Changes in Human Fecal Microbiota Due to
Chemotherapy Analyzed by TaqMan-PCR, 454
Sequencing and PCR-DGGE Fingerprinting
Jutta Zwielehner1, Cornelia Lassl1, Berit Hippe1, Angelika Pointner1, Olivier J. Switzeny1, Marlene
Remely1, Elvira Kitzweger2, Reinhard Ruckser2, Alexander G. Haslberger1*
1Department of Nutritional Sciences, Vienna, Austria, 2Sozialmedizinisches Zentrum Ost, Vienna, Austria
Background: We investigated whether chemotherapy with the presence or absence of antibiotics against different kinds of
cancer changed the gastrointestinal microbiota.
Methodology/Principal Findings: Feces of 17 ambulant patients receiving chemotherapy with or without concomitant
antibiotics were analyzed before and after the chemotherapy cycle at four time points in comparison to 17 gender-, age-
and lifestyle-matched healthy controls. We targeted 16S rRNA genes of all bacteria, Bacteroides, bifidobacteria, Clostridium
cluster IV and XIVa as well as C. difficile with TaqMan qPCR, denaturing gradient gel electrophoresis (DGGE) fingerprinting
and high-throughput sequencing. After a significant drop in the abundance of microbiota (p=0.037) following a single
treatment the microbiota recovered within a few days. The chemotherapeutical treatment marginally affected the
Bacteroides while the Clostridium cluster IV and XIVa were significantly more sensitive to chemotherapy and antibiotic
treatment. DGGE fingerprinting showed decreased diversity of Clostridium cluster IV and XIVa in response to chemotherapy
with cluster IV diversity being particularly affected by antibiotics. The occurrence of C. difficile in three out of seventeen
subjects was accompanied by a decrease in the genera Bifidobacterium, Lactobacillus, Veillonella and Faecalibacterium
prausnitzii. Enterococcus faecium increased following chemotherapy.
Conclusions/Significance: Despite high individual variations, these results suggest that the observed changes in the human
gut microbiota may favor colonization with C.difficile and Enterococcus faecium. Perturbed microbiota may be a target for
specific mitigation with safe pre- and probiotics.
Citation: Zwielehner J, Lassl C, Hippe B, Pointner A, Switzeny OJ, et al. (2011) Changes in Human Fecal Microbiota Due to Chemotherapy Analyzed by TaqMan-
PCR, 454 Sequencing and PCR-DGGE Fingerprinting. PLoS ONE 6(12): e28654. doi:10.1371/journal.pone.0028654
Editor: Markus M. Heimesaat, Charite ´, Campus Benjamin Franklin, Germany
Received September 20, 2011; Accepted November 11, 2011; Published December 14, 2011
Copyright: ? 2011 Zwielehner et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: The work was supported by the Austrian Science Fund (FWF). The funders had no role in study design, data collection and analysis, decision to publish,
or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: firstname.lastname@example.org
The human intestinal ecosystem can be pictured as a microbial
organ within a host organism involving a dynamic interplay
between food, host cells and microbes . The microbiota plays
several significant roles in the digestion of food, energy regulation,
generation of short-chain fatty acids, vitamin synthesis, prevention
of colonization by pathogens and protection against cell injury
[2,3,4]. Moreover, the gut microbiota influences the host by
directing intestinal epithelial cell proliferation and differentiation,
pH, and the development of the immune system . Recent
culture-independent molecular studies on healthy individuals have
shown that the intestinal microbiota is specific to the host  and
resilient to modifications over time, as it is able to form an
alternative stable state after disruption [6,7]. A healthy microbiota
contains a balanced composition of many classes of bacteria .
The fecal microbiota is dominated by three groups of anaerobic
bacteria: the Clostridium coccoides group -clostridial cluster XIVa
(reclassified as Blautia coccoides ), the Clostridium leptum group -
Clostridium cluster IV, and the Bacteroides [10,11]. All three groups
are known to positively affect the gut health through nutrient
absorption, production of short chain fatty acids (SCFAs) and
epithelial cell maturation [12,13]. Moreover, the subgroup
bifidobacteria seems to be an important part of the gastrointestinal
tract (GI) microbiota, being involved in the prevention of atopic
disease, obesity and insulin resistance via enhanced barrier
function of the gut epithelium .
To prevent the invasion of endogenous bacteria from oral cavity
and the GI tract into the blood stream, three defense mechanisms
are considered to be relevant: innate immunity, mechanical
mucosal barrier, and colonization resistance . However,
chemotherapy damages the rapidly generated mucosal cells of
the GI and the use of antibiotics disrupts the ecological balance,
allowing pathogens such as Clostridium difficile to grow [16,17]. This
bacterium is thought to be the causative agent in up to 20% of
antibiotic-associated diarrhea (AAD) cases . It is evident that
the intestinal microbial ecosystem has an important but incom-
pletely defined role in mucosal protection .
PLoS ONE | www.plosone.org1 December 2011 | Volume 6 | Issue 12 | e28654
Mucositis is a major oncological problem,causedby the cytotoxic
effects of cancer chemotherapy and radiotherapy . Approxi-
mately 40% of patients receiving standard dose chemotherapy and
up to 100% of patients receiving high dose chemotherapy and stem
cell or bone marrow transplantation suffer from abdominal pain,
ulceration, bloating and vomiting [21,22]. Although gastrointestinal
disturbances (mucositis, diarrhea and constipation) and immuno-
suppression are well recognized side-effects of cancer treatment,
very little research has been conducted into the underlying
mechanisms and the changes in the composition of the microbiota.
Because of these changes, nutrient absorption and other intestinal
functions involving the microbiota may also be altered .
receiving cancer chemotherapy with or without antibiotics in
comparison to healthy control individuals. Prescription of antibiotics
may become necessary in some individuals due to bacterial infection
. Samples were taken at four time points before and after
chemotherapy to study changes infecal microbiota over the courseof
time. In this study, we aimed to clarify how chemotherapy agents
influence total fecal bacteria, Bacteroides, bifidobacteria, Clostridium
cluster IV, Clostridium cluster XIVa and C. difficile using culture-
independent methods assessing abundance and diversity. Four
samples were also analyzed with 454 high-throughput sequencing.
PCR-DGGE fingerprinting analysis shows decreased
diversity of Clostridium clusters IV and XIVa in response to
medical treatment compared to healthy individuals
DGGE fingerprinting analyses of all bacteria, Clostridium cluster
IV and Clostridium cluster XIVa indicate a highly diverse dataset
between individuals and uniqueness of fecal microbiota. Table 1
shows the average number of bands in cancer patients at the three
time points and for controls over all time points. It becomes
apparent that the average number of bands within Clostridium
cluster IV declined immediately after chemotherapy (T1), followed
by a recovery at T2. The average number of Clostridium cluster
XIVa bands decreased after onset of chemotherapy and remained
low also at T2. The datasets were subjected to principal
component analysis (PCA). PCA extracts underlying components
of samples according to their variance. Figure 1A illustrates the
bacterial fingerprints of sample P01 over time. Figure 1B displays
the PCA analysis of all bacteria. Most samples taken after
chemotherapy are grouped together with all other samples.
Patients who receive antibiotics are indicated as black symbols.
