NATURE REVIEWS | GASTROENTEROLOGY & HEPATOLOGY 	 VOLUME 9  |  OCTOBER 2012  |  577
Microbiology group,
Rowett Institute of
Nutrition and Health,
University of Aberdeen,
Greenburn Road,
Bucksburn, Aberdeen
AB21 9SB, UK
(H. J. Flint, K. P. Scott,
P. Louis, S. H. Duncan).
Correspondence to:
H. J. Flint
h.flint@abdn.ac.uk
The role of the gut microbiota in nutrition
and health
Harry J. Flint, Karen P. Scott, Petra Louis and Sylvia H. Duncan
Abstract | The microbial communities that colonize different regions of the human gut influence many
aspects of health. In the healthy state, they contribute nutrients and energy to the host via the fermentation
of nondigestible dietary components in the large intestine, and a balance is maintained with the host’s
metabolism and immune system. Negative consequences, however, can include acting as sources of
inflammation and infection, involvement in gastrointestinal diseases, and possible contributions to
diabetes mellitus and obesity. Major progress has been made in defining some of the dominant members
of the microbial community in the healthy large intestine, and in identifying their roles in gut metabolism.
Furthermore, it has become clear that diet can have a major influence on microbial community composition
both in the short and long term, which should open up new possibilities for health manipulation via
diet. Achieving better definition of those dominant commensal bacteria, community profiles and system
characteristics that produce stable gut communities beneficial to health is important. The extent of
interindividual variation in microbiota composition within the population has also become apparent, and
probably influences individual responses to drug administration and dietary manipulation. This Review
considers the complex interplay between the gut microbiota, diet and health.
Flint, H. J. et al. Nat. Rev. Gastroenterol. Hepatol. 9, 577–589 (2012); published online 4 September 2012; doi:10.1038/nrgastro.2012.156
Introduction
The relationship between the mammalian host and
microorganisms that colonize the intestinal tract is
the outcome of a lengthy and complex coevolution.1
The primary imperative for the host must be to defend
against the constant threat of infection that is posed by
microorganisms in the gut. On the other hand, mammals
have gained the ability to benefit from nutrients supplied
by the resident microbiota, and the develop­ment of the
gut and of the immune system is attuned to the presence
of a complex microbiota.2,3
Research into infectious
diseases has always sought to identify single causative
agents wherever possible, in most cases with remarkable
success. Understanding the role of our gut microbiota
in nutrition and the maintenance of health, however,
represents a very different challenge that necessarily
involves different approaches.4
Certain organisms, such
as bifidobacteria and Faecalibacterium prausnitzii,5
are
considered beneficial for health, although the supporting
evidence and mechanistic basis for these benefits
remain incomplete and in some cases equivocal. At the
same time, the gut community also harbours organisms
that have the capacity for adverse effects, via their metabolic
outputs and gene products, or potential for patho-
genicity.6
The balance of benefit and harm for the host
therefore depends on the overall state of the microbial
community in terms of its distribution, diversity, species
composition and metabolic outputs (Figure 1). In this
Review, we focus mainly on the interplay between diet,
the species composition of the microbial community and
microbial metabolism in the healthy state.
The gut environment
The gut environment differs markedly between different
anatomical regions in terms of physiology, digesta
flow rates, substrate availability, host secretions, pH and
oxygen tension. The human intestinal microbiota should
therefore be viewed as a collection of semidiscrete communities.
The large intestine, which is characterized by
slow flow rates and neutral to mildly acidic pH, harbours
by far the largest microbial community (dominated by
obligate anaerobes) that will be the main subject of this
article. Important differences in gut environment occur
between proximal and distal regions, and more locally
between the gut lumen and surfaces (Box 1; Figure 2).
By comparison, the small intestine provides a more
challenging environment for microbial colonizers given
the fairly short transit times (3–5 h) and high bile con-
centrations.7,8
Molecular analysis has revealed that the
jejunal and ileal microbiota consists mainly of facultative
anaerobes, including Gram-positive streptococci,
lactobacilli and enterococci species and Gram-negative
Proteobacteria and Bacteroides.7,8
The major micro­bially
produced short-chain fatty acids (SCFAs) detected in
ileal effluents from individuals with an ileostomy were
acetate, propionate and butyrate in the molar proportions
of 20:1:4,8
compared with approximately 3:1:1 in a
typical faecal sample. Indications, however, exist that the
Competing interests
The authors declare no competing interests.
FOCUS ON GUT MICROBIOTA
© 2012 Macmillan Publishers Limited. All rights reserved
578  |  OCTOBER 2012  |  VOLUME 9 www.nature.com/nrgastro
microbiota of the intact terminal ileum might be differ-
ent9
and closer to that of the proximal colon.10
Given the
importance of the ileum as a site for interactions with
the immune system and with pathogens, having more
information on these communities is clearly desirable.
The ‘normal’ human colonic microbiota
Most of the information that is available on the composition
of the gut microbiota derives from faecal samples that
mainly reflect the community present in the lumen of the
distal large intestine. Extensive analysis of small subunit
(16S) ribosomal RNA (rRNA) sequences amplified
from faecal samples11–15
has been supplemented by data
Key points
■■ Molecular surveys have revealed remarkable diversity within the human gut
microbiota, but certain dominant species are detected in faecal samples from
most healthy adults
■■ Dietary intake, especially of nondigestible carbohydrates, alters the species
composition of the gut microbiota both in the short term and in the long term
■■ Interindividual variation in colonic microbiota composition influences responses
to dietary manipulation
■■ The gut microbiota potentially influences the host’s energy balance through
multiple mechanisms, including supplying energy from nondigestible dietary
components and influences on gut transit, energy intake and energy expenditure
■■ Whether variation in gut microbiota composition is a major factor that
influences obesity and metabolic disease in humans is not yet clear
■■ The latest research has suggested new candidate organisms among the
healthy gut microbiota that might be beneficial to gut health and new strategies
for correcting dysbiosis associated with certain disease states
from metagenomic sequencing16
to produce a broad consensus
on microbial diversity; thus, the dominant bacterial
phyla in the healthy state in humans are the Firmicutes,
Bacteroidetes and Actinobacteria, with Proteobacteria and
Verrucomicrobia also present in lower numbers.
Descriptions at a more detailed taxonomic level reveal
many hundreds of species (or ‘phylotypes’, to include
noncultured variation) in a typical faecal sample. These
findings lead us to a series of important questions: to
what extent can each individual be considered to carry
a unique collection of gut microbiota (or, conversely, is
there a ‘core’ set of gut bacteria that is common to everyone);
to what extent do samples taken from the same
individual vary in microbiota composition with time as
a result of changes in diet, environment or other influences
(for example, antibiotics); how much does gut
microbiota composition change with life stage? Ideally,
we need answers to all these questions in relation to the
microbiota of the healthy gut before addressing the question
of how microbiota changes might be associated with
disease states. Fortunately, studies in the past few years
have provided at least partial answers.
Dominant bacterial species in the colon
Despite the diversity at the level of phylotypes, it is
clear that some species are commonly detected in
high numbers in most adult faecal samples. Tap et al.12
reported 66 particularly abundant phylotypes among
17 healthy individuals; it was noted that most of the same
dominant phylotypes were common to those reported in
Normal gastrointestinal
immune function
Supply of nutrients
and energy
Cancer prevention
Inhibition of pathogens
Normal gut motility
Cardiovascular health
Obesity and
metabolic syndrome
Cancer promotion
Source of pathogens
IBD
Cardiovascular disease
IBS (constipation,
diarrhoea, bloating)
■ SCFA production, vitamin synthesis
■ Influences on energy supply, gut hormones, satiety,
energy expenditure
■ Lipopolysaccharide, inflammation
■ Butyrate production, phytochemical release
■ Toxins, carcinogens, inflammation
■ SCFA production, intestinal pH, bacteriocins,
competition for substrates and/or binding sites
■ Toxin production, tissue invasion, inflammation
■ Balance of proinflammatory versus anti-inflammatory
signals, development
■ Inflammation, immune disorders
■ Metabolites (SCFA, gases) from nondigestible carbohydrates
■ Lipid, cholesterol metabolism
Health Microbial products or activities Disease
Figure 1 | Influence of gut microbial communities on health. Most of the microbial activities indicated in the centre column
are functions of the whole community of gut microbiota rather than being attributable to a single species. The balance of the
community and its output determines the net contribution to health or disease. Abbreviation: SCFA, short-chain fatty acid.
REVIEWS
© 2012 Macmillan Publishers Limited. All rights reserved
NATURE REVIEWS | GASTROENTEROLOGY & HEPATOLOGY 	 VOLUME 9  |  OCTOBER 2012  |  579
four previously published studies. Walker et al.13
found
50 dominant phylotypes that each represented >0.5% of
total 16S rRNA sequences across six obese male individuals;
interestingly, 62% of these corresponded to cultured
species, whereas only 28% of the remaining 270
phylotypes had cultured representatives. Five of the top
10 species (Bacteroides vulgatus, Eubacterium rectale,
F. prausnitzii, Colinsella aerofaciens and Ruminococcus
bromii; Figure 3) corresponded to the top five most abundant
bacteria detected in an entirely culture-dependent
study.17
It should be no surprise that the most abundant
phylotypes have been isolated preferentially, but
this finding also suggests that the remaining microbial
diversity might not have been cultured mainly because
they are relatively less abundant and consequently occur
sporadically among the most dominant bacteria in the
community, rather than being intrinsically unculturable.
Goodman et al.18
also concluded that anaerobic culturing
in principle enabled the recovery of the majority of diversity
that was detected by sequencing at the species level in
faecal samples. Evidence, however, indicates substantial
geographical variation in dominant phylotypes,19,20
suggesting
that, for some human populations, many highly
abundant species will not have been cultured. It should
also be recognized that less abundant (or subdominant)
species can have critical roles in the microbial community.
Although much of this phylogenetic variation might
be functionally redundant,21
it could also include species
that possess unique functional properties (for example,
as ‘keystone’ species that release energy from recalcitrant
substrates, or as pathogens) and that could contribute
to major interindividual variation in health outcomes.
