Environmental Microbiology Laboratory Department of - Repositorio

Environmental Microbiology Laboratory

Department of Biology

University of Puerto Rico at Rio Piedras

Enterococcus faecalis-infecting phages (enterophages) as markers of human fecal pollution

and as reservoirs of antibiotic-resistance and virulence genes

Tasha M. Santiago Rodriguez

A dissertation submitted to the Biology Intercampus doctoral program in

Partial fulfillment of the requirements for the degree of

Doctor of Philosophy

April, 2013

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This dissertation has been accepted by the faculty of the

Biology Intercampus Doctoral Program

University of Puerto Rico

Rio Piedras Campus and Medical Sciences Campus

In partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

In the subject of BIOLOGY

Gary A. Toranzos, Ph.D.

Thesis Advisor

Elvira Cuevas, Ph.D.

Committee Member

Maite Muniesa, Ph.D.

Committee Member

Carlos Gonzalez, Ph.D.

Committee Member

Steven E. Massey, Ph.D.

Committee Member

San Juan, Puerto Rico. April 2013.

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TABLE OF CONTENTS

List of Figures 8-9

List of Tables 10-11

Abbreviations 12

Author’s biography 13

Thesis abstract 14

Dedication 15

Acknowledgments 16-17

Chapter 1: Introduction

General introduction 18

Bacterial and viral indicators

Total and thermotolerant coliforms 19

Enterococci 19-20

Bacteriophages 20-21

Molecular techniques 21-22

Ribotyping and Pulsed –Field gel electrophoresis (PFGE) 22

Denaturing Gradient Gel Electrophoresis (DGGE) and Terminal 22-23

Restriction Fragment Length Polymorphism (T-RFLP)

PCR-based techniques 23-24

Correlations of indicators with pathogens and illness 24-25

Antibiotic-resistance and virulence genes in enterococci 25-26

Tetracycline mode of action and resistance genes 26-27

Vancomycin mode of action and resistance-genes 28-29

Enterococcal surface protein 29-30

Horizontal Transfer of antibiotic-resistance and virulence genes 30-31

Thesis direction and general objectives 32

Novel microbial indicator of human fecal pollution 32

Enterophages as vectors of antibiotic resistance and virulence genes 32-33

Literature cited 34-43

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Chapter 2: Enterophages, a group of phages infecting Enterococcus faecalis, and their

potential as alternate indicators of human faecal contamination (as published in Water

Science and Technology).

Abstract 44

Introduction 45

Alternate microbial indicators of fecal water contamination 46-47

Materials and Methods

Host strains 47-48

Host bacteria 48

Samples 48

Optimal conditions for viral replication 48-49

Single Layer Plaque assays 49

Plaque isolation 49

Transmission Electron Microscopy 50

Determination of Burst Sizes 50

Survival 51

Prevalence 51

Results and Discussion 51-52

Viral morphology 52

Burst sizes 53

Survival 53-54

Prevalence 54-55

Conclusions 55-56

Acknowledgements 56


Chapter 3: Characterization of Enterococcus faecalis-infecting phages (enterophages) as

markers of human fecal pollution in recreational waters (as published in Water Reseach).

Abstract 60-61

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Introduction 61-63


Detection of enterophages in animal and human feces 63

Enterophages isolation and purification 64

Morphological characterization of enterophages 64-65

Nucleic acid analyses 65

Prevalence in raw sewage 65

Detection of enterophages and other viral and bacterial indicators 65-66

in a large watershed

Survival of enterophages and coliphages in waters and sand 67

Results and Discussion

Enterophages and coliphages in animal and human feces 67-68

Differences in enterophage isolates according to morphology, genetic 68-69

material and ability to replicate at different temperatures

Prevalence of enterophages in sewage and in a large watershed in Puerto 70-74

Rico

Survival of enterophages in fresh waters 74-75

Survival of enterophages in marine recreational water and sand 75-78

Conclusions 78-79

Acknowledgments 79


Chapter 4: Microbial quality of tropical inland waters and effects of rainfall events (as

published in Applied and Environmental Microbiology).

Abstract 85

Introduction 86-87


Sample collection and sampling sites 87-89

Enumeration of indicators by culture methods 90

DNA extraction and PCR conditions 90-92

USGS precipitation data 92

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Statistical analyses 92-93

Results

Detection of indicators by culture-based methods 93-94

Detection of host-specific Bacteroides by PCR and enterococci by 94-96

qPCR

Correlations of indicators with rainfall and detection methods 96-98

Discussion

Prevalence of the bacterial and viral indicators detected by culture 98-100

methods

Monitoring indicator bacteria by molecular-based techniques 100-101

Conclusions 102

Acknowledgments 102

Supplementary Information 103-108


Chapter 5: Evaluation of Enterococcus phages as indeces of fecal pollution (as published in

Journal of Water and Health).

Abstract 113

Introduction 114-115


Detection of Enterococcus phages in feces and sewage 115-116

Isolation and characterization of Enterococcus phages 116

Inactivation rates and survival studies 116-117

Statistical analyses 117

Results

Enterococcus phages in animal feces and domestic sewage 117-121

Replication and morphology of Enterococcus-infecting phages 121-122

Inactivation rates and survival of Enterococcus phages 123

Survival of enterophages and coliphages in fresh waters 123-126

Enterophages in chlorinated and dechlorinated tap water and sterile 126-127

distilled, tap and wastewater

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Discussion

Enterococcus-infecting phages in feces and domestic sewage 127-128

Inactivation rates and survival of Enterococcus phages 128-130

Conclusions 130-131

Acknowledgments 131

Supplementary information 131-138


Chapter 6: Antibiotic-resistance and virulence genes in tropical environmental

Enterococcus spp. (as accepted in Journal of Water and Health).

Abstract 144

Introduction 145-146


Study sites 146

DNA extraction of the bacterial fraction and enterococci 146-148

Virus concentration and DNA extraction 148

Lysogen induction 149

PCR amplification conditions 149-151

Statistical analyses 151

Results

Bacterial fraction and enterococci isolates 151-153

Viral fraction and induced phages 153-154

Discussion 154-156

Conclusions 156-157

Acknowledgments 157


Chapter 7: General Conclusions and Future Directions

General conclusions 163-165

Future directions 165-166

Literature cited 167

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LIST OF FIGURES____________________________________________________________

Chapter 1

Figure 1: General mode of action of vancomycin.

Figure 2: Integration of a bacteriophage genome into a bacterial genome.

Chapter 2

Figure 1: Transmission Electron Microscopy image (TEM) of enterophages in this study.

Figure 2: Survival of enterophages at 22 °C (A) and at 37 °C (B).

Chapter 3

Figure 1: Shows study site. Sample points are ordered according to their position in the

watershed, localized in the central region of Puerto Rico.

Figure 2: Transmission Electron Microscopy image (TEM) of an enterophage isolate in this

study.

Figure 3: The genetic material of several enterophage isolates in a 0.7% agarose gel.

Figure 4: Enterophage concentration/mL of four different enterophage isolates among four

different temperatures.

Figure 5: Concentrations of enterophages (A), coliphages (B), enterococci (C) and

thermotolerant coliforms (D) at ten different fresh water sample points with different impacts.

Figure 6: Survival of enterophages (A) and coliphages (B) in fresh water.

Figure 7: Survival of enterophages (A) and coliphages (B) in marine water.

Chapter 4

Figure 1: Rio Grande de Arecibo watershed in Puerto Rico.

Figure 2: Credible intervals (CIs) for thermotolerant coliforms (A), enterococci (B), coliphages

(C), and enterophages (D) in the Rio Grande de Arecibo watershed.

Figure 3: Presence of human-, cattle-, and chicken-specific markers in the Rio Grande de

Arecibo watershed.

Figure 4: qPCR for enterococci and correlation between qPCR and culture-based techniques for

enterococci.

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Figure 5: Correlation between rainfall and microbial indicators.

Supplementary Figure 1: Enterococci and thermotolerant coliforms CFU/100mL in the Rio

Grande de Arecibo watershed period of high (November to January) and low rainfall events

(February to April).

Supplementary Figure 2: Enterophages in the Rio Grande de Arecibo watershed according to the

incubation temperature and rainfall period.

Supplementary Figure 3: Coliphages in the Rio Grande de Arecibo watershed according to the

incubation temperature and rainfall period.

Chapter 5

Figure 1: Prevalence of Enterococcus phages in raw domestic sewage in three wastewater

treatment plants (WTP) in Puerto Rico.

Figure 2: Enterococcus faecalis-infecting phages isolated from domestic sewage.

Figure 3: Survival of enterophages and coliphages across a tropical watershed in Puerto Rico.

Figure 4: Survival of enterophages in various sterile water types at 37 ºC.

Chapter 6

Figure 1: Sampled sites in this study.

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LIST OF TABLES_____________________________________________________________

Chapter 1

Table 1: Host specificity of different groups of F + RNA coliphages.

Table 2: Classification of most of the tet determinants according to their mode of action.

Table 3: Vancomycin-resistance in enterococci. Five phenotypes have been identified so far and

enterococci are classified according to the levels of vancomycin and teicoplanin it exhibit

resistant.

Chapter 2

Table 1: Concentrations of coliphages in sewage treatment plants in Puerto Rico (ND-Not

detected).

Table 2: Concentrations of enterophages detected at sewage treatment plants in Puerto Rico.

Chapter 3

Table 1: Prevalence of enterophages in raw and treated sewage at different domestic wastewater

treatment plants in Puerto Rico (PR) and Portugal (Port).

Chapter 4

Table 1: Summary of oligonucleotide primers and probes for PCR and TaqMan qPCR.

Table 2: Reported rainfall during the sampling period (November 2009 to April 2010).

Chapter 5

Table 1: Enterococcus and E. coli-infecting phages per gram of feces in chicken (n=30), cattle

(n=30), dogs (n=12) and pigs (n=10).

Table 2: Mean removal percents of Enterococcus phages in primary effluent from three WTP

(PR-A, PR-B and PR-C) in Puerto Rico.

Table 3: Host range of Enterococcus-infecting phages at different incubation temperatures.

Table 4: T90 values for enterophages and coliphages in three fresh water samples.

Table 5: T90 values (days) for enterophages and coliphages in chlorinated and dechlorinated

drinking water.

Supplementary Table 1: Average of the decay percent day -1

and T90 values for phages infecting

E. faecalis, E. faecium, E. casseliflavus and E. coli in raw sewage at 4°C.

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Supplementary Table 2: Decay values (log·day-1

) for enterophages and coliphages in three fresh

water samples.

Supplementary Figure 1: Survival of enterophages and coliphages in three sites in a tropical

watershed during a period of high rainfall events.


watershed during a period of low rainfall events.

Chapter 6

Table 1: Primers in this study.

Table 2: Prevalence of antibiotic-resistance and virulence-encoding genes in the bacterial

fractions of tropical marine (OP, CBB) and fresh waters (LC, RP) and wet and dry sands (OP,

CBB).

Table 3: Prevalence of the esp variants present in E. faecalis and E. faecium and int in tropical

marine (OP, CBB) and fresh waters (LC, RP) and wet and dry beach sands in the Enterococcus

isolates. Percents are presented in parenthesis.

Table 4: Enterococcus isolates tested for tetracycline (16 μg/mL) and vancomycin (20 μg/mL)

resistance isolated from tropical samples.

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Abbreviations

RNA: Ribonucleic Acid

DNA: Deoxyribonucleic Acid

PCR: Polymerase Chain Reaction

CFU: colony forming units

PFU: plaque forming units

ºC: degree Celsius

mL: milliliters

qPCR: quantitative Polymerase Chain Reaction

SD: standard deviation

μg: micrograms

BLAST: Basic Local Alignment Search Tool

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Author’s Biography

Tasha M. Santiago-Rodriguez was born on September 4, 1985 in Cayey, Puerto Rico. She

attended the public schools Salvador Brau (elementary), Ramon E. Betances (junior high) and

Miguel Melendez Muñoz (high school) in Cayey. As a junior high school student, Tasha always

had a special interest in science and developed a fascination for microbes. Her academic

achievements and interest in science gave her the opportunity to study Natural Sciences in the

University of Puerto Rico, making Tasha the first of her family to pursue a college education. As

an undergraduate, she had an interest to in medicine. As one requirement for Medical School was

experience in research, she had the opportunity to do summer research at the University of

Medicine and Dentistry of New Jersey during the summer of 2005. She worked under the

supervision of Dr. Vincianne Gaussin and Dr. Boudewijn Kruithof, experts in the field of cardiac

valve development. In 2007, Tasha graduated Magna-Cum Laude, but was not accepted into

Medical School. During this time, Tasha strongly considered graduate school as an alternative to

pursue a career in one of her fields of interest, microbiology. In 2008, Tasha was accepted into

the graduate Biology Department at the University of Puerto Rico at Rio Piedras. Tasha has

presented in both local and international meetings including the Health-Related Water

Microbiology and the American Society of Microbiology meetings. This has given her the

opportunity to visit countries such as Greece and New Zealand. She has published 5 articles in

important journals related to water quality including Water Research and Applied and

Environmental Microbiology. Her fields of interest are not limited to environmental

microbiology or the microbial quality of waters, as other interests include ancient DNA and

mechanisms of microbial communication. She has contributed in the characterization of the

microbial communities of coprolites from Pre-Columbian cultures as a way to elucidate dietary

habits and cultural traditions. Data from this project have been accepted for publication in

PLosOne. Tasha has also contributed in the characterization of amber bacteria dating to millions

of years in collaboration with Dr. Raul Cano. She also has an interest in identifying genes

involved in ligninocellulose degradation as alternative sources of energy, resulting in at least 2

manuscripts. Her academic achievements in graduate school make her the student with the

highest number of publications in the history of the Biology Program at the UPR. She has been

accepted as a postdoctoral fellow at the University of California at San Diego, School of

Medicine.

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THESIS ABSTRACT

The introduction of human fecal material into water sources represents a concern to public health

since pathogens could be introduced. Not all laboratories possess the facilities to detect human

enteric pathogens and thus microbial indicators are used. An ideal microbial indicator of human

fecal pollution should be: (i) enumerated using simple laboratory methods, (ii) present in fecally

polluted waters and absent in pristine waters, (iii) associated with the source of the

contamination, (iv) able to survive and inactivate similarly to the pathogen of concern and (v) be

detected in various geographical regions. No microbial indicator of fecal pollution satisfies all

these characteristics. Therefore, we have developed a relatively simple culture technique to

characterized bacteriophages that infect a specific type strain of Enterococcus faecalis, which we

call enterophages, as markers of human fecal pollution. Enterophages were detected exclusively

in domestic wastewaters in Puerto Rico and Portugal, tropical recreational waters and possess a

survival and inactivation rates similar to human pathogenic enteric viruses. Enterophages possess

non-contractile tails of 60 to 200nm, icosahedral capsids of 12 to 80nm and dsDNA genomes of

30 to 40kb. Given that their role as vectors of genes conferring antibiotic-resistance and

virulence to the bacterial host remains largely unknown, the complete genomes of two phages

infecting E. faecalis (one lytic and one lysogenic) are currently being sequenced. Lysogenic

Enterococcus phages, as well as their host isolated from marine and fresh waters, dry and wet

sands in Puerto Rico, were tested for tetracycline (tetM) and vancomycin-resistance (vanA and

vanB) genes. The prevalence of the enterococcal surface protein (Esp), a virulence factor, was

also determined given that it has received great attention as a marker of human fecal pollution.

The prevalence of an integrase-encoding gene (int) specific for E. faecalis phages was

determined since integrases are markers of lysogeny and are responsible for the integration of the

phage genome into the bacterial host. tetM, vanA, vanB, esp and int were only detected in the

bacterial fraction and enterococci, with the exception of int, which was detected in the viral

fractions and lysogenic enterococci phages.

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DEDICATION

This thesis is dedicated to my mother and grandmother, for their unconditional support to pursue

this journey.

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ACKNOWLEDGMENTS

Special thanks to my advisor Dr. Gary A. Toranzos and my thesis committee, Dr. Elvira Cuevas,

Dr. Maite Muniesa, Dr. Carlos Gonzalez and Dr. Steve Massey, for their guidance, support,

inputs, comments and suggestions for the present dissertation.

To all the coauthors of the publications resulting from the present project: Natasha Bonilla,

Catalina Davila, Dr. Patricia Marcos, Manuela Cadete, Sylvia Monteiro, Jessica Rivera, Mariel

Coradin, Joel Gonzalez, Dr. Carlos Toledo, Dr. Raymond Tremblay, Dr. Jorge Santo-Domingo,

Miguel Urdaneta, Dr. Hodon Ryu and Dr. Ricardo Santos.

To all the undergraduate students of the Environmental Microbiology Laboratory (2008-2013),

of which I had the opportunity to teach something and learn from: Gwendolyn Arguello, Jean F.

Ruiz, Gabriela Tirado, Alfredo Gonzalez, Dashari Colon, Jose Soto and Alex Vermont.

To the Autoridad de Acueductos y Alcantarillados for collecting sewage samples and Marisol

Rodriguez (my mom) for collecting fresh water samples from el Rio La Plata. To the US

Geological Survey and the Department of Natural Resources of Puerto Rico for data used in the

present project.

I would like to thank Dr. Pablo A. Ortiz Pineda who taught me several of the bioinformatic

programs and molecular techniques, and for support to pursue this project. I would also like to

thank Natasha Bonilla who taught me the culture techniques for the detection of enterophages

and Ana Rita Patricio and Silvia Planas who taught me DNA extraction, PCR and

bioinformatics.

Special thanks to the following personnel of the Biology Department: Dr. Garcia-Arrarás, Millie

Viera, Aidamarie Perez, Diana Rosario, Gladys Ramos and Jose Fontánez for their excellent

work.

I would like to give special thanks to Dr. Raul Cano for support, guidance, and for giving me the

opportunity to work in other projects (amber bacteria, El Yunque and the coprolites) which have

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given me the opportunity to learn many cool things. Special thanks for teaching me lessons

which I know are very valuable as a scientist and as a person.

I would like to thank the Environmental Protection Agency, DEGI, the Biology graduate

program and the RISE program for financial support. Without this support, I would not have had

the opportunity to travel to present, network and learn new techniques.

To my brother Pedro Santiago and my aunts: Evelyn and Sonia Rodriguez, Johanna and Pichi

Vazquez, for their unconditional support. Most importantly, I would like to thank my mom,

Marisol Rodriguez and grandmother, Enriqueta Marrero, to who I owe this and at least 10 more

theses.

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CHAPTER 1

General Introduction

Clean water sources are vital for drinking and recreational purposes, but many can get

contaminated by fecal material. Direct contact with and ingestion of waters contaminated by

fecal material can affect millions of people every year as a result of the gastrointestinal and

respiratory illnesses and eye and skin infections caused by enteric pathogens. Among the

possible types of fecal contamination, that from a human source may represent a major concern

to our health given that human enteric pathogenic bacteria, protozoans and enteric viruses are

among those that can cause most of the mentioned diseases. Thus, the identification of the fecal

contamination in waters is necessary to eliminate the source and to minimize the potential risk

that enteric pathogens represent to public health. Other concerns with human enteric pathogens is

that these can cause illness with infectious doses as low as 1 to 10 [3, 4], can exhibit resistance to

removal and inactivation treatments (e.g. chlorination) [5-7] and can be reintroduced from

sediments to surface waters after disturbances (e.g. precipitation events) [8-11].

To remediate the source of the fecal contamination and avoid disease, enteric pathogens can be

detected using culture and/or molecular methods. However, culture methods may be expensive

and time-consuming, and not all enteric pathogens are cultivable. Molecular methods may

circumvent culturing microorganisms, and although current water quality guideline standards are

implementing molecular methods, several drawbacks have been identified, such as the detection

of noninfectious and unviable enteric pathogens. In addition, detection of human enteric

pathogens using culture or molecular methods may be relatively expensive, require the right

personnel and equipment. Moreover, given that many different human enteric pathogens can be

introduced into water sources, it would be impractical trying to detect them simultaneously [3].

In order to infer fecal contamination and the possible presence of enteric pathogens in water

sources, other enteric microorganisms (e.g. enterococci, thermotolerant coliforms and

bacteriophages) are used, and these are known as microbial indicators of fecal pollution [3, 12-

14]. These microorganisms should satisfy several characteristics in order to be considered

reliable indicators of fecal contamination: (i) be detected using simple laboratory methods, (ii) be

present in fecally polluted waters and absent in pristine waters, (iii) be associated with the source

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of the contamination, (iv) possess a survival similar to that of the pathogen of concern and (v) be

unable to replicate outside the host. However, as discussed below, currently used microbial

indicators fail to fulfill many of these characteristics and thus studies are currently being

conducted in order to identify and characterize novel markers of fecal pollution [7].

Bacterial and viral indicators

Total and thermotolerant coliforms

Total coliforms have long been used to assess the microbial quality of water sources since they

are present in the intestinal tract of animals. These bacteria can ferment lactose and produce gas

within 24 to 48 h at 35 ºC and include the genera Escherichia, Klebsiella, Enterobacter and

Citrobacter. Similarly, thermotolerant coliforms are part of the intestinal microbiota of warm-

blooded animals and fecal discharges contain large numbers of bacteria belonging to this group,

but unlike total coliforms, can grow at 45 °C [15]. Thermotolerant coliforms are currently being

used as an index of fecal contamination in many water sources; however, total and

thermotolerant coliforms may also originate from non-fecal sources such as run-off [16]. In

addition, some members of the coliform group can replicate in subtropical and tropical waters

and have been found to be part of the environmental microbiota of many of these water sources

[17, 18]. Those members that have been linked to fecal pollution possess survival times and

inactivation rates that cannot be correlated with those of many human enteric pathogens under

similar conditions. In addition, coliforms have been detected in the absence of enteric pathogens

and viceversa, and thus there is a lack of correlation between the number of these indicator

bacteria and enteric pathogens [13, 19].

Enterococci

Enterococci are gram-positive bacteria that include members of the genus Enterococcus, which

can grow at a wide temperature range (10 to 45 ºC), basic pH (9.6) and high salinities (6.5 %

NaCl). These have been accepted as indicators of fecal pollution since are commonly found in

the feces of warm blooded animals and their prevalence seems to be similar to that of many

bacterial pathogens [20]. However, confirmation methods are often needed when detecting

enterococci and these can be relatively laborious and results require are obtained within 24 h

[21]. Enterococci may not be used to track the source of the fecal contamination given that these

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are present in the intestinal tracts of different warm-blooded animals and interestingly, these

bacteria have also been detected in the intestines of pigeons and in flies [22, 23]. In addition,

enterococci have been detected in pristine waters and have been detected when pathogens are

absent and viceversa [13, 24].

Bacteriophages

Bacteriophages are viruses that infect bacteria and are present in the intestinal tract of warm-

blooded animals. Phages are very specific in terms of the bacteria they infect and source (i.e.

those isolated from the feces of humans have not been isolated from the feces of other animals)

[25]. They have been specifically proposed and used as indicators of the virological quality of

waters since the 1970’s [26]. Because of this source specificity, the use of phages as indicators of

fecal pollution arose. Other reasons for considering phages as indicators of fecal contamination

included the need to find models of human enteric viruses and a reliable method to assess the

virological quality of waters. The similar morphology, structure and “behavior” of

bacteriophages to that of many human enteric viruses, suggests that they should be more reliable

indicators of the virological quality of water sources than indicator bacteria. In addition, the

bacteriophage method possesses several advantages compared to the bacteriological methods: are

relatively less expensive, results are obtained in less time (4 to 6 h after incubation) and do not

require laborious confirmation methods [24, 27]. In addition, bacteriophages are as or more

resistant to removal and water disinfection processes compared to many enteric viruses [28-30].