They cluster together with the samples taken after chemotherapy
and also with the majority of samples before chemotherapy and
healthy controls. There are two exceptions though: Two samples
from P07 after chemotherapy under antibiotic treatment are
outliers in the lower right part of the PCA plot. P07 received blood
stem cell transplantation resulting in a sharp decline in bacterial
abundances as measured with quantitative PCR. The first two
principal components explain 17.4% of variance.
Figure 1C shows the principal components analysis of Clostridium
found to be less variable than healthy controls and patients before
onset oftreatment.Althoughoverlapping,PCAresulted ingrouping
of band patterns before and after chemotherapy. Additional effects
by antibiotic treatment became evident: Antibiotic treatment
significantly reduced the diversity within the Clostridium cluster IV
(p=0.00003) with Shannon diversity index being 1.460.7 com-
pared to patients under chemotherapy alone 2.160.6. In the PCA
plot, samples affected by antibiotics are found in the lower right
corner of the plot. This means that they are grouped according to
their variance along principal component (PC) 1 and 2. These two
PCs explain 17.9% and 9.15% of the variance in the dataset,
underlining the validity of this interpretation. Principal components
analysis of Clostridium cluster XIVa is not shown.
Chemotherapeutic treatment with or without antibiotics
decreases absolute bacterial numbers in comparison to
To study whether chemotherapy with or without antibiotics
changes the human GI microbiota composition in contrast to
healthy individuals and over time, we investigated absolute numbers
and relative percentages of bacterial subgroups. Absolute numbers
give an indication about the direct antimicrobial effects of the
treatments. Relative quantification is able to identify which bacterial
subgroups are particularly affected and helps to describe the
community disruption induced by chemotherapy with or without
antibiotics. In absolute numbers, oncology patients harbored
significantly less bacteria (p,0.05) than healthy control (figure 2).
From already low bacterial counts before chemotherapy, bacterial
abundance significantly declined further (p=0.037) immediately
after chemotherapy (T1) and recovered 5–9 days later (T2) in
comparison to time points before treatment (T0). Absolute numbers
of bacteria in different time points of healthy controls are following a
lognormal distribution in contrast to microbiota abundances in
oncology patients. The decrease in total bacteria following
chemotherapy (p=0.037) was significantly greater than any
variation in copy numbers observed in healthy controls (p=0.027).
The observed decrease after chemotherapy affected the Bacteroides
(p=0.044), the bifidobacteria (p=0.034) and Clostridium cluster IV
(p=0.049) as shown in figure 3. There were also fewer absolute
numbers of Clostridium cluster XIVa, but this difference was not
significant. All patients with fever showed an increase in total fecal
microbiota (see figure 3). In sample P07 a sharp decline affecting all
bacteria and bacterial subgroups was observed at T1 (figure 3),
following blood stem cell transplantation and medical intervention.
Patients who received antibiotics had highest abundances of all
bacteria (p=0.000003) amongst all patients (data not shown). This
bacterial overgrowth affected the Bacteroides, the bifidobacteria and
Clostridium clusters IV and XIVa, since relative abundances of those
subgroups did not stand out significantly. Thus, patients were
grouped according to their chemotherapeutic cycle regardless
whether or not they received antibiotics. The influence of
antibiotics on the species composition as assessed with PCR-
DGGE fingerprinting is discussed in the previous section.
Clostridium cluster XIVa shows great alterations due to
chemotherapeutical interventions, while the Bacteroides
and bifidobacteria seem to be marginally affected
Relative quantification of Clostridium cluster XIVa as percentage
of total bacterial DNA showed that oncology patients harbored
significantly less Clostridium cluster XIVa (p=0.047) than healthy
controls. The mean proportion of Bacteroides in stool samples was
26612% in chemotherapy patients and 22614% in healthy
individuals. The mean percentage of bifidobacteria in patients was
0.861.4% and 0.360.6 in controls. Patients harbored on average
1669% of Clostridium cluster IV and 18612% of Clostridium cluster
XIVa, while controls harbored 20612% and 24615% of
clostridial clusters IV and XIVa.
Clostridium cluster XIVa higher before chemotherapy
The mean percentage of Clostridium cluster XIVa before
chemotherapy was 22613% compared to after chemotherapeutic
Chemotherapy Changes Fecal Microbiota
PLoS ONE | www.plosone.org2 December 2011 | Volume 6 | Issue 12 | e28654
cycles with 19612%. The average amount of Bacteroides,
bifidobacteria and Clostridium cluster IV were 26611%, 1.462%
and 1669% at time points before chemotherapy and 28614%,
0.561.2% and 18612% after chemotherapy. Figure 3 illustrates
the development of the microbiota in the course of antibiotic
treatment. Data were normalized for clarity, so that changes in
abundances from time point T0 (before onset of treatment) to T1
(1–4 days after chemotherapy) and T2 (5–9 days after chemo-
therapy) rather than relative abundances are shown. It can be seen
that chemotherapy causes a dramatic reduction of microbiota
abundance immediately after chemotherapy, affecting all sub-
groups. As mentioned above, the significant decrease in all
bacteria following chemotherapy was significantly greater than
any variation in copy numbers observed in healthy controls
C. difficile colonization found in individuals receiving
chemotherapeutic and antibiotic treatment
To find out whether the chemotherapeutic and antibiotic
disruption favors the growth of pathogens, we investigated the
abundance of C. difficile. Three out of seventeen patients receiving
chemotherapy harbored C. difficile (data not shown). Patient P09
harbored C. difficile at all time points investigated. Mean
proportion over all four samples of P09 was recorded as
0.460.7%, yet the highest level (1.22% of total bacteria) occurred
at sampling point T1 immediately after chemotherapeutic and
antibiotic treatment. C. difficile was detected in P11 (3.90% of all
analyzed bacteria) after chemotherapeutic intervention at time
point T1. P14 carried C. difficile in low abundance directly after
onset of chemotherapy (0.003% of all analyzed bacteria). Samples
of patients P09 and P11 at T0 and T1 were further analyzed in
Figure 1. A PCR-DGGE fingerprinting of 16S rRNA coding regions of dominant bacteria over time. Bands that become stronger or nearly
disappear following a single chemotherapeutic treatment are indicated with arrows. B Principal components analysis (PCA) based on dominant
bacteria PCR-DGGE fingerprinting. The two outliers in the lower right corner of the plot are two samples of P07 following blood stem cell
transplantation. C PCA illustrating the development of Clostridium cluster IV diversity in the course of chemotherapy and antibiotic treatment. Cluster
IV diversity drops right after chemotherapy, causing a grouping of samples. Samples under antibiotic treatment (indicated as grey dots) group even
closer, indicating a strong influence of antibiotics on Clostridium cluster IV diversity. A, sample of P01 before chemotherapy B, C and D, samples of P01
after chemotherapy; E, healthy control; SL, unrelated standard lane; black symbols… patients under chemotherapy and antibiotic treatment.
Table 1. Number of bands observed in PCR-DGGE
fingerprinting in oncology patients before chemotherapy (T0),
immediately after chemotherapy (T1) and 5–9 days after
chemotherapy (T2) and healthy controls averaged over all
19.663.6 1566.0 5.262.6
control 19.263.512.065.0 8.963.0
Chemotherapy Changes Fecal Microbiota
PLoS ONE | www.plosone.org3 December 2011 | Volume 6 | Issue 12 | e28654
High throughput sequencing
High throughput sequencing showed a dramatic increase in
sequences within the Peptostreptococcaceae towards sequences 98.9–
100% similar to C.difficileT(figure 4): Clostridium bartletti related
sequences (98.1–100% similarity) were only detected before
chemotherapy (T0). After chemotherapy (T1), 63 sequences
98.9–100% similar to C.difficile appeared in samples P11 and
P09. In accordance with the phylogenetic classification by the
ribosomal database project, they are shown as ‘unclassified
Peptostreptococcaceae’ in figure 5.