For practical reasons, most gut bacterial diversity
is likely to remain uncultured and descriptions of the
gut microbial communities will continue to depend
almost entirely on rapid culture-independent molecular
approaches. The arrival of high-throughput sequencing
technologies offers the alternative approach of analysing
the gene content of the community through metagenomics
(Box 2) rather than focussing on phylogenetic
groups.16,20,22
Not surprisingly, as many core functions
are conserved across species, gene content shows less
interindividual variability than phylogenetic composi-
tion.23
As the microbial cell is the basic unit of replication
and metabolism, it still is essential to understand the
coevolved collections of genes that represent individual
genomes, together with the interactions of individual
microbes with each other, and with the host.
Phylogenetic analyses have demonstrated marked
variation in the phylotypes present between individuals
within populations.13,24,25
Large-scale sequence analysis
has also suggested that the human intestinal community
might exist in a small number of discrete states or
‘enterotypes’,26,27
although the degree of discontinuity for
interindividual variation within the human microbiota
is not yet clear.28
Impact of diet upon the gut microbiota
Faecal microbiota profiles in healthy adults seem to
have substantial stability over time.29–31
The influence
of particular dietary components can, however, be seen
in carefully controlled human dietary studies. Although
many studies have documented the response of selected
groups to prebiotics,32
only a few have examined temporal
changes in the whole gut microbial community in
response to dietary change (Table 1). In one 2011 study
in which obese male volunteers were given controlled
diets differing in the type and content of nondigestible
carbohydrates for 3‑week periods, faecal microbiota profiles
tended to group by individual more than by diet.13
On the other hand, there were marked changes in the
relative abundance of several dominant phylotypes in
response to the dietary shifts, especially increased intake
of resistant starch (Figure 3). These changes occurred
within a few days, and were reversed equally quickly
by a subsequent dietary switch. The species affected
were mainly those already shown to utilize starch, but
the same species did not respond in the same manner
in all individuals. In other studies, supplementation
with galacto-­oligosaccharides or inulin was shown to
increase the relative abundance of bifidobacteria on
average, but again certain volunteers were found to be
‘non­responders’.33,34
Thus, changes in the intake of nondigestible
carbo­hydrates clearly affect faecal microbiota
composition, but these responses are not universal and
are influenced by the initial composition of an individual’s
gut microbiota. It should also be noted that many
dominant groups of bacteria, perhaps those that possess
a greater degree of nutritional diversity or flexibility,
remained unaffected by dietary change.
Box 1 | Influence of the colonic environment upon gut microbiota composition
Intestinal pH gradients
Colon pH varies from mildly acidic conditions in the proximal colon to more neutral
pH distally. Growth of Bacteroides spp. is curtailed by pH values <6.0 at short-chain
fatty acid concentrations typical of the colon (50–100 mM).89
Many Firmicutes are
more tolerant of acidic pH, giving them a competitive advantage at the low pH that
can result from active substrate fermentation. A major shift in species composition
and metabolic outputs of the human intestinal microbiota has been seen between
pH 5.5 and pH 6.5 in a continuous culture model in vitro.60
Intestinal oxygen gradients
Another factor that influences the spatial distribution of the microbiota is
oxygen.144
The colonic lumen becomes highly anaerobic (Eh of ~250 mV) largely
because facultative anaerobes consume available oxygen. Most colonic bacteria
are obligate anaerobes that fail to grow >5 × 10–3
atm oxygen, but Bacteroides
spp. can be ‘nanaerobes’; B. fragilis possesses a cytochrome bd oxidase that
allows growth in nanomolar oxygen concentrations.145
Although most colonic
Firmicutes are considered strict anaerobes that become inviable in air within
minutes,58
growth of Faecalibacterium prausnitzii is actually stimulated by very
low oxygen concentrations because of its ability to shuttle electrons to oxygen via
flavins and thiols,83
which suggests that F. prausnitzii, like B. fragilis, might exploit
niches close to the mucosa that involve some exposure to oxygen.144
Bile acids
Bile acids are derived from cholesterol in the liver and secreted as conjugated
bile acids into the small intestine, followed by deconjugation by microbial bile
salt hydrolases.146
Reabsorption in the small intestine also contributes to
gradients of bile acid concentration. Bile acids have strong antimicrobial activity;
feeding cholic acid to rats caused a major shift in gut microbiota composition
towards Firmicutes and against Bacteroidetes.147
In the large intestine, bile acids
are modified by the gut microbiota via 7‑α-dehydroxylation to form potentially
carcinogenic secondary bile acids.99
FOCUS ON GUT MICROBIOTA
© 2012 Macmillan Publishers Limited. All rights reserved
580  |  OCTOBER 2012  |  VOLUME 9 www.nature.com/nrgastro
Interestingly, a correlation was reported in 2011 by
Wu et al.27
between two enterotypes defined in 96 adults
and long-term dietary habits. Thus, a ‘Prevotella-type’
community was associated with fibre intake and a
‘Bacteroides-type’ community with high protein intake,
suggesting that enterotypes might reflect discrete patterns
of habitual dietary intake within the study population.
An earlier study by de Fillippo et al.19
detected
major differences in faecal microbiota between Italian
and African children, which they ascribed to differences
in dietary intake. 16S rRNA sequences corresponding
to Prevotella spp. were more abundant in the African
children than in the Italian children and their overall
intake of vegetable fibre was higher than in the Italian
children. The Italian children showed higher proportions
of Bacteroides spp. and Firmicutes than the African children,
together with higher intakes of starch and protein.
This finding suggests that, in addition to short-term
changes induced by dietary shifts, long-term consequences
of habitual diet upon the composition of the
gut microbiota exist; although less evidence is available,
it seems possible that such long-term changes would also
be reversible by dietary change.
Development of the gut microbiota
The composition of the microbiota changes substantially
at three stages in life: from birth to weaning; from
weaning to attaining a ‘normal’ diet; during old age.
The first bacteria to colonize the gut at birth are facultative
anaerobes;35
these bacteria in turn create anaerobic
conditions that promote the growth of obligate anaerobes
(initially Bifidobacterium and Bacteroides spp.)
within about 2 weeks. Infants born naturally become
inoculated by the mother’s vaginal and faecal microbiota
during birth,36
although those born by caesarian
section are initially colonized by bacteria from the
environment and skin.37
At 3 days, naturally delivered
newborn babies harbour a greater abundance and variety
of Bifidobacterium spp. than those born by caesarian
section.38,39
Babies that are solely breastfed until weaning
tend to have a more stable, less diverse, bacterial com-
munity,40,41
with higher proportions of bifidobacteria
than formula-fed babies.40,42,43
Studies using PCR amplification
and sequencing44
have not always reflected the
abundance of Bifidobacterium spp. reported in other
studies, although improved primers for 16S rRNA gene
amplification are now available.45
An early meta­genomic
study found that the faecal microbiota in Japanese infants
was distinctly different to that of weaned children and
adults, and that 80% of the infant sequences matched
Bifidobacterium-derived sequences.22
Some evidence
indicates geographical variation in the composition of
the gut microbiota, with bifidobacteria dominating in
northern Europe and Bacteroides spp. and lactobacilli in
Gut mucosa
Inner mucin layer
Outer mucin layer
Oxygen
Digesta Undigested
food
particles
Lumen
1
3 4
2
Starch
Plant cell
wall fibre
Figure 2 | Microbial microenvironments within the large intestine. Several microenvironments exist within the large intestine
in which microorganisms can reside: 1) epithelial surface and inner mucin layer (minimal colonization in the healthy state);
2) diffuse mucin layer (specialist colonizers, for example, Akkermansia muciniphila); 3) gut lumen–liquid phase (diverse
microbial community); and 4) gut lumen–substrate particles (specialized primary colonizers e.g. Ruminococcus spp.).
REVIEWS
© 2012 Macmillan Publishers Limited. All rights reserved
NATURE REVIEWS | GASTROENTEROLOGY & HEPATOLOGY 	 VOLUME 9  |  OCTOBER 2012  |  581
southern Europe,40
although bifidobacteria dominated in
breastfed babies both in Malawi and Finland.46
Bacterial
strains found in breast milk have also been detected in
faecal samples from the corresponding babies.47,48
These
bacteria are postulated to translocate from the mother’s
intestine to the mammary gland via the mesenteric
lymph nodes.49
The early gut microbiota in preterm
infants was reported to be dominated by facultative
anaerobes;50
this finding might be, in part, because of
the necessary medical interventions involved in preterm
birth, including the administration of antibiotics.
After the introduction of solid food, gut microbiota
composition develops towards the adult pattern with
increased diversity43,51
and increased abundance of
anaerobic Firmicutes.52
The microbiota of breastfed and
formula-fed babies converge gradually, becoming indistinguishable
by around 18 months of age,43
and resembles
that of an adult by age 3 years.20
Changes in the genetic
capacity of the microbiome with human development
include changes in the abundance of genes involved
in vitamin biosynthesis.20
Early colonization of the gut
has been shown to influence maturation of the immune
system,53
and there might be a link between aberrant gut
microbiota and atopic diseases such as eczema.54,55
A decline in microbiota diversity has been reported in
old age,56
with reduced numbers of bifidobacteria and an
increase in Enterobacteriaceae.57
Bacteroidetes become
more abundant and Firmicutes less abundant in elderly
adults (aged >65 years) compared with younger adults
(28–46 years) as controls.25
How these changes correspond
to changes in health status is not yet clear, as is
to what extent they are driven by altered dietary intake,
physical activity or altered immune function.
Microbial metabolism in the gut
The metabolic activities of gut microorganisms have
major consequences for the host that can be both beneficial
and harmful. Metabolism in anaerobic microbial
communities is highly interactive, with crossfeeding
between different organisms an important and widespread
phenomenon.58
In a few cases, it is possible
to associate particular metabolic products with one,
or a few, species, as with the conversion of oxalate by
Oxalobacter formigenes.59
The situation is generally far
more complex, with particular metabolites being produced
by many members of the microbial community
and being consumed or transformed by others. Crucially,
although the species composition of the microbiota
clearly has a role,60
it is the substrates that are available
to the microbiota that largely determine the metabolic
outputs from the community.61
As discussed earlier,
dietary substrates have a major influence on the species
composition of the microbiota, but their structures
also determine which metabolic pathways are used for
fermentation by individual bacterial species.