Among the bacteriophages proposed as indicators of fecal contamination are those infecting

Bacteroides fragilis. Bacteroides fragilis phages have been linked to human fecal pollution [31-

33], since they are absent when enteric viruses are absent, do not replicate under environmental

conditions, seem to be more resistant to various water treatments than human enteric viruses and

have been proposed as a tool for viral waterborne disease control [34]. However, they have been

detected only at certain geographical regions of USA and Europe. In addition, the techniques

used to detect B. fragilis bacteriophages are difficult since the conditions must be anaerobic [35,

36].

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Table 1: Host specificity of different groups of F + RNA coliphages. Modified from [1].

Somatic and F (male)-specific coliphages have also been proposed and used as indicators of fecal

pollution [35, 37, 38]. Coliphages infect bacteria belonging to the Enterobacteriaceae family,

being E. coli the most widely host used for their detection. Somatic coliphages, like other

somatic phages, infect bacteria by attaching to specific surface receptors. In contrast, F(male)-

specific coliphages only infect conjugating bacteria by recognizing receptors in the pilli. F

(male)-specific coliphages possess genomes of + RNA or + DNA and belong to the Leviviridae

and Inoviridae family, respectively. F + RNA coliphages have been well characterized as

markers of fecal pollution and the possible sources, but less is known about F + DNA coliphages

[39]. However, the use of coliphages as indicators of specific sources of fecal contamination may

also have several disadvantages. The reason for this is that, even thought it has been suggested

that serotypes II and III of F (male)-specific coliphages could be used to infer human fecal

contamination [1, 40] (Table 1), other studies have found that the same serotypes can also be

found in animal feces [41]. In addition, coliphages are rarely detected in human feces, their

concentrations and survival cannot be correlated with that of many enteric viruses and certain

studies have found that somatic coliphages can replicate in the environment [35, 42].

Molecular techniques

Over the past years, molecular methods have been developed and tested to identify possible fecal

sources [43-45]. Currently developing methods generally involved the amplification of the

microbial indicator’s nucleic acids by using specific primers, although other methods have been

tested and will be discussed below. For some researchers, molecular methods offer the advantage

of obviating the need for cultivation. Culturing microorganisms is considered for some a

laborious procedure, which involves selective media and enrichment in order to isolate a specific

microbial indicator. Culturing microbial indicators may underestimate their numbers as some

Group Host

I Non-human animals.

II Humans and occasionally pigs.

III Exclusively human.

IV Non-human origin with rare human associations.

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may be unviable and uncultivable, and thus molecular methods may circumvent this. However,

molecular methods may not always be as ideal as expected. In this section, the advantages and

disadvantages of several molecular methods that have been used and are currently accepted to

infer fecal pollution are discussed.

Ribotyping and Pulsed-Field Gel Electrophoresis (PFGE)

Ribotyping consists of a fingerprint pattern resulting from differences in the size of DNA

fragments. Total genomic DNA is extracted from pure cultures and is treated with enzymes,

resulting in digested DNA fragments. The fragments are separated using agarose gel

electrophoresis and then transferred to nylon membranes. Southern blot hybridization is then

performed using rDNA probes, which results in a pattern composed of several bands. Various

restriction enzymes can be used in further analyses if one wants to increase the specificity of the

results [46]. The technique has been used to distinguish E. coli of human and non-human origins.

However, the technique is poor when analyzing samples containing multiple sources of fecal

contamination [47, 48]. Similarly, in the PFGE technique, pure bacterial cultures are placed in

agarose plugs and the DNA is digested using restriction enzymes. The digested plugs are

embedded into electrophoresis gels with alternating currents. Although PFGE has been used for

the characterization of environmental E. coli and enterococci, specialized equipment is

necessary, the technique may be time-consuming and the number of isolates which can be

processed simultaneously is limited [49].

Denaturing Gradient Gel Electrophoresis (DGGE) and Terminal-Restriction Fragment

Length Polymorphism (T-RFLP)

The DGGE is a technique able to differentiate between PCR products having similar sizes and

this is due to the melting properties of the DNA fragments and their mobility in the gel. DGGE

has been used to characterize fecal bacteria populations of both animals and humans. Yet, one

disadvantage of this technique is that the gene targeted must possess enough sequence variability

among the bacteria in order to identify the possible fecal source [50]. PCR products, like those of

the 16S rRNA gene, can also be detected using a DNA sequencer when fluorescently labeled, a

technique known as T-RFLP. The method is used to determine differences in the lengths of gene

fragments and offers the advantage of obviating culturing bacteria from the environmental

23

samples. It has been tested in the feces of farm animals and humans, but a more studies are

needed to determine if T-RFLP is suitable for other types of fecal contamination [51, 52].

PCR-based techniques

PCR-based methods have acquired great interest as these have shown to identify many sources of

fecal contamination. Moreover, water quality guidelines may incorporate PCR-based techniques

to routinely assess the microbial quality of waters since these are relatively simple and results are

obtained in less time compared to most culturing methods. Among the most accepted PCR-based

methods are the host-specific. Bernhard and Field developed PCR primers for the amplification

of conserved 16S rRNA gene sequences present in Bacteroides species specifically present in

cattle and humans. Detection of these genera possess several advantages: (i) in phylogenetic

analyses, members have shown to group according to the host, (ii) are restricted to warm-

blooded animals, (iii) are among the most numerous microorganisms in feces and (iv) possess

short survival times once introduced into water sources, thus replication in the environment is

not likely. There was the need to design PCR primers for the detection of Bacteroides because

these microorganisms are difficult to culture due to their strictly anaerobic nature. Interestingly,

differences were noted in the 16S rRNA gene sequences of bacteria present in human and cattle

fecal material and accordingly, primers targeting these sequences can be used to identify possible

sources of fecal contamination [53].

Studies by Bernhard and Field are among the pioneer studies about primer design for the

detection of host-specific bacteria. However, the presence/absence of indicator bacteria may not

reflect the extent of the fecal contamination. For this reason, most PCR-based techniques aim to

quantify the nucleic acids of indicator bacteria, being quantitave PCR (qPCR) one of the most

accepted and currently tested in human and animal fecal materials and water sources. qPCR

assays for the identification of human fecal contamination have shown to be specific and

sensitive, as in the case of the human Bacteroides markers HF-183 and BacHum-UCD. Markers

of animal fecal contamination, such as BacCow-UCD (cattle) and BacCan-UCD (dog) have

shown to be specific, but not necessarily sensitive since these amplify a fraction of the

corresponding fecal material and in some cases, cross reactions with horse fecal matter (as in the

case of BacCow-UCD) and human feces (as in the case of BacCan-UCD) can occur [54]. In

24

addition, it has been suggested that qPCR may overestimate the concentrations of indicator

bacteria in different water sources, and thus many studies have correlated qPCR with culture-

based methods.

Correlation results between qPCR and culture methods depend in the water type tested. For

instance, in temperate marine waters, positive correlations between the two methods have been

noticed [55]. In subtropical marine waters, the detection of E. coli using uidA and enterococci

using the 23S rRNA gene have positively correlated with their respective culture methods.

Interestingly, in subtropical climates, the strength of the correlations also depends in the sample

site and the time at which the samples are collected. Accordingly, a stronger correlation between

qPCR and culture methods for enterococci are noticed when samples are collected in the

morning [56]. In the present thesis, the correlation between qPCR and culture methods for the

detection of enterococci in tropical inland waters is presented in Chapter 4. Correlations

between molecular and culture-based methods may suggest that either technique could be used.

This represents, for some researchers, an opportunity to substitute culturing microorganisms with

molecular techniques. Results are often obtained in less time and thus immediate actions can be

taken to remediate the source of the fecal contamination and minimize the possible risks to

public health. However, for some researchers, correlations between both molecular and culture

techniques may represent a disadvantage. The reason for this is that it has been suggested that a

toolbox of methods may be more appropriate to infer fecal pollution.

Correlations of indicators with pathogens and illness

Ideally, the characterization of indicators of fecal pollution should include correlation analyses

with enteric pathogens. Studies of this type are limited since not all laboratories possess the

facilities to detect enteric pathogens [57]. The presence of specific microbial indicators has

positively correlated with the presence of bacterial pathogens, as in the case of total coliforms

and Clostridium perfringes with Salmonella, but in other cases, coliforms do not correlate with

enteric pathogens [58, 59]. Possible reasons for this include differences in the prevalence and

survival times, and the ability of coliforms to become part of the environmental microbiota of

waters [60]. In terms of the enterococci, their prevalence has positively correlated with human

enteric viruses [61]. Other studies, however, have found no correlation between enterococci

25

detected by molecular or culture methods and Bacteroides spp. with Campylobacter spp. (a

pathogenic bacteria that causes gastroenteritis) [62]. Differences in the correlation analyses

between indicators and pathogens may be due to the water types tested, as aquatic ecosystems

may be differently impacted by ecological factors (e.g. salinity, transport of microbes from other

sites, turbidity and rainfall). Other influential variables may include differences in the type and

numbers of indicators and pathogens (as the latter are often detected in lower concentrations),

pathogen source, sample size and statistical methods. Statistical methods may have a great

influence on the results as these depend on the sample size and recent studies have suggested that

discrepancies between correlations between indicators and pathogens may be due to the

insufficient data for assessing these correlations [63]. It has also been suggested that microbial

indicators of fecal pollution may not necessarily infer the presence of pathogens, rather, it is a

probability of their co-occurrence [64].

Correlations studies also involve the epidemiological aspects of the health effects associated with

microbial indicators of fecal pollution. These studies aim to determine symptoms associated with

gastrointestinal and respiratory illness and ear, eye and skin infections after swimming in

possibly fecally contaminated waters and the correlation with the presence of microbial

indicators. Reports often involve threshold values of indicator-bacteria and symptoms associated

with the mentioned illnesses and how variations in the severity of the health effects correlate

with the extent of the fecal contamination [65]. Most of the positive correlations are observed

between gastrointestinal illness and enterococci, thermotolerant coliforms and E. coli and

interestingly, most of the thresholds are lower to those of current water quality guidelines [66].

Interestingly, other studies have reported a correlation between enterococci and skin illness [67],

but these variations may be due to differences in the experimental designs (e.g. the indicators

tested, participants and water type).

ANTIBIOTIC-RESISTANCE AND VIRULENCE GENES IN ENTEROCOCCI

Fecal contamination of water sources can also result in the introduction of bacteria harboring

antibiotic-resistance and virulence genes, but the possible risks to public health remain largely

unknown [68-70]. Specific antibiotic resistance phenotypes have been proposed as a tool for

microbial source tracking (MST) [71]. This is based on the hypothesis that bacteria exposed to

26

antibiotics will develop resistance and this selective pressure can be a mechanism for

discriminating fecal microorganisms from different sources. The potential problem when using

antibiotic resistance as a tool for MST is that bacteria can transfer the resistance genes to other

bacteria as these can be found on a variety of mobile genetic elements (e.g. plasmids and

transposons) [72-74]. Similarly, virulence factors have also been proposed as a means to infer

specific sources of fecal pollution. Virulence factors are ways that bacteria have developed to

circumvent the host defenses. However, as discussed below, many of these studies focus on the

presence/absence of the genes conferring virulence and most ignore the ecology of the

microorganisms harboring these genes.

One of the most studied bacteria harboring antibiotic-resistance and virulence genes is

Enterococcus spp., which can carry genes encoding for resistance to tetracycline and

vancomycin, a last resource drug, and the enterococcal surface protein (Esp), a virulence factor

[75, 76]. Tetracycline and vancomycin have been prescribed to treat infections caused by

Enterococcus [77], but the excessive use of these antibiotics has lead to an increase in

Enterococcus isolates exhibiting resistance to both antibiotics. Many of these antibiotic-resistant

enterococci can also harbor Esp, which enables bacteria to form biofilms, often difficult to treat

with antibiotics. In the next sections, the molecular aspects of tetracycline and vancomycin-

resistance genes and esp are discussed. Chapter 6 presents the prevalence of these gene in the

environment.

Tetracycline mode of action and resistance genes

Tetracyclines are among the wide-spectrum antibiotics commonly used for treating infections.

These belong to a family of antibiotics which inhibit protein synthesis by binding to the

ribosomal acceptor site (A-site) and thus inhibit the association of the tRNA with the bacterial

ribosome. Tetracycline-resistant bacteria can carry at least one of the 38 known tetracycline-

resistant genes. Most of these genes encode for the formation of efflux pumps, which are

membrane-associated proteins that move tetracycline out of the bacterial cell. Genes encoding

for efflux pumps can be found in both gram-positive and gram-negative bacteria. This efflux

activity in bacteria is the result of the expression of two genes: one that encodes for the

membrane-associated protein and another that encodes for a repressor protein. In the absence of

27

tetracyclines, the repressor protein inhibits the expression of the gene encoding for the formation

of the efflux pumps. Other tetracycline resistance genes encode for ribosomal protection

proteins, which attach for a short time to the tetracycline-binding site, inhibiting the association

of tetracycline with the A-site. Other data have suggested that ribosomal protection proteins

bind to the ribosomes and change their conformation [75, 76, 78]. Tetracycline resistance genes

also include those encoding for inactivating enzymes, which inactivate tetracycline (tet(X)) and

tet(U), whose function remains unknown (Table 2).

Table 2: Classification of most of the tet determinants according to their mode of action.

Modified from [75].

Prevalence of tetracycline-resistance genes has been determined in the feces of humans and

animals, clinical settings and waters in contact with swine-production facilities [79, 80]. These

have been identified in mobile elements, such as plasmids and transposons and thus their lateral

transmission has been reported, even between bacteria of different genera [81-83]. Most of the

tetracycline-resistance genes have been identified in gram-negative bacteria, but others are

present in gram-positive bacteria (e.g. tet(M), tet(K), tet(L)) and are not structurally similar to

those of gram-negative bacteria. Among the tetracycline-resistance genes found in gram-positive

bacteria, tet(M) is the most disperse and is found in many Enterococcus spp. (E. faecalis, E.

faecium and E. gallinarum) and has received greater attention [75, 83, 84]. For this reason, its

prevalence has been determined in clinical settings, sewage and environmental settings [79, 80,

85]

Efflux proteins Ribosomal protection Inactivating enzymes Unknown

tet(A), tet(B), tet(C),

tet(D), tet(E), tet(G),

tet(H), tet(J), tet(L),

tet(V), tet(Y), tet(Z)

tet(M), tet(O), tet(S),

tet(W), tet(Q), tet(T)

tet(X) tet(U)

28

Vancomycin mode of action and resistance-genes

Vancomycin is used to treat infections caused by gram-positive bacteria and it is often used to

treat Enterococcus infections exhibiting resistance to other antibiotics. It is, therefore, a last-

resource drug as it possesses hazardous side effects. It acts by inhibiting the synthesis of the cell

wall and thus bacterial replication. Briefly, vancomycin binds to the ᴅ-Ala-ᴅ-Ala moieties of the

N-acetylmuramic acid (NAM) and N-acetylglucosamine (NAG) peptides, preventing the

backbone polymers to form, resulting in the destabilization of the cell wall (Figure 1). However,

the use of this antibiotic has resulted in the isolation of vancomycin-resistant enterococci from

clinical settings and from the environment to a lesser extent [78, 86-88].

Figure 1: General mode of action of vancomycin. Vancomycin attacks the ᴅ-Ala-ᴅ-Ala moieties

of the cell wall, causing instability and cell death.

Five vancomycin-resistance phenotypes have been identified in enterococci so far: VanA, VanB,

VanC, VanD and VanD. Vancomycin-resistant enterococci are classified according to the

exhibited resistance to various concentrations of vancomycin and teicoplanin, an antibiotic

mainly used in Europe. Bacteria exhibiting the VanA phenotype are resistant to a wide

concentration of vancomycin and teicoplanin. The VanB phenotype is characterized by bacteria

showing resistance to different concentrations of vancomycin, but unlike the VanA phenotype,

bacteria exhibiting the VanB phenotype, are resistant to lower levels of teicoplanin. Enterococci

exhibiting the VanC phenotype are resistant to lower levels of vancomycin compared to the

VanA and VanB phenotypes. The VanD phenotype is characterized by bacteria resistant to

intermediate levels of vancomycin and has been described in E. faecium. The VanE phenotype is

characterized by E. faecalis strains exhibiting resistance to lower doses of vancomycin and

teicoplanin compared to the VanA phenotype (Table 3).

NAM/NAG

peptides

D-Ala-D-Ala

Moieties

Vancomycin

29

Among the vancomycin-resistance phenotypes in enterococci, the VanA and VanB have received

greater attention. These phenotypes are the result of the expression of operons which include

vanA or vanB, vanRAB, vanSAB, vanHAB, vanXAB and vanZAB. VanA and VanB are ligases which

produce ᴅ-Ala-ᴅ-Lac instead of ᴅ-Ala-ᴅ-Ala. VanH is a ᴅ-hydroxy acid dehydrogenase which

creates a pool of ᴅ-lactate for use in the previous reaction. VanX is a ᴅ, ᴅ-dipeptidase lacking

activity against ᴅ-Ala-ᴅ-Lac and reduces the availability of ᴅ-Ala- ᴅ-Ala produced by the

enterococcal ligase, thereby minimizing the competing synthesis of normal pentapeptide [89].

Among the genes discussed, those encoding for the ligases, vanA and vanB, are considered when

detecting specific vancomycin-resistance phenotypes and may be present in the bacterial

chromosome, plasmids or transposons [89]. The prevalence of vancomycin-resistant genes have

been mainly determined in hospitals, sewage and in aquatic ecosystems [90, 91].

Characteristic Phenotype

VanA VanB VanC VanD VanE

Vancomycin MIC

(mg/mL)

64->1000 4-1024 2-32 128 16

Teicoplanin MIC

(mg/mL)

16-512 < 0.5 < 0.5 4 0.5

Most frequent

enterococcal species

E. faecalis, E.

faecium

E. faecalis, E.

faecium

E. gallinarum, E. casseliflavus,

E. flavescens

E.

faecium

E.

faecalis

Table 3: Vancomycin-resistance in enterococci. Five phenotypes have been identified so far and

enterococci are classified according to the levels of vancomycin and teicoplanin it exhibit

resistant. Modified from [89].

Enterococcal surface protein

Enterococci exhibiting resistance to antibiotics may also harbor virulence factors [92]. Virulence

factors are mechanisms that bacteria have evolved to avoid the host defenses and thus can result

in pathogenicity. Among the virulence factors present in enterococci is the enterococcal surface

protein, encoded by esp. This gene has acquired great attention since nosocomial infections,

including urinary tract infections, endocarditis and bacteremia, are caused by enterococci

harboring esp [93, 94]. The major problem with enterococci harboring esp is that these can form

30

aggregates or biofilms which exhibit resistance to antimicrobial treatments [95]. The gene was

first discovered in E. faecalis strains, but a variant of the gene was later discovered in E. faecium.

Further studies detected the esp variant present in E. faecalis in both human and animal feces,

and the E. faecium variant was detected uniquely in human fecal material [96]. This resulted in

the characterization of esp as a tool for MST purposes.

Conflicting MST studies have tried to determine the specificity of esp. In these studies, PCR

primers were designed based on available sequences, and their ability to amplify enterococci

DNA in fecal matter, sewage and surface waters impacted by fecal contamination. Many studies

have been successful in the amplification of the gene from human feces and these same studies

have determined the absence of the gene in animal fecal material [97, 98]. However, other

studies have amplified the gene in environmental enterococci and thus the specificity of the gene

has been questioned. This is the case of studies performed by Byappanahalli and colleagues,

which have detected the gene in enterococci isolated from sources with no apparent input of

fecal material. The same studies have detected esp in Cladophora, one main source of

enterococci in water sources. In addition, environmental factor can affect the detection of esp,

including rainfall. It has been found that the prevalence of esp is higher after precipitation events

[99].

HORIZONTAL TRANSFER OF ANTIBIOTIC-RESISTANCE AND VIRULENCE

GENES

It is well known that tetracycline and vancomycin resistance genes are present in mobile

elements, and can be transferred between bacteria [76]. These genes can also be present in the

bacterial chromosome, and this is also the case of esp, but few studies have determined the

transferability of esp to other bacterial species [94, 100]. Among the possible vehicles mediating

the transmission of antibiotic-resistance and virulence genes, bacteriophages are often not

considered. Phages can harbor genes which are not indispensable for their “life cycles”, but

could be transferred to the recipient bacteria and result in an increased fitness. Such is the case of

the lateral transmission of antibiotic-resistance and virulence genes mediated by temperate

phages, but few studies are still available [101]. Lysogenic phages may have a greater impact in

the evolution of bacteria than what is believed.

31

Phages are classified as strictly virulent (lytic) or temperate (lysogenic). Lytic bacteriophages

lyse their host bacterium, within a couple of minutes to hours after infection, by producing

hundreds or thousands of phages. On the other hand, a temperate phage may not lyse its host

immediately. Rather, it integrates its genome into the bacterial chromosome, until environmental

conditions are unfavorable, triggering the expression of the phage lytic genes [102, 103]. The

state of a phage to remain “dormant” in a bacterial chromosome is known as lysogeny, and it is

governed by the expression or repression of the phage genes. Specifically, the protein known as

CI, which is codified by the cI gene is involved in the repression of the expression of the phage

genes involved in the lytic cycle. Expression of the cI gene is in turn regulated by the CII and

CIII proteins, encoded by the cII and cIII genes. It is possible that CI-like proteins in E. faecalis-

infecting phages possess the same function as those found in the λ bacteriophage. Lysogenic

phages also harbor anti-repressor genes, which encode for proteins that interfere with the

function of repressors, promoting the expression of the phage genes and thus inducing the lytic

cycle [104].

The lysogeny state in λ and lysogenic E. faecalis phages is governed by a module of genes which

include those encoding for integrases [104]. Integrases are responsible for the integration of the

phage genome into the bacterial genome and this is due by the recognition of specific nucleotide

sequences in both the phage and bacterial genome (Figure 2). The recognition sites in the phage

and bacterial genomes are known as the attP and attB sites, respectively [2, 105].

Figure 2: Integration of a bacteriophage genome into a bacterial genome. Integrases recognize

specific nucleotide sequences in the phage and bacterial genomes (attP and attB sites,

respectively). Modified from [2].

32

THESIS DIRECTION AND GENERAL OBJECTIVES

Novel microbial indicator of human fecal pollution

Given that no microbial indicator of human fecal pollution satisfies most of the characteristics

mentioned previously, there is a need of identifying and characterizing novel indicators of this

type of contamination. Therefore, we have proposed a group of phages that infect a specific type

strain of Enterococcus faecalis, which we call enterophages, as indicators of human fecal

pollution. The present project presents data of the characterization of enterophages as markers of

human fecal pollution. Although results are promising, there is still the need of testing

enterophages in other geographical areas as only domestic wastewaters, marine and fresh waters

in Puerto Rico and domestic wastewaters in Portugal have been tested.