Furthermore pronounced reductions of Faecalibacterium spp. as
well as lactobacilli, Veillonella spp., bifidobacteria (in P09) and
E.coli/Shigella became apparent in response to chemotherapy
(figure 5). The abundance of lactobacilli decreased in both patients
after chemotherapy, in P09 from already low levels. Individual
P11 did not receive concomitant antibiotics, whereas P09 did. In
P09 and P11 Faecalibacterium spp. decreased dramatically from
9.5% and 8.3% to 0.07% and 0.00%, respectively. In both
individuals Enterococcus faecium increased following chemotherapy.
Furthermore, less abundant sequences appeared that were
attributable to bacterial genera not detected before chemotherapy.
These genera are: Eggerthella, Megasphaera, Parvimonas (only in P11),
Anaerostipes, Eubacterium, Anaerococcus, Methylobacterium, Holdemania,
Turicibacter, Akkermansia, Sutterella (only in P09), Sphingomonas,
Anaerotruncus, Coprococcus, Streptococcus and Dorea. Species with
abundance ,0.01% of all sequences are not shown in figure 5.
The number of Blautia species from Clostridium cluster XIVa
remained constant in the 454 sequencing datasets before and after
Chemotherapeutic and antibiotic use is associated with severe
side effects such as mucositis, diarrhea and constipation. These
side effects increase the cost of health services and are often life
threatening . Chemotherapeutic and antibiotic treatment has
a detrimental impact on the host microbial ecosystem, which is
important for host mucosal protection  and thereby increases
the risk of infection . Overgrowth of species with potential
pathogenicity such as toxigenic C. difficile and inflammatory
complications are among the most common serious complications
of chemotherapy and antibiotic treatment among patients with
We investigated how the use of cancer chemotherapy (in some
individuals together with antibiotic treatment) perturbs the fecal
microbial ecosystem during the course of therapy. We assessed if
the microbiota is able to return to its original profile after
chemotherapeutic and antibiotic intervention with special interest
in the abundance of C. difficile. We used a combination of
molecular methods including high-throughput sequencing to
compare diversity (PCR-DGGE) and abundance (qPCR) of all
Figure 2. TaqMan qPCR quantification of bacterial 16S rRNA
coding regions showing lower abundance in patients under-
going chemotherapy and antibiotic treatment (P) than healthy
controls (C). T0, samples taken before a single shot of chemotherapy;
T1, 1–2 days after chemotherapy; T2, 5–9 days after chemotherapy;
Asterisk indicates a significant difference at p,0.05.
Figure 3. Abundances of bacterial 16S rRNA coding regions
over time in oncology patients (P) and healthy controls (C). The
declined abundances of bacteria, Bacteroides, Clostridium cluster XIVa,
Clostridium cluster IV and bifidobacteria immediately after chemother-
apy (T1) were observed to recover several days after treatment (T2).
Patients P04, P08 and P13 had never received chemotherapy before;
P04, P05, P07, P08, P09 and P10 took antibiotics. Values were z-scored
for presentation in this heatmap showing changes over time rather
than absolute abundances. T0, before chemotherapy; T1, 1–2 days after
chemotherapy; T2, 5–9 days after chemotherapy; F, fever; S, blood stem
Chemotherapy Changes Fecal Microbiota
PLoS ONE | www.plosone.org4 December 2011 | Volume 6 | Issue 12 | e28654
bacteria, Bacteroides, bifidobacteria, Clostridium cluster IV, Clostridium
cluster XIVa and C. difficile between groups and different time
points of chemotherapy. The majority of previous studies on the
effect of chemotherapy on human fecal microbiota used standard
microbiological culture techniques [16,22]. Other studies have
focused on the colonization of pathogenic bacteria [17,25] in
patients with cancer and chemotherapy-induced diarrhea [22,26].
As mentioned above, we used feces as source of information. Fecal
microbial communities are composed of autochthonous gut
members and transient bacteria. Even though the fecal microbiota
might be different from the adherent microbiota, we chose fecal
samples to investigate the microbial composition of the intestinal
microbiota because they are easy to collect, are less invasive and
reflect shifts in microbial population composition .
In this study, we assessed species richness using PCR-DGGE
fingerprinting. Each lane of a PCR-DGGE gel represented a
microbial fingerprint of a fecal sample; each band within a lane
corresponded to one bacterial species, although different species
may sometimes be represented by the same band . It has also
been observed that one bacterial strain may form several bands
due to multiple 16S rRNA operons, e.g. E.coli (figure 4A). The
limitations of DGGE in microbial analysis have been previously
described . Nevertheless, substantial information about species
composition can be obtained from very complex microbial
communities such as the gut microbiota . We found decreased
species richness immediately after the chemotherapeutic shot,
especially within Clostridium cluster IV where the number of
different bands decreased from 1467 before chemotherapy (T0) to
1066 bands shortly after (T1). The microbiota recovered to a
richness of 1566 Clostridium cluster IV bands per individual, but at
a different composition, as evidenced by the grouping of samples
in principal components analysis.
For quantification of fecal microbiota we used the strains
Bacteroides thetaiotaomicronT, Bifidobacterium longum ssp. longumTand C.
difficile as well as the clones CL16 and CC34 as standards.
However, a mixture of different strains for qPCR standards might
result in a more accurate image of the human microbiota.
Therefore absolute amounts should be considered as semi-
Grouping oncology patients with and without antibiotic
treatment poses a risk to falsely interpret the effects of antibiotic
treatment as effects of chemotherapy. Patients who received
antibiotics had significantly higher bacterial abundances than
patients without antibiotics. This observation might be the reason
Figure 4. Phylogenetic tree showing the Peptostreptococcaceae found in samples from two oncology patients before and after
chemotherapy. Identical sequences were grouped; the table on the right hand side shows their abundances in the 454 sequencing dataset.
Sequences with .98.9% similarity to Clostridium difficile appeared only in samples taken immediately after chemotherapeutic cycles. Numbers
indicate bootstrap values after 100 resamplings.
Chemotherapy Changes Fecal Microbiota
PLoS ONE | www.plosone.org5 December 2011 | Volume 6 | Issue 12 | e28654
for antibiotic treatment rather than its effect . The abundance
of bacterial subgroups, also Clostridium cluster IV, changed together
with total bacteria both in patients with and without antibiotics.
The sharp reduction of bacteria immediately after chemotherapy
equally affected patients with and without antibiotics. In PCR-
DGGE analysis we found that the all bacteria and Clostridium
cluster XIVa fingerprints did not differ significantly in patients with
or without antibiotics. This indicates that the use of antibiotics does
not fully explain the observed changes. Previous work  has also
found additional effects of chemotherapy in cases under
prophylactic antibiotic treatment. Although the Clostridium cluster
IV abundance did not differ significantly due to antibiotics, PCR-
DGGE fingerprints showed grouping of patients under antibiotic
treatment in principal components analysis.
