The Effects of Oral Probiotic Supplements on the Human Gut Microbiome" (2016)

University of Mississippi eGrove Honors College (Sally McDonnell Barksdale Honors Theses Honors College)

2016 The ffecE ts of Oral Probiotic Supplements on the Human Gut Microbiome Erin Hudnall University of Mississippi. Sally McDonnell Barksdale Honors College

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Recommended Citation Hudnall, Erin, "The Effects of Oral Probiotic Supplements on the Human Gut Microbiome" (2016). Honors Theses. 328. https://egrove.olemiss.edu/hon_thesis/328

This Undergraduate Thesis is brought to you for free and open access by the Honors College (Sally McDonnell Barksdale Honors College) at eGrove. It has been accepted for inclusion in Honors Theses by an authorized administrator of eGrove. For more information, please contact [email protected]. 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

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 (short- chain 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-the- counter 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 Date Cycle 1 22-Mar Off 2 29-Mar Off 3 5-Apr Off 4 12-Apr On 5 19-Apr On 6 26-Apr On 7 3-May On 8 10-May On 9 17-May On 10 24-May On 11 31-May Off 12 7-Jun Off 13 9-Aug On 14 16-Aug On 15 23-Aug On 16 30-Aug On 17 6-Sep On 18 13-Sep On 19 20-Sep Off 20 27-Sep Off 21 4-Oct Off 22 11-Oct Off 23 18-Oct Off 24 25-Oct Off 25 1-Nov On 26 8-Nov On 27 15-Nov On 28 22-Nov On 29 29-Nov On

8 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, having been combined during amplification due to incomplete extension against one 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 twenty- seven 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.

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 Phylum Firmicutes

69.94% 69.05% 58.07% 65.7%

3,066 3,663 5,856 4,195 Phylum Bacteroidetes

23.71% 21.49% 31.30% 25.5%

317 793 1275 795

Phylum Proteobacteria

2.45% 4.65% 6.82% 4.6%

498 786 693 659

Phylum Actinobacteria

3.85% 4.61% 3.70% 4.1%

(a) Pecentages of Phylum Firmicutes 100 90 80 70

60 50

Percentage 40 30 20 10 0 5-Jul 7-Jun 4-Oct 5-Apr 6-Sep 2-Aug 9-Aug 12-Jul 19-Jul 26-Jul 1-Nov 8-Nov 3-May 14-Jun 21-Jun 28-Jun 11-Oct 18-Oct 25-Oct 13-Sep 20-Sep 27-Sep 12-Apr 19-Apr 26-Apr 16-Aug 23-Aug 30-Aug 15-Nov 22-Nov 29-Nov 22-Mar 29-Mar 10-May 17-May 24-May 31-May Date

(b) Percentages of Families Within Order Clostridiales Percentages of Families Within Order Clostridia 80

30 Percentage 20 10 0 5-Jul 7-Jun 4-Oct 5-Apr 6-Sep 2-Aug 9-Aug 12-Jul 19-Jul 26-Jul 1-Nov 8-Nov 3-May 14-Jun 21-Jun 28-Jun 11-Oct 18-Oct 25-Oct 13-Sep 20-Sep 27-Sep 12-Apr 19-Apr 26-Apr 16-Aug 23-Aug 30-Aug 15-Nov 22-Nov 29-Nov 22-Mar 29-Mar 10-May 17-May 24-May 31-May 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 Percentages of Phylum Bacteroidetes 70

30 Percentage

0 5-Jul 7-Jun 4-Oct 5-Apr 6-Sep 2-Aug 9-Aug 12-Jul 19-Jul 26-Jul 1-Nov 8-Nov 3-May 14-Jun 21-Jun 28-Jun 11-Oct 18-Oct 25-Oct 13-Sep 20-Sep 27-Sep 12-Apr 19-Apr 26-Apr 16-Aug 23-Aug 30-Aug 15-Nov 22-Nov 29-Nov 22-Mar 29-Mar 10-May 17-May 24-May 31-May 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 (a) Percentages of Bacteroides unclassiﬁed 100

50 Percentage 40

0 5-Jul 7-Jun 4-Oct 6-Sep 5-Apr 2-Aug 9-Aug 12-Jul 19-Jul 26-Jul 1-Nov 8-Nov 3-May 14-Jun 21-Jun 28-Jun 11-Oct 18-Oct 25-Oct 12-Apr 19-Apr 26-Apr 13-Sep 20-Sep 27-Sep 16-Aug 23-Aug 30-Aug 15-Nov 22-Nov 29-Nov 22-Mar 29-Mar 10-May 17-May 24-May 31-May Date

(b) Percentages of Bacteroides caccae 25

15 Percentage 10

20 (c) Percentages of Bacteroides ovatus 20

10 Percentage 8

(d) Percentages of Bacteroides fragilis 10

5 Percentage 4

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 (a) Percentages of Phylum Proteobacteria 45

20 Percentage 15

(b) Percentages of Classes Within Proteobacteria 100

Percentage 40

0 5-Jul 7-Jun 4-Oct 6-Sep 5-Apr 2-Aug 9-Aug 12-Jul 19-Jul 26-Jul 1-Nov 8-Nov 3-May 14-Jun 21-Jun 28-Jun 11-Oct 18-Oct 25-Oct 12-Apr 19-Apr 26-Apr 13-Sep 20-Sep 27-Sep 23-Aug 30-Aug 16-Aug 15-Nov 22-Nov 29-Nov 22-Mar 29-Mar 24-May 31-May 10-May 17-May Date

Betaproteobacteria Deltaproteobacteria Gammaproteobacteria

(c) Percentages of Orders Within Class Gammaproteobacteria 100 90 80 70 60 50 40 Percentage 30 20 10 0 5-Jul 7-Jun 4-Oct 6-Sep 5-Apr 2-Aug 9-Aug 12-Jul 19-Jul 26-Jul 1-Nov 8-Nov 14-Jun 21-Jun 28-Jun 3-May 25-Oct 11-Oct 18-Oct 12-Apr 19-Apr 26-Apr 13-Sep 20-Sep 27-Sep 16-Aug 23-Aug 30-Aug 15-Nov 22-Nov 29-Nov 22-Mar 29-Mar 10-May 17-May 24-May 31-May Date

Enterobacteriales Pasteurellales Pseudomonadales

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 (unshaded areas) and taking (shaded areas) a probiotic supplement.

22 (a) Percentages of Phylum Ac4nobacteria 10 9 8 7 6 5

Percentage 4 3 2 1 0 5-Jul 7-Jun 4-Oct 5-Apr 6-Sep 2-Aug 9-Aug 12-Jul 19-Jul 26-Jul 1-Nov 8-Nov 3-May 14-Jun 21-Jun 28-Jun 11-Oct 18-Oct 25-Oct 13-Sep 20-Sep 27-Sep 12-Apr 19-Apr 26-Apr 16-Aug 23-Aug 30-Aug 15-Nov 22-Nov 29-Nov 22-Mar 29-Mar 10-May 17-May 24-May 31-May Date

(b) Percentages of Species Within Order Biﬁdobacteriales 100

40 Percentage

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.

Percentage of Bacillus Percentage of Bacillus Cycle coagulans in Clostridia 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.

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