Enterophages as vectors of antibiotic resistance and virulence genes

Since tetracycline and vancomycin-resistance and esp are present in enterococci, it is fair to

believe that lysogenic phages infecting Enterococcus possess genes similar to those found in the

bacterial hosts. This has been previously found in coliphages, which possess a module of genes

known as R factors. R factors can be transferred and confer tetracycline and streptomycin-

resistance to the host bacteria [106]. Other studies have found that the viral DNA fraction of

sewage and environmental samples harbor methicillin-resistance [107]. Similarly, certain

coliphages can harbor Shiga toxins-encoding genes, a virulence factor, which can be transferred

to specific E. coli strains and cause serious risks to health [108-110]. It was unknown if phages

infecting Enterococcus harbor the mentioned genes, and although none were detected in

Enterococcus phages or the viral DNA fractions of environmental samples, tetracycline and

vancomycin resistance genes, and esp, were detected in the bacterial fractions and enterococci

isolates. However, it should not be ignored that Enterococcus phages can harbor other antibiotic-

resistance and virulence genes, and thus future studies are still needed.

It was also unknown if enterococci phage isolates harbored integrase-encoding genes. One

importance of this gene is that it enables one to determine if a phage is temperate and thus

amplification of this gene in Enterococcus-infecting bacteriophages could be used as a marker of

lysogeny [104]. This in turn represents an opportunity to focus on those phages harboring

integrases to further detect antibiotic-resistance and virulence-encoding genes. Another

33

importance of detecting integrases, although this hypothesis needs to be further tested, is that

integrases may be used to determine the presence of mobile elements in samples and therefore,

by determining the prevalence of microorganisms harboring integrases in the environment, future

studies could determine the possible increased risks to public health as a result of lateral gene

transfer in the environment.

Finally, the complete genomes of two Enterococcus phages are currently being determined: one

strictly lytic to E. faecalis strain ATCC 19433 and the other was induced from an environmental

enterococci isolate. Sequencing of these two phage types will enable us to describe the genomes

of novel Enterococcus phages and may open the opportunity to further develop the molecular

techniques for their detection in sewage and fecally contaminated waters.

34

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44

CHAPTER 2

Enterophages, a group of phages infecting Enterococcus faecalis and their potential as

alternate indicators of human fecal contamination

N. Bonilla1, T. M. Santiago-Rodriguez

1, P.

Marcos

2, M. Urdaneta

1, J. W. Santo Domingo

3 and G.

A. Toranzos1

1Environmental Microbiology Laboratory, Department of Biology, University of Puerto Rico, P.O.

Box 23360, San Juan, PR 00931-3360, 2Biological Sciences Department, General Studies,

University of Puerto Rico, San Juan, PR 00931, 3U.S. Environmental Protection Agency,

Cincinnati, Ohio

ABSTRACT

We have developed a method for the detection of viruses in environmental samples that we have

called enterophages, which specifically infect Enterococcus faecalis. This method has allowed us

to determine the prevalence and to study the ecology of this group of phages. The enterophages

replicate at 35ºC, and at 41ºC. The presence of NaN3 in the media inhibits the growth of

background microbiota and allow for an accurate, specific and rapid detection of these viruses.

Enterophages were present in raw domestic sewage at lower concentrations (average 1.8 x 102

PFU/100ml) than those of coliphages (average 1.7 x 105

PFU/100ml). Phages were characterized

by transmission electron microscopy showing icosahedral capsids, some with non-contractile

tails as well as icosahedral non-tailed capsids. Different isolates had capsid sizes ranging from 20

nm to about 75 nm in diameter. These data describe a new group of phages that may serve as

alternate indicators of human fecal pollution, especially in recreational waters. The ecology of

these enterophages indicates that these may be strictly of human origin.

Keywords: enterococci; enterophages; faecal contamination; indicators; recreational waters

Reference: Bonilla, N., et al., Enterophages, a group of phages infecting Enterococcus faecalis,

and their potential as alternate indicators of human faecal contamination. Water Sci Technol,

2010. 61(2): p. 293-300.

45

INTRODUCTION

Pathogenic microorganisms are present in recreational waters and other water sources as a result

of the presence of feces of warm-blooded animal. Some examples of these pathogens are

Salmonella, Shigella, Campylobacter spp., as well as several different types of enteric viruses

and pathogenic protozoa. The low infectious dose (as low as 1 to 50 in some cases) of many of

these pathogens is a cause for concern if they are released into the water environment, since

humans who are in contact with these contaminated waters, are bound to ingest the contaminated

water. Human rotaviruses, astroviruses and noroviruses viruses may be found in sewage-

contaminated waters and in fact, among the waterborne pathogens, these are of greatest concern,

since they may be more resistant to environmental conditions than pathogenic bacteria and they

may be more resistant to chlorination and UV disinfection [1], and have perhaps the lowest

infectious dose of any of the waterborne pathogens.

Indicator organisms have been used for the last century as surrogates for the presence of

pathogenic microorganisms in waters. In the past, bacterial indicators such as the thermotolerant

coliform group, Escherichia coli or the Enterococcus group to determine the microbiological

safety of waters have been tremendously useful. The levels of these indicators are monitored in

treated drinking waters as well as recreational waters and the concentrations at which they are

found enables (e.g.) beach managers in making decisions that involve closing beaches or keeping

them open to the public.

Escherichia coli and Enterococcus spp. have been used as indicators of the presence of

pathogenic microorganisms in different types of water. High concentrations of these indicators

have previously been found to correlate with risk to bathers; concentrations of E. coli have been

correlated with risk to bathers in freshwater beaches [2, 3], whereas concentrations of

Enterococcus spp. have been correlated with risk in marine beaches. As such, they are now

considered the most useful indicators of risk to bathers as a result of primary contact exposure to

bathing waters. However, bacterial indicators (E. coli and Enterococcus spp.) have been isolated

from pristine sites in a tropical rain forest [4] and are routinely detected as part of the

environmental microbiota in Puerto Rico and other tropical areas. Fecal indicators such as E. coli

and enterococci may also be able to multiply and become a significant environmental source in

46

tropical regions such as Hawaii, Guam, Puerto Rico and South Florida [5]. Thus their role as

microbial indicators of fecal contamination is not entirely clear and alternative indicators of

waterborne enteric viruses are needed.

Alternate microbial indicators of fecal water contamination

Thermotolerant and total coliforms tend to be poor indicator microorganisms because of their

short survival and susceptibility to water treatments [1]. Moreover, these microbial indicators

have the ability to grow in natural waters and there is a lack of correlation between the number of

coliforms and that of infectious microorganisms [6]. Also, pathogenic viruses have been found in

waters where the number of coliforms had not exceeded the standards [7].

Bacteriophages, especially coliphages, have also been used as model viruses for water quality

control [8]. Coliphages have been proposed as indicators of fecal water contamination [9-12], as

have those phages that infect Bacteroides fragilis [13], since many have similar morphological

characteristics to the enteric viruses as well as similar survival times in aquatic environments.

However, their usefulness has been questioned, since the group is made up of different viruses

belonging to several different families, each having different ecological and biological

characteristics. A more specific group seems to be the male-specific RNA coliphages, but their

usefulness is still being debated.

Out of all methods available for the determination of the microbiological water quality, the

bacteriophage methods take the least amount of time, and are the most reliable methods for the

detection of the target since they may not require confirmation procedures. Additionally, they

may be amongst the least costly methods once they are implemented in an environmental

monitoring laboratory.

A useful bacterial indicator is Enterococcus faecalis, because of its apparent inability to multiply

in sewage contaminated waters and their survival time in the environment seems to be greater

than E. coli [14]. E. faecalis shows a closer correlation with risk to bathers in point-source

impacted recreational waters [2]; however, we know very little about the ecology of this group of

47

microorganisms. Although E. faecalis has been described as having a limited host range, the

genus may be more cosmopolitan than previously thought. The usual hosts include humans,

chickens and dogs [3], but they are also present in the intestines of other animals, such as

pigeons [15] and have also been found as part of the microbiota of insects, including flies [16-

18]. In Puerto Rico we have detected Enterococcus spp. as part of the microbiota in sand flies,

and even in laboratory strains of Drosophila spp. (in preparation).

The detection of pathogenic microorganisms or reliable indicator microorganisms is important

for the determination of the microbiological quality of recreational waters. However, currently

used detection methods have several disadvantages: they can be expensive or the detection time

can take too long, taking on the average 24 to 48 hours before results can be obtained. This is

certainly not the best manner to manage possible risk in recreational waters. Additionally,

bacterial indicators are not reliable indicators of the possible presence of enteric viruses. We

need a method that will give reliable results in less than 12 to 18 hours, and the only non-

molecular method that seems to lend itself for this time interval is one that detects infectious

bacteriophages.

Molecular methods are fast and reliable, but dead bacteria, naked DNA or non-infectious viruses

may be detected, so the question of viability (in the case of bacteria) and infectivity (in the case

of viruses) come into play. Thus, the detection of infectious bacteriophages, when using the

plaque assay, if they can be correlated to risk, would perhaps be the most appropriate manner of

managing risk in recreational waters.

In the present paper we describe a method we developed for the detection of a group of phages

that infect specifically E. faecalis. Our goal is to eventually determine if this group of viruses can

be used as a reliable alternative indicator of risk to bathers as a result of the presence of human

pathogens in recreational waters.

MATERIALS AND METHODS

Host strains

48

Several isolates were obtained from raw domestic sewage from several sewage treatment plants

in Puerto Rico. These were obtained using membrane filtration using mEnterococcus media

prepared as indicated by the manufacturer. Isolated colonies were purified and isolated in pure

culture using the same media. Over 100 individual isolates were tested using the single and

double layer methods [19] for the detection of phages. Additionally, several Enterococcus spp.

type strains were tested as possible hosts, these were: E. pseudoavium, E. faecium, E. durans, E.

casseli, E. hirae, E. dispar, E. gallinarum and E. faecalis.

Host bacteria

Once the best host was determined to be E. faecalis, it was kept streaked for isolation in

mEnterococcus agar plates and incubated overnight at 35 ºC. The Petri dishes were kept at 4-7 ºC

for up to one week. To prepare the host bacteria for plaque assays, individual colonies were

picked and inoculated into sterile (Difco) Dextrose Azide Broth medium and incubated overnight

at 35 ºC. Five-mL of this inoculated medium was used for every 50 mL of sample added to

50mL of 1X regular-strength (Difco) Tryptic Soy Broth (TSB).

Samples

Sewage and recreational water samples were collected in sterile plastic bottles. Samples were

shipped to the laboratory at 2 to 8ºC using cold packs and stored at 4 ºC. Additionally, bird, and

several mammal fresh fecal samples were tested for the presence of enterophages.

Optimal conditions for viral replication

Several concentrations of calcium chloride (CaCl2·2H20 Fisher Scientific Co., NJ, USA) and

sodium azide (NaN3 MCB, OH) were tested, using as the base those concentrations used by other

authors [19] for the detection of coliphages (in the case of calcium chloride) and commercially

available mEnterococcus medium (in the case of sodium azide).

To determine the optimal concentration of medium, several media were tested, such as: Nutrient

Broth, Azide Dextrose and mEnterocuccus, at different concentrations. Similarly, 2X

concentration of TSB and 1.5 % agar was tested as was 1X TSB containing different

concentrations of agar. Agar concentrations ranging from 0.375 to 1.5 % were tested. Phage

49

plates were incubated at 22, 37, 41 and 45°C. Viral plaques were counted at 6, 18, 24 and 48

hours.

Single Layer Plaque Assays

Once optimal conditions were determined for enterophage plaque formation, a 50 mL volume of

1X TSB was prepared and agar added to a final concentration of 0.375 or 0. 75 % where mixed

with an equal volume of the sample. The medium was autoclaved and kept at 50 ºC. To a 50

mL-volume of sample, CaCl2 was added to a final concentration of 5.2 mg/mL, as was NaN3 to a

final concentration of 0.4 mg/mL plus 5ml of a freshly grown culture (overnight culture) of E.

faecalis. Solution was mixed carefully with the liquefied 50 mL volume of 1X TSB, and poured

into four sterile 100-mm diameter Petri dishes. After the agar solidified, the Petri dishes were

incubated at 22, 35 or 41 °C, in order to determine if different phage populations that replicate at

different temperatures are present. Viral plaques were counted after 3, 8, 24 and 48 hours of

incubation.

Plaque isolation

Discrete viral plaques were plucked using standard procedures. Briefly, sterile glass Pasteur

pipettes were used to obtain a plug which was then placed in an Eppendorf tube containing 500

μL of sterile 1X Phosphate Buffer Solution (PBS). Many of these phages are tailed and rough

treatment may break the tails rendering the viruses non-infective. The plug was gently dislodged

and broken by pipetting up and down. The tubes were centrifuged at 14,000 rpm for 10 min at 10

ºC. The supernatant was transferred into a sterile Eppendorf tube, and the supernatant tittered

using serial dilutions using the double layer method using the same media as above. Briefly, the

dilutions were mixed with 4 mL of 1X TSB containing the same concentrations of CaCl2 and

NaN3 as above and 0.75 % agar and then poured into Petri dishes containing 20 mL of bottom

agar media containing CaCl2 and NaN3 and 1.5 % agar. Plates were incubated at 35ºC and 5mL

of PBS or physiological saline (0.85 % NaCl) was added to those plates showing complete viral

lysis and slowly agitated rotationally for 10 min. The top agar was transferred to a sterile

Oakridge tube, pipetted up and down to break down the agar, and then centrifuged at 14,000 rpm

for 10 min at 7 ºC. The supernatant containing the viruses were kept in a sterile tube at 4-7 °C.

50

Transmission Electron Microscopy

The resulting supernatant from above was loaded into dialysis tubes (12,000-14,000 MW cutoff,

Spectrapor, Los Angeles, CA, USA) for hydroextraction and covered with crystalline

polyethylene glycol (PEG, Mol.wt. 8,000,Sigma Chem. Co. MO; APHA, 1989) clamped and

placed at 4-7 °C overnight. The hydroextracted solution was recovered, the inside of the dialysis

tubing washed with 100 μL of sterile 0.85 % NaCl transferred into a sterile Eppendorf tube and

kept at 4 °C till electron-microscopic analyses.

An aliquot of the concentrated portion was placed on carbon Type-B 200 mesh copper grids or

ultra thin carbon film/holey carbon 400 mesh copper grids and stained with uranyl acetate (UA)

2%, pH 4.5 or potassium phosphotungstate (PTA) 2%, pH 7.2. Alternatively, 30 μL of each

phage concentrate was placed on parafilm and the grids floated on top for 15 minutes at room

temperature prior to staining. All specimens were examined using a Karl Zeiss Leo 922 energy

filtered transmission electron microscope operated at 200 KV. At least 5 phage particles of each

type observed were measured. The phages were measured directly on the images, which had the

magnifications previously calibrated. Capsid sizes were measured between opposite apices.

Determination of Burst Sizes

To determine the burst sizes of each of phage isolate, E. faecalis was grown to mid-log phase in

Azide Dextrose Broth and the concentration determined. One-mL of the proper enterophage

dilution was added in order to have a Multiplicity of Infection (MOI) of 1.0. The phage was

allowed to come in contact with the bacteria by incubating at 35 °C for 5 minutes. The solution

was then centrifuged at 14,000rpm at 10°C for 10 minutes. The supernatant was eliminated, and

10 mL of Azide Dextrose were added to the tube and the pellet resuspended to the original

volume. Aliquots were obtained every 15 min for 3 hours, and the aliquots serially diluted with

1X TSB containing the previously indicated concentrations of NaN3 and CaCl2. A volume of the

resulting dilution was transferred into 4 mL of top agar (0.375 %) and poured into agar plates

containing the bottom agar described above. The plates were incubated at 35 °C for 24 hours.

Plaques were counted and burst sizes determined using the point in the graph where the values

remained constant.

51

Survival

A sewage sample containing about 103 enterophages/100ml was inoculated into 1L of unsterile

sea water. Samples were incubated at 23, 35 and 41 ºC and aliquots obtained twice of

thrice/week and analyzed for the presence of enterophages using the single-layer method as

described above.

Prevalence

Grab samples were obtained from the influent sites at several different domestic sewage

treatment plants in Puerto Rico. Samples were kept refrigerated till processing. Fifty-mL

volumes were analyzed as described above using the single-layer method.

RESULTS AND DISCUSSION

Sewage and surface water grab samples were originally analyzed using commercially available

enterococci media (mEnterococcus, mE, mEI, Azide Dextrose), which did not result in any

visible plaques using either the single or double layer method. This was surprising, since the

media mention are standard for the isolation and detection of Enterococcus spp., but none were

useful for the detection of viral plaques (data not shown). We also tested different concentrations

of the TSB, and though we got some plaques, the growth of the host was not very conducive to

the detection of viral plaques. When testing different concentrations of the media, we started out

with 2X concentrations, since that is the standard procedure for the detection of coliphages;

however, we were surprised to see that E. faecalis seems to be susceptible to the osmotic

pressures created by even the final 1X resulting concentration after mixing with an equal volume

of the sample. This phenomenon was even more obvious when testing marine waters, possibly as

a result of the increased osmotic pressure; this opens up other questions as to the survivability of

Enterococcus spp. in marine waters. Thus we decided to use a final concentration of 0.5 % of

TSB. Though viral plaques were visible when using only 0.5 % TSB under laboratory

conditions, when processing environmental samples, it was impossible to see any plaques in the

absence of NaN3. Of several concentrations of CaCl2, the optimal concentration was shown to be

5.2 mg/mL and the optimal concentration of sodium azide 0.4 mg/mL. Viral plaques were visible

at 0.75 % agar at 37 °C, but somehow clearer at 0.375 % agar at 41 °C (data not shown).

52

We tested fecal samples from several different animals, including birds, dogs and cats as well as

sand flies for the presence of enterophages and none were positive; though all of them were

positive for the presence of Enterococcus spp. (data not shown)

Viral Morphology

The morphology of the virions (Figure 1) show tailed phages and they are different from the

round capsid morphology observed by others [20] in enterophages isolated from human saliva;

though we also detected some isolates that were non-tailed, with icosahedral capsids about 20-25

nm in diameter (data not shown). Our best studied phage isolate to date showed an icosahedral

75 nm diameter capsid with a long non-contractile tail measuring 240 nm long. This morphology

is similar to the classical Bradley’s basic morphology Group B belonging to the Siphoviridae

family [21]. Further studies are needed to characterize other groups of enterophage isolates. It

should be noted that viruses infecting E. faecalis have been described previously, but not those

from environmental samples. Previous isolation and use of these phages was for typing of E.

faecalis clinical isolates [22] going back several decades. The fact that enterophages are also

found in human saliva [20] may indicate that they are in fact unique to humans, and as humans

usually spit when they immerse their heads into the water while swimming, the concentrations of

these enterophages may show to be a good indicator of risk to bathers.

(A) (B)

Figure 1: Enterophage (A) with an icosahedral head of 13 nm and a 56 nm non-contractile tail

(Bar = 20 nm). Enterophage group (B) with an icosahedral head of 75 nm with a non- contractile

240 nm length (Bar = 200 nm).

53

Burst Sizes

When viruses successfully infect cells, results in a certain number of progeny virions called the

burst size. It would seem apparent that the phages most likely to be found in sewage and fecally-

contaminated waters are those that have a high burst size. Our data indicated that the burst size

varies depending on the isolate and the numbers go from 102 to 10

5 virions/cell (data not shown).

We will focus our future characterization efforts on those enterophages that have the highest

burst size, since they may be more likely to be detected in low volumes of water samples.

Survival

Any indicator of risk should be at least as reliable as those currently in use and should meet

several criteria, and among these the ability to survive for at least as long as the pathogens under

the environmental conditions is a must. Our data (Figure 2) show that there may be different

populations of enterophages present in raw sewage, as indicated by the different numbers

obtained in these experiments in plates incubated at three different temperatures. However, all

three populations survived similarly and their numbers decreased by over two log10 over a

period of 7 days, which is similar to the survival times of enteric viruses as reported by others

[23]. Figure 2 (A) shows that although the enterophages survived better at ambient temperature

where they showed about a 0.5 log10 drop after 3 days, whereas there was at least 1-1.5 log10 drop

in concentrations at 37 ºC after the same length of time. It should be noted that there was no

replication of the enterophages at ambient temperature in spite of having native E. faecalis

present in the sewage inoculum which would have been shown by an increase in the number of

viruses. It is also noteworthy to see different groups of enterophages capable of replicating at

different temperatures (namely 22, 37 and 41 ºC). These data will guide future studies as to the

types of enterophages to focus on as the possible best indicators of human fecal contamination.

We are in the process of carrying out similar survival experiments in marine waters, freshwaters,

beach sand and soils in the presence of the E. faecalis host we have been using.

54

0

0.5

1

1.5

2

2.5

0 2 4 6 8 10 12

Nu

mb

er o

f p

ha

ges

/10

0m

L (

Lo

g)

Time (Days)

22°C

37°C

41 C

(A)

(B)

Figure 2: Survival of enterophages at 22 °C (A) and at 37 °C (B). Aliquots were tested and Petri

dishes were incubated at the three different temperatures shown.

Prevalence

When measuring coliphages and enterophages in treated and untreated sewage from Puerto Rico

at concentrations ranging from 1.4 x 105 to 2.6 x 10

5 and from 55 to 363 Plaque Forming Units

(PFU)/100ml for coliphages and enterophages, respectively (Tables 1 and 2). It is also

noteworthy that the enterophage plaques could be easily detected even in raw sewage samples

after 8 hours. Coliphage analyses of the same samples showed luxuriant growth of background

microbiota which masked the viral plaques; even in the presence of antibiotics. These data also

showed that sewage treatment plants are able of removing/inactivating from 87 to >99.9 % of

55

influent enterophage concentrations, and the removal/inactivation of coliphages was always >90

% in the same samples.

Sewage Treatment

Plant

Influent

(PFU/100mL)

Effluent

(PFU/100mL)

% of

removal

Puerto Nuevo 2.6 X 105 1.6 x 10

4 94.0

Carolina 1.4 X 105 3.5 x 10

3 98.0

Bayamon 1.5 X 105 1.4 x 10

3 99.0

Caguas 1.4 X 105 ND 99.0

ND=Not detected

Table 1: Concentrations of coliphages in sewage treatment plants in Puerto Rico (ND-Not

detected).

Sewage Treatment

Plant

Influent

(PFU/100mL)

Effluent

(PFU/100mL)

% of

removal

Puerto Nuevo 336 2 >99.0

Carolina 184 20 89.0

Bayamon 55 7 87.0

Caguas 139 0 >99.0

Table 2: Concentrations of enterophages detected at sewage treatment plants in Puerto Rico.

CONCLUSIONS

In conclusion, we described a method developed for the detection of enterophages in

environmental samples. This method lends itself for any type of sample such as water, sewage,

soils, food and sand. We have also detected these phages in beach sand samples, but are working

on an elution procedure that may also be used for food samples. These viruses need to be studied

in order to determine if in fact, they can be used as alternate indicators of human fecal

contamination. We urge those readers who may try to determine the prevalence of enterophages

56

under their own conditions to try to use the same procedures as well as the same host in order to

be able to compare results in the future. One of the problems we have encountered with

coliphages is that many different hosts were used over the years, making it difficult to determine

if in fact the same coliphages are being detected. We hope to avoid this problem with this group

of viruses. We have started an international collaboration with colleagues from several different

geographical areas encompassing several continents, to determine the prevalence of

enterophages in raw sewage and recreational waters. This will let us know if these phages can be

used as alternate indicators of human fecal contamination in waters and other media at a global

scale. Further work also needs to be done to further characterize enterophages not only

morphologically, but also molecularly before they are taken into consideration as indicators of

human fecal contamination.