Despite high individual variations, we show a significantly lower
absolute bacterial load in feces of patients receiving chemotherapy
in comparison to healthy controls. These findings are in line with
data from van Vliet et al. (2009) who reported 100-fold lower total
bacterial numbers during chemotherapy than in healthy controls.
The abundance of fecal microbiota decreased after a single
cycle of chemotherapy. After the end of chemotherapeutic
administration the bacterial abundance recovered within a few
days, sometimes even showing a ‘‘rebound-effect’’ with numbers
elevating above initial levels. Relative numbers of Clostridium
cluster IV and XIVa showed great alterations due to chemother-
apeutical interventions, while the bifidobacteria seemed to be less
affected. In agreement with previous results  increased counts
of Bacteroides spp. were found in patients undergoing chemother-
apy. Nyhle `n et al. (2007) also reported significant increases in yeast
in patients, making it a focus for further research in immunocom-
promised patients. Samples taken immediately after chemotherapy
had a lower diversity within Clostridium cluster IV. Antibiotics
strongly contributed to the reduced diversity of cluster IV but were
not alone responsible for this effect. A few days later we observed a
quantitative recovery, but not a recovery of the composition as
evidenced by clustering of DGGE fingerprints.
The incidence of C. difficile in subjects P09 and P11 immediately
after chemotherapy is accompanied by a decrease of the genera
Bifidobacterium, Lactobacillus and Clostridium cluster IV. Sequences
attributable to Faecalibacterium prausnitzii decreased dramatically
from 9% to zero. The anti-inflammatory F. prausnitzii was
associated with dietary fiber in colonic fermentation of healthy
subjects  and found at low abundance in individuals suffering
from inflammatory bowel diseases  [30,31]. Enterococcus faecium
increased following chemotherapy, possibly filling the ecological
niches vacated by the lactobacilli and bifidobacteria. Enterococcus
faecium is a facultative pathogenic bacterium causing life-threaten-
ing infections especially in nosocomial settings . Enterococcus
faecium has previously been found to increase in wastewater upon
treatment . The acquisition of multi-resistant E. faecium strains
has been described in hospital environments under high selective
antibiotic pressure. Under such conditions probiotic strains were
demonstrated as unable to prevent nosocomial infection .
After chemotherapy less abundant sequences appeared that
were not detected before treatment. These genera are: Eggerthella,
Megasphaera, Parvimonas, Anaerostipes, Eubacterium, Anaerococcus, Methy-
lobacterium, Holdemania, Turicibacter, Akkermansia, Sutterella, Sphingomo-
nas, Anaerotruncus, Coprococcus, Streptococcus and Dorea. Eggerthella lenta
was described to convert dietary lignans to the bioactive
enterolactone . Megasphaera spp. have been described as
propionate-producers that utilize lactate  comparable to
Veillonella spp. that were no longer detected by 454 sequencing
after chemotherapy. The butyrate-producing Anaerostipes caccae and
Eubacterium hallii utilize lactate as well. They were suggested to
compete for lactate with sulfate-reducing bacteria such as
Desulfobacter piger whose preferred co-substrate is lactate. High
concentrations of sulfate are toxic for the gut epithelium and may
contribute to bowel disease . Microorganisms of the genus
Methylobacterium are facultative methylotrophic, gram-negative rods
that are ubiquitous in nature and rarely cause human disease,
except in subjects with pre-existing immunosuppression. For
Figure 5. Heatmap showing abundances within the 454
sequencing dataset on the genus level. High throughput
sequencing of samples P09 and P11 before (T0) and after therapy (T1)
further helped to characterize the influence of a single chemothera-
peutic cycle on the GI-microbiota. P11 was treated with chemotherapy
alone and P09 also received antibiotic treatment.
Chemotherapy Changes Fecal Microbiota
PLoS ONE | www.plosone.org6December 2011 | Volume 6 | Issue 12 | e28654
instance, in 2010, a case of M. fujisawaense infection was described
in a patient with relapsed acute leukemia undergoing unrelated
allogeneic hematopoietic stem cell transplantation . Turicibacter
is a poorly known genus previously found in weaned piglets,
known to be susceptible to chlortetracycline . In humans,
Turicibacter spp. have been found in the ileal pelvic pouch of a
former ulcerative colitis patient . Akkermansia muciniphila is a
common mucin-degrading bacterium of the human GI. Its
prevalence has been described to be 108cells/g feces in adults,
decreasing with age . Dorea spp. are mucosa-associated
bacteria of the human GI that are members of the Clostridium
coccoides rRNA group of organisms [42,43].
Further research is needed to elucidate if there is a causal
relationship between growth of C. difficile and decreased abundance
of lactobacilli, bifidobacteria and Clostridium cluster IV, especiallythe
anti-inflammatory Faecalibacterium prausnitzii. The increase of mucus-
degrading bacteria might be a result of C.difficile and probably also
E. faecium associated inflammation of the gut epithelium. Mucus
hypersecretion is a common symptom of irritable bowel syndrome,
ulcerative colitis and bacterial infections of the gut epithelium
[44,45]. The lactate-utilizing microbiota shifted from Veillonella spp.
to Anaerostipes, Eubacterium and Megasphaera spp. This change may be
interpreted as a beneficial adaptation, because lactate could
otherwise be used as a co-substrate for sulfate-reduction. Sulfate-
reducing bacteria, however, were not detected here.
The oncology patients assessed here suffered from a variety of
malignancies and received different chemotherapy treatment
regimes. Only two participants (P01 and P08) had never received
any cancer therapy before, while all others had a history of
chemotherapeutic treatment. Therefore, the observed changes are
likely to be influenced by previous cycles of chemotherapy. For
example, the significantly lower bacterial abundance in cancer
patients before chemotherapy in comparison to control could be a
consequence of previous treatments. The results presented here
illustrate changes due to a single chemotherapeutic cycle, but cannot
rule out, that these changes occurred as a consequence of several
chemotherapeutic cycles over the course of several years. Six cancer
patients also received antibiotics. These patients were characterized
by significantly elevated abundances of bacteria. This finding
confirms the diagnosis ‘bacterial infection’ for which antibiotic
treatment was prescribed. Clostridium cluster IV PCR-DGGE profiles
revealed a shift in species composition by chemotherapy, and even
more so by antibiotics. Thus we conclude that antimicrobial
treatment significantly reduces the species richness of the Clostridium
the most abundant representative. Van Vliet et al. (2009) tested the
effect of chemotherapy in vitro and showed a direct bacteriostatic
effect of chemotherapeutics on bacterial growth.
Further research is needed to show whether changes in bacterial
colonization play a role in the development and maintenance of
mucosal barrier function, infection and inflammation .
The use of prebiotics, probiotics and bacterial products, such as
butyrate to prevent mucosal barrier injury and its complications
could be a promising concept in restoring impaired functions or
enhancing specific desirable functions of the microbiota. The use
of pre- and probiotics to affect the composition and metabolic
activity of the fecal microbiota in times of cancer chemotherapy
and immunosuppression might be part of future research.
In conclusion, chemotherapy treatment causes changes in fecal
microbiota, which coincide with the development of C. difficile
infection in some patients. These changes in microbiota may have
systemic effects and may contribute to the development of
chemotherapy-induced mucositis, influencing important beneficial
functions of the microbial ecosystem.
Materials and Methods
The Viennese Human Ethics committee (3., Thomas-Klestil-
Platz 8/2) under the chair of Dr. Karin Spacek, approved the
proposal of the project ‘‘Analysis of microbiota in feces of patients
with immunosuppression’’. Votum: EK 07-153-VK, 2008. From
all participants involved in the study written consent was obtained.
