THE EFFECTS OF ORAL PROBIOTIC SUPPLEMENTS ON THE

THE EFFECTS OF ORAL PROBIOTIC SUPPLEMENTS ON THE HUMAN GUT
MICROBIOME
By
Erin Kyle Hudnall
A thesis submitted to the faculty of The University of Mississippi in partial fulfillment of
the requirements of the Sally McDonnell Barksdale Honors College.
Oxford
May 2016
Approved by
X
Advisor: Dr. Colin Jackson
X
Reader: Dr. Sarah Liljegren
X
Reader: Dr. Erik Hom
©2016
Erin Kyle Hudnall
ALL RIGHTS RESERVED
ii
ACKNOWLEDGEMENTS
First, I would like to thank my advisor, Dr. Colin Jackson. Your willingness in
allowing me to work in your lab and your patience and guidance during this process was
essential to its success. I am beyond grateful for the time and effort you spent on this
project. Next, I want to thank the faculty of The University of Mississippi, particularly
those of the Sally McDonnell Barksdale Honors College, for going above and beyond the
call of duty to contribute to my education. I want to thank my fiancé, Jordan Baldwin,
who has kept me motivated throughout my time at Ole Miss and who refused to allow me
to abandon my dream of attending medical school. Most especially, I would like to thank
my parents, Charlie and Merrill Hudnall, for supporting me throughout my life, and for
their continuing support as I pursue a career in medicine. The love, patience, and
encouragement each of you has shown to me are the foundation of who I am.
iii
ABSTRACT
Erin Kyle Hudnall: The Effects of Oral Probiotic Supplements on the Human Gut
Microbiome
(Under the direction of Colin R. Jackson, Ph.D.)
A diverse bacterial community makes up the human gut microbiome and is
essential not only to digestion, but also to other physiological processes. This bacterial
community is subject to change due to external factors including, but not limited to, age,
diet, and lifestyle. This study observed the effects of oral probiotic supplements on this
bacterial community. Fecal samples were collected from a single subject once each week
during alternating two- to six-week cycles of taking and abstaining from the probiotic
supplement. The samples were purified, the bacterial cells within lysed, and the enclosed
DNA was collected. A portion of the 16S rRNA gene was amplified and sequenced.
Sequences were identified and compared to determine whether there were measurable
effects of taking the probiotic on the subject’s gut microbial community. Firmicutes and
Bacteroidetes were the dominant bacterial Phyla, with smaller proportions of
Proteobacteria and Actinobacteria. Fluctuations in the subject’s gut microbiome were
seen at the phylum level and at multiple taxonomic levels within these phyla from the
start to the end of the study, but did not generally correspond to cycles of taking the
probiotic. Furthermore, the relative abundance of Bacillus coagulans, the bacterial
species within the probiotic supplement, did not change dramatically. This study
iv
showed that taking this particular probiotic supplement did not cause substantial changes
in the relative abundances of bacterial taxonomic groups in the human gut, and that the
fluctuations seen were likely due to other factors – for example, normal and expected
variations in the individual’s gut bacteria, intestinal permeability caused by the subject’s
food allergies, or other external influences.
v
TABLE OF CONTENTS
LIST OF TABLES AND FIGURES…………………………………………………….vii
INTRODUCTION…………………………………………………………………….…..1
METHODS………………………………………………………………………………..6
RESULTS………………………………………………………………………………..13
DISCUSSION……………………………………………………………………………25
LIST OF REFERENCES………………………………………………………………...29
vi
LIST OF TABLES AND FIGURES
Table 1
Schedule of fecal sample collection for the subject involved in the
study.………………………………………………………………………8
Table 2
Summary of the various commands within the software package, mothur,
that were used to analyze data in this study, and their intended purpose for
analysis..………………………………………………………………….12
Figure 1
Image of an agarose gel demonstrating the presence of DNA isolated from
representative stool samples……………………………………………..16
Table 3
Bacterial community composition of fecal samples obtained from a human
subject, prior to taking probiotic supplements…………………………...17
Figure 2
Changes in the relative abundance of Phylum Firmicutes (a) and the
families within Order Clostridia (part of Phylum Firmicutes; b) in bacterial
communities in stool samples taken from an individual during cycles of
abstaining from and taking a probiotic supplement……………………...18
Figure 3
Changes in the relative abundance of Phylum Bacteroidetes in bacterial
communities in stool samples taken from an individual during cycles of
abstaining from and taking a probiotic supplement……………………...19
vii
Figure 4
Changes in the relative abundance of the species within Family
Bacteroidaceae (Phylum Bacteroidetes). Percentages of an unclassified
species (a), of Bacteroides caccae (b), of Bacteroides ovatus (c), and of
Bacteroides fragilis (d) in bacterial communities in stool samples taken
from an individual during cycles of abstaining from and taking a probiotic
supplement……………………………………………………………….20
Figure 5
Changes in the relative abundance of Phylum Proteobacteria (a), the
Classes within the phylum (b), and the orders within one of those classes
(Gammaproteobacteria; c) in bacterial communities in stool samples taken
from an individual during cycles of abstaining from and taking a probiotic
supplement……………………………………………………………….22