Dietary nondigestible polysaccharides
Much of the undigested dietary residue that arrives
in the large intestine is in the form of insoluble particles
(especially plant cell walls and resistant starch).62
Evidence from in vitro model systems suggests that specialized
groups of bacteria are involved in accessing these
structures.63
In 2012, Ze et al.64
presented evidence that
R. bromii, species belonging to the Firmicutes, might
act as a key primary degrader of resistant starch particles
in the human colon, making the substrate available
to other amylolytic bacteria. Analysis of faecal samples
from healthy volunteers had previously revealed a markedly
higher proportion of Ruminococcaceae sequences
associ­ated with the particulate fraction (12.2%) than with
the liquid fraction (3.3%),65
whereas the Gram-negative
Bacteroides sequences tended to partition more with the
liquid phase. This finding suggests that certain bacteria
are preferentially associated with insoluble digesta in
the gut and might represent specialist primary degraders
of these substrates. Although many human colonic
Bacteroides spp. are found to possess large genomes
extremely rich in diverse glycoside hydrolase genes,66
these bacteria seem better equipped to utilize soluble
rather than insoluble carbohydrates.67
The parts played by
different human gut bacteria in carbohydrate breakdown
are only now beginning to be understood.68
Utilization of host-derived substrates
Mucin provides a protective barrier for the gut epithelium,
but is also a potential growth substrate for
intestinal bacteria.69
The specialized mucin-degrader
Akkermansia muciniphila is an important member of the
healthy colonic microbiota that has been found to modulate
immune responses in a mouse model.69,70
Other
colonic species, notably Bacteroides spp., have the ability
to utilize a variety of host-derived glycans.66,68
Short-chain fatty acid metabolism
Under the anaerobic conditions of the large intestine,
undigested carbohydrates are fermented mainly to SCFAs
M NSP
Total16SrRNAsequences(%)
RS WL
Bacteroides vulgatus
Colinsella aerofaciens*
Clostridium clostridioforme
Anaerostipes hadrus
Eubacterium hallii
Eubacterium rectale‡
Ruminococcus bromii‡
Faecalibacterium prausnitzii
Diet type
0
30
25
20
15
10
5
35
Figure 3 | Influence of diet upon dominant human colonic bacteria determined by
16S rRNA gene sequencing. The graph shown is based on data from means for six
obese male volunteers in a controlled dietary trial reported by Walker et al.13
Weight
maintenance diets were M (control); NSP, high in wheat bran (3 weeks); RS, high in
type 3 resistant starch (3 weeks); and WL, high in protein and reduced in
carbohydrates (3 weeks). Data are from faecal samples analysed at the end of each
dietary period. Statistically significant differences in the percentage abundance of a
given species compared with the other diets are indicated (*P <0.05, ‡
P <0.001).
The eight most abundant bacterial species shown here together accounted for 29%
of the total 16S rRNA sequences detected in these samples. Abbreviations: M,
maintenance; NSP, nonstarch polysaccharide; RS, resistant starch; WL, weight loss.
FOCUS ON GUT MICROBIOTA
© 2012 Macmillan Publishers Limited. All rights reserved
582  |  OCTOBER 2012  |  VOLUME 9 www.nature.com/nrgastro
(such as butyrate and acetate) and gases (hydrogen,
carbon dioxide, methane and hydrogen sulphide). SCFAs
have multiple effects on the host, as the major anions
in the colon and as energy sources for the host, with
butyrate being consumed mainly by the colonic epithelium
and acetate becoming available systemically.71
It has
also been recognized that SCFAs signal to the gut receptors
free fatty acid receptor 2 (FFAR2, formerly known as
GPR43) and free fatty acid receptor 3 (FFAR3, formerly
known as GPR41).72
These receptors are involved in
controlling anorectic hormones—including peptide YY
(PYY) and glucagon-like peptide 1 (GLP1)—that have
roles in appetite control, thus providing a potential link
between microbial SCFA formation and food intake.72
Other reported influences include anti­cancer effects
(especially for butyrate), anti-inflammatory proper-
ties73,74
and changes in gut motility75,76
and energy
expenditure.77
Therefore, changes in the relative production
rates of the major SCFAs by the colonic microbiota
are likely to have important physio­logical consequences.
Considerable progress has been made in defining
the dominant groups of bacteria that have key roles
in anaerobic metabolism on the basis of cultured isolates,
16S rRNA-based molecular detection and new
approaches targeted at particular functionally relevant
genes61
(Box 2). Two butyrate-producing Firmicutes
F. prausnitzii and Eubacterium rectale, for example,
are among the most abundant bacteria in the healthy
colonic community.13,21
Although these two species use
similar routes for butyrate synthesis, relying on butyryl
CoA:acetate CoA transferase,78
evidence indicates that
they have distinct ecological niches. E. rectale and the
closely related Roseburia spp. are flagellated bacteria with
the ability to utilize a range of dietary polysaccharides,
especially starch.79–81
F. prausnitzii is nonflagellated and
fails to utilize many dietary polysaccharides, including
starch.82
Furthermore, the stimulation of F. prausnitzii by
low concentrations of oxygen (Box 1) was not observed
for the close E. rectale relative Roseburia inulinivorans.83
In human volunteer trials, weight-loss diets low in total
carbohydrate have been shown to decrease the percentage
of butyrate among faecal SCFAs,84,85
correlating
with decreased populations of the butyrate-producing
Roseburia plus E. rectale group.84,86
By contrast, F. prausnitzii
showed little change in its representation among
the faecal microbiota with low-carbohydrate diets.13,84
Faecal butyrate concentrations have been shown to
increase with total SCFA concentrations under conditions
of more rapid gut transit,75,87,88
an effect that might
be mediated partly via the influence of pH on the gut
community60,89
(Box 1).
Acids such as lactate, succinate and formate normally
behave as intermediates in microbial metabolism in the
gut because of onward conversion (Figure 4). Certain
Firmicutes that are dominant in the healthy colon,
Eubacterium hallii and Anaerostipes spp., have the ability
to convert lactate and acetate into butyrate.90
E. hallii
numbers were shown to increase in faecal slurries in
the presence of lactate in vitro,91
and substantial flow of
label has been noted from 13
C lactate to butyrate in the
mixed community in isotope labelling studies to monitor
SCFA production.92–94
Label was also found in propionate
in these studies, assumed to be due to conversion of
lactate to propionate by members of the Veillonellaceae.
Given the widespread formation of lactate and its low
pKa (acid dissociation constant), the activities of these
lactate-utilizing bacteria are likely to have an important
role in maintaining homeostasis within the community.
Lactate can accumulate under conditions of disturbance
(or dysbiosis)—for example in severe colitis95
—probably
due to the curtailment of the growth of lactate-utilizing
bacteria at reduced pH.91
Less widely recognized is that
acetate, although almost invariably reaching the highest
concentration among faecal SCFAs, is also an intermediate
that is consumed by the major butyrate-producing
bacteria78
(Figure 4).
Although most sugars are fermented by common or
converging pathways to yield SCFAs,96
some require
alternative routes. The deoxyhexose sugars fucose and
rhamnose, for example, are fermented by some intestinal
bacteria via propanediol to yield propionate and
propanol.97
Propionate production from hexose sugars
is thought to be mainly associated with the Bacteroidetes
and the Veillonellaceae (Firmicutes); most seem to
use the succinate pathway for propionate formation,
with only a few bacterial species known to use the
acrylate pathway.61
Fermentation of amino acids derived from dietary or
host-derived proteins yields a much wider range of prod-
ucts.98
Faecal branched-chain fatty acids are indicative of
fermentation of branched-chain amino acids and their
faecal concentrations increase on high-protein diets.86
Box 2 | ‘Meta-omics’ analysis of gut microbial communities
High-throughput metagenomic sequencing potentially provides information on the
full complement of functional genes in the microbial community, in contrast with
the phylogenetic information that is obtained from amplification of ribosomal RNA
genes.16
The currently available technologies can already yield several gigabases of
sequence per sample, and throughput is expected to increase further. Rather less
attention has been paid to metagenomic approaches that target specific functional
genes, although these enable in-depth analysis of functional groups within the
microbiota at lower cost and without the need for massive bioinformatic capability.
Phylotypes detected using such a targeted approach to amplify β‑glucuronidase
genes from human faecal DNA were found to correlate well with a large
metagenomic dataset.108
Another example of this approach targeted the butyrylCoA:acetate
CoA-transferase gene for butyrate formation;21
88% of sequences
from 12 healthy volunteers were closely related to 12 cultured isolates, whilst
only 12% belonged to novel uncultured phylotypes, suggesting that the dominant
butyrate-producing bacteria are well represented by cultured species. Degenerate
primers have also been developed for functional genes of hydrogenotrophic
microbial groups, acetogens, sulphate-reducing bacteria and methanogenic
Archaea148–150
and for genes involved in xylan breakdown.151
There is a clear need
for more functional analysis, however, as a major proportion of genes identified
during nontargeted metagenomic analyses remain of unknown function. Screening
of metagenomic libraries provides one approach for detecting novel genes with
specific functions of interest,152
but the availability of genome sequences for
cultured isolates of human gut bacteria that enable functions to be confirmed—for
example, by gene knockouts—remains crucial. ‘–Omics’ approaches can also
be used to reveal those gene products that are most highly expressed within
particular gut environments, either at the RNA (metatranscriptomic) or protein
(metaproteomic) level.153
Metatranscriptomics has been applied to patients with an
ileostomy to gain information on the small intestinal community.8
REVIEWS
© 2012 Macmillan Publishers Limited. All rights reserved
NATURE REVIEWS | GASTROENTEROLOGY & HEPATOLOGY 	 VOLUME 9  |  OCTOBER 2012  |  583
Potentially toxic or carcinogenic products of protein
fermentation include N‑nitroso compounds, amines
and cresol.99
Hydrogen disposal
Hydrogen has a key role in anaerobic ecosystems. Its
disposal reduces the levels of gaseous compounds produced
in the colon and also affects the metabolism of
hydrogen-producing fermentative bacteria by enabling
a shift in the relative production of different fermentation
products.100
Which hydrogenotrophic microbial
group dominates potentially also has important health
effects. Hydrogen sulphide generated by sulphatereducing
bacteria (SRB) is generally regarded as a toxic
product,101
although it is also produced by the human
body and influences host functions, including promoting
the healing of ulcers and anti-inflammatory
effects.102
Methanogenesis is associated with a slower
transit time,87
which might reflect the slow growth of
methanogenic Archaea that use hydrogen and carbon
dioxide, or formate, to form methane. Intriguingly, evidence
exists that methane might actively be involved in
prolonging intestinal transit.103
Acetogenic bacteria can
produce acetate fromhydrogenand carbon dioxide, or
formate, as well as from carbohydrates.104
A 2012 study investigating human biopsy samples
from the left and right side of the colon and the rectum
of 25 healthy individuals found that, using PCR primers
that target functional genes, all individuals carried all
three groups of hydrogenotrophs, with methanogenic
Archaea comprising around 50% of hydrogen utili­zers
in each of the three regions of the colon. SRB were more
abundant than acetogens in the right colon and acetogens
more abundant in the left colon and rectum.105
SRB are able to use lactate as a cometabolite to produce
hydrogen sulphide and acetate106
(Figure 4).
