![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
|
|
|
|
|
||
*
Department of Animal Nutrition and Physiology and
Department of Agricultural Systems, Danish Institute of Agricultural Sciences, Research Center Foulum, DK-8830 Tjele, Denmark
4To whom correspondence should be addressed.
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
KEY WORDS: • sweetener • sugar • fermentation • adaptation • pigs • D-tagatose
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
). However, the
metabolism of D-tagatose is unclear. Earlier results in
rats have shown that 43% of intravenously administered carbon of
14C-labeled D-tagatose was excreted
in urine and 37% in expirate (Levin et al. 1995
,
Saunders et al. 1999
). However, when administered
orally, only 5–6% was excreted in the urine, 49–68% was excreted in
expiration and 11–29% with feces. Gastrointestinal adaptation to
D-tagatose seems to be important for metabolism because the
extent of urinary and fecal excretion was dependent on whether the rats
were adapted to D-tagatose in the diet. We previously found
that <26% of D-tagatose was absorbed in the proximal
two-thirds of the small intestine
(SI)5
of pigs fed a D-tagatose–containing diet for 20 d
(Lærke and Jensen 1999
). However, no
D-tagatose could be detected in feces, which suggests that
the malabsorbed D-tagatose was fully fermented in the large
intestine. Thus to assess the energy value and compliance of diets
containing D-tagatose, it is of great interest to study the
capability of unadapted and adapted gut microbiota to degrade
D-tagatose. The aims of this investigation were as follows: 1) to study the influence of D-tagatose on the short chain fatty acid (SCFA) and lactic acid production in the gastrointestinal tract determined by in vitro incubations of gut contents from control pigs or pigs adapted to D-tagatose, 2) to estimate the degradation of D-tagatose and formation of fermentation products from D-tagatose during in vitro incubation of D-tagatose with slurries of bacteria harvested from various regions of the gastrointestinal tract and 3) to investigate the population size of D-tagatose–degrading bacteria in feces from control pigs or pigs adapted to ingestion of D-tagatose.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
At 7-d intervals, two groups of eight (2 x 4 littermates and 4
x 2 littermates) 62- to 75-d-old castrated male Landrace x
Yorkshire pigs, obtained from the swine herd at the Danish Institute of
Agricultural Sciences, Foulum, Denmark, were given a low-fiber
basal diet containing either 150 g/kg sucrose (control diet) or 50 g/kg
sucrose + 100 g/kg of D-tagatose (tagatose diet) for
18 d. The pigs had an adaptation period of 2 d during which
they were fed a traditional Danish pig’s diet (d -4 and d
-3), followed by 2 d on a 1:1 mixture of the standard
feed and the experimental diet (d -2 and d -1). Subsequently, the
pigs were fed the experimental diet for 18 d (d 0 to d 17). The
pigs were fed twice a day at 0700 and 1500 h. The feed intake was
restricted to a total amount of 40 g/(kg body weight · d). For
details about the experimental diets see Lærke and Jensen (1999)
. The protocol used in this experiment complied with the
Danish Ministry of Justice, law no. 726 (December 9, 1993) concerning
animal experimentation and care of experimental animals.
Sampling during the experiment.
Fecal samples were taken directly from the anus in the morning when the pigs were fed the standard diet (d -3) and after 1, 8 and 15 d of consuming the experimental diets for determination of total anaerobic bacteria, D-tagatose–degrading bacteria, dry matter and pH.
Collection of samples after slaughter.
On d 17, the pigs were killed 3 h after the morning meal with a
lethal injection of pentobarbital sodium (200 g/L). Immediately after
slaughter the gastrointestinal tract was removed and divided into
segments by ligatures as previously described (Lærke and Jensen 1999
). Samples from the stomach, the distal third of the small
intestine (distal SI), the cecum and the mid third of the colon (mid
colon) were used for in vitro incubations.
In vitro fermentation of unfractionated gut contents.
The rate of production of SCFA and lactic acid in the gastrointestinal
tract of pigs on the two diets was determined by incubation of a 20
g/100 g slurry of gastrointestinal contents from the stomach, distal
SI, cecum or mid colon in 100 mmol/L Na-phosphate buffer (pH 6.5).
The incubations were essentially carried out as described by
Jensen and Jensen (1993)
, with an incubation time of
2 h in serum bottles. Samples were taken for analysis after 0, 15,
30, 60 and 120 min.
In the contents of the stomach and the distal SI, a linear increase in lactic and acetic acid production was detected up to 1 h of incubation, followed by a steeper increase in concentration (presumably due to bacterial multiplication). With the contents from the cecum and large intestine, constant SCFA production was detected during the first 30 min (cecum) to 1 h (mid colon). After this time, the SCFA production leveled off. As a consequence, the rate of SCFA production was calculated using the slope of the lines between 0 and 30 min.