Figure 6
Changes in the relative abundance of Phylum Actinobacteria (a) and the
species within Order Bifidobacteriales (part of Phylum Actinobacteria; b)
in bacterial communities in stool samples taken from an individual during
cycles of abstaining from and taking a probiotic supplement……………23
Table 4
Average percentage of Clostridia and of total bacteria made up of Bacillus
coagulans in fecal samples obtained from a human subject, during
alternating cycles “off” probiotic supplement and “on” probiotic
supplement……………………………………………………………….24
viii
INTRODUCTION
There are trillions of bacteria, representing more than a thousand species, living
on and within the human body. It is estimated that bacterial cells out number human cells
in the body by a factor of ten to one (Ackerman 2012). These bacteria perform many
important functions in metabolism and defense (Hopkins et al. 2001). For example,
bacteria typical of the ‘normal’ human intestinal community assist the body in metabolic,
nutritional, and immunological functions including degradation of certain food
components, production of vitamins, and production of certain digestive enzymes
(Holzapfel et al. 1998). These bacteria exert crucial metabolic activities by fermenting
non-digestible polysaccharides such as fibers and starches into energy substrates (shortchain fatty acids) for the benefit of both the microbes themselves as well as the host
(Blaser 2014). These metabolic activities also lead to the production of vitamin K,
vitamin B12, folic acid, and amino acids, substances which humans are unable to produce
themselves. In addition, the intestinal microbiome participates in defending the body
against pathogens through colonization resistance, the production of antimicrobial
compounds, and contributing to the mucosal immune system (Gerritsen et al. 2011).
The large intestine is by far the most heavily colonized portion of the digestive
12
tract, containing up to 10 bacterial cells per gram of gut contents (Gibson et al. 1995).
The bacteria within the large intestine base their metabolism on the nutrients a person
consumes, causing a diverse bacterial community to be present in this organ. It is
1
estimated that the large intestine of healthy adults is host to approximately 500 different
species belonging to more than 190 genera (Ansell 2011).
Bacterial communities in the human gut are predominately composed of members
of Phylum Bacteroidetes, which typically account for 17-60% of identified sequences and
Phylum Firmicutes, which account for approximately 35-80% (Shoaie et al. 2013).
Phylum
Bacteroidetes
consists
of
four
classes:
Bacteroidia,
Flavobacteria,
Sphingobacteria, and Cytophagia (Thomas et al. 2011). These classes range in metabolic
type from the strictly anaerobic Bacteroidia to the solely aerobic Flavobacteria (Thomas
et al. 2011). Bacteroidia are the dominant class of Bacteroidetes found in the human large
intestine, because their obligate anaerobic metabolism allows them to thrive in the
oxygen-poor environmental conditions (Thomas et al. 2011).
The other major phylum that dominates the human large intestinal community,
Firmicutes, is currently the largest bacterial phylum, containing more than 200 genera.
The majority of the Firmicutes detected in the human intestinal tract fall primarily into
two groups, Clostridium coccoides (also known as Clostridium cluster XIVa) and
Clostridium leptum (also known as Clostridium cluster IV). The bacterial species falling
into each of these groups are obligate anaerobes capable of producing endospores
(Gerritsen et al. 2011). A larger concentration of Firmicutes in the gut has been found to
be associated with obesity. It is theorized that this correlation is due their production of
excess energy from nutrients that have been consumed (Fujimura et al. 2010). Other
important bacterial phyla in the human gut include the Actinobacteria (Fujimura et al.
2010) and various Proteobacteria. Each of these organisms plays some role as part of the
microbial community in the human large intestine (Shoaie et al. 2013).
2
The variable nature of gut bacteria allows individuals to be studied for differences
and similarities on various levels such as age, geography, or diet. Diet and environmental
factors begin to affect the gut microbial community beginning from birth. Vaginally
delivered infants show higher levels of Bifidobacterium and Bacteroides while infants
born via Cesarean section exhibited a gut microbial community dominated by
Staphylococcus, Streptococcus, and Clostridium difficile (Fujimura et al. 2010). In one
study, infants who were exclusively fed a formula diet showed a greater abundance of
Clostridium difficile and Escherichia coli than those infants that were breastfed only. The
two groups maintained similar abundances of Bifidobacteria; however, the breastfed
infants showed greater gene expression within these Bifidobacteria, which may have
allowed them to metabolize a greater variety of complex oligosaccharides (Fujimura et al.
2010).
External factors continue to contribute to the gut microbiota throughout human
life. A change in diet at any time can result in changes in the relative abundances of
major phyla of gut bacteria, which could lead to diseases such as obesity (Turnbaugh et
al. 2009). The ratio of Firmicutes to Bacteroidetes in lean subjects (3:1) has been reported
to be enhanced by a factor of ten in obese subjects (up to 35:1; Fujimura et al. 2010).