ACKNOWLEDGMENTS

We thank Ing. Capeles and Ms. Cruz Minerva Ortiz from PRASA for their help with access to

sewage treatment plant samples. We also acknowledge the help of Margerie Rivera, Nataly

Montes, Veronica Marcantoni, and Leiribel Tavarez for their help in the analyses of the samples.

We also thank Camilo and French guy for their help with the TEM analyses.

57

LITERATURE CITED

1. Moe, C.L., Waterborne Transmission of Infectious Agents in Manual of Environmental

Microbiology, R.L. Crawford, Editor. 1996, ASM: Washington, D.C. p. 22-240.

2. Dufour, A.P., Bacterial indicators of recreational water quality. Can J Public Health,

1984. 75(1): p. 49-56.

3. Wheeler, A.L., et al., Potential of Enterococcus faecalis as a human fecal indicator for

microbial source tracking. J Environ Qual, 2002. 31(4): p. 1286-93.



5. Byappanahalli, M. and R. Fujioka, Indigenous soil bacteria and low moisture may limit

but allow faecal bacteria to multiply and become a minor population in tropical soils.

Water Sci Technol, 2004. 50(1): p. 27-32.


drinking waters., in Manual of Environmental Microbiology, R.L. Crawford, Editor.



environments: health risks, detection, and potential water quality assessment tools.

Microbiol Mol Biol Rev, 2005. 69(2): p. 357-71.

8. Stetler, R.E., Coliphages as indicators of enteroviruses Appl Environ Microbiol, 1984.

48(3): p. 668-670.

9. Havelaar, A.H. and W.M. Hogeboom, A method for the enumeration of male-specific

bacteriophages in sewage. J Appl Bacteriol, 1984. 56(3): p. 439-47.

10. Loh, C.L., et al., Development of a field test kit for coliphage detection in natural waters

Tox. Assess, 1988. 5: p. 451-460.

58

11. Hernandez-Delgado, E.A., M.L. Sierra, and G.A. Toranzos, Coliphages as alternate

indicators of fecal contamination in tropical waters. Environ. Toxicol, 1991. 6: p. 131-

143.

12. Moce-Llivina, L., F. Lucena, and J. Jofre, Enteroviruses and bacteriophages in bathing

waters. Appl Environ Microbiol, 2005. 71(11): p. 6838-44.

13. Tartera, C. and J. Jofre, Bacteriophages active against Bacteroides fragilis in sewage-

polluted waters. Appl Environ Microbiol, 1987. 53(7): p. 1632-7.

14. Muñiz, I., et al., Survival and activity of Streptococcus faecalis and Escherichia coli in

tropical fresh water. Microbial Ecology, 1989. 18: p. 125-134.

15. Baele, M., Composition of enterococcal and streptococcal flora from pigeon intestines Applied

and Environmental Microbiology, 2002. 92(2): p. 348-51.

16. Martin, J.D. and J.O. Mundt, Enterococci in insects. Appl Microbiol, 1972. 24(4): p. 575-

80.

17. Macovei, L. and L. Zurek, Ecology of antibiotic resistance genes: characterization of

enterococci from houseflies collected in food settings. Appl Environ Microbiol, 2006.

72(6): p. 4028-35.

18. Cox, C.R. and M.S. Gilmore, Native microbial colonization of Drosophila melanogaster

and its use as a model of Enterococcus faecalis pathogenesis. Infect Immun, 2007. 75(4):

p. 1565-76.



20. Bachrach, G., et al., Bacteriophage isolation from human saliva. Lett Appl Microbiol,

2003. 36(1): p. 50-3.

21. Classification and nomenclature of viruses. Fourth report of the International Committee

on Taxonomy of Viruses. Intervirology, 1982. 17(1-3): p. 1-199.

59

22. Pleceas, P. and H. Brandis, Rapid group and species identification of enterococci by

means of tests with pooled phages. J Med Microbiol, 1974. 7(4): p. 529-33.

23. Ward, R.L., D.R. Knowlton, and P.E. Winston, Mechanism of inactivation of enteric

viruses in fresh water. Appl Environ Microbiol, 1986. 52(3): p. 450-9.

60

CHAPTER 3

Characterization of Enterococcus faecalis-infecting phages (enterophages) as markers of

human fecal pollution in recreational waters

T.M. Santiago-Rodríguez1, C. Dávila

1, J. González

1, N. Bonilla

1, P. Marcos

2, M. Urdaneta

1,

M. Cadete 3, S. Monteiro

3, R. Santos

3, J.W. Santo-Domingo

4 and G. A. Toranzos

1

1 Environmental Microbiology Laboratory, Department of Biology, University of Puerto Rico,

San Juan, PR, 2

Biological Sciences Department, General Studies, University of Puerto Rico, San

Juan, PR, 3

Instituto Superior Tecnico, Laboratorio de Analises, Lisboa, Portugal, 4

US

Environmental Protection Agency, Cincinnati, Ohio, USA

ABSTRACT

Enterophages are a novel group of phages that specifically infect Enterococcus faecalis and have

been recently isolated from environmental water samples. Although enterophages have not been

conclusively linked to human fecal pollution, we are currently characterizing enterophages to

propose them as viral indicators and possible surrogates of enteric viruses in recreational waters.

Little is known about the morphological or genetic diversity which will have an impact on their

potential as markers of human fecal contamination. In the present study we are determining if

enterophages can be grouped by their ability to replicate at different temperatures, and if

different groups are present in the feces of different animals. As one of the main objectives is to

determine if these phages can be used as indicators of the presence of enteric viruses, the

survival rate under different conditions was also determined as was their prevalence in sewage

and a large watershed. Coliphages were used as a means of comparison in the prevalence and

survival studies. Results indicated that the isolates are mainly DNA viruses. Their morphology as

well as their ability to form viral plaques at different temperatures indicates that several groups

of enterophages are present in the environment. Coliphage and enterophage concentrations

throughout the watershed were lower than those of thermotolerant coliforms and enterococci.

Enterophage concentrations were lower than coliphages at all sampling points. Enterophages

showed diverse inactivation rates and T90 values across different incubation temperatures in both

61

fresh and marine waters and sand. Further molecular characterization of enterophages may allow

us to develop probes for the real-time detection of these alternative indicators of human fecal

pollution.

Keywords: enterococci; enterophages; fecal pollution; indicators

Reference: Santiago-Rodriguez, T.M., et al., Characterization of Enterococcus faecalis-infecting

phages (enterophages) as markers of human fecal pollution in recreational waters. Water Res,

2010. 44(16): p. 4716-25.

INTRODUCTION

Contamination of water sources by sewage is a health-related risk because of the possible

presence of pathogenic microorganisms such as Salmonella, Campylobacter, enteric viruses as

well as pathogenic protozoa [1-4]. Thermotolerant coliforms, enterococci [5, 6], Bacteroides

fragilis phages [7, 8] and somatic and F(male)-specific coliphages [9-13] are used and have been

proposed to infer the presence of these pathogens in water sources. However, these

microorganisms fail to fulfill the criteria of an ideal microbial indicator. For instance, the

prevalence and survival of thermotolerant coliforms is short and cannot be correlated with that of

pathogens [14, 15], human pathogens are not always accompanied by enterococci and vice versa

[16-18] and Bacteroides fragilis phages are found only in some geographical areas [16]. The

survival of somatic and F+ RNA and F+ DNA coliphages is approximately 20-100 days at 20 °C

in fresh water, depending on the group, which cannot be correlated with the survival of some

enteric viruses [10, 19]. Yet, other studies have found that F+ RNA coliphages could be active as

long as 3 days and somatic coliphages as long as 7 days [20]. Even though serotypes II and III of

F+ RNA coliphages have been correlated with human fecal contamination [19, 21, 22], their

presence in cow, pig, horse and bird feces makes them unreliable markers of human fecal

pollution [23, 24]. Also, currently used indicators are very susceptible to standard chlorination

treatments [24], while enteric viruses like Coxsackie virus, Rotavirus and Norwalk viruses are

susceptible to higher chlorine concentrations and have been found in treated sewage [25].

Because the presence of enteric viruses in water sources is of great concern due to their

resistance to removal treatments, new and reliable indicators of human fecal pollution and the

62

possible presence of these pathogens are still needed. Therefore, we are proposing

bacteriophages that specifically infect Enterococcus faecalis, which we call enterophages, as

viral indicators of human fecal contamination. Enterophages are a new group of phages that have

been recently isolated from recreational waters. Most isolates are tailed-phages, with icosahedral

capsids of 80 nm in diameter and tails of 200 nm in length [15], while others have smaller

capsids (12 nm) and shorter tails (60 nm) (this study), which accordingly could belong to the

Siphoviridae family [15]. These isolates differ in size from other previously characterized

Enterococcus faecalis-infecting phages, which have bigger capsids and tails (93 nm and 204 nm,

respectively) [26]; other isolates do not possess tails and have sphere-shaped, spiked structures of

approximately 70 nm [26, 27]. We have previously developed the methods for their detection in

environmental samples and results showed that these are promising markers of human fecal

pollution. The survival of enterophages in marine recreational waters was seen to be

approximately 7-10 days [15], similar to that of enteric viruses under similar environmental

conditions [15, 28] and unlike bacterial fecal indicators, enterophages have not been detected in

pristine aquatic ecosystems [15].

Even though results show that enterophages possess some of the characteristics of an ideal viral

indicator of water contamination by human feces, further characterization is still needed. It still

remains unknown how many groups of enterophages can be present in waters contaminated with

feces, though it has been suggested that at least three groups exist [15]. Previous studies have

found that different groups of coliphages exist and can be grouped according to their ability to

replicate at specific temperatures [29]. Consequently, the aims of the present study are to

determine if enterophages are present in animal and human fecal samples, to characterize

enterophages by means of their morphology, composition of their genetic material and their

ability to replicate at some temperatures and not others; to determine the prevalence of

enterophages in a fresh water gradient in the central region of Puerto Rico as well as to

determine if there are temporal variations in sewage in Puerto Rico and Portugal, and to

determine their survival in fresh and marine waters. Because coliphages have been used and

proposed as water quality controls of fecal contamination, their detection in both animal and

human feces, as well as the prevalence and survival studies were done in parallel with that of

enterophages as a means of comparison. The survival of enterophages in sand was also

63

performed because of the possible introduction of pathogens by feces. Most bacteria are not

reliable indicators of fecal contamination in beach sand because these are part of the

environmental microbiota [30].


Detection of enterophages in animal and human feces

Cattle were selected as a study model to detect enterophages because E. faecalis is the most

commonly enterococci found in their feces [31]. Eight grams of cattle feces from 15 different

animals were collected from 4 different farms and stored at -20 °C until processed. To detect

enterophages, 1 g of each sample, 0.1 mL of Tween-20 [32], were added to 50 mL of sterile

distilled water and processed using the single layer method. The suspension was added to an

equal volume of liquefied regular strength (1X) Tryptic Soy Broth (TSB) (Difco) + agar (0.75 %

w/v) (Difco) which had CaCl2·2H20 (Fisher Scientific Co. NJ, USA) and NaN3 (MCB, OH,

USA) (final concentration of 2.6 mg/mL and 0.4 mg/mL, respectively). Additionally, 5 mL of an

overnight culture of E. faecalis grown in Azide Dextrose Broth (Difco) were added to the

mixture. This method was similarly conducted for the detection of coliphages. Briefly, the

suspension was added to molten 2X-strenght TSB and agar (1.5 % w/v). One mL of a fresh

culture of Escherichia coli in TSB was used for their detection. To test the presence of different

groups of coliphages and enterophages (according to their ability to replicate at different

temperatures), the mixtures were poured into sterile Petri dishes and incubated at 22, 37 or 41 °C

for 24 h [15].

To detect enterophages and coliphages in humans, fresh feces from 5 healthy individuals were

processed as follows: 0.1-0.5 g of fresh feces were suspended in 3 mL of a sterile 0.85 % NaCl

solution. 0.1 and 1 mL of the suspension were separately processed using the double layer

method. Briefly, the volumes were added to 4 mL of molten 1X TSB and agar (final 0.75 %

w/v), which had CaCl2 and NaN3 as described. Three-mL of an overnight culture of E. faecalis

were added to the mixture and poured into Petri dishes containing 20 mL of 1X TSB and agar

(final 1.5 % w/v). For the detection of coliphages, 1 mL of an overnight culture of E. coli was

added to the mixture. Plates were incubated at 37 °C for 24 h.

64

Enterophage Isolation and Purification

To isolate enterophages for further characterization, a 50 mL volume sample of raw domestic

sewage was processed as described [15]. The solution was mixed, poured into sterile Petri dishes

and plates were incubated at different temperatures, namely 22, 37, 41 and 45 °C for 24 h.

Individual viral plaques detected at each of the incubation temperatures were isolated. Different

plaque size and translucency were used as criteria for the isolation. Plaques were plucked using

sterile Pasteur pipettes and the plug was placed in a sterile Eppendorf tube containing 500 μL of

sterile Phosphate Buffer Solution (PBS). The plugs were dislodged and emulsified using the

same pipette. The tubes were centrifuged at 14,000 rpm for 10 min at 10 ºC to remove cellular

debris and agar and the supernatant transferred into a sterile tube. The isolates were then

propagated using 100 μL of the supernatant and processed using the double layer method as

described. Then, for those plates showing complete viral lysis, 5 mL of PBS was added and

slowly agitated rotationally for 20 min. The top agar was transferred to a sterile centrifuge tube,

broken down and then centrifuged at 14,000 rpm for 10 min at 10 ºC. The supernatant containing

the viruses was kept in a sterile tube at 4-7 °C for further use. To determine if the isolates were

capable of replicating at other temperatures, an E. faecalis inoculum was spread throughout a

Petri dish containing 1X TSB, 0.75 % agar, CaCl2 and NaN3 using a sterile swab. An aliquot of

10 μL of the purified supernatant was placed on top of the Petri dishes and incubated at 22, 37,

41 and 45 °C for 24 h. Further titration of each isolate was done using the double layer method

as described above.

Morphological Characterization of Enterophages

Prior to the examination of the morphology of the enterophage isolates, hydroextraction was

used to concentrate the viruses; briefly, a dialysis tube (12,000-14,000 MW cutoff, Spectrapor,

Los Angeles, CA) was loaded with 5 mL of the enterophage isolate, covered with crystalline

polyethylene glycol (PEG, Mol.wt. 8, 000, Sigma Chem. Co. MO; APHA, 1989), clamped and

placed at 4-7 °C overnight. The dialysis tube was washed with 100 μL of sterile 0.85 % NaCl.

The resulting solution was transferred into a sterile Eppendorf tube and kept at 4 °C until

analysis. An aliquot of the final solution was placed on carbon Type-B 200 mesh copper grids or

ultra thin carbon film/holey carbon 400 mesh copper grids and stained with uranyl acetate 2 %,

pH 4.5. All specimens were examined using a Karl Zeiss Leo 922 energy filtered transmission

65

electron microscope operated at 200 KV. At least 5 phage particles of each type were observed

[15].

Nucleic Acid Analyses

Extraction of enterophages nucleic acids was conducted using standard techniques of proteinase

K and phenol:chloroform treatments, followed by precipitation with ethanol [33]. To determine

if the enterophage isolates genetic material was DNA or RNA, aliquots of the isolated nucleic

acids were treated with either 1 U/μl DNase (Promega, Madison, WI, USA) [34] or 10 μg/μl

RNase (Sigma-Aldrich, Co. St. Louis, MO, USA). Agarose gel electrophoresis was performed as

previously described [33], using 0.7 % of agarose (Sigma-Aldrich, Co. St. Louis, MO, USA) gels

in TAE buffer. Bands were visualized after staining with an ethidium bromide solution (final

concentration of 0.5 μg/mL).

Prevalence in raw sewage

To determine the prevalence of enterophages in raw and treated domestic sewage from Puerto

Rico and Portugal, samples were collected monthly for 6 months and processed using the single

layer method as described [15]. For the detection of coliphages, serial dilutions were done and

processed using the double layer method as described. Plates were incubated at 37 °C for 24 h.

Detection of enterophages and other viral and bacterial indicators in a large watershed

Ten sampling sites subjected to different environmental conditions in the Rio Grande de Arecibo

watershed, localized in the central region of Puerto Rico, were selected to determine the

prevalence of enterophages in a natural setting (Figure 1). One liter samples from each point

were collected in sterile plastic bottles every week for 2.5 months and kept at 8 °C until

processing. Samples were analyzed for thermotolerant coliforms, enterococci, coliphages and

enterophages [15, 35]. For bacteria enumeration, 100 mL of water per sample site were filtered

using 47 mm polycarbonate membrane filters (GE Water & Process Technologies, pore size

0.45μm). This was done separately for the enumeration of both bacterial indicators. For the

enumeration of thermotolerant coliforms, filters were placed on Difco m-FC agar and incubated

at 45 °C for 24 h. Enumeration of enterococci was done by placing the membrane filter on Difco

m-Enterococcus agar and incubated at 37 °C for 48 h. Coliphage and enterophage were

66

enumerated by processing 100 mL of the water samples using the single layer method as

described in section 2.1 [15]. The solution was poured into Petri dishes and incubated at 22, 37,

41 or 45 °C for 24 h to test for the possible presence of different groups of enterophages across

the watershed.

Figure 1: Shows study site. Sample points are ordered according to their position in the

watershed, localized in the central region of Puerto Rico. Rio Grande de Arecibo watershed

localized in the central region of Puerto Rico. Sample points in this study are ordered according

to their position in the watershed (See legend).

67

Survival of enterophages and coliphages in waters and sand

To determine the survival of enterophages and coliphages in fresh and marine waters as well as

in beach sand, 2 L of fresh or marine water samples and 2 kg of sand were collected in sterile

plastic bottles and kept at 8 °C till used. At the laboratory, the water and sand samples were

placed in a sterile covered glass beaker. In order to simulate the die-off rate of these

bacteriophages under natural conditions [36, 37], raw domestic sewage with a coliphage

concentration of 104/100 mL

was added to the samples; the same concentration of an

enterophage isolate from a laboratory stock was also added to the samples. The samples were

kept in the dark at 22 °C for up to 12 days. Every two days, 50 mL aliquots were separately

processed for the detection of enterophages and coliphages using the single layer method as

described. Similarly, 50 g of sand and 0.1 mL of Tween-20 were added to 50 mL of sterile

distilled water and vigorously shake. The suspensions were processed as previously described

[15].

In order to make comparisons, decay constants (kd) from both enterophages and coliphages in

fresh and marine waters and sand were determined using the slopes of linear regressions made on

the semilog plots (PFU versus time). Decay rates (percent/ 2 days) were calculated by

multiplying the decay constants by 100. The time to reach a 90 % reduction in PFU densities

(T90) was also determined by dividing ln (0.1)/kd [36, 38-40].

RESULTS AND DISCUSSION

Enterophages and coliphages in animal and human feces

Coliphages have been used as viral indicators of fecal pollution, but their host specificity

includes cows, swine, humans, etc. In addition, those groups found in humans can also be found

in animal feces [23]. Similarly to these studies, we found coliphages in 12 of the 15 cattle feces

samples at an incubation temperature of 22 °C, but not at other incubation temperatures. Their

concentrations ranged from 8 Plaque Forming Units (PFU) to 326 PFU/g (data not shown). On

the other hand, enterophages were not detected in any of the cattle feces. Even though

enterophages were not detected, additional fecal samples from different animal species must be

processed to confirm enterophages specificity. Coliphages were detected in 3 out of the 5 human

fecal samples (approximately 80 PFU/g; data not shown), but enterophages were not detected.

68

These results could be due to the low concentrations of enterophages found in human feces.

Also, other incubation temperatures must be tested for their detection in fecal samples (namely

22 and/or 41 °C). Regarding the method used, the single layer method could be more appropriate

for their detection in human feces. These results are similar to those found in other studies which

showed that bacteriophages may be present in human feces at concentrations ranging from 0 to

105

CFU/g [41-44]. These studies have found that many of the bacteriophages in the human

intestine could be those infecting gram-positive bacteria, as in the case of enterophages [41]. The

absence of enterophages in cattle feces and other animals is encouraging, since it suggests that

these viruses are restricted to human hosts.

Differences in enterophage isolates according to morphology, genetic material and ability

to replicate at different temperatures

Both the morphology and genetic material of the enterophage isolates in this study correspond to

those viruses belonging to the Siphoviridae family. The morphology of the enterophage isolates

described here indicates that these are tailed phages, with icosahedral capsids of 12 nm in

diameter and non-contractile tails of 60 nm long (Figure 2). This morphology is different from

other virions we have described recently [15], which showed a bigger capsid and a longer tail,

similar to the classical Bradley’s basic morphology Group B belonging to the Siphoviridae

family [45].

Figure 2: Transmission Electron Microscopy image (TEM) of an enterophage isolate in this

study. The image shows that the isolate possesses an icosahedral head of 12 nm and a 60 nm

non-contractile tail (Bar =20 nm).

69

Figure 3: The genetic material of several enterophage isolates in a 0.7% agarose gel. The

molecular weight marker is shown in lane M (Lambda DNA HindIII digest, SigmaAldrich Co.

St. Louis, MO, USA). Lanes 1 and 4 contain untreated nucleic acids from two different isolates.

Lanes 2 and 5 contain DNase treated samples and lanes 3 and 6 contain RNase-treated samples.

In terms of the genetic material of enterophages, high molecular weight bands were observed in

the Rnase-treated samples. Enterophage nucleic acid molecular weight appears to be more than

23 kb and be in a super coiled conformation. Fragments in lanes 2 and 5 were degradated by

DNase, showing that the genetic material of the enterophage isolates described here is composed

of DNA (Figure 3). However, it still remains the possibility that the morphology and genomes of

other isolates could be different from those described. It also remains unknown if the isolates in

this study are double stranded or single stranded DNA viruses.

Enterophages isolated at 22 °C were capable of replicating at 22, 37, 41 and 45 °C. However,

those enterophages that were isolated at 37 and 41 °C replicated at 22, 37 and 41 °C, but not at

45 °C (Figure 4). Though further characterization of enterophages is still needed, these results

suggest that at least two groups of enterophages exist. Because enterophages can be isolated at

environmental temperatures, they can be detected in places without the facility of an incubator

by incubating the plates at ambient temperatures.

70

0

2

4

6

8

10

12

14

1 2 3 4

En

tero

ph

ag

e c

on

cen

tra

tio

n/m

L (

Lo

g)

Phage groups

22°C

37°C

41°C

45°C

Figure 4: Enterophage concentration/mL of four different enterophage isolates among four

different temperatures. Group 1 and 2 represent phages isolated from 22 ºC, group 3 represent an

isolate from 37 ºC and group 4 represents the isolate from 41 ºC. Results represent the geometric

mean of three replicas and standard deviations are represented by error bars.

