The Fecal Bacteria

THE Fecal Bacteria Edited by Michael J. Sadowsky Department of Soil,Water, and Climate and BioTechnology Institute, Un...

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THE

Fecal Bacteria Edited by

Michael J. Sadowsky Department of Soil,Water, and Climate and BioTechnology Institute, University of Minnesota, St. Paul, Minnesota

Richard L. Whitman Lake Michigan Ecological Research Station, Great Lakes Science Center, U.S. Geological Survey, Porter, Indiana

ASM

PRESS

Washington, DC

Address editorial correspondence to ASM Press, 1752 N St. N W , Washington, DC 20036-2904, USA Send orders to ASM Press, P.O. Box 605, Hemdon, VA 20172, USA Phone: 800-546-2416; 703-661-1593 Fax: 703-661-1501 E-mail: [email protected] Online: estore.asm.org Copyright 0 2011 ASM Press American Society for Microbiology 1752 N St. N W Washington, DC 20036-2904

Library of Congress Cataloging-in-Publication Data The fecal bacteria / edited by Michael J. Sadowsky, Richard L. Whitman. p. ; cm. Includes bibliographicalreferences and index. ISBN 978-1-55581-608-7 (hardcover : alk. paper) 1. Enterobacteriaceae. I. Sadowsky, M. J. (MichaelJ.) 11. Whitman, Richard Lincoln. [DNLM: 1. Enterobacteriaceae. 2. Environmental Monitoring-methods. 3. Feces-microbiology. 4. Fresh Wateranalysis. Q W 1381 QR82.E6F43 2011 579.3'46~22 2010029320 Current printing (last digit) 10 9 8 7 6 5 4 3 2 1

All Rights Reserved Printed in the United States of America Cover illustration: Interaction of Salmonella enterica serovar Typhimurium with mouse intestinal tissue. Bacteria are green, and mouse intestine epithelial tissue is blue and red. (Photo courtesy of Barbel Stecher and Wolf-Dietrich Hardt, ETH Zurich, Switzerland.)

CONTENTS

Contributors vii Pre$ace ix

1. The Fecal Environment, the Gut / 1 Denis 0.Krause and Ehsan Khajpour

2.

Taxonomy, Phylogeny, and Physiology of Fecal Indicator Bacteria / 23 Militza Cawero-Coldn, Gene S . Wickham, and Ronald F. Turco

3.

The Gut Microbiota: Ecology and Function / 39 Benjamin P. Willing and Janet K. Jansson

4.

Animals and Humans as Sources of Fecal Indicator Bacteria / 67 Christopher K. Yost, Moussa S . Diawa, and Edward Topp

5.

Environmental Sources of Fecal Bacteria / 93 Muruleedhara N. Byappanahalli and Satoshi Ishii

6.

Physical and Biological Factors Influencing Environmental Sources of Fecal Indicator Bacteria in Surface Water / 111 Richard L. Whitman, Meredith B. Nevers, Katarzyna Przybyla-Kelly, and Muruleedhara N.Byappanahalli

7.

Impacts of Fecal Bacteria on Human and Animal Health-Pathogens and Virulence Genes / 135 TimothyJ.Johnson

8.

Modeling Fate and Transport of Fecal Bacteria in Surface Water / 165 Meredith B. Nevers and Alexandria B. Boehm

V

vi

CONTENTS

9.

Microbial Source Tracking / 189 ValerieJ. Harwood, Hodon Ryu, andJorge Santo Doming0

10. Prevalence and Fate of Gut-Associated Human Pathogens in the Environment / 217 Katherine G. McElhany and Suvesh D. Pillai 11. Classical and Molecular Methods To Measure Fecal Bacteria / 241 Thomas A. Edge and Alexandria B. Boehm 12. Fecal Bacteria and Foods / 275 Francisco Diez- Gonzalez 13. Conclusions and Future Use of Fecal Indicator Bacteria for Monitoring Water Quality and Protecting HumanHealth / 295 MichaelJ. Sadowsky and Richard L. Whitman Index / 303

CONTRIBUTORS

Alexandria €3. Boehm Environmental and Water Studies, Department of Civil & Environmental Engineering, Jerry Yang & Akiko Yamazaki Environment & Energy Building, Stanford, CA 94305

Muruleedhara N. Byappanahalli U.S. Geological Survey, Great Lakes Science Center, Lake Michigan Ecological Research Station, Porter, IN 46304

Militza Carrero-Col6n Department of Agronomy, Purdue University, West Lafayette, IN 47907

Moussa S. Diarra Pacific Agn-Food Research Centre, Agriculture and Agr-Food Canada, Agassiz, BC, Canada VOM IAO

Francisco Diez-Gonzalez Department of Food Science and Nutrition, University of Minnesota, St. Paul, M N 55108

Thomas A. Edge Water Science & Technology Directorate, National Water Research Institute, Environment Canada, Burlington, O N , Canada L7R 4A6

Valerie J. Hanvood Department of Integrative Biology, University of South Florida, Tampa, FL 33620

Satoshi Ishii Department of Applied Biologcal Chemistry, Graduate School of Agricultural and Life Sciences, University of Tokyo, Tokyo, 113-8657, Japan

Janet K. Jansson Department of Ecology, Earth Sciences Division, Lawrence Berkeley National Laboratory, Berkeley, CA 94720

Timothy J. Johnson Department of Veterinary and Biomedical Sciences, University of Minnesota, St. Paul, M N 55108

vii

viii

CONTRIBUTORS

Ehsan Khafipour Department of Animal Science, University of Manitoba, Winnipeg, MB, Canada R3T 2N2

Denis 0. Krause Department of Animal Science and Department of Medical Microbiology, University of Manitoba, Winnipeg, MB, Canada R3T 2N2

Katherine G. McElhany Food Safety & Environmental Microbiology Program, Texas A&M University, College Station, TX 77843

Meredith B. Nevers U.S. Geological Survey, Great Lakes Science Center, Lake Michigan Ecological Research Station, Porter, IN 46304

Suresh D . Pillai Food Safety & Environmental Microbiology Program, Texas A&M University, College Station, T X 77843

Katarzyna Przybyla-Kelly U.S. Geological Survey, Great Lakes Science Center, Lake Michigan Ecological Research Station, Porter, IN 46304

Hodon Ryu ORD/NRMRL/WSWRD/MCCB U.S. Environmental Protection Agency, Cincinnati, OH 45268

Michael J. Sadowsky BioTechnology Institute and Department of Soil, Water, and Climate, University of Minnesota, St. Paul, MN 55108

Jorge Santo Domingo ORD/NRMRL/WSWRD/MCCB U.S. Environmental Protection Agency, Cincinnati, OH 45268

Edward Topp Agriculture and Agr-Food Canada, London, ON, Canada N5V 4T3

Ronald F. Turco Laboratory for Soil Microbiology, Department of Agronomy, Purdue University, West Lafayette, IN 47907

Richard L. Whitman U.S. Geological Survey, Great Lakes Science Center, Lake Michigan Ecological Research Station, Porter, IN 46304

Gene S . Wickham The Center for Environmental Biotechnology, University of Tennessee, Knoxville, T N 37996

Benjamin P. Willing Michael Smith Laboratories, University of British Columbia, Vancouver, BC, Canada V6T 124

Christopher K. Yost Department of Biology, University of Regina, Regina, SK, Canada S4S OA2

PREFACE

A

lthough often maligned, the fecal indicator bacteria, commonly found in the gastrointestinal tract of humans and warm-blooded animals, represent a broad group of enteric bacteria that have had much uthty over the last 40 years as a means to assess water quality and food safety. These bacteria serve as the focal point of “the fecal indicator concept,” used worldwide to monitor water quality and protect people from swimming- and water-related Illnesses. The fecal indicator concept stems, in part, from the pioneering work of Von Fritsch and Escherich who, in the 1800s, made the logical connections between the presence of Klebsiella pneumoniae and K . rhinoscleromatis and sewage contamination and the use of Bacillus coli (subsequently renamed Escherichia colq as an indicator of fecal pollution. Their work eventually led to the development of the inhcator bacterial parahgm as a means to assess microbial aspects of water quality, and the concept was promulgated by the U.S. Public Health Service in their 1914 standards for drinking water that were based on total coliforms. The regulations were based, in large part, on the belief that if these organisms were present in water there was a likelihood that human pathogens would also be present and that microorganisms causing diseases in humans were more likely to originate from human sources (e.g., human feces) than from other animals. Over the years, however, the fecal inmcator concept has moved away from use of t h s complex and large group of organisms, collectively referred to as the coliform bacteria, many of which are members of the f d y Enterobacten’aceae,to reliance on measurements of E. coli and noncoliform bacteria such as enterococci. These bacteria are better related, based on epidemiologicalstuhes, to health risks associated with swimming and recreational activities in fresh and marine waters. It is important to note, however, that the fecal indicator bacteria represent just a small portion of the total fecal bacteria present in the gastrointestinal tracts

ix

x

PREFACE

of warm-blooded animals. While other potential indicators for microbiological surface water and food quality have been suggested, incluhng Clostridiumpegingens, B$dobacteriurn spp., Bacteroides+agilis, coliphages, and enteric viruses, there is currently much debate as to which organism or organisms might best serve this purpose. This is being driven by the realization that some of the currently used bacteria do not meet the ideal requirements of a fecal indicator bacterium, in that they can occur and grow in secondary environmental habitats and persist for a long time in soils, sand, and sedments and in association with plants, algae, and cold-blooded animals. Some of the recent &scoveries have been heled by molecular microbiological methods that not only have provided insight into the taxonomic status of many previously uncultured intestinal bacteria (specifying who is there) but also have yielded a variety of technologies that can be used to idenhf) sources of these organisms in the environment (microbial source tracking methods) and rapidly quantif;l these organisms in secondary habitats (quantitative PCR). One of the primary goals of this book is to provide the reader with a broad understanding and appreciation of the fecal bacteria in terms of their taxonomy, relation to the physiology and anatomy of the host animal, ecology, and use in water quality and food safety issues. This book is organized into 13 chapters that we hope represent the breadth of the fecal bacteria. Whenever possible, the authors have linked these &verse microorganisms to issues of water and food quality. Chapters 1 and 2 provide background on what is thought to be the primary habitat for these microbes, the gut, and how these organisms have been characterized with respect to their taxonomy, phylogeny, and physiology. Since the fecal bacteria in natural habitats do not live in isolation but rather interact with their abiotic and biotic environments, chapter 3 presents information on the ecology of fecal bacteria in the host, obtained by new metagenomic approaches. This technology also reveals that the gut environment is complex and contains many more organisms that have not been revealed by traditional culture-based approaches. Aside from humans, fecal bacteria are also added to natural environments via other hosts; chapters 4 and 5 discuss the contributions of animals and secondary habitats to environmental sources and sinks of fecal bacteria. Chapter 4 specifically examines factors that influence the abundance, composition, and characteristics of fecal bacteria and how these are influenced by the use of antibiotics and other pervasive anthropogenic inputs to watersheds. Little is known about how different environmental factors affect the survival, persistence, and growth of bacteria in ambient environments. These variables, including hydrometeorological factors and transport phenomena, will in turn have significant impacts on water quality. Chapter 6 specifically addresses how these physical and biological factors influence environmental source and fates of fecal indicator bacteria and how this impacts water quality and the use of water for recreational purposes. Along these lines, chapter 8 discusses, in detail, how deterministic and empirical statistical modeling of fecal bacteria in surface waters can be used to better understand sources, fate, and transport of fecal bacteria in the environment. This chapter also dmusses how this information

can be used to predict and noti@ the public of the microbiological quality of water and beaches. While many of the chapters discuss fecal bacteria in relation to their use as indicators of water quality, this is ultimately being done to garner information about the possible presence of enteric pathogens. However, many of the traditional indicator bacteria themselves can be pathogens for humans. Chapter 7 presents a review of what we know about the subset of the Enterobacteriaceae that are bacterial pathogens and how pathogenicity is shaped by the presence of an array of virulence genes in many of these microorganisms. Moreover, chapter 10 examines the available information concerning the prevalence, survival, and fate of these enteric pathogens in different environments. Historically, many of the bacteria that are currently used as inhcators of fecal pollution were chosen based on their ease of growth on traditional laboratory culture media. While this approach has led to the development of some useful selective media that can accurately determine numbers of viable fecal indicator bacteria, the presence of viable but not culturable bacteria and the need for rapid assessment methods have led to the development of new methods, many of which are based on PCR and other molecular tools. Chapter 11 discusses the classical and molecular methods in detail; many of these methods are further discussed in chapter 9 for use in microbial source tracking. A comprehensive review of microbial source tracking, a more recently defined term to describe a variety of phenotypic and genotypic methods that can be used to determine the sources of fecal bacteria in waterways and sand, soil, and sediment matrices, is also presented in chapter 9. Although a whole book on this subject area was published in 2007 (J. W. Santo Doming0 and M. J. Sadowsky, ed., Microbial Source Tracking, ASM Press, Washington, DC), chapter 9 presents a fresh look at this research area and focuses on new tools that can be used for determinations of total maximum daily load and epidemiological analyses. Lastly, although the majority of chapters in this book are centered on fecal bacteria in the environment and waterways, many of the same organisms are also used as indicators of contamination of ready-to-eat foods and fresh produce which may have been impacted by fecal material during postprocessing handling. Consequently, chapter 12 discusses how the selection of appropriate fecal bacteria for assessing food contamination has paralleled those criteria used for water quality assessment. The chapter also hscusses how contamination of foods with fecal bacteria continues to be a major concern for the transmission of food-borne diseases, as has been witnessed in recent food-borne outbreaks. Moreover, the chapter discusses how our requirements for a safe food supply have led to a better understanding of the routes of contamination and the development of more sensitive and reliable detection methods. Many of the same tools and technologies also have direct application to questions concerning contamination of the environment with fecal bacteria. In summary, it is our hope that this book will provide the reader with a more comprehensive understanding of the fecal bacteria. Although much has been learned about this important group of organisms over the last 200 years, much is still unknown. This is especially true of the ecological aspects of many of the fecal bacteria in primary and secondary habitats. While some of

xii

PREFACE

this information has recently been revealed using genomic and metagenomic technologies, we have a lot more to understand with respect to how these bacteria function in the host (what they are doing), how they modulate the host’s immune response and overall well-being, and how they interact with other microorganisms in foods and extraorganismal environments. We thank Greg Payne of ASM Press for his advice and guidance throughout all phases of the editorial process. We also thank Brian Badgley, Meredith Nevers, Dawn Shively, and Muruleedhara Byappanahalli for their useful suggestions, comments, and discussion. MICHAEL J. SADOWSKY RICHARD L. WHITMAN

THE FECAL ENVIRONMENT, THE GUT Denis 0. Krause and Ehsan Kha$pour

(Dinalo and Relman, 2009). It also has significant implications for the sheddmg of pathogens by animals. For example, Escherichiu coli 0157:H7 is thought to reside primarily at the recto-anal junction in cattle and the nature of the det, animal physiology, and disease status influences the shedding of these microorganisms into the manure (Sheng et al., 2006). In this chapter we examine the ecology of the digestive tract of humans and animals with particular reference to the association between anatomical structures, their related physiologies, the mode of feeding, and the functional association of specific microbial communities in different gut compartments of the digestive tract. The reason for spending some time exploring these fascinating hostmicrobial symbioses is to give the reader an appreciation of the anatomical diversity of digestive tract systems and how closely aligned the microbial communities are with these anatomical structures (Fig. 1).

Traditionally, veterinary, animal science, and m e d d students take courses in digestive tract physiology and anatomy. In these courses the mechanical and chemical breakdown of feed and food is examined in great detail and includes a review of mastication in the buccal cavity, the enzymes of the pancreas, and the mucosa. Further, the absorption of nutrients in both the small and large intestines is discussed. One topic that is generally not discussed in great detail is the role that microorganisms play in the digestion, absorption, and general homeostasis of the digestive tract. This omission is fairly surprising when one considers that the number of microorganisms in the digestive tract may be as high as whereas all the eukaryotic cells associated with the human and animal body are in the region of lo9 (Tuohy et al., 2009). Of the whole human and animal microbiome the largest, by far, is the gut microbiome. The gut microbiome is in an intimate symbiosis with the host and this interaction has profound effects on the host

OVERVIEW OF THE ANATOMY AND PHYSIOLOGY OF THE

DIGESTIVE TRACT The purpose of this section is to provide the reader who is not familiar with the digestive tract a brief, nonexhaustive summary of

Denis 0. Krause, Department of Animal Science and Department of Medical Microbiology, University of Manitoba, Winnipeg, MB, Canada R3T 2N2. Ehsan Khafipour, Department of Animal Science, University of Manitoba, Winnipeg, MB, Canada R3T 2N2.

The Fecal Bacten'a, Edited by M. J. Sadowsky and R. L. Whitman, 0 2011 American Society for Microbiology, Washington, DC

1

2 W KRAUSE AND KHAFIPOUR

A Case

I

Diet

Gastric

I1

Diet

Gastric

-

i t

Colonic

Feces

Dog Humans

Colonic

Feces

Pig Horse

Cecal

Ceca I

B

tl

FIGURE 1 (A) Sequence of digestion in nonruminants. The simplest case (little or no cecal digestion) occurs in most carnivoresand in humans. Cecal digestionoccursin large herbivoresbut is secondaryto colon digestion (Stevens and Hume, 1988). (B) Sequence of digestion in rodents, most of which practice coprophagy; cecal fermentation is dominant over secondary colonic fermentation. Some rodents (hamsters and voles) exhibit pregastric digestion (see Fig. 2). Reprinted from Van Soest (1994) with permission from Cornell University Press.

the major features of the digestive tract. An in-depth analysis of this topic is covered in Dukes’ Physiology of Domestic Animals (Dukes and Reece, 2004). The digestive tract in mammals lies within the thoracic and abdominal cavities and stretches from the mouth to the anus. Differentiation between the thoracic and abdominal cavities is made with reference to the diaphragm, which lies at the base of the ribcage. The thoracic cavity consists of the space between the diaphragm and the base of the neck and contains the heart, lungs, and esophagus, plus some other organs like the thymus. The abdominal cavity is circumscribed by the space between the base of the diaphragm and the pelvic rim. The organs of the digestive tract contained within this cavity include the gastric stomach and small

intestines, as well as the large intestines, the rectum, and anus. Also within this cavity are the accessory organs to the digestive tract, including the liver and pancreas. Additional organs in the abdominal cavity are the spleen, which is an organ of the immune system, and the kidneys. The kidneys are not strictly part of the digestive tract but are important for water and mineral homeostasis in the body and come into play when diarrhea occurs. Organs of the reproductive tract are also within the abdominal cavity (Dukes and Reece, 2004). Food or feed is usually broken down into smaller particles in the oral cavity by mastication and grinding by the teeth (Beauchemin, 1991; Allen, 1997). A small amount ofamylase is produced in the saliva so that starch digestion begins in the mouth, but no lipases or proteases

1. THE FECAL ENVIRONMENT, THE GUT

are produced (Pedersen et al., 2002). It is generally accepted that the microorganisms in the oral cavity have little role in food digestion. In recent years, however, the microbial ecology of the oral cavity has been well-studied in the human and it is now recognized that there is a very intricate regional biogeography in the oral cavity (Parahitiyawa et al., 2010). Although this has not been studied in animals to any large extent, this is likely also the case for animals. We do not have clear evidence of the importance of oral microflora on digestive processes, but we do know that microorganisms in the mouth have a role in inoculating the rest of the digestive tract (Yilmaz, 2008). In ruminants, the production of saliva in the mouth can be highly significant and can be as much as 100 liters per day (Beauchemin, 1991; Allen, 1997). The saliva is rich in enzymes and contains bicarbonate. Saliva production is stimulated by chewing and buffers the rumen against low pH conditions. Once food, or digesta, is swallowed it passes along the esophagus, through the diaphragmatic hiatus, and into the gastric stomach. The gastric stomach is generally divided up into the cardiac region (adjacent and distal to the esophagus), the fundus (body of the gastric stomach), and the pyloric region (adjacent to the duodenum) (Dukes and Reece, 2004). Three muscle layers (oblique, circular, and longitudinal) form the wall of the gastric stomach, are rich in nerve endings, and their coordinated contractions ensure proper mixing and passage of digesta (Dukes and Reece, 2004). In the gastric stomach, cells along the inner lining of the stomach produce various secretions. Mucus is produced by mucus cells, pepsinogen is secreted by chief cells, and HC1 is produced by parietal cells. The production of HC1 is important because it not only has antimicrobial properties, but also stimulates proteolytic activity in the gastric stomach. The pH can vary within the gastric stomach from as high as 7.0 in the cardiac region to as low as 2.0 in the pyloric region (Dukes and Reece, 2004).

3

Another function of the gastric stomach is to communicate with the liver, pancreas, and duodenum via stretch receptors in the gastric muscle layers and via the secretion of hormones like gastrin (Raybould, 2008; Williams, 2009). For example, if a fatty meal has being consumed, the gastric stomach w d fill, stretch receptors in the . muscle layers of the gastric stomach will be stimulated, and hormones will be released. This results in “priming” of the liver and pancreas to produce lipases and proteases. The pH in the duodenum is usually much higher than in the gastric stomach and is largely influenced by the secretion of bicarbonate from the gall bladder (Raybould, 2008; Williams, 2009). There are substantial variations and anatomical adaptations in the gastric and pregastric regons of the animals (Stevens and Hume, 1998). These adaptations are discussed elsewhere in this chapter and their significance in relation to food acquisition and microbial fermentation is explored. Likewise, there are significant adaptations in the large intestines of mammals, primarily the cecum and colon, which is dscussed later (Stevens and Hume, 1998). The small intestine is made up of the duodenum, jejunum, and ileum (Dukes and Reece, 2004). It is incorrect to assume that the small intestines or, in fact, any of the abdominal organs float freely within this cavity. The intestines are encased by a tough doublelayered connective tissue sheet called visceral peritoneum. Between the sheets of peritoneal tissue there are nerves and blood vessels that connect the intestinal tissue to the rest of the body. This is important for transporting nutrients to the liver, etc., and for nerve attachment to the muscle layers of the intestines (Dukes and Reece, 2004). The most important muscular structures of the intestines are the longitudmal and circular muscle layers, which coordinate peristaltic movement and propel digesta down the tract (Dukes and Reece, 2004). Fingerlike projects, called villi, increase the surface area of the small intestines and also secrete various

4 W KRAUSEANDKHAFIPOUR

enzymes that aid in the digestion and absorption of carbohydrates, protein, and lipids (Groschwitz and Hogan, 2009). Mammalian enzymes cannot digest p-1 ,Cglycosidic linkages found in plant cells’ wall carbohydrates like cellulose (Flint et al., 2008). Thus, the cellulosic-rich foods can only be fermented by microorganisms (Flint et al., 2008). Moving lstally down the small intestines there is an increase in the follicular-associated lymphoid tissue (Singh et al., 2009). These are regions of immune activity characterized by cell specialization and dense populations of immune active cells. Histologically, they are noticeable for their dark staining attributes. Various antimicrobial products are produced by these follicles, including antibodies, which aid in the control of microbial populations in the bowel (Singh et al., 2009). Almost all digestible food components are absorbed when the digesta reaches the terminal ileum (Fahey et al., 2008; Saulnier et al., 2009). The remaining undigested food particles pass through the ileo-cecal junction into the large intestines, which, in most mammals, are made up of the cecum, colon, rectum, and anus (Fahey et al., 2008; Saulnier et al., 2009). As mentioned previously, adaptations in the large intestines are numerous and are discussed in greater detail later. In general, a rich population of microorganisms ferments any feed particles that reach the large intestines, and the volatile fatty acids (VFA) and ammonia are absorbed. It is important to note that, in most animals, the ability to absorb simple sugars, lipids, and amino acids in the large intestines is limited or nonexistent. This is significant because, if substantial amounts of sugars, lipids, or amino acids are present in the large intestines, the animals cannot normally gain the benefit from them unless there is some sort of adaptation (Fahey et al., 2008; Saulnier et al., 2009). The large intestines do not have villi, but do have colonic cuffs, which increase the surface area for absorption (Hauck et al., 2005). Follicular-associated lymphoid tissue is also

present and increases in density as one moves distally toward the rectum. Lastly, one of the most important functions of the large intestines is water balance (Hauck et al., 2005). The large intestines are well-adapted to extract water from the digesta and, in arid-adapted mammals (e.g., eland, goats, camels), the feces are almost dry, whereas in animals like domesticated cattle the feces are quite wet and soft (Stevens and Hume, 1998). INTRODUCTION TO HOSTMICROBIAL SYMBIOSES Differences between animals essentially arise because of specialization in different regions of the digestive tract (Flint et al., 2008). These differences are reflected in the various host microbial symbioses that exist (Table 1) and the fermentative capacity in different regons of the gut. In general, regions of high fermentative capacity have a greater volume as a percentage of the total gut volume (Table 2). For example, the rumen in many herbivores is a specialized pregastric fermentation chamber that facilitates the degradation of cellulose by the animal. On the other hand, the human has a very limited ability to lgest cellulose, which is exemplified by the absence of pregastric fermentation (Flint et al., 2008). In almost all animals studied, there are microorganisms associated with the lgestive tract (Flint et al., 2008). The only known exceptions are some asteroidechinoderms:“spiny-skinned” creatures with a star- or asteroid-shaped body, for example starfish (Stevens and Hume, 1998). Another example of animals without a gut are bivalve mollusks, like mussels, that feed by filtering sea water through their gills (Stevens and Hume, 1998). The gut is largely anaerobic and although many bacteria, like the Proteobacteria, can tolerate oxygen, the majority cannot. Primarily, prokaryotes, archaea, and protozoa populate the gut, but fungi and viruses are also present. Based largely on the anatomical structure seen in animals and humans it is possible to &vide host-microbial symbioses into three models (Mackie, 2002). This is not the only

1. THE FECAL ENVIRONMENT, THE GUT H 5

TABLE 1 Mammals classified by gastrointestinal anatomy‘ Class

Species

Foregut fermenters Ruminants

A A

Hoatzin

Grazing herbivores Selective herbivores, including folivores and frugivores Selective herbivore Selective herbivores Grazing and selective herbivores Folivore

Capybara Rabbit (lemming) Rat, mice

Grazer Selective herbivores Omnivores

B B D

Elephant, horse, zebra New world monkeys Pig, human

Grazers Folivores Omnivores

Giant and red panda Dog, cat

Herbivore Carnivore

B C D E E

Cattle, sheep Deer, antelope, camel

Nonruminants

Colobine monkey Hamster, vole Kangaroo, hippopotamus

Hindgut fermenters Cecal digesters

Colonic digesters Sacculated

Unsacculated

Microbiome community typeb

Dietary habit

C A A C

“Adapted from Van Soest (1994) with permission tiom Cornell University Press, and Ley et al. (2008)with permission from Macmillan Publishers Ltd. bFecal bacterial community types were clustered into five groups based on the diet and gastrointestinal morphology: A, foregut fermenter herbivores; B, hindgut fermenter herbivores; C, folivores or omnivores; D, omnivores or frugivores; E, carnivores.