While a higher ratio of Firmicutes to Bacteroidetes tends to be correlated with obesity
(Fujimura et al. 2010), it may be that a decrease in the amount of Bacteroidetes, rather
than an actual increase in Firmicutes, could be responsible for the increased ratio between
these phyla in obese subjects (Armougom et al. 2009). Aging also plays a role in the
variability of intestinal bacteria, causing structural changes in the microbiota. This
process affects the proportion of protective Bifidobacteria (members of the
3
Actinobacteria; Garrity et al. 2004) leading to major effects on innate colonization
resistance (Hopkins et al. 2001). Parameters, such as nutrient consumption or antibiotic
use, can be manipulated for comparison among individuals or within a single individual.
Analyzing the connection between gut bacteria and the human host has lead to
discoveries that may further advances in human wellness (Kinross et al. 2011).
Each bacterial species has a specific 16S rRNA gene sequence (Woese 1987).
Modern approaches to microbial ecology rely on sequencing this variable region to
identify the specific types of bacteria present and to estimate overall bacterial diversity
(De Santis et al. 2006). The use of this 16S rRNA gene sequencing technique is important
to the study of microbial diversity in the human gut, as this technique removes the need
to culture bacteria in order to identify them. This aspect is particularly important because
many of the intestinal bacteria, such as species of Class Bifidobacteria, are anaerobic and
therefore difficult to culture using standard approaches (Turnbaugh et al. 2009). Thus,
16S rRNA gene sequencing is a valuable tool for the identification and comparison of
intestinal bacteria, allowing for diverse studies concerning the gut microbiome; next
generation sequencing of regions of this gene have helped increase our knowledge of the
gut microbial community over the last decade.
In recent years, there has been interest in improving intestinal health and digestion
by modulating the composition of the gut bacterial community through introduction of a
potentially remedial community (Collins et al. 1999). Attempts have been made to
modulate the indigenous intestinal microbiome through the ingestion of live microbial
adjuncts, called ‘probiotics.’ Although various definitions have been proposed to describe
these products, Havenaar et al. (1992) first suggested a definition according to which
4
probiotics are defined as “mono- or mixed cultures of live microorganisms which, when
applied, beneficially affect the host by improving the properties of the indigenous
microflora” (Holzapfel et al. 1998). The most widely used and accepted definition,
however, describes probiotics as “live microbial food supplements that beneficially affect
the host animal by improving its intestinal microbial balance” (Collins et al. 1999).
These definitions encompass over-the-counter preparations that contain
lyophilized bacteria for adult human use. The microorganisms included in these
preparations are often lactic acid producers such as Lactobacilli and Bacilli (Phylum
Firmicutes) as well as Bifidobacteria (Collins et al. 1999). Some of the most important
functional effects of the bacterial species introduced into the gut through probiotic
supplements include aspects such as immune modulation and strengthening the gut
mucosal barrier (Holzapfel et al. 1998). An effective probiotic should meet certain
criteria. These include exertion of a beneficial effect on the host, containing a large
number of viable cells, and capability of surviving and metabolizing in the gut. In
addition, probiotic supplements must remain viable during storage and use and be
nonpathogenic and nontoxic (Collins et al. 1999).
This study monitored the composition of the large intestinal bacterial community
of a single individual in an attempt to determine the effects of consuming an over-thecounter probiotic supplement on the gut microbiota. Stool samples were obtained once a
week over a period of eight months, within which the subject alternated between
approximately six-week “on” cycles of consuming the supplement and “off” cycles of
abstaining. The gut microbiome composition was characterized by 16S rRNA gene
sequencing, facilitating analysis of this diverse community.
5
METHODS
Background Information
This study was conducted on a single subject – a 21-year-old female
approximately 1.57 meters tall. The subject lives a lightly active lifestyle and has a diet
limited by allergies, thus avoiding large quantities of wheat, rice, and soy. The
commercial probiotic supplement taken was Schiff® Digestive Advantage® Probiotic
Gummies, distributed by Reckitt Benckiser (Parsippany, New Jersey) which were
acquired over-the-counter at a Wal-Mart store in Oxford, Mississippi, on March 18, 2015.
The bacterial species contained in this supplement was listed to be Bacillus coagulans,
with each gummy containing 250 million viable cells at the time of manufacture. The
subject cycled through two- to six-week periods of consuming and abstaining from the
supplement. During “on” cycles, the subject consumed the maximum dose (four
gummies) each morning at approximately 9:00 AM.
Sample Collection
Stool samples (approximately 0.1 g) were collected from the subject each Sunday,
beginning March 22 and ending November 29, 2015. Each sample was collected
immediately following regular defecation using a sterile swab. In order to reduce possible
contamination of the sample by bacteria on the skin, care was taken to avoid skin contact.
6
The sample was then aseptically transferred to a sterile collection tube, and immediately
frozen (-20°C) and stored until all samples had been collected.
7
Table 1. Schedule of fecal sample collection for the subject involved in the study. For
each sample, the distinction between “on” cycles of consuming probiotic supplements
and “off” cycles of abstaining from the supplements is noted.
