoculation,
Clos is clostridia numbers at sample time divided by num-
bers at inoculation, and total is total bacteria number at
TABLE 2. Changes in bacterial populations in batch cultures after 8 and 24 h of incubation
No. (log 10 cells/ml) with:
Control FOS FG DG
Total bacteria 0 9.20 ⫾ 0.03
8 9.17 ⫾ 0.04 9.36 ⫾ 0.11 9.39 ⫾ 0.05
9.23 ⫾ 0.02
24 9.19 ⫾ 0.02 9.47 ⫾ 0.01
9.40 ⫾ 0.01
9.21 ⫾ 0.01
Biﬁdobacteria 0 8.03 ⫾ 0.03
8 8.14 ⫾ 0.06 8.81 ⫾ 0.05
8.54 ⫾ 0.02
8.22 ⫾ 0.05
24 7.99 ⫾ 0.04 8.67 ⫾ 0.01
8.60 ⫾ 0.11
8.01 ⫾ 0.11
Bacteroides 0 8.23 ⫾ 0.01
8 8.30 ⫾ 0.04 8.39 ⫾ 0.01 8.31 ⫾ 0.02 8.31 ⫾ 0.06
24 8.34 ⫾ 0.12 8.42 ⫾ 0.04
8.24 ⫾ 0.01
8.38 ⫾ 0.06
Clostridia 0 7.47 ⫾ 0.09
8 7.54 ⫾ 0.07 7.18 ⫾ 0.04
7.29 ⫾ 0.07
7.51 ⫾ 0.03
24 7.54 ⫾ 0.02 7.37 ⫾ 0.04 7.52 ⫾ 0.02 7.57 ⫾ 0.07
Eubacterium rectale 0 8.28 ⫾ 0.01
8 8.33 ⫾ 0.01 8.64 ⫾ 0.13 8.97 ⫾ 0.08
8.45 ⫾ 0.04
24 8.34 ⫾ 0.08 8.64 ⫾ 0.08
8.91 ⫾ 0.02
8.46 ⫾ 0.08
Lactobacilli 0 7.58 ⫾ 0.02
8 7.72 ⫾ 0.09 8.02 ⫾ 0.01
7.73 ⫾ 0.04 7.66 ⫾ 0.05
24 7.68 ⫾ 0.07 7.90 ⫾ 0.04
7.63 ⫾ 0.01 7.66 ⫾ 0.08
Bacterial counts were obtained by using FISH. Values are the means ⫾ standard deviations of the results.
Signiﬁcantly different from control at8h(P ⬍ 0.05).
Signiﬁcantly different from control at 24 h (P ⬍ 0.05).
4266 MANDALARI ET AL. APPL.ENVIRON.MICROBIOL.
sample time divided by numbers at inoculation. The PI
represents a comparative relationship between the growth
of “beneﬁcial” bacteria, such as biﬁdobacteria, lactobacilli,
and Eubacterium rectale, and that of the “less desirable”
ones, such as clostridia and bacteroides, in relation to the
change in the total number of bacteria (Fig. 2). For both
substrates, the PI values obtained at8hofincubation were
higher than those at 24 h. The FOS fraction produced the
highest PI value after8hofincubation, 6.36, with that of FG
being 4.98, whereas FG produced the highest PI value at the
24-h time point, 4.43, with that of FOS being 3.49. Low PI
values were obtained with DG and the control at both the 8-
and 24-h incubation time points.
Short-chain fatty acid production during fermentation. The
concentrations of lactic, acetic, propionic, and butyric acids
produced during fermentation are shown in Table 3. FOS gave
the highest total short-chain fatty acid production at all time
points tested. However, butyrate production signiﬁcantly in-
creased after8hofincubation with FG and peaked at 24 h,
coinciding with the highest number of Eubacterium rectale bac-
teria. Fermentation with FOS resulted in the highest produc-
tion of lactic and acetic acids: their concentrations increased at
4 h and remained elevated up to 24 h. These increases corre-
lated with changes in the numbers of biﬁdobacteria and lacto-
bacilli. The concentrations of propionic and butyric acids were
higher after 8 and 24 h of fermentation with FG and DG, again
correlating with Eubacterium rectale population changes. In the
absence of the added carbon source, an increase in acetic acid
was observed after 24 h, although the amounts of the other
organic acids did not change signiﬁcantly.