TABLE 2

Ruminant Cattle Sheep Camel Nonruminant Elephant Horse Pig Rabbit Rodent Capybara Guinea pig Rat Vole

Gastrointestinal volume of mammal species“

13-1 8 12-19 -

0.5 0.7-1.6 -

0.9-2.3 1.O-1.6 -

0.8 0.9-1.6 0.1-0.3

0.8-1.5 0.54.7 1.0-2.2

-

-

16.4 10.4 7-18

1.3 3.6 2-7

2.6 1.9 0.6-1.8

3.2 2.4 1.6 2.5-7.8

8.8 8.8 3.4 0.7-1.3

16.5 8.0 3.1 -

1.8 1.9 0.4 -

1.0 0.9 0.9 -

11.7 3.7 1.o 5.9

1.3 1.6 0.9 -

“Reprinted from Van Soest (1994) with permission from Comell University Press. Volume is weight ofliquid contents as percentage of body weight.


way to classi@ animals, but it is a useful one because it allows one to appreciate the relationships between host and microbe. MODELS OF HOST-MICROBIAL SYMBIOSES

Competition Model The first of these models is called the competition model (Mache, 2002). It is exemplified by the carnivore digestive tract that is essentially a straight tube with little specialization. Many of these animals, like the dog and members of the cat family, have little fermentative specialization (Fig. 1A and 2A). Host and A

Diet

PregaStriC fermentation

A I

I I

microbiome compete for the same substrate. The host retards the ability of the microbiome to utilize the food by the secretion of acids and other antimicrobial mechanisms. It was once thought that the gastric stomach was almost sterile and that this was the reason that the host could extract most of the nutrients from the food. We now know this is not true because the gastric stomach and small intestine are well-populated with microorganisms. The real competitive advantage of the carnivore digestive system is that the rate of passage of food is relatively high. Because the proteindominant diet is highly digestible, the host quickly extracts nutrients.

-1

I I

I I I

Cecal

-

I

Pregastric rmentation

Rumination

I

I

I

~

Feces I

w/coprophagy

Diet

langur monkeys, hoatzin, some I

2. Vole, hamster

B

hippos, Colobine, and

Gastric

Colonic

I

1. Kangaroos,

I

Cecal

dinosaurs

I

Omasal N sieve

1

~ -b Gastric

Bypass

peccaries, and (probably) large herbivorous

I -

Colonic

~

Feces

FIGURE 2 (A) Sequence of digestion in nonruminants with pregastric digestion. (B) Sequence of digestion in ruminants. The sieving system of the omasum is much more developed in grazing species. Reprinted from Van Soest (1994) with permission from Cornell University Press.

1. THE FECAL ENVIRONMENT. THE GUT

Combination Model The combination model is one in which the inefficiencies of foregut fermentation are translocated to the hindgut (Mackie, 2002). A typical combination model digestive tract is essentially nonruminant, but animals have a capacious hindgut that accommodates fermentation. These animals have some cecal fermentation abilities and often practice coprophagy (Fig. 1B and 2B). Alternatively, some animals in this group have very extensive colonic fermentative ability as in the case of the horse (Fig. 1A and 2A). In this manner, the inefficiencies of pregastric microbial fermentation of protein are avoided and the animal has the opportunity to digest and absorb amino acids. Any residual protein enters the large intestine and is fermented by microorganisms. The dfference between the competition and combination models is the extent to which fermentation occurs in the cecum and colon. In the competition model, exemplified by the meat eater, there is little hindgut fermentation but, in the combination model (exemplified by the horse, rabbit, and dugong), extensive fermentation in the cecum and colon is possible. However, one of the inefficiencies of this system is that the animal cannot gain any benefit from the microbial protein produced during fermentation in the hindgut because there is no ability to absorb protein post-ileum (Maclue, 2002). This makes animals like the horse inherently inefficient because they consume substantial amounts of forage and, because the most prominent fermentation site is the cecum, the microbial protein produced cannot be absorbed. At the same time, there is no ability to degrade the cellulose and hemicellulose in the forage by the mammalian enzymes and the only place for that to happen is in the cecum. Some animals, like the rabbit, practice cecotrophy in which the microbial protein rich fecal pellets are reconsumed (Fig. 1A and 2A). Cooperation Model The ruminant animal, in which there is extensive fermentation proximal to the gastric stomach, exemplifies the cooperation model

7

(Mackie, 2002). Ruminant animals can convert grass/forage/cellulose to calories, but this process is dependent of the cellulolytic microbiota that exist in the rumen (Fig. 1B and 2B). The animal provides a readily available substrate (cellulose) and a thermostable environment. O n the other hand, the microbiota provides cellulases and hemicellulases not produced by mammalian cells that can degrade forage. The end products of these processes are VFA (acetate, propionate, and butyrate) that are absorbed across the rumen wall and form an energy source for the animal. In addition, the microbial protein that is produced in the rumen flows to the lower gut and supplies all the protein requirements of the animal. The largest cost in this model is that potentially useful nutrients that enter the pregastric fermentation chamber are destroyed (Mackie, 2002). An example of this is protein, which is rapidly fermented by microorganisms to produce ammonia, VFA, and microbial protein. Excess ammonia is absorbed across the gut wall and lost in urine. Also, the conversion of feed protein into microbial protein is inherently inefficient because of ATP transaction costs during microbial metabolism (Russell and Wells, 1997). ANATOMY AND PHYSIOLOGY OF THE FOREGUT Moir (1965, 1968) listed a number of herbivores that have muscular constrictions dividing the pyloric and cardiac regions of the gastric stomach into two sacs. The animals in this grouping of Rodentia are mice and rats. Mice include Muscardinus (dormice), Notomys (kangaroo mice), Peromyscus (deer mice), and Perognathus (pocket mice). Rats include Neotoma (wood rats), Dipodomys (kangaroo rats), and Cricetomys (giant pouched rats). Beavers (Castor) and voles (Microtus) also fall into this grouping. Camain et al. (1960) evaluated the gastric stomach of the Gambian, or giant pouched rat (Cricetomys gambianus). It has a cardiac chamber (7.25 X 3.0 cm) that is connected by a narrow orifice (1 cm) to a pyloric chamber (3.5 X 2.5 cm).

8

KRAUSEANDKHAFIPOUR

An interesting foregut fermentative adaptation is found in animals that practice coprophagy or reingestion of feces (Fig. lB, 2A, and 3). This practice is largely restricted to rodents and lagomorphs (rabbits),but it also occurs in some pnmates and marsupials (Bauchop, 1977). These animals have an extended hndgut but are largely herbivorous and do not have extensive pregastric fermentative abihties. Fiber in plants that they consume is therefore not dgested extensively untd it gets to the large intestine, where cellulolytic microorganisms can break it down and convert the carbon into microbial protein. The problem is that the microbial protein is produced in a region of the gut where protein is usually not broken down and absorbed. These Cellulose

animals usually excrete a pellet that is soft, rich in microbial protein, and often mucoid. Animals then reconsume the pellet so that the microbial protein-rich pellet passes through the hgestive tract a second time. Ths process is called cecotrophy (Gnffiths and Davies, 1963). The development of pregastric fermentation in herbivores is a very successful adaptation for animals consuming diets dominated by forages. However, it has its own inefficiencies-the most important being the fermentation of mdk and other intact proteins (Black and Sharkey, 1970). When & is dxectly infused into the abomasums (gastric stomach) of lambs, the growth rates of the animals are superior than if the milk was inhsed into the rumen, but not

Starch

Sugar

Hexose

NADH

NAD Lactate I I I

I I I I

I

v Acetate

Acetate

+ Propionate

Butyrate

Propionate

FIGURE 3 Schematic showing the major pathways of carbohydrate fermentation by ruminal bacteria. “X’ denotes alternative electron carrier (e.g., ferredoxin). In some mminal bacteria, pyruvate decarboxylation is coupled to formate production, but most of this formate is converted to hydrogen and carbon dioxide by hydrogen formate lyase. The dashed lines show pathways that occur in other organisms. Reprinted from Russell and Rychlik (2001) with permission fkom AAAS.


if the lambs drank the milk by suckhng @lack, 1970). In ruminant and ruminant-like herbivores the reticular groove contracts when young animals suckle (Black and Sharkey, 1970). This muscular groove forms a channel that allows the milk to pass through the rumen directly to the abomasum. The reason for the superior growth rates of the animals is that the rumen microorganisms wastefully ferment the milk. Black and Sharkey (1970) surveyed the literature and concluded that all animals that have a major foregut fermentative adaptation also have a reticular groove. Apart from domesticated ruminants, members of the orders Marsupialia (kangaroos), Primates (coloboid monkeys), Edentata (sloths), and Artiodactyla (bovids, camelids, deer, hippos) are included. In contrast, herbivorous animals that do not have extensive foregut fermentation do not have a reticular groove. Exceptions to this appear to be the elephant and lemmings, which have a rudimentary reticular groove (Black and Sharkey, 1970). MULTICHAMBERED FORESTOMACH WITH MINIMAL FERMENTATION Several mammals have multichambered stomachs, like the golden hamster (Mesocricetus auratus) and the rock hyrax (Procavia habessinica), s;

which have a two compartment stomach divided into pregastric and gastric regions (Hume and Warner, 1980) (Fig. 4). There is clearly microbial fermentation in these compartments because there are significant concentrations of VFA. Even aquatic mammals like bowhead and gray whales, spotted dolphins, and black finless porpoises have multichambered stomachs (Morii, 1972; Morii and Kanazu, 1972; Morii, 1979; Herwig et al., 1984). There is also microbial fermentation in these animals as evidenced by the high concentrations of VFA found. Two examples are discussed in more detail in the following text.

Golden Hamster In the golden hamster (Mesocricetus auratus) there is a sphincter-like muscular ring that clvides the stomach into two compartments and it is likely that contraction of this muscle layer regulates the flow of digesta between the two compartments (Hoover et al., 1969) (Fig. 4). The esophagus joins the pregastric chamber in close proximity to this sphincter and most of the ingested food is deposited in this pregastric chamber (Ehle and Warner, 1978). If the sphincter is removed and the dgesta moves freely across the anastomosis margin, there is apparently little effect on the digestion of

Max cellulose

40 7

4 ATP 30

20

: : 10

2 ATP

1 -

W

9

E

0

I

0

10

20

9

30

40

50

Rate of fermentation Pk/h)

FIGURE 4 The amount of microbial protein produced per unit of feed fermented in the rumen affects the rate of fermentation. Reprinted with permission from Van Soest et al. (1991).

10

KRAUSE AND KHAFIPOUR

nutrients; thus, the actual role of the sphincter is not clear (Ehle and Warner, 1978). It is clear that the VFA concentration in the pregastric chamber approaches 160 mmol/liter, which is similar to that in the rumen (Hungate, 1966; Hoover et al., 1969).

Rock Hyrax The rock hyrax (Procavia habessinica)is unique in that it has three fermentation compartments that are completely separated f?om one another anatomically (Fig. 4). There is a gastric fermentation chamber in whch high levels of VFA are found (Hungate, 1966) but which has a low rate of digesta flow and may take as long as 8 h to empty. Between the terminal ileum and the colon there is an enlarged sac, called the midgut sacculus, where extensive fermentation occurs. The last site of fermentation is in the paired ceca. In both of these last two sites VFA concentration is considerably less than in the cranial sac, and is in the region of 60 to 80 mmol/liter.

region allowing for a reduction in pH (2 to 4), but, because it is anatomically separated from the proximally located saccus and presaccus, fermentation (PH 6 to 7) can occur in these two forechambers without the antimicrobial effects oflow pH (Kuhn, 1964;Bauchop and Martucci, 1968; Ohwaki et al., 1974).The stomach capacity is between 11% and 32% of body weight, whch results in a slow rate of passage (Kuhn, 1964; Bauchop and Martucci, 1968; Ohwaki et al., 1974).Microbial fermentation is extensive and a majority of animal’s energy requirements comes &om VFA production in the foregut (Bauchop and Martucci, 1968).

Herbivorous Macropodid Marsupials

Colobid Monkeys

In the family Macropodidae all species are foregut fermenters and are divided into two groups, the Macropodinae (kangaroos and wallabies) and the Potoroinae (rat-kangaroos), most of which are found in Australia (Hume, 1982) (Fig. 4). The stomach is divided into three sacs, although the anatomical distinction between these regions is not always obvious. Close to the esophageal opening there is a sacciform and tubiform forestomach, which is divided by a ventral fold (Dellow, 1979). It is in these two regions that most of the microbial fermentation of fibrous feedstuffs takes place (Langer, 1980). There is also a third chamber, which is glandular, and in which acid secretion takes place. The VFA concentrations in this third chamber are fairly low in comparison to that found in the two cranial sacs. From this one can conclude that the degradation of the predominantly fiber diets these animals consume takes place in the two cranial sacs (Dellow, 1979; Hume, 1982).

There are three genera of monkeys in the family Cercopithecidae that are leaf-eating herbivores (Ha, 1952; Bauchop, 1977, 1978). The esophagus enters into two large chambers, the presaccus and the saccus (Fig. 4). An esophageal groove connects these two chambers by way of the lesser curvature of the stomach, via a tubular portion, to an almost anatomically distinct pyloric region. Acid secretion occurs in this pyloric

PSEUDORUMINANTS These constitute animals in the suborder Tylopoda and in the family Camelidae, and are made up of one-humped camels (Camelus dromedarius), two-humped camels (C. bactrianus), as well as llamas (Lama peruna) (Fig. 4), alpacas (L. pacos), and vicunas (Vicugna vicugna). They are similar to true ruminants in that they ruminate

RUMEN-LIKE! ADAPTATIONS The largest difference between animals with rumen-like adaptations, and those without, is that the microbial activity is very extensive and the VFA produced is a major source of energy for the animal. As discussed previously, these animals, which fit into the combination model described by Hungate (1976, 1984), are adapted to microbial breakdown of the plant cell wall and converting plant carbon into microbial protein. There are a large number of animals in this group and a number of examples will be dscussed.


and remasticate digesta but have three distinct compartments but not the fourth found in true ruminants. The first of these three compartments makes up 83% of the total gastric volume and is the site where most of the degradation of fiber occurs. Consequently, the VFA concentrations are high, indicating active microbial fermentation (Vallenas et al., 1971). It is often thought that this first compartment is similar to the rumen, and it may be in terms of fermentative function, but it most certainly is not histologically and anatomically similar. First, there is an esophageal groove that runs from the first compartment to the third compartment. The comparable structure in true ruminants, the reticulo-omasal orifice, has two muscular lips, while the one in the Camelidae only has one. Second, in the first compartment, there is a cranial and cardial sacculation, both of which are glandular. There is no glandular tissue in the rumen. These sacculations are covered by a mucosal diaphragm that prolapses into the first compartment during contractions. Also, the rest of the first compartment is covered by stratified squamous epithelium, whereas the surface of the rumen is covered by papilla. Last, the esophagus in the Camelidae joins the upper right part of the first compartment, whereas in ruminants the esophagus joins at the atrium ventriculi shared by the rumen and reticulum. The second compartment is analogous to the reticulum in true ruminants but has some obvious differences. It makes up about 6% of the gastric volume and is thus much smaller than the first compartment (Vallenas et al., 1971). It also has papdlated gastric glands in the region of the greater curvature. The rest of the surface is covered with stratified squamous epithelium. The third compartment (11% of gastric volume) is an elongated chamber and is connected to the second compartment by a thick muscular tube which is probably a continuation of the esophageal groove. Gastric mucosa covers the entire surface of this compartment and the cells in the terminal region produce acid. It is thought that the proximal

11

four-fifths of the compartment are analogous to the omasum in true ruminants and the distal one-fifth is analogous to the abomasums (Vallenas et al., 1971). The extent of fiber digestion and fermentation appears to be much less than in the first compartment (Vallenas and Stevens, 1971). PRERUMINANTS Chevrotains can be categorized into water chevrotains, or Hyemoschus aquaticus, and Tragulusjavanicus, the mouse deer. The mouse deer weighs between 2.5 and 4.5 kg and has a stomach with three compartments, the first two of which are completely covered with papillated squamous epithelia (Walker, 1964). The ventrocaudal blind sac extends to give the rumen an S-shape, which is not typical in the true ruminants (Moir, 1968). Langer (1974) conducted a series of studies with fetal ruminants and concluded that the S-shaped rumen in the chevrotain is actually a primitive stomach. It was further suggested that the suborder Ruminanti further divided into an infraorder without an omasum (Tragulina) and one with an omasum (Pecora). There is no reticulo-omasal orifice in the Tragulina so that the passage of feed into the abomasum would be expected to be quicker than in true ruminants (Langer, 1974; Janis, 1976). The disadvantage of this anatomical adaptation is that the deer mouse cannot retain fiber in the ventrocaudal sac to enable complete dgestion of plant cell walls, because cell wall digestion is a relatively slow process. There is evidence that the Tragulina select for immature forage that would be easier to digest (Grzimek, 1974). RUMINANTS True ruminants are found in four families, the Cervidae, Giraffidae, Antilocapridae, and Bovidae (Romer, 1966). The Cervidae contain 17 genera and at least 53 species including musk deer, muntjacs, fallow deer, axis deer, red deer, and elk. In the Giraffidae there are two species, the giraffe and okapi.


The Antilocapridae only has one species, the pronghorn antelope. In the Bovidae there are 49 genera and 115 species and the most common members are domesticated cattle, sheep, and goats (Walker, 1964). The ruminant stomach consists of four stomach compartments: rumen, reticulum, omasum, and abomasums (Fig. 4). Stratified squamous epithelium lines the first three compartments and the rumen in particular is papillated. The abomasum is covered with glandular tissue and is analogous to the gastric stomach in humans. In general, the overall anatomy of these four stomachs is very similar when comparing members of these four families except that the omasum in the Cervidae tends to be much smaller. There are also some differences in epithelial structure, papilla, and number of omasal leaves (Church, 1976). All animals in this order appear to have an esophageal groove that allows milk to be shunted from the esophageal antrum to the abomasums (Church, 1976). The most important advantage is that milk is not fermented in the rumen by suckling animals and wasteful fermentation is avoided (Black and Sharkey, 1970). The capacity of the rumen and the various compartments is difficult to measure because of the influence that age, diet, and physiological state have on the volume of each chamber. In sheep, the rumen ranges from 3 to 15 liters and the abomasums is about 2 liters. In cattle, the rumen volume ranges from 35 to 100 liters and the abomasum ranges from 3 to 8 liters (Church, 1976; Hofhan, 1988a, 1988b). Microbial fermentation in the rumen is extensive and was first demonstrated by Von Tappeiner in 1884 (Hungate, 1966). This investigator demonstrated that, by inactivating bacteria, cellulose digestion could be terminated. ANATOMY AND PHYSIOLOGY OF THE CECUM AND PROXIMAL COLON The anatomical variety in the hindgut (cecum and colon) of mammals is extensive and is largely related to the ability of the animals to chgest feed. For example, the carnivore

hindgut is a small simple tube with essentially no sacculation. This small size inchcates low fermentative ability, placing these animals in the competitive model category. Carnivorous marsupials (quolls, dunnarts, wombats, Tasmanian devil) and several eutherian carnivores such as the mink have no cecum (Hume and Warner, 1980). It should also be noted that the human has a very small cecum, or even nonexistent cecum, ending in an appendix. In contrast the capacity of the hindgut of herbivores, in general, is quite extensive. For example, the horse has a hindgut capacity that reaches 12% of body mass (Engelhardt et al., 1983).

CECUM FERMENTERS These are animals with a body mass of less than 10 kg and, in these herbivores, the principle site of mgestion and fermentation of fibrous material is the cecum (Hume and Warner, 1980). In many cases the fermentative activities extend into the colon-the best example of which is the koala, in which the cecum and colon have similar capacities (Hume, 1982). Examples of these animals also include the greater glider, ringtail possums, bushtail possums (eucalyptus consuming marsupials), rodents, lagomorphs (rabbits, hares, and pika), and some small monkeys (e.g., tamarins, marmosets) (Fig. 4). Bjornhag (1987) summarized the various colonic separation mechanisms employed by animals to selectively retain fluid and particles in the hindgut. As is the case with the horse, there are strong retroperistaltic muscular waves in rabbits that run in the direction of tail to head, emanating from the junction of the proximal and distal colon. Larger particles tend to be caught up in the haustrated architecture of the colonic wall, whereas small particles (including bacteria) are washed down the tract to the rectum. In rabbits there is net secretion of water into the proximal colon, but a net absorption of water from the cecum (Clauss, 1984). This process allows rabbits to regulate cycles of hard and soft feces formation. When soft feces are formed, there is no net absorption of water from the proximal colon.

1. THE FECAL ENVIRONMENT, THE GUT W 13

The overall effect of selective retention of fluid and fine particles in the cecum is to maximize the ability of the microbiota to ferment the nutrients, which results not only in an increase in microbial protein yield but also VFA, which is absorbed (Bjornhag, 1987). In addition, the cycling in the cecum and colon allows for the rapid expulsion of larger particles that are less fermentable. The formation of soft feces is also accompanied by decreased motility in the proximal colon and increased motility in the hstal colon. The ability to remove large particles rapidly is advantageous because it allows the animals to maintain a higher feed intake. For example, the rabbit is able to &gest only 10% of the neutral detergent fiber from alfalfa but the guinea pig can digest 34% (Sakaguchi et al., 1987). The guinea pig has a much less effective separation mechanism in the cecum and colon and retains larger particles for much longer than the rabbit. Foley and Hume (1987) demonstrated that the bushtail possum (Trichosuvus vulpeculu) is not able to survive on eucalyptus forage because it retains the larger particles in the cecum for fairly long periods. In contrast, the common ringtail possum (Pseudocheirus peregrinus) can survive exclusively on eucalypt forage and does not retain large particles for much shorter periods in the cecum (Chilcott and Hume, 1985).

Colon Fermenters These are animals in which the cecum acts as an extension of proximal colon and there is extensive mixing between these two compartments. This is typically the case in large hindgut fermenters or animals with a body mass of greater than 10 kg (Hume and Sakaguchi, 1991). In some colon fermenters, like the marsupial wombat, the cecum is almost completely absent. Other colon fermenters with a body mass of more than 10 kg are primates, humans, rhinos, dugong, equids, and pigs (Hume and Sakaguchi, 1991). Colon fermenters tend to be more adapted to fiber digestion than cecum fermenters (Hume and Sakaguchi, 1991).