Metabolism of phytochemicals and xenobiotics
Colonic bacteria are also involved in the release and
transformation of a wide range of non-nutrient, but
potentially bioactive, compounds of plant origin, including
a wide variety of aromatic compounds.107
Some of
these derive from the degradation of plant-cell-wall
structures. For example, ferulic acid, an important
component of cereal bran, is largely converted to 4‑OH
phenyl­propionic acid in faecal samples.86
Other aromatic
Table 1 | Diet-driven changes in gut microbial community composition in humans*
Dietary intervention Duration
(weeks)‡
Volunteers§
Molecular profiling
methods (16S rRNA)
Bacterial changes detected Reference
Controlled diet composition
Resistant starch (RS3) 3 14 obese, M Sequencing; qPCR Ruminococcus bromii, Eubacterium rectale,
Roseburia spp. and Oscillibacter spp.
Walker et al. (2011)13
Nonstarch polysaccharides
(wheat bran)
3 14 obese, M Sequencing; qPCR No major changes Walker et al. (2011)13
Weight-loss diet||
3 14 obese, M Sequencing; qPCR Collinsella aerofaciens, E. rectale
and Roseburia spp.
Walker et al. (2011)13
Weight-loss diets||
4 18 obese, M FISH E. rectale, Roseburia spp.
and Bifidobacterium spp.
Duncan et al. (2007)84
Weight-loss diets||
4 17 obese, M FISH E. rectale, Roseburia and
Bifidobacterium spp.
Russell et al. (2011)86
Dietary supplementation
Resistant starch (RS2) 3 10 healthy Sequencing; qPCR R. bromii and E. rectale Martínez et al. (2010)154
Resistant starch (RS4) 3 10 healthy Sequencing; qPCR Bifidobacterium spp. and Parabacteroides
distasonis
Martínez et al. (2010)154
Resistant starch
(Hi Maize)
4 46 healthy DGGE; qPCR R. bromii Abell et al. (2008)155
Inulin and oligofructose 2.3 12 healthy qPCR Faecalibacterium prausnitzii
and Bifidobacterium spp.
Ramirez-Farias et al.
(2009)34
Inulin (long chain) 3 31 healthy FISH Bifidobacterium spp., Lactobacilli spp.
and Atopobium spp.
Bacteroides spp. and/or Prevotella spp.
Costabile et al. (2010)156
Inulin 2 30 healthy FISH Bifidobacterium spp.
Bacteroides and/or Prevotella and
Clostridium histolyticum
Kleessen et al. (2007)157
Galacto-oligosaccharides 3 18 healthy Sequencing F. prausnitzii and Bifidobacterium spp.
 Bacteroides
Davis et al. (2011)33
Raffinose 3 12 healthy Sequencing; qPCR F. prausnitzii, Bifidobacterium spp. Fernando et al. (2010)158
*Recent studies (within previous 5 years) that have attempted to analyse the whole gut microbe community (from faecal samples) and provide detailed information on dietary intake are shown.
Many additional studies have shown stimulation of specific groups, especially bifidobacteria (reviewed32
). ‡
Most studies employed a crossover design comparing the influence of different diets
within individuals. Duration refers to a single dietary period. §
Adults of both sexes unless otherwise stated (M = males only). ||
High protein, reduced levels of carbohydrates. Abbreviations:
DGGE, denaturing gradient gel electrophoresis; FISH, fluorescence in situ hybridization; qPCR, quantitative PCR.
FOCUS ON GUT MICROBIOTA
© 2012 Macmillan Publishers Limited. All rights reserved
584  |  OCTOBER 2012  |  VOLUME 9 www.nature.com/nrgastro
compounds derive from hydrolysis of soluble glycoside
conjugates present in the plant.107
Many plant compounds, and also drugs, are treated as
xenobiotics and conjugated to form glucuronides in the
liver, then to be released into the gut; these glucuronides
are subject to hydrolysis by microbial β‑glucuronidase,
releasing the original compound. A targeted analysis
has shown that a fairly small number of bacterial
phylo­types account for most copies of the bacterial gus
(β-glucuronidase) gene found present in the human
colon.108
Thus, variation in the populations of these few
species would be expected to result in considerable interindividual
variation in gus activity and the cleavage of
glucuronide conjugates in the gut. Although a possible
second gene responsible for β‑glucuronidase activity has
been identified in human gut bacteria,109
its contribution
is not yet clear.108
A wide range of metabolites formed
by microbial activity in the gut can be detected in the
bloodstream, and a number of these metabolites have
potential as biomarkers of health or disease.110
Energy, obesity and metabolic health
As noted above, absorption of microbially produced
SCFAs provides energy to the host from dietary components
that have remained undigested in the small
intestine. Gut microbes, therefore, contribute to the
‘energy harvest’ from the diet,111
and this contribution
might be vital under conditions of food scarcity. The
microbial contribution to the host’s energy supply will
depend on many factors, including the nondigestible
carbohydrate content of the diet and upon gut transit,
which affects SCFA absorption and the extent of digestion
and fermentation of dietary carbohydrates.112,113
The species composition of the gut microbiota also has
the potential to influence energy harvest; for example,
through variation in keystone species responsible for
the breakdown of recalcitrant substrates64
or the proportions
of different metabolic groups involved in
forming SCFAs or gaseous products. The calorific value
per mole of nondigestible carbohydrate is considerably
less than for a fully digestible carbohydrate (Figure 5)
and clearly depends on the extent of fermentation and
of SCFA absorption.114
This finding means that directly
replacing digestible carbohydrate by nondigestible
carbo­hydrate in the diet should reduce the net delivery
of calories to the host, assuming equal intake. Some evidence
indicates that dietary nondigestible carbohydrate
might contribute to satiety.115
Much speculation has been made over the possible
contribution of gut bacteria to obesity in humans, initially
focussing on the possible influence of microbiota
composition on energy harvest. The balance of evidence
does not show a consistent phylum level shift in microbiota
composition in obese humans.13,116–118
Although a
lower Bacteroides:Firmicutes ratio was reported in obese
individuals in one human study and in ob/ob mice,24,111
other human studies have reported the opposite result,116
or no difference.117
More subtle changes might occur in
Carbohydrate fermentors
■ Bacteroidetes
■ Firmicutes
■ Actinobacteria
Propionate producers
■ Bacteroidetes
■ Veillonellaceae
Carbohydrates
(hexoses)
Pyruvate
Butyryl CoA
Butyrate
Propionate
Lactate
Acetate
Succinate
Formate/H2 +CO2
SO4
H2S
CH4
Methanogens
■ Methanobrevibacter
smithii (Archaea)
Acetogens
■ Blautia
hydrogenotrophica
■ Marvinbryantia
formatexigenes
(Firmicutes)
Sulphate reducers
■ Desulfovibrio piger
(Proteobacteria)
Butyrate producers
■ Faecalibacterium
prausnitzii
■ Eubacterium rectale,
Roseburia spp.
■ Eubacterium hallii
(Firmicutes)
■ Anaerostipes spp.
Acetyl CoA
Figure 4 | Functional and phylogenetic groups of gut bacteria involved in the metabolism of short-chain fatty acids. Figure
shows a schematic of the gut microbiota involved in the metabolism of short-chain fatty acids. Acetate and lactate are
shown as intermediates. Representative species and phyla are indicated based on information from cultured
microorganisms. Whereas most colonic bacteria use the Embden–Meyerfhof pathway for hexose metabolism,
bifidobacteria (Actinobacteria) use the bifid shunt pathway (not shown here).96
REVIEWS
© 2012 Macmillan Publishers Limited. All rights reserved
NATURE REVIEWS | GASTROENTEROLOGY & HEPATOLOGY 	 VOLUME 9  |  OCTOBER 2012  |  585
obese individuals at the species level, but these changes
could be the result either of different dietary habits84
or
altered host physiology. Changes in faecal microbiota
profile (towards Bacteroidetes) have been reported for
individuals with type 2 diabetes mellitus.119
Small animal studies continue to suggest intriguing,
but complex, links between gut microbiota composition
and adiposity.120
Germ-free rodents can show markedly
greater, or lesser, gains in adiposity and body weight than
conventional animals when fed high-fat diets.121,122
These
different outcomes depend on the exact composition of
the diet supplied and seem to correlate with influences
on energy expenditure more than energy harvest (these
diets in any case contain little fibre).122
A number of
studies have demonstrated a major influence of highfat
diets on microbiota composition in rodents.123,124
Nevertheless, a series of studies involving transfer of
gut microbiota from obese animals to germ-free lean
animals have suggested that the composition of the gut
microbiota can influence adiposity, with concomitant
changes in either energy harvest, energy intake or energy
expenditure.111,125,126
Evidence has also been obtained
from small animal studies that increased passage of
bacterial lipopolysaccharide (LPS) into the bloodstream
occurs during consumption of high-fat diets. It has been
proposed that this increase in LPS might be involved in
the development of insulin resistance that is triggered
by high-fat diets.127
Proinflammatory LPS is assumed to
originate mainly from Gram-negative Proteobacteria in
the small intestine.