Sample ID
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
Date
22-Mar
29-Mar
5-Apr
12-Apr
19-Apr
26-Apr
3-May
10-May
17-May
24-May
31-May
7-Jun
9-Aug
16-Aug
23-Aug
30-Aug
6-Sep
13-Sep
20-Sep
27-Sep
4-Oct
11-Oct
18-Oct
25-Oct
1-Nov
8-Nov
15-Nov
22-Nov
29-Nov
8
Cycle
Off
Off
Off
On
On
On
On
On
On
On
Off
Off
On
On
On
On
On
On
Off
Off
Off
Off
Off
Off
On
On
On
On
On
DNA Extraction
Samples were allowed to thaw to room temperature. Bacterial DNA was then
extracted from the samples using a Mo Bio PowerFecal™ DNA Isolation kit, following
the detailed protocol provided by the manufacturer (Mo Bio Laboratories; Carlsbad,
California), and described as follows. Thawed samples were loaded into tubes containing
sterile garnet beads and a buffer solution to aide in dissolving and dispersing the sample
particles during homogenization and cell lysis. Sodium dodecyl sulfate (SDS) was added
to help break down fatty acids and lipids in bacterial cell membranes, and the tubes were
heated at 65°C for 10 minutes. The tubes were then agitated for 10 minutes to cause the
beads to collide with the cells and help lysis (“bead beating”). Tubes were then
centrifuged and the supernatant retained and sequentially treated with proprietary
inhibitor removers (Mo Bio Laboratories). A high concentration salt solution was then
added to the final DNA solution, which was then bound to a silica filter. The bound DNA
was rinsed with ethanol to further remove impurities. A sterile elution buffer was then
added to the filter to solubilize and release the bound DNA from the silica filter. The
presence of DNA was verified by agarose gel electrophoresis (220 V, 25 minutes) and the
purified DNA samples transferred to sterile tubes, and immediately frozen (-20°C) and
stored until all samples were prepared for sequencing.
DNA Sequencing
The hypervariable V4 region of the bacterial 16S rRNA was amplified and
sequenced using paired-end, barcoded Illumina MiSeq next generation sequencing
(Kozich et al. 2013). This sequencing procedure was conducted at the Molecular and
9
Genomics Core Facility at the University of Mississippi Medical Center (UMMC) in
Jackson, MS. The resulting 16S rRNA sequence data were subsequently downloaded as
FASTQ files and assessed via the bioinformatics software package, mothur (Schloss et al.
2009) using the recommended procedures according to Schloss et al. (2011) and Kozich
et al. (2013) as follows.
Raw FASTQ data resulting from two separate reads (R1 and R2) during the
sequencing process of each sample were merged to generate high quality sequence data.
Multiple copies of identical unique sequences were consolidated to enable faster data
processing. The sequences were then compared against pre-aligned reference sequences
found in the SILVA database (Quast et al. 2013), and screened based on their length and
on potential ambiguity between bases. This alignment process was used to account for
potential gaps and obtain a better classification, verifying that the sequences were truly
collected from a portion of the 16S rRNA gene. Sequences containing runs of greater
than eight identical sequential bases were removed, as they could be indicators of
potential sequencing error. Sequences differing by two or fewer bases were then clustered
together to remove potential amplification artifacts, as these could represent noise from
sequencing or PCR error rather than actual genetic variation. Chimeras were checked
for and eliminated using the incorporated UCHIME software (Edgar et al. 2011).
These are unreal sequences that originated from more than one initial sequence,
havingbeencombinedduringamplificationduetoincompleteextensionagainstone
template then completion of the extension against a different template. The
remaining sequences were then classified using the Greengenes database (Desantis et al.
10
2006). Contaminant sequences – those from chloroplasts or mitochondria, for example –
were removed from the dataset.
11
Table 2. Summary of the various commands within the software package, mothur, that
were used to analyze data in this study, and their intended purpose for analysis.
Command
Function
Make.contigs
Initial Processing
Screen.seqs
Screen data for length errors
Unique.seqs
Filters out identical sequences to reduce processing time
Count.seqs
Compress unique sequences and samples together
Align.seqs
Aligns sequences to an established database
Filter.seqs
Filters out non-informative gaps
Pre.cluster
Clusters almost identical sequences together
Chimera.uchime
Identifies chimeras within sequences
Remove.seqs
Removes chimeras
Classify.seqs
Classifies remaining sequences according to GreenGenes
12
RESULTS
Stool samples were obtained and bacterial DNA extracted and analyzed according
to the aforementioned procedures. The presence of DNA in select samples was verified
via gel electrophoresis (Figure 1), and the samples proved suitable for 16S rRNA gene
sequencing. Sequencing was successful and yielded a total of 539,134 valid bacterial
sequences, which classified into 518 species belonging to 435 genera, across twentyseven samples. Prior to the addition of any probiotic supplement, Phylum Firmicutes
dominated the dataset, accounting for 65.7% of the total sequences, followed by Phylum
Bacteroidetes, accounting for 25.5%. Sequences identified as Proteobacteria and
Actinobacteria were also fairly prevalent and represented 4.6% and 4.1% of the total
number of sequences, respectively (Table 3).