HORSE The horse is the best-studied colon fermenter and is primarily a herbivore that is well adapted to fiber fermentation (Fig. 4). Argenzio et al. (1974) found that the retention times of particles, as is the case in true ruminants, is greater for particulates than for particle-free material. As the length of the particles increases, so does the retention time. The increase in retention is because of the semiluminal folds in the colon and, secondly, the retroperistaltic muscular waves that travel from tail to head and originate at the colonic-pelvic flexure (Sellers et al., 1982). However, the difference in retention times between particle and fluid is only about 3 to 5 h, which is less than one may have expected, and is the result of contraction of the distal colon in the direction of head to tail (Bjornhag, 1987). In general, large herbivorous colon fermenters do not dgest fiber as well as true ruminants largely because of the lower retention times (Foose, 1982). WOMBATS Barboza (1993) demonstrated that in wombats the mean retention time of particles was 57 h and 33 h for that of fluid (Fig. 4). These times are greater than in the horse, which has mean particle retention times of 27 h and mean fluid retention times of 22 h (Orton et al., 1985). This discrepancy appears to be related to low nutrient requirements, likely related to the low metabolizable energy needs of the wombat (Barboza and Hume, 1992). The digestive strategy of the wombat appears to revolve around a combination of low intakes of poor quality forages, from which the maximum amount of energy is extracted by allowing for extended retention times in the colon (Barboza and Hume, 1992).

i

HOST-MICROBIAL SYMBIOSES In the precedmg sections we discussed the morphological variation in both foregut- and hindgut-fermenting mammals. Each of the a n i m a l s discussed can generally be assigned to the competition, combination, or cooperation

14


models. The question then becomes whether these models of host-microbial symbiosis can predict the phylogenetic composition of the microbiome. Many questions stdl need to be answered but some titillating clues are emerging. Rappt and Giovannoni (2003) analyzed the phylogenetic diversity in the biosphere based on available 16s rDNA sequences and could divide sequences into 52 phyla. Of these 52 phyla, only 26 had cultured members, whereas the remaining phyla had not one single cultured member represented. The &gestive tracts of mammals contain roughly 12 of these phyla and are dominated by the Firmicutes and Bacteroidetes (Ley et al., 2008). This restricted distribution means that the digestive tract is not simply colonized by random microorganisms that pass through the digestive tract. For example, cattle continually consume soil in the process of eating forage. One of the major phyla in soil is Nitrospira, but this organism never occurs in the rumen (Cole et al., 2009). On the other hand, the phylum Firmicutes is found in both soil and the rumen, but at the family level there are a large number of taxa that occur in soil and not in the rumen (Cole et al., 2009). One of the problems leading to these discrepancies is that molecular taxonomies derived from 16s rDNA sequences are not always reflective of differences in the function or phenotype of bacteria. We know that 16s rRNA classification is often not coherent with the function (Staley, 2006). In other words, simply because two bacteria have the same 16s rDNA sequence does not mean they have the same function. Classic examples are Escherichia coli and Salmonella spp., which only differ by one base in their 16s sequence but have an entirely different disease etiology (Cole et al., 2009). However, having noted these caveats, 16s rDNA analysis can still be used to shed light on the relationships between hostmicrobial symbioses. Ley et al. (2008) compared 16s rDNA sequences of fecal microbiota samples from humans to 17 nonhuman primates and to

42 animals from 11 taxonomic orders, many of which are represented by the gut morphological groups dscussed previously. As might be expected, the human-associated microbiomes were more simdar to each other than to any of the other mammals. The authors suggested that &et type had a major influence in microbial clustering (Table 1). Humans clustered with other omnivores like the ring-tailed lemur, black lemur, bonobo, and spider monkey. Other hominids that consumed &ets dominated by plant material clustered in a position intermedate between omnivorous primates and nonprimate herbivores (Ley et al., 2008). In general, the phylogenetic clustering of gut microbiota could be explained by met (herbivore, omnivore, or carnivore) and whether the animals were fore- or hindgut fermenters (Ley et al., 2008). But there are exceptions that may lead us to a deeper understanding of host-microbial symbioses. The giant and red pandas consume a diet dominated by forage (bamboo shoots), making them herbivores. These animals have a digestive tract that is relatively short and does not have areas of fermentative specialization, thus placing them with the carnivores based on digestive tract morphology (Wei et al., 2007). However, their microbiota has a composition making it much more similar to carnivores (Ley et al., 2008). In the analysis of Ley et al. (2008), two groups of carnivores are described. For the purposes of this discussion we will call them true carnivores because they have a fairly simple digestive tract with little specialized fermentation areas and eat meat primarily. This group includes the hyena, bushdog, cheetah, and lion. The other group, which we call omnivorous carnivores, includes animals like the panda, the polar bear, spectacled bear, and echidna. These animals also have a fairly simple digestive tract with little fermentative specialization, but they consume mostly plant material. Another test of the diet hypothesis is to take mammals and place them on entirely different diets and then ask whether there is a difference

1. THE FECAL ENVIRONMENT, THE GUT W 15

in microbial composition. In a recent study, Khafipour et al. (2009) placed dairy cattle on a high grain diet and on a diet dominated by plantdforage. The high grain diet was designed to induce a metabolic disease called subacute ruminal acidosis whereas the high forage diet l d not. In both studies the distribution of phyla represented was the same, but there were differences at the genus level in relative abundance. FUNCTIONAL ROLES OF MICROORGANISMS OF THE DIGESTIVE TRACT In the preceding section, significant attention is given to host-microbial symbioses. However, microbial-microbial symbioses are just as important. A major function of microbial symbioses with the mammalian host is the byproducts of fermentation. O n the one hand, VFA are produced and can be a major source of energy for the host, particularly in ruminants, but they also are a source of microbial protein for the host. The extent to which the host can derive benefit from these end-products largely depends on whether the digestive tract fits into the competition, cooperation, or combination models (see previous text). An exhaustive review of the bacteria, protozoa, and fungi is not given in this chapter and interested readers are encouraged to refer to other published works (Orpin and Joblin, 1997; Stewart et al., 1997; Williams and Coleman, 1997). Bacteria in the lgestive tract require carbohydrates as well as nitrogen to grow optimally and, to enable maximum microbial yield in the gut, these two substrates need to be provided in synchrony. The amount of microbial protein produced per unit of feed, or substrate, affects the rate of fermentation in the digestive tract (Van Soest, 1994). The highest rates of fermentation are cellulose < unprocessed starch < processed starch < pectin (Fig. 4). Cellulolytic bacteria are very efficient because their maintenance cost is low compared to other bacteria and this gves them an advantage in the digestive tract. The reason for this is that

the rate of lgestion dilutes the maintenance cost for bacteria and carbohydrates that ferment more quickly improve efficiency. When large amounts of starch are added to the diet, starch-fermenting bacteria like Streptococcus bovis switch from acetate to lactate from which they get an ATP yield of two instead of four (Russell and Strobel, 1989). The increased lactic acid decreases gut pH and gives s. bovis a competitive advantage. Table 3 provides the major bacterial groups found in the gut of ruminants and humans, their major substrates, as well as end products. Figure 5 provides a schematic of the major fermentation pathways in the digestive tract. The digestive tract is generally a highly anaerobic environment but substrates entering are only partially oxilzed. A major challenge of anaerobic fermentation. in the digestive tract is to produce enough ATP, which first requires the disposal of hydrogen. The major mechanism of reducing equivalent disposal is via NADH (Russell and Wells, 1997). Hydrogen is almost always the major end-product in gut fermentation and is dependent on the presence of hydrogenases to convert the hydrogen to methane, a role reserved for methanogens. However, not all mammals produce methane (Hackstein and Stumm, 1994). If carbohydrates are converted to propionate, butyrate, or lactate, then dehydrogenases in various bacterial species can carry out these reactions (Russell and Rychlik, 2001). When hydrogenase activity is coupled with the production of acetate then the ATP yield is 4 mol per mole of glucose, for butyrate it is 3 mol, and for lactate it is 2 mol. Anaerobic gut bacteria were once thought not to possess respiratory chains but fumarate reductases have been coupled to cytochromes and the ATP yield from propionate can be as high as that of acetate (Russell and Rychlik, 2001). Although there are many substrates in the dgestive tract, it can be argued that the major difference between mammahan dgestive tracts is the manner in which carbohydrates are metabolized. In h s section we pay special attention

16 W KRAUSE A N D KHAFIPOUR

TABLE 3

Characteristics of predominant bacteria in human large intestine a n d in the rurnena’b

Ruminal niche

Species

Fermentation products

Bacteroidetes

Bacteroides thetaiotaomicron Bacteroides ovatus Bacteroides cellulosilyticus Bacteroides sp. nov.

ST XY, ST

cu XY

Firmicutes

Rosebutia intestinalis Roseburia inulinivorans Ruminococcus bromii Ruminococcus sp. nov. Eubacterium rectale

XY, ST IN, ST ST cu, XY ST

Actinobacteria

B$dobacterium adolescentis

ST

Rumen Bacteroidetes

Prevotella ruminicola, albensis, brevis, and bryantii

ST, PC, XY, SU, PR

Firmicutes

Ruminococcus albus Ruminococcusflavefaciens Anaerovibrio lipolytica Megasphaera elsdenii Selenomonas ruminantiurn Butyrivibriofibrosolvens Lachnospira multiparus Peptostreptococcus anaevobius Clostridium aminophilum Clostridiurn sticklandii Streptococcus bovis Eubacterium ruminantiurn Eubacterium cellulosolvens

CU, HC CU, HC GL, TG, SU, L, AA L, SU, GL, PP ST, DX, SU, L, S, GL, PR ST, CU, HC, PC, SU, PR PC, su

AA AA, PR AA,PR ST, SU, PR HC, DX, SU cu, XY, PP

Proteobacteria

Ruminobacter amylophilus Succinomonas amylolytica Succinivibrio dextn’nosolvens Wolinella succinogenes Fibrobacter

Fibrobacter succinogenes Fibrobacter intestinalis

ST ST PC, DX, SU O A , Hz, F

cu cu

“Adapted from Russell and Rychlik (2001) with permission from AAAS, and Flint et al. (2008) with permission &om Macmillan Publishing Ltd. bAbbreviations: A, acetate; AA, amino acids; B, butyrate; Br, branched-chain volatile fatty acids; CU, cellulose; DX, dexmns; E, ethanol; F, formate; GL, glycerol; Hz, hydrogen; HC, hemicellulose; IN, inulin; L, lactate; OA, organic acids; P, propionate; PC, pectin; PP, peptides; PR, protein; S, succinate; ST, starch; SU, sugars; TG, triglyceride; XY, xylans.

to comparative carbohydrate degradation by the bacteria ofthe digestive tract. The digestive tracts of both humans and animals are populated by bacteria that produce glycosyl hydrolases that are involved in the degradation of polysaccharides, but only a few bacteria are directly involved in

t h s process in the sense that they actually adhere to the substrate (Flint et al., 2008).

In herbivores, carbohydrates are usually in the form of plant cell walls and primarily

constitute cellulose, hemicellulose, pectin, and lignin (Van Soest, 1994). These substrates

THE FECAL ENVIRONMENT, THE GUT W 17

1.

-Carbohydrate fermentation (VFAs) - - - Protein degradation -----Ammonia level I A

0

6

12

18

24

Hours postfeeding

FIGURE 5 Model relation in which release of nitrogen fractions parallels fermentation curves for respective fist- and slow-degrading carbohydrates and optimizes microbial protein output. Peptide production, which is important for the fast pool, depends on the presence of degradable true protein. Ammonia can come kom the nonprotein nitrogen pool and from peptide degradation. The ammonia level does not rise much because it is competitively recycled into microbial protein. A: Nonprotein nitrogen and peptides; B1: true soluble protein; B2: neutral detergent soluble protein. Reprinted from Van Soest (1994) with permission from Comell University Press.

are recalcitrant to digestion by mammalian enzymes. Bacterial systems have specialized mechanisms by which plant cell walls are broken down, but this process tends to be quite slow; the major rate-limiting factor is the presence of lignin in the cell wall. Lignin is make up of phenolic ring structures and, in gut ecosystems, the free energy required to achieve ring cleavage is deficient (Van Soest, 1994). The ability to degrade plant cell walls is an important factor regulating the composition of the gut microbiota. Without the reduction of large particles to small ones there is a retardation of passage of material through the digestive tract and a reduction of feed intake (Van Soest, 1994). If feed intake declines, so

does the ability of the animal to extract energy from the feed. In humans, the consumption of plant cell walls is not nearly as common as in herbivores, but complex carbohydrates in the form of starches are consumed and, in fact, encouraged because of the benefit to gut heath (Flint et al., 2008). In Table 3 a comparison is made of the microorganisms involved in carbohydrate degradation in humans and ruminants (Russell and Rychlik, 2001; Flint et al., 2008). Although their taxonomy is dfferent, there is a great deal of overlap in function between these two groups. In ruminants the major cellulolytic species are Ruminococcus albus, R.jlavefaciens, Eubacterium cellulosolvens, and Fibrobacter succinogenes.

is


TABLE 4

Genome sequences concerned with plant polysaccharide breakdown in four species of gut bacteria

Location Genome size

Number of g lycoside hydrolases' Number of polysaccharide lyases' Number of carbohydrate esterases' Number of carbohydratebinding modules" Reference(s)

Bacteroides thetaiotaomicron

B$dobacterium longum

5482

NCC2705

Human colon 6.26 Mb (complete genome) 236 (40)

Ruminococcus jlavefaciens FD 1

Human colon Rumen, cellulolytic 2.26 Mb (complete Approximately 4 Mb genome) (partial genome; dockerinencoding genes only) 47 (17) 65 (14)

Fibrobacter succinogenes S85 Rumen, cellulolytic 3.8 Mb (complete genome)

104

15 (7)

0

12 (4)

4

20 (9)

1

23 (5)

14

16 (3)

10 (5)

61 (12)

Limited information available

Hooper et al. (2002)

MacFarlane and Englyst (1986)

Rincon et al. (2005)

Pegden et al. (1998) and Morrison and Miron (2000)

"The number of enzyme families that are represented is shown in parentheses (CAZY [carbohydrate-active enzymes] database). Mb, megabases. Reprinted from Flint et al. (2008) with permission from Macmillan Publishers Ltd.

However, in humans, the major cellulolyticspecies are Bactevoides cellulosilyticus and Ruminococcus spp. There seems to be little species overlap between the rumen and the human hindgut. A similar situation occurs for the xylanolytic and especially the starch dgesters, whch seem to be quite important in humans. The discussion on polysacchande digestion and the species involved in both humans and ruminants is important because it points to the already mentioned notion that the function of bacteria are not necessarily matched by the taxonomy. In Table 4 the plant polysaccharide digesting apparatus of four species of gut bacteria are given, two from the human gut and two from the rumen (Flint et al., 2008). All are specialized in the degradation of complex plant material although the human isolates are more adept at degrading starch. All of these bacteria have multiple glycosyl hydrolases, polysaccharide lyases, and esterases (Flint et al., 2008). An important feature is that they

all have carbohydrate-binding domains. These bacteria produce their carbohydrate-degrading arsenal as multienzyme complexes and the overall enzymatic motifi have a fair degree of similarity (Flint et al., 2008). CONCLUSION This chapter surveys the immense diversity in the anatomical structures in the digestive tracts of mammals. In particular, special attention is given to adaptations in the digestive tract that allow for fermentative digestion and how various animals have evolved to utilize plant material. It is fascinating to note that these adaptations are not possible without the symbiotic relationships with the microorganisms that inhabit the digestive tract because they provide enzymes to break down plant cell walls that mammals do not have. Although still in the early stages of investigation, it now also appears that the relationship between the taxa inhabiting the digestive


19

Church, D. C. 1976. Digestive Physiology and Nutrition ofRuminants, 2nd ed., vol. I. O&B Books, Corvallis, OR. Clauss, W. 1984. Circadian rhythms in Na’ transport, p. 273-283. In E. Skadhauge and K. Heintze (ed.), Intestinal Absorption and Secretion. MTP Press, Lancaster, UK. Cole, J. R., Q. Wang, E. Cardenas, J. Fish, B. Chai, R. J. Farris, A. S. Kulam-SyedMohideen, D . M. McGarrell, T. Marsh, REFERENCES G. M. Garrity, and J. M. Tiedje. 2009. The Allen, M. S. 1997. Relationship between fermentaRibosomal Database Project: improved alignments tion acid production in the rumen and the requireand new tools for rRNA analysis. Nucl. Acids Res. 37:D141-145. ment for physically effective fiber. J. Dairy Sci. Dellow, D . W. 1979. Physiology of digestion in the 80: 1447-1462. macropodine marsupials. PhD thesis, University of Argenzio, R. A., J. E. Lowe, D. W. Pickard, New England, Armidale, Australia. and C. E. Stevens. 1974. Digesta passage and Dinalo, J. E., and D. A. Relman. 2009. Cross-talk water exchange in equine large-intestine. Am. J . Physiol. 226:1035-1042. in the gut. Genome Bid. 10:203. Barboza, P. S. 1993. Digestive strategies of the Dukes, H. H., and W. 0. Reece. 2004. Dukes’ Physiology .fDomestic Animals, 12th ed. Comstock wombats - feed-intake, fiber digestion, and digesta passage in two grazing marsupials with hindgut ferPub. Associates, Ithaca, NY. mentation. Physiol. Zool. 66:983-999. Ehle, F. R., and R. G. Warner. 1978. Nutritional Barboza, P. S., and I. D. Hume. 1992. Digestiveimplications of the hamster forestomach. J. Nutr. tract morphology and digestion in the wombats 108:1047-1053. (Marsupialia, Vombatidae). J . Comp. Physiol. BEngelhardt, W. V., G . Rechkemmer, L. Luciano, Biochem. Syst. Environ. Physiol. 162:552-560. and E. Reale. 1983. The hindgut: lessons for Bauchop, T. 1977. Foregut fermentation,p. 223-250. man from other species, p. 3-17. In L. Barbara, In R. T. J. Clarke and T. Bauchop (ed.), Microbial M. Miglioli, and S. F. Phillips (ed.), New Trends Ecology ofthe Gut. Academic Press, London. in Pathophysiology and Therapy of the Large Bowel. Bauchop, T. 1978. The significance of microElsvier, Amsterdam, The Netherlands. organisms in the stomach of non-human primates. Fahey, G. C., Jr., K. A. Barry, and K. S. SwanWorld Rev. Nutr. Diet 32:198-212. son. 2008. Age-related changes in nutrient utiliBauchop, T., andR. W. Martucci. 1968.Ruminantzation by companion animals. Annu. Rev. Nu6. like digestion of the langur monkey. Science 161: 28:425-445. Flint, H. J., E. A. Bayer, M. T. Rincon, R. 698-700. Lamed, and B. A. White. 2008. Polysaccharide Beauchemin, K. A. 1991. Ingestion and mastication of feed by dairy cattle. Vet. Clin. North. A m . Food utilization by gut bacteria: potential for new inAnim. Pract. 7:439463. sights from genomic analysis. Nut. Rev. Microbiol. Bjornhag, G. 1987. Comparative aspects of digestion 6:121-131. in the hindgut ofmammals - the colonic separation Foley, W. J., and I. D. Hume. 1987. Passage of dimechanism (CSM) (A review). Dtsch. Tierarztl. gesta markers in two species of arboreal folivorous Wsschr. 94:33-36. marsupials - the greater glider (Petauroides, volans) Black, J. L. 1970. Nutritional significance of the reticand the brushtail possum (Trichosurus, vulpecula). P h y ~ i ~2001. l . 60~103-113. ular groove (sulcis reticuli).Aus.J. Sci. 32:332-334. Black, J. L., and M. J. Sharkey. 1970. Reticular Foose, T. J. 1982. Trophic strategies of ruminants versus nonmminants ungulates. PhD thesis, University of groove (sulcis reticuli): an obligatory adaptation in ruminant-like herbivores. Mammalia 34:294-302. Chicago, Chicago, IL. Camain, R., A. Quenum, J. Kerrest, and S. Griffiths, M., and D . Davies. 1963. The role ofthe Goueffon. 1960. Considerations sur l’estomac de soft pellets in the production of lactic acid in the Cricetomys gambianus. Bull. Mem. Ecole. Natl. Med. rabbit stomach.]. Nutr. 80:171-180. Groschwitz, K. R., and S. P. Hogan. 2009. IntestiPhaum. Dakar 8:134-142. Chilcott, M. J., and I. D . Hume. 1985. Conal barrier function: molecular regulation and disease prophagy and selective retention of fluid digesta pathogenesis.J. Alletgy Clin. Immunol. 1 2 4 5 2 0 . their role in the nutrition of the common ringGrzimek, B. 1974. Animal Lij Encyclopedia, vol. 13 tail possum, Pseudochei~s-peregrinus.Aust. J . 2002. (Mammals IV). Van Nostrand Reinhold, New 33: 1-1 5. York. NY.

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TAXONOMY, PHYLOGENY, AND PHYSIOLOGY OF FECAL INDICATOR BACTERIA Militxa Cawero-Coldn, Gene S. Wickharn, and Ronald F. Turco

Although natural environments contain millions of different microorganisms, most of these pose little threat because they are not pathogenic for humans. Most environmental microorganisms are saprophytic and free-living (i.e., they do not require a host to function) and are able to utilize the carbon, nutrients, and cofactors they encounter in soil and water. On the other hand, pathogenic microorganisms require a host organism to proliferate and grow optimally. Inside their requisite hosts, pathogens encounter the most appropriate environment (required nutrients, pH, and temperature, to mention a few) that will sustain their growth and reproduction. Environmental pathogens are microorganisms that spend a portion of their life outside of a host but, when introduced to an acceptable host, may cause a disease. While in the environment pathogens can survive for various lengths of time, but growth and reproduction may not often occur. Pathogens can be present in water, soil, air, and food. Although the drrect detection of

pathogens in the environment would provide the best evidence for their presence, this is a hfficult task as they are usually present in low numbers (Straub and Chandler, 2003). Many protocols for sample collection, microbial concentration, and identification are not sensitive enough to detect the pathogenic agents within the larger nonpathogenic population. Because of the challenges with drect pathogen detection, scientists and the regulatory community have trahtionally focused on the use of “indicators,” organisms whose presence signals the possible presence of pathogens in the environment. Indicators are “living tracers” and should have similar behavioral characteristics and responses to environmental perturbations as pathogens (Armon and Kott, 1996). Fecal indicators have been defined as “bacteria utilized to determine the sanirary and microbiologcal quality of water” (Haack et al., 2009). The indicators occur in higher numbers and represent the pathogens that may be transmitted through the same environmental transmission route. In any given watershed or water supply, point sources (such as poorly functioning wastewater treatment plants, combined sewer overflow events, improperly managed animal confinement operations) or nonpoint sources (such as septic systems, application of animal

Milifza Carrero-Colo’n, Department of Agronomy, Purdue University, West Lafayette, IN 47907. Gene S. Wickham, The Center for Environmental Biotechnology, University of Tennessee, Knoxville, TN 37996. Ronald F. Turco, Laboratory for Soil Microbiology, Department of Agronomy, Purdue University, West Lafayette, IN 47907.

The Fecal Bacteria, Edited by M. J. Sadowsky and R. L. Whitman, 0201 1 American Society for Microbiology,Washington, DC

23

24

rn CARRERO-COLONET AL.

waste to soils, and wildlife) may be sources of indicator bacteria. Regardless of source, however, the occurrence of fecal pollution should restrict the use of surface waters for either direct consumption or full body contact (Scott et al., 2002; Bitton, 2005; Okabe and Shimazu, 2007). PHYLOGENETICS OF FECAL INDICATORS In studies of environmental quality, there are several different groups of bacteria frequently used as fecal indicators. The most widely applied approach is to use the group represented by the family Enterobucteriaceae (coliforms and fecal coliforms). Although Escherichia coli is a primary fecal indicator organism, it has been suggested that other bacteria with specific characteristics related to their original source could also be used. These other fecal indicators include members of the genus Enterococcus (formally fecal streptococci) and the anaerobic bacteria Clostridium perfringens, BiJidobacteria, and the some members of the

genus Bacteroides (Table 1 and Fig. 1). Because of their widespread use as indictors of the quality of water and other environments, this chapter covers the phylogeny, taxonomy, and physiology of these key fecal indicator bacteria. In order to define more clearly the relationship between fecal indicators and pathogens we conducted a small-subunit rRNA sequence comparison, using 35 widely reported species of fecal pathogens and their cognate indicator organisms. This analysis is represented in the phylogenetic reconstruction depicted in Fig. 1. Clearly these findings are limited to only a few regions in the tree of life. Of the more than 50 phyla (Rappe and Giovannoni, 2003; Pruesse et al., 2007; Cole et al., 2009) and innumerable families examined within the domain Bucteria, the pathogens and indcators fall into four phyla-the Proteobacteriu, the Firmicutes (formerly the low G + C gram-positive bacteria), the Actinobacteriu (formerly the high G + C gram-positive bacteria), and the Bucteroidetesand 12 fadies.