In vitro fermentation of D-tagatose with slurries of bacteria harvested from different segments of the gastrointestinal tract.
The rate of microbial fermentation and the composition of fermentation
products after in vitro fermentation of D-tagatose were
determined with slurries of bacteria harvested from the stomach, distal
SI, cecum and mid colon. A 10 g/100 g suspension of gastrointestinal
contents was homogenized for 2 min and pressed through a double layer
of cheesecloth to remove feed and other particulate material. Bacteria
were harvested by centrifugation (20,000 x g for
15 min). The pellet was resuspended in 100 mmol/L phosphate buffer (pH
6.5) to the original volume. The bacterial suspension was divided into
two bottles and incubated anaerobically with and without the addition
of 10 g D-tagatose/L. The incubations were carried out
in serum bottles as described by Jensen and Jensen (1993)
, with an incubation time of 4 h and sampling at 0,
1, 2 and 4 h.
The rate of SCFA production from D-tagatose, calculated as the difference between incubations with and without addition of D-tagatose, was constant during the whole incubation period. The rate of SCFA production from D-tagatose was therefore calculated from the slope of the lines between 0 and 4 h.
Total counts of anaerobic bacteria and D-tagatose–degrading bacteria.
The numbers of total anaerobic and D-tagatose–degrading
bacteria were determined by the most probable number technique (MPN).
Feces were serially diluted 10-fold in anaerobic mineral salt medium
(Jensen and Jørgensen 1994
) supplemented with 20 g/100
g rumen liquid, 10 g/L yeast extract and 10 g/L peptone. A mixture of
four carbohydrates, mannose, xylose, glucose and cellubiose each at a
final concentration of 2.5 g/L each (total anaerobic bacteria), or
D-tagatose at a final concentration of 10 g/L
D-tagatose (tagatose-degrading bacteria) was added to the
tubes through a sterile filter (0.45 µm) after the rest of the media
was sterilized by autoclaving. For each type of bacteria, five parallel
dilution series were prepared. The MPN counts were estimated after a
7-d incubation at 38°C. Tubes were considered positive if the pH was
<5.0. Tubes without added carbohydrates were used as controls. The MPN
counts were calculated from the dilution factors and MPN tables for
five parallel dilutions.
Determination of SCFA, lactic acid and gasses.
The concentration and composition of SCFA and the concentration of
lactic acid were determined as described by Jensen et al. (1995)
. The gas composition in the headspace was determined as
described by Jensen and Jørgensen (1994)
.
Determination of D-tagatose.
D-tagatose remaining after 4 h of in vitro incubation
with bacterial slurries was determined after centrifugation of the
suspension at 2,000 x g for 10 min and addition of
an internal standard solution (arabinose 2 g/L) at a sample to standard
ratio of 1:1. The sample was filtered on a Sep-Pak C18
filter (Waters Corporation, Milford, MA) using a Vac Elut SPS 24
(Analytichem International, Harbor City, CA). The filter was pretreated
with 1 v 100% methanol and 3 v water prior to filtration of
the sample. The sample was subsequently filtered through a 0.2-µm
Sartorius filter (Sartorius AG, Goettingen, Germany).
D-tagatose was quantified by injection of 25 µL of
filtrate onto a sodium-based sulfonated polystyrene column (Shodex
Ionpak KS-801, Showa Denko, Tokyo, Japan) at 85°C using a
Waters HPLC LC Module1 (Waters Corporation) at a flow rate of 0.6
mL/min and a Waters 410 RI detector, internal temperature 45°C,
sensitivity 64 (Johansen et al. 1996
). The in vitro
samples were calibrated against a standard consisting of 3.9 g/L
D-tagatose and 1 g/L arabinose where D-tagatose
was quantified by height and arabinose by area.
The degradation of D-tagatose during incubation was determined as the difference between the added amount (10 g/L) and the concentration of D-tagatose determined after 4 h of incubation.
Calculations.
The energy in D-tagatose recovered as SCFA after microbial
fermentation (E) was calculated according to the
following formula:
 |
Here a is the mean rate of SCFA production per gram
of degraded D-tagatose for bacteria in the cecum or colon,
respectively, mi is the molar proportion
(mol/100 mol) of the specific SCFA and Hi
is the heat of combustion of the same SCFA. The values used for heat of
combustion were as follows: D-tagatose, 15.6 kJ/g; formic
acid, 255 kJ/mol; acetic acid, 877 kJ/mol; propionic acid, 1533 kJ/mol;
butyric acid, 2185 kJ/mol; valeric acid, 2758 kJ/mol; caproic acid,
3501 kJ/mol; and heptanoic acid, 4137 kJ/mol (Chemical Rubber Company 1983
).
Statistical analysis.
