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Applied and Environmental Microbiology, October 1998, p. 4084-4088, Vol. 64, No. 10
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Effective Recovery of Bacterial DNA and
Percent-Guanine-Plus-Cytosine-Based Analysis of Community Structure
in the Gastrointestinal Tract of Broiler Chickens
Juha H. A.
Apajalahti,1,*
Laura K.
Särkilahti,1
Brita R. E.
Mäki,1
J. Pekka
Heikkinen,1
Päivi H.
Nurminen,1 and
William
E.
Holben2
Cultor Corporation Technology Center,
FIN-02460, Kantvik, Finland,1 and
Division of Biological Sciences, The University of Montana,
Missoula, Montana 59812-10022
Received 13 May 1998/Accepted 23 July 1998
 |
ABSTRACT |
A DNA-based, direct method for initial characterization of the
total bacterial community in ileum and cecum of the chicken gastrointestinal (GI) tract was developed. The efficiencies of bacterial extraction and lysis were >95 and >99%, respectively, and
therefore the DNA recovered should accurately reflect the bacterial
communities of the ileal and cecal digesta. Total bacterial DNA samples
were fractionated according to their percent G+C content. The profiles
reflecting the composition of the bacterial community were reproducible
within each compartment, but different between the compartments of the
GI tract. This approach is independent of the culturability of the
bacteria in the consortium and can be used to improve our understanding
of how diet and other variables modulate the microbial communities of
the GI tracts of animals.
| |
TEXT |
The microbiology of the
gastrointestinal (GI) tracts of farm animals has long been of interest
for reasons of both food safety and animal nutrition and health.
Attempts are being made to bolster host defenses by using feed
ingredients which favor the growth of bacteria generally regarded as
beneficial. Typical modulators of GI tract ecology are prebiotics
(e.g., oligosaccharides [25]) and probiotics (e.g.,
lactobacilli and bifidobacteria [5, 24, 25]).
Understanding of the mode of action and development of effective
products would be greatly assisted by reliable tools with which to
monitor the composition of the microbial community in the GI tract.
Molecular techniques have been employed to monitor for the presence of
specific bacterial pathogens in chicken digesta samples (2, 6, 16,
26). In order to place findings from such single-species studies
in perspective, some knowledge of the total bacterial community
composition of this system must be obtained. However, direct analyses
(i.e., analyses that are not based on culturability) of bacterial
community structure in the various compartments of the chicken GI tract
have not yet been performed. In other complex environments, DNA-based
community-level molecular analyses have been used to obtain information
on microbial community diversity, structure, and function (10, 12,
20, 21, 27, 30, 36). In this report, we show for the first time
essentially quantitative recovery and lysis of the bacterial fraction
from the distal part of broiler chicken small intestine (ileum) and cecum and also purification of total DNA representing the bacterial community, a requisite step for DNA-based community analyses. We also
provide an initial depiction of the total bacterial community in these
two chicken GI tract compartments obtained by DNA-based profiling.
Bacterial extraction.
Broiler chickens were raised in cages
and unless otherwise indicated were fed a standard wheat-based diet
with no antibiotics. The birds were killed by cervical dislocation, and
the ileum and cecum were immediately removed and dissected. All digesta
samples were kept on ice and processed further within 2 h.
Quantitative bacterial recovery and lysis are prerequisites for all
subsequent comprehensive DNA-based community analyses. Therefore, these
protocols were carefully developed, and their efficiencies were
demonstrated. Digesta samples from the different compartments of the
broiler chicken digestive tract were found to vary in composition, the amount of material present, and bacterial density. Bacterial numbers in
the ileum, the distal part of small intestine, were typically between
107 and 109 per g of digesta, and the bulk of
the dry matter was undigested feed particles, which had to be
eliminated prior to bacterial lysis. This was accomplished through the
five-cycle differential centrifugation process. Seven grams of ileal
digesta was suspended in 200 ml of wash buffer (50 mM sodium phosphate
buffer [pH 8], 0.1% Tween 80). The suspensions were then shaken for
20 min on a reciprocating horizontal platform shaker at 100 oscillations/min at room temperature. Undigested feed particles were
removed from the suspension by low-speed centrifugation at 200 × g for 15 min. The supernatant was carefully transferred to a
clean flask and kept on ice. The digesta pellet was again suspended in
wash buffer, the differential centrifugation process was repeated for a
total of five rounds, and samples of both the suspended digesta and the
low-speed supernatant were taken before each round for microscopic enumeration of bacteria. The bacteria in the pooled supernatants were
collected by centrifugation at 30,000 × g for 15 min
at room temperature. In the dual cecal compartments, solid feed
particles were absent and bacterial densities were typically between
1010 and 1011 per g of digesta. For efficient
bacterial recovery and lysis from this compartment, it was important to
remove the viscous polysaccharides and other soluble compounds
interfering with these procedures. This was achieved by a four-cycle
dilution and washing process. Cecal samples (1 g) were suspended in 30 ml of wash buffer and then shaken for 10 min on a reciprocating
horizontal platform shaker at moderate speed at room temperature. The
resulting suspension was subjected to centrifugation at 30,000 × g for 15 min to collect the bacterial fraction. The pellet
was resuspended and washed three more times in 30 ml of fresh wash
buffer, and samples of the suspended bacteria were taken at each step
for direct microscopic enumeration. Microscopic enumeration of bacteria
was accomplished by fluorescence microscopy of DAPI
(4',6-diamidino-2-phenylindole)-stained cells essentially as described
previously (32). An automated stage-manipulation system and
digital camera using Image Pro Plus (Media Cybernetics, Silver
Spring, Md.) and Amcas (Cultor Technology Center, Kantvik, Finland)
software were used to enumerate cells.