TABLE 1 Key fecal indicator bacteria used in determining fecal contamination in different environments‘ Indicator Total coliforms

Bacteria

E. coli, Enterobacter, Klebsiella, and Citrobacter

Fecal coliforms

Suggested use as indicator for: Best indicators for treatment efficiency of water treatment plants, drinking water, and ground water Recreational water and shellfish

E. coli

E. coli

Best indicator of presence of fecal material from warmblooded animals in coastal recreational waters and ground water

Fecal streptococci

S. faecalis, S. bovis, S. equinus, and S. avium

Recreational waters

Enterococci

Suggested as indicator of viruses, particularly in biosolid and marine environments

Clostridiumpefingens

E. faecalis, E. faecium, E . durans, E . gallinarum, and E. avium C. petjiingens

Bacteroides spp.

B. fragilis

Suggested as good indicators because restricted to warm-blooded animals and do not survive long in the environment

Bifidobacteria

B. bijidum, B. adolescentis, B. infantis, and B. dentium

Because of their host specificity it is useful to differentiate human from animal fecal contamination and it is good indicator of recent contamination

“Adapted from Bitton, 2005.

Better indicator of past pollution, also used as indicator of and the quality of recreational waters (viruses and parasitic protozoan)

2.

TAXONOMY, PHYLOGENY, AND PHYSIOLOGY OF FECAL BACTERIA

Over half of the fecal bacteria are Proteobacteria, and all but one of these proteobacterial species is a Garnmaproteobacterium. The lone exception to this relationship is the epsilonproteobacterial pathogen Campylobacter jejuni. Of the 12 families of fecal bacteria, the Proteobacteria comprise four: the Campylobacteraceae, with its one member mentioned above; the Aeromonadaceae, which comprises three pathogens that belong to the genus Aeromonas; the Legionellaceae, containing one species, Legionella pneumophila; and the Enterobacteriaceae. The Enterobacteriaceae are comprised of the colifom and fecal coliform bacteria, and these represent the most frequently used fecal indicator bacteria. In turn, the bacteria from the coliform grouping that is most relied on as indicators of fecal contamination are E. coli, Citrobacterfreundii, Enterobacter cloacae, and Klebsiella pneumoniae (Stewart et al., 2007). E. coli and K. pneumoniae are specifically classified as fecal coliforms. The indicator species mentioned (E. coli, C. freundii, E. cloacae, and K. pneumoniae) can be both pathogens as well as indicators. The rest of the fecal bacteria in the Enterobacteriaceae, three species of Salmonella, and two species each of Shigella and Yersinia are pathogens but not typically used as indicators. The Firmcutes comprise five farmlies of fecal bacteria: (i) the Enterococcaceae consisting of Enterococcus durans, Enterococcusfaecium, E. avium, E. gallinarum, and E.faecalis; (ii) the Streptococcaceae, comprising Streptococcus bovis and S. equinus; (iii) the Listeriaceae containing Listeria monocytogenes; (iv) the Clostridiaceae comprising Clostridium botulinum and C. pe@ngens; and (v) the Staphylococcaceae consisting of Staphylococcus aureus. The Firmicutes comprise the largest number of indicators of any of the three fecal bacteria-containing phyla. E. durans, E. faecium, E. avium, E. gallinarum, S . equinus, and S. bovis are all fecal indcators. E. faecalis and C. pefringens are both fecal indicators and fecal pathogens. S. aureus and C. botulinum are fecal pathogens but are not used as inmcators of fecal contamination. The third phylum containing fecal bacteria is the Actinobacteria. This phylum contains two families, the Bijidobacteriaceae and the Mycobac-

25

teriaceae. Three species in the genus Bijidobacterium are fecal inmcators. The Mycobacteriaceae contains a single fecal bacterium, the pathogen Mycobacterium tuberculosis. The fourth phylum depicted in Fig. 1 is the Bacteroidetes, which contains the bacterium Bacteroides fragilis, a bacterium that is both a fecal pathogen as well as an indicator organism. COLIFORMS AND FECAL COLIFORMS Domain: Bacteria Phylum: Proteobacteria Class: Gammaproteobacteria Order: Enterobacteriales Family: Enterobacteriaceae To understand the terms total coliform (TC) and fecal coliform (FC) requires an understanding of the taxonomic complex that make up these operational groups. The family Enterobacteriaceae contains many coliform bacteria and, as is indicated in Fig. 1, the term coliform can be used to represent (indicate)a variety of other near and phylogeneticdy distant bacteria. In general, coliform bacteria are thought to originate in the intestine of warm-blooded animals. However, the f d y also contains many bacteria that may or may not have an animal or fecal origin. In general, members of the f d y Enterobacteriaceae are rod-shaped, gram-negative, facultative anaerobic bacteria. They ferment sugar, producing lactic acid and other products; they are catalase positive; and most reduce nitrate. They lack cytochrome c oxidase and many have flagella, although some are nonmotile (Murray et al., 2009). Many members of this family are part of the normal flora of the intestines of humans and other warm-blooded animals and are typically found in concentrations ranging from lo6 to lo9 coliforms per gram of feces. Some of these bacteria, however, also freely exist in secondary or environmental habitats including water, soil, sand, and sediments. The major set of phenotypic characteristics defining the Enterobacteriaceae family include the following: small gram-negative rods, facultative but preferring aerobic metabolism, oxidase

Entembacter cloacae, subsp. cloacae, (Pll) Klebsiella pneumoniae,

(P)

inia entercolitica subsp. entercolitica, (P) rsinia pestis, (P) reundii, (Pll)

sp. Enterica Serovar Gallin, (P)

phila subsp hydrophila, (PI tatasubsp punctata, (P)

LEnterococcus avium, (I) Enterococcusgallinarum, (I)

I

terococcus faecalis, (Pll)

I

Streptococcusbows, (I) genes, (P) hylococcus aureus subsp aureus, (P)

Demococcus radiodurans R 1

0.04

2. TAXONOMY, PHYLOGENY, AND PHYSIOLOGY OF FECAL BACTERIA W 27

negative, catalase positive, can reduce nitrates to nitrites, do not require Na’ for growth, can ferment D-glucose, contain enterobacterial common antigens, and have a mol% G+C content of 38 to 60 (Jandaand Abbott, 2006). Coliforms are aerobic or facultative, gramnegative, non-spore-forming rods that inhabit the intestinal track of all vertebrates. However, Klebsiella pneumoniae and K. rhinoscleromatis, the first members of the coliform group as described by Friedlander and Frisch in 1882 (Leclerc et al., 2001), are also commonly found in soil. More recently, the principal members of the TC group are considered to be E. coli, Enterobacter aerogenes, E. cloacae, and Citrobacter jieundii (Leclerc et al., 2001) As indcated by Leclerc et al. (2001), the classification methods for coliform bacteria are complex and have many problems. Originally, classification of coliform bacteria was based on fermentation of sucrose and dulcitol, production of indole and acetylmethylcarbinol, and gelatin liquefaction. Using this system some 128 coliform types were originally defined by MacConkey, and t h s was later expanded by Bergey and Deehan (1908) to 256 microorganisms (Kabler et al., 1964; Leclerc et al., 2001). Other

differentiation methods for this group of bacteria where subsequently added, and included the ratio of H2:COz produced from glucose, an inverse correlation between Voges-Proskauer and methyl red reaction, and the assidation of citrate. This prompted the introduction of the IMViC formula (indole production, methyl red reaction, Voges-Proskauer, and citrate utilization). Using the data resulting from the use of these traits in the classification of coliforms, there are 16 possible combinations of classification, with 3 belonging to the E. coli group, 3 belonging to the Aerobacter aerogenes group, and 10 belonging to the “intermediate” group (Leclerc et al., 2001). It should be noted that Parr (1939), as referenced by Leclerc et al (2001),concluded that the coliform bacterial group is composed of a collection of “highly intergrading and somewhat unstable bacteria.” Because of the complexity and the size of the coliform group and reoccurring problems with classification of these organisms, DNADNA relatedness has now been used as a defining group characteristic where strains had to be 70% or higher with less than 5°C A T, (Leclerc et al., 2001) to be considered similar. This requirement resulted in a redefinition of

FIGURE 1 Evolutionary relationships of fecal bacteria. The small-subunit, or 16S, rRNA-based evolutionary relationships of representative fecal bacteria are shown. A total of 36 (35 fecal bacteria sequences and 1 outgroup sequence) nearly full-length 16s rRNA sequences from the Ribosomal Database Project (Larsen et al., 1993; Cole et al., 2009) and/or the SILVA database (Pruesse et al., 2007) were aligned in ARB (Ludwig et al., 2004). A modified version of the Lane mask (Lane, 1991) was used to define 1344 unambiguously aligned nucleotide positions for comparative analysis by quartet puzzling (Strimmer and vonHaeseler, 1996).Each sequence is denoted as an indicator (I), pathogen (P), or both (P/I). Deinococcus vadioduvan R 1 was used to root the tree. The scale bar represents 0.4 fixed mutations per nucleotide position. Whenever available, sequences from type strains were used. In instances where sequences from non-type strains were not available, non-type strain sequences were chosen based on their length and quality. The accession identifier for each sequence is shown in parentheses below. Type strains: Aeromonas caviae (X60408),Bacteroidesjagilis (X83935),Bijidobacterium adolescentis (M58729),Bijidobacterium bijidum (M38018), Bifidobacteriurn dentium @86183), Campylobacter jejuni (L14630), Citrobacterjeundii (AJ233408), Clostridium botulinum A (L37585), Clostridium pefingens (M59103),Enterobacter aerogenes (AB004750),Enterobacter cloacae (AJ251469), Enterococcus avium (AF133535),Enterococms durans (AJ420801), Enterococcusfaecium (AJ276355), Escherichia coli (X80725),Klebsiella pneumoniae (Y17656), Listeria monocytogenes (X56153), Salmonella enterica subsp. enterica serovar Typhi (Z47544), Streptococcus bovis (AB002482),Streptococcus equinus (X58318),Aevomonas hydrophila (X74677), and Legionella pneumophila (M59157). Non-type strains: Aevomonas veronii (X74684), Citvobacter kosevi (AF025372), Enterococcus faecalis (AJ301831), Enterococcus gallinavuvn (AF039898), Klebsiella oxytoca (AY292868), Mycobacterium tuberculosis (AJ131120), Salmonella enteritidis (EU118104), Salmonella enterica subsp. enterica serovar Gallinarum (EU073018), Shigella dysenten’ae (X96966), Shigella sonnei (EU009199), Staphylococcus auveus (D83357), Yersinia entercolitica (M59292),and Yersinia pestis (AF366383).

28

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CARRERO-COLONET AL.

the coliform group to 19 genera and 80 species from the Enterobacteriaceae and o-nitrophenylP-galactopyranoside-using organisms. All bacteria that belong to this group are members of the family Enterobacteriaceae, and have the ability to ferment lactose to gas within 48 hours when incubated at 35°C (Geldreich et al., 1958). Moreover, all strains contain the enzyme P-galactosidase and are inclusive of the genera Escherichia, Klebsiella, Enterobacter, and Citrobacter (Edberg et al., 2000; Leclerc et al., 2001). Members can be routinely found in the environment, but are not always associated with fecal pollution. The group includes all aerobic and facultative anaerobic, gramnegative, non-spore-forming rod-shaped bacteria that produce a red colony with a metallic sheen within 24 hours at 35°C on an endotype medium containing lactose (Romprt et al., 2002). They are also capable of growth in the presence of bile salts, are oxidase negative, and display P-galactosidase activity (Leclerc et al., 2001). Lactose fermentation is dependent on the enzyme formic-hydrogenase, which converts formic acid to CO;! and H2. In contrast, the term FC is given to those members of the TC group that are able to ferment lactose at 44.5"C; at this temperature the growth and activity of environmental coliform-type bacteria are suppressed (Turco, 1994). Genera belonging to the fecal coliform group include E . coli and Klebsiella. However, as mentioned previously, Klebsiella is not always of fecal origin; as a result, the occurrence of Klebsiella is not a strong indicator of fecal contamination because it is generally not present in high numbers in feces and is found to occur widely in soil and water. Klebsiella is also found in high concentrations in some non-feces-related industrial wastes and is able to grow in nutrient-rich waters (Leclerc et al., 2001). Other genera of the FC group that do not always originate in feces include Enterobacter and Citrobacter (Leclerc et al., 2001). Clearly, the terms coliform and fecal coliform provide a widely applied operational separation but are not true taxonomic definitions.

E . coli E. coli was originally named Bacterium coli commune by Theodor Escherich in 1885. After much taxonomic controversy, it was renamed E. coli and is now situated in the family Enterobacteriaceae (Jandaand Abbott, 2006). This bacterium can be subclassified into three groups: commensal, darrheagenic, and extraintestinal. The commensal E. coli is nonpathogenic and is often considered to be a normal inhabitant of the gastrointestinal tract of warm-blooded animals. Although these bacteria are considered beneficial to the host, some can cause hsease in immunocompromised patients. Within the darrheagenic E. coli, six pathogenic groups can cause infection in the gastrointestinal tract. The extraintestinal E. coli is composed of strains capable of causing hsease outside of the intestine. E. coli is a thermotolerant organism that is able to produce indole fiom tryptophan at a 44 2 0.5"C, will not use citrate as its sole carbon source, does not produce acetyl methyl carbinol, and is methyl red positive (Rompr2 et al., 2002). E. coli is part of the normal flora of mammals and is a common gram-negative facultative anaerobe found in human feces. However, the bacterium can be found in sewage, treated effluent, natural waters, sand, sediment, and soils. Since the 1980s, this bacterium has been widely used as inhcator of water treatment safety, replacing the less well defined FC and TC groups (Edberg et al., 2000). Its presence in the environment is used to indicate a threat to public health. E. coli is found in mammahan feces in numbers as high as lo9 per gram of feces and often comprises -1% of the total bacterial biomass. More importantly, E. coli are considered a stable member of intestinal community, as they are not as sensitive to dietary considerations as bacteria in other genera (Leclerc et al., 2001). Approximately 95% of E. coli isolates can metabolize the complex substrate 4-methylumbeWerone-~-~-glucuronide, using P-glucuronidase resulting in the formation of a flourescent product 4-methylumbelhferone. Ths particular characteristic, when combined with its abhty to grow at 44.5"C, is used for the defined substrate technology (DST) to identify E. coli. The DST is included in commercially

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TAXONOMY, PHYLOGENY, AND PHYSIOLOGY OF FECAL BACTERIA W 29

available methods such as Colilert and Colisure (Idem, Westbrook, ME). Lactose fermentation in E. coli is affected by media, temperature, and incubation time and it is interesting to note that 10% of strains are genetically not capable of producing gas from t h s fermentation (Leclerc et al., 2001). The survival of E. coli in the environment is influenced by many factors includmg temperature, protozoa grazing, exposure to UV ra&ation, and the availability of nutrients. Studies have shown that E. coli can survive in water containing a reasonable amount of microflora and nutrients at 15 to 18°C for -4 to 12 weeks (Edberg et al., 2000). Coliforms (total or fecal) have been used extensively to determine the presence of fecal pollution in: drinking well water (Lavoie, 1983; Edberg et al., 2000; van Lieverloo et al., 2007), surface waters (Blanch et al., 2004; Park et al., 2006; Haack et al., 2009), pulp and paper mill water (Gauthier and Archibald, 2001), storm water (Noble et al., 2004), reclaimed water (Harwood et al., 2005), seawater (Efstratiou et al., 2009), estuaries (Berthe et al., 2008), river water palters et al., 2007; Gronewold et al., 2008), and watersheds systems (Dorner et al., 2007; Plummer and Long, 2007). Differences in ratio of T C to FC bacteria have been suggested as an indicator of the sanitary quality of soil and water environments, and as a possible source traclung method (Leclerc et al., 2001). However, although the potential of using dfferences in TC and FC for microbial source trackmg stuhes has been extensively investigated, it has been concluded that the ratios are not capable of differentiating between host sources (human, animal, etc.) (Toranzos and McFeters, 1997). ENTEROCOCCI Domain: Bacteria Phylum/Division: Firmicutes Class: Bacilli Order: Luctubacillales Family: Enterococcaceae Genus : En terucoccus

The classification of fecal streptococci was previously used to describe those streptococci associated with the gastrointestinal tract (Leclerc et al., 1996). As of 1937, these bacteria were classified into four main groups: pyogenic, viridans, lactic, and enterococcus. Streptococcusfaecalis, S.faecium, S. durans, and S. avium were initially used to describe the four major groups in the genus Enterococcus. This classification was based on morphology and biological and serological differences (Morrison et al., 1997). These organisms were subsequently split into three genera: Lactococcus, Enterococcus, and Streptococcus (Godfree et al., 1997). At that time, the majority of the fecal species were assigned to the genera Enterococcus or Streptococcus, but advanced taxonomic studies using 16s rRNA sequence analysis show that most of the species actually belonged to the genus Enterococcus. Early on, the genus Enterucoccus was defined as lactic acid bacteria and as gram-positive, catalase negative, and nonhemolytic diplococci that occur in the gastrointestinal tract (Hartman et al., 1966) and contain the Lancefield group D antigen, but there are some exceptions (Devriese et al., 1993; Leclerc et al., 1996). Enterococcus can be found in soil, water, dairy products, food, and plants (Fisher and Phillips, 2009), but the common commensal forms in the human intestine are E . faecalis (90 to 95%) and E. faecium (5 to 10%). Members have the ability to grow at temperatures between 10 and 45"C, at pH 9.6, in 6.5% NaCl broth (Domig et al., 2003; Tendolkar et al., 2003), grow in 0.1% methylene blue milk (Slanetz and Bartley, 1964), produce ammonia in peptone broth (Hartman et al., 1966; Leclerc et al., 1996; Godfree et al., 1997), and survive at 60°C for 30 minutes. The enterococci are gram-positive, elongated cocci that may occur indvidually, in pairs, or in chains and can occur in the human colon at levels reaching 10' CFUs per gram feces, although they wdl typically only represent 1% of the human intestinal flora (Tendolkar et al., 2003). In addition, enterococci do not have cytochrome enzymes and, for this reason, most species are catalase negative, but some

30

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wdl express a pseudocatalase activity. They are also benzidme negative, esculin positive, and all species produce leucine aminopeptidase. Enterococci are nonfilamentous and some strains are reported to be motile (FacMam et al., 2002). Although they can tolerate oxygen, they are facultative anaerobes that follow the EmbdenMeyerhof-Parnas homofermentative pathway for metabolism. These bacteria produce L (+) lactic acid from glucose fermentation at an optimal growth temperature of 35°C. Their cell wall is associated with glycerol teichoic acid antigen (D antigen), although detection of this antigen in some species is hfficult. Their membrane is composed of long-chain fatty acids (saturated or monounsaturated) with lysineD-asparagine linkages in their peptidoglycan (Domig et al., 2003). Based on DNA-DNA hybrihzation, 16s r RNA sequencing, and cell protein analysis, this genus is comprised of 23 species with a G+C composition of 38 to 45 mol% (Hardie and Whiley, 1997). Functionally, enterococci are hvided into five groups based on their ability to produce acid from mannitol and sorbose, and hydrolyze arginine (Facklam et al., 2002). Group I is composed of E. avium, E. malodoratus, E. ra8nosus, E. pseudoavium, E. saccharolyticus, E. pallens, and E. gilvus. They do not hydrolyze arginine, but produce acid fiom carbohydrate broth, especially mannitol, sorbose, and sorbitol. They also utilize pyruvate and sucrose as growth substrates, and are sometimes pigmented. Group I1 is composed of E. faecalis, and the Lactococcus spp. E. faecium, E. caselijlavus, E. mundtii, and E. gallinarum. They hydrolyze arginine and will produce acid from mannitol, but not fiom sorbose. Group I11 is composed of E. durans, E. porcinus (E. villomm), E. hivae, E. dispau, mannitol-negative variants of E.faecalis and E.faecium, and E. ratti. Ths group does not produce acid from mannitol or sorbose, but they do hydrolyze argmine. There are three species found in group IV: E. asini, E. sulfurem, and E. cecorum. They also do not produce acid fiom mannito1 and sorbose and, in contrast to Group 111, they do not hydrolyze arginine. Members of

group V include variant strains of E. cassel$lavus, E. gallinarum, and E. faecalis that are unable to hydrolyze arginine (Facklam et al., 2002). In general, the enterococci can successfully inhabit harsh environments and express a metabolic flexibility that gives the group the ability to catabolize a myriad of energy sources, including glycerol, lactate, malate, citrate, arginine and agmatine, and many a-keto acids. They are resistant to 40% bile salts, azide, detergents, sodium hypochlorite, heavy metals, ethanol, and, although not being a sporeforming bacteria, long periods of desiccation (Huycke, 2002). In addition, they are Pglucosidase positive and urease negative. With the exception of E. saccharolyticus, they are Voges-Proskauer positive, leucin arylamidase positive, P-glucuronidase negative (with the exception of E. cecomm), and will hydrolyze aesculin. Most members of this genus can produce acid from N-acetylglucosamine, amygdalin, arbutin, cellobiose, D-fructose, galactose, P-gentiobiose, D-glucose, lactose, maltose, Dmannose, methyl-P-D-glucopyranoside, ribose, salicin, and trehalose (Devriese et al., 1993; Huycke, 2002). This ability likely enhances their competitiveness in various environments, especially in the intestinal tract. It is also believed this group feeds on nonabsorbed sugars in the intestine. However, they cannot produce acid from D-arabinose, cellobiose, glycogen, D-fucose, L-fucose, a-methyl-D-xyloside, pullulan, and L-xylose (Devriese et al., 1993). Members of this genus are resistant to cephalosporins and aminoglycosides (except for gentamicin and streptomycin) and they have reduced sensitivity toward penicillin, carbapenems, and other p-lactams. They are also resistant to lincosamides, trimethoprim, and sulphonamides and have acquired resistance to chloramphenicol, tetracyclines, macrolides, streptogramins, and lincosamides through gene transfer including sex pheromones, plasmids, and transposons (Morrison et al., 1997). Enterococci are useful indicators of fecal contamination in aquatic environments because they are always present in animal feces,

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are usually unable to multiply in the environmentalthough they can show some reproductive capability under nonextreme environmental conmtions-and have survival rates simdar to waterborne pathogenic bacteria (Lleo et al., 2005). Although some species have been found in secondary nonfecal habitats (soil, sand, water, and sediments), they are still thought to be good indicators of fecal contamination for recreational marine waters. Several stumes have investigated the potential use of enterococci as indicators of fecal contamination of frozen sea food (Raj et al., 1961), freshwater fish (Geldreich and Clarke, 1966), drinlung water (Edberg et al., 1997), water stored in traditional vessels (Tandon et al., 2007), estuaries (Morrison et al., 2008), recreational waters (Leskinen and Lim, 2008; Wade et al., 2008; Wong et al., 2009), surface waters (Ibekwe and Lyon, 2008; Haack et al., 2009), and sea water (Efitratiou et al., 2009). The enterococci have been used in watershed studies to evaluate different areas including: lake water, stream water, near shore sand, backshore sand, Cladophorales, and periphyton (Byappanahalli et al., 2008). The potential of this genus as a fecal indicator for microbial source tracking has also been stumed, especially because its host range is limited mostly to humans and several other animals (Simpson et al., 2002; Wheeler et al., 2002). Some examples of recent microbial source tracking studies include urban watershed and lake studes (Tiang et al., 2007; Ahmed et al., 2008) and estuaries (Korajkic et al., 2009). ANAEROBIC FECAL INDICATORS

Clostridium perfringens Domain: Bacteria Division: Firmicutes Class: Clostridia Order: Clostridiales Family: Clostridiaceae Genus: Clostridium Based on 23s rRNA homologies and 16s rRNA classification, the genus Clostridium is

31

classified into four groups, I to IV. This eubacteria group is a low G+ C content subgroup of bacteria in the gram-positive phylum (Canard et al., 1992). Clostridium pefingens (formerly known as Clostridium welhii) is a gram-positive, oblige anaerobic, nonmotile (Rood and Cole, 1991), rod-shaped bacterium that produces a heat-resistant endospore. The bacterium is ubiquitous in soil, but is commonly associated with feces (Bisson and Cabelli, 1980). It was linked to food poisoning as early as 1899 (Nakamura and Schulze, 1970). C. pefingens is found in the gastrointestinal tract ofboth humans and animals, and is routinely detected in sewage (Rood and Cole, 1991; Canard et d., 1992).This bacterium is a member of the sulfite reducing clostrida, an auxotroph, and fermentative microorganism that can grow in mema containing carbohydrates, 11 amino acids, and vitamins. During growth, the bacterium produces H2 and COz, helping to ensure an anaerobic atmosphere. Under the right conmtions, C. pefiingens has a rapid doubling time (20 min) that results in a high growth yield. Although it is a commensal organism for both humans and animals, under the appropriate conditions, it can cause a variety of diseases (Canard et al., 1992) includmg gangrene, food poisoning, necrotizing enterocolitis of ihnts, enteritis necroticans in humans, and lamb dysentery ovine enterotoxemia. Spores of t h s bacterium are resistant to environmental stress and the vegetative cells are resistant to the antibiotics tetracycline and erythromycin. They are, however, sensitive to chloramphenicol and penicillin, which is the drug of choice for treatment. C. pefringens can produce five major types oftoxins-A, B, C, D, and E-that are directly linked to the mseases they cause. In the case of secretory diarrhea, the disease-causing mechanisms is linked to the release of enterotoxins that causes intestinal damage, this is accompanied with acute pain and abdominal cramping in the host (Meyer and Tholozan, 1999). In Europe, C. pefiingens has been used as an indcator of fecal contamination, but because of a lack of reliable detection method for this

32

CARRERO-COLONET AL.

anaerobe it was not used in the United States until the 1980s. The value of C. perfvingens as an indicator is based on its ability to occur in both vegetative and spore forms; it is suggested that the spores are acting as a conservative tracer of long-term contamination. Therefore, its detection can indicate both past and current pollution depending on the ratio of spores to vegetative cells (Bisson and Cabelh, 1980). C. pefiingens has been studied as an indicator of fecal contamination in marine waters (LaBelle et al., 1980; Fujioka, 2001; Shibata et al., 2004), surface water (Horman et d., 2004; Wilkes et d., 2009), aquifers (Paul et al., 1995), wetlands (Song et al., 2008), wastewater treatment (Barrett et al., 2001), river water (Desmarais et al., 2002), viral pathogens in shellfish (Muniain-Mujika et al., 2003), drinkmg water (Lisle et al., 2004), lakes and reservoirs (Brookes et al., 2005), recreational waters (Wiedenmann et al., 2006), tropical streams (Plancherel and Cowen, 2007), and storm water runoff (Cizek et al., 2008). Most studies concluded that C. pefingens can be use as a “surrogate” indicator of fecal contamination. However, because of the environmental resistance of its spores, this bacterium has been suggested as an indcator of past pollution.