Intestinal health
Substantial evidence exists for modification of the
faecal and colonic microbiota in certain forms of IBD,
especially ileal Crohn’s disease,128,129
which are discussed
in detail elsewhere in this Focus issue.130
Some
Firmicutes, notably F. prausnitzii, are reported to show
decreased representation in patients with ileal Crohn’s
disease. Furthermore, patients with low F. prausnitzii
abundance had a greater likelihood of relapse following
surgical resection than those with higher abundance.5
Combined with evidence that F. prausnitzii produces an
anti-­inflammatory product, this finding has led to strong
interest in this organism as a potentially beneficial component
of the healthy gut microbiota. It has also been
noted that patients can recover despite having very low
numbers of F. prausnitzii,131
and it remains to be established
whether F. prausnitzii is simply an indicator of
the gut environment or an agent that actively influences
gut health. The contribution of potentially pathogenic
Proteobacteria that are suspected to have a key role in
causation of IBD has been reviewed in detail.132
In the case of IBS, several studies have found that
imbalance in the gut microbiota can be detected in dominant
bacterial ribotypes133
or in functional groups.134
As yet, no clear consensus exists on the changes that
occur in different forms of IBS or their clinical significance
in terms of aetiology.135
In general, it can be difficult
to disassociate the effects of the active disease state
and of treatment regimes upon the microbiota from
compositional changes that might be causative or protective.
Studies that include patients at first presentation
or in remission can, therefore, provide valuable insights.
The gut microbiota is considered to have an important
role in the prevention of sporadic colorectal cancer
through the production of butyrate and the transformation
of certain dietary phenolics.99
On the other hand,
cancer-promoting compounds can also be generated by
microbial activity, and the balance of procarcinogenic
and anticarcinogenic actions is highly dependent on
diet and xenobiotic intake.136
Bacterial changes have
been noted between patients with cancer and healthy
control groups; a 2012 study based on next-generation
sequencing found, amongst other changes, a decrease in
butyrate-producers in patients with colorectal cancer,137
but, again, the causal relationship between microbiota
profile and cancer development remains unclear.
Some gut pathogens seem able to modify host
responses and the gut environment—so as to favour
their own proliferation138
—by producing major changes
(dysbiosis) in the resident gut microbial community.
Little is known about how gut microbial communities
recover following episodes of diarrhoea, or indeed after
anti­biotic treatment.139
Future perspectives
The latest reports indicate that replacement of disturbed
gut microbiota by faecal microbiota from a healthy
indivi­dual can be a highly successful approach to treating
Clostridium difficile infection.140–143
This approach is
also being considered for other disease states in which
the causation is less well understood, including gut disorders
such as IBD and IBS, and autoimmune diseases.143
In future, the idea of developing a restorative ‘cocktail’
of beneficial bacteria normally present in the healthy
colonic microbiota might look increasingly attractive as
High
Energy
content
Low
Carbon dioxide + water
Energy to the host
(small intestine)
Fermentation—
energy for
bacterial growth
Energy to the host
(large intestine)
Organic acids
(acetic, propionic, butyric)
Anaerobic
Aerobic
Nondigestible
carbohydrates
■ Energy to the host:
0–2.5kcal/g*
Digestible
carbohydrates
■ Energy to the host:
3.9kcal/g
Figure 5 | Contribution of ingested carbohydrates to dietary energy supply to the
host. Fermentation of dietary substrates by anaerobic microorganisms in the large
intestine enables the recovery of only a fraction of the initial energy content for
microbial growth. This step allows the host to absorb and oxidize the SCFAs that
are produced as microbial fermentation products. The energy yield to the host from
nondigestible carbohydrates through this route will vary depending on the
efficiencies of fermentation and of absorption of the SCFA products. *Estimates
from Roberfroid study.114
Abbreviation: SCFA, short-chain fatty acid.
FOCUS ON GUT MICROBIOTA
© 2012 Macmillan Publishers Limited. All rights reserved
586  |  OCTOBER 2012  |  VOLUME 9 www.nature.com/nrgastro
an alternative to using somewhat undefined faecal preparations.
Furthermore, evidence showing that the composition
of the gut microbiota responds to diet indicates that
prebiotic approaches for delivering health benefits can be
made more targeted and effective. These goals will require
a far more detailed characterization of key members of
the healthy gut community and their interactions with
each other, and with the host, than is currently available.
Such information would also assist greatly in interpreting
the large metagenomic datasets that are currently being
produced with the aim of understanding the role of the
gut microbiota in the aetiology of human diseases. The
combination of gut microbiology, gastroenterology and
epidemiology with developments in the rapid analysis
of metabolites, microbial markers and molecular signals
promises exciting progress in the coming years.
Conclusions
Molecular analyses have revealed remarkable diversity
within the colonic microbiota of adult humans. Although
certain dominant bacterial species are detected in most
healthy adults, there is also substantial interindividual
variation in microbial community composition. Dietary
intake determines the metabolic outputs of the microbial
community at the same time as modifying the
species composition. Diet, therefore, offers a potential
route to delivering health benefits through manipulation
of the microbial community. It remains challenging,
however, to define those states of the community
that are most beneficial to health and those that pose a
long-term risk to health. Achieving this feat will depend
both on more detailed functional analysis of representative
cultured species, and on information from new
‘‑omics’ approaches that examine shifts in overall gene
complement and expression within the community.
Review criteria
Multiple literature searches were made using Web of
Science and Scopus (1950–2012) using as search
terms: “gut bacteria/microbiota/microflora/flora” AND,
for example, “bile, phytochemicals, metabolites, obesity
OR infant”. The broad scope of this article, together
with limitations on length, however meant that not all
papers could be cited. The resulting selection inevitably
represents the authors’ own assessment of relevance to
the topic, with some bias towards recent papers. Where
recent comprehensive reviews were already available on
topics of current interest (for example, bacteriotherapy,
microbiota involvement in IBD and IBS) we have provided
these rather than reporting the primary literature.
1.	 Ley, R. E., Lozupone, C. A., Hamady, M.,
Knight, R. & Gordon, J. I. Worlds within worlds:
Evolution of the vertebrate gut microbiota.
Nat. Rev. Microbiol. 6, 776–788 (2008).
2.	 Sekirov, I., Russell, S. L., Antunes, L. C. &
Finlay, B. B. Gut microbiota in health and
disease. Physiol. Rev. 90, 859–904 (2010).
3.	 Hooper, L. V. & MacPherson, A. J. Immune
adaptations that maintain homeostasis with the
intestinal microbiota. Nat. Rev. Immunol. 10,
159–169 (2010).
4.	 Cho, I. & Blaser, M. J. The human microbiome:
at the interface of health and disease. Nat. Rev.
Genet. 13, 260–270 (2012).
5.	 Sokol, H. et al. Faecalibacterium prausnitzii
is an anti-inflammatory commensal bacterium
identified by gut microbiota analysis of Crohn
disease patients. Proc. Natl Acad. Sci. USA 105,
16731–16736 (2008).
6.	 Blaser, M. J. & Kirschner, D. The equilibria that
allow bacterial persistence in human hosts.
Nature 449, 843–849 (2007).
7.	 Booijink, C. C. G. M. et al. High temporal and
inter-individual variation detected in the human
ileal microbiota. Environ. Microbiol. 12,
3213–3227 (2010).
8.	 Zoetendal, E. G. et al. The human small
intestinal microbiota is driven by rapid uptake
and conversion of simple carbohydrates. ISME J.
6, 1415–1426 (2012).
9.	 Hartman, A. L. et al. Human gut microbiome
adopts an alternative state following small bowel
transplantation. Proc. Natl Acad. Sci. USA 106,
17187–17192 (2009).
10.	 Wang, M., Ahrné, S., Jeppsson, B. & Molin, G.
Comparison of bacterial diversity along the
human intestinal tract by direct cloning and
sequencing of 16S rRNA genes. FEMS Microbiol.
Ecol. 54, 219–231 (2005).
11.	 Eckburg, P. B. et al. Microbiology: diversity of the
human intestinal microbial flora. Science 308,
1635–1638 (2005).
12.	 Tap, J. et al. Towards the human intestinal
microbiota phylogenetic core. Environ. Microbiol.
11, 2574–2584 (2009).
13.	 Walker, A. W. et al. Dominant and diet-responsive
groups of bacteria within the human colonic
microbiota. ISME J. 5, 220–230 (2011).
14.	 Suau, A. et al. Direct analysis of genes encoding
16S rRNA from complex communities reveals
many novel molecular species within the human
gut. Appl. Environ. Microbiol. 65, 4799–4807
(1999).
15.	 Hold, G. L., Pryde, S. E., Russell, V. J., Furrie, E.
& Flint, H. J. Assessment of microbial diversity in
human colonic samples by 16S rDNA sequence
analysis. FEMS Microbiol. Ecol. 39, 33–39
(2002).
16.	 Qin, J. et al. A human gut microbial gene
catalogue established by metagenomic
sequencing. Nature 464, 59–65 (2010).
17.	 Moore, W. E. C. & Moore, L. H. Intestinal floras of
populations that have a high risk of colon cancer.
Appl. Environ. Microbiol. 61, 3202–3207 (1995).
18.	 Goodman, A. L. et al. Extensive personal human
gut microbiota culture collections characterized
and manipulated in gnotobiotic mice. Proc. Natl
Acad. Sci. USA 108, 6252–6257 (2011).
19.	 De Filippo, C. et al. Impact of diet in shaping gut
microbiota revealed by a comparative study in
children from Europe and rural Africa. Proc. Natl
Acad. Sci. USA 107, 14691–14696 (2010).