The overall percentages of Phylum Firmicutes showed a decreasing trend during
each successive cycle of taking the probiotic supplement (Figure 2a); the mean
percentage of Firmicutes to overall bacteria during the first “on” cycle was 59.5%, during
the second was 45.4%, and was 21.2% during the third. Class Clostridia dominated this
phylum, making up an average of 99.11% of the Firmicutes throughout all cycles. The
order Clostridiales made up 100% of this class, and was made up of four families. Family
Lachnospiraceae was the dominant family within the Clostridiales except for the sample
taken on August 16, while Ruminococcaceae was the second most dominant family
except for the same sample, within which the relative abundances of these two families
were reversed. Family Veillonellaceae and an unclassified family also made up a small
13
portion of this class (Figure 2b).
The Bacteriodetes exhibited no major changes in their proportional abundance
over the course of the study (Figure 3). The Bacteroidetes were dominated by Class
Bacteroidia (99.7%), which was made up entirely of Order Bacteroidales, of which
99.9% of the bacteria fell into Family Bacteroidaceae and 100% of those belonged to the
genus Bacteroides. Four main species made up this genus – an unclassified species
(Figure 4a), Bacteroides caccae (Figure 4b), Bacteroides ovatus (Figure 4c), and
Bacteroides fragilis (Figure 4d). These species showed no major changes in relative
abundance throughout any of the cycles.
Phylum Proteobacteria showed a clear surge in their relative abundance during the
final cycle of taking the probiotic supplement (Figure 5a). Three classes made up nearly
the entirety of this phylum – Class Betaproteobacteria, Class Deltaproteobacteria, and
Class Gammaproteobacteria. Betaproteobacteria dominated the phylum during the first
and second “on” cycles, except for a dip on May 17, but showed a marked decrease
during the third. The Deltaproteobacteria remained at a relatively stable, low level
throughout the course of the study. The Gammaproteobacteria were low during the first
two “on” cycles, except for a spike on May 17, but showed a dramatic increase to
dominate the phylum during the final cycle of taking the probiotic supplement (Figure
5b). The three orders within Gammaproteobacteria were Order Enterobacteriales,
composed entirely of Escherichia coli, Order Pasteurellales, composed of 93.0%
Haemophilus parinfluenzae, and Order Pseudomonadales, made up 66.8% by an
unclassified species belonging to the genus Pseudomonas and 33.2% by Acinetobacter
guilloulae (Figure 5c).
14
Phylum Actinobacteria showed a possible slight decreasing trend in overall
percentages with each successive cycle, but overall did not show a distinguishable trend
based on the cycles (Figure 6a). The two major classes, Actinobacteria (93.53%) and
Coriobacteria (6.47%), were slightly variable but again showed no trend based on the
probiotic cycles. Within Class Actinobacteria the dominant order was Bifidobacteriales
(96.5%), with 100% of the bacteria within falling into Family Bifidobacteriaceae, and
100% of that family made up of the genus Bifidobacterium. An unclassified species
dominated the genus at 94.99%, and the species Bifidobacterium bifidum made up 0.05%;
among these species there was no distinguishable trend, save for a spike in
Bifidobacterium bifidum in the November 22 sample (Figure 6b).
During the “off” cycles of abstaining from the probiotic supplement, the species
Bacillus coagulans made up 0.00% of the Clostridia and of the total gut bacteria,
increasing slightly during the “on” cycles (Table 4). This species made up 92.39% of the
bacterial sequences in the probiotic supplement, with the remaining portion composed of
20 other sequence types; however, Bacillus coagulans was never very prevalent in the gut
community. Overall, there was no clear distinction in the gut microbiome between cycles
of taking and not taking the probiotic, and the specific bacteria within the probiotic never
became dominant members of the gut bacterial community.
15
Figure 1. Image of an agarose gel demonstrating the presence of DNA isolated from
representative stool samples (samples 5-12, noted by numbers above each lane).
16
Table 3. Bacterial community composition of fecal samples obtained from a human
subject, prior to taking probiotic supplements. Samples are listed by date (Day-Month
2015). Numbers represent the number of sequences obtained that classified into that
taxon, with corresponding percentages of the total from that sample below them.
Taxon
22-March
29-March
5-April
Mean
9,043
11,771
10,864
10,559
69.94%
69.05%
58.07%
65.7%
3,066
3,663
5,856
4,195
23.71%
21.49%
31.30%
25.5%
317
793
1275
795
2.45%
4.65%
6.82%
4.6%
498
786
693
659
3.85%
4.61%
3.70%
4.1%
Phylum Firmicutes
Phylum Bacteroidetes
Phylum Proteobacteria
Phylum Actinobacteria
17
(a)
PecentagesofPhylumFirmicutes
100
90
80
Percentage
70
60
50
40
30
20
8-Nov
22-Nov
29-Nov
22-Nov
29-Nov
1-Nov
1-Nov
15-Nov
25-Oct
25-Oct
15-Nov
18-Oct
18-Oct
8-Nov
4-Oct
11-Oct
11-Oct
27-Sep
4-Oct
20-Sep
6-Sep
13-Sep
30-Aug
23-Aug
9-Aug
16-Aug
2-Aug
26-Jul
19-Jul
5-Jul
12-Jul
28-Jun
21-Jun
7-Jun
14-Jun
31-May
24-May
17-May
3-May
10-May
26-Apr
19-Apr
5-Apr
12-Apr
29-Mar
0
22-Mar
10
Date
(b)
PercentagesofFamiliesWithinOrderClostridiales
PercentagesofFamiliesWithinOrderClostridia
80
70
Percentage
60
50
40
30
20
27-Sep
20-Sep
13-Sep
6-Sep
30-Aug
23-Aug
16-Aug
9-Aug
2-Aug
26-Jul
19-Jul
12-Jul
5-Jul
28-Jun
21-Jun
14-Jun
7-Jun
31-May
24-May
17-May
10-May
3-May
26-Apr
19-Apr
12-Apr
5-Apr
29-Mar
0
22-Mar
10
Date
Lachnospiraceae
Ruminococcaceae
Veillonellaceae
unclassified
Figure 2. Changes in the relative abundance of Phylum Firmicutes (a) and the families
within Order Clostridiales (part of Phylum Firmicutes; b) in bacterial communities in
stool samples taken from an individual during cycles of abstaining from (unshaded areas)
and taking (shaded areas) a probiotic supplement.