Direct microscopic counts were used to enumerate bacterial numbers in
the ileal and cecal digesta for the starting samples and during the
bacterial extraction. Methods that have been used for bacterial
extraction from other solid matrices (9, 11) gave relatively
poor recovery of bacterial cells (<50%) when applied to the digesta
samples (data not shown), indicating the need to optimize these
procedures for the samples being analyzed. The differential
centrifugation approach resulted in extraction of approximately 96% of
the cells that were present in the starting ileum digesta samples. The
first cycle recovered 50% of the bacteria, whereas a minimum of four
cycles were needed for 90% recovery (Fig.
1). Similarly, cell counts in the
bacterial fraction from the cecum indicated quantitative recovery, with
>97% of the bacterial cells present in the starting sample being
recovered by the four-cycle dilution and washing process. Microscopic
examination of the resuspended bacterial pellets from the high-speed
centrifugation step indicated that bacteria in the supernatant pelleted
quantitatively at the g forces used. Lower g
forces resulted in incomplete pelleting and poor bacterial recovery,
probably due to the presence of small bacterial cells in these samples
(data not shown). It is worth noting that the efficiency of bacterial
recovery from these samples exceeds values obtained from other types of
matrices, such as soils and sediments, which typically range between 33 and 42% (11, 15). Presumably these high recoveries reflect
both the importance of multiple rounds of extraction of cells from
environmental samples (four or five for each protocol) and a lack of
tight adhesion between bacteria and the digesta matrix.
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FIG. 1.
Cell removal from digesta material
( ) and
cumulative recovery
( ) of
bacteria from ileal digesta samples during the bacterial recovery
process. The data were obtained by direct microscopic enumeration of
bacterial cells in the supernatant following each round of
differential, low-speed centrifugation. Percent removal is based on the
number of cells present in the starting ileal digesta sample. Error
bars indicate 1 standard error of the mean (n = 4).
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Bacterial lysis and DNA recovery.
Effective lysis of complex
mixtures of bacteria often requires a combination of various treatments
known to be effective for lysing individual bacterial populations
(11). To maximize the lysis efficiency for the bacterial
populations present in chicken digesta, the salient steps from a
variety of cell lysis protocols were combined to obtain the general
bacterial lysis protocol. Samples of the bacterial fractions (0.3 g of
cecal bacterial pellet and 0.7 g of ileal bacterial pellet) were
resuspended in 3 ml of TE buffer (10 mM Tris [pH 8], 1 mM EDTA).
These suspensions were subjected to five freeze-thaw cycles of
incubation at 
70°C for 60 min followed by 40°C for 15 min.
Lysozyme was then added to each sample as 0.7 ml of a 200-mg/ml stock
solution (in TE buffer) followed by incubation at 37°C for 3 h.
After the addition of 0.2 ml of sodium dodecyl sulfate (10%
[wt/vol]) and 20 µl of proteinase K solution (20 mg/ml in TE
buffer), the mixture was incubated at 37°C for an additional hour.
Following this incubation, 0.72 ml of 5 M NaCl, 0.6 ml of 10% CTAB
(hexadecyltrimethyl ammonium bromide) in 0.7 M NaCl, and 1 g of
1,000-µm-diameter glass beads (Sigma Chemical Company, St. Louis,
Mo.) were added. The mixtures were then incubated at 65°C for 20 min
with vortexing for 30 s after every 5 min. Samples of the
suspended bacterial fraction were taken before and after lysis to
assess lysis efficiency by direct microscopic enumeration of bacteria.