FIG. 1. Differences in the bacterial population sizes (black bars, FOS; white bars, predigested FG) compared to the total numbers of bacteria
counted at 8 h [(selected bacterial numbers at 8 h/total bacteria counted at 8 h) ⫺ (selected bacterial numbers at 0 h/total bacteria counted at 0 h)]
(A) and 24 h [(selected bacterial numbers at 24 h/total bacteria counted at 24 h) ⫺ (selected bacterial numbers at 0 h/total bacteria counted at
0 h)] (B). Error bars show standard deviations.
FIG. 2. Prebiotic index (PI) scores from batch cultures at 8 h (black
bars) and 24 h (white bars) using FOS, predigested FG, DG, and
untreated (control) cultures. Error bars show standard deviations.
VOL. 74, 2008 PREBIOTIC EFFECT OF ALMOND SEEDS 4267
In the present study, we have demonstrated the prebiotic
potential of almond seeds. As far as we are aware, this is the
ﬁrst study that has used combined models of human digestion
which include gastric and duodenal digestion followed by co-
lonic fermentation to study the effects of almond extracts on
the modulation of gut microbiota. Commercially available pre-
biotics (such as FOS and galactooligosaccharides) are not sen-
sitive to gastric acid and do not serve as substrate for hydrolytic
enzymes in the upper digestive tract. The evaluation of novel
prebiotic compounds should take into account the available
ingredients which can be digested by human enzymes and
adsorbed, thus entering into intermediary metabolism. On the
contrary, food components able to reach the colon can poten-
tially provide the body with additional energy via microbial
fermentation, and the production of short-chain fatty acids
may have potential prebiotic functionality.
In our previous study, we showed that relatively small
amounts of almond lipids and proteins are bioavailable during
gastric and small intestinal digestion and that nutrient encap-
sulation by cell walls is likely to prevent their digestion in the
upper GIT (27). However, bioaccessibility is improved when
the number of fractured cells is increased by processing or by
increased residence time in the gut. Evidence of bacterial fer-
mentation of the almond tissue was provided by micrographs
of fecal samples collected from healthy subjects consuming an
almond-rich diet (13). By using transmission electron micros-
copy, it was possible to document the presence of bacteria both
on the cell wall surface and within the cells. Therefore, almond
intracellular lipids, together with nonstarch polysaccharides
from cell walls, which are known to be metabolized to a vari-
able degree in the large intestine, could represent a suitable
carbon source for bacterial fermentation (12, 13). The erosion
of the middle lamella observed in fecal samples was considered
to be further evidence of pectic degradation by gut microbiota.
Our data presented in this study show that when almond lipids
are available for fermentation in the large bowel, modulation
of the composition of the bacterial population is observed, with
a signiﬁcant increase in the numbers of biﬁdobacteria and
Eubacterium rectale (Fig. 1). However, when the almond lipid
source was removed (defatted almond product) no signiﬁcant
changes in the bacterial population were detected (Table 1).
These results suggest that the lipid component of almond seeds
is relevant in the alteration of bacterial growth and metabo-
The unique role of dietary ﬁber, essentially the plant cell
wall, has been evaluated in many physiological processes and in
disease prevention (7). Pectins with different degrees of ester-
iﬁcation have previously been shown to increase Eubacterium
rectale numbers, and this group of gut bacteria is known to
produce relatively large amounts of butyrate (1, 11). The anti-
inﬂammatory (4) and antineoplastic (25, 39) properties of bu-
tyrate have been demonstrated on cell tissue cultures in vitro,
increasing the interest in the potential effect of butyrate in
inﬂammatory bowel disease and colorectal cancer. However,
butyrate production from oligofructose fermentation was
shown to be mainly derived from interconversion of extracel-
lular lactate and acetate (30). In the present study, FG pro-
duced more butyrate than FOS and can therefore be consid-
ered a butyrogenic prebiotic.