Prevalence of enterophages in sewage and in a large watershed in Puerto Rico

Although enterophages were isolated at lower concentrations than coliphages, they were detected

in raw and treated domestic sewage in this study (Table 1). Interestingly, even though the same

host strain was used to detect enterophages in both Portugal (Port) and Puerto Rico (PR)

treatment plants, they were found in higher concentrations in raw domestic sewage from

Portugal, but coliphage concentrations were lower than those found in Puerto Rico (data not

shown). Prevalence studies in Portugal indicate that these viruses are not restricted to particular

geographical areas and could be tested as viral indicators in different waters types. Both

enterophages and coliphages (data not shown) were also detected in treated sewage in Puerto

Rico. Treated sewage was not processed for the enumeration of coliphages from Portuguese

treatment plants. Primary treatment is performed on raw sewage from domestic wastewater

treatment plants PR-A, PR-B and PR-C, while raw sewage from PR-D receives tertiary

treatment. In raw sewage receiving primary treatment, approximately 56 to 97 % of enterophages

were removed (Table 1), while a 91 % removal was seen in sewage from wastewater treatment

71

plant PR-D. Coliphages, on the other hand, were removed from a 95 to a 98 % in PR-A, PR-B

and PR-C, whereas more than 99 % of coliphages were removed from raw sewage in PR-D (data

not shown). Approximately 60 to > 99 % of enterophages were removed from raw sewage in

Portuguese treatment plants. Prevalence results from Puerto Rican treatment plants suggest that

enterophages may be more resistant to removal treatments than coliphages and could be

potentially more useful as indicators of the presence of enteric viruses even in treated sewage.

Water Treatment

Plant

Enterophages in inffluent

(PFU/100 mL)

Enterophages in effluent

(PFU/100 mL)

PR-A 82 ± 51.5 56 ± 55.6

PR-B 60 ± 46.1 14 ± 14.5

PR-C 115 ± 140.1 18 ± 1.2

PR-D 53 ± 53.5 6 ± 10.7

Port-A 17 ± 3.2 1 ± 0.9

Port-B 634 ± 15.7 253 ± 4.6

Port-C 695 ± 21.6 3 ± 0.9

Port-D 22 ± 2.8 2 ± 0.6

Port-E 575 ± 22.9 5 ± 1.3

Port-F 212 ± 7.0 12 ± 1.3

Port-G 405 ± 6.6 11 ± 1.4

Port-H 173 ± 10.9 21 ± 1.7

Port-I 774 ± 11.2 3 ± 0.9

Port-J 487 ± 6.5 4 ± 0.9

Table 1: Prevalence of enterophages in raw and treated sewage at different domestic wastewater

treatment plants in Puerto Rico (PR) and Portugal (Port). All data represent the average

concentration and standard deviation of viral plaques over a six-month period (n = 6).

In the large watershed, coliphages were found at higher concentrations than enterophages in all

sample points, but both bacteriophages were found at lower concentrations than thermotolerant

coliforms and enterococci (Figure 5). The high concentrations of bacterial indicators in the

watershed suggest that these may also be from animal origin, consequently, using these to track

the source of the contamination may not be reliable [46, 47]. No fluctuations in enterococci and

thermotolerant coliforms concentrations were found throughout the sampling period (Figure 5C

and 5D, respectively). Coliphages were detected at all incubation temperatures in most of the

sample points. These results suggest that different groups of coliphages were introduced into the

72

water sources, which may not necessarily be of human origin. Coliphage concentrations were

found to fluctuate throughout the sampling period; however, fluctuations were not as prominent

as those of enterophages (Data not shown).

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Figure 5: Concentrations of enterophages (A), coliphages (B), enterococci (C) and

thermotolerant coliforms (D) at ten different fresh water sample points with different impacts.

Sample points are numbered according to their position in the watershed: (1) Garza lake, (2)

74

Vaca river, (3) Before WTP Adjuntas, (4) After WTP Adjuntas, (5) Rio Grande de Arecibo, (6)

Before WTP Utuado, (7) After WTP Utuado, (8) Mouth of Rio Grande de Arecibo, (9)

Criminales river and (10) Caguana river at Jayuya. Numbers represent the geometric mean of a

2.5 month sampling period. Standard deviation is represented by error bars.

We are in the process of further characterizing the enterophage isolates molecularly, including

the sequencing of the viral genomes, which may allow us to determine unique sequences to be

used for the development of real-time methods for their detection in recreational and other water

sources. Also, future epidemiological studies should be focused on determining the relationship

between the prevalence of enterophages and enteric viruses in recreational waters and the

symptoms associated with ingesting water testing positive for the presence of enterophages.

Survival of enterophages in fresh waters

Enterophages were detected at 37 and 41 °C at 4 h after the incubation of the Petri dishes, but not

at 22 °C (data not shown). This indicates that these temperatures can be used to detect

enterophages in water samples in less time than bacterial indicators at similar temperatures.

Enterophages have a survival similar to that of human enteric viruses in fresh waters (Figure

6A) [28] and are able of replicating at 22, 37 and 41 °C. On the other hand, the survival of

coliphages in this study was more than 12 days, which cannot be correlated with that of certain

enteric viruses (Figure 6B). Even though temperature does not affect the formation of viral

plaques, it seems to affect the decay rates of enterophages. At 22 and 37 °C, the decay rates were

33.7 and 38.5 % every 2 days, respectively and at 41 °C the inactivation rate was 35.9 % every 2

days. T90 values also differed among the different temperatures. At 22 and 41 °C, 90 % of the

initial inoculum was inactivated at approximately 6.8 and 6.6 days, respectively and 6.0 days at

37 °C. Although it is not new that temperature does affect the decay rates of microorganisms [19,

36], our experiments on enterophages decay rates are part of their characterization as markers of

human fecal pollution and further experiments are still needed. Temperature also affected the

inactivation rates of coliphages. At 22 °C, approximately 11.6 % of the initial inoculum was

inactivated every 2 days. At 37 °C, coliphages had an inactivation rate of 9.9 % every 2 days; but

the highest inactivation rate was seen at 41 °C (26.2 % every 2 days). The time required to

inactivate 90 % of the initial coliphage concentration at 22 °C was 20.0 days, 23.3 days at 37 °C

75

and 9.0 days at 41 °C (Figure 6B). The long die-off rate of coliphages indicates that some

groups may not be used to track recent fecal contamination in water sources. Future survival

studies of enterophages in fresh waters will be done in order to determine the survival of

enterophages and coliphages in different points across the watershed described here and those

factors that could potentially affect their survival.

Survival of enterophages in marine recreational water and sand

As in the survival study of enterophages in fresh waters, plaques were detected at 37 and 41 °C

at 4 h after the incubation, but not at 22 °C, which suggests that these temperatures are favorable

to detect enterophages in short incubation periods. This could be due to diminished adsorption

rates of enterophages at low temperatures possibly because of a reduced affinity between phages

and bacterial surface receptors. Also, the adsorption rate is usually higher at the optimum

temperature for the growth of the host bacteria, suggesting that actively functioning bacteria are

required for a successful infection [48]. Nevertheless, after 11 days viral plaques were not

detected at 4 hr of incubation to any of the temperatures, but they were still detected after 24

hours. This may indicate that the adsorption rates were reduced with time.

The enterophage isolate used in this study has a survival time in both marine waters and sand of

11 to 13 days (Figure 7A and 8A), which is less than coliphages (Figure 7B and 8B). The decay

rates were 27, 29.3 and 30.7 % every 2 days at 22, 37 and 41 °C, respectively. The calculated T90

values for each temperature were 8.5, 7.9 and 7.5 days correspondingly. In the study of

enterophage survival in sand, the decay rates were 17, 20.6 and 21.4 and the calculated T90

values for each temperature, were 13.5, 11.2 and 10.8 respectively. Regarding coliphages, the

decay rates in marine water were 5, 4.5 and 7.7 % every 2 days. As in the study of coliphages in

fresh water, the higher decay rate occurred at 41 °C. The decay rates for coliphages in sand were

3.6, 4.1 and 6.2 and the T90 values were 64, 56.2 and 37.1.

Given that the isolate used in both survival studies (fresh water and marine water and sand) was

the same, it is interesting that the observed survival time was higher in marine water and sand

environments. Factors that could cause these longer survival periods include the high

76

concentration of inorganic salts, due to the fact that phage adsorption is strongly dependent on

salt concentrations [49].

These results and previous survival experiments using coliphages have found that it would be

impossible to correlate their survival with recent fecal contamination [50]. The survival of

enterophages in fresh and marine waters could indicate the recent introduction of human feces

into water sources. Molecular methods, like those currently tested on other microorganisms [51],

could also be developed to determine the die-off rate of enterophages in recreational waters. In

addition, because the effect of sunlight has been extensively studied on bacteria and

bacteriophages [36, 38], the inactivation of enterophages by sunlight has to be tested in future

studies.

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77

Figure 6: Survival of enterophages (A) and coliphages (B) in fresh water. Numbers represent the

arithmetic mean of three replicas. Standard deviations are represented by error bars.

Figure 7: Survival of enterophages (A) and coliphages (B) in marine water. Numbers represent

the average of three replicas. Standard deviations are represented by error bars.

78

Figure 8: Survival of enterophages (A) and coliphages (B) in sand. Numbers represent the

average of three replicas. Standard deviations are represented by error bars.

CONCLUSIONS

The absence of enterophages in cattle feces could indicate their specificity to the human colon

and could be potentially used to determine the presence of human enteric viruses in recreational

waters. Differences in morphology, genome and replication temperatures are factors that should

be taken in consideration when characterizing a new viral indicator, as in the case of

enterophages. These characteristics should be taken in consideration when choosing which

79

enterophage isolates have the potential of being molecularly characterized for their detection in

water sources. Enterophages could be used as surrogates of enteric viruses in recreational waters

due to their resistance to primary and tertiary treatments and similarity in die-off rates in fresh

and marine waters. Unlike proposed viral indicators, enterophages are not constrained to specific

regions and could be used to infer human fecal pollution in different water sources.

Few indicators have been proposed to infer the introduction of pathogens into beach sand. The

die-off rate of enterophages could also indicate recent fecal contamination, not only in water

sources, but also in beach sand.

ACKNOWLEDGMENTS

We thank Carlos Toledo for reviewing the manuscript at its early and late stages, Marisol

Rodriguez for collecting samples, Jean Frances Ruiz and Gwendolyn Argüello for processing the

samples.

80

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85

CHAPTER 4

Microbial quality of tropical inland waters and effects of rainfall events

T.M. Santiago-Rodriguez1, R.L. Tremblay

2, 3, C.Toledo-Hernandez

4, J.E.Gonzalez-Nieves

4, H.

Ryu4, J.W. Santo-Domingo

4 and G.A. Toranzos

1

1Environmental Microbiology Laboratory, Department of Biology, University of Puerto Rico,

Rico, San Juan, Puerto Rico 00931, 2Department of Biology, Call Box 860, University of Puerto

Rico, Humacao, Puerto Rico 00792, 3Crest-Catec, Center for Applied Tropical Ecology and

Conservation, PO Box 23341, University of Puerto Rico, Rio Piedras, Puerto Rico, USA 00931,

4US Environmental Protection Agency, Cincinnati, OH, USA

ABSTRACT

Novel markers of fecal pollution in tropical waters are needed since conventional methods

recommended for other geographical regions may not apply. To address this, the prevalence of

thermotolerant coliforms, enterococci, coliphages and enterophages was determined by culture

methods across a watershed. Additionally, human-, chicken- and cattle-specific PCR assays were

used to identify potential fecal pollution sources in this watershed. An enterococci quantitative

PCR (qPCR) assay was tested and correlated with culture methods at three sites since water

quality guidelines could incorporate this technique as a rapid detection method. Various rainfall

events reported prior to sample collection at three sites, were considered in the data analyses.

Thermotolerant coliforms, enterococci, coliphages and enterophages were detected across the

watershed. Human-specific Bacteroides, unlike the cattle and chicken-specific bacteria, were

detected mostly at sites with the corresponding fecal impact. Enterococci were detected by

qPCR as well, but positive correlations with the culture method were noted at two sites,

suggesting that either technique could be used. However, no positive correlations were noted for

an inland lake tested, suggesting that qPCR may not be suitable for all water bodies.

Concentrations of thermotolerant coliforms and bacteriophages were consistently lower after

rainfall events, pointing to a possible dilution effect. Rainfall positively correlated with

enterococci detected by culturing and qPCR, but this was not the case for the inland lake. The

toolbox of methods and correlations presented here could be potentially applied to assess the

microbial quality of various water types.

86

Keywords: bacterial indicators, bacteriophages, enterophages Microbial Source Tracking,

rainfall

Reference: Santiago-Rodriguez, T.M., et al., Microbial quality of tropical inland waters and

effects of rainfall events. Appl Environ Microbiol, 2012. 78(15): p. 5160-9.

INTRODUCTION

Monitoring microbial indicators of fecal pollution in tropical waters remains an issue of concern

since these may not accurately indicate the presence of microbes associated with fecal matter.

Indicators of fecal contamination used in different geographical areas include the thermotolerant

coliforms and enterococci [1]. These indicator bacteria are present in the intestinal tract of warm-

blooded animals and therefore, their presence in waters may indicate fecal pollution. However, it

has been shown that thermotolerant coliforms and enterococci may be part of the environmental

microbiota of tropical waters [2]. In addition, many bacterial indicators cannot be used to

indicate the time and source of the fecal contamination since they can replicate outside their host

and are present in the feces of different animals [3-8]. These shortcomings with regard to

indicator bacteria have prompted the use of alternate indicators, such as bacteriophages, which

show promising characteristics. For instance, coliphages, which are normally isolated from feces

and fecally-contaminated waters, do not appear to replicate outside their host and certain groups

have similar survival characteristics to enteric viruses in waters [9-12]. However, certain

coliphage groups are present in the feces of different warm-blooded animals and therefore, may

not be used to discriminate the source of the fecal pollution [13].

Recently, isolated phages that infect a specific Enterococcus faecalis type strain (enterophages)

have been proposed as good indicators of human-specific fecal contamination. Enterophages

have a similar survival time as human enteric viruses in marine and fresh waters and sand [14-

17], have been detected in raw and treated domestic sewage in Puerto Rico and Portugal and

have been detected neither in pristine waters nor in animal feces (e.g. pigs, dogs, cattle and

chickens) [16, 17]. However, more data are needed in order to accept enterophages as alternate

indicators of human fecal contamination. One way to further test enterophages as indicators of

87

fecal pollution is to compare them with currently used indicators and emerging molecular

methods in various water types.

Microbial Source Tracking (MST) methods can complement traditional methods used to assess

microbial water quality [18]. Specifically, host-specific assays have been tested in various water

sources and several have been successfully used to discriminate among the sources of fecal

pollution. Many Bacteroides species make up the intestinal microbiota of warm-blooded animals

their primary habitat and some species have shown high levels of host-specificity. In addition,

Bacteroides cannot replicate outside the intestinal tract as most species are strictly anaerobic

bacteria [19, 20]. Quantitative PCR (qPCR) is also perhaps one of the most promising methods

that indicate the levels of the target contaminant. However, as with culture-based methods,

molecular techniques suffer from shortcomings, such as the inability to distinguish between

viable and dead cells or the infectivity status of the target microorganism.

Detection of bacterial and viral indicators by culture or molecular-based techniques may be

influenced by rainfall events. It has been shown that rainfall may lead to higher numbers of

indicators and pathogens into surface waters and thus may represent an increased risk to human

health [21, 22]. Runoff, resuspension of sediments and sewage overflows, resulting from rainfall

events, may contribute to an increment of indicators and pathogens in surface waters [23, 24].

Nonetheless, rainfall is often not considered when monitoring the microbial quality of waters.

One main question to address is whether sampling under dry conditions alone is sufficient to

infer the presence of fecal indicators and pathogens or if results differ under wet conditions.

However, it is relatively difficult to confirm that an increase in microbial indicators and

pathogens after precipitation events truly represents a recent input of fecal matter [23]. In the

present study, we assessed the microbial quality of a tropical watershed using currently used

(thermotolerant coliforms, enterococci and coliphages) and proposed indicators (enterophages),

as well as molecular methods (host-specific assays and qPCR), considering rainfall as a possible

influential variable.


Sample collection and sampling sites

88

In all cases, single one-liter grab samples were collected in sterile plastic bottles with sodium

thiosulfate (final concentration of approximately 10.0 mg/L) [25], up to four times per month

from November 2009 to April 2010. A total of 13 samples per site were collected and tested,

kept at 6-8 °C and processed within 4 to 6 h. Samples were collected from the Rio Grande de

Arecibo watershed in Puerto Rico, characterized by sporadic rainfall throughout the year.

Watersheds, unlike those found in many regions, are composed of small creeks. One of the

interesting characteristics of the Rio Grande de Arecibo watershed is that it has very distinct land

uses that contribute to human and/or animal fecal matter loadings. The watershed possesses both

urban and rural areas inhabited by more than 100,000 people, with waters that are used for

recreational activities and as a drinking water source. The watershed is linked to two of the

biggest dams on the island, providing approximately 380 million liters of drinking water daily to

the north coast [26].

Ten sampling sites were selected based on the potential different fecal pollution sources

associated with them. Site 1 (Largo Garza) is bordered at the southwestern part by the Reserva

Forestal de Toro Negro, thus the number of households are limited and farm animals, such as

poultry and cattle, are scarce or absent. Site 2 (Rio Vaca) is located downstream from site 1 and

residencies, poultry and cattle are also limited. Site 3 and 4 (Río Cidra) are located downstream

from the urban nucleus of the municipality of Adjuntas. Therefore, both sites are presumably

influence by urban runoff and local poultry. Site 3 is located upstream of the domestic

wastewater treatment plant (WTP) at Adjuntas and site 4 is located downstream from this WTP.

Site 5 (Río Grande de Arecibo) is lined by houses that drain their wastewaters directly into the

river and poultry and horses were also frequently observed nearby or in the river. Site 6 is

located at the southern entrance of the urban nucleus of the municipality of Utuado. It is

bordered by houses that manage their wastewater through septic tanks and by grass-fields, where

cattle were frequently observed. Site 7 is located 200 m downstream from the domestic WTP at

Utuado and poultry and cattle were frequently observed. Site 8 is located where the Rio Grande

de Arecibo discharges into the Atlantic Ocean, is impacted by cattle grazing and used for

recreational activities. The Río Criminales (site 9) is located between a fenced-farm with more

than 100 cows and human residences with septic tanks. Site 10 is located at the Rio Caguana and

receives the input of a WTP. This place is also impacted by cattle and horses (Figure 1).

89

Figure 1: Rio Grande de Arecibo watershed in Puerto Rico. Water samples were collected from

Lago Garza (site 1), Rio Vaca (2), upstream from the WTP at Adjuntas (3), downstream from the

WTP at Adjuntas (4), Rio Grande de Arecibo (5), upstream from the WTP at Utuado (6),

downstream from the WTP at Utuado (7), the estuary (8), Rio Criminales (9), and Rio Caguana

(10). Reproduced from reference 13 with permission.

90

Enumeration of indicators by culture methods

Thermotolerant coliforms were enumerated using m-FC agar incubated at 45 °C for 24 h and

enterococci using m-Enterococcus agar incubated at 37 °C for up to 48 h [27]. Coliphages and

enterophages were quantified using the single layer method as described previously [16]. The

type strains used for phage enumeration were Escherichia coli (ATCC 15597) and E. faecalis

(ATCC 19433). Plates were incubated at 22, 37, 41 and 45 °C to detect bacteriophages that

replicate at different temperatures. However, data presented in this study correspond to those

phages that replicate at 22 ºC since preliminary analyses suggested that this may be the optimal

temperature.

DNA extraction and PCR conditions

Water samples (100mL) were filtered through polycarbonate membranes (0.4 μm pore size, 47

mm diameter) (GE Water and Process Technologies, Trevose, PA). Total DNA was extracted

from the membranes using Mo Bio PowerSoil kits (MO BIO Laboratories, Carlsbad, CA)

according to the manufacturer’s protocol. DNA concentration was estimated using a NanoDrop

ND-1000 UV spectrophotometer (NanoDrop Technologies, Wilmington, DE). DNA extracts

were stored at -20 °C until further processing and analyzed using host specific PCR assays

commonly used in fecal source tracking studies and qPCR assay for general enterococci [28]

(Table 1). PCR was performed using cattle and human specific Bacteroides assays targeting 16S

rRNA genes and two chicken-specific assays targeting functional genes [29-31]. For

convenience, cattle and human-specific Bacteroides will be referred as CSB and HSB,

respectively. PCR amplifications were performed in 25 μL using the polymerase TaKaRa Ex

TaqTM (Takara Bio Inc.) in a Bio-Rad Tetrad2 Peltier Thermal Cycler (Bio-Rad, Hercules, CA)

under the following cycling conditions: an initial denaturation step at 95 °C for 5 min, followed

by 35 cycles of 1 min at 95 °C, 1 min at optimum annealing temperature, and 1 min at 72 °C.

PCR products were visualized in 1.5 % agarose gels using GelStar Nucleic Acid gel stain

(Lonza, Rockland, ME, USA).

The Taqman qPCR assay targeting the 23S rRNA gene of Enterococcus spp. (Entero1) was

performed in 25 μL reaction mixtures containing 1× TaqMan universal PCR master mix with

AmpErase uracil-N-glycosylase (Applied Biosystems, Foster City, CA), 0.2 mg/mL bovine

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serum albumin (Sigma), 200 nM of each primer and FAM-labeled TaqMan probe. The qPCR

assays were performed using a 7900 HT Fast Real-Time Sequence Detector (Applied

Biosystems). All reaction mixtures were prepared in triplicate in MicroAmp Optical 96-well

reaction plates with MicroAmp Optical Caps (Applied Biosystems). The amplification protocol

involved incubation at 50 °C for 2 min to activate uracil-N-glycosylase, followed by 10 min of

incubation at 95 °C, and 40 cycles of 95 °C for 15 s and 60 °C for 1 min. PCR data were

analyzed using ABI’s Sequence Detector software (version 2.2.2). Duplicate serial dilutions of E.

faecalis genomic DNA (10-8

to 10-12

g/reaction) were used to generate standard curves. Percent

amplification efficiencies were calculated following the instrument’s manufacturer instructions

(Applied Biosystems). No-template controls were used to check for cross-contamination (two per

PCR plate). Assays were performed with 2 µL DNA extracts in a total volume of 100 µL, and

ten-fold dilutions of each DNA extract were used to test for PCR inhibition. Based on the

standard curve, qPCR intensities (QI) were expressed as a unit of pg/2 µL (i.e., genomic DNA

mass/reaction volume). Subsequently, the concentrations of the target gene (C) in water samples

were calculated by the following equation C (pg/100 mL) = QI (pg/2 µL) × CF × DF, where CF

(µl/mL) is a conversion factor of 1 and DF is a dilution factor of 50 [32].

Assay Primer and probe sequences (5’ to 3’)* Ta

(°C)**

Size

(bp)

Reference

General Bacteroides Bac32F: AACGCTAGCTACAGGCTT 53 694 Bernhard &

Bac708R: CAATCGGAGTTCTTCGTG Field (2000a)

Human-specific

Bacteroides (HSB)

HF183: ATCATGAGTTCACATGTCCG 63 543 Bernhard &

Field (2000b)

Cattle-specific

Bacteroides (CSB)

CF128: CCAACYTTCCCGWTACTC 62 598 Bernhard &

Field (2000b)

Chicken-specific

Bacteroides

CP2-9F: GTAAGACAGCAACCCCATGTA 56 245 Lu et al. (2007)

CP2-9R: ACCTATGGTTCAACACGCTTTA

Chicken-specific

Clostridium

CP3-49F: GTCCAGCGCCTCATTGAT 57 329 Lu et al. (2007)

CP3-49R: TGGTGATCGACTTTTCCAAT

General Enterococcus

qPCR (Entero1)

ECST748F: AGAAATTCCAAACGAACTTG 60 92 Ludwig &

Schleifer (2000) ENC854R: CAGTGCTCTACCTCCATCATT

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GPL813:6FAM-

TGGTTCTCTCCGAAATAGCTTTAGGGCTA-

TAMRA

GPL813TQ:6FAM-

TGGTTCTCTCCGAAATAGCTTTAGGGCTA-

TAMRA

Table 1: Summary of oligonucleotide primers and probes for PCR and TaqMan qPCR.