The effect of diet in a given intestinal segment, including data
obtained from in vitro incubations, was tested using a simple ANOVA
based on the following general linear model (GLM):
 |
Here Ydi is the dependent variable,
µ is the overall mean, 
d is the effect of
diet, d = 1, 2 and 
di ~
N(0,
2) represents the unexplained
random error (Snedecor and Cochran 1973
).
Data concerning the effect of diet on the development over time were
analyzed using multivariate ANOVA with diet as the between animal
effect and time (d 1, d 8 and d 15) as the within animal effect
according to the following model:
 |
Here d denotes the effect of diet,
d = 1, 2, ßs is the effect of
time, t = 1, 8, 15, ßds is
the interaction between diet and segment and i refers to an
individual pig. The variance component Udi
~ N(0,
2) accounts for the
repeated measurements that were made on the same individual, thereby
rendering these observations correlated, whereas the term
dti ~
N(0,2) represents the unexplained
random error (SAS Institute 1989
).
The effect of litter and series was tested and found not to be significant, therefore series and litter were not included in the statistical analyses presented here. The analyses were performed with SAS for Windows version 6.12 (SAS Institute, Cary, NC).
Statistical analysis of bacterial counts was performed after logarithmic conversion of the data. Values are means ± SEM.
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
The two experimental diets did not create significant differences in
the dry matter content of the feces during the experimental period
except at d 15, when the dry matter content was significantly lower in
pigs fed the tagatose diet than in those fed the control diet
(Fig. 1A
). The dry matter content of feces increased from d -3 to d 1 as a
consequence of changing the diet from the standard pig diet to the
experimental diets. There was a further increase with time
(P = 0.0001) of consuming the experimental diets,
without any effect of diet (P = 0.11) or interaction
between diet and time (P = 0.09).
|
, and was significantly lower in pigs fed
the tagatose diet on d 1 and d 15, but not on d 8. Differences in the
time courses for the two experimental groups led to a significant
interaction between time and diet in the multivariate analysis
(P = 0.03).
The shift of diet from d -3 to d 1 increased the total number of
anaerobic bacteria in feces (Fig. 1C)
. The change in bacterial counts
was evident on d 1 and no further change in numbers during the
experimental period was noted (P = 0.29). Neither in
the univariate analyses nor in the multivariate analysis (P
= 0.47) was there any effect of dietary treatment.
The number of D-tagatose–degrading bacteria in feces from
the tagatose-fed pigs was approximately ten times the value found
in the feces of the pigs fed the control diet (Fig. 1D)
, which led to a
significant effect of experimental diet in the multivariate analysis
(P = 0.0001). A slight further increase in numbers was
seen on d 8, resulting in a significant time effect (P
= 0.046) in the experimental period (d 1 to d 15).
Production of SCFA during in vitro fermentation of unfractionated digesta.
For both dietary groups, the predominant fermentation products produced
in the stomach and distal SI were lactic acid followed by acetic and
formic acids. In addition, a little butyric acid was formed in the
distal SI (Fig. 2
).
|
), there were no significant differences in the total
productions of these acids (P > 0.35). The total SCFA
and lactic acid production in the stomach, accounting for differences
in digesta volumes, was 7.4 ± 2.2 and 8.2 ± 2.2 mmol/h for
the control and tagatose group, respectively (P = 0.8),
whereas the total production in the distal SI was 8.0 ± 1.7
mmol/h for pigs fed the control diet and 6.4 ± 1.7 mmol/h for
those fed the tagatose diet (P = 0.5). The rate of SCFA production from the cecal material of the pigs in the tagatose group consisted in decreasing order of propionic, acetic, butyric, valeric and formic acid, whereas the main acid for the control pigs was acetic acid, with lesser amounts of propionic, butyric and valeric acid. For both dietary groups, the main SCFA produced with colonic material consisted in decreasing order of acetic, propionic, butyric and valeric acid.
The rate of propionic acid production in the cecum was more than twice
as high with the tagatose-fed pigs compared with the pigs fed the
control diet (P = 0.006) and there was a twofold higher
rate of valeric acid production (P = 0.006). Allowing
for differences in digesta volume (Lærke and Jensen 1999
), this led to a total production of valeric acid with the
tagatose diet that was almost six times as high (1.27 ± 0.35
mmol/h) as the control diet (0.22 ± 0.35 mmol/h, P
= 0.049). Although the rate of valeric acid production with the
colonic material was twice as high with the tagatose diet compared with
the control diet, the total valeric acid production with the control
diet (0.33 ± 0.09 mmol/h) was not significantly different
(P = 0.18) from that obtained with the tagatose diet
(0.52 ± 0.09 mmol/h). The total SCFA production in the cecum was
3.0 ± 2.7 and 10.7 ± 2.7 mmol/h for the control and
tagatose group, respectively (P = 0.06), whereas the
productions in the mid colon were 5.1 ± 1.4 and 5.9 ± 1.4
mmol/h, respectively (P = 0.7).