20.	 Yatsunenko, T. et al. Human gut microbiome
viewed across age and geography. Nature 486,
222–227 (2012).
21.	 Louis, P., Young, P., Holtrop, G. & Flint, H. J.
Diversity of human colonic butyrate-producing
bacteria revealed by analysis of the butyrylCoA:acetate
CoA-transferase gene. Environ.
Microbiol. 12, 304–314 (2010).
22.	 Kurokawa, K. et al. Comparative metagenomics
revealed commonly enriched gene sets in
human gut microbiomes. DNA Res. 14, 169–181
(2007).
23.	 Gill, S. R. et al. Metagenomic analysis of the
human distal gut microbiome. Science 312,
1355–1359 (2006).
24.	 Ley, R. E., Turnbaugh, P. J., Klein, S. &
Gordon, J. I. Microbial ecology: human gut
microbes associated with obesity. Nature 444,
1022–1023 (2006).
25.	 Claesson, M. J. et al. Composition, variability,
and temporal stability of the intestinal
microbiota of the elderly. Proc. Natl Acad. Sci.
USA 108, 4586–4591 (2011).
26.	 Arumugam, M. et al. Enterotypes of the human
gut microbiome. Nature 473, 174–180 (2011).
27.	 Wu, G. D. et al. Linking long-term dietary patterns
with gut microbial enterotypes. Science 334,
105–108 (2011).
28.	 Huse, S. M., Ye, Y., Zhou, Y. & Fodor, A. A. A core
human microbiome as viewed through 16S rRNA
sequence clusters. PLoS ONE 7, e34242 (2012).
29.	 Franks, A. H. et al. Variations of bacterial
populations in human feces measured by
fluorescent in situ hybridization with groupspecific
16S rRNA-targeted oligonucleotide
probes. Appl. Environ. Microbiol. 64, 3336–3345
(1998).
30.	 Zoetendal, E. G., Akkermans, A. D. L. &
De Vos, W. M. Temperature gradient gel
electrophoresis analysis of 16S rRNA from
human fecal samples reveals stable and hostspecific
communities of active bacteria. Appl.
Environ. Microbiol. 64, 3854–3859 (1998).
31.	 Costello, E. K. et al. Bacterial community
variation in human body habitats across space
and time. Science 326, 1694–1697 (2009).
32.	 Roberfroid, M. et al. Prebiotic effects: Metabolic
and health benefits. Br. J. Nutr. 104, S1–S63
(2010).
33.	 Davis, L. M. G., Martínez, I., Walter, J., Goin, C. &
Hutkins, R. W. Barcoded pyrosequencing reveals
that consumption of galactooligosaccharides
results in a highly specific bifidogenic response
in humans. PLoS ONE 6, e252000 (2011).
REVIEWS
© 2012 Macmillan Publishers Limited. All rights reserved
NATURE REVIEWS | GASTROENTEROLOGY & HEPATOLOGY 	 VOLUME 9  |  OCTOBER 2012  |  587
34.	 Ramirez-Farias, C. et al. Effect of inulin on the
human gut microbiota: stimulation of
Bifidobacterium adolescentis and
Faecalibacterium prausnitzii. Br. J. Nutr. 101,
541–550 (2009).
35.	 Eggesbø, M. et al. Development of gut
microbiota in infants not exposed to medical
interventions. APMIS 119, 17–35 (2011).
36.	 Karlsson, C. L. J., Molin, G., Cilio, C. M. &
Ahrné, S. The pioneer gut microbiota in human
neonates vaginally born at term‑A pilot study.
Pediatr. Res. 70, 282–286 (2011).
37.	 Dominguez-Bello, M. G. et al. Delivery mode
shapes the acquisition and structure of the
initial microbiota across multiple body habitats
in newborns. Proc. Natl Acad. Sci. USA 107,
11971–11975 (2010).
38.	 Biasucci, G. et al. Mode of delivery affects the
bacterial community in the newborn gut. Early
Hum. Dev. 86 (Suppl. 1), 13–15 (2010).
39.	 Huurre, A. et al. Mode of delivery—effects on gut
microbiota and humoral immunity. Neonatology
93, 236–240 (2008).
40.	 Fallani, M. et al. Intestinal microbiota of
6‑week‑old infants across Europe: Geographic
influence beyond delivery mode, breast-feeding,
and antibiotics. J. Pediatr. Gastroenterol. Nutr.
51, 77–84 (2010).
41.	 Klaassens, E. S. et al. Mixed-species genomic
microarray analysis of fecal samples reveals
differential transcriptional responses of
bifidobacteria in breast- and formula-fed infants.
Appl. Environ. Microbiol. 75, 2668–2676 (2009).
42.	 Harmsen, H. J. M. et al. Analysis of intestinal
flora development in breast-fed and formula-fed
infants by using molecular identification and
detection methods. J. Pediatr. Gastroenterol.
Nutr. 30, 61–67 (2000).
43.	 Roger, L. C. & McCartney, A. L. Longitudinal
investigation of the faecal microbiota of healthy
full-term infants using fluorescence in situ
hybridization and denaturing gradient gel
electrophoresis. Microbiology 156, 3317–3328
(2010).
44.	 Palmer, C., Bik, E. M., DiGiulio, D. B.,
Relman, D. A. & Brown, P. O. Development of the
human infant intestinal microbiota. PLoS Biol. 5,
e177 (2007).
45.	 Sim, K. et al. Improved detection of
bifidobacteria with optimised 16S rRNA-gene
based pyrosequencing. PLoS ONE 7, e32543
(2012).
46.	 Grzes´kowiak, Ł. et al. Distinct gut microbiota in
South Eastern African and Northern European
infants. J. Pediatr. Gastroenterol. Nutr. 54,
812–816 (2012).
47.	 Solís, G., de los Reyes-Gavilan, C. G.,
Fernández, N., Margolles, A. & Gueimonde, M.
Establishment and development of lactic acid
bacteria and bifidobacteria microbiota in breastmilk
and the infant gut. Anaerobe 16, 307–310
(2010).
48.	 Martín, R. et al. Isolation of bifidobacteria from
breast milk and assessment of the
bifidobacterial population by PCR-denaturing
gradient gel electrophoresis and quantitative
real-time PCR. Appl. Environ. Microbiol. 75,
965–969 (2009).
49.	 Perez, P. F. et al. Bacterial imprinting of the
neonatal immune system: lessons from
maternal cells? Pediatrics 119, e724–e732
(2007).
50.	 Magne, F. et al. Low species diversity and high
interindividual variability in faeces of preterm
infants as revealed by sequences of 16S rRNA
genes and PCR-temporal temperature gradient
gel electrophoresis profiles. FEMS Microbiol.
Ecol. 57, 128–138 (2006).
51.	 Favier, C. F., Vaughan, E. E., De Vos, W. M. &
Akkermans, A. D. L. Molecular monitoring of
succession of bacterial communities in human
neonates. Appl. Environ. Microbiol. 68, 219–226
(2002).
52.	 Fallani, M. et al. Determinants of the human
infant intestinal microbiota after the introduction
of first complementary foods in infant samples
from five European centres. Microbiology 157,
1385–1392 (2011).
53.	 Martin, R. et al. Early life: gut microbiota and
immune development in infancy. Benef. Microbes
1, 367–382 (2010).
54.	 Penders, J. et al. Gut microbiota composition
and development of atopic manifestations in
infancy: The KOALA birth cohort study. Gut 56,
661–667 (2007).
55.	 Kalliomäki, M. Pandemic of atopic diseases—
a lack of microbial exposure in early infancy?
Med. Chem. Rev. Online 2, 299–302 (2005).
56.	 O’Toole, P. W. & Claesson, M. J. Gut microbiota:
Changes throughout the lifespan from infancy to
elderly. Int. Dairy J. 20, 281–291 (2010).
57.	 Woodmansey, E. J. Intestinal bacteria and
ageing. J. Appl. Microbiol. 102, 1178–1186
(2007).
58.	 Flint, H. J., Duncan, S. H., Scott, K. P. & Louis, P.
Interactions and competition within the microbial
community of the human colon: links between
diet and health: Minireview. Environ. Microbiol. 9,
1101–1111 (2007).
59.	 Allison, M. J., Dawson, K. A., Mayberry, W. R. &
Foss, J. G. Oxalobacter formigenes gen. nov., sp.
nov.: oxalate-degrading anaerobes that inhabit
the gastrointestinal tract. Arch. Microbiol. 141,
1–7 (1985).
60.	 Walker, A. W., Duncan, S. H., McWilliam
Leitch, E. C., Child, M. W. & Flint, H. J. pH and
peptide supply can radically alter bacterial
populations and short-chain fatty acid ratios
within microbial communities from the human
colon. Appl. Environ. Microbiol. 71, 3692–3700
(2005).
61.	 Louis, P., Scott, K. P., Duncan, S. H. & Flint, H. J.
Understanding the effects of diet on bacterial
metabolism in the large intestine. J. Appl.
Microbiol. 102, 1197–1208 (2007).
62.	 Van Wey, A. S. et al. Bacterial biofilms
associated with food particles in the human
large bowel. Mol. Nutr. Food Res. 55, 969–978
(2011).
63.	 Leitch, E. C. M., Walker, A. W., Duncan, S. H.,
Holtrop, G. & Flint, H. J. Selective colonization of
insoluble substrates by human faecal bacteria.
Environ. Microbiol. 9, 667–679 (2007).
64.	 Ze, X., Duncan, S. H., Louis, P. & Flint, H. J.
Ruminococcus bromii is a keystone species for
the degradation of resistant starch in the human
colon. ISME J. 6, 1535–1543 (2012).
65.	 Walker, A. W. et al. The species composition of
the human intestinal microbiota differs between
particle-associated and liquid phase
communities. Environ. Microbiol. 10,
3275–3283 (2008).
66.	 Martens, E. C., Koropatkin, N. M., Smith, T. J. &
Gordon, J. I. Complex glycan catabolism by the
human gut microbiota: The Bacteroidetes suslike
paradigm. J. Biol. Chem. 284, 24673–24677
(2009).