Study participants and study design
Seventeen subjects receiving ambulant chemotherapy with or
without antimicrobial therapy (aged 59613 y, BMI 2766) from
the Sozialmedizinisches Zentrum Ost (SMZ Ost) in Vienna and
seventeen healthy individuals (aged 65618 y, BMI 2465) joined
this study. Four fecal samples within two weeks were collected of
each ambulant oncology patient in order to collect samples before
and after a single immune-suppressive chemotherapy cycle. The
four samples obtained from every patient were grouped into three
groups: samples taken before the day of chemotherapy (T0),
samples taken 1–4 days after chemotherapy (T1) and samples
taken 5–9 days after chemotherapy (T2). Healthy individuals also
donated four samples during two weeks. Gender ratio among
healthy controls was 56% female, 44% male. Oncology patients
were 47% female and 53% male. Three out of seventeen patients
(P04, P08, P13) had never received any chemotherapy before,
while the others had a history of chemotherapy. Anonymous
medical records reported types of malignancies as well as
chemotherapeutic and antimicrobial treatment as shown in table 2.
We interviewed all study participants assessing age, gender,
body length, weight, health status (chronic and acute diseases), and
life-style aspects such as alcohol consumption and physical activity.
Dietary habits were assessed using a food frequency questionnaire.
Exclusion criteria for healthy controls were (a) antimicrobial
medication (b) chemotherapeutic treatment and (c) pre- and
probiotics at least three months before sample collection. Approval
for this study was obtained from the Viennese Human Ethics
committee (3., Thomas-Klestil-Platz 8/2).
Stool sample processing
After collection, study participants immediately stored their
samples at -18uC in their homes. Stool samples were still frozen
when brought to the laboratory and then immediately stored at
270uC. A 200 mg aliquot of each sample was treated twice for
45 s in a bead-beater (Mini-Beadbeater-8). Thereafter DNA was
extracted using the QIAampH DNA Stool Mini Kit (QIAGEN)
following the manufacturer’s protocol. The DNA was stored at
220uC until analysis.
We used type strains, known to be part of the human
gastrointestinal microbiota and cloned sequences to design a
DGGE standard lane marker. Type strains Bacteroides thetaiotaomi-
cron DSM 2079T, Enterococcus faecium DSM 20477T, Lactobacillus
reuteri ATCC 55730T, Bifidobacterium longum ssp. longum DSM
20097T, Escherichia coli IMBH 252/07 and clones CL16 and CC34
(see below) were used for creating a comparable standard lane
marker for DGGE gels analyzing all bacteria. E.coli IMBH 242/07
gave 4 bands due to its multiple operons for the 16S rRNA gene.
To create a standard lane marker for DGGE analysis and to
identify members of the Clostridium cluster XIVa we constructed
clone libraries from two stool samples of healthy volunteers. For
this purpose PCR products amplified with primers 195-F  and
Chemotherapy Changes Fecal Microbiota
PLoS ONE | www.plosone.org7December 2011 | Volume 6 | Issue 12 | e28654
Ccocc-R  were inserted into a p-GEM Easy Vector (Promega)
following the instructions of the manufacturer. Nucleotide
sequences were corrected for primer and vector sequences in
CodonCodeAligner (www.codoncode.com) and taxonomically
identified using the online tools of the ribosomal database project
(http://rdp.cme.msu.edu/). The clone library used for creating a
standard lane marker for DGGE analysis of Clostridium cluster IV
has previously been described . Clones CL16 (Clostridium leptum
16) and CC34 (Clostridium coccoides 34) were also used as positive
controls in Taqman qPCR.
Polymerase chain reaction (PCR)
PCR was carried out amplifying 16S rRNA gene sequences
from bacteria in fecal samples, type strains and cloned sequences
for DGGE analysis as well as for creation of the clone library using
group- and kingdom- specific primers (table 3). The PCR reaction
mixture consisted of ready-to-use mastermix (Promega) with
1.5 mM MgCl2, 500 nM of primers and 2 ml of template DNA.
When amplifying fecal samples, bovine serum albumin (Fermen-
tas) was added to a final concentration of 400 mg/ml. We used a
Robocycler (Stratagene) for all amplifications.
DGGE was performed as previously described . Primer
pairs and annealing temperatures to analyze the diversity of (a) all
bacteria, (b) Clostridium cluster IV and (c) Clostridium cluster XIVa
are described in table 3. PCR products were separated by
polyacrylamide gels with a denaturing gradient of 30–60% for
predominant bacteria, 30–50% for Clostridium cluster IV and 35–
50% for Clostridium cluster XIVa. Electrophoresis was performed
for 9 h at 129 V at 60uC (predominant bacteria), 5 h at 200 V at
60uC (Clostridium cluster IV) and 7 h at 200 V at 60uC (Clostridium
cluster XIVa). Standard lane markers were created for each DGGE
analysis assay to ensure reliable gel-to-gel comparison. These
standard lane markers (described above) were loaded in triplicate
on each gel to adjust for gradient-variations between gels. We
analyzed PCR-DGGE fingerprints using GelComparII (www.
applied-maths.com). When generating the band comparison, a 1%
tolerance was selected. Principal components analysis (PCA) was
applied on quantitative band comparison datasets in ‘R’ (www.r-
project.org) using the default settings. Shannon diversity index was
calculated on quantitative band information as well with the
default settings implemented in the ‘vegan’ package in ‘R’ (www.r-
project.org). Shannon index is defined as H=2g pi ln pi, where
pi is the proportional abundance of species i. In short, the higher
the Shannon index is, the higher is the diversity. For interpretation
of results, samples were grouped into three groups: samples taken
before the day of chemotherapy (T0), samples taken 1–4 days after
chemotherapy (T1) and samples taken 5–9 days after chemother-
Quantitative TaqMan qPCR
The abundance of bacteria and bacterial subgroups was
measured by 16S rRNA gene-targeting TaqMan qPCR in a
Rotorgene 3000 (Corbett Life Science). Primers, annealing
temperatures and expected product sizes are shown in table 4.
Each sample was analyzed in duplicate. Amplifications were
carried out in a total volume of 10 ml consisting of 5 ml Taq-Man
SensiMix DNA Kit (Quantance), 1 ml of each primer and Taq-
Man probe (concentrations see table 4) and 10 ng of bacterial
Table 2. Relevant clinical data of study participants undergoing immunesuppressive chemotherapy.
name diagnosischemotherapeutic treatmentantimicrobial treatment other condition
P01 urothel carcinoma gemcitabine, cisplatinum
P02 plasmocytoma, multiple myelomabortezomib, dexamethasonerheumatism
fever at 4thsampling
P03Non-Hodgkin lymphoma bendamustinediabetes II, adipositas,
P04 ovarian fibroma taxol, carboplatinlevofloxacin
P05 multiple myeloma bortezomib, doxorubicin, dexamethasonecotrimoxazoleosteoporosis
P06 mamma carcinoma pegylated liposomal doxorubicin hydrochloride,
P07Non-Hodgkin lymphomahigh dose radiation therapy and PBSCT cotrimoxazole, piperacillin,
fever at 2ndand 3rdsampling
P08 monozytic leukemiacytarabine, idarubicin cotrimoxazole, piperacillin,
fever at 2ndsampling
P09 acute leukemiahigh dose Ara-C, radiated erythrocyte concentrate piperacillin, tazobactam fever at 2ndsampling
P10Non-Hodgkin lymphomaifosamid, etoposid, methotrexat levofloxacin
P11Acute lymophoblastic leukemiacytarabine, methotrexatadipositas, hypertension
P12small intestinal tumorcetuximab
P13rectal tumorcapecitabine, oxaliplatin
P14 thymus tumor taxol, carboplatin, bevacizumab, radiation
P15 Acute lymophoblastic leukemiacyclophosphamide, methotrexate, doxorubicin,
P16Acute lymophoblastic leukemiacytarabine, mitoxantrone
P17colon tumor oxaliplatin, capecitabine, bevacizumab, irinotecan,
PBSCT… peripheral blood stem cell transplant.