Rate of production and composition of SCFA during in vitro fermentation with slurries of bacteria with and without addition of D-tagatose.
The presented D-tagatose caused no increase in SCFA or
lactic acid production in bacterial slurries from the stomach and
distal SI, indicating that the bacteria from these segments of the
gastrointestinal tract were not able to degrade D-tagatose
in vitro. The rate of SCFA production from D-tagatose
obtained with bacteria from the cecum (Table 1
) was generally higher for the tagatose group than from the
control group, which was significant for formic, propionic, butyric,
valeric and caproic acid but not for acetic acid (P = 0.42). The bacteria from the mid colon from the control group had a
slower SCFA production than the bacteria obtained from the
tagatose-fed pigs. The production of formic, acetic, propionic,
butyric, valeric and caproic acids was higher in the latter group than
in the former.
|
, whereas no effect of diet was seen in
the proportion of propionic and butyric acid produced from
D-tagatose. There was a similar picture with the bacteria
from the mid colon, but the proportion of acetic acid from the control
group was much higher. On the other hand, D-tagatose
induced a much bigger increase in the contribution of butyric acid to
the SCFA production in the colon than in the cecum. Furthermore, the
proportion of propionic acid produced by cecal bacteria was generally
higher than that with bacteria from the mid colon irrespective of the
dietary treatment. SCFA production related to the degradation of D-tagatose.
D-tagatose was not degraded when incubated with bacteria
isolated from the stomach and distal SI, independent of the dietary
treatment of the pigs (values ranged from -1.5 to 0.1 g/(h ·
kg). Figure 3
illustrates the SCFA production by bacteria harvested from the cecum
and colon of pigs fed the control or tagatose diet plotted against
their D-tagatose degradation. Generally, there was a very
low degradation of D-tagatose with bacteria harvested from
the pigs in the control group. Thus the degradation of
D-tagatose was significantly higher for the bacteria
isolated from the mid colon of pigs fed D-tagatose [15.3
± 1.2 g/(h · kg), n = 6] compared with the degradation
seen with bacteria from the control pigs [1.2 ± 1.2 g/(h ·
kg), n = 5, P = 0.0001]. There was no significant
difference in the capacity of the microbiota from the cecum to degrade
D-tagatose between the control diet [2.7 ± 1.0
g/(h · kg), n = 6] and the tagatose diet [4.4
± 1.1 g/(h · kg), n = 6, P = 0.30].
|
Gas production during in vitro fermentation of isolated bacteria with and without addition of D-tagatose.
Major increases in hydrogen production were seen during the in vitro
incubation of D-tagatose with bacteria isolated from the
cecum and colon of tagatose-fed pigs compared with the pigs fed the
control diet (Table 2
). Significant effects of
D-tagatose on methane production were not seen until mid
colon where the production in the tagatose group was more than five
times the production in the control group.
|
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
The increase in density of the population of total anaerobic bacteria per gram of fresh feces and the increase in dry matter content of feces at the beginning of the experiment could be explained by a reduction in the dietary fiber content of the experimental diets (~5 g/100 g of dry matter) compared with traditional pig feed (18–20 g/100 g of dry matter).
Although the response to D-tagatose in terms of
number of D-tagatose–degrading bacteria was rapid, there
was a reduction in the pH of the feces on d 1 (equivalent to d 3 after
the first exposure to D-tagatose), which was very clear for
the pigs fed the tagatose diet. Presumably, at this stage the
microbiota in the proximal part of the colon was not adapted to
digestion of D-tagatose, and their ability to degrade
D-tagatose was limited. Consequently, more
D-tagatose would pass on to the more distal parts of the
colon before it was completely fermented, which would lead to a larger
SCFA production in this area and a lower pH of the excreta. After
adaptation (on d 8 and d 15), D-tagatose could be fully
fermented in the proximal large intestine, and less substrate would be
available for the microbiota in the distal large intestine, leading to
a higher pH compared with the period just after introduction of
D-tagatose to the diet. These results suggest that a
certain period of adaptation is necessary before the gastrointestinal
microbiota can digest D-tagatose. This was confirmed in the
in vitro fermentation studies with isolated bacteria, where the rate of
D-tagatose degradation and SCFA production was larger for
the adapted bacteria (from the pigs in the tagatose group) than the
unadapted bacteria. The results also indicate that the large intestinal
bacteria have the capacity to metabolize D-tagatose at much
higher levels than the 10% added to the diet of this experiment. The
lack of difference between dietary groups in the SCFA production during
in vitro incubation of digesta from the mid colon of pigs after a 20-d
adaptation period suggests that at this point all
D-tagatose was fermented in the proximal part of the colon
and did not reach the mid colon. Indeed, the in vitro incubations
showed that the bacteria in the distal colon had a high capacity to
digest D-tagatose, although the microbial activity in the
distal colon was lower than in the cecum and proximal colon
(Lærke and Jensen 1999
). However, the possibility
remains that a higher microbial activity caused by fermentation of
D-tagatose would have been seen in the more distal part of
the large intestine if the pigs had been killed later than 3 h
after the morning meal. At 3 h after a meal, dietary material
present in the large intestine of pigs mostly stems from the preceding
meal fed 19 h earlier, where D-tagatose, which was
found to be a readily fermentable substrate, would have been fermented.