The cell lysate was extracted with an equal volume of
chloroform-isoamyl alcohol (24:1) and then was subjected to
centrifugation at 6,000 × g for 10 min at room
temperature to separate the aqueous and organic phases. After transfer
of the aqueous phase to a clean Corex test tube, the DNA was
precipitated by addition of 0.6 volume of 100% isopropanol and
incubation for 1 h at room temperature. The precipitated DNA was
collected by centrifugation at 10,000 × g for 15 min
at room temperature. The DNA pellet was washed briefly with 70%
ethanol, vacuum dried, and dissolved in 2 ml of TE buffer. The
extracted DNA was then purified by two rounds of cesium
chloride-ethidium bromide equilibrium gradient centrifugation as
described by Holben (9). The DNA bands from these gradients
were subjected to ethidium bromide extraction, desalting, and
concentration of DNA by ethanol precipitation, also as described by
Holben (9). DNA concentration and purity were estimated
based on A260/280.
Microscopic analysis was used to monitor the efficiency of lysis of the
bacterial fractions obtained from ileal and cecal
digesta. Direct
microscopic counts before and after the lysis
procedure indicated that
>99% of the bacteria in both the ileal
and cecal bacterial fractions
were lysed (data not shown). Combined
with the highly efficient
recovery of bacteria from the digesta
samples, the efficient lysis
achieved with this protocol results
in bacterial community DNA samples
that are presumed to be representative
of the bacterial populations
present in the digesta from the chicken
GI tract. Thus, the procedures
developed here provide sound samples,
not only for the community
analysis described below, but also
for any other DNA-based analysis
(PCR, hybridizations, etc.) that
might be used for the characterization
of bacterial communities
in the chicken GI tract.
Percent G+C profiles of the digestal communities.
To obtain a
profile of ileal and cecal digesta bacterial communities based on
percent G+C content, 100 µg of each DNA sample was subjected to
cesium chloride-bisbenzimidazole gradient analysis as described
previously (11). Since there was no residual protein in
these highly purified DNA preparations, DNA quantitation was based on
A280, which minimizes background absorbance
resulting from the cesium chloride gradient itself and unbound
bisbenzimidazole. Determination of the percent G+C content represented
by each gradient fraction was accomplished by regression analysis
(r2 > 0.99) of data obtained from gradients
containing standard DNA samples of known percent G+C composition
(Clostridium perfringens, Escherichia coli, and
Micrococcus lysodeikticus). Fractionation of total bacterial
community DNA based on percent G+C content has previously been used to
analyze bacterial communities and how they respond to changing
conditions in soils (10), bioreactors (12), and
cricket hindgut (30). The data reported here extend that
analysis to studies of the bacterial communities in the GI tracts of
broiler chickens. To assess variability and reproducibility in this
system, replicate samples of bacterial community DNA isolated from
ileal and cecal digesta of 4-week-old broiler chickens were subjected
to cesium chloride-bisbenzimidazole gradient analysis. The data
indicate that replicate samples from each compartment show similar
profiles, while the bacterial communities in the ileal and cecal
compartments were substantially different from one another (Fig.
2). While the pooling of digesta from two
birds may have resulted in some averaging of differences in bacterial communities between individual birds, the replicate samples analyzed were independent replicates, and the data thus suggest that birds raised together under identical conditions have similar bacterial community compositions in their GI tracts (Fig. 2).
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FIG. 2.
Profiles of the bacterial community in ileal (A) and
cecal (B) digesta material from 4-week-old chickens. Data were obtained
as a continuous stream from the UV absorbance flow cell and are
presented as percent G+C content versus relative abundance. The solid
line in each panel indicates the mean of two replicate samples. The
shaded areas in each panel indicate the standard error of the mean.
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Numerous bacterial species have been characterized from the GI tract of
the chicken by selective plating and subsequent identification
by
biochemical profiles (
1,
3,
4,
7,
8,
14,
17,
23,
28,
29,
35). Such studies clearly demonstrate the
presence of the
selected bacterial groups, but probably do not
accurately depict the
total bacterial community in question. Figure
3 shows ranges of percent G+C content in
the DNA of some of the
genera which have been reported to be present in
the GI tract
of the chicken. The ileal and cecal bacterial communities
illustrated
by the percent G+C profiles (Fig.
2 and
4) may be, but are not
necessarily,
composed of the members of the bacterial genera listed.
Speculation
regarding the identity of organisms represented by
these profiles is
useful in guiding future studies, but to truly
demonstrate whether a
particular organism or group of organisms
is present in a given region
of the bacterial community profile
requires additional DNA
hybridization or ribosomal DNA analyses
as reported previously
(
12,
33).
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FIG. 3.
Ranges of percent G+C content in bacterial genera
present in the GI tract of the chicken. Boxes indicate ranges, which
accommodate 80% of the species within a given genus, and the vertical
line in each box is the median of that genus. The values in parentheses
show the number of species included in the survey. The figure is based
on the literature data (13, 19, 22, 34, 37).
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FIG. 4.