Chemotherapy Changes Fecal Microbiota
PLoS ONE | www.plosone.org8December 2011 | Volume 6 | Issue 12 | e28654
DNA. Amplification programs included an initial denaturation at
95uC for 10 min followed by 40 cycles consisting of denaturation
at 95uC for 30 s, annealing at 55uC (all bacteria, Clostridium cluster
IV), 56uC (Clostridium cluster XIVa), 58uC (C. difficile) or 60uC
(Bacteroides, bifidobacteria) for 30 s and extension at 72uC for 50 s.
We used tenfold serial DNA dilutions of type strains Bacteroides
thetaiotaomicronT, Bifidobacterium longum ssp. longumTand C. difficile as
well as cloned sequences and one fecal sample to construct
standard curves for comparison of PCR reaction efficiencies
among different experiments.
We quantified DNA of Bacteroides thetaiotaomicronT, Bifidobacterium
longum ssp. longumTand C. difficile, using the nanodrop method
and calculated DNA copies/ml through mean G+C content of
each strain. Clones CL16 and CC34 were amplified with the SP6
Promoter Primer (Promega, Cat.# Q5011) and the T7 Promoter
Primer (Promega, Cat.# Q5021) and the PCR product quantified
using a nanodrop machine. Knowing the sequences of these two
PCR products and their flanking vector sequences we could
quantify the copy numbers and use it as standards. Relative
percentages of bacterial subgroups were calculated in relation to
total rRNA gene copies amplified with primer pair BAC-338-F
and BAC-805-R .
We reviewed sensitivity of PCR reactions with stepwise dilutions
of standard curve DNA until we achieved sensitive detection levels
of PCR. The specificity was confirmed using non-target DNA.
High throughput sequencing
In total, four samples (P09-T0, P09-T1, P11-T0, P11-T1) were
amplified with primer 525F (59- TCAGCAGCCGCGGTAATAC
-39) and 926R (59-TCCGTCAATTCCTTTGAGTTT -39) using
a high-fidelity DNA polymerase (PhusionH, Finnzymes, Thermo
Fisher Scientific) and submitted to 454 barcode sequencing
(AGOWA, Berlin, Germany), resulting in a total of 113 000
reads. The sequences were trimmed and aligned using the pyro
Table 3. 16S rRNA gene primers used for PCR-DGGE fingerprinting.
Target organism Primer Sequence (59-39) Ann. temp (6C)Reference
All bacteria27f GTGCTGCAGAGAGTTTGATCCTGGCTCAG57 [52 T., Blocker 1989]
985r GTAAGGTTCTTCGCGTT 57 
341f-GC CCT ACG GGA GGC AGC AG 55
518r ATT ACC GCG GCT GCT GG55 
Clostridium cluster IVsg-Clept-F-GCGCA CAA GCA GTG GAG T55
sg-Clept-R3CTT CCT CCG TTT TGT CAA
Clostridium cluster XIVaCcocc-F-GC AAATGACGGTACCTGACTAA55
Ccocc-R CTTTGAGTTTCATTCTTGCGAA 
Table 4. Primers and probes used for TaqMan qPCR quantification of 16S rRNA genes.
Target organism Primer and probeSequence (59 - 39) Size (bp)Conc. (nM)Reference
Bifidobacterium spp.Forward primer GCG TGC TTA ACA CAT GCA AGT C125300
Reverse primerCAC CCG TTT CCA GGA GCT ATT 300
Probe (FAM)- TCA CGC ATT ACT CAC CCG TTC GCC -(BHQ-1)150 
BacteroidesAllBac296f GAG AGG AAG GTC CCC CAC 106300
AllBac412rCGC TAC TTG GCT GGT TCA G300
AllBac375Bhqr (FAM)-CCA TTG ACC AAT ATT CCT CAC TGC TGC CT-(BHQ-1)100
All bacteria BAC-338-F ACT CCT ACG GGA GGC AG468 1000
BAC-805-R GAC TAC CAG GGT ATC TAA TCC1000
BAC-516-P(FAM)-TGC CAG CAG CCG CGG TAA TAC-(BHQ-1) 200
Clostridium cluster IV sg-Clept-F GCA CAA GCA GTG GAG T239 400
CTT CCT CCG TTT TGT CAA400
(FAM)-AGG GTT GCG CTC GTT-(BHQ-1) 200 This study
Clostridium cluster XIVa 195FGCA GTG GGG AAT ATT GCA 500
CcoccRCTT TGA GTT TCA TTC TTG CGA A 500
Clostridium diffficile CdiffFTTG AGC GAT TTA CTT CGG TAA AGA 1000
CdiffRTGT ACT GGC TCA CCT TTG ATA TTC A1511000
CdiffP(6-FAM)-CCA CGC GTT ACT CAC CCG TCC G-(BHQ-1) 200
Chemotherapy Changes Fecal Microbiota
PLoS ONE | www.plosone.org9December 2011 | Volume 6 | Issue 12 | e28654
pipeline of the ribosomal database project (http://rdp.cme.msu.
edu/). Only sequences longer than 150 bp were retained, resulting
in 3886 to 6811 sequences per sample with the average lengths of
366 to 368 bp. All analyses were performed using the online tools
of the ribosomal database project pyro pipeline (http://rdp.cme.
msu.edu/). Results of the phylogenetic classification are shown as a
heatmap . The Peptostreptococcaceae, harbouring also C.difficile,
from all four datasets were analyzed in more detail using the online
tools of the ribosomal database project. 100% similar sequences
were grouped and their abundances shown together with a
Statistical evaluation of differences between groups (chemother-
apy and control) and changes within the chemotherapy group (all
time points before and after chemotherapy) was carried out using
the OriginPro version 8 (OriginLab, Northampton, MA). For two
group comparisons of independent ordinal and interval values we
used the two-sample t-test and the nonparametric Mann-Whitney
U-test. For the analysis of related data we used the paired sample
t-test or the non-parametric Wilcoxon signed-rank test. P values
,0.05 were considered statistically significant. To show the
decline in abundance immediately after chemotherapy qPCR
results were plotted in heatmaps . Values were z-scored for
presentation in this heatmap showing changes over time rather
than absolute abundances.
We assessed the participants’ dietary habits using a food
frequency questionnaire. All study participants (patients and
controls) were omnivores and showed similar consumption
patterns of liquids, alcohol, fruits, vegetables, grains and milk
products. Healthy controls stated more frequent consumption of
fruits, whole grain products and alcohol several times a week
compared to patients receiving chemotherapy.