The rates of SCFA production in digesta from the cecum and mid colon
found in this investigation were from three (acetic acid) to seven
(propionic acid) times higher than the rates reported by Holtug et al. (1992)
for pigs calculated from 6 h in vitro
incubations. This difference is probably due to the shorter incubation
time used in this investigation. The rates of SCFA production would
have been underestimated in this study if they had been based on
incubation times >30 min, because the rate of SCFA production
decreased after prolonged fermentation.
Strikingly, no D-tagatose degradation and SCFA or
lactic acid production took place with bacteria isolated from the
stomach and distal small intestine, neither from the adapted nor from
the unadapted pigs, suggesting that the microbiota in the upper
gastrointestinal tract were not able to adapt to utilization of
D-tagatose as substrate for microbial fermentation.
Consequently, the SCFA produced in the upper gastrointestinal tract
would be expected to stem from other components of the diet, which was
supported by the observation that the total amount and composition of
SCFA and lactic acid present at slaughter was not significantly
different in the two dietary groups (Lærke and Jensen, 1999
). Other easily fermentable carbohydrates such as
raffinose-oligosaccharides and mannitol (Canibe and Bach Knudsen 1997
, Gdala et al.1997
, Saunders and Wiggins 1981
), inulin (Bach Knudsen and Hessov 1995
, Graham and Åman 1986
) and ß-glucans
(Bach Knudsen and Hansen 1991
, Bach Knudsen et al. 1993
, Johansen et al. 1997
, Sundberg et al. 1996
) have been found to be degraded to variable extents
at the terminal ileum in healthy ileostomy patients, ileum cannulated
and intact pigs. Our finding that D-tagatose, a chemically
modified monosaccharide, is totally unavailable for the microbiota in
the small intestine of the pigs is therefore unexpected. Lactulose is
one of few other easily fermentable carbohydrate sources, which has
been found fully recovered from ileostomies (Saunders and Wiggins 1981
).
An interesting observation was the high production of valeric
acid obtained by in vitro fermentation of digesta from the cecum and
colon of the pigs fed D-tagatose and also by incubation of
the isolated bacteria with D-tagatose. The results indicate
that D-tagatose may particularly stimulate the production
of this acid but whether this is due to a shift in metabolism of the
normal gut flora or proliferation of specific groups of bacteria is not
known. Very few bacteria have been shown to produce valeric acid as a
major fermentation product (Holdeman et al. 1977
), and
it has even been suggested that it is exclusively a product of amino
acid fermentation (Rasmussen et al. 1988
). However,
enhanced amino acid catabolism in the tagatose-fed pigs is not a
likely explanation for the difference seen between the two experimental
diets in this study because the productions of isobutyric and
isovaleric acid, which also are products of protein degradation, were
very small and comparable in the two experimental diets.
The bacterial slurries, especially from the colon of the
tagatose-adapted pigs, also produced significant amounts of butyric acid, which is believed to be clinically important because
of its metabolic role in the health of the colonic mucosa
(Roediger 1980
) and its protective role against
colon-rectal cancer (Young and Gibson 1994
).
Previous reports of carbohydrates that stimulate butyric acid production in the lower gut of monogastrics include starch
(Englyst et al. 1987
, Mathers and Smith 1993
, Silvester et al. 1995
, Weaver et al. 1992
), oat bran (Bach Knudsen et al. 1993
, Christensen et al. 1999
), sorbitol,
galacturonic and glucuronic acid (Mortensen et al. 1988
), neosugar (Berggren et al. 1993
) and
fructooligosaccharides (Campbell et al. 1997
,
Gibson and Wang 1994
, Le Blay et al. 1999
). From this study it appears that D-tagatose
can be added to the list of components that may beneficially enhance the production of butyric acid in the large intestine.
It is also noteworthy that the bacteria isolated from the cecum
of both dietary groups produced high proportions of propionic acid,
whereas the bacteria harvested from the mid colon did not. Previous
studies have concluded that the site of collection and adaptation to
the diet had a significant influence on the composition of the SCFA
produced in vitro and in vivo in rats (Berggren 1996
,
Monsma and Marlett 1995
). Our results support this for
pigs but are in contrast to the results of Christensen et al. (1999)
, who found that adaptation to the diet was less
important if the contribution of SCFA from the fecal inoculum produced
during in vitro incubation was subtracted from the total production.