67.	 Flint, H. J., Bayer, E. A., Rincon, M. T., Lamed, R.
& White, B. A. Polysaccharide utilization by gut
bacteria: Potential for new insights from
genomic analysis. Nat. Rev. Microbiol. 6,
121–131 (2008).
68.	 Flint, H. J., Scott, K. P., Duncan, S. H., Louis, P.
& Forano, E. Microbial degradation of complex
carbohydrates in the gut. Gut Microbes
http://dx.doi.org/10.4161/gmic.19897.
69.	 van Passel, M. W. J. et al. The genome of
Akkermansia muciniphila, a dedicated intestinal
mucin degrader, and its use in exploring
intestinal metagenomes. PLoS ONE 6, e16876
(2011).
70.	 Derrien, M. et al. Modulation of mucosal immune
response, tolerance, and proliferation in mice
colonized by the mucin-degrader Akkermansia
muciniphila. Front. Microbiol. 2, 166 (2011).
71.	 Pomare, E. W., Branch, W. J. & Cummings, J. H.
Carbohydrate fermentation in the human colon
and its relation to acetate concentrations in
venous blood. J. Clin. Invest. 75, 1448–1454
(1985).
72.	 Sleeth, M. L., Thompson, E. L., Ford, H. E., ZacVarghese,
S. E. K. & Frost, G. Free fatty acid
receptor 2 and nutrient sensing: a proposed role
for fibre, fermentable carbohydrates and shortchain
fatty acids in appetite regulation. Nutr.
Res. Rev. 23, 135–145 (2010).
73.	 Hamer, H. M. et al. Review article: the role of
butyrate on colonic function. Aliment. Pharmacol.
Ther. 27, 104–119 (2008).
74.	 Gassull, M. A. Review article: the intestinal
lumen as a therapeutic target in inflammatory
bowel disease. Aliment. Pharmacol.Ther. 24,
90–95 (2006).
75.	 Lewis, S. J. & Heaton, K. W. Increasing butyrate
concentration in the distal colon by accelerating
intestinal transit. Gut 41, 245–251 (1997).
76.	 Scheppach, W. Effects of short chain fatty acids
on gut morphology and function. Gut 35,
S35–S38 (1994).
77.	 Gao, Z. et al. Butyrate improves insulin
sensitivity and increases energy expenditure in
mice. Diabetes 58, 1509–1517 (2009).
78.	 Louis, P. & Flint, H. J. Diversity, metabolism and
microbial ecology of butyrate-producing bacteria
from the human large intestine. FEMS Microbiol.
Lett. 294, 1–8 (2009).
79.	 Aminov, R. I. et al. Molecular diversity, cultivation,
and improved detection by fluorescent in situ
hybridization of a dominant group of human gut
bacteria related to Roseburia spp. or
Eubacterium rectale. Appl. Environ. Microbiol. 72,
6371–6376 (2006).
80.	 Scott, K. P. et al. Substrate-driven gene
expression in Roseburia inulinivorans:
Importance of inducible enzymes in the
utilization of inulin and starch. Proc. Natl Acad.
Sci. USA 108, 4672–4679 (2011).
81.	 Ramsay, A. G., Scott, K. P., Martin, J. C.,
Rincon, M. T. & Flint, H. J. Cell-associated
α‑amylases of butyrate-producing Firmicute
bacteria from the human colon. Microbiology
152, 3281–3290 (2006).
82.	 Lopez-Siles, M. et al. Cultured representatives
of two major phylogroups of human colonic
Faecalibacterium prausnitzii can utilize pectin,
uronic acids, and host-derived substrates for
growth. Appl. Environ. Microbiol. 78, 420–428
(2012).
83.	 Khan, M. T. et al. The gut anaerobe
Faecalibacterium prausnitzii uses an
extracellular electron shuttle to grow at oxicanoxic
interphases. ISME J. 6, 1578–1585
(2012).
84.	 Duncan, S. H. et al. Reduced dietary intake of
carbohydrates by obese subjects results in
decreased concentrations of butyrate and
butyrate-producing bacteria in feces. Appl.
Environ. Microbiol. 73, 1073–1078 (2007).
85.	 Brinkworth, G. D., Noakes, M., Clifton, P. M. &
Bird, A. R. Comparative effects of very lowcarbohydrate,
high-fat and high-carbohydrate, lowfat
weight-loss diets on bowel habit and faecal
short-chain fatty acids and bacterial populations.
Br. J. Nutr. 101, 1493–1502 (2009).
FOCUS ON GUT MICROBIOTA
© 2012 Macmillan Publishers Limited. All rights reserved
588  |  OCTOBER 2012  |  VOLUME 9 www.nature.com/nrgastro
86.	 Russell, W. R. et al. High-protein, reducedcarbohydrate
weight-loss diets promote
metabolite profiles likely to be detrimental to
colonic health. Am. J. Clin. Nutr. 93, 1062–1072
(2011).
87.	 El Oufir, L. et al. Relations between transit time,
fermentation products, and hydrogen consuming
flora in healthy humans. Gut 38, 870–877
(1996).
88.	 McOrist, A. L. et al. Fecal butyrate levels vary
widely among individuals but are usually
increased by a diet high in resistant starch.
J. Nutr. 141, 883–889 (2011).
89.	 Duncan, S. H., Louis, P., Thomson, J. M. &
Flint, H. J. The role of pH in determining the
species composition of the human colonic
microbiota. Environ. Microbiol. 11, 2112–2122
(2009).
90.	 Duncan, S. H., Louis, P. & Flint, H. J. Lactateutilizing
bacteria, isolated from human feces,
that produce butyrate as a major fermentation
product. Appl. Environ. Microbiol. 70, 5810–5817
(2004).
91.	 Belenguer, A. et al. Impact of pH on lactate
formation and utilization by human fecal
microbial communities. Appl. Environ. Microbiol.
73, 6526–6533 (2007).
92.	 Morrison, D. J. et al. Butyrate production from
oligofructose fermentation by the human faecal
flora: What is the contribution of extracellular
acetate and lactate? Br. J. Nutr. 96, 570–577
(2006).
93.	 Bourriaud, C. et al. Lactate is mainly fermented
to butyrate by human intestinal microfloras but
inter-individual variation is evident. J. Appl.
Microbiol. 99, 201–212 (2005).
94.	 Belenguer, A. et al. Rates of production and
utilization of lactate by microbial communities
from the human colon. FEMS Microbiol. Ecol. 77,
107–119 (2011).
95.	 Vernia, P. et al. Fecal lactate and ulcerative
colitis. Gastroenterology 95, 1564–1568 (1988).
96.	 Macfarlane, G. T. & Gibson, G. R. in
Gastrointestinal Microbiology Vol. I (eds
Mackie, R. I. & White, B. A.) 269–318 (Chapman
and Hall, London, 1997).
97.	 Scott, K. P., Martin, J. C., Campbell, G., Mayer, C.
& Flint, H. J. Whole-genome transcription
profiling reveals genes up-regulated by growth on
fucose in the human gut bacterium “Roseburia
inulinivorans”. J. Bacteriol. 188, 4340–4349
(2006).
98.	 Smith, E. A. & Macfarlane, G. T. Enumeration of
amino acid fermenting bacteria in the human
large intestine: Effects of pH and starch on
peptide metabolism and dissimilation of amino
acids. FEMS Microbiol. Ecol. 25, 355–368
(1998).
99.	 Gill, C. I. R. & Rowland, I. R. Diet and cancer:
Assessing the risk. Br. J. Nutr. 88, S73–S87
(2002).
100.	Macfarlane, S. & Macfarlane, G. T. Short-chain
fatty acids. Regulation of short-chain fatty acid
production. Proc. Nutr. Soc. 62, 67–72 (2003).
101.	Attene-Ramos, M. S., Wagner, E. D., Plewa, M. J.
& Gaskins, H. R. Evidence that hydrogen sulfide
is a genotoxic agent. Mol. Cancer Res. 4, 9–14
(2006).
102.	Medani, M. et al. Emerging role of hydrogen
sulfide in colonic physiology and
pathophysiology. Inflamm. Bowel Dis. 17,
1620–1625 (2011).
103.	Sahakian, A. B., Jee, S. R. & Pimentel, M.
Methane and the gastrointestinal tract. Dig. Dis.
Sci. 55, 2135–2143 (2010).
104.	Rey, F. E. et al. Dissecting the in vivo metabolic
potential of two human gut acetogens. J. Biol.
Chem. 285, 22082–22090 (2010).
105.	Nava, G. M., Carbonero, F., Croix, J. A.,
Greenberg, E. & Gaskins, H. R. Abundance and
diversity of mucosa-associated hydrogenotrophic
microbes in the healthy human colon. ISME J. 6,
57–70 (2012).
106.	Marquet, P., Duncan, S. H., Chassard, C.,
Bernalier-Donadille, A. & Flint, H. J. Lactate has
the potential to promote hydrogen sulphide
formation in the human colon. FEMS Microbiol.
Lett. 299, 128–134 (2009).
107.	Possemiers, S., Bolca, S., Verstraete, W. &
Heyerick, A. The intestinal microbiome: a
separate organ inside the body with the
metabolic potential to influence the bioactivity of
botanicals. Fitoterapia 82, 53–66 (2011).
108.	McIntosh, F. M. et al. Phylogenetic distribution of
genes encoding β‑glucuronidase activity in
human colonic bacteria and the impact of diet
on faecal glycosidase activities. Environ.
Microbiol. 14, 1876–1887 (2012).
109.	Gloux, K. et al. A metagenomic β‑glucuronidase
uncovers a core adaptive function of the human
intestinal microbiome. Proc. Natl Acad. Sci. USA
108, 4539–4546 (2011).
110.	Wikoff, W. R. et al. Metabolomics analysis
reveals large effects of gut microflora on
mammalian blood metabolites. Proc. Natl Acad.
Sci. USA 106, 3698–3703 (2009).
111.	Turnbaugh, P. J. et al. An obesity-associated gut
microbiome with increased capacity for energy
harvest. Nature 444, 1027–1031 (2006).
112.	Blaut, M. & Klaus, S. Intestinal microbiota and
obesity. Handb. Exp. Pharmacol. 209, 251–273
(2012).