Bacteroides Domain: Bacteria Phylum: Bacteroidetes Class: Bacteroidetes Order: Bactevoidales Family: Bacteroidaceae Genus: Bacteroides Bacteroides spp. are gram-negative, nonpigmented, motile or nonmotile bacilli that are part of the normal flora of the gut warmblooded animals (Fiksdal et al., 1985) and are one of the five predominant bacteria in the intestinal tract of humans (Duerden, 1980). Bacteroides spp. are present in much greater numbers than E. coli, and can account for up to 30% of the total fecal isolates, with the most common species being B. vufgatus, B. distasonis, and B. thetaiotaomicron (10” per dry

weight of feces). Whereas B. fragilis, B. ovatus, B. eggerthii, B. un$ormis, and Bactevoides B5-21 are also present, but less numerous (lo9per dry weight of feces) (Duerden, 1980). Organisms in the genus Bactevoides are classified based on the ability to ferment a variety of carbohydrates and their tolerance to oxygen, although their nutritional requirements are not complex. They are able to grow in meha containing carbohydrates, ammonia, hemin, vitamin B12, inorganic salts, and a reducing agent (Salyers, 1984). They are able to ferment the sugars glucose and mannose and their products of fermentation include acetate (decarboxylation ofpyruvate), succinate (NADH oxidation), and propionate (decarboxylation of succinate). They have high growth yields but slow growth rates because of the limited resources in the gastrointestinal tract. In the colon, their main source of energy comes from of polysaccharides because sugars are limited. Some example polysaccharides include chondroitin sulfate and arabinogalactan, which are not absorbed in the gastrointestinal tract. Some species can also ferment polysaccharides produced by the host organism and plant polysaccharides, which gves them some competitive advantage in the colon. There is limited uptake of amino acids and proteins, and Bacteroides cannot use them as a sole source of energy especially when growing on carbohydrates. Although this bacterium is an obligate anaerobe, it can survive extended exposure to oxygen (Salyers, 1984). This genus is characterized by its resistant to kanamycin, neomycin, penicillin, aminoglycosides, and p-lactam antibiotics. Some are also resistant to tetracycline (Salyers, 1984) and tolerant to taurocholate and Victoria blue 4 R (Duerden, 1980). They are susceptible to chloramphenicol, metronidazole, clindamycin (Salyers, 1984), and rifampicin (Duerden, 1980). The abihty of Bacteroides to grow in natural environments is limited by factors including grazing by protozoa, bacteriophage infection, cell death and breakdown of DNA, temperature, pH, initial cell concentration, and cell aggregate size (Okabe and S h a m , 2007). The persistence

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of ths indcator bacterium in the environment seems to be related to protozoan grazing, with a longer persistence at lower temperatures due to low number of predators (Bell et al., 2009). Some species belonging to the Bacteroides include B. fragilis, B. vulgatus, B. thetaiotaomicron, B. un$ortnis, B. variabilis/egqerthii, and B. splanchnicus. Cells in this group are surrounded by a membrane composed of branched, long chain fatty acids, sphingolipids, lipopolysaccharide, capsular material, and inner and outer membranes (Salyers, 1984). Originally, B. fvagilis contained a number of subspecies, hence their apparent high numbers in the gastrointestinal tract. This was clarified by further DNA-DNA homology and other tests that reveal that some of the subspecies were serologically distinct with different polysaccharide utilization patterns (Salyers, 1984). B.jagilis has a mol% G + C composition of 40 to 48%. Since the 1990s, B. fvagilis has been suggested as an alternative fecal indicator. As was discussed previously, because B. fvagilis does not survive well in aerobic conditions such as freshwater, the use of this organism as an indicator shows recent contamination events or extensive contamination. However, antigen from bacteria can be detected for long periods after the organism has died (Fiksdal et al., 1985), confusing results from serological methods for quantification. Recent molecular tools to detect members of the Bacteroides have provided a fast and reliable approach for the detection of the bacteria (Kildare et al., 2007). Other studies have concluded that Bacteroides spp. are useful in discriminating human and nonhuman sources of fecal pollution (Gawler et al., 2007).

Bifidobacterium Domain: Bacteria Phylum: Actinobacteria Class: Actinobacteria Subclass: Actinobacteridae Order: Bijidobacteriales Family: Bijidobacteriaceae Genus: Bijidobacteriurn

Described in the 1900s by Tissier, the Bijdobacterium genus consists of gram-positive, multiple branching rods that are nonmotile, non-spore-forming, non-gas-producing, anaerobic, catalase negative, and saccharoclastic bacteria that are among the most abundant microorganismsin the large intestine. They have an optimum pH range from 6.5 to 7.0, but do not grow below pH 4.5 and above pH 8.5. Members of this genus can catabolize hexose, using the enzyme fructose-6-phosphate phosphoketolase, and produce acetate, lactate ethanol, and formate. This mechanism of hexose catabolism is unique for this genus, as it is not found in other gram-positive intestinal bacteria. Their typical habitat includes the human and insect intestine, the oral cavity, food, the animal gastrointestinal tract, and the sewer (Ventura et al., 2004). Originally proposed as belonging to the family Lactobacillaaceae, the bacterium was first named Lactobacillus bijidus (Biavati et al., 2000). It was later found that the differences in carbohydrate utilization patterns separate Bijidobacterium from the Lactobacillaaceae. The genus Bijidobacterium belongs to the order Bijidobacteriales. The family Bijidobacteriaceae and has 32 recognized species, including: B. adolescentis, B. bijidum, B. infantis, B. dentium, and B. longum (Biavati et al., 2000). Bifidobacteria are found in the gastrointestinal tract of humans and animals where they coexists with other anaerobic bacteria. Bijidobacterium is identified based on cell morphology, sugar fermentation reactions, complex polysaccharides and mucin profiles, electrophoretic mobility of enzymes, and DNADNA hybridization (Ventura et al., 2004). Using rRNA sequencing, more specific classification methodology has been introduced (Holzapfel et al., 2001). Species belonging to the genus Bijidobacteriurn exhibit a 93% rRNA sequence identity with other members of the genus. They have a genome of 2,256,646 base pairs, with a mol% G + C composition of 55 to 67 (Biavati et al., 2000). The cell wall of the bifidobacteria is composed of a thick peptidoglycan envelope containing polysaccharides,

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proteins, and teichoic acids. These organisms have been shown to be antagonistic toward the growth of some intestinal pathogens, such as E. coli and Shigelh, possibly due to their secretion of the antimicrobial agent bifidocin B (Biavati et al., 2000) Bifidobacteria are resistant to a wide variety of antibiotics (Biavati et al., 2000; Ventura et al., 2004) and have complex nutritional requirements for nitrogen, vitamins, other growth factors, and metals. They also belong to the probiotic group of microorganisms that are thought to aid the host animal in improving the balance of the intestinal microflora. Bifidobacteria are also saccharolytic and are important contributors of sugar fermentation in the colon. The microorganisms use sucrosephosphorylase to metabolize sucrose, resulting in the formation of glucose-1-phospate and fructose, and use P-galactosidase to reduce lactose content. These pathways lower the intestinal pH and are thought to have bactericidal effects toward both gram-negative and -positive, possibly pathogenic, bacteria (Biavati et al., 2000). Ferric iron is converted to ferrous iron at the cell surface and then is transported into the cell by a divalent metal permease. This mechanism of iron uptake by the Bifidobacteria is known to hinder proliferation of other organisms because it creates an iron deficiency in the environment wentura et al., 2004). They also produce a variety of vitamins-thiamine, pyridoxine, folic acid, and nicotinic acid-and use ammonium salts as their sole source of nitrogen. Because of the specificity of this genus in relation to their ecological origin, isolates exhibit different optimal temperature range. Isolates from humans can grow at 36 to 38"C, whereas isolates from animals are able to grow at higher temperatures ranging from 41 to 43°C. With the exception of B. psychroaerophilum, bifidobacteria can grow at a minimum temperature of 20°C (Biavati et al., 2000; Ventura et al., 2004). It has also been shown to have some antimutagenic and anticarcinogenic properties in laboratory studies (Ventura et al., 2004).

Several studies have suggested that bifidobacteria are a good indicator of recent fecal contamination and may be useful to determine the source(s) of fecal pollution (animal or human) in raw sewage (Resnick and Levin, 1981; Lynch et al., 2002; Nebra et al., 2003; Bonjoch et al., 2005), human and animal wastewater (Bonjoch et al., 2004), meat and meat products (Beerens, 1998), the sheep meat production chain (Delcenserie et al., 2008), estuaires (King et al., 2007); watershed systems (Lamendella et al., 2008), and tributaires and drinking water reservoirs (Long et al., 2005). REFERENCES Ahmed, W., M. Hargreaves, A. Goonetilleke, and M. Katouli. 2008. Population similarity analysis of indicator bacteria for source prediction of fecal pollution in a coastal lake. Mar. Pollut. Bull. 56:1469-1475. Armon, R., and Y. Kott. 1996. Bacteriophages as indicators of pollution. Crit. Revs. Environ. Sci. Technol. 26:299-335. Barrett, E. C., M. D . Sobsey, C. H. House, and K. D . White. 2001. Microbial indicator removal in onsite constructed wetlands for wastewater treatment in the southeastern U.S. Water Sci. Technol. 44: 177-182. Beerens, H. 1998. Bijidobacteria as indicators of fecal contamination in meat and meat products: detection, determination of origin and comparison with Escherichia coli. Int. J . Food Micro. 40:203-207. Bell, A., A. C. Layton, L. McKay, D . Williams, R. Gentry, and G . S. Sayler. 2009. Factors influencing the persistence of fecal bacteroides in stream water. J. Environ. Qual. 38:1224-1232. Bergey, D . H., and S. J. Deehan. 1908. The colon-aerogenes group of bacteria. J . Med. Res. 19:175-200. Berthe, T., A. Touron, J. Leloup, J. Delofie, and F. Petit. 2008. Fecal-indicator bacteria and sedimentary processes in estuarine mudflats (Seine, France). Mar. Pollut. Bull. 57:59-67. Biavati, B., M. Vescovo, S. Torriani, and V. Bottazzi. 2000. Bijidobacteria: hstory, ecology, physiology and applications. Ann. Miuobiol. 50:117-131. Bisson, J. W., and V. J. Cabelli. 1980. Clostridium pe$ringens as a water pollution indicator. J . Water Pollut. Control Fed. 52:241-248. Bitton, G. 2005. Microbial indicators of fecal contamination, p. 153-171. In G. Bitton (ed.), Wastewater Microbiology. John Wiley & Sons, Inc., Hoboken, NJ.

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2. Microbiological and Biochemical Properties. Soil Science Society of America, Madison, WI. van Lieverloo, J. H., E. J. Blokker, and G. Medema. 2007. Quantitative microbial risk assessment of distributed drinking water using faecal indicator incidence and concentrations.J. Water Health Suppl 5. 1~131-149. Ventura, M., S. D. van, G. F. Fitzgerald, and R. Zink. 2004. Insights into the taxonomy, genetics and physiology of bifidobacteria. Antonie Van Leeuwen. 86:205-223. Wade, T. J., R. L. Calderon, K. P. Brenner, E. Sams, M. Beach, R. Haugland, L. Wymer, and A. P. Dufour. 2008. High sensitivity of children to swimming-associated gastrointestinal illness: results using a rapid assay of recreational water quality. Epidemiology 19:375-383. Walters, S. P., V. P. Gannon, and K. G. Field. 2007. Detection of Bacteroidales fecal inhcators and the zoonotic pathogens E. coli 0157:H7, Salmonella, and Campylobacter in river water. Enoiron. Sci. Technol. 41: 1856-1862. Wheeler, A. L., P. G. Hartel, D. G. Godfrey, J. L. Hill, and W. I. Segars. 2002. Potential of Enterococcus faecalis as a human fecal indicator for microbial source tracking. J . Environ. Qual. 31:1286-1293. Wiedenmann, A., P. Kruger, K. Dietz, J. M. Lopez-Pila, R. Szewzyk, and K. Botzenhart. 2006. A randomized controlled trial assessing infectious disease risks from bathing in fi-esh recreational waters in relation to the concentration of Escherichia coli, intestinal Enterococci, Clostridium pe$?ingem, and somatic coliphages. Environ. Health Perspect. 114:228-236. Willces, G., T. Edge, V. Gannon, C. Jokinen, E. Lyautey, D. Medeiros, N. Neumann, N. Ruecker, E. Topp, and D. R. Lapen. 2009. Seasonal relationships among indicator bacteria, pathogenic bacteria, Cryptosporidium oocysts, Giardia cysts, and hydrological indices for surface waters within an agriculturallandscape. Water Res. 43:2209-2223. Wong, M., L. Kumar, T. M. Jenkins, I. Xagoraraki, M. S. Phanikumar, and J. B. Rose. 2009. Evaluation of public health risks at recreational beaches in Lake Michigan via detection of enteric viruses and a human-specific bacteriological marker. Water Res. 43:1137-1149.

THE GUT MICROBIOTA: ECOLOGY AND FUNCTION Benjamin P. Willing andJanet K. Jansson

The gastrointestinal (GI) tract is teeming with an extremely abundant and &verse microbial community. The members of this community have coevolved along with their hosts over millennia. Until recently, the gut ecosystem was viewed as black box with little knowledge of who or what was there or their specific functions. Over the past decade, however, this ecosystem has become one of fastest growing research areas of focus in microbial ecology and human and animal physiology. This increased interest is largely in response to studies tying microbes in the gut to important diseases afflicting modern society, including obesity, allergies, inflammatory bowel diseases, and diabetes. Although the importance of a resident community of microorganisms in health was first hypothesized by Pasteur over a century ago (Sears, 2005), the multiplicity of physiological changes induced by commensal bacteria has only recently been recognized (Hooper et al., 2001). The term “ecological development” was recently coined to support the idea that development of the GI tract is a product of the

genetics of the host and the host’s interactions with resident microbes (Hooper, 2004). The search for new therapeutic targets and disease biomarkers has escalated the need to understand the identities and functions of the microorganisms inhabiting the gut. Recent studies have revealed new insights into the membership of the gut microbial community, interactions within that community, as well as mechanisms of interaction with the host. This chapter focuses on the microbial ecology of the gut, with an emphasis on information gleaned from recent molecular studies. MEMBERSHIP (WHO IS THERE) In most animals studied to date, the microbial community in the most densely populated regions of the GI tract is dominated by bacteria with total cell counts reaching as high as a hundred billion cells per gram of digesta (Hooper and Gordon, 2001a). In humans, this number is equivalent to approximately 1 kg of the human mass. The gut is also colonized by microbes from the domains Archaea and Eukarya including protozoa, yeast, and fungi. However, only an estimated 20% to 25% of the gut inhabitants have been cultured to date. Therefore, most recent studies have used molecular approaches to identi@ community members in the gut. This research area

Benjnmin P. Willing, Michael Smith Laboratories, University of British Columbia, Vancouver, BC, Canada V6T 124. Janet K. Jansron, Department of Ecology, Earth Sciences Division, Lawrence Berkeley National Laboratory, Berkeley, CA 94720.

The Fecal Bacteria, Edited by M. J. Sadowsky and R. L. Whihnan, 0 2011 American Society for Microbiology, Washington, DC

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40 W WILLING AND JANSSON

has resulted in the terms microbiome (referring to collective microbial genes in the community) and microbiotu (referring to the collective microorganisms present in the community). Most attention has been devoted to bacterial components of the gut microbiota and, thus, they are the focus of this chapter. The indigenous microbiota that colonizes the GI tract comprise the resident, autochthonous members of the community and are also referred to as the commensal microbiota. The commensal relationship implies that one member of the association benefits while the other is unaffected. However, in most cases, this is a mutualistic relationship. The microbiota benefit from the host that provides a nutrient rich and hospitable environment, while the microbiota provide nutrients that would not otherwise be available to the host, regulate immune development, and create a barrier to infection from pathogens. There are also transient (allochthonous)bacteria that are regularly ingested, but are unable to colonize the gut under normal conditions (Berg, 1996). This is not to say that allochthonous organisms cannot have an effect on the host. The majority of probiotic bacteria would be considered allochthonous-when the host ceases to consume the live organisms they are transient and do not persist at high levels.

Methods To Analyze the Microbiota Until the late 199Os, the microbiota of the GI tract was nearly exclusively studied using selective culture techniques. The inability to culture the majority of gut microbes because of unknown growth requirements and potential dependence on co-colonizing bacteria or host factors (Zoetendal et al., 2004), spurred the establishment of molecular-based approaches as the new standard. Most molecular approaches utilize the 16s rRNA gene as a phylogenetic marker because of its genetic stability and composition of conserved and variable regions (Woese et al., 1990). The establishment of public databases of 16s rRNA gene sequences (http://rdp.cme.msu.edu/ and http://www .ncbi.nlm.nih.gov/) from cultured organisms

allows for taxonomic classification of uncultured bacteria. Because of the complexity of the bacterial community in the gut, the method used for its study depends on the question being asked. An overview of different molecular “omics” approaches to study the gut microbiota is shown in Fig. 1. To examine overall changes in the microbial community a number of fingerprinting approaches have been developed. These techniques are also collectively referred to as “microbiomics,” which attempts to study the identities of members of complex microbial communities. Most microbiomics approaches for bacterial identification are based on sequences from the variable regions of the 16s rRNA gene. The most frequently used techniques include terminal restriction fragment length polymorphism (T-RFLP), denaturing gradient gel electrophoresis, and cloning and sequencing (Fig. 1). These methods take advantage of the conserved regions of the 16s rRNA gene as they utilize PCR primer binding sites that are present in all bacteria. Denaturing gradient gel electrophoresis separates PCR amplified DNA molecules based on their ability to migrate through a polyacrylamide gel that contains a denaturant. The extent of migration is dependent on the DNA sequence of amplicons produced via PCR. As the strands become denatured and separate, they become less mobile in the gel. One end of each amplicon is held together with a GC clamp to prevent complete strand separation. The resulting output is a series of bands on a gel that can subsequently be excised and sequenced to determine the identities of the populations corresponding to individual bands. In T-RFLP, the variability in lengths of restriction-dgested PCR products is visualized by capillary-based DNA sequencing. One primer, normally at the 5’ end of the amplicon, has a fluorescent label to enable the terminal restriction fragment to be detected by the automated sequencer. Because of its relatively high throughput and high reproducibility, this technique is often the method of choice for comparison of multiple samples over time or

3.

eta

THE GUT MICROBIOTA: ECOLOGY AND FUNCTION

41

its

FIGURE 1 Overview of “omics” approaches to study the gut microbiota.

different treatments (Dicksved et al., 2007). Although both methods give insight into overall changes in the microbial population, their taxonomic resolution is relatively poor. Clone libraries of PCR amplified 16s rRNA genes have greater taxonomic resolution; however, they are limited by the sequencing costs, although some large studes have achieved as many as 10,000to 20,000 sequences (Eckburg et al., 2005). The recently developed 454 pyrosequencing approach, using multiple sequence tags or barcodes, allows for relatively deep sequencing of multiple samples in parallel at fairly high taxonomic resolution, although not at a level equal to that of full-length 16s rRNA sequencing (Sogin et al., 2006; Anderson et al., 2008). The power of this method was recently demonstrated in a sequencing study of human fecal samples that resulted in 440,000 reads with an average read length of 59 bp (Dethlefien et al., 2008). This study revealed

5,600 operational taxonomic units (OTUs) at 99% identity from only 3 individuals. Furthermore, advances in pyrosequencing enable longer read lengths-approximately 250 bp on the 454 GS-FLX platform and approximately 450 bp on the 454 GS-Titanium platform. The Roche 454 pyrosequencing technology is continuing to improve with longer read lengths, thus improving taxonomic resolution. Other promising second generation sequencing approaches that have yet to be applied for detailed studies of the human gut include the Illumina Solexa platform and the SOLiD platform by Applied Biosystems. Another recent fingerprinting approach to assess the gut microbial community composition uses a phylogenetic microarray. The Phylochip is a microarray that contains oligonucleotide probes designed based on all environmental sample sequences submitted to the RDP website. These sequences have passed a quality check and are deposited

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in the Greengenes database, http://greengenes .lbl.gov/cgi-bin/nph-index.cgi(Yergeau et al., 2009). The HITChip is another phylogenetic microarray that specifically contains probes for sequences isolated from the human intestinal tract (Rajilic-Stojanovic et al., 2009). Although much improved over previous culture-based methods, all of the microbiomics approaches introduce biases because of differences in efficiencies of cell lysis and DNA extraction, primer design and binding, PCR, sequencing (e.g., homopolymers produced during 454 pyrosequencing), and/or cloning. For the quantification of specific bacterial groups, or species, fluorescence in situ hybridization (FISH) and quantitative real-time PCR have most commonly been applied. Both approaches utilize the variabhty within the 16s rRNA gene for the development of specific probes or primers, respectively,that only amplifj or hybridize to the regons of the 16s rRNA gene that distinguish a specified target group or species. Primers and probes for both methods can be generated using software such as the ARB Project (http://www.arb-home.de). FISH can also be used to visuahze the localization of bacteria within the gut, giving insight into which bacteria are in close proximity with the mucosal surface (Johansson et al., 2008). Another advantage of FISH over other molecular approaches is that it does not depend on DNA purification or amplification, thus avoidmg many of the biases mentioned previously. FISH, however, does have its own disadvantages. For example, because only a few probes can be used simultaneously, it is limited in its ability to target many different bacteria in the same sample. Also, FISH is a relatively technically demanding method and there can be problems in &stinguishing fluorescent cells over background fluorescence in samples. Quantitative PCR overcomes many of the &sadvantages of using FISH and can be used for accurate quantification of a specific 16s rRNA target. Group- or species-specific primers can be designed to specifically quantiG certain members of the microbiota. For

example, Willing et al. (2009a) used specific primers to quantify populations of Escherichia coli and Fuecalibacterium sp. in fecal samples and intestinal biopsies. Although the methods described here provide a means to determine the composition of members of the gut microbiota, there is also interest in the functional characteristics of the community. Although a certain level of predction can be achieved by extrapolating to physiological properties of related cultured species, this approach is challenging and, in many cases, inaccurate. Metagenomic analysis of the microbiota gives sequence information from the collective genomes of all members of the microbiota (microbiome). The first metagenomic study of the human gut resulted in 78 mdlion base pairs of DNA sequences from two American indwiduals (Gill et al., 2006). This study cataloged the combined gene complement of the microbiome, including functional genes. However, it is important to keep in mind that some of the DNA analyzed may have been derived from inactive, dormant, or even dead members of the community. Also, not all genes are expressed at any given time and condition. Therefore, metagenomic approaches only provide information about functions that have the potential to be expressed. Other “ o ~ ~ capproaches, s” such as metatranscriptomics, metaproteomics, and metabolomics, enable dfferent levels of expression to be studied and a more direct assessment of which processes are active and functioning in the gut under a given condition. Some results using these different approaches wdl be discussed later in the context of relevant subsections.