Based on the in vitro incubations we estimated that ~51% of
the energy in D-tagatose was recovered as SCFA after
microbial fermentation. It is important to note that this value was
obtained from actively metabolizing bacteria, which were not limited in
the amounts of available D-tagatose. It is well known that
bacteria growing in a medium with excess carbohydrate will accumulate
the carbohydrate and store it in the bacterial cells (Dawes and Senior 1973
, Herbert 1961
). It is likely that an
extended fermentation until all the substrate was used, which is a
situation similar to the in vivo conditions, would lead to higher
values for recovery of energy as SCFA than obtained in this study. In
comparison, Englyst et al. (1987)
found that the amount
of available energy in SCFA from fermentation of pure–cell-wall
polysaccharides (arabinogalactans and pectins) was 35–54%, whereas
starch fermentation yielded ~60%. Mikkelsen and Jensen (1997)
reported an energy yield of 59% from fermentation of
fructo-oligosaccharides. According to Hungate (1966)
, the energy content in SCFA is ~60% of that of the
substrate during the fermentation of carbohydrates in the rumen,
whereas Cummings (1994)
has suggested an upper limit of
60% energy yield for carbohydrate fermentation in the colon and a
fixed energy yield of 75% was suggested by McNeil (1984)
.
Some of the energy in D-tagatose was lost in the form of
various gasses. Substantial amounts of hydrogen could be produced by
the fermentation of D-tagatose with bacteria isolated from
the cecum and colon, while large amounts of methane were produced
during incubation of D-tagatose with bacteria isolated from
the mid colon. In that respect, D-tagatose resembles other
easily fermentable carbohydrates such as fructo-oligosaccharides,
galacto-oligoosaccharides and lactulose, which have also been shown to
increase hydrogen production (Cristl et al. 1992
,
Djouzi and Andrieux, 1997
). A high gas production in the
large intestine can have several negative effects on the host,
including flatulence, abdominal pain and diarrhea. Increased flatulence
after continuous ingestion of large amounts of D-tagatose
(30 g/d) has been reported in man (Buemann et al. 1999
).
In conclusion, the proportion of D-tagatose–degrading bacteria in feces was dependent on the presence of D-tagatose in the diet. The rate of D-tagatose fermentation was higher for bacteria adapted to D-tagatose compared with unadapted bacteria. D-tagatose was not utilized as substrate for the microbiota in the stomach and small intestine of pigs, whereas it was readily fermented by the microbiota in the cecum and colon. Consequently, D-tagatose contributed energy to the host. The proportion of butyric and valeric acid produced by microbiota in the colon of pigs adapted to D-tagatose was two to three times higher than from the unadapted pigs. In addition to short chain fatty acids, D-tagatose could also be metabolized by the bacteria of the large intestine to form hydrogen and methane.
|
ACKNOWLEDGMENTS |
|---|
|
FOOTNOTES |
|---|
2 Supported by MD Foods Ingredients A.M.B.A., Netherlands Vium, Denmark and The Danish Institute of Agricultural Sciences, Denmark.
3 Submission of this paper was approved by the Danish Institute of Agricultural Sciences.
5 Abbreviations used: Ace, acetic acid; but,
butyric acid; Cap, caproic acid; for, formic acid; Hep, hepatonic acid;
Iso-but, isobutyric acid; Iso val, isovaleric acid; Lac, lactic
acid; MPN, most probable number; Pro, propionic acid; SCFA, short chain
fatty acids; SI, small intestine; Val, valeric acid.
Manuscript received October 6, 1999. Initial review completed November 23, 1999. Revision accepted March 24, 2000.
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
1. Bach Knudsen K. E., Hansen I. Gastrointestinal implications in pigs of wheat and oat fractions. 1. Digestibility and bulking properties of polysaccharides and other major constituents. Br. J. Nutr. 1991;65:217-232[Medline]
2. Bach Knudsen K. E., Hessov I. Recovery of inulin from Jerusalem artichoke (Helianthus tuberosus L.) in the small intestine of man. Br. J. Nutr. 1995;74:101-113[Medline]
3. Bach Knudsen K. E., Jensen B. B., Hansen I. Oat bran but not a ß-glucan-enriched oat fraction enhances butyrate production in the large intestine of pigs. J. Nutr. 1993;123:1235-1247
4. Berggren A. Formation, pattern and physiological effects of short-chain fatty acids. Doctoral thesis 1996 University of Lund Lund, Sweden
5. Berggren A. M., Björck I.M.E., Nyman G. L. Short-chain fatty acid content and pH in cecum of rats given various sources of carbohydrates. J. Sci. Food. Agric. 1993;63:397-406
6. Buemann B., Toubro S., Astrup A. Human gastrointestinal tolerance to D-tagatose. Reg. Tox. Pharmacol. 1999;29:S71-S77[Medline]
7.