18
PercentagesofPhylumBacteroidetes
70
60
Percentage
50
40
30
20
29-Nov
22-Nov
8-Nov
15-Nov
1-Nov
25-Oct
18-Oct
4-Oct
11-Oct
27-Sep
20-Sep
6-Sep
13-Sep
30-Aug
23-Aug
9-Aug
16-Aug
2-Aug
26-Jul
19-Jul
5-Jul
12-Jul
28-Jun
21-Jun
7-Jun
14-Jun
31-May
24-May
17-May
3-May
10-May
26-Apr
19-Apr
5-Apr
12-Apr
29-Mar
0
22-Mar
10
Date
Figure 3. Changes in the relative abundance of Phylum Bacteroidetes in bacterial
communities in stool samples taken from an individual during cycles of abstaining from
(unshaded areas) and taking (shaded areas) a probiotic supplement.
19
0
Date
20
29-Nov
22-Nov
15-Nov
8-Nov
1-Nov
25-Oct
18-Oct
11-Oct
4-Oct
27-Sep
20-Sep
13-Sep
6-Sep
30-Aug
23-Aug
16-Aug
9-Aug
2-Aug
26-Jul
19-Jul
12-Jul
5-Jul
28-Jun
21-Jun
14-Jun
7-Jun
31-May
24-May
17-May
10-May
3-May
26-Apr
19-Apr
12-Apr
5-Apr
29-Mar
(b)
29-Nov
22-Nov
15-Nov
8-Nov
1-Nov
25-Oct
18-Oct
11-Oct
4-Oct
27-Sep
20-Sep
13-Sep
6-Sep
30-Aug
23-Aug
16-Aug
9-Aug
2-Aug
26-Jul
19-Jul
12-Jul
5-Jul
28-Jun
21-Jun
14-Jun
7-Jun
31-May
24-May
17-May
10-May
3-May
26-Apr
19-Apr
12-Apr
5-Apr
29-Mar
22-Mar
0
22-Mar
Percentage
Percentage
(a)
100
PercentagesofBacteroidesunclassified
90
80
70
60
50
40
30
20
10
Date
PercentagesofBacteroidescaccae
25
20
15
10
5
PercentagesofBacteroidesovatus
(c)
20
18
16
Percentage
14
12
10
8
6
4
8-Nov
15-Nov
22-Nov
29-Nov
22-Nov
29-Nov
1-Nov
1-Nov
15-Nov
25-Oct
25-Oct
8-Nov
18-Oct
18-Oct
4-Oct
11-Oct
27-Sep
20-Sep
6-Sep
13-Sep
30-Aug
23-Aug
9-Aug
16-Aug
2-Aug
26-Jul
19-Jul
5-Jul
12-Jul
28-Jun
21-Jun
7-Jun
14-Jun
31-May
24-May
17-May
3-May
10-May
26-Apr
19-Apr
5-Apr
12-Apr
29-Mar
0
22-Mar
2
Date
PercentagesofBacteroidesfragilis
(d)
10
9
8
Percentage
7
6
5
4
3
2
11-Oct
4-Oct
27-Sep
20-Sep
13-Sep
6-Sep
30-Aug
23-Aug
16-Aug
9-Aug
2-Aug
26-Jul
19-Jul
12-Jul
5-Jul
28-Jun
21-Jun
14-Jun
7-Jun
31-May
24-May
17-May
10-May
3-May
26-Apr
19-Apr
5-Apr
12-Apr
29-Mar
0
22-Mar
1
Date
Figure 4. Changes in the relative abundance of the species within Family Bacteroidaceae
(Phylum Bacteroidetes). Percentages of an unclassified species (a), of Bacteroides caccae
(b), of Bacteroides ovatus (c), and of Bacteroides fragilis (d) in bacterial communities in
stool samples taken from an individual during cycles of abstaining from (unshaded areas)
and taking (shaded areas) a probiotic supplement.