113.	Flint, H. J. Obesity and the gut microbiota. J. Clin.
Gastroenterol. 45, S128–S132 (2011).
114.	Roberfroid, M. B. Caloric value of inulin and
oligofructose. J. Nutr. 129, 1436S–1437S
(1999).
115.	Parnell, J. A. & Reimer, R. A. Prebiotic fibres
dose-dependently increase satiety hormones
and alter Bacteroidetes and Firmicutes in lean
and obese JCR:LA-cp rats. Br. J. Nutr. 107,
601–613 (2012).
116.	Schwiertz, A. et al. Microbiota and SCFA in lean
and overweight healthy subjects. Obesity 18,
190–195 (2010).
117.	Duncan, S. H. et al. Human colonic microbiota
associated with diet, obesity and weight loss.
Int. J. Obes. 32, 1720–1724 (2008).
118.	Jumpertz, R. et al. Energy-balance studies reveal
associations between gut microbes, caloric load,
and nutrient absorption in humans. Am. J. Clin.
Nutr. 94, 58–65 (2011).
119.	Larsen, N. et al. Gut microbiota in human adults
with type 2 diabetes differs from non-diabetic
adults. PLoS ONE 5, e9085 (2010).
120.	Ravussin, Y. et al. Responses of gut microbiota
to diet composition and weight loss in lean and
obese mice. Obesity 20, 738–747 (2012).
121.	Bäckhed, F. et al. The gut microbiota as an
environmental factor that regulates fat storage.
Proc. Natl Acad. Sci. USA 101, 15718–15723
(2004).
122.	Fleissner, C. K. et al. Absence of intestinal
microbiota does not protect mice from dietinduced
obesity. Br. J. Nutr. 104, 919–929
(2010).
123.	Hildebrandt, M. A. et al. High-fat diet determines
the composition of the murine gut microbiome
independently of obesity. Gastroenterology 137,
1716–1724e2 (2009).
124.	Murphy, E. F. et al. Composition and energy
harvesting capacity of the gut microbiota:
relationship to diet, obesity and time in mouse
models. Gut 59, 1635–1642 (2010).
125.	Turnbaugh, P. J. et al. The effect of diet on the
human gut microbiome: a metagenomic analysis
in humanized gnotobiotic mice. Sci.Transl. Med.
1, 6ra14 (2009).
126.	Vijay-Kumar, M. et al. Metabolic syndrome and
altered gut microbiota in mice lacking toll-like
receptor 5. Science 328, 228–231 (2010).
127.	Cani, P. D. et al. Metabolic endotoxemia initiates
obesity and insulin resistance. Diabetes 56,
1761–1772 (2007).
128.	Willing, B. et al. Twin studies reveal specific
imbalances in the mucosa-associated
microbiota of patients with ileal Crohn’s disease.
Inflamm. Bowel Dis. 15, 653–660 (2009).
129.	Sokol, H. et al. Low counts of Faecalibacterium
prausnitzii in colitis microbiota. Inflamm. Bowel
Dis. 15, 1183–1189 (2009).
130.	Manichanh, C., Borruel, N., Casellas, F. &
Guarner, F. The gut microbiota in IBD. Nat. Rev.
Gastroenterol. Hepatol. http://doi.dx.org/
nrgastro.2012.152.
131.	Jia, W. et al. Is the abundance of
Faecalibacterium prausnitzii relevant to Crohn’s
disease? FEMS Microbiol. Lett. 310, 138–144
(2010).
132.	Mukhopadhya, I., Hansen, R., El-Omar, E. M.
& Hold, G. L. IBD—what role do proteobacteria
play? Nat. Rev. Gastroenterol. Hepatol. 9,
219–230 (2012).
133.	Rajilic´-Stojanovic´, M. et al. Global and deep
molecular analysis of microbiota signatures in
fecal samples from patients with irritable bowel
syndrome. Gastroenterology 141, 1792–1801
(2011).
134.	Chassard, C. et al. Functional dysbiosis within
the gut microbiota of patients with constipatedirritable
bowel syndrome. Aliment. Pharmacol.
Ther. 35, 828–838 (2012).
135.	Simrén, M. et al. Intestinal microbiota in
functional bowel disorders: a Rome Foundation
working team report. Gut http://dx.doi.
org/10.1136/gutjnl‑2012‑30267.
136.	Boleij, A. & Tjalsma, H. Gut bacteria in health
and disease: A survey on the interface between
intestinal microbiology and colorectal cancer.
Biol. Rev. 87, 701–730 (2012).
137.	Wang, T. et al. Structural segregation of gut
microbiota between colorectal cancer patients
and healthy volunteers. ISME J. 6, 320–329
(2012).
138.	Stecher, B. et al. Salmonella enterica serovar
Typhimurium exploits inflammation to compete
with the intestinal microbiota. PLoS Biol. 5,
2177–2189 (2007).
139.	Jernberg, C., Löfmark, S., Edlund, C. &
Jansson, J. K. Long-term impacts of antibiotic
exposure on the human intestinal microbiota.
Microbiology 156, 3216–3223 (2010).
140.	Khoruts, A., Dicksved, J., Jansson, J. K. &
Sadowsky, M. J. Changes in the composition of
the human fecal microbiome after
bacteriotherapy for recurrent Clostridium difficileassociated
diarrhea. J. Clin. Gastroenterol. 44,
354–360 (2010).
141.	Guo, B., Harstall, C., Louie, T., Veldhuyzen Van
Zanten, S. & Dieleman, L. A. Systematic review:
faecal transplantation for the treatment of
Clostridium difficile-associated disease.
Aliment. Pharmacol. Ther. 35, 865–875 (2012).
142.	Mattila, E. et al. Fecal transplantation, through
colonoscopy, is effective therapy for recurrent
Clostridium difficile infection. Gastroenterology
142, 490–496 (2012).
143.	Borody, T. J. & Khoruts, A. Fecal microbiota
transplantation and emerging applications. Nat.
Rev. Gastroenterol. Hepatol. 9, 88–96 (2012).
144.	Swidsinski, A., Loening-Baucke, V.,
Verstraelen, H., Osowska, S. & Doerffel, Y.
Biostructure of fecal microbiota in healthy
subjects and patients with chronic idiopathic
REVIEWS
© 2012 Macmillan Publishers Limited. All rights reserved
NATURE REVIEWS | GASTROENTEROLOGY & HEPATOLOGY 	 VOLUME 9  |  OCTOBER 2012  |  589
diarrhea. Gastroenterology 135, 568–579e2
(2008).
145.	Baughn, A. D. & Malamy, M. H. The strict
anaerobe Bacteroides fragilis grows in and
benefits from nanomolar concentrations of
oxygen. Nature 427, 441–444 (2004).
146.	Jones, B. V., Begley, M., Hill, C.,
Gahan, C. G. M. & Marchesi, J. R. Functional
and comparative metagenomic analysis of bile
salt hydrolase activity in the human gut
microbiome. Proc. Natl Acad. Sci. USA 105,
13580–13585 (2008).
147.	Islam, K. B. M. S. et al. Bile acid is a host
factor that regulates the composition of the
cecal microbiota in rats. Gastroenterology 141,
1773–1781 (2011).
148.	Gagen, E. J. et al. Functional gene analysis
suggests different acetogen populations in the
bovine rumen and tammar wallaby forestomach.
Appl. Environ. Microbiol. 76, 7785–7795
(2010).
149.	Scanlan, P. D., Shanahan, F. & Marchesi, J. R.
Culture-independent analysis of desulfovibrios
in the human distal colon of healthy, colorectal
cancer and polypectomized individuals. FEMS
Microbiol. Ecol. 69, 213–221 (2009).
150.	Mihajlovski, A., Doré, J., Levenez, F., Alric, M. &
Brugère, J. F. Molecular evaluation of the human
gut methanogenic archaeal microbiota reveals
an age-associated increase of the diversity.
Environ. Microbiol. Rep. 2, 272–280 (2010).
151.	Hayashi, H. et al. Direct cloning of genes
encoding novel xylanases from the human gut.
Can. J. Microbiol. 51, 251–259 (2005).
152.	Tasse, L. et al. Functional metagenomics to
mine the human gut microbiome for dietary
fiber catabolic enzymes. Genome Res. 20,
1605–1612 (2010).
153.	Verberkmoes, N. C. et al. Shotgun
metaproteomics of the human distal gut
microbiota. ISME J. 3, 179–189 (2009).
154.	Martínez, I., Kim, J., Duffy, P. R., Schlegel, V. L.
& Walter, J. Resistant starches types 2 and 4
have differential effects on the composition of
the fecal microbiota in human subjects. PLoS
ONE 5, e15046 (2010).
155.	Abell, G. C. J., Cooke, C. M., Bennett, C. N.,
Conlon, M. A. & McOrist, A. L. Phylotypes related
to Ruminococcus bromii are abundant in the
large bowel of humans and increase in response
to a diet high in resistant starch. FEMS Microbiol.
Ecol. 66, 505–515 (2008).
156.	Costabile, A. et al. A double-blind, placebocontrolled,
cross-over study to establish the
bifidogenic effect of a very‑long‑chain inulin
extracted from globe artichoke (Cynara
scolymus) in healthy human subjects. Br. J. Nutr.
104, 1007–1017 (2010).
157.	Kleessen, B. et al. Jerusalem artichoke and
chicory inulin in bakery products affect faecal
microbiota of healthy volunteers. Br. J. Nutr. 98,
540–549 (2007).
158.	Fernando, W. M. et al. Diets supplemented with
chickpea or its main oligosaccharide component
raffinose modify faecal microbial composition in
healthy adults. Benef. Microbes 1, 197–207
(2010).
Acknowledgements
The authors receive support from the Scottish
Government Rural and Environment Science and
Analysis Service.
Author contributions
H. J. Flint researched data and content for the article.
H. J. Flint and P. Louis reviewed and/or edited the
manuscript before submission. All authors
contributed to writing the article.
FOCUS ON GUT MICROBIOTA
© 2012 Macmillan Publishers Limited. All rights reserved