USGS precipitation data

Precipitation data reported 24 and 48 h and one week prior to sample collection were obtained

from the US Geological Survey (USGS) Caribbean Water Science Center

(http://pr.water.usgs.gov/) for sites 1, 6 and 7 (USGS stations 50020100, 50021700 and

50024950, respectively) (Table 2). Precipitation data was collected from these sites due to their

proximity to a USGS station.

Site USGS station 24 h 48 h 1 week

Lago Garza (1) 50020100 1.6±0.4 5.7±0.4 64.3±40.3

Before WTP Utuado (6) 50021700 13.3±15.3 18.5±15.7 60.2±35.6

After WTP Utuado (7) 50024950 8.3±10.2 8.6±10.3 40.6±25.6

Table 2: Reported rainfall (mm; mean ± SD) for indicated period (November 2009 to April

2010) before sample collection. Rainfall data was collected from the USGS stations indicated.

Statistical analyses

The mean concentrations of cultivable thermotolerant coliforms, enterococci, coliphages and

enterophages and enterococci detected by qPCR were calculated using a Bayesian approach as

described previously [33]. Estimates were calculated assuming normal distribution of response.

Analysis of comparison among concentration at different temperatures, sample dates and sample

sites were analyzed using a one-way or two-way ANOVA [33]. Interaction effects were not

considered in these analyses and in all Bayesian analyses, credible intervals (CI) were calculated.

In addition, to determine correlations between rainfall and the concentration of the indicators (24

h, 48 h and one week) prior to collecting the sample, a Cross-correlation analysis was used.

http://pr.water.usgs.gov/

93

However, prediction of the response or effect of rainfall to microbial concentrations was

estimated using non-linear regression as described by [33-37]. In all cases, the error around the

point estimate, the 95% CI, is calculated. The 95% CI is the area under the curve of the posterior

distribution, as compared to the 95% confidence interval, that is the probability that the point

estimate would be between the lower and upper bound [36]. All analyses were performed with

the software package IBM SPSStatistics v.19.

RESULTS

Detection of indicators by culture-based methods

Thermotolerant coliforms showed the lowest counts at sites 1 and 9, with mean concentrations of

0.0 and 11.6 CFU/100 mL, respectively. The highest concentrations of thermotolerant coliforms

correspond to sites 4 and 7, with mean concentrations of 817.8 and 1053.0 CFU/100 mL,

respectively (Figure 2A). The variance of thermotolerant coliforms did not differ across the

sampled points (354.0-356.0 CFU/100 mL), except for site 10, in which the SD was 32.3

CFU/100 mL. Enterococci exhibited lower counts at sites 1 and 10, with mean concentrations of

0.0 and 9.6 CFU/100 mL, respectively, and higher counts at site 9 (499.2 CFU/100 mL) (Figure

2B). The variance of enterococci did not differ across most of the sampled points, exhibiting a

standard deviation (SD) of 99.23-99.67 CFU/100 mL, except for site 10, which showed a SD of

21.2 CFU/100 mL. In terms of the phages, these were detected in lower concentrations

compared to the bacterial indicators. The lowest and highest means for coliphages correspond to

sites 1 and 9 (32.7 PFU/100 mL and 314.0 PFU/100 mL, respectively) (Figure 2C). The

variance for the coliphages differed across the sampling sites, in which the lowest and highest

SD correspond to sites 1 (6.3 PFU/100 mL) and 9 (139.1 PFU/100 mL), respectively. In terms of

the enterophages, the highest mean concentrations correspond to sites 3 and 7 (Figure 2D) and

variances did not differ across the sampling sites (SD = 4.9). CI for the bacterial and viral

indicators are represented in Figure 2.

94

Figure 2: Credible intervals (CIs) for thermotolerant coliforms (A), enterococci (B), coliphages

(C), and enterophages (D) in the Rio Grande de Arecibo watershed. Results represent the means

of the sampling period (n=13), and credible intervals, representing positive values, were drawn

as well. Except for enterophages, CIs could not be calculated for site 10 since most data are

zeros.

Detection of host-specific Bacteroides by PCR and enterococci by qPCR

Of the 13 samples tested per site, HSB were not detected at sites 1 and 2, were detected once at

sites 3, 6, 8 and 9, twice at sites 4 and 5 and three times at sites 7 and 10. CSB were detected

once at sites 1 and 9 and twice at sites 7 and 8, but were not detected in samples collected from

95

sites 2 to 6 and 10. Chicken-specific bacterial markers were detected once with both CP2-9 and

CP3-49 at sites 7 and 8, respectively (Figure 3).

Figure 3: Presence of human-, cattle-, and chicken-specific markers in the Rio Grande de

Arecibo watershed. Numbers represent the frequency of detection of the HF183, CF128, CP2-9,

and CP3-40 markers during the sampling period (n=13). Both chicken-specific markers were

detected only once during the sampling period.

The range of quantification (ROQ) for the enterococci qPCR was 10-8

to 10-12

g of genomic

DNA per reaction. The qPCR amplification efficiency ranged from 94.2 to 98.4 %, with R2

values ≥ 0.994. No signals were detected in the negative controls (i.e., no-template reactions)

indicating the absence of cross-contamination in this study. None of the samples showed

increases of signal intensity compared to the undiluted DNA templates, suggesting that PCR

inhibition did not interfere with the amplification efficiency. Enterococci were detected by qPCR

across all sites (Figure 4A). The lowest means correspond to sites 1, 2 and 3 (0.0 pg/100 mL)

and the highest correspond to site 7 (30.3 pg/100 mL). Variance was the highest at site 1

(SD=275.9 pg/100 mL), but remained constant throughout the rest of the sampled sites

(approximately 98.0 pg/100 mL). In terms of possible correlations between qPCR and culture

techniques for the detection of enterococci, a positive correlation between both methods was

found in this study (Figure 4B). This was the case for sites 6 and 7 (p= < 0.0001, R2= 0.78,

DF=12; p= 0.013, R2=0.39, DF=12), but no correlation was noted for samples collected from site

1.

96

Figure 4: qPCR for enterococci and correlation between qPCR and culture-based techniques for

enterococci. (A) Results show the mean genomic mass (pg)/100 mL (n=13) for enterococci

across the Rio Grande de Arecibo watershed and the corresponding credible intervals. (B)

Correlation between qPCR and culture- based methods for the detection of enterococci (site 7).

Correlations of indicators with rainfall and detection methods

At site 1, a positive correlation was noted with enterococci detected by culture methods and

rainfall reported 24 h, for 48 h, but not for one week prior sample collection. At sites 6 and 7, a

positive correlation was noted with the precipitation reported 24 h (Figure 5A), for 48 h and one

week prior sample collection. For enterococci qPCR data from sites 6 and 7, a positive

correlation was found at 24 h (Figure 5B), for 48 h and one prior sample collection, but this was

not the case for site 1 (Figure 5C). Results from the correlation analyses between enterococci

detected by culture methods or qPCR are shown in Table 3. No correlation was found with

thermotolerant coliforms and precipitation in any of the study sites (Figure 5D). Similarly, no

correlations were found with coliphages or enterophages and precipitation (Figure 5E and F). In

97

addition, positive correlations between thermotolerant coliforms and coliphages were found at

sites 1 and 7 (Nonparametric Spearman’s test: ρ= 0.45; p=0.014 and ρ= 0.85; p=0.049,

respectively). No other correlations between any of the microbial indicators detected by culture

methods were noted.

98

Figure 5: Correlation between rainfall and microbial indicators. Precipitation data were collected

from sites 1, 6, and 7 due to the proximity of a USGS station. Results show the possible effect of

precipitation reported 24 h before collection of the samples on enterococci detected by culture

methods (site 7) (A), 24 h before collection of the samples on enterococci detected by qPCR (site

7) (B), 24 h before collection of the samples on enterococci detected by qPCR (site 1) (C), 48 h

before collection of the samples on thermotolerant coliforms (site 1) (D), 24 h before collection

of the samples on coliphages (site 7) (E), and 24 h before collection of the samples on

enterophages (site 7) (F).

DISCUSSION

Prevalence of the bacterial and viral indicators detected by culture methods

Detection of thermotolerant coliforms throughout the watershed suggests an input of fecal

matter, although it is relatively difficult to identify the possible sources. High concentrations of

these indicator bacteria at sites 4 and 7 may suggest an inefficient removal by sewage treatment

or an additional input of fecal matter, but future studies are needed to confirm this. Moreover,

rainfall may play an important role in the loading dynamics. For instance, when no precipitation

was reported at site 1, thermotolerant coliforms ranged between 0 to ~40 CFU/100mL and 1

CFU/100mL was detected when 10.2 mm of rain were reported 24 h prior to collecting the

samples (data not shown). This may suggest that rainfall may have a possible dilution effect on

the thermotolerant coliforms. Interestingly, 0 to ~ 35 CFU/100mL were detected when no

precipitation was reported, but ~ 40 CFU/100mL were detected when >25.4 mm of rain were

reported for 48 h prior to sample collection at site 1. This may suggest that tropical sediments

may be a source of thermotolerant coliforms and these may have been resuspended due to the

rainfall reported in the previous 48 h. However, future studies are still needed to determine the

prevalence of these indicator bacteria in tropical sediments and other possible loading

mechanisms after rainfall events. A similar pattern was noted with thermotolerant coliforms and

rainfall reported for 24 and 48 h prior to collecting the samples at sites 6 and 7, but unlike site 1,

high numbers of these bacteria could also be associated with their transport from higher sites of

the watershed. Additional studies are needed to determine the possible transport mechanisms of

these indicator bacteria in tropical watersheds. No differences in the concentrations of

thermotolerant coliforms were noted when correlated with the rainfall reported for one week

99

prior sample collection. This suggests that different periods of precipitation may differently

influence the numbers of these bacteria when detected by culture methods.

It has been suggested that some enterococci are naturally-occurring in tropical waters [2]. Results

presented here suggest that many Enterococcus spp detected by culture methods are from a fecal

origin. The reason for this is that site 1 is one of the highest sites of the watershed, possibly one

of the less impacted by human or animal activities and very low numbers were detected. These

results are consistent with previous studies in which lower numbers of indicator bacteria have

been detected in inland lakes [38]. Higher enterococci numbers at sites 4 and 7 suggests that

these sites are point-sources of fecal pollution. Interestingly, site 10 represents a point-source of

fecal contamination as well, but low numbers may be due to an efficient removal by sewage

treatment. Also, the possibility remains that rainfall may also contribute to the input of

enterococci into tropical surface waters. Even though rainfall did not correlate with enterococci

detected at site 1, a positive correlation was noted with rainfall reported for one week prior to

sample collection at site 6. In addition, a positive correlation between enterococci and rainfall

reported 24 and for 48 h and one week prior to sample collection at site 7 suggests that loading

of enterococci into surface waters may be influenced by precipitation events. Based on previous

studies, it is possible that sediments and runoff could contribute to the input of these bacteria into

surface waters, but future studies are still needed in order to confirm if this is the case for waters

in Puerto Rico. Results presented here are consistent with previous studies in which higher

numbers of enterococci are detected after rainfall events in Hawaiian marine waters [39].

However, previous studies have not considered the effect of various precipitation periods prior to

sample collection. The present study suggests that various rainfall episodes may be considered

prior to inferring a fecal input in tropical surface waters. However, the possibility remains that,

although positive correlations were noted, rainfall may not always result in an increase in

enterococci numbers.

Both coliphages and enterophages were detected throughout the watershed, indicating that the

sampled sites may be impacted by human or animal fecal matter at some extent. The high

concentrations of coliphages (compared to enterophages) suggest that more than one source

could be contributing to their input and/or that they are present in higher numbers in feces. The

100

low concentrations of enterophages throughout the watershed may suggest that: (i) these are

present in low concentrations in human feces, (ii) few sources of human fecal pollution are

contributing to the input of these bacteriophages (e.g. septic tanks or domestic WTP), (iii)

enterophages are able to survive for short periods in tropical fresh waters and/or (iv) may be

diluted as a result of precipitation events. Even though enterophage concentrations were

relatively low, further studies need to compare their numbers with those of enteric viruses under

similar conditions since a reliable indicator should be at least as abundant as the pathogen of

concern. Also, the possibility remains that rainfall may have a possible dilution effect on both

phages. The reason for this is that up to 1000 and 50 coliphage and enterophage PFU/100mL,

respectively, were detected when no rainfall was reported and approximately 300 and 0 PFU/100

mL, respectively, were detected when >38.1 mm of rain were reported.

Monitoring indicator bacteria by molecular-based techniques

Human and animal fecal contaminations are among the major concerns to public health since

pathogens could be present [40-42]. Therefore, it is important to validate source-specific

markers, particularly in the tropics, as most studies were done in temperate regions. One

promising tool for indicating human fecal contamination is the Bacteroides marker HF183 [43,

44]. In the present study, HSB were mostly detected in samples collected downstream of all three

domestic WTP, suggesting that their presence is the result of the input of human fecal matter.

This supports the specificity of the HF183 marker in tropical inland waters with human fecal

impact. In terms of the cattle-specific markers, these have been shown to be a promising assay to

detect ruminant pollution in environmental waters. However, some studies have shown that CF

128 cross reacts with DNA extracts from various farm animals [45]. In the present study, CSB

were detected at sites 1, 2 and 3, in which cattle is not present, but it was expected to detect CSB

at site 6 since cattle grazing takes place. Site 5, which could be impacted by horse fecal matter,

tested positive for CSB by using the CF128 marker and this is comparable with previous studies

in which CF128 amplified CSB DNA in horse fecal matter [20]. Similarly, chicken-specific

Bacteroides were not detected using CP2-9 or CP3-49 in waters impacted by chicken fecal

matter. Sample sites 4, 5 and 7 were considered to be impacted by chicken-fecal matter, but

chicken-specific Bacteroides were detected once at site 7 during the sampling period and

chicken-specific Clostridium were detected once in samples collected from site 8. However, it

101

should be noted that the number of chickens in these sites are relatively low, suggesting that the

assays may not be sensitive enough to detect low levels of poultry pollution.

qPCR is currently being considered as a possible rapid detection method to be incorporated into

USEPA guidelines; therefore, it is important to test this technique in different water types prior

to implementation. Positive correlations between qPCR and culture methods in temperate waters

suggest that detection of enterococci by qPCR could also be used as a tool for determining

health-related risks and that both techniques may be reliable for the detection of enterococci [46].

In the present study, enterococci genomic mass detected by qPCR was found to be higher at sites

with known point-sources of fecal pollution. However, detection of the multiple copies of the

23S rRNA gene present in the bacterial genome, viable-non-cultivable bacteria, as well as DNA

from dead cells may add to the apparent increase in enterococci genomic mass [47]. Rainfall may

also contribute to the increment of enterococci detected by qPCR. At sites 6 and 7, a positive

correlation with rainfall may suggest that it may have an indirect contribution to the input of

Enterococcus into tropical surface waters (e.g. septic tanks overflows and animal droppings).

However, as with culture methods, it is difficult to predict that rainfall may always have a

positive correlation with enterococci detected by qPCR.

Correlations between culture-based and qPCR methods in this study may suggest that either of

the techniques may be used to detect enterococci in tropical inland waters. Correlation results in

this study are among the few performed for tropical inland waters, but similar outcomes have

been reported across the US [38]. Specifically, both methods were positively correlated in

Hawaiian marine waters, but this correlation was not noted for fresh waters and estuaries [39].

Results suggest that qPCR may not be suitable for all water types since different outcomes are

obtained from both methods and a correlation may not be expected in various water types at all

times. Future studies need to determine if correlations between qPCR and culture methods in

tropical inland waters exist throughout the year. Correlations between molecular and culture

techniques and between thermotolerant coliforms and coliphages suggest that the bacteriological

and virological quality of tropical inland waters should be evaluated using diverse MST tools.

This opens up the possibility of using a toolbox whenever determining the microbiological

quality of tropical inland waters.

102

CONCLUSIONS

Few studies have evaluated the microbial quality of tropical inland waters. One problem is the

need to develop appropriate water quality standards for tropical waters since guidelines used in

other areas may not be applicable. The present study tested combinations of markers to assess the

microbial quality of a tropical watershed in Puerto Rico, but results presented here need to be

further tested in other tropical watersheds. Even though a different combination of MST

techniques was tested in this study, there is still a need of developing a more robust toolbox to

infer fecal pollution and the possible sources in various tropical water types. In terms of the

bacterial indicators, these alone may not be suitable when inferring fecal contamination or its

source in tropical inland waters. Additional studies are needed to further examine the loading

dynamics of indicator bacteria into tropical fresh waters as a result of rainfall events.

Enumeration of coliphages and enterophages may be a more appropriate way to infer fecal

contamination since these are not ubiquitous in tropical waters and their numbers do not increase

after rainfall events. In terms of the host-specific markers in this study, only HF183 seemed a

reliable means of inferring the targeted source of fecal contamination in tropical waters. Cattle

and chicken-specific Bacteroides and Clostridium markers may not distinguish the source of the

fecal pollution in tropical waters; therefore, additional markers that could more reliably detect

cattle and chicken-specific sources of fecal pollution in Puerto Rico remain an important need.

Positive correlations between culture and molecular methods in this study should be considered

prior to implementing qPCR as a rapid method for inferring fecal pollution in the tropics since it

may not apply to all water bodies, specifically to inland lakes. qPCR assays targeting enterococci

in tropical waters must be carefully considered and further tested in other tropical areas since

rainfall may overestimate the numbers of enterococci detected by this method. The methods and

results presented here could be potentially useful in other water types.

ACKNOWLEDGMENTS

We thank the USGS for the precipitation data, the Department of Natural Resources of Puerto

Rico for information about the Rio Grande de Arecibo watershed and Gwendolyn Argüello and

Jean F. Ruiz for processing the samples. We also thank Dr. Pablo A. Ortiz-Pineda for managing

the figures at the modification stage. This research was partially supported by MBRS-RISE (NIH

Grant Number 2R25GM061151-09) and the US Environmental Protection Agency.

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SUPPLEMENTARY INFORMATION

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Supplementary Figure 1: Enterococci and thermotolerant coliforms CFU/100mL in the Rio

Grande de Arecibo watershed. Panels (A) and (B) represent the concentration of enterococci

during a period of high and low rainfall events, respectively. Panels (C) and (D) represent the

concentration of thermotolerant coliforms during the same periods of rainfall. Minimum and

maximum values are represented by error bars.

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Supplementary Figure 2: Enterophages in the Rio Grande de Arecibo watershed according to

the incubation temperature and rainfall period. Figure shows enterophages that replicate at 22

(A), 37 (B), 41 (C) and 45°C during the periods of November to January and February to April.

Standard deviations are shown by error bars.

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/10

0m

L (

Lo

g1

0)

Sample sites

Nov-Jan

Feb-Apr

22ºC

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ha

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L (

Lo

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37ºC

108

(C)

(D)

Supplementary Figure 3: Coliphages in the Rio Grande de Arecibo watershed according to the

incubation temperature and rainfall period. Figure shows enterophages that replicate at 22 (A),

37 (B), 41 (C) and 45°C during the periods of November to January and February to April.

Standard deviations are shown by error bars.

0.00

0.50

1.00

1.50

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1 2 3 4 5 6 7 8 9 10

Co

lip

ha

ges

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00

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109

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113

CHAPTER 5

Evaluation of Enterococcus-infecting phages as indeces of fecal pollution

T. M. Santiago-Rodriguez1, P. Marcos

2, S. Monteiro

3, M.Urdaneta

1, R. Santos

3 and G. A.

Toranzos1


Rico, San Juan, Puerto Rico, 2Biological Sciences Department, General Studies, University of

Puerto Rico, San Juan, Puerto Rico, 3Laboratório de Análises, Instituto Superior Técnico, Lisboa,

Portugal

ABSTRACT

No microbial source tracking tool satisfies all the characteristics of an ideal indicator of human

fecal pollution. For this reason, the potential of Enterococcus faecalis phages (enterophages) as

markers of this type of contamination was tested by using eight Enterococcus type strains as the

possible hosts. The prevalence of enterophages in animal feces and domestic sewage were

determined, as were the inactivation rates in raw sewage at 4 ºC and surface and tap waters at 22

ºC. Enterophages were exclusively detected in raw sewage (up to 66.0 PFU/100 mL), suggesting

a strictly human origin; and exhibited inactivation rates of approximately 0.002 to 0.05, 0.3 to

0.5 and 0.4 to 1.4 log/day in raw sewage and surface and tap waters, respectively, similar to

those of previous reports on human enteric viruses under similar conditions. Interestingly, phages

infecting other Enterococcus type strains were detected in both animal feces and domestic

sewage in concentrations of up to 335.8 and PFU/g and 96.0 PFU/100 mL, and certain phage

isolates infected several of the strains tested. This clearly indicates the possible promiscuous

nature of some Enterococcus phages and thus opens up the opportunity to further characterize

these as indices of specific fecal sources.

Keywords: enterococci, enterophages, fecal pollution, microbial indicators, Microbial Source

Tracking

Reference: Santiago-Rodriguez TM, et al. Evaluation of Enterococcus phages as indeces of

fecal pollution. Journal of Water and Health. 2013. 11(1): p. 51-63.

114

INTRODUCTION

Human pathogens can be introduced into water sources as a result of fecal pollution. Among

these, enteric viruses are of great concern since they: (i) can cause disease with infectious doses

as low as 1 to 10 particles, (ii) affect 5 to 18 million people every year and (iii) can be

resuspended from sediments to surface waters after disturbances [1-4]. Enteric viruses are often

detected by culture and molecular techniques [5-8]. However, culture methods are notoriously

expensive and laborious and noninfectious viral particles or naked viral nucleic acids can be

detected by molecular techniques [9, 10]. Since not all laboratories have the facilities for enteric

virus detection, microbial indicators have been used to infer the presence of these pathogens in

waters and sewage; however, many microbial indicators fail to fulfill the characteristics of a

reliable marker of fecal pollution [11]. For instance, relatively large numbers of both enterococci

and thermotolerant coliforms have been detected in pristine waters and are able to replicate in

these environments [12-14]. Algae and sediments could also be sources of indicator bacteria and

although certain type strains have been linked to fecal pollution, their presence in the feces of

different warm-blooded animals makes them unreliable for microbial source tracking (MST)

purposes [15]. In addition, enteric viruses can be present in waters that meet bacteriological

standards and absent from waters failing to meet this criteria [1].

Bacteriophages have also been proposed as markers of fecal contamination and as models of

human enteric viruses. Such is the case of phages infecting Bacteroides fragilis and Escherichia

coli (coliphages). However, B. fragilis phages have been detected only at certain geographical

regions [16-19] and the techniques used for their detection are relatively difficult due to the strict

anaerobic nature of the bacterial host [20, 21]. Coliphages may not discriminate the source of the

fecal contamination since these have been detected in the feces of different animals [22, 23]. In

addition, survival experiments have been performed with these bacteriophages [20, 24-26].