We thank all the study participants at the SMZO for their cooperation. We
thank Dr. Viviana Klose and Mag. Varity-Ann Sattler of IfA Tulln for
their guidance on using the GelCompareII program for the analysis of
DGGE fingerprinting. Furthermore we would like to thank Dr. Guadalupe
Pinar and Dr. Katja Sterflinger for their input and Dr. Konrad Domig for
cultivation of type strains.
Conceived and designed the experiments: JZ RR AGH. Performed the
experiments: JZ AP CL BH OJS. Analyzed the data: JZ CL BH AP OJS
RR AGH. Contributed reagents/materials/analysis tools: EK MR RR.
Wrote the paper: JZ AGH.
1. Turroni F, Ribbera A, Foroni E, van Sinderen D, Ventura M (2008) Human gut
microbiota and bifidobacteria: from composition to functionality. Antonie Van
Leeuwenhoek 94: 35–50.
2. Vrieze A, Holleman F, Zoetendal EG, de Vos WM, Hoekstra JB, et al. (2010)
The environment within: how gut microbiota may influence metabolism and
body composition. Diabetologia 53: 606–613.
3. Chung H, Kasper DL (2010) Microbiota-stimulated immune mechanisms to
maintain gut homeostasis. Curr Opin Immunol 22: 455–460.
4. Backhed F, Ley RE, Sonnenburg JL, Peterson DA, Gordon JI (2005) Host-
bacterial mutualism in the human intestine. Science 307: 1915–1920.
5. Ley RE, Hamady M, Lozupone C, Turnbaugh PJ, Ramey RR, et al. (2008)
Evolution of mammals and their gut microbes. Science 320: 1647–1651.
6. Dethlefsen L, Relman DA (2011) Incomplete recovery and individualized
responses of the human distal gut microbiota to repeated antibiotic perturbation.
Proc Natl Acad Sci U S A Mar 15;108 Suppl 1: 4554–61.
7. De La Cochetiere MF, Durand T, Lepage P, Bourreille A, Galmiche JP, et al.
(2005) Resilience of the dominant human fecal microbiota upon short-course
antibiotic challenge. J Clin Microbiol 43: 5588–5592.
8. Qin J, Li R, Raes J, Arumugam M, Burgdorf KS, et al. (2010) A human gut
microbial gene catalogue established by metagenomic sequencing. Nature 464:
9. Liu C, Finegold SM, Song Y, Lawson PA (2008) Reclassification of Clostridium
coccoides, Ruminococcus hansenii, Ruminococcus hydrogenotrophicus, Rumi-
nococcus luti, Ruminococcus productus and Ruminococcus schinkii as Blautia
coccoides gen. nov., comb. nov., Blautia hansenii comb. nov., Blautia
hydrogenotrophica comb. nov., Blautia luti comb. nov., Blautia producta comb.
nov., Blautia schinkii comb. nov. and description of Blautia wexlerae sp. nov.,
isolated from human faeces. Int J Syst Evol Microbiol 58: 1896–1902.
10. Maukonen J, Matto J, Suihko ML, Saarela M (2008) Intra-individual diversity
and similarity of salivary and faecal microbiota. J Med Microbiol 57:
11. Eckburg PB, Bik EM, Bernstein CN, Purdom E, Dethlefsen L, et al. (2005)
Diversity of the human intestinal microbial flora. Science 308: 1635–1638.
12. Louis P, Scott KP, Duncan SH, Flint HJ (2007) Understanding the effects of diet
on bacterial metabolism in the large intestine. J Appl Microbiol 102: 1197–1208.
13. Schiffrin EJ, Blum S (2002) Interactions between the microbiota and the
intestinal mucosa. Eur J Clin Nutr 56 Suppl 3: S60–64.
14. Cani PD, Neyrinck AM, Fava F, Knauf C, Burcelin RG, et al. (2007) Selective
increases of bifidobacteria in gut microflora improve high-fat-diet-induced
diabetes in mice through a mechanism associated with endotoxaemia.
Diabetologia 50: 2374–2383.
15. Guarner F, Malagelada JR (2003) Gut flora in health and disease. Lancet 361:
16. Nyhlen A, Ljungberg B, Nilsson-Ehle I, Nord CE (2002) Impact of combinations
of antineoplastic drugs on intestinal microflora in 9 patients with leukaemia.
Scand J Infect Dis 34: 17–21.
17. van Vliet MJ, Tissing WJ, Dun CA, Meessen NE, Kamps WA, et al. (2009)
Chemotherapy treatment in pediatric patients with acute myeloid leukemia
receiving antimicrobial prophylaxis leads to a relative increase of colonization
with potentially pathogenic bacteria in the gut. Clin Infect Dis 49: 262–270.
18. Koning CJ, Jonkers DM, Stobberingh EE, Mulder L, Rombouts FM, et al.
(2008) The effect of a multispecies probiotic on the intestinal microbiota and
bowel movements in healthy volunteers taking the antibiotic amoxycillin.
Am J Gastroenterol 103: 178–189.
19. Croswell A, Amir E, Teggatz P, Barman M, Salzman NH (2009) Prolonged
impact of antibiotics on intestinal microbial ecology and susceptibility to enteric
Salmonella infection. Infect Immun 77: 2741–2753.
20. Sonis ST (1998) Mucositis as a biological process: a new hypothesis for the
development of chemotherapy-induced stomatotoxicity. Oral Oncol 34: 39–43.
21. Stringer AM, Gibson RJ, Logan RM, Bowen JM, Yeoh AS, et al. (2009)
Irinotecan-induced mucositis is associated with changes in intestinal mucins.
Cancer Chemother Pharmacol 64: 123–132.
22. Stringer AM, Gibson RJ, Logan RM, Bowen JM, Yeoh AS, et al. (2007)
Chemotherapy-induced diarrhea is associated with changes in the luminal
environment in the DA rat. Exp Biol Med (Maywood) 232: 96–106.
23. Gibson RJ, Keefe DM (2006) Cancer chemotherapy-induced diarrhoea and
constipation: mechanisms of damage and prevention strategies. Support Care
Cancer 14: 890–900.
24. Bergmans DC, Bonten MJ, Gaillard CA, van Tiel FH, van der Geest S, et al.
(1997) Indications for antibiotic use in ICU patients: a one-year prospective
surveillance. J Antimicrob Chemother 39: 527–535.
25. Schalk E, Bohr UR, Konig B, Scheinpflug K, Mohren M (2001) Clostridium
difficile-associated diarrhoea, a frequent complication in patients with acute
myeloid leukaemia. Ann Hematol 89: 9–14.
26. Abd El-Atti S, Wasicek K, Mark S, Hegazi R (2009) Use of Probiotics in the
Management of Chemotherapy-Induced Diarrhea: A Case Study.
JPEN J Parenter Enteral Nutr Sep–Oct 33(5): 569–70.
27. Muyzer G, Smalla K (1998) Application of denaturing gradient gel
electrophoresis (DGGE) and temperature gradient gel electrophoresis (TGGE)
in microbial ecology. Antonie Van Leeuwenhoek 73: 127–141.
28. Benus RF, van der Werf TS, Welling GW, Judd PA, Taylor MA, et al. (2010)
Association between Faecalibacterium prausnitzii and dietary fibre in colonic
fermentation in healthy human subjects. Br J Nutr. pp 1–8.
29. Cucchiara S, Iebba V, Conte MP, Schippa S (2009) The microbiota in
inflammatory bowel disease in different age groups. Dig Dis 27: 252–258.