Identification of Microorganisms in the Gut Microbiomic studies, primarily those targeting bacteria, have been used to study the composition of the gut microbiota in humans and some animals. A recent study examined the microbial composition of 60 divergent animal species found in zoos and in the wild (Ley et al., 2008). Of more than a 100 known bacterial phyla

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TABLE 1 Representation of classified bacterial phyla from 60 mammalian species' Phylum

% of classified sequences

Fimicutes Bacteroidetes

65.7 16.3

Proteobacteria Actinobacteria Vemcomicrobia Fusobacteria Spirochaetes

8.8 4.7 2.2 0.67 0.46

DSSl Fibrobacteres TM7 Cyanobacteria Planctomycetes

0.35 0.13 0.13 0.1 0.08

Dderribacteres Lentisphaerae Chloroflexi SR1 Deinococcus- Themus

0.05 0.04 0.005 0.005 0.005

"Adaptedfrom Ley et al., 2008.

(divisions), representatives of only 17 were detected in this diverse set of fecal samples (Table 1). This relatively restricted representation of bacterial phyla suggests that the gut microbiota coevolved with their mammalian hosts. The Fimicutes phylum was the only phylum detected in all animal species and was also the most abundant, representing nearly 67% of all classified sequences. Other highly represented phyla included the Bactevoidetes, Pvoteobacteria, Actinobacteria, Vewucomicvobia, and Fusobactevia. The remaining 11 phyla represented less than 0.5% of the classified sequences. The functions of most of these less prevalent species have not been specifically identified and it is not known if the rare members of the microbiota have important roles. Current studies suggest, however, that the dominant phyla are best adapted to the gut environment. Although, in the study by Ley et al. (2008), there were many similarities between animal species and within species-

43

each individual studied also harbored at least one unique OTU (at 96% similarity) that was not found in any other sample. O n average, 62% of the OTUs from a given animal species were unique to that species. There have been additional studies of the bacterial composition of the gut in other animals, including chickens (Dumonceaux et al., 2006), mice (Yap et al., 2008), pigs (Hill et al., 2005), and cows (Hernandez et al., 2008). These studies have revealed that each animal species has a gut microbiota that is unique and representative of that species. This is exemplified by a recent study of the horse microbiota that found that less than 33% of 16s rRN A clones had matches in databases at the species level, with a species being defined at 98% 16s rRNA sequence homology (Willing et al., 2009b). Because it is not feasible to discuss the microbiota of all mammals in the scope of this chapter, we will focus on the GI microbiota of humans, including specific examples from other species that have been relatively well studied, such as the mouse and pig. The human fecal microbiota is dominated by two bacterial phyla: the Fivmicutes and Bacteroidetes (Eckburg et al., 2005). Approximately 800 to 1,000 different bacterial species and more than 7,000 strains inhabit the human GI tract, based on estimates of 16s r RN A libraries from fecal samples (Backhed et al., 2005). The application of second generation sequencing approaches, such as 454 pyrosequencing of 16s rRN A genes, has provided a greater depth of view of the composition of the gut microbiota (Andersson et al., 2008). One pyrosequencing study revealed as many as 3,300 unique OTUs in a single individual (Dethlefien et al., 2008). As fecal samples from more individuals are being sequenced, it is becoming apparent that each individual has a relatively unique gut microbiota. This, in turn, presents a major challenge when trying to design studies and trying to apply general ecological rules to better understand the functional roles of the gut microbiota.


Core Microbiome Given the great variability in gut microbiota between individuals, a question has been raised whether there is a “core microbiome,” or a set of microorganisms that is found in all individuals. One study of 17 subjects that looked to define the phylogenetic core found that there were no OTUs, defined at 2% dssirmlarity, that could be found in all subjects sampled, suggesting that the concept of a core microbiome is likely invalid (Tap et al., 2009). However, other sequencing studies suggest that a core microbiome exists at the gene level, including important components of various metabolic functions (Tumbaugh et al., 2009), and a “functional core” can be potentially defined. Although the study by Turnbaugh et al. (2009) revealed substantial variation in the relative amounts of the key phyla-Firmicutes, Bacteroidetes, Actinobactevia, and Proteobactevia-there was very little variation in function, based on the categories of gene function, in the same samples analyzed (Fig. 2). These results suggest that, although the composition of the gut microbiota is variable, there likely remains functional redundancy in the community, which may be a protective mechanism to perturbation.

a

Bacterial ohylwn

The Archaea Members of the Archaea, particularly methanogens, have been found in the intestinal tract of many vertebrate species (Lin and Miller, 1998). In humans, methanogens can represent as much as 10% of the anaerobic microorganisms in the adult colon and are dominated by a single species, Methanobrevibacter smithii (Eckburg et al., 2005). The genome sequence of this microbe has revealed that it is specifically adapted to the human gut environment (Samuel et al., 2007). Other minor representatives of Archaea in the gut include Methanosphaera stadtmanae and the Crenarchaeotes (Rieu-Lesme et al., 2005). These methanogens play a vital role in the gut by removal of H2 via anaerobic metabolic processes. However, not all humans produce methane gas in their intestines but, rather, utilize other Hz disposal routes, as discussed later (Walker, 2007). The gut environment from other hostsincluding the horse, pig, cow, rat and goose-have been shown to contain Archaea in the genus Methanobrevibacter that have high sequence similarity to the 16s r R N A gene from M. smithii. This suggests that methanogens inhabiting the gut environment share a

b

COG c a t m e s

FIGURE 2 Functional redundancy of the gut microbiota suggested by large variations in community composition between individuals: (a) compared to community functions or (b) categories of gene function. Reprinted from Turnbaugh et al. (2009), with permission.

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THE GUT MICROBIOTA ECOLOGY AND FUNCTION H 45

common descent. However, sheep isolates from one study were sufficiently divergent to suggest a separate line of descent within the genus Methanobrevibacter (Lin and Miller, 1998). Methanogens are of particular importance in ruminant microbiology. The production of methane by gut fermentation reactions results in reduced metabolic efficiency, with up to 13% of the energy from the diet being lost to methane production, as well as an environmental concern because of the role of methane as a greenhouse gas (Ohene-Adjei et al., 2008). Efforts are currently underway to limit the abundance and activity of methanogenic Archaea to reduce the impact of beef production on global warming (Hook et al., 2009).

Intestinal Fungi Recent studies have begun to reveal the identities of fungi in the intestinal tract. One study identified a wide variety of fungi in mouse feces, but it was not determined whether they were colonizing commensals or simply transient organisms attached to dietary particles (Scupham et al., 2006). Repeated sampling of the human gut indicated that some fungi were temporally stable and actually colonized the intestine (Scanlan and Marchesi, 2008). Similar to bacteria, there is a great disparity between what can be cultivated and what can be detected using DNA-based methods. The cultivable fraction of the fungal microbiota in human feces is dominated by Candida spp. strains (Scanlan and Marchesi, 2008), whereas culture-independent analysis revealed that members of the genera Gloeotinia/Paecilomyces and Galactomyces were the most abundant (Scanlan and Marchesi, 2008). However, the diversity and abundance of fungi in the GI tract is quite low relative to that seen with the bacteria. Although relatively unexplored, specific fungal populations may play important roles in the gut. This is suggested by a recent study that found an increased abundance of host antibodies specific to Saccharomyces cevevisiue in patients with Crohn's dsease (Dossopoulos et al., 2007).

LOCALIZATION OF GUT MICROBIOTA Microbes survive within many microhabitats in the digestive tract, including the intestinal lumen, the mucus layer lining, and other varying compartments of the GI tract (Maclue et al., 1999). The bacterial population ofthe GI tract increases in density and complexity from proximal to distal regions, with the exception of the mouth, and changes composition with location and time (Fig. 3). The stomach and duodenum have a relatively low microbial density, with lo3 to lo5 CFUs per gram of contents, and a population dominated by the Lactobacillus, Streptococcus, and Enterococcus genera. This community is limited in density and diversity because of an acidic environment and the presence of bile secretions. Because the rate of digesta passage is rapid through this region, the ability to attach to and colonize the mucosal layer appears to be important in the proximal small intestine (SI). The rate of digesta passage slows in the dstal SI and bacterial counts increase to lo8 to lo9 CFU per gram of contents accordingly. Phylogenetic representation in the dstal SI also increases and includes those genera indicated previously, plus members of genera within the Clostridiales and Bacteviodetes families (Hill et al., 2004; Richards et al., 2005). Complexity of the microbial community is greatest in the large intestine, where the density increases to lo1' to 10l2 CFU per gram and the community is dominated by gram-positive genera, including Clostridium, Bacillus, Ruminococcus, and Fusobacterium (Hill et al., 2002; Leser et al., 2002). Variation within bacterial species among locations in the GI tract also substantially contributes to overall gut microbial diversity and function (Dixit et al., 2004). For example, fermentation of carbohydrates and amino acids are functions that become more prominent in the distal locations of the gut than in proximal regions (Cummings and Macfarlane, 1991). The mucous layer covering the epithelial layer is made up of two dstinct layers, referred to as loosely and firmly bound mucous layers. In situ hybridization studies have revealed that


-Bacteria

Loosely Adherent Firmly Adherent

ipithelial Cells

ESD09045

FIGURE 3 Bacterial colonization of the GI tract. Bacterial numbers increase in abundance from proximal to distal regions. Bacteria reside in close proximity to the intestinal epithelium. A firmly adherent layer of mucous keeps bacteria at a safe distance so as to prevent continual mucosal stimulation and inflammation and a loosely adherent layer provides a habitat for abundant microbial colonization.

bacteria do not reside in the firmly bound mucous layer (Johansson et al., 2008). In contrast, the loosely bound mucosal layer contains abundant bacteria (Fig. 3). Attachment to the mucosa has been shown to be dependent on a variety of bacterial cell surface ligands (Sillanpaa et al., 2000; Roos and Jonsson, 2002), as well as specific receptors such as sugar chains of glycolipids or glycoproteins produced by host cells. The firmly bound mucous layer is extremely important for animal health as evidenced by stuhes of mice lackmg the MUC2 gene, the major mucin of the colon mucus. These mice have bacteria in direct contact with the epithelial layer that results in inflammation and cancer development (Johansson et al.,

2008). Although potentially not in continuous direct contact with the intestinal epithelium, substantial bacterial populations live in close proximity in the mucin layer and likely have intermittent direct contact. The abundance of bacteria in close proximity to the intestinal epithelium has been demonstrated by measuring microbial populations present in homogenized whole tissues after the intestinal contents have been eradcated by washing (Frece et al., 2005). The discrepancy observed between attachment in vitro and in vivo may be a consequence of the static nature of in vitro cell lines and the fact that tissue cultures lack other important host factors that keep the intestinal microbiota at bay. Conversely, the recent observation that

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THE GUT MICROBIOTA: ECOLOGY AND FUNCTION W 47

bacteria are not dn-ectly associated with the intestinal epithelium may be a consequence of tissue collection conditions and assay sensitivity. Although the extent to which commensal bacteria come in l r e c t contact with the intestinal epithelium is not clear, these bacteria clearly have major effects on the host. SUCCESSIONAL CHANGES At birth, the neonate leaves the sterile environment of the uterus and is immediately introduced to a population of bacteria in the vaginal tract and, subsequently, in the surrounlng environment (Tannock et al., 1990a, 1990b). Analogous to the gradual establishment of pioneer microorganisms and species succession after a forest fire, there are early colonizers that quickly colonize the vacant system. These early colonizers are facultative anaerobes that create a reduced environment that is more favorable to growth by anaerobic species (Stark and Lee, 1982) and are eventually supplanted by their successors. The ecological succession of the commensal microbiota is relatively poorly understood in most species. The distal colon of the pig is colonized with lo9 to 10" bacteria per gram of digesta within 12 h of birth (Swords et al., 1993). Clostridium spp. become dominant after 5 days of age, begin to decline at weaning, and are eventually replaced by Bacteroides spp., the dominant adult colonic microbiota in the pig (Swords et al., 1993). The diversity and activity of given genera also change through time, particularly at weaning (Janczyk et al., 2007). It is not clear whether microbial succession patterns in the proximal SI mirror those observed in the colon and feces. However, molecular based profiling studies in the pig have demonstrated marked differences in microbial composition between proximal and distal intestinal locations (Konstantinov et al., 2004; Hill et al., 2005; Richards et al., 2005). There have also been some studies of succession in humans. However, they are limited to the fecal microbiota for obvious reasons. As discussed in the section about &et, some

dramatic differences have been observed in the gut microbiota of infants that are fed formula compared to infants that are breast-fed (Harmsen et al., 2000). Other factors affecting succession patterns of the microbiota include method of delivery (vaginal vs. Caesarean section), postnatal hygiene, environment, and antibiotic use (Gronlund et al., 1999; Schwiertz et al., 2003; Lofinark et al., 2006). IMPACT OF ANTIBIOTICS Although there is great variation in the composition of the gut microbiota between individuals, within a given individual the composition is relatively stable and resilient over time. However, antibiotic administration can cause major disruptions in the gut microbiota. Of course, the reason that antibiotics are administered in the first place is to reduce levels of pathogenic populations that are detrimental to the host. Different specific bacterial populations may be targeted depending on the antibiotic that is administered. However, antibiotics also have a positive or negative impact on other nontarget members of the commensal microbiota. Microbes that are resistant to the antibiotic used have a selective advantage and may increase in numbers relative to susceptible strains. Alternatively, nontarget susceptible populations may be reduced in numbers upon antibiotic treatment. In general, the diversity of the gut microbiota is reduced following administration of antibiotics (Jernberg et al., 2007), but this reduction is short lived and usually lasts only a matter of weeks or days. Relatively soon after antibiotic treatment is hscontinued the commensal gut microbiota present before disruption return, for the most part, relatively quickly (Dethlefien et al., 2008). However, some members of the bacterial community remain disrupted for long periods of time, even several years after antibiotic treatment (Jernberg et al., 2007). This has been demonstrated for bacterial populations that are targeted by clindamycin (Lofinark et al., 2006; Jernberg et al., 2007). For example, a conventional

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WILLING AND JANSSON

7-day administration of clindamycin was found to alter the ecology of the gut microbiota up to 2 years posttreatment (Jernberg et al., 2005; Lofmark et al., 2006; Jernberg et al., 2007). Our recent 454 pyrosequencing studies have also shown that the gut microbiota is dramatically perturbed for up to 4 years posttreatment after taking the common triple therapy for Helicobacter pylori (clarithromycin, metronidazole, and omeprazole) Uakobsson et al., 2010). In addition to dsturbances in the ecology of the microbiota, antibiotics can also select for persistence of antibiotic resistance genes in the host. Studies have shown that several antibiotic resistance genes are stably maintained in the GI tract once they are acquired (Shoemaker et al., 2001). These genes are often carried on conjugative elements and, under selection pressure, they have the potential to be transferred between dfferent species within the densely populated gut microbial community. Jernberg et al. (2007) used real-time PCR to quanti@ dfferent resistance genes in DNA extracted from feces. They reported that clindamycin exposure led to significant increases of erm(F), erm(G), and erm(B) in feces. High levels of these resistance genes could still be detected 2 years after the antibiotic treatment had ceased. This is disconcerting considering the growing problem with increasing prevalence of antibiotic-resistant strains that are not susceptible to current arsenals of antibiotics. ADAPTATIONS OF THE COMMENSAL MICROBIOTA T O THE GUT ENVIRONMENT Increasing evidence suggests that the human gut, no matter the &et or region of the world, is consistently colonized by the same bacterial phyla and only a single &vision of Archaea. This indicates that only certain divisional members have evolved close associations with the human gut (Eckburg et al., 2005). What are the mechanisms that govern membership of the community as well as relative abundances of the members? Microbes colonizing the GI tract are not simply there as a matter of

chance. Based on studies of a wide variety of animal species, it is apparent that the microbiota has truly coevolved along with the mammalian host and is highly specialized to thrive in the gut environment. The complement of genes represented in the gut microbiome is estimated to be 100 times the number of genes found in the host (Backhed et al., 2005) and is enriched in genes for glycan, amino acid, and xenobiotic metabolism; methanogenesis; and synthesis of vitamins and isoprenoids (Gdl et al., 2006). The gut microbiome also has a diverse repertoire of environmental sensors and cognate regulators (Xu et al., 2007). Interestingly, motility and chemotaxis are two characteristics that seem to be underrepresented among gut microbiota (Turnbaugh et al., 2009; Verberkmoes et al., 2009). Recently, the proteins in the feces of two healthy individuals were determined using a community proteomics, or “shotgun metaproteomics” approach (Verberkmoes et al., 2009). Thousands of microbial and host-produced proteins were identified using this technique. A group of approximately 700 equally abundant proteins in the two individuals were suggested to represent the core proteome. This core contained proteins for housekeeping functions, translation, energy production, protein turnover, and carbohydrate metabolism. The proteins present in the core were also representative of common gut bacteria, such as Bacteroides, Bijidobacterium, and Clostridium. Another key feature of the microbiota that exemplifies coevolution is communication with the host. Some bacteria are able to shape the host to fit their ecological requirements. For example, some bacteria in the gut produce specific effector molecules that function to stimulate targeted host functions that benefit the bacteria. Although relatively little is understood about effector molecules secreted by commensal bacteria, the use of effector molecules by pathogenic bacteria has been intensely studied and serves a variety of functions in pathogen establishment. For example, enteropathogenic E. coli inject effector proteins into enterocytes causing actin rearrangement

3. THE GUT MICROBIOTA ECOLOGY AND FUNCTION

in the host that leads to the formation of a pedestal-like structure thereby allowing intimate microbial-host contact (Dean and Kenny, 2009). Some commensal bacteria secrete effector molecules that stimulate the host to provide a nutrient source, as well as inhibit the inflammatory response (Hooper and Gordon, 2001a, 2001b; Sougioultzis et al., 2003). The gut environment is also the site of specific intimate metabolic interactions between the host and the microbiota. For example, some members of the gut microbiota have evolved to take advantage of specific host-derived substrates, such as fucose, which is produced by epithelial cells in the gut lining. The best known example of bacterial signaling the host to create a beneficial nutrient environment is demonstrated by Bacteroides thetaiotaomicron. This bacterium signals the host to manufacture fucosylated glycans in place of sialylated glycans because it uses fucose as a main energy source. Segmented filamentous bacteria and a normal microbiota are also able to induce the expression of this glycolipid (Umesaki et al., 1995). The mechanism by which fucosylated glycan production is up-regulated is not yet clear, although the signaling molecule produced by B. thetaiotaomicron has been found. It has been established that transcription of fucosyltransferase genes are activated by this molecule (Hooper and Gordon, 2001b). FACTORS AFFECTING THE GUT MICROBIOME Major factors affecting microbial number and composition along the length of the intestine appear to be age, l e t , and host genotype (van der Wielen et al., 2000; Apajalahti et al., 2001; Hill et al., 2005). Comparisons between the zoo animals described previously revealed that diet was the strongest determining factor in shaping microbial composition, where animals were grouped based on whether they were carnivores, herbivores, or omnivores. Gut microbial diversity is generally highest in herbivores, intermediate in omnivores, and lowest in carnivores (Ley et al., 2008). There

49

were, however, intriguing similarities between animal species that indxate the importance of lineage. For example, pandas and bears had relatively similar microbiotas, although their diets are extremely different (Ley et al., 2008). Some of these factors will be discussed in more detail later.

Effect of Diet The nutrient requirements for bacteria are highly variable. Therefore, it is not surprising that the composition of the diet is very important in shaping microbial composition. This is particularly true for components of the det, such as fiber, that escape digestion and absorption by the host in the proximal part of the intestine. The composition of the gut microbiota can change dramatically in response to changes in &et; however, these changes do not always happen quickly. It can take several weeks for activators of microbial degradation enzymes, such as xylanase and cellulose, to adjust to a new diet (Castdlo et al., 2006). In the case of omnivores and herbivores, the introduction of plant carbohydrates favors the growth of species that are adapted to the fermentation of detary fiber. One of the most dramatic changes in microbial composition occurs during the weaning process, where there is a switch from a highly digestible simple liquid diet (mdk) to a solid &et that is much more complex and less digestible (Jensen, 1998). Components of human rmlk contain nonabsorbable oligosaccharides, nucleotides, and gangliosides, which affect colonization by bifidobactena (Kunz and Rudloff, 1993; Balmer et al., 1994; Rueda et al., 1998). It has been shown that bifidobacteria are dominant in feces of breast-fed infants, compared to a more diverse fecal microbiota dominated by Bacteroides spp. in formula-fed infants (Stark and Lee, 1982; Harmsen et al., 2000). Several studies have examined the impact of obesity on the gut microbiota. The overall diversity of the microbiota is reduced in obese individuals that have an increased food intake (Turnbaugh et al., 2009). There is an intriguing relationship between obesity and composition of the microbiota. By analyzing the microbiota


of obese mice ob/ob, lean ob/+, and wild-type siblings, and their ob/+ mothers, Ley et al. (2005) found that ob/ob mice have a reduced abundance of Bacteroidetes and an increase in Firmicutes. The efficiency of fermentation may also be improved in ob/ob mice because of an increased abundance of methanogenic Archaea, which were hscussed previously, are responsible for Hz removal (Turnbaugh et al., 2006). Some specific lifestyle factors can also impact the composition and diversity of the gut microbiota. For example, an anthroposophic lifestyle that entails the consumption of organically produced and fermentedfoods and the restricted use of antibiotics and vaccinations were correlated with a higher bacterial diversity as compared to a conventional lifestyle or to being raised on a farm (Dicksved et al., 2007). It is, however, difficult to define a causal relationship between diet and microbial composition because many lifestyle factors that coincide with det also play a role in influencing the microbial composition in the gut. Similar observationshave been made in studes examining the effect of diet on cultivable bacteria (Mai, 2004).

The Host Interaction with the Gut Microbiome There remains a hgh variabihty in the composition of the gut microbiota between indviduals, even with s i d a r diets and lifestyles. An individual’s microbiota is, however, surprisingly stable. The host itself is thus one of the contributing factors in the ecology of the fecal microbiome. It not only creates the abiotic environment, including temperature, pH, and moisture, it also provides nutrients that are utilized by the members of the microbiota and is active in regulating the microbial abundance and composition. Paneth cells are known to produce a-defensins that protect the proliferative stem cell compartment of the intestinal crypt from colonization (Ayabe et al., 2000). The expression and secretion of RegIIIy, a bacteriocidal protein effective against only gram-positive bacteria, is induced by host recognition of gram-negative bacteria (Cash et al., 2006). A proteomic study of feces

revealed that the third most abundant group of human proteins, after dgestive enzymes and structural cell adhesion and cell-cell interaction proteins, were human innate immunity proteins. These included antimicrobialpeptides, intellectin, resistin, and others (Verberkmoeset al., 2009). These data therefore provided a ghmpse into the protein arsenal that is involved in hostmicrobe interactions in the gut. An interesting example of the importance of host physiology in shaping the composition of the microbiota was shown in reciprocal transplantations of gut microbiota between mice and zebrafish (Rawls et al., 2006). These authors found that when the microbiota from a mouse was introduced to a germ-free zebrafish, the relative abundance of community members shifted so as to resemble the normal microbial composition of the zebrafish. Recently, it was shown that the composition of a recipient’s fecal microbiota could be altered by fecal transplantation from a donor (Khoruts et al., 2010). This particular example involved the case of a woman who suffered from severe diarrhea caused by Clostridium d@cile-associated disease. The patient had undergone a series of antibiotic treatments, all of which were unsuccessful. As a last resort the attending physician prescribed a fecal transplantation from a healthy donor. Interestingly, not only did this cure the disease, but the patient adopted the fecal microbial composition of the donor, at least for several weeks. These results indicate that the host programmed microbiome may be altered in some cases, at least temporarily, and suggest a therapy for severe cases of C. dficile-associated disease. Studies comparing the microbial composition of twins, related individuals,and unrelated individuals indicate an important role of host genotype in shaping microbial composition. Twins living in different environments share a more similar microbiota than do twins and their spouses who share the same environment and d e t (Zoetendal et al., 2001). A more recent study that examined the gut microbiota of obese and lean twins and their mothers, using

3.


clone libraries and pyrosequencing, revealed that related individuals had more bacterial species in common and a more similar bacterial community structure than unrelated individuals, regardless of obesity status (Turnbaugh et al., 2009). We have performed extensive studies on a Swedish twin cohort that include healthy individuals and individuals with different inflammatory bowel diseases (see more detail later). Our initial T-RFLP data revealed a high degree of similarity in the gut microbiota of identical twins, even when they lived apart for decades (Dicksved et al., 2008). More recently, 454 pyrosequencing data revealed that, although the healthy twins shared some degree of similarity at an OTU/species level, the similarity was much more apparent at the genus level (Willing et al., unpublished data).