Profiles of the cecal bacterial community in 10-day-old
chickens. Data were obtained as a continuous stream from the UV
absorbance flow cell and are presented as percent G+C content versus
relative abundance. Each line represents the profile obtained from
pooled cecal samples of six broiler chickens. Solid line ( ),
uninoculated control; dotted line (........),
inoculation with 108 cecal bacteria from commercially
raised, free-range chickens; dashed line (---),
inoculation with 105 C. perfringens cells.
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Changing digestal communities by live-fed bacteria.
To
determine whether it is possible to alter the community in the GI tract
and detect such changes in community structure by using this DNA-based
community analysis, newly hatched chicks were inoculated with a single
dose either of bacteria from the cecal contents of commercially raised,
free-range chickens or C. perfringens. C. perfringens (1:1
mixture of ATCC 13124 and ATCC 3626) was grown overnight anaerobically
at 37°C in RCM medium (Difco, Detroit, Mich.). The bacterial cells
were collected by centrifugation at 20,000 × g and
then resuspended in anaerobic 0.9% NaCl solution. Approximately
105 bacteria in 0.1 ml of solution were introduced into the
crop of the broiler chicks with a 1-ml syringe. In another treatment, broiler ceca were dissected from 30 5-week-old broiler chickens, and
the contents (digesta) were pooled in an anaerobic glove box and then
stored frozen at 70°C in 1-ml aliquots. An aliquot was used to
inoculate 50 ml of anaerobic brain heart infusion medium (LAB M; Bury,
United Kingdom) in a tightly stoppered serum bottle. The culture was
grown static overnight at 37°C. The bacteria were harvested by
centrifugation at 20,000 × g and then resuspended in
anaerobic 0.9% NaCl solution. Approximately 108 bacteria
were introduced as 0.1 ml of an appropriately diluted bacterial
suspension directly into the crop of the broiler chicks as described
above. Bacterial numbers in the inocula were determined by direct
microscopy. Following bacterial inoculation, the birds were raised
under identical conditions for 10 days on a standard, antibiotic-free,
corn-based diet, and then the cecal bacterial community DNA was
recovered and analyzed. The bacterial community profiles from the ceca
(cecal digesta of six chickens were pooled) of differentially
inoculated broiler chickens were different from each other and from
those of the uninoculated control birds (Fig. 4).
Compared to the percent G+C profile from the uninoculated broiler
chickens, those inoculated with
C. perfringens showed an
increase in the relative abundance of DNA, and hence bacteria,
having
25 to 40% G+C content, and a decrease in the relative abundance
of
bacteria whose DNA is in the 42 to 55% G+C range (Fig.
4).
Although no
subsequent confirming analyses were performed, this
is consistent with
an increase in the abundance of clostridia,
campylobacteria, and
enterococci or perhaps unculturable bacterial
populations having the
indicated percent G+C content. The cecal
bacterial community from
chickens inoculated with the mixed bacterial
populations from
commercially raised birds showed a relatively
large increase in
bacterial populations having 55 to 75% G+C content
and a corresponding
decrease in the relative abundance of bacterial
populations having 30 to 55% G+C content. These data are consistent
with an increase in
bifidobacteria and propionibacteria (Fig.
3 and
4), although no
confirming analysis was performed.
Profiling of the DNA from the total GI tract bacterial community
according to its percent G+C content as described here is
totally
independent of the culturability of the component bacteria.
Further
studies employing DNA hybridization or quantitative PCR
with specific
primers would help confirm the presence and abundance
of specific
bacterial groups in the total microbial community
of the chicken GI
tract (
11,
18,
31,
33). However, these
approaches are
dependent on reference bacteria and therefore on
previous strain
isolation under laboratory conditions. Assessment
of the relative
contribution of uncultured organisms to the total
bacterial community
is an arduous task requiring extensive molecular
analyses, including
primer and probe design, PCRs, cloning, sequencing,
and nucleic acid
hybridization experiments. With the approaches
described here, we have
at hand a relatively rapid community-level
analysis which generates
data in the form of DNA abundance versus
percent G+C content. This
technique provides a window on population
dynamics in the community to
guide future experiments related
to the microbial ecology of the GI
tracts of chickens.
| |
ACKNOWLEDGMENTS |
This work was financially supported by the Cultor Corporation,
Helsinki, Finland, and Finnfeeds International, Ltd., Marlborough, United Kingdom.
We thank Andrew Morgan and Mike Bedford for useful comments and
stimulating discussions. We also thank Osmo Siikanen, Hilkka Heikkinen,
Liisa Leppä, and Esa Wainio for excellent technical assistance;
Hanna Jatila for preparing the starter cultures; and Hannele Kettunen
for animal maintenance and recovery of intestinal specimens.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Cultor
Corporation Technology Center, FIN-02460, Kantvik, Finland. Phone:
358-9-297-4684. Fax: 358-9-298-2203. E-mail:
juha.apajalahti{at}cultor.com.
| |
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