Campbell J. M., Fahey G. C., Bryan W. W. Selected indigestible oligosaccharides affect large bowel mass, cecal and fecal short-chain fatty acids, pH and microbiota in rats. J. Nutr. 1997;127:130-136
8. Canibe N., Bach Knudsen K. E. Digestibility of dried and toasted peas in pigs. 1. Ileal and total tract digestibilities of carbohydrates. Anim. Feed Sci. Tech. 1997;64:293-310
9. Chemical Rubber Company West R. C. eds. CRC Handbook of Chemistry and Physics 64th ed. 1983 CRC Press, Boca Raton FL.
10. Christensen D. N., Bach Knudsen K. E., Wolstrup J., Jensen B. B. Integration of ileum cannulated pigs and in vitro fermentation to quantify the effect of diet composition on the amount of short-chain fatty acids available from fermentation in the large intestine. J. Sci. Food Agric. 1999;79:755-762
11. Cristl S. U., Murgatroyd P. R., Gibson G. R., Cummings J. H. Production, metabolism, and excretion of hydrogen in the large intestine. Gastroenterology 1992;102:1269-1277[Medline]
12. Cummings J. H. Quantitating short chain fatty acid production in humans. Binder H. J. Cummings J. Soergel K. H. eds. Short Chain Fatty Acids 1994:11-19 Kluwer Academic Publishers Dordrecht, Germany.
13. Dawes E. A., Senior P. J. The role of regulation of energy reserve polymers in micro-organisms. Adv. Microbial. Physiol. 1973;10:135-267[Medline]
14. Djouzi Z., Andrieux C. Compared effects of the three oligosaccharides on metabolism of intestinal microbiota in rats inoculated with a human faecal flora. Br. J. Nutr. 1997;78:313-324[Medline]
15. Englyst H. N., Hay S., Macfarlane G. T. Polysaccharide breakdown by mixed populations of human fecal bacteria. FEMS Microbiol. Ecol. 1987;95:163-171
16. Gdala J., Johansen H. N., Bach Knudsen K. E., Knap I. H., Wagner P., Jørgensen O. B. The digestibility of carbohydrates, protein and fat in the small and large intestine of piglets fed non-supplemented and enzyme supplemented diets. Anim. Feed. Sci. Tech. 1997;65:15-33
17. Gibson G., Wang X. Enrichment of bifidobacteria from human gut contents by oligofructose using continuous culture. FEMS Microbiol. Lett. 1994;118:121-128[Medline]
18. Graham H., Åman P. Composition and digestion in the pig gastrointestinal tract of Jerusalem artichoke tubers. Food Chem 1986;22:1-10
19. Herbert D. The chemical composition of micro-organisms as a function of their environment. Microbial reaction to environment. Symp. Soc. Gen. Microbiol. 1961;11:391-416
20. Holdeman L. V., Cato E. P., Moore E. C. Anaerobe laboratory manual 1977 Virginia Polytechnic Institute and State Univ Blacksburg, VA.
21. Holtug K., Rasmussen H. S., Mortensen P. B. An in vitro study of short-chain fatty acid concentrations, production and absorption in pig (Sus scrofa) colon. Comp. Biochem. Physiol. 1992;103A:189-197
22. Hungate R. E. The Rumen and Its Microbes 1966 Academic Press New York.
23. Jensen B. B., Jensen M. T. In vitro measurements of microbial production of skatole in the digestive tract of pigs. Bonneau M. eds. Measurements and Prevention of Boar Taint in Entire Male Pigs, Les colloques no 60 1993:99-105 Institute Nationale de la Recherche Agronomique Roskilde, Denmark.
24.
Jensen B. B., Jørgensen H. Effect of dietary fiber on microbial activity and microbial gas production in various regions of the gastrointestinal tract of pigs. Appl. Environ. Microbiol. 1994;60:1897-1904
25. Jensen M. T., Cox R. P., Jensen B. B. Microbial production of skatole in the hind gut of pigs fed different diets and its relation to skatole deposition in backfat. Anim. Sci. 1995;61:293-304
26. Johansen H. N., Bach Knudsen K. E., Glitsø L. V. Influence of extraction solvent and temperature on the quantitative determination of oligosaccharides from plant materials by HPLC. J. Agric. Food Chem. 1996;44:1470-1474
27. Johansen H. N., Bach Knudsen K. E., Wood P. J., Fulcher R. G. Physico-chemical properties and the degradation of oat bran polysaccharides in the gut of pigs. J. Sci. Food Agric. 1997;73:81-92
28.