Coliphages can survive for long periods in pristine waters and may not necessarily indicate

recent fecal pollution [24, 26, 27].

There is a lack of microbial indicators of human fecal pollution. For this reason, we have

proposed phages that infect a specific type strain of Enterococcus faecalis (enterophages) as

markers of human fecal contamination. Most enterophage isolates characterized in our laboratory

115

are tailed phages, with icosahedral capsids measuring between 13 to 80 nm and DNA genomes

>23 kbp [28, 29]. In terms of their reliability as indicators of human fecal pollution,

enterophages: (i) have been detected in raw and treated domestic sewage [28, 29], (ii) possess a

survival similar to that of many enteric viruses in fresh waters [2, 30] and (iii) have not been

detected in cattle feces. Even though previous results have been promising, more data are needed

to fully determine the potential of enterophages as indicators of human fecal contamination. The

present study focused on the use of various Enterococcus species (spp.) as the bacterial hosts,

and fecal material from different animals and domestic sewage were tested for the presence and

prevalence of enterophages. We also determined the inactivation rates and survival times of

enterophage isolates under various conditions, and compared results to those studies using

human enteric viruses. Therefore, the aims of the present study were to: determine the presence

of enterophages in animal feces, their prevalence in raw and treated domestic sewage,

inactivation rates and survival times under various conditions, including raw sewage at 4 ºC,

fresh and tap waters at 22 ºC and various sterile water types at 37 ºC.


Detection of Enterococcus phages in feces and sewage

Poultry, cattle, pigs and humans are among the most common sources of fecal pollution [31, 32].

For this reason, the presence of enterophages in animal feces and domestic sewage was tested by

using Enterococcus type strains from the American Type Culture Collection (ATCC) and

included: E. faecalis (ATCC 19433), E. faecium, E. gallinarum, E. durans (ATCC 19432), E.

dispar (ATCC 51266), E. hirae (ATCC 8043), E. casseliflavus (ATCC 25788) and E.

pseudoavium (ATCC 49372) as the possible bacterial hosts. In addition, the presence of

Enterococcus phages in dog feces was tested since close contact with humans and lateral

transmission of microorganisms can occur [33]. Samples were also processed for coliphages

using E. coli C3000 (ATCC 15597) as the bacterial host as a means of comparison. Composites

of the fecal samples were processed by using the single layer method: Briefly, one gram of

chicken (n=30), cattle (n=30), pig (n=10) and dog (n=12) feces were eluted in 50 mL of a saline

solution (0.85% w/v) per gram of feces and mixed with an equal amount of liquefied media as

described previously [29]. In order to detect the possible presence of different bacteriophage

thermal groups, the mixture was poured into four Petri dishes and incubated at 22, 37 and 41 °C

116

and viral plaques were enumerated at 24 h. The presence of Enterococcus phages was tested in

raw sewage and primary effluent from three wastewater treatment plants (WTP) in Puerto Rico.

Samples were collected and processed monthly from August 2010 to August 2011 using the

single layer method with a modification [28]. Sixty-mL aliquots were added to equal amounts of

molten media and processed individually by using the Enterococcus type strains described above

as the bacterial hosts. Petri dishes were incubated at 22, 37 and 41 °C, viral plaques were

enumerated at 24 h and the percent removal of each phage group was calculated.

Isolation and characterization of Enterococcus phages

Individual viral plaques were isolated as described previously [28]. Several of the isolates were

tested against different Enterococcus type strains using a spot test to determine the host

specificity and replication temperature. Briefly, 1 μL of the bacteriophage isolate was placed on

top of a Petri dish containing Trypticase Soy Broth (TSB) (Difco), agar (1.5% w/v), CaCl2·2H2O

(Fisher Scientific Co. NJ, USA) and NaN3 (MCB, OH, USA) (final concentration of 2.6 mg/mL

and 0.4 mg/mL, respectively) and a lawn of the bacteria tested. A total of three dishes were

incubated at 22, 37 or 41 °C for 24h. In addition, the morphology of two E. faecalis phages was

determined by using Transmission Electron Microscopy, as described previously [28].

Inactivation rates and survival studies

Current regulations require that water and sewage samples be processed within 6 h and stored at

4-7 °C prior to analysis. To test if storage time could affect the detection of Enterococcus

phages, we determined the inactivation rates of phages infecting E. faecalis, E. casseliflavus, E.

faecium and E. coli in raw sewage. One-liter of raw domestic sewage was seeded with a

concentration of 104

plaque forming units (PFU)/100 mL from prepared laboratory stocks and

kept at 4-7 ºC. Aliquots were obtained daily and processed using the single layer method as

indicated above. To simulate the survival of enterophages and coliphages under environmental

conditions, raw domestic sewage was added to three fresh water samples collected from the Rio

Grande de Arecibo watershed in Puerto Rico [34, 35]. Experiments were performed during a

period of low and higher rainfall events and included: (i) the highest and thus one of the less

polluted sites of the watershed (site 1), (ii) immediately after a WTP (site 2), since it represents a

point-source of fecal contamination and (iii) the estuary (site 3). The survival experiments were

117

conducted as described above and seeded samples were kept at 22 °C, average temperature of

waters in Puerto Rico [29]. All precipitation data was obtained from the US Geological Survey

(USGS) Caribbean Water Science Center, stations 50020100, 50024950 and 50021700

(http://pr.water.usgs.gov/). Similarly, for the survival of enterophages and coliphages in tap

water, 2 L were collected in a sterile container and seeded with enterophages and coliphages and

aliquots were processed as described [28]. In order to determine the possible effect of chlorine in

the survival of enterophages and coliphages, one of the samples was dechlorinated using sodium

thiosulfate (final concentration of 10.0 mg/L) and kept at 22 °C. Similarly, survival experiments

with an enterophage isolate were performed at 37 °C (the optimal growth temperature of the host

and the isolation temperature of the enterophage tested) in sterile tap, distilled and wastewater, as

described previously.

Results for all inactivation rates and survival experiments were plotted as semi-log plots of time

(days) versus bacteriophage PFU/100 mL. Decay constants (kd) were calculated by using the

slopes of the linear regressions of the semi-log plots (-log10 PFU·day-1

) [29, 36]. T90 values, or

the time to reach a 90 % reduction in PFU densities, were calculated as ln (0.1)/kd [34, 37, 38].


Non-parametric one-way analyses of variances (Kruskal-Wallis) were used to determine

differences in the prevalence of Enterococcus phages in raw sewage as influenced by the

bacterial host tested and incubation temperature. The same analyses were used to determine

differences in the inactivation rates of the Enterococcus phages (E. faecalis, E. faecium and E.

casseliflavus) and coliphages in raw sewage at 4 ºC [23, 39]. For the survival experiments in

fresh waters, the analyses were used to determine differences in the inactivation rate of

enterophages and coliphages as influenced by the incubation temperature and sampled site.

Statistical analyses were performed using the R statistical software (version 2.11.1) [40].

RESULTS

Enterococcus phages in animal feces and domestic sewage

Interestingly, phages infecting E. faecium, E. casseliflavus and E. pseudoavium were detected in

chicken feces at 37 °C (Table 1). Phages infecting E. gallinarum, E. durans, E. dispar and E.

http://pr.water.usgs.gov/

118

hirae were not detected in chicken, cattle or pig feces. Coliphages were detected in chicken

(1.0±0.0 PFU/g) and cattle feces at 22 and 37 °C (40.3±28.0 and 335.81±4.0 PFU/g,

respectively) and in dog feces at 41 °C (120.79±29.45 PFU/g). Neither coliphages nor

Enterococcus phages were detected in pig feces. Enterophages were detected at all temperatures

tested in raw sewage collected from all WTP and at 22 and 37 °C in the primary effluents from

two WTP. Interestingly, E. faecium-infecting phages were also detected in raw and treated

sewage from two WTP at all temperatures tested.

Host

Source E. coli E. faecalis E. faecium E. casseliflavus E. pseudoavium

Chicken 1.0±0.0a ND 107.5±10.6

b 65.0±0.0

b 3.8±1.8

b

Cattle 40.3±28.0a ND ND ND ND

Dogs

335.81±4.0b,

120.79.±29.45c ND ND ND ND

Pigs ND ND ND ND ND

ND=Not detected

a=22 ºC

b=37 ºC

c=41 ºC

Table 1: Enterococcus and E. coli-infecting phages per gram of feces in chicken (n=30), cattle

(n=30), dogs (n=12) and pigs (n=10). Enterococcus spp, included: E. faecalis, E. faecium, E.

gallinarum, E. hirae, E. durans, E. dispar, E. casseliflavus and E. pseudoavium. Only hosts

exhibiting positive results in at least one fecal source were included in the table.

Higher concentrations of both E. faecalis and E. faecium phages were detected in domestic

sewage compared to other enterococci phages and exhibited removal percents of 37.0 to ≥ 99.0

% and 47.0 to ≥ 99.0 %, respectively. Other Enterococcus phages exhibited various removal

percents depending on the WTP and incubation temperature (Table 2). Phages infecting E.

casseliflavus were detected at 22 and 37 °C in raw sewage collected from all WTP and at 22 and

37 °C in the primary effluents. Enterococcus pseudoavium-infecting phages were also detected at

22 °C in raw sewage from all WTP and at 37 °C in two WTP, but were not detected in the

primary effluents.

119

PR-A PR-B

Host 22ºC 37 ºC 41 ºC 22 ºC 37 ºC 41 ºC

E. faecalis 37.0±44.0 44.0±33.0 ≥99.0±0.9 83.0±16.8 80.0±15.9 ≥99.0±0.09

E. faecium 63.0±16.0 86.0±5.8 ND 47.0±42.0 60.0±37.0 ≥99.0±0.09

E. gallinarum ND ND ND ND ND ND

E. hirae ND ND ND ≥99.0±0.09 ≥99.0±0.9 ND

E. durans ND ND ND ≥99.0±0.09 ≥99.0±0.09 ND

E. dispar ≥99.0±0.09 ND ND 48.0±25.2 ≥99.0±0.45 ≥99.0±0.09

E. casseliflavus 27.0±32.4 ≥99.0±0.9 ND 81.0±12.1 ≥99.0±0.09 ≥99.0±0.09

E. pseudoavium ≥99.0±0.09 ≥99.0±0.9 ND 87.0±1.0 ≥99.0±0.09 ≥99.0±0.09

PR-C

Host 22 ºC 37 ºC 41 ºC

E. faecalis ≥99.0±0.09 ≥99.0±0.09 ≥99.0±0.09

E. faecium ≥99.0±0.09 ≥99.0±0.09 ≥99.0±0.09

E. gallinarum ND ND ND

E. hirae ≥99.0±0.09 ND ND

E. durans ND ND ND

E. dispar ≥99.0±0.09 ND ND

E. casseliflavus 73.0±20.4 ≥99.0±0.09 ND

E. pseudoavium ≥99.0±0.09 ND ND

Table 2: Mean removal percents of Enterococcus phages in primary effluent from three WTP

(PR-A, PR-B and PR-C) in Puerto Rico. Numbers were calculated for different incubation

temperatures, namely 22, 37 and 41 °C. Phages infecting several Enterococcus were not detected

(ND).

Other phages detected in raw sewage were those infecting E. hirae, E. durans and E. dispar, but

these were not constantly detected in primary effluent (Figure 1).

120

(A)

(B)

-1.00

-0.50

0.00

0.50

1.00

1.50

2.00

2.50

3.00

3.50

faecalis faecium gallinarum hirae durans dispar casseliflavus pseudoaviumBacte

rio

phage P

FU

/10

0m

L(L

og

10)

Enterococcus host

22ºC

-1.00

-0.50

0.00

0.50

1.00

1.50

2.00

2.50

3.00

3.50

faecalis faecium gallinarum hirae durans dispar casseliflavus pseudoaviumBacte

rio

phage P

FU

/10

0m

L(L

og

10)

Enterococcus host

37ºC

121

(C)

Figure 1: Prevalence of Enterococcus phages in raw domestic sewage in three wastewater

treatment plants (WTP) in Puerto Rico. Petri dishes were incubated at 22 (A), 37 (B) and 41°C

(C) and each column represent a WTP. Enterococcus spp included E. faecalis, E. faecium, E.

gallinarum, E. hirae, E. durans, E. dispar, E. casseliflavus and E. pseudoavium. Results

represent the log transformed geometric mean of n=12 samples and standard deviations are

represented by error bars.

Replication and morphology of Enterococcus-infecting phages

Most of the Enterococcus phage isolates in this study did not infect a specific bacterial host, with

the exception of those infecting E. faecalis, which replicated at all three replication temperatures

tested as shown previously [29]. Table 3 shows the host range and replication temperature of the

tested phages. In terms of the morphology of the E. faecalis phages in this study, these exhibited

long non-contractile tails of approximately 180 nm and icosahedral capsids of approximately 80

nm (Figure 2). This morphology is similar to the enterophages characterized previously [29].

-1.50

-1.00

-0.50

0.00

0.50

1.00

1.50

2.00

2.50

3.00

3.50

faecalis faecium gallinarum hirae durans dispar casseliflavus pseudoavium

Bact

erio

phag

e P

FU

/10

0m

L (L

og

10)

Enterococcus host

41ºC

122

a=22 ºC

b=37 ºC

c=41 ºC

Table 3: Host range of Enterococcus-infecting phages at different incubation temperatures.

Table shows the source of the isolated phages and the Enterococcus type strains used. Stocks

tested contained only one phage type and were tested against other Enterococcus spp., as shown

in the following columns.

Figure 2: Enterococcus faecalis-infecting phages isolated from domestic sewage. Isolates

exhibited long non-contractile tails of approximately 200nm and icosahedral capsids of 80nm.

Bar=100nm.

Host

Range

Source

Host for

Isolation

E.

faecalis

E.

faecium

E.

gallinarum

E.

hirae

E.

durans

E.

dispar

E.

casseliflavus

E.

pdeudoavium

Sewage E. faecalis + a,b,c

- - - - - - -

E. faecium + a,b,c

+ a,b

- - - + a,b

- + a,b

E. hirae - - - + a,b,c

- + c - +

b,c

E. dispar + a,b,c

- - - - + b,c

- + a,b

E. casseliflavus - - - + b,c

- + b +

a +

a,b,c

E. pseudoavium + b - - +

c - +

b - +

a,b

Fresh

water E. faecalis + a,b,c

- - - - - - -

E. faecium - + b - - - - - +

a,b

Poultry E. faecium - + b - - - +

a,b - +

a,b

E. casseliflavus - + a,b

+ a,b,c

+ a,b,c

+ a,b,c

+ a,b,c

+ a,b,c

+ a,b,c

E. pseudoavium - + a,b,c

+ a,b,c

+ a,b,c

+ a +

a,b,c - +

a,b,c

123

Inactivation rates and survival of Enterococcus phages

The initial titer of E. faecalis, E. faecium, E. casseliflavus and E. coli phages in raw sewage at 4

ºC remained constant for 6 days. Calculated decays for E. faecalis phages were 0.050 ± 0.030,

0.030 ± 0.030 and 0.080 ± 0.030 log·day-1

at 22, 37 and 41 °C, respectively. Enterococcus

faecium phages exhibited a decay of 0.0040 ± 0.0020 and 0.0020 ± 0.0020 log·day-1

at 22 and 37

°C, respectively. For this specific E. faecium phage isolate, 41 °C was inhibitory. Similarly, the

E. casseliflavus phage isolate in these experiments was able to replicate only at 22 °C and

exhibited a decay of 0.020 ± 0.030 log·day-1

. Coliphages exhibited decays of 0.010 ±0.020,

0.040 ±0.020 and 0.020 ± 0.050 log·day-1

at 22, 37 and 41 °C, respectively.

Survival of enterophages and coliphages in fresh waters

During the period of low rainfall events, up to 1.01 mm of rain was reported at site 1and no

precipitation was reported at sites 2 or 3 24 h prior to collecting the samples. During this period,

enterophages exhibited an average decay of 0.47 ± 0.03 log·day-1

at all sites. Coliphages

exhibited an average decay of 0.47 ± 0.03 log·day-1

at sites 1 and 2 and 0.33 ± 0.07 log·day-1

at

site 3. During the period of higher rainfall events, up to 2.54 mm of rain were reported at sites 1

and 3 and 0.20 mm were reported at site 2. During this period, enterophages exhibited an average

decay of 0.40 ± 0.00 and 0.30 ± 0.00 log·day-1

at sites 1 and 2 and 0.37 ± 0.03 log·day-1

at site 3.

Coliphages exhibited and average decay of 0.19 ± 0.12, 0.08 ± 0.03 and 0.20 ± 0.10 log·day-1

at

sites 1, 2 and 3, respectively. Calculated T90 values for enterophages and coliphages differed

during the period of low and high rainfall and across the incubation temperatures as well (Table

4). The survival of enterophages and coliphages during both rainfall periods are shown in Figure

3.

124

Table 4: T90 values for enterophages and coliphages in three fresh water samples. Samples were

collected from the Rio Grande de Arecibo watershed in Puerto Rico during a period of low and

higher rainfall events. Sites 1, 2 and 3 correspond to the unpolluted site, the point-source of fecal

pollution and the estuary, respectively.

(A)

Enterophages

Low rainfall High rainfall

Site 22°C 37°C 41°C 22°C 37°C 41°C

1 4.8±0.8 4.8±0.8 3.7±1.0 4.0±1.0 3.1±0.7 2.9±0.5

2 4.6±0.3 4.6±0.2 5.2±0.4 7.2±2.3 7.2±2.2 6.6±1.7

3 3.9±1.4 3.2±0.6 3.2±0.9 5.3±0.9 5.3±1.0 4.8±1.5

Coliphages


Site 22°C 37°C 41°C 22°C 37°C 41°C

1 5.2±0.5 4.6±1.4 4.7±0.9 32.2±2.6 10.3±1.3 6.9±0.9

2 5.1±0.002 5.0±1.4 4.6±0.7 45.3±9.2 22.2±13.0 25.0±5.0

3 8.1±4.1 7.0±2.2 5.8±3.3 24.0±15.2 12.2±5.0 6.5±2.0

-1.50

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Enterophages: None or low rainfall

125

(B)

(C)

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Enterophages: Higher rainfall

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Coliphages: None or low rainfall

126

(D)

Figure 3: Survival of enterophages and coliphages across a tropical watershed in Puerto Rico.

Sampled sites included the highest site (1), after a waste treatment plant (2) and the estuary (3).

Samples were collected during a period of low rainfall events and processed for the detection of

enterophages (A) and coliphages (B). The survival of enterophages (C) and coliphages (D)

during a period of higher rainfall events was also determined. Data represent those phages that

replicated at 37 ºC.

Enterophages in chlorinated and dechlorinated tap water and sterile distilled, tap and

wastewater

The enterophage isolate in this study survived approximately 12 days in chlorinated tap water

with an initial free chlorine concentration of 0.13 ± 0.02 ppm and more than 12 days in

dechlorinated tap water (data not shown). These exhibited decays of 0.23 ± 0.16, 1.05 ± 0.72 and

1.42 ± 0.75 log·day-1

in chlorinated tap water and 0.14 ± 0.045, 0.70 ± 0.38 and 1.02 ± 0.12

log·day-1

in dechlorinated tap water at 22, 37 and 41 ºC, respectively. Coliphages survived for

more than 12 days in both chlorinated and dechlorinated tap water (data not shown) and

exhibited a decay of 0.60 ± 0.0062, 0.83 ± 0.26 and 1.10 ± 0.20 log·day-1

in chlorinated tap water

at 22, 37 and 41 ºC, respectively. In dechlorinated tap water, coliphages showed a decay of 0.13

± 0.045, 0.70 ± 0.38 and 1.02 ± 0.12 log·day-1

at 22, 37 and 41 ºC. Calculated T90 values for

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Coliphages: Higher rainfall

127

enterophages and coliphages in both chlorinated and dechlorinated tap water are described in

Table 5. In terms of the survival experiments with the enterococci phage isolate at 37 ºC, kd

values for wastewater, tap and distilled water were 0.038, 0.36 and 0.37 log·day-1

, respectively.

A decay of approximately 2 log10 was observed when testing sterile wastewater during 43 days

(Figure 4A). On the other hand, when testing sterile tap and distilled water, enterophages

exhibited a notable decay, surviving approximately 11 days (Figure 4B and C). T90 values were

approximately 6 days for both tap and distilled water and 60 days for wastewater.

Table 5: T90 values (days) for enterophages and coliphages in chlorinated and dechlorinated

drinking water.

Figure 4: Survival of enterophages in various sterile water types at 37 ºC. Samples include (A)

wastewater, (B) tap (chlorinated) and (C) distilled waters and standard deviation is represented

by error bars.

Enterophages Coliphages

Temperature

(°C) Chlorinated Dechlorinated Chlorinated Dechlorinated

22 9.4±7.0 18.1±7.7 18.1±7.7 39.1±4.3

37 8.9±7.0 28.3±6.0 32.1±6.0 43.3±8.3

41 6.9±1.7 28.3±3.7 28.3±3.7 33.0±3.9

(A) (B)

(C)

128

DISCUSSION

Enterococcus-infecting phages in feces and domestic sewage

The source (human) and host-specificity exhibited by enterophages suggest that the E. faecalis

strain in this study may detect human-specific phages. The relative wide range of Enterococcus

spp. infected by specific phages showed that these have a promiscuous nature, as shown with the

coliphages elsewhere [41]. This in turn suggests that future studies are needed in order to

confirm if Enterococcus phages may recognize receptors which are shared by the enterococci

tested in the present study. Several of the Enterococcus strains tested also exhibited immunity to

certain phages, suggesting the presence of prophages, prophage remnants or Clustered Regularly

Interspaced Short Palindromic Repeats (CRISPR) [42, 43]. It is also possible that results may be

influenced by the culture media. Host range and replication temperature may be used to group

enterococci phages as those of an animal or human origin; this in turn suggests their potential for

MST purposes [44]. Results also suggest that humans are reservoirs of various enterococci phage

groups, as seen by the influence of the bacterial host used (x2 = 27.50, df = 6, p = 0.00012) and

temperature tested (x2 = 24.27, df = 2, p = 5.37 e-06). In addition, given that there is the need of

characterizing markers of chicken-fecal pollution, enterococci phages detected in chicken fecal

matter could be further characterized and tested in various water types impacted by this source of

fecal contamination. Future studies need to determine potential hosts for the detection of

Enterococcus phages in other animal feces.

In terms of the coliphages, previous studies have shown that their concentrations vary according

to each individual and can range from 100 to 10

7 PFU/g in cows, pigs and humans and < 10

PFU/g in dog feces [45, 46]. In the present study, coliphage concentrations in cows and humans

are within the range reported previously, but higher concentrations were detected in dog feces.

Although coliphages were not detected in pig feces in the present study, 0 to < 10 PFU/g have

been reported elsewhere [45].

Inactivation rates and survival of Enterococcus phages

Our results showed that titers of E. faecalis, E. faecium, E. casseliflavus and E. coli phages do

not decrease significantly at 4 °C (~0.002 to 0.05 log·day-1

, depending on the bacterial host),

129

suggesting that samples may be stored longer at refrigeration temperatures prior to analyses.