30. Sokol H, Pigneur B, Watterlot L, Lakhdari O, Bermudez-Humaran LG, et al.
(2008) Faecalibacterium prausnitzii is an anti-inflammatory commensal
bacterium identified by gut microbiota analysis of Crohn disease patients. Proc
Natl Acad Sci U S A 105: 16731–16736.
31. Willing B, Halfvarson J, Dicksved J, Rosenquist M, Jarnerot G, et al. (2009)
Twin studies reveal specific imbalances in the mucosa-associated microbiota of
patients with ileal Crohn’s disease. Inflamm Bowel Dis 15: 653–660.
Chemotherapy Changes Fecal Microbiota
PLoS ONE | www.plosone.org10 December 2011 | Volume 6 | Issue 12 | e28654
32. Grobner S, Fritz E, Schoch F, Schaller M, Berger AC, et al. (2010) Lysozyme Download full-text
activates Enterococcus faecium to induce necrotic cell death in macrophages.
Cell Mol Life Sci Oct 67(19): 3331–44.
33. Ferreira da Silva M, Tiago I, Verissimo A, Boaventura RA, Nunes OC, et al.
(2006) Antibiotic resistance of enterococci and related bacteria in an urban
wastewater treatment plant. FEMS Microbiol Ecol 55: 322–329.
34. de Regt MJ, Willems RJ, Hene RJ, Siersema PD, Verhaar HJ, et al. (2010)
Effects of Probiotics on Acquisition and Spread of Multi-resistant Enterococci.
Antimicrob Agents Chemother Jul 54(7): 2801–5.
35. Woting A, Clavel T, Loh G, Blaut M (2010) Bacterial transformation of dietary
lignans in gnotobiotic rats. FEMS Microbiol Ecol Jun 72(3): 507–14.
36. Flythe MD, Aiken GE (2010) Effects of hops (Humulus lupulus L.) extract on
volatile fatty acid production by rumen bacteria. J Appl Microbiol Oct 109(4):
37. Marquet P, Duncan SH, Chassard C, Bernalier-Donadille A, Flint HJ (2009)
Lactate has the potential to promote hydrogen sulphide formation in the human
colon. FEMS Microbiol Lett 299: 128–134.
38. Fanci R, Corti G, Bartoloni A, Tortoli E, Mariottini A, et al. (2010) Unusual
Methylobacterium fujisawaense Infection in a Patient with Acute Leukaemia
Undergoing Hematopoietic Stem Cell Transplantation: First Case Report. Case
Report Med 2010: 313514.
39. Rettedal E, Vilain S, Lindblom S, Lehnert K, Scofield C, et al. (2009) Alteration
of the ileal microbiota of weanling piglets by the growth-promoting antibiotic
chlortetracycline. Appl Environ Microbiol 75: 5489–5495.
40. Falk A, Olsson C, Ahrne S, Molin G, Adawi D, et al. (2007) Ileal pelvic pouch
microbiota from two former ulcerative colitis patients, analysed by DNA-based
methods, were unstable over time and showed the presence of Clostridium
perfringens. Scand J Gastroenterol 42: 973–985.
41. Collado MC, Derrien M, Isolauri E, de Vos WM, Salminen S (2007) Intestinal
integrity and Akkermansia muciniphila, a mucin-degrading member of the
intestinal microbiota present in infants, adults, and the elderly. Appl Environ
Microbiol 73: 7767–7770.
42. Taras D, Simmering R, Collins MD, Lawson PA, Blaut M (2002)
Reclassification of Eubacterium formicigenerans Holdeman and Moore 1974
as Dorea formicigenerans gen. nov., comb. nov., and description of Dorea
longicatena sp. nov., isolated from human faeces. Int J Syst Evol Microbiol 52:
43. Nomura T, Ohkusa T, Okayasu I, Yoshida T, Sakamoto M, et al. (2005)
Mucosa-associated bacteria in ulcerative colitis before and after antibiotic
combination therapy. Aliment Pharmacol Ther 21: 1017–1027.
44. Goodman MJ, Kent PW, Truelove SC (1977) Glucosamine synthetase activity of
the colonic mucosa in membranous colitis. Gut 18: 229–231.
45. Guilmeau S, Flandez M, Bancroft L, Sellers RS, Tear B, et al. (2008) Intestinal
deletion of Pofut1 in the mouse inactivates notch signaling and causes
enterocolitis. Gastroenterology 135: 849–860, 860 e841–846.
46. Meier H, Amann R, Ludwig W, Schleifer KH (1999) Specific oligonucleotide
probes for in situ detection of a major group of gram-positive bacteria with low
DNA G+C content. Syst Appl Microbiol 22: 186–196.
47. Matsuki T, Watanabe K, Fujimoto J, Takada T, Tanaka R (2004) Use of 16S
rRNA gene-targeted group-specific primers for real-time PCR analysis of
predominant bacteria in human feces. Appl Environ Microbiol 70: 7220–7228.
48. Zwielehner J, Liszt K, Handschur M, Lassl C, Lapin A, et al. (2009) Combined
PCR-DGGE fingerprinting and quantitative-PCR indicates shifts in fecal
population sizes and diversity of Bacteroides, bifidobacteria and Clostridium
cluster IV in institutionalized elderly. Exp Gerontol 44: 440–446.
49. Muyzer G, de Waal EC, Uitterlinden AG (1993) Profiling of complex microbial
populations by denaturing gradient gel electrophoresis analysis of polymerase
chain reaction-amplified genes coding for 16S rRNA. Appl Environ Microbiol
50. Yu Y, Lee C, Kim J, Hwang S (2005) Group-specific primer and probe sets to
detect methanogenic communities using quantitative real-time polymerase chain
reaction. Biotechnol Bioeng 89: 670–679.
51. Symons S, Zipplies C, Battke F, Nieselt K (2010) Integrative systems biology
visualization with MAYDAY. J Integr Bioinform Mar 25 7(3.
52. Edwards U, Rogall T, Blocker H, Emde M, Bottger EC (1989) Isolation and
direct complete nucleotide determination of entire genes. Characterization of a
gene coding for 16S ribosomal RNA. Nucleic Acids Res 17: 7843–7853.
53. Heuer H, Hartung K, Wieland G, Kramer I, Smalla K (1999) Polynucleotide
probes that target a hypervariable region of 16S rRNA genes to identify bacterial
isolates corresponding to bands of community fingerprints. Appl Environ
Microbiol 65: 1045–1049.
54. Neefs JM, Van de Peer Y, De Rijk P, Goris A, De Wachter R (1991)
Compilation of small ribosomal subunit RNA sequences. Nucleic Acids Res 19
55. Shen J, Zhang B, Wei G, Pang X, Wei H, et al. (2006) Molecular profiling of the
Clostridium leptum subgroup in human fecal microflora by PCR-denaturing
gradient gel electrophoresis and clone library analysis. Appl Environ Microbiol
56. Penders J, Vink C, Driessen C, London N, Thijs C, et al. (2005) Quantification
of Bifidobacterium spp., Escherichia coli and Clostridium difficile in faecal samples of
breast-fed and formula-fed infants by real-time PCR. FEMS Microbiol Lett 243:
Chemotherapy Changes Fecal Microbiota
PLoS ONE | www.plosone.org11 December 2011 | Volume 6 | Issue 12 | e28654