21
Percentage
100
90
80
70
60
50
40
30
20
10
0
Enterobacteriales
Figure 5. Changes in the relative abundance of Phylum Proteobacteria (a), the Classes
Pasteurellales
22
29-Nov
22-Nov
15-Nov
8-Nov
1-Nov
25-Oct
18-Oct
11-Oct
4-Oct
27-Sep
20-Sep
13-Sep
6-Sep
30-Aug
23-Aug
Deltaproteobacteria
16-Aug
9-Aug
2-Aug
26-Jul
19-Jul
12-Jul
Betaproteobacteria
5-Jul
28-Jun
21-Jun
14-Jun
7-Jun
31-May
90
80
70
60
50
40
30
20
10
13-Sep
6-Sep
30-Aug
23-Aug
16-Aug
9-Aug
2-Aug
26-Jul
19-Jul
12-Jul
5-Jul
28-Jun
21-Jun
14-Jun
7-Jun
31-May
24-May
17-May
10-May
3-May
26-Apr
19-Apr
12-Apr
5-Apr
29-Mar
22-Mar
22-Nov
29-Nov
22-Nov
29-Nov
1-Nov
8-Nov
1-Nov
25-Oct
15-Nov
25-Oct
18-Oct
8-Nov
18-Oct
11-Oct
15-Nov
4-Oct
11-Oct
4-Oct
27-Sep
PercentagesofClassesWithinProteobacteria
20-Sep
100
27-Sep
Date
20-Sep
13-Sep
6-Sep
30-Aug
23-Aug
16-Aug
9-Aug
2-Aug
26-Jul
19-Jul
12-Jul
5-Jul
28-Jun
21-Jun
14-Jun
7-Jun
31-May
24-May
17-May
10-May
3-May
26-Apr
19-Apr
12-Apr
5-Apr
(b)
24-May
17-May
10-May
3-May
26-Apr
19-Apr
(c)
12-Apr
5-Apr
29-Mar
22-Mar
0
29-Mar
0
22-Mar
Percentage
Percentage
(a)
PercentagesofPhylumProteobacteria
45
40
35
30
25
20
15
10
5
Date
Gammaproteobacteria
PercentagesofOrdersWithinClassGammaproteobacteria
Date
Pseudomonadales
within the phylum (b), and the orders within one of those classes (Gammaproteobacteria;
c) in bacterial communities in stool samples taken from an individual during cycles of
abstaining from (unshaded areas) and taking (shaded areas) a probiotic supplement.
PercentagesofPhylumAc4nobacteria
(a)
10
9
8
Percentage
7
6
5
4
3
2
29-Nov
22-Nov
8-Nov
15-Nov
1-Nov
25-Oct
18-Oct
4-Oct
11-Oct
27-Sep
20-Sep
6-Sep
13-Sep
30-Aug
23-Aug
9-Aug
16-Aug
2-Aug
26-Jul
19-Jul
5-Jul
12-Jul
28-Jun
21-Jun
7-Jun
14-Jun
31-May
24-May
17-May
3-May
10-May
26-Apr
19-Apr
5-Apr
12-Apr
29-Mar
0
22-Mar
1
Date
(b)
PercentagesofSpeciesWithinOrderBifidobacteriales
100
Percentage
80
60
40
29-Nov
22-Nov
15-Nov
8-Nov
1-Nov
25-Oct
18-Oct
11-Oct
4-Oct
27-Sep
20-Sep
13-Sep
6-Sep
30-Aug
23-Aug
16-Aug
9-Aug
2-Aug
26-Jul
19-Jul
12-Jul
5-Jul
28-Jun
21-Jun
14-Jun
7-Jun
31-May
24-May
17-May
10-May
3-May
26-Apr
19-Apr
12-Apr
5-Apr
29-Mar
0
22-Mar
20
Date
Bifidobacteriumbifidum
Bifidobacteriumunclassified
Figure 6. Changes in the relative abundance of Phylum Actinobacteria (a) and the species
within Order Bifidobacteriales (part of Phylum Actinobacteria; b) in bacterial
communities in stool samples taken from an individual during cycles of abstaining from
(unshaded areas) and taking (shaded areas) a probiotic supplement.
23
Table 4. Average percentage of Clostridia and of total bacteria made up of Bacillus
coagulans in fecal samples obtained from a human subject, during alternating cycles
“off” (unshaded areas) probiotic supplement and “on” (shaded areas) probiotic
supplement. Samples are listed by their Sample ID.
Cycle
Percentage of Bacillus
coagulans in Clostridia
Percentage of Bacillus
coagulans in total Bacteria
Samples 1-3
0.000%
0.0000%
Samples 4-10
0.030%
0.0002%
Samples 11-12
0.000%
0.0000%
Samples 13-18
0.001%
0.0000%
Samples 19-24
0.00%
0.0000%
Samples 25-29
0.025%
0.0001%
24
DISCUSSION
This study examined the effects of an over-the-counter probiotic supplement on
the human gut microbiome. A total of twenty-nine samples were taken over
approximately six-week cycles “on” and “off” the supplement. For twenty-seven of these
samples, 16S rRNA sequencing was used to determine the bacterial taxa present and their
relative proportions; the remaining two samples were unsuitable for analysis, possibly
due to low DNA yield from the extraction procedure. This data suggested that changes in
the gut bacterial community did indeed occur, however, these changes were not
immediately attributable to the use of the probiotic supplement. Fluctuations in the
relative abundances of different bacterial populations were expected, although it was
uncertain which taxa would be affected, and at what taxonomic levels the fluctuations
would be seen. It was expected that samples taken during the “on” cycles would differ
from those taken during the “off” cycles, and that the changes in abundances would
follow a trend throughout the course of the study. To some extent, this was the case for
certain bacterial phyla and for various taxa within these phyla; however, fluctuations did
not occur in the proportions of Bacillus coagulans, the live species contained within the
probiotic supplement. It is quite possible that the changes seen were due to expected
variability in gut bacteria or to other external factors, such as those attributable to the
subject’s food allergies.