Impact of Disease There is increasing evidence linkmg the composition of the gut microbiota to the health of the host, both positively and negatively. Some examples of diseases where the gut microbiota is implicated to have a role, although still largely undefined, include inflammatory bowel diseases (IBDs) such as irritable bowel syndrome, Crohn’s disease (CD), ulcerative colitis, and colon cancer. In this chapter we primarily discuss CD because of the large number of recent reports that have focused on the correlation of the gut microbiota to this particular disease. Different molecular fingerprinting techniques, including T-RFLP (Dicksved et al., 2008; Willing et al., 2009a), TGGE (Seksik et al., 2005), qPCR (Willing et al., 2009a), and 454 pyrosequencing (Willing et al., unpublished data), have revealed that the gut microbiota is different in individuals with CD compared to healthy individuals. This altered gut microbiota has been referred to as a state of “dysbiosis” (Seksik et al., 2005; Tamboli et al., 2004). The theory of dysbiosis is that there is a breakdown in the balance between protective versus harmful intestinal bacteria that is either a cause, or a consequence, of disease (Tamboli et al., 2004).

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Recent findings have honed in on specific members of the gut microbiota that are more or less prevalent in individuals with CD. For example, a T-RFLP study of identical twins that were discordant for CD found that there were differences in relative amounts of some Bacteroides species in the twins with CD, compared to healthy twins (Dicksved et al., 2008). Individuals with CD in the ileum also had a lower bacterial diversity in the mucosal lining of the intestine, compared to those with disease localized in the colon (Willing et al., 2009a). Several studies have implicated E. coli (Darfeuille-Michaud et al., 2004; Baumgart et al., 2007; Willing et al., 2009a) or Mycobacterium paratuberculosis (Naser et al., 2004) as potential virulent species that may have a specific role in the pathogenesis of CD, but these microbes remain to be confirmed. One striking example of dysbiosis is the dramatic reduction in abundance of some members of the Clostridium leptum group (Cluster IV), such as Faecalibacterium pruusnitzii, in fecal samples and biopsies of individuals with CD (Swidsinski et al., 2008; Willing et al., 2009a). F. prausnitzii is considered to be a beneficial member of the commensal microbiota because of its ability to produce butyrate and provide therapeutic properties in a mouse IBD model (Sokol et al., 2008). One hypothesis is that a reduction in F. prausnitzii would result in a reduced amount of butyrate that could trigger gut inflammation, or even induce the expression of antimicrobial peptides, thus allowing pathogenic E. coli to proliferate. Recent 454 pyrosequencing data (Willing et al., unpublished data) have revealed that the situation is even more complex and that several adhtional bacteria differ in individuals with different IBD phenotypes. These differentiating populations might serve as diagnostic targets of IBD in the future and obviate the current need for invasive endoscopic tests.

Methods to Alter the Microbiota In animal production, the search for a means to manipulate the microbiota to promote growth and prevent disease has been a major


focus of animal nutritionists. Antibiotics have been used to regulate the GI microbiota of livestock for the last half of the 20th century. The development of antibiotic resistance and “superbugs,” however, has resulted in a ban of in-feed antibiotics in the European Union. The growing awareness of the importance of the microbiota in health and disease has also resulted in an increased interest in manipulating the microbiota in human nutrition. Alternative strategies to regulate the GI microbiota that have become commonplace include the use of prebiotics and probiotics as supplements. Prebiotics are dietary additives that are directed at promoting health through their effects on the microbiota (Gibson and Roberfroid, 1995), whereas a probiotic is defined as a live microbial feed supplement that beneficially affects the host by improving its intestinal balance (Fuller, 1989). The majority of probiotic organisms used are lactic acid bacteria, which include a variety of lactobacilli, enterococci, and bifidobacteria. Other probiotics include bacteria within the genus Bacillus and yeasts such as Saccharomyces cerevisiae. The probiotic mode of action is generally accredited to competitive exclusion of pathogens and strengthening of the mucosal barrier (Servin, 2004; Saxelin et al., 2005). In contrast, prebiotics are largely concentrated on the use of indigestible carbohydrates that can be utilized by bacteria in the large bowel. The effect of these carbohydrates on the microbiota is dependent on how available they are to bacterial fermentation, which is dependent on water solubility and the degree of polymerization and lignification. The prebiotics inulin and fmctooligosaccharides have been shown to stimulate the growth of beneficial gut bacteria, such as bifidobacteria and lactobacilli (Van Loo, 2004). MICROBIAL METABOLISM IN THE GUT The different microbes residing in the gut carry out specific metabolic processes that can directly or indirectly impact the host. Metabolites produced by gut microbiota can be

transferred across the epithelial barrier and reach systemic sites of the host. This was evident, for example, when comparing the plasma metabolic profiles of germ-free and conventional mice. Numerous circulating metabolites were only present when the mice were colonized by microorganisms and there were significant differences in the abundance of several metabolites in germ-free and conventional mice (Claus et al., 2008; Wikoff et al., 2009). Claus et al. (2008) found that the microbiome has an impact on the metabolism of bile acids and kidney homeostasis. These data confirm that microbes modulate the expression of metabolites both locally in the gut and in other body sites. One of the goals of current research is to identie metabolites, including those of microbial origin, that are diagnostic biomarkers of health or disease ofthe host. This goal may soon be realized because the availability of new and advanced technologies that produce detailed information about the metabolic composition in human samples, including feces, urine, and blood. These technologies include gas chromatography-mass spectrometry (GC-MS), ‘H nuclear magnetic resonance (NMR) spectroscopy, and ion cyclotron resonance Fourier transform mass spectrometry (ICR-FT/MS). These methods have recently been used to determine hundreds to thousands of metabolites in human samples, collectively referred to as the “metabolome” and the field of “metabolomics” or “metabonomics” (Nicholson et al., 2005; Jansson et al., 2009).

Beneficial Metabolites In return for a hospitable environment and provision of nutrients, the microbiota provide the host with major services. The cumulative microbiota have a vast capacity to mediate a diverse set of beneficial roles for the host (Marchesi and Shanahan, 2007). These roles include synthesis of vitamins, degradation of xenobiotics, metabolism of bile and host hormones (Hooper et al., 1998), immune development, and the competitive exclusion of pathogens.

3. THE GUT MICROBIOTA: ECOLOGY AND FUNCTION W 53

One of the most important roles that the microbiota provides is the provision of energy for the host’s colonic epithelium. The energy provided is primarily in the form of short chain fatty acids (SCFAs) that are produced by bacterial fermentation of undigested carbohydrates and fiber (Karah et al., 2006). Accordingly, SCFA production is highly affected by diet. Dependmg on the animal species and the nature of the &et, SCFA can make up as much as 70% of the energy that the host derives from the &et and, of these, butyrate is the preferred energy source for colonocytes (Bergman, 1990). Interestingly, different types of SCFAs are produced in the gut of bottleversus breast-fed infants. The SCFAs produced by the gut microbiota in formula-fed babies have less lactic acid and more propionic acid than in breast-fed babies (Edwards et al., 1994). Not all populations ofbacteria are equal in their ability to liberate energy from the diet. A reduction in Bacteroidetes and an increase in Firmicutes, for example, has been shown to increase SCFA production resulting in increased energy for the host and, in some cases, contribute to obesity (Turnbaugh et al., 2006). The fermentation of undgested dietary components results in the production of an extensive combination of metabolites, many of which remain to be characterized. Studes comparing the blood metabolites of gem-free to conventional animals indicate that the metabolites produced by bacteria can have systemic effects (Wikoff et al., 2009). M o w n g the composition of the microbiota using nutritional changes can modulate host lipid, carbohydrate, and amino acid metabolism (Martin et al., 2009). These effects may improve health because, for example, some bacteria cause reduced plasma lipoproteins (Martin et al., 2009). The composition of the microbiota has been shown to correlate with components of the fecal metabolome as well as metabolites found in the urine (Marchesi et al., 2007; Jansson et al., 2009). The variation in abundance of Faeculibacteriumprausnitxii has been associated with the modulation of at least eight urinary metabolites of diverse structure (Li et al., 2008).

Biomarkers of Disease Shifts in the gut microbiota and their metabolites have been correlated to several diseases, including diabetes, obesity, and IBD. Because fecal extracts of patients with IBD have reduced levels ofbutyrate, acetate, methylamine, and trimethylamine, changes in metabolite profiles associated with changes in microbial populations likely play an important role in intestinal health (Marchesi et al., 2007). Using ICR-FT/MS, we recently found that the situation is even more complex-thousands of metabolites differentiated individuals with different CD phenotypes and healthy individuals (Jansson et al., 2009). Pathways with differentiating metabolites include those for metabolism of amino acids, fatty acids, bile acids, and arachidonic acid. The mass spectrum corresponding to glycocholic acid in the bile acid biosynthesis pathway was more prevalent in individuals with CD than in healthy individuals. Increased bile in the feces may be one of the causes leading to symptoms associated with inflammation in CD. This example illustrates the important contribution of the metabolic processes carried out by the gut microbiota on human health. In addition to the mass spectra that could be identified using this large metabolic survey, there were several hundred that could not be identified because of their absence in current metabolic databases. Therefore, there remains a tremendous amount to be discovered about the metabolic processes occurring in the gut.

Toxic Metabolites The intestinal microbiota can also produce a variety of metabolites such as NH3, H2S, and deconjugated bile and phenolics, which are toxic to the host both locally in the small intestine and systemically. The deamination of amino acids by microorganisms leads to the production of ammonia, a toxic metabolite that has been considered in the etiology of IBD. This was hypothesized because in vitro studies have shown that the microbiota of IBD patients produce more ammonia compared to those from healthy individuals

54

WILLING AND JANSSON

(Van Nuenen et al., 2004). Perfusion of ammonia into the rat colon showed that the life span of colon cells is shortened by ammonia and it also induces proliferation of mucosa (Lin and Visek, 1991). Because ammonia is used as a preferential nitrogen source by carbohydrate-fermenting bacteria (Bryant, 1974), ammonia concentration in the feces can be reduced by feeding fermentable carbohydrates (Awati et al., 2006). Hydrogen sulfide (HzS) is produced by bacteria as a result of the fermentation of sulfurcontaining amino acids and other dietary and host-derived sulfur sources, including sulfurcontaining mucins (Florin et al., 1991; Magee et al., 2000). H2S reduces overall metabolic activity of the intestinal epithelial cells leading to a reduction in the oxidation of glutamine, butyrate, and acetate (Leschelle et al., 2005). This reduction in overall metabolic activity and slow down of all cell cycle phases leads to a reduction in mitotic activity. Deconjugation of bile may s e c t fat &gestion and fat soluble vitamin absorption (Knarreborg et al., 2002a) and production of lithocholic acid may be toxic to enterocytes (Jharreborg et al., 2002b; Wanitschke and Ammon, 1978). Lactobacilli are responsible for the majority ofbile salt hydrolysis, although Bacteroides, Bijidobacterium, and Clostridium spp. all have bile salt hydrolase activity (Gaskins, 2001). Changes in the microbiota have been correlated to increased dehydroxylation of cholic acid, a main bile acid, to deoxycholic acid (Kitahara et al., 2004). Increased levels of bile acid metabolites have also been detected in Crohn’s patients, which may play a role in disease progression (Jansson et al., 2009). Microbial production of phenolics can also lead to the production of toxic metabolites, although this is highly variable between individuals, most likely because the variation in the composition of gut microbiota (Van Nuenen et al., 2004). For example, metabolism of tyrosine leads to the production of several toxic compounds including p-cresol (Bakke and Midtvedt, 1970). The p-cresol is absorbed and conjugated to p-cresylsulphate, which has been

shown to increase the percentage of leukocytes displaying oxidative burst activity (Schauser and Larsson, 2005). It has been suggested that microbiologically produced phenolics reduce growth in weanling pigs and antibiotic treatment reduces urinary and fecal excretion of aromatic compounds (Yokoyama et al., 1982). Bacteroides, Lactobacillus, Clostridium, and Bijidobacterium species strains are major producers of phenolic compounds from aromatic amino acids (Macfarlane and Macfarlane, 1995).

Bioactive Metabolites Bacteria along with digestive enzymes break down dietary proteins, converting them into peptides, some of which are bioactive. The extent of in vivo production of bioactive peptides by luminal bacteria has yet to be directly studied. It is known, however, that bioactive peptides have diverse effects on the host, ranging from local effects in the intestine (such as mucin production) to systemic effects (such as reduced blood pressure). Many of the effects of bioactive peptides are receptormediated and, although some of those receptors have been identified, there still remains much to be defined. P-casomorphin-7, a bioactive peptide that is a product of milk fermentation, causes a dramatic increase in mucin production in rat intestinal cells (Trompette et al., 2003). This bioactive peptide acts directly on the intestinal epithelium by activation of specific receptors (Zoghbi et al., 2006). This interaction likely leads to improved intestinal protection and may have implications for improved intestinal health. Some probiotic microbes, such as Lactobacillus casei (Thoreux et al., 1998) also produce bioactive metabolites that specifically interact with the host epithelial cells in the intestine. L. joknsonnii appears to produce a metabolite that lowers blood pressure via a histaminergc receptor present in the intestinal epithelium (Tanida et al., 2005). These different metabolites may, at least in part, be responsible for some of the beneficial properties of ingestion of probiotic bacteria.

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Immune Development and Protection The effects of the microbiota on the mucosal immune system are vast, ranging from the enhancement of barrier integrity by stimulating the expression of tight junction proteins (Danielsen et al., 2007), to the balancing of the T-helper (Th) cell population (Mazmanian et al., 2005). For example, B. thetaiotaomicron colonization leads to differential transcription of host genes responsible for nutrient absorption, mucosal barrier fortification, xenobiotic metabolism, angiogenesis, and intestinal maturation (Hooper et al., 2001). A healthy microbiota is important not only in the development of immunity, but also in maintained protection against specific pathogens, such as Salmonella. A normal mouse microbiota confers resistance to Salmonellainduced gastroenteritis and shifting of the normal microbiota creates increased susceptibility to Salmonella (Sekirov et al., 2008). This function is not simply a matter of total bacterial abundance. The type of bacteria present is important because disruption of the composition of the microbiota without a reduction in total levels led to increased susceptibility to enteric infection by Salmonella and other enteric pathogens (Sekirov et al., 2008). The mechanistic explanation for this is yet to be defined; however, it is expected that it is either the result of competitive exclusion by commensal bacteria or the result of immune regulation by other bacteria. In a separate study, Brandl et al. (2008) reported that the normal microbiota seemingly inadvertently prevented infection by enterococci by causing the host to secrete the antimicrobial peptide, RegIIIy. When the normal microbiota was removed by use of antibiotics, the expression of this peptide dropped, and the host became susceptible to infection. However, protection could be maintained by adding bacterial lipopolysaccharide (LPS) during the course of antibiotic treatment, indicating the importance of maintaining appropriate immune stimulation. The ability of the microbiota to regulate the host immune system is extremely important because, if the homeosta-

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sis established between the host and microbiota is disrupted, overt inflammation can occur lealng to IBD as discussed previously. MICROBIAL INTERACTIONS

Syntrophy Bacterial dependency on co-colonizers is exemplified by cross-feedmg. Synergism can be exemplified by the relationship between Eubacterium hallii and Anaerostipes caccae. These organisms cannot grow separately on starch; however, they are able to do so when cocultured with Bifidobacterium adolescentis. B. adolescentis provides these organisms with lactate to grow and to produce butyrate. A second mechanism is demonstrated by the inability of Roseburia sp. A2-183 to grow on fructooligosaccharides (FOS). When co-cultured with B. adolescentis, Roseburia is able to utilize the partially degraded FOS (Belenguer et al., 2006). The syntrophic interactions between bacteria in the metabolism of carbohydrates are shown in Fig. 4. The removal of reducing units (H2) is very important for overall microbial community dynamics and is a key process in many syntrophic interactions. As mentioned previously, about 30% to 50% of individuals in the Western population dispose of hydrogen via methanogenesis. These individuals can be identified using a simple breath test for methane production. The remaining human population use different H2 disposal routes, via sulfate reduction or reductive acetogenesis. Gut metagenome studies of two American individuals (Gill et al., 2006) found that the genes involved in methanogenesis were abundant, suggesting that methanogenesis in their gut environments occurred via the H2 disposal route. By contrast, the metagenomes of Japanese individuals (Kurokawa et al., 2007) and the proteomes of Swedish individuals (Verberkmoes et al., 2009) contained genes and proteins, respectively, that were typical of the acetogenic H2 disposal route. In most animal species, H2-utilizing organisms are dominated by methanogens (Morvan


S

Systemic Circulation ESDO9.047

FIGURE 4 Synergisticaction ofbacteria in degradation of carbohydrates. Different bacterial species work together in the metabolism of dietary carbohydrate, each contributing to the process.

et al., 1996). Other known Hz-utilizing organisms found in the gut include homoacetogens and sulfate-reducing bacteria (Christ1 et al., 1992; Strocchi et al., 1994). In an experiment where M . smithii and B. thetaiotaomicron were used to inoculate mice alone or together, cocolonization resulted in a 100- to 1,000-fold increase in the abundance of both organisms. This increased colonization indicated that the

methanogen was more successful with a source of H2 and that the accumulation of Hz in the environment was inhibiting the growth of B. thetaiotaomicron. Because there is a great deal of competition for Hz in the gut between acetogens, sulfite-reducing bacteria, and methanogens, it seems likely that in a normal gut environment there is never a buildup of H2 to inhibit bacterial NADH dehydrogenases.

3. THE GUT MICROBIOTA: ECOLOGY AND FUNCTION

Bacteriocins One bacterial competitive tactic is to produce bacteriocins that kill neighboring bacterial cells. This mechanism, however, does not always result in the expected outcome so that it remains competitive. For example, one might expect that, in a system where there is a bacteriocin producing bacterium and one susceptible to that bacteriocin, the producing strain would have a growth advantage. In a simple system this would likely be the case, but, as described previously, the gut microbiota is extremely complex. The complexity of this situation was demonstrated by introducing three E. coli strains that included an isolate that produced colicins (narrow-spectrum antibiotics produced by and active against E . cofi), one that was susceptible to colicins, and another that was resistant to colicins (Kirkup and Riley, 2004). The result of this experiment was a bacterial game of “rock-paperscissors,” where each strain took its turn at being the dominant E. coli found in the mouse gut. Interacting through the Host The host also plays an important role in the competition between bacteria. B. thetaiotamicron (gram-negative) stimulates host expression of genes responsible for antimicrobial activity targeting gram-positive bacteria, whereas Bifidobacterium fongum (gram-positive) suppresses the expression of these same genes (Sonnenburg et al., 2006). Another interaction through the host immune system is indicated by a mechanism where the commensal microbiota protects the host from Enterococcus infection by inducing host expression of a c-type lectin that kills gram-positive bacteria (Brand1 et al., 2008). Model Systems There are two in vivo model systems that have been employed to study microbial interactions in the intestine. In the first system, a germ-free (or gnotobiotic) animal is used as a host for a defined bacterial population or community.

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An alternate approach is to remove individual components of the indigenous microbiota and to observe losses of function or changes in response microbial loss. A number of attempts have also been made to model the GI microbiota in vivo (Kovatcheva-Datchary et al., 2009), but this is challenged by the inability to accurately mimic the contributions of the host. A recent study demonstrated the complexity of the relationships in the GI ecosystem by creating a model system made up of single representatives from each of the two main phyla present in the human GI tract (Mahowald et al., 2009): Eubacterium rectale, a prominent member of Fimicutes, and B. thetaiotaomicron, a common and well-characterized species of Bacteroidetes. These two bacteria were then introduced into germ-free mice alone (mono-association) and together (di-association). Transcriptional profiling of both bacteria and the host revealed that these two bacteria changed their behavior depending on whether they are colonizing the host alone or with other bacteria. Host response to these bacteria was not simply additive and revealed that the interactions between bacteria resulted in amplified and modified microbial-host interactions (Willing and Finlay, 2009) (Fig. 5). Co-colonization with E. rectale caused B. thetaiotaomicron to increase the expression of glycan-degrading enzymes, presumably as a consequence of competition for dietary carbohydrate. As a result of the lack of nutrients, B. thetaiotaomicron also began sending signals to the host to produce glycans that it, but not E. rectale, could utilize. Based on genome analysis of B. thetaiotaomicron and other sequenced Bacteroidetes, these bacteria appear to have a surplus of glycan degradmg enzymes compared to members of Firmicutes. The ability of B. thetaiotaomicron to utilize hostdrived glycans in the face of competition by Fimicutes may therefore be a common adaptation used by Bacteroidetes to remain competitive. Although B. thetaiotaomicron increased its ability to degrade carbohydrate in the face of competition, E. rectale decreased expression


of glycan-degrading enzymes and increased expression of a number of amino acid and peptide transporters. Results of this study also demonstrated that there were synergistic interactions between these two bacteria. Although E. rectale was able to produce butyrate from the metabolism of carbohydrates, this bacterium was able to produce much more butyrate when co-colonized with B. thetaiotaomicron. Moreover, butyrate was more efficiently produced in a situation where acetyl-CoA produced by

A

B. thetaiotaomicron was utilized and converted to butyrate by E. rectale. As dscussed previously, butyrate regulates many host functions. Therefore, the increased production of butyrate resulted in greater host responses to colonization. Although this model gave some new insights into the complex ecology of the gut microbiota, it is yet unclear whether the interactions observed between E. rectale and B. thetaiotaomicron are representative of common interactions between Bacteroidetes and Firmicutes.

B

Dietary Carbohydrate

Dietary Carbohydrate

High Glycoside HydrolaseActivity

Bactenal Signals

Small Host Response

C

Increased Glycoside HvdrolaseActivitv

Small Host Response

Low Glycoside HvdrolaseActivitv

Large Host Response ESD09-046

FIGURE 5 A representaaon of B. thetaiotaomicron and E. rectale colonizing the gastrointeshnal tract of the mouse alone (A, B) or together (C). Interactions between bacteriaresultinchangesinbactellalphysiologyandhowtheyaffectthe host. Compemon with E. rectale for nutrients causes B. thetaiotaomicron to stimulate the production of glycans &om the host that it, but not E. rectale, can ut&ze. E. rectale utihzes acetylCoA produced by B thetaiotaomicron resulting in an increased produchon of butyrate. The interactions between bacteria, including competiaon and synergistic interactions, result in an amphfied host response. Adapted &om W&ng and Finlay (2009).

3 . THE GUT MICROBIOTA: ECOLOGY AND FUNCTION W 59

CONCLUSION Although we are gaining increasing appreciation of the important role that the gut microbiota play in health and disease, we are also becoming aware of the complexity of the gut ecosystem. The advent of “omics” tools has enabled us to obtain vast amounts of data about species, genes, proteins, and metabolites in the gut. However, we are lagging behind in the development of bioinformatics and biostatistics tools that enable us to mine these enormous data sets and to perform correlations between them. It is currently a massive puzzle to determine which microbial species and/or processes are essential for normal gut function, for different disease states, and other physiological conditions, such as obesity. These puzzles are further confounded by the large variation in microbial species composition among individuals that makes generalizations Ifficult. The study of genetically matched twins and defined model systems are examples of approaches that have promise to help define diagnostic targets and disease biomarkers. Eventually, we will need to combine information gleaned from molecular approaches with detailed biochemical and physiological studies of the effects of specific microbes, proteins, and metabolites on health (e.g., in a gnotobiotic animal model) to fulfill the requirements of Koch’s postulates. Current evidence also suggests that these simple models will need to be modified to take into account interactions that occur between microbes and the host in the gut. The hope for the future is that we will be able to design diet plans and drugs that optimize beneficial microbial functions in the gut and that these will result in health benefits to individuals. REFERENCES Andersson, A. F., M. Lindberg, H. Jakobsson, F. Backhed, P. Nyren, and L Engstrand. 2008. Comparative analysis of human gut microbiota by barcoded pyrosequencing. PLOS One 3:e2836. Apajalahti, J. H. A., A. Kettunen, M. R. Bedford, and W E. Holben. 2001. Percent G + C profiling accurately reveals diet-related dfferences in the gastrointestinal microbial community of broiler chickens. Appl. Environ. Microbiol. 67:5656-5667.