The kinetics of almond lipid digestion and absorption is an
important factor for postprandial lipemia and has implications
for the regulation of body weight (41). The results of a dose-
response study have shown that almond consumption im-
TABLE 3. Concentrations at 0, 4, 8, and 24 h of short-chain fatty acids and lactate produced during fermentation
Control FOS FG DG
Total fatty acids 0 4.19 5.58 4.10 4.22
4 7.44 40.35 14.44 11.29
8 14.80 74.02 39.13 33.21
24 24.20 91.23 62.36 61.36
Lactic acid 0 0.53 ⫾ 0.03 0.54 ⫾ 0.05 0.25 ⫾ 0.02 0.22 ⫾ 0.05
4 0.51 ⫾ 0.29 12.23 ⫾ 0.36 4.96 ⫾ 0.03 1.18 ⫾ 0.20
8 0.68 ⫾ 0.21 16.58 ⫾ 1.36 4.79 ⫾ 0.27 0.45 ⫾ 0.38
24 0.69 ⫾ 0.25 16.91 ⫾ 2.02 7.18 ⫾ 0.44 1.58 ⫾ 0.17
Acetic acid 0 1.42 ⫾ 0.22 2.15 ⫾ 0.28 1.24 ⫾ 0.32 1.16 ⫾ 0.28
4 3.38 ⫾ 1.03 21.97 ⫾ 0.36 5.25 ⫾ 1.06 7.00 ⫾ 0.88
8 8.81 ⫾ 1.89 46.15 ⫾ 1.21 14.09 ⫾ 0.16 20.77 ⫾ 1.72
24 15.05 ⫾ 2.43 50.77 ⫾ 3.91 26.88 ⫾ 0.01 34.79 ⫾ 2.90
Propionic acid 0 0.90 ⫾ 0.21 1.05 ⫾ 0.16 1.27 ⫾ 0.04 1.21 ⫾ 0.06
4 1.80 ⫾ 0.56 2.67 ⫾ 0.36 2.44 ⫾ 1.06 1.04 ⫾ 0.06
8 2.42 ⫾ 0.19 4.00 ⫾ 0.62 8.09 ⫾ 1.31 7.06 ⫾ 1.60
24 3.90 ⫾ 0.70 11.11 ⫾ 2.27 12.09 ⫾ 2.23 14.77 ⫾ 0.70
Butyric acid 0 1.34 ⫾ 0.04 1.85 ⫾ 0.21 1.35 ⫾ 0.04 1.63 ⫾ 0.08
4 1.75 ⫾ 0.66 3.47 ⫾ 0.36 1.79 ⫾ 0.54 2.07 ⫾ 0.09
8 2.89 ⫾ 0.25 7.29 ⫾ 0.71 12.16 ⫾ 0.33 4.93 ⫾ 0.24
24 4.56 ⫾ 0.17 12.44 ⫾ 4.73 16.21 ⫾ 0.87 10.22 ⫾ 1.07
Values for individual acids are means ⫾ standard deviations of the results.
4268 MANDALARI ET AL. APPL.ENVIRON.MICROBIOL.
proved the serum proﬁle of healthy and middle-hypercholes-
terolemic adults, thus reducing risk factors for coronary heart
disease (38). The predominant fatty acid of almond triacyl-
glycerols is oleic acid, comprising more than 65% of the total
oil fraction and contributing to the high monounsaturated fat
content of almonds. Prebiotics have also been reported to
indirectly lead to a reduction in serum triglyceride levels (44),
and short-chain fatty acid production can modulate the expres-
sion of multiple genes involved in the atherosclerosis process
(32). An in vivo murine study investigating the effects of pre-
biotics on atherosclerotic plaques showed that inulin and FOS
were able to reduce plasma and hepatic cholesterol (34). In the
present study, we have shown that almond extracts can act as
prebiotics, and this may add functionality to almonds in im-
proving serum cholesterol levels.
In this study, we have not investigated whether oleic acid,
which is present in the FG fraction, was utilized by the gut
bacteria. However, there is evidence to indicate that oleic acid
may be metabolized by the ruminal bacterium Selenomonas
ruminantium and strains of Streptococcus, Enterococcus, and
Lactobacillus (20, 21). Linoleic acid, another unsaturated fatty
acid, was also metabolized in the human colon by a number of
Roseburia species (8). In addition, by being incorporated into
the bacterial membrane of speciﬁc gut bacteria, oleic acid may
contribute to their increased survival rate in gastric juice (6).
In conclusion, we have shown that almond seeds exhibited
the potential to be used as a novel source of prebiotics, in-
creasing the populations of biﬁdobacteria and Eubacterium
rectale with the subsequent increase in butyrate concentrations.
More-detailed studies on the digestibility of almonds and the
role played by lipids in the potential prebiotic effect need to be
performed using human volunteers.
We gratefully acknowledge the help provided by Yvan Lemarc
(IFR) with statistic analyses. We thank Karen Lapsley (ABC) for
providing the almond products and for useful discussions.
This research was funded by the Almond Board of California
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