Sequences classified as members of Phyla Firmicutes and Bacteroidetes were
25
dominant within the samples, followed by sequences identified as Proteobacteria and
Actinobacteria. Bacteroidetes are commonly found in the human gut, and may be
important in digestion as well as interacting with the immune system to limit pathogenic
colonization of the gut (Thomas et al. 2011). This phylum is known for its symbiotic
activity in degrading biopolymers and polysaccharides in the large intestine (Mahowald
et al. 2009, Thomas et al. 2011). Members of Firmicutes are also common with the gut;
these bacteria are Gram-positive, sometimes anaerobic bacteria. The most prevalent
sequences were identified as being in the Class Clostridia, genus Faecalibacterium. The
large representation of this typical intestinal genus supports that the samples included
only intestinal bacteria not those bacteria present on skin of the anus (Miquel et al. 2013).
Sequences identified as being in Phyla Proteobacteria and Actinobacteria were prevalent
in smaller proportions. Specifically, one species of Proteobacteria found in every sample
was Escherichia coli. Because E. coli are well adapted to the conditions of the human
large intestine and well documented as occurring within (Rajilic-Stojanovic et al. 2007),
the presence of this species also supports correct sampling and handling.
Many previous studies have found that diet and other factors affect gut microbe
composition (Muegge et al. 2011, Wu et al. 2011, David et al. 2014). Distribution
between and within the major phyla for the particular community studied was likely
affected by the restrictive nature of the subject’s diet due to allergy limitations on
consumption of common dietary staples wheat, rice, and soy. The lining of a normal
intestinal tract is nearly leak proof and only fully digested food molecules are permitted
to pass through this lining into the bloodstream and lymph vessels. However, this leak
proof lining is only one cell layer thick and can be easily damaged as its cells have a short
26
life span, extremely high metabolic activity, and intense nutritional demands. Damage to
the intestinal lining is detrimental, as the intestines are full of hostile digestive enzymes
and trillions of microorganisms that would cause irritation if allowed to escape into the
bloodstream. If the diet does not contain enough nutrients – as is the case of those with
certain food allergies – to repair the intestinal permeability, it can become a persistent
problem. This permeability, often termed “Leaky Gut Syndrome,” can also be due to
inflammation of the lining that occurs when an allergen is inadvertently ingested
(Brodhead 2002). A combination of the usually expected variability in gut bacteria and a
leaky gut is the most probable cause of the relative changes in abundances of gut
bacterial taxa observed in this study.
Findings of a connection between food allergies and intestinal permeability have
provided the foundation for various intervention studies designed to modify gut microbial
composition for the treatment of allergic disease and for aiding digestion. These studies
were based on the speculation that the flora within probiotic supplements would aid in
digestion, generate important nutrients, stimulate the immune system, and diminish
allergic reactivity, thus diminishing the effects of intestinal permeability. Studies have
shown that breastfed infants have higher amounts of Bifidobacteria in their gut; further
research now suggests that children with adequate quantities of Bifidobacteria are less
likely to develop allergic diseases. Bifidobacteria are commomly included in probiotic
supplements. A strain taken from Swiss cheese, Propionibacterium freudenreichii,
stimulates the growth of various strains of Bifidobacteria, and also shows promise for
delivery through oral supplements (Brodhead 2002). Lactobacillus rhamnosus is another
species that contains some of the most impressive strains of scientifically
27
validated probiotics and can markedly diminish symptoms in those with food allergies
(Brodhead 2002).
The effects of a variety of other beneficial bacteria in probiotic supplements have
been studied, but more stringent causality assessments should be applied to demonstrate
the consistency of the assumed linkage between the gut microbiome and allergies (Yao et
al. 2010). Because little to no detectable increase in Bacillus coagulans was seen
throughout any of the cycles of supplementing with that species, this study determined
that the use of this particular probiotic supplement is not likely to increase the proportion
of Bacillus coagulans within the gut. Further studies may reveal, however, that by
introducing this species, other taxa are enabled to increase or caused to decrease within
the microbial community. Relative abundances of bacteria at various taxonomic levels
showed both positive and negative fluctuations. While possible explanations for these
changes were outlined above, the specific reasons are outside the scope of this study.
Further work could be done to determine whether normal and expected variations in the
individual’s gut bacteria, a leaky gut caused by the subject’s food allergies, or other
external factors caused these fluctuations or whether the introduction of Bacillus
coagulans by means of the supplement indeed caused a “domino effect” on other taxa
within the gut community.
28
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