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Turnbaugh, P. J., M. Hamady, T. Yatsunenko, B. L. Cantarel, A. Duncan, R. E. Ley, M. L. Sogin, W. J. Jones, B. A. Roe, J. P. AtTourtit, M. Egholm, B. Henrissat, A. C. Heath, R. Knight, and J. I. Gordon. 2009. A core gut microbiome in obese and lean twins. Nature 457~480-484. Turnbaugh, P. J., R. E. Ley, M. A. Mahowald, V. Magrini, E. R. Mardis, and J. I. Gordon. 2006. An obesity-associated gut microbiome with increased capacity for energy harvest. Nature 444: 1027-1031. Umesaki, Y., Y. Okada, S. Matsumoto, A. Imaoka, and H. Setoyama. 1995. Segmented filamentous bacteria are indigenous intestinal bacteria that activate intraepithelial lymphocytes and induce MHC class-I1 molecules and fucosyl asialo Gml glycolipids on the small-intestinal epithelialcells in the ex-germ-free mouse. Microbiol. Immunol. 39:555-562. van der Wielen, P. W. J. J., S. Biesterveld, S. Notermans, H. Hofstra, B. A. P. Urlings, and F. van Knapen. 2000. Role of volatile fatty acids in development of the cecal microflora in broiler chickens during growth. Appl. Environ. Microbiol. 66:2536-2540. Van Loo, J. A. 2004. Prebiotics promote good health: the basis, the potential, and the emerging evidence. J. Clin. Gastuoenterol. 38:S70-75. Van Nuenen, M., K. Venema, J. Van der Woude, and E. Kuipers. 2004. The metabolic activity of fecal microbiota from healthy individuals and patients with inflammatory bowel hsease. Digest. Dis. Sci. 49:485-491. Verberkmoes, N. C., A. L. Russell, M. Shah, A. Godzik, M. Rosenquist, J. Halfvarson, M. G. Lefsrud, J. Apajalahti, C. Tysk, R. L. Hettich, and J. K. Jansson. 2009. Shotgun metaproteomics of the human distal gut microbiota. ISMEJ. 3:179-189. Walker, A. 2007. Genome watch - say hello to our little friends. Nut. Rev. Miuobiol. 5:572-573. Wanitschke, R., and H. Ammon. 1978. Effects of dihydroxy bile-acids and hydroxy fatty-acids on absorption of oleic-acid in human jejunum.J. Clin. Invest. 61:178-186. Wikoff, W. R., A. T. Anfora, J. Liu, P. G. Schultz, S. A. Lesley, E. C. Peters, and G. Siuzdak. 2009. Metabolomics analysis reveals large effects of gut microflora on mammalian blood metabolites. Puoc. Natl. Acad. Sci. USA 106:3698-3703. Willing, B., J. Halfvarson, J. Dicksved, M. Rosenquist, G. Jarnerot, L. Engstrand, C. Tysk, and J. K. Jansson. 2009a. Twin studies reveal specific imbalances in the mucosa-associated microbiota of patients with ileal Crohn’s disease. Inflarnrn. Bowel Dis. 15:653-660.


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ANIMALS AND HUMANS AS SOURCES OF FECAL INDICATOR BACTERIA Christopher K. Yost, Moussa S . Diarra, and Edward Topp

In mixed activity watersheds, water can be compromised by fecal pollution from a variety of sources including domesticated livestock, humans, and wildlife. The density and the specific types of fecal bacteria will depend on the feces originating from each potential source and, in the case of humans, confined livestock, and poultry production systems, changes in fecal populations that accompany sewage or manure storage and treatment prior to release into the broader environment. Enteric bacteria that are pathogenic for humans may be found in the intestinal tracts of various wildlife or domesticated animal hosts. They are generally only sporadically present in individuals and distributed at low densities in populations at a landscape scale. The ability to quantitatively detect all pathogens in a given environmental sample is impractical if not intractable, and, thus, attempting to do so has not been used as a standard approach for validating water quality. Rather, fecal indicator bacteria are used to detect fecal contamination of water, to mandate water quality, and to act as surrogates for studies evaluating the environmental fate of

bacterial enteric pathogens (National Research Council, 2004). Indicator bacteria must ideally meet a number of criteria in order to be useful robust indicators of fecal pollution (Buttiaux and Mossel, 1961; Yates, 2007). They should be ubiquitously distributed in high density in the gastrointestinal (GI) tract of humans and animals, absent from environmental matrices uncontaminated with fecal material, and readily detected by reasonably inexpensive means. Indicator bacteria should be reasonable surrogates of pathogenic bacteria with respect to growth requirements, survival characteristics, and susceptibility to environmental stress. The intestinal microflora is a diverse community that plays a key role in the maintenance of health and the efficient utilization of nutrients (O’Hara and Shanahan, 2006). The population structure and phenotypic characteristics of shed enteric bacteria are influenced by a variety of anthropogenic factors, including diet and antibiotic use. The veterinary use of antibiotics and other interventions used to enhance feed conversion in commercial agricultural practices influences the composition of gut bacteria (Diarra et al., 2007; Lefebvre et al., 2006; Bonnet et al., 2009). This chapter considers the distribution of the major indicator bacteria, Escherichia coli, Enterococcus spp., and a variety of anaerobes

Christopher K. Yost,Department ofBiology, University ofRegina, Regina, SK, Canada S4S OA2. Moilssa S. Diawa, Pacific Agri-Food Research Centre, Agriculture and Agri-Food Canada, Agassiz, BC, Canada VOM IAO. Edward Topp, Agriculture and Agri-Food Canada, London, ON, Canada N5V 4T3.

The F e d Bacteria, Edited by M. J. Sadowsky and R. L. Whitman, 0 2011 American Society for Microbiology, Washington, DC

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TABLE 1

Indicator bacteria

Indicator bacterium

General characteristics

Concentration (CFU/g feces)a

Bactevoidales

Non-spore-forming gram-negative rod, obligate anaerobe, dificult to culture

Human: l o t 1 (Finegold et al., 1974) Bovine: NA6 Swine: NA* Poultry: NA"

B9dobacterium

Non-spore-forming grampositive rod, obligate anaerobe, used as probiotic supplements in dairy products

Human: 10' (Resnick and Levin, 1981) Bovine: 104-10' (Resnick and Levin, 1981) Swine: 10' (Resnick and Levin, 1981) Poultry: lo5 (Baurhoo et al., 2007)

Clostridium pe$iingens

Spore-forming gram-positive rod

Human: lo6 (Carman et al., 2008) Bovine: 10' (Tschirdewahn et al., 1991) Swine: 10' (Cox et al., 2005) Poultry: lo3 (Cox et al., 2005)

Escherichia coli

Non-spore-forming gram-negative rod, facultative aerobe, easy to culture

Human: 10' (Adlerberth and Wold, 2009) Bovine: 105-108 (Lefebvre et al., 2006) Swine: lo7 (Duriez and Topp, 2007) Poultry: 107-108 (Diarra et al., 2007)

Entevococcus spp.

Non-spore-forming pi-positive cocci, facultative anaerobe, easy to culture

Human: 105-108 (Tannock and Cook, 2002) Bovine: 104-10' (Lefebvre et al., 2006) Swine: lo8 (Fava et al., 2007) Poultry: 106-107 (Diarra et al., 2007)

"The numbers provided are general averages and it is important to note that for particular indicators (especially C. pey'?i"ngens) the abundance in individuals can vary greatly. 'No cultural enumerationsare readily available, possibly due to difficulty in culturing. However, quantitative PCR for enumeration of Bacteroidules indicates a high prevalence in bovine and swine feces with 10'" and 10" gene copiedgr feces, respectatively (Okabe and Shiinazu, 2007). The prevalence in poultry is thought to be considerably lower (please refer to text for more detail).

(including Clostridium p e f i n g e n s , B$dobacteria spp., and members of the Bactevoidales) in humans, in domesticated animals and birds, and in undomesticated animal hosts (Table 1).Factors that can influence the abundance, composition, and characteristics of fecal bacteria, notably antibiotic use, are presented. Finally, practices that can attenuate or m o d @ environmental exposure to indcator bacteria once shed, namely manure storage and treatment, are considered. ESCHERICHLA COLI

The fecal indicator bacterium E. coli is ubiquitously distributed in the digestive tract of homeothermic animals. Gut populations can include resident and transient clones and, depending on the complement of virulence genes, can have a commensal or pathogenic (either intestinal or extraintestinal) host interactions (Janda and Abbott, 2006; Lefebvre et al., 2006). Typical E. coli densities in mammalian

fecal matter are >lo7 CFU/gr feces. E. coli may be carried by a variety of heterothermic hosts, but likely only transiently, and the distribution in populations is heterogeneous. The organism can be harbored, survive, and potentially proliferate in other secondary hosts and environmental matrices (Anderson et al., 2008). E. coli is frequently used to establish host sources of fecal contamination in environmental studies (Bartosch et al., 2004).

E . cold in Humans E. coli is one of the first colonizers of the human GI tract. At birth, the vaginally delivered neonate can be colonized with E. coli via exposure to the maternal flora and subsequently through exposure to other people (Adlerberth and Wold, 2009). By 12 months of age, 100% ofinfants are colonized and, thus, E. coli is ubiquitous in adults. The abundance of E. coli in feces (CFU/g) can range up to lo9 in l-weekold infants and is on the order of lox in adults

4.

(Adlerberth and Wold, 2009). Humans can also shed E. coli harbored external to the GI tract; for example, when ill with an infection of the urinary tract (Donnenberg, 2000). The human gut microbiome (chapter 3), influences of pathogenic E. coli on human health (chapter 7), and fate of pathogenic E. coli in the environment (chapter 10) are reviewed in detail elsewhere in this book.

E . coli in Domesticated Mammals and Birds Both monogastric and polygastric (ruminant) livestock carry E. coli. In some cases they are asymptomatic carriers of serotypes that are pathogenic for humans should they be exposed to contaminated meat products, water, or field crops. Cattle, sheep, and goats are the most important reservoirs for enterohemorrhagic E. coli (EHEC) 0157:H7, but swine as well as d d animals can also carry EHEC (La Ragione et al., 2009; Yoon and Hovde, 2008). Various pathotypes of E. coli can also be associated with a wide variety of serious illness in animals, includmg GI infections accompanied by diarrhea in calves, lambs, and piglets; mastitis in b r y cattle; and colibacdlosis in poultry (Gyles, 1994). Overall, animals f?om a variety of livestock and poultry operations wdl shed commensal E. coli, and potentially zoonotic pathotypes, with or without overt clinical symptoms (Bonnet et al., 2009). The temporal and spatial patterns of E. coli shedding by cattle can be influenced by the distribution and dynamics of bacterial populations within a herd and within the digestive tract of individual animals. Genotyping studies have revealed animal-to-animal transmission of specific E. coli clones between cattle stocked at commercial densities in feedlots (Sharma et al., 2008). In a Scottish study of pastured animals, 8% of cattle excreted 99% of the E. coli 0157:H7 detected (Chase-Topping et al., 2008). Those animals that excreted high rates of toxigenic E. coli over several months (“super-shedders”) were found to have the mucosal epithelium of the terminal rectum colonized by the bacterium. In contrast, those animals that were not rectally colonized excreted the pathogen at lower rates and much

SOURCES OF FECAL INDICATOR BACTERIA W 69

more transiently. The E. coli 0157:H7 phage type PT 21/28 was often associated with colonized animals,but the basis for differential rectal colonization remains to be elucidated. The diet fed to livestock wdl influence the gut flora and sheddmg rates of specific bacteria (Bach Knudsen, 2001). For example, switching cattle from a grain-based diet to hay reduces shedding of generic E. coli and E. coli 0157:H7 (Callaway et al., 2009). Household dogs and cats are also reservoirs of E. coli, including pathogenic types (Beutin, 1999; Damborg et al., 2009a). E. coli was detected in 101 of 168 cloaca1 swabs of cockatoos, a common pet, housed within a single facility ( F l a m e r and Drewes, 1988).

E. coli in Undomesticated Animals, Birds, and Other Hosts Gordon and Cowling (2003) undertook an extensive survey of E. coli in Australian fauna, sampling 2,300 individuals drawn from over 350 undomesticated vertebrate hosts living across the continent. Overall, this important study established the distribution of E. coli in a range of hosts and explored some key factors specific to the host, the climate, and proximity to humans. Key findings of their studies included the following: E. coli was more commonly detected in mammals, less frequently in birds, and infrequently in fish, frogs, and reptiles. E. coli was less prevalent in carnivores than herbivores or omnivores. Proximity to humans increased the likelihood of E. coli being detected in wildlife. Animals living in desert climates were less likely to harbor E. coli than if they lived in temperate or tropical regions. Organisms with large body mass or with a GI tract that included a hindgut fermentation chamber had a higher prevalence of E. coli. A number of reports indicate that reptiles and amphibians are carriers of E. coli. In a study undertaken in Indonesia, E. coli was isolated from wdd, but not fkom captive, Komodo dragons (Vauanus komodoensis) (Montgomery et al., 2002). Various species of fecal coliforms were isolated fiom mamondback terrapin turtles (Malaclemys terrapin centrata) captured in estuarine waters off of

70 W YOSTETAL.

Florida, with E. coli detected the most frequently (41% ofanimals sampled) (Hanvood et al., 1999). In a survey of reptiles in the Trinidad Emperor Valley Zoo, 52% of fecal samples and cloacal swabs fi-om snakes contained E. coli (Gopee et al., 2000). Animals carrying the bacterium included the American crocodde (Crocodylus acutus) and the spectacled cayman crocodile (Cayman crocodilus), and a variety of snakes and tortoises. Eight percent of cloacal swabs taken fi-om green turtles (Chelonia mydas) nesting on a Costa Rican beach had E. coli (Santoro et al., 2006). However, the authors pointed out that the samples may have been contaminated by beach sand. Waturangi et al. (2003) reported the antibiotic resistance characteristics of 28 E. coli fecal isolates obtained from monitor lizards in Indonesia. The sampling method and number of animals and species (Varanus spp.) sampled were not specified. E. coli and Enterococcus spp. were detected in 6 and 8 of 23 rectal swabs, respectively, obtained from Ricord's iguanas (Cyclura yicordii) captured in a national park in the Dominican Republic (Maria et al., 2007). Tadpoles of the American bullfrog (Rana catesbeiana) maintained at 26 to 28OC carried E. coli 0157:H7 for 2 weeks following inoculation (Gray et al., 2007). A survey of turtles, snakes, and lizards in Austraha detected E. coli in 10% or fewer of the indwiduals sampled (Gordon and Cowling, 2003). Taken together, these reports indicate that heterothemnic reptiles in wam-climate marine and terrestrial systems can harbor E. coli. Whether GI colonization is transient or becomes established, whether gut residents are generally commensal or pathogenic, and the fiequency of E. coli carriage in reptiles and amphibians in temperate and colder climates all remain to be determined. Animals living in the polar regions carry E. coli. Weddell seal (Leptomychotes weddellii) scats obtained near McMurdo Station in Antarctica contained up to 1.2 X lo4 CFU/g dry weight E. coli (Lisle et al., 2004). E. coli were isolated from rectal swabs in a survey of arctic fox (Alopex lagopus) in northern Sweden, but the authors did not specifj the frequency of detection (Aguirre et al., 2000). Marine inaminals living in temperate or tropical waters carry

E. coli. The anuses and feces of 254 individual fi-ee-ranging bottlenose dolphins (Tursiops trtrncates) captured in US coastal waters were sampled (Buck et al., 2006) and 41% of the animals carried E. coli. The intestinal contents of the freshwater fish jenynsia multidentata (livebearer) recovered from the Suquia River in Argentina were found to contain E. coli at varying concentrations (Guzmin et al., 2004). Aquarium experiments with fish immersed in water containing varying densities ofan E. coli inoculum revealed that higher levels of water contamination resulted in higher levels of E. coli in gut contents. About 20% of 32 benthic fish (common carp, Cyprinus cayio; white sucker, Catostomus commersonii; brown bullhead, Ameiuvus nebulosus; ruffe, Gymnocephalus cevnuus; and round goby, Neogobius melanostomus) sampled in the Duluth-Superior Harbor on Lake Superior harbored E . coli (Hansen et al., 2008). Pelagic species ( n = 25) were also sampled (white perch, Morone amevicana; yellow perch, Perca Javescens; pumpkinseed, Lepomis gibbosus; and rock bass, Ambloplites vupestvis) and found to harbor E . coli at much lower frequencies. Fish isolates were compared with environmental and mammalian isolates by rep-PCR and those whose source could be identified were most similar to isolates from sediments, wildfowl, and wastewater. O n this basis the authors suggest that fish can be considered as a vector for E . coli rather than a reservoir. ENTEROCOCCUS SPP. Enterococci are gram-positive bacteria ubiquitously found in the GI tracts of animals, birds, and humans, as well as in soil and water. Aspects of the ecology and the pathogenicity of some Enterococcus spp. have been reviewed elsewhere (Facklam et al., 2002; Hancock and Gilmore, 2006; Scott et al., 2005). Prior to 1984, members of the genus Entevococcus were classified as group D streptococci. There are now about 40 recognized Enterococcus species, including E. aquimarinus, E. asini, E . avium, E. caccae, E . camelliae, E . canintestini, E. canis, E. casselijlavus, E. cecorum, E . colum-

4.

bae, E. devriesei, E. dispar, E. durans, E. faecalis, E. faecium, E. Javescens, E. gallinarum, E. gilvus, E. haemoperoxidus, E. hevmanniensis, E. hirae, E. malodoratus, E. moraviensis, E. mundtii, E. pallens, E. phoeniculicola, E. porcinus (villorum), E. ratti, E. rafinosus, E. saccharominimus (italicus), E. saccharolyticus, E. solitarius, E. pseudoavium, E. seriolicida, E. sulfureus, E. silesiacus, E. thailandicus, and E. termites (http://www.bacterio .cict.fr/e/enterococcus.html).The various Enterococcus spp. strains share many similar phenotypic and biochemical characteristics, thus making precise species level identification very difficult. Consequently, it is important to develop methods that allow unambiguous differentiation between strains with a high virulence potential and the presumably harmless, or even beneficial, strains used in the food industry and that are found in the environment. Some Enterococcus strains, particularly of E. faecium and E. faecalis, are of importance to public health because of their resistance to multiple antibiotics and the presence of virulent strains (Hancock and Gilmore, 2006; Ogier and Serror, 2008; Weaver, 2006). Furthermore, the ability of Enterococcus to acquire and transfer antibiotic resistance through plasmids and transposons, chromosomal exchange, or mutation presents a significant challenge for therapeutic intervention (Conly, 2002). Numerous mechanisms of antibiotic resistance have been described in the enterococci. The main resistance mechanisms include those that involve target modification or substitution, antibiotic modification or destruction, antibiotic efflux, and restricted permeability to antibiotics (Leclercq and Courvalin, 2005). The aim of all of these resistance mechanisms, which are encoded by specific resistance genes, is to allow the enterococci to grow in the presence of antibiotics, even at high concentrations. Because of their inherent ability to acquire and disseminate antibiotic resistance determinants, enterococci have been proposed as indicator bacteria for antimicrobial resistance, as well as indicators for the hygienic quality of food and water (Pierson et al., 2007; Simjee et al., 2006; Wheeler et al., 2002).

SOURCES O F FECAL INDICATOR BACTERIA W 71

Enterococci in Humans Enterococcus spp. are among the most abundant gram-positive cocci in the human intestine, with densities ranging from lo5 to 10' CFU/g of intestinal content (Tannock and Cook, 2002). The most frequently detected species are E. faecalis and E. faecium, varying in abundance fkom lo2 to 10' CFU/g (Ogier and Serror, 2008). In humans, enterococci can cause urinary tract infections, bactereinia, intra-abdominal infection, and endocardh. About 95% of all these infections are caused by E. faecalis and E. faecium (Hancock and G h o r e , 2006; U b i et al., 2007). The virulence of enterococci appears to be multifactorial and associated with several genes including ace (collagen binding cell wall protein), acm (surface-exposed antigen), agg (aggregative pheromone inducing adherence to extra-matrix protein), agrB$ (AgrBfi protein of faecahs), esp (enterococcal surface protein), hyl (hyaluronidase), cad2 (pheromone cADl precursor lipoprotein), CAM373 (sex pheromone cAM373 precursor), cCF2 0 (pheromone cCFlO precursor lipoprotein), cob (pheromone COB1 precursor/lipoprotein, YaeC fBrmly), cpdl (pheromone cPDl lipoprotein), cylABLM (hemolysin), efaAfs (endocardtis specific antigen), sagA (secreted antigen), and gelE (gelatinase). The esp gene may be a specific marker for the detection of human fecal pollution, although some sourceslack this marker gene (Byappanahalh et al., 2008). In Southeast Queensland (Australia), t h s marker was able to quantiG enterococci to about lo3 CFU per 100 mL ofwater, suggesting that esp may be a reliable marker to detect human fecal pollution in surface waters (Ahmed et al., 2008). O n the other hand, Byappanahah et al. (2008) reported that the esp gene was not in fact human-specific and was thus of limited use for ascribing sources. Enterococci in Domesticated Mammals and Birds To date no single enterococcal species has been clearly identified as being primarily associated with one livestock or animal type. Such host-dependent specificity would be useful for identi6ing livestock sources of fecal pollution

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in food and environments. In a longitudinal study of feedlot cattle (Bos taurus), Enterococcus was excreted at high frequency for at least 186 days with fecal counts ranging from lo4 to l o 6 CFU/g (Lefebvre et al., 2006). At day 186, Enterococcus was shed by almost all the steers and its prevalence was higher than that observed in younger steers. Entevococcus was shed by 50% of the animals on day 46 and by more than 94% on day 109. Such a high prevalence (95%) was also found in the colon at slaughter (Lefebvre et al., 2006). Survival rates of indcator bacteria in bovine feces have recently been reported (Sinton et al., 2007) and Moriarty et al. (2008) showed that counts and prevalence of enterococci was 1.3 X lo4 per g of wet weight sample of fresh bovine feces in New Zealand. However, all counts were highly variable within and between samples. Nevertheless, their results clearly indicate that fresh bovine feces are a significant source of these indicator bacteria on New Zealand pastures (Moriarty et al., 2008). A great genetic diversity (different genotypes) among isolates of enterococci, including E. faecium, E. casselgavus, and E. faecalis, from bovine origin has been reported (Peterson-Wolfe et al., 2008). The GI tract ofpigs (Sus domesticus) is populated with bacteria whose abundance (cells/g of fresh material) increases from lo7 in the stomach to lo9 in the distal small intestine to lo1' to lo1' in the colon (Bach Knudsen, 2001). The majority of the cultivable bacteria in the colon are gram-positive bacteria, including streptococci and lactobacilli. Fluorescence in situ hybridization studes indicated that, of the four groups of bacteria examined (Bijidobacterium, Bacteroidales, lactobacilli/enterococci, Clostridium pe?7ryingens/histolyticum), the lactobacilli/ enterococci were the most dominant in the intestine, cecum, and the colon with densities ranging from 1.6 X lo8 to 6.8 X 10' CFU/g of contents (Fava et al., 2007). A survey of 122 horses (Equusferus caballus) from Slovahan farms revealed enterococcal densities ranging from lo1 to lo5 CFU per gram of feces (Laukovi et al., 2008). Forty-three enterococci were isolated, 14 (32.5%) of which were of

unknown identity, 25 (58.1%) were identified as E.faecium, three (6.9%) were E. mundtii, and one was identified as E. faecalis. In chickens (Gallus gallus, or G. gallus domesticus), Enterococcus spp. densities were higher in the cecum (lo8 to lo6 CFU/g) than in the cloaca (lo7 to lo6 CFU/g) and Enterococcus spp. were more abundant in 7-day-old birds than in 35-day-old birds (Diarra et al., 2007). Surveys of meat products at the retail level reveal insights into the presence and type of various Enterococcus spp. in commercial livestock and poultry that may have become contaminated during processing. Hayes et al. (2003) analyzed 981 samples of retail raw meats (chicken, turkey, pork, and beef) obtained from 263 grocery stores in Iowa for the presence of Enterococcus spp. They recovered 1,357 enterococci isolates with contamination rates ranging from 97% of pork samples to 100% of ground beef samples. E. faecium was the predominant species recovered (61%), followed by E.faecalir (29%), and E. hirae (5.7%). E.faecium was the predominant species recovered from ground turkey (60%), ground beef (65%), and chicken breast (79%), whereas E. faecalis was the predominant species recovered from pork chops (54%).

Enterococci in Undomesticated Animals, Birds, and Other Hosts Enterococci are widely distributed among undomesticated mammals and birds, reptiles, and insects. Some Enterococcus spp. display some degree of host-specificity. For example, E. columbae has been found mostly in pigeons (Columba livia domestica) and E. asini is found mostly in donkeys (Equus ajiricanus asinus) (Aarestrup et al., 2002). The density of fecal enterococci found in glaucous-winged gull (Larus glaucescens) has been reported in the range of 4.0) was considered indicative of human fecal pollution, whereas low ratios were considered to indicate animal sources. Geldreich (1976) subsequently suggested that a FC/ FS ratio of

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