Lærke H. N., Jensen B. B. D-tagatose has low small intestinal digestibility, but high fermentability in the large intestine of pigs. J. Nutr. 1999;129:1002-1009
29.
Le Blay G., Michel C., Blottière H. M., Cherbut C. Prolonged intake of fructo-oligosaccharides induces a shert-term elevation of lactic acid-producing bacteria and a persistent increase in cecal butyrate in rats. J. Nutr. 1999;129:2231-2235
30.
Levin G. V., Zehner L. R., Saunders J. P., Beadle J. R. Sugar substitutes. Their energy values, bulk characteristics, and potential health benefits. Am. J. Clin. Nutr. 1995;62(suppl.):1161S-1168S
31. Mathers J. C., Smith H. Factors influencing cecal butyrate in rats fed on raw potato starch. Proc. Nutr. Soc. 1993;52:376A(abs.)
32.
McNeil N. The contribution of the large intestine to energy supplies in man. Am. J. Clin. Nutr. 1984;39:338-342
33. Mikkelsen L. M., Jensen B. B. Effect of fructo-oligosaccharide (FO) on the intestinal microbiota of rats; and determination of the fermentative transformation and energy transmission of FO. Proceedings of the International Symposium ‘Non-digestible oligosaccharides: healthy food for the colon?’ December 4–5, 1997 1997:158 Wageningen Netherlands.
34. Monsma D. J., Marlett J. A. Rat cecal inocula produce different patterns of short-chain acids than fecal inocula in in vitro fermentations. J. Nutr. 1995;125:2463-2470
35. Mortensen P. B., Holtug K., Rasmussen H. S. Short-chain fatty acid production from mono and disaccharides in a fecal incubating system: implications for colonic fermentation of dietary fiber in humans. J. Nutr. 1988;118:321-325
36. Rasmussen H. S., Holtug K., Mortensen P. B. Degradation of amino acids to short-chain fatty acids in humans. An in vitro study. Scand. J. Gastroenterol. 1988;23:178-182[Medline]
37.
Roediger W.E.W. Role of anaerobic bacteria in the welfare of the colonic mucosa in man. Gut 1980;21:793-798
38. SAS Institute Inc SAS® User’s Guide: Statistics, Version 6 4th ed. 1989 SAS Institute Inc Cary, NC.
39.
Saunders D. R., Wiggins H. S. Conservation of mannitol, lactulose, and raffinose by the human colon. Am. J. Physiol. 1981;241:G397-G402
40. Saunders J. P., Zehner L. R., Levin G.V.B. Disposition of D-[U-14C]tagatose in the rat. Regul. Toxicol. Pharmacol. 1999;29:S46-S56[Medline]
41.
Silvester K. R., Englyst H. N., Cummings J. H. Ileal recovery of starch from whole diets containing resistant starch measured in vitro fermentation of ileal effluent. Am. J. Clin. Nutr. 1995;62:403-411
42. Snedecor G. W., Cochran W. G. Statistical Methods 6th ed. 1973 Iowa State University Press Ames, IA.
43.
Sundberg B., Wood P., Lia Å., Andersson H., Sandberg A.-S., Hallmans G., Åman P. Mixed-linked beta-glucan from breads of different cereals is partly degraded in the human ileostomy model. Am. J. Clin. Nutr. 1996;64:878-885
44.
Weaver G. A., Krause J. A., Miller T. L., Wolin M. J. Corn starch fermentation by the colonic microbial community yields more butyrate than dose cabbage fiber-fermentation: corn starch fermentation rates correlates negatively with methanogenesis. Am. J. Clin. Nutr. 1992;55:70-77
45. Young G. P., Gibson R. P. Butyrate and the colorectal cancer cell. Binder H. J. Cummings J. Soergel K. H. eds. Short Chain Fatty Acids 1994:148-160 Kluwer Academic Publishers Dordrecht, Germany.
This article has been cited by other articles:
|
|
 |
|
  Y. Huang, M. Suyemoto, C. D. Garner, K. M. Cicconi, and C. Altier Formate Acts as a Diffusible Signal To Induce Salmonella Invasion J. Bacteriol., June 15, 2008; 190(12): 4233 - 4241. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 N. Canibe, O. Hojberg, S. Hojsgaard, and B. B. Jensen Feed physical form and formic acid addition to the feed affect the gastrointestinal ecology and growth performance of growing pigs J Anim Sci, June 1, 2005; 83(6): 1287 - 1302. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 O. Hojberg, N. Canibe, B. Knudsen, and B. B. Jensen Potential Rates of Fermentation in Digesta from the Gastrointestinal Tract of Pigs: Effect of Feeding Fermented Liquid Feed Appl. Envir. Microbiol., January 1, 2003; 69(1): 408 - 418. [Abstract] [Full Text] [PDF]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