Similar outcomes have been reported for adenovirus 40 and 41 and poliovirus 1 in sewage at 4

ºC, which have exhibited a decrease of 2.5, 2.0 and 2.2 log after 50 days [47]. In fresh waters, the

similar replication of enterophages at all temperatures tested suggests that a group predominated

in the analyses. Unlike enterophages, coliphages exhibited differences in their replication at

different temperatures (x2 = 11.46, df = 2, p = 0.0033), suggesting that various groups were

present, as previously suggested [48]. In terms of the inactivation rates, enterophages did not

exhibit differences across the sampled sites during the period of low or high rainfall events. This

is comparable with the inactivation rates of poliovirus, coxsackievirus and rotaviruses SA11,

which exhibit similar inactivation rates in unpolluted and polluted sites (~ 0.5 to 1.0 log·day-1

)

[27]. Similarly, coliphages did not show differences in their inactivation rates across the sampled

sites during the period of low rainfall, but this was not the case for the period of higher rainfall

events (x2

= 10.11, df = 2, p = 0.0064). This is comparable with echovirus 7, which exhibits

different inactivation rates in unpolluted (1.0 log·day-1

) and polluted sites (0.5 log·day-1

) [27].

Although there are no hard data that may explain this, several factors may be involved in these

differences in survival times during both rainfall periods including pH changes, dilution of

proteolytic enzymes and antiviral chemicals, as described elsewhere [49-51]. Another possibility

is that due to the relatively long survival times of coliphages in surface waters (20 to 160 days)

and sediments (30 days) at 20 ºC, it is reasonable to believe that the opportunity to attach to

sediment particles is greater [26, 35]. Once attached, coliphages may be desorbed after rainfall

events, and thus their numbers may be higher. In terms of the estuary, polioviruses,

coxsackieviruses, rotaviruses SA11 and echoviruses can survive approximately 3 days in

estuaries, similar to enterophages during the period of low rainfall events in the present study,

but exhibit a more rapid decay compared to both enterophages and coliphages (~ 0.5 to 2.5

log·day-1

) [27].

Survival of the tested bacteriophages may also be influenced by chlorine and the environmental

microbiota. Few studies have determined the survival of enteric viruses in waters with chlorine

levels similar to those found in tap water [52]. Therefore, comparing the survival of enterophages

and coliphages with that of enteric viruses in chlorinated tap water is relatively difficult. Certain

enteric viruses can survive up to 30 minutes while others can survive 16 h and exhibit decays of

130

~ 1 log·h-1

in waters with free chlorine levels of 0.10 ppm at 22 ºC [53]. This suggests that

enteric viruses are more susceptible to chlorine than both enterophages and coliphages, which

exhibited T90 values of ~7 to 9 and18 to 32 days, and decays of ~ 0.20 to 1.40 and 0.60 to 1.10

log·day-1

, respectively. This in turn suggests that chlorine may have a more visible effect on the

survival of enterophages compared to coliphages. In dechlorinated tap water, T90 values for

enterophages detected at 37 and 41 ºC were similar to that of Poliovirus 1 at 23 °C

(approximately 30 days); however, neither the enterophages nor the coliphages possess a

survival similar to that of adenoviruses 40 and 41, under similar conditions [47]. These studies,

however, did not consider the effect of the environmental microbiota. Results from using

different sterile water types in the present study suggest that the absence of other microorganisms

enhances the survival of enterophages; therefore, future studies should consider the

environmental microbiota in the survival of enteric viruses and bacteriophages in various water

types.

CONCLUSIONS

This is one of the few studies in which the potential of enterophages as markers of human fecal

pollution has been further tested. The presence of enterophages in chicken, cattle, dog and pig

fecal materials and their prevalence in domestic sewage were determined by testing various

Enterococcus spp. as the bacterial hosts. This, however, may represent one limitation to the

present study as phages infecting other Enterococcus spp. may also be present in fecal material

and fecally contaminated waters and thus future studies are needed in order to determine this.

Humans and animals are reservoirs of Enterococcus phages and those specifically infecting E.

faecalis seem promising indeces of human fecal pollution. The present study also determined the

inactivation rates and survival times of enterophages and compared results with those of human

enteric viruses reported elsewhere under similar conditions; however, the present studies must

also be performed in situ. Enterophages could be considered models of certain human enteric

viruses in various water types since their inactivation rates and survival times are similar to those

reported previously. Phages infecting E. faecium, E. casseliflavus and E. pseudoavium

replicating at 37 °C may be used to infer the presence of chicken-fecal matter in fresh water

sources, but more studies need to determine if these phages are present in waters impacted by

chicken-fecal matter. The present study opens the possibility to further characterize

131

Enterococcus phages as indeces of specific fecal sources, especially in various geographical

regions and to predict their behavior in different areas by using specific mathematical models,

for example. Also, future studies need to develop the techniques for the detection of

enterophages using molecular methods and compare results with culture techniques.

ACKNOWLEDGMENTS

We thank Carlos Toledo-Hernandez, Cruz Minerva Ortiz of the Autoridad de Acueductos y

Alcantarillados for collecting the fresh water and sewage samples. We also thank Jorge W.

Santo-Domingo for providing the Enterococcus spp. type strains and the USGS for the

precipitation data. This research was partially supported by MBRS-RISE, NIH Grant Number

2R25GM061151-09, and the Center for Renewable Energy and Sustainability with a grant from

the Department of Defense to the University of Puerto Rico.

SUPPLEMENTARY INFORMATION

Decay (%)·day

-1 T90 (days)

Host 22°C 37°C 41°C 22°C 37°C 41°C

E. faecalis 4.7±3.1 3.4±2.7 8.3±3.6 49.0±29.7 67.1±154.3 27.8±13.8

E. faecium 0.4±0.2 0.2±0.2 N/D 579.0±300.5 1158.8±712.0 N/D

E. casseliflavus 1.9±2.7 N/D N/D 121.8±59.3 N/D N/D

E. coli 1.0±1.5 4.1±2.0 2.1±5.3 224.9±120.8 55.6±49.1 111.0±82.4

Supplementary Table 1: Average of the decay percent day -1

and T90 values for phages infecting

E. faecalis, E. faecium, E. casseliflavus and E. coli in raw sewage at 4°C. Both values were

calculated for phages detected at 22, 37 and 41 °C. Not all phages were detected at all incubation

temperatures and thus this is represented by N/D (not detected).

132

Enterophages


Site 22°C 37°C 41°C 22°C 37°C 41°C

1 0.5±0.04 0.5±0.04 0.4±0.06 0.4±0.1 0.4±0.1 0.4±0.1

2 0.5±0.03 0.5±0.02 0.5±0.02 0.3±0.1 0.3±0.1 0.3±0.1

3 0.4±0.006 0.5±0.02 0.5±0.02 0.4±0.1 0.4±0.2 0.3±0.1

Coliphages


Site 22°C 37°C 41°C 22°C 37°C 41°C

1 0.4±0.07 0.5±0.1 0.5±0.1 0.07±0.004 0.2±0.03 0.3±0.03

2 0.5±0.02 0.5±0.1 0.5±0.04 0.05±0.01 0.1±0.04 0.09±0.02

3 0.3±0.2 0.3±0.1 0.4±0.1 0.1±0.04 0.2±0.06 0.3±0.05

Supplementary Table 2: Decay values (log·day-1

) for enterophages and coliphages in three

fresh water samples. Samples were collected from the Rio Grande de Arecibo watershed in

Puerto Rico during a period of low and higher rainfall events. Sites 1, 2 and 3 correspond to the

unpolluted site, the point-source of fecal pollution and the estuary, respectively.

(A)

-0.50

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22ºC

133

(B)

(C)

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134

(D)

(E)

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135

(F)

Supplementary Figure 1: Survival of enterophages and coliphages in three sites in a watershed.

Samples were collected during a period of high rainfall events and included: a relatively

unpolluted site (1), after a WTP (2) and the estuary (3). Panels A, B and C correspond to the

survival of enterophages detected at 22, 37 and 41°C, respectively. Panels D, E and F correspond

to the coliphages detected at 22, 37 and 41°C, respectively. Results represent the average of three

replicas and standard deviations are represented by error bars.

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136

(A)

(B)

(C)

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(F)


watershed. Samples were collected during a period of low rainfall and include a relatively

unpolluted site (1), after a WTP (2) and the estuary (3). Panels A, B and C correspond to the

survival of enterophages detected at 22, 37 and 41°C, respectively. Panels D, E and F correspond

to the coliphages detected at 22, 37 and 41°C, respectively. Results represent the average of three

replicas and standard deviations are represented by error bars.

-0.50

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139

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144

CHAPTER 6

Antibiotic-resistance and virulence genes in Enterococcus isolated from tropical

recreational waters and sand

T. M. Santiago-Rodriguez1, J. I. Rivera

1, M. Coradin

1 and G. A. Toranzos

1


Rico, San Juan, Puerto Rico 00931-3360

ABSTRACT

The prevalence of enterococci harboring tetracycline and vancomycin-resistance genes, as well

as the enterococcal surface protein (esp) has mostly been determined in clinical settings, but their

prevalence in tropical recreational waters remains largely unknown. The present study

determined the prevalence of tetM (tetracycline-resistance), vanA and vanB (vancomycin-

resistance) in the bacterial and viral fractions, enterococci and their induced phages isolated from

tropical recreational marine and fresh waters, dry and wet sands. Since lysogenic phages can act

as vectors for antibiotic-resistance and virulence factors, the prevalence of the mentioned genes,

as well as that of an integrase-encoding gene (int) specific for Enterococcus faecalis phages was

determined. Up to 60 % and 54 % of the bacterial fractions and enterococci harbored at least one

of the tested genes, respectively, suggesting that bacteria in tropical environments may be

reservoirs of antibiotic-resistance and virulence genes. int was detected in the viral fractions and

in one Enterococcus isolate after induction. This study opens the opportunity to determine if the

presence of bacteria harboring antibiotic-resistance and virulence genes in tropical recreational

waters represents a threat to public health.

Keywords: antibiotic-resistance, bacteriophages, enterococci, enterococcal surface protein

Reference: Santiago-Rodriguez, et al. Antibiotic-resistance and virulence genes in Enterococcus

isolated from tropical recreational waters and sand. 2013. Journal of Water and Health.

Accepted.

145

INTRODUCTION

There is an increasing concern towards bacteria harboring antibiotic-resistance and virulence

genes in recreational waters. One of the main reasons is that it remains unknown if

microorganisms harboring these genes represent a risk to bathers. Among these, enterococci have

received great attention since these are ubiquitous to many aquatic ecosystems [1-5]. However,

certain Enterococcus spp. are also important opportunistic pathogens and infections have been

treated with tetracyclines. This has resulted in tetracycline-resistant enterococci and these may

harbor at least one of the tet determinants, being tetM one of the most studied in gram-positive

bacteria [6-8]. Resistant Enterococcus infections can be treated with vancomycin, but only as a

last resort due to the hazardous side effects [9]. However, enterococci have also acquired

resistance to vancomycin due to genes such as vanA and vanB, commonly described in E.

faecalis and E. faecium. In addition, enterococci exhibiting resistance to antibiotics can harbor

genes encoding for virulence factors, as in the case of the enterococcal surface protein (Esp),

described in E. faecalis and E. faecium [10-13]. The esp variant present in E. faecium was

proposed as a marker of human fecal contamination and there have been conflicting studies on

the usefulness of this gene as a species-specific marker [14-17].

Another concern is the transmission of antibiotic-resistant and virulence genes by mobile

elements, such as plasmids and transposons, but bacteriophages can also act as vectors of these

genes [18, 19]. Lysogenic phages can carry additional bacterial genes which could promote

fitness and virulence in the recipient bacteria [20-22]. These genes are often encoded by the host

and may be accidentally packaged into the phage capsid. It has been shown that the viral DNA

fraction of sewage and environmental waters harbor β-lactamase-encoding genes and that

lysogenic phages infecting E. faecalis may harbor genes that influence virulence in the bacterial

host [23, 24]. Most lysogenic phages infecting enterococci described so far belong to the

Siphoviridae family (dsDNA) and one possible approach to identify these is by amplifying

integrase-encoding genes (int). Integrases are considered markers of lysogeny and are involved

in the integration of the phage genome into that of the bacterial host [25]. Detection of phage

integrases within an ecosystem may provide insights as to what possible risks exist within a

specific ecosystem [26].

146

The prevalence of tetM, and vanA and vanB has mostly been determined in animal husbandry

facilities, and temperate and subtropical recreational waters, respectively [5, 27]; and although

the prevalence of esp has been determined in tropical regions, few studies are still available [28].

In addition, the presence of antibiotic-resistance and virulence genes in the bacteriophage

fraction of recreational waters, and integrases as a way to identify these, remains largely

unknown. Therefore, the main aim of the present study was to determine the prevalence of tetM,

vanA, vanB, and esp variants present in E. faecalis and E. faecium, as well as int, in the bacterial

and DNA viral fractions, enterococci and their lysogenic phages in tropical recreational waters.


Study sites

Samples were collected from OP, CBB, LC and RP in Puerto Rico (Figure 1). OP and CBB are

beaches located in the Atlantic Ocean, alongside the northern area of the island and are heavily

used by bathers year-round. LC is a man-made lake whose influents are three of the island’s

major rivers, serves as a major water reservoir for the San Juan metropolitan area and is also

used for recreational purposes. RP receives the input of minor rivers and streams used for

recreational activities. One-liter grab samples of marine (OP, n=35; and CBB, n=13), and fresh

(LC, n=8; RP, n=10) waters were collected in sterile plastic bottles from July 2011 to May 2012.

In addition, 100 g of wet and dry sands from OP (n=16 and n=15, respectively) and CBB (n=14

and n=11, respectively) were collected since sand may act as a reservoir of bacteria harboring

antibiotic-resistance and virulence genes [29, 30]. All water and sand samples were stored at 6-8

ºC and processed within 6 h.

DNA extraction of the bacterial fraction and enterococci

To determine the presence of the mentioned genes in the bacterial fraction, 100 mL were

processed by membrane filtration (0.4 μm pore size, 47 mm diameter) (GE Water and Process

Technologies, Trevose, PA). The filter was placed in 25 mL of Azide Dextrose Broth (Difco)

and incubated at 37 ºC for 24 h. DNA was extracted from 1 mL by using the Fermentas GeneJet

Genomic DNA Purification Kit following the manufacturer’s instructions. Alternatively, filters

were placed in tubes containing 0.3 g of acid-wash glass beads (Sigma) and stored at -20 ºC until

processed. For this, 600 μL of AE buffer were added, placed on a bead-beater for 60 s at

147

maximum speed and centrifuged for 1 min at 14,000 rpm. The supernatant was collected, re-

centrifuged for 5 min at 14,000 rpm to remove any remaining debris and stored at -20 ºC until

processed [31]. For the sand samples, 10 g were directly added into 20 mL of Azide Dextrose

Broth, incubated at 37 °C for 24 h and 1 mL was used for DNA extraction using the Fermentas

GeneJet Genomic DNA Purification Kit. Alternatively, 10 g were eluted in 120 mL of 0.1%

Tween20, 100 mL of the elution were filtered and DNA from the membranes was extracted as

described above.

Figure 1: Sampled sites in this study. Sites included CBB ( ), OP ( ), RP ( a), and LC ( ) in

Puerto Rico. Marine water and dry and wet sands were collected from CBB and OPB. Fresh

water was collected from RP and LC.

Atlantic Ocean

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Enumeration of enterococci was performed by using membrane filtration and m-Enterococcus

agar (Difco) as previously described [32, 33]. For the detection of enterococci in sand, 10 g were

eluted in 0.1% Tween20 followed by filtration of the resulting solution and membranes were

placed on m-Enterococcus agar as described. After 24-48h incubation at 37 ºC, individual

colonies were picked and transferred onto m-Enterococcus plates containing tetracycline (final

concentration 16 μg/mL) or vancomycin (final concentration 20 μg/mL). Colonies growing in the

presence of these antibiotics were picked and enriched in 1 mL of Azide Dextrose Broth for up

to 48 h at 37 ºC and DNA was extracted as described above. DNA quality and concentration was

estimated by using a NanoDrop® (ND-1000) spectrophotometer.

Virus concentration and DNA extraction

Filtrates from above were recovered, concentrated and purified for the detection of the

mentioned genes in the viral DNA fractions. Briefly, a final concentration of 0.2 % chloroform

(v/v) was added to the viral suspensions and kept at room temperature for 30 min to eliminate

any remaining viable bacteria. Solid NaCl (Fisher Scientific, enzyme grade ≥ 99.9 %, NJ, USA)

was added to the suspensions at a final concentration of 0.5 M and stored at 4 °C for 1 h. Any

remaining bacterial debris was removed by centrifugation at 8,500 rpm for 10 min at 4 °C and

the supernatant was transferred to sterile Oakridge tubes. PEG 8000 (Promega) was dissolved in

the phage suspension in a final concentration of 10 % and stored at 4 °C for 24 h. Bacteriophages

were sedimented at 14,000 rpm at 4 °C for 15 min and the supernatant was discarded. The phage

containing pellet was resuspended in 0.5 mL of PBS and left overnight at 4 °C. The residual PEG

and bacterial debris were removed by adding an equal volume of chloroform to a Phase Lock Gel

tube (5 Prime Inc., MD, USA) and centrifuging at 9500 rpm for 15 min [34]. Prior to viral DNA

extraction, samples were treated using DNAse (Sigma) (final concentration 1U/μL). Viral DNA

was extracted by using phenol:chloroform followed by precipitation with 95 % ethanol or the

Wizard DNA Clean up system following the manufacturer’s instructions. In order to confirm the

size and purity of the viral DNA, 4 μL were visualized in 0.7 % agarose gels using GelStar

Nucleic Acid Gel Stain (Lonza, Rockland, ME, USA).

Lysogen induction

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One-hundred-μL of an overnight culture of 50 randomly selected enterococci isolates harboring

one or more of the genes of interest, were added to 5 mL of Azide Dextrose (Difco) containing

mitomycin C (Sigma) (final concentration of 1 μg/mL). Samples were incubated in a water bath

at 37 °C for 4-6 h and centrifuged at 14,000 rpm for 15 min at 4 ºC [35]. Supernatants were

tested for the presence of lysogenic phages exhibiting the formation of viral plaques by using the

double layer method and E. faecalis (ATCC 19433), E. faecium, E. gallinarum, E. hirae (ATCC

8043), E. durans (ATCC 19432), E. dispar (ATCC 51266), E. casseliflavus (ATCC 25788), E.

pseudoavium (ATCC 49372) and Staphylococcus aureus (ATCC 25923) as the bacterial hosts.

Briefly, 100 μL of the possible phage lysates and 1 mL of a fresh culture of each bacterial host

were added to 4 mL containing Trypticase Soy Broth (TSB) (Difco) and agar (0.75 % w/v) and

poured onto a Petri dish containing TSB, agar (1.5 % w/v), CaCl2·2H2O (Fisher Scientific Co.

NJ, USA) and NaN3 (MCB, OH, USA) (final concentration of 2.6 mg/mL and 0.4 mg/mL,

respectively). Two temperatures were tested for formation of viral plaques (22 and 37 ºC) and

enumerated after 24 h. Phages exhibiting the formation of viral plaques were isolated,

propagated using E. faecalis (ATCC 19433) as the bacterial host and concentrated as described

[36]. A composite of the phage lysates lacking the formation of viral plaques using the

mentioned bacterial strains was concentrated as described. DNA was extracted from phages

exhibiting and lacking the formation of viral plaques as described above. One of the induced

phage isolates was characterized morphologically by using type-B 200 mesh copper grids placed

on top of individual viral plaques and stained with uranyl acetate (UA) 2%, pH 4.5.

Bacteriophages were visualized by using a Karl Zeiss Leo 922 energy filtered transmission

electron microscope operated at 200 KV.

PCR amplification conditions

All primers used in the present study are described in Table 1. Primer design in this study was

performed using Primer 3 (http://frodo.wi.mit.edu/primer3/) and verified for the formation of

secondary structures using NetPrimer (http://www.premierbiosoft.com/netprimer/). Detection of tetM

was performed by designing primers targeting the tetM sequence found in Gene Bank (accession

number AY304474.1). Primers used for the detection of vanA and vanB were those previously

described [5, 37]. For the detection of the esp variant of E. faecalis, primers were designed using

the gene sequence found in Gene Bank (accession number DQ845099.1). Detection of the esp

http://frodo.wi.mit.edu/primer3/

http://www.premierbiosoft.com/netprimer/

150

variant of E. faecium was performed using primers described elsewhere [14]. For the detection of

lysogenic phages, primers were designed using the integrase gene sequence of the Enterococcus

phage phiFL1B found in Gene Bank (accession number GQ478082.1).

Target gene Primer sequence Direction Reference

tetM GGAAAATACGAAGGTGAACA Forward This study

GAATCCCCATTTTCCTAAGT Reverse This study

vanA GGGAAAACGACAATTGC Forward Roberts et. al, 2009

GTACAATGCGGCCGTTA Reverse Roberts et. al, 2009

vanB TTGCATGGACAAATCACTGC Forward Roberts et. al, 2009

GCTCGTTTTCCTGATGGATG Reverse Roberts et. al, 2009

esp E. faecalis CACAAATGGGTGAAGGAAGA Forward This study

AGACGAATTTCCCAGTTTGC Reverse This study

esp E. faecium TATGAAAGCAACAGCACAAGTT Forward Scott et. al, 2005

ACGTCGAAAGTTCGATTTCC Reverse Scott et. al, 2005

int TATTAGGAAAACCTCCGTCA Forward This study

ATATCTTGGGCGTAAGTGAA Reverse This study

16S rRNA AGAGTTTGATCCTGGCTCAG Forward Amann et al., 1995

ACGGGCGGTGTGTRC Reverse Amann et al., 1995

Table 1: Primers in this study. Target genes include those encoding for resistance to tetracycline

(tetM) and vancomycin (vanA and vanB), the enterococcal surface protein (esp) variants found in

E. faecalis and E. faecium and an integrase-encoding gene specific for E. faecalis phages (int).

The 16S rRNA gene amplification was used as a control in the viral fractions and lysogenic

Enterococcus phages.

Amplifications were performed in a total volume of 8μL per reaction, including 1 μL of genomic

DNA at a concentration of approximately 10 ng/μL, 4 μL of Fermentas DreamTaq PCR Master

Mix, 2.2 μL of milliQ water and 0.4μL of each primer at 10 μM. PCR conditions for tetM were:

an initial denaturation of 94 ºC for 5 min, followed by 25 cycles of 94 ºC for 30 s, annealing of

55 ºC for 30 s, followed by an extension of 72 ºC for 30 s and a final extension of 72 ºC for 7

min. Both vanA and vanB PCR conditions were: an initial denaturation of 96 ºC for 3 min,

followed by 35 cycles of 96 ºC for 30 s, annealing of 60 ºC for 1 min, extension of 72 ºC for 2

min and a final extension of 72 ºC for 10 min. For the detection of the esp variant found in E.

faecalis the following conditions were used: an initial denaturation of 95 ºC for 2 min, followed

151

by 30 cycles of 95 ºC for 45 s, annealing of 57 ºC for 45 s and an extension of 72 ºC for 1 min.

Detection of the esp variant found in E. faecium was done by using an initial denaturation of 95

ºC for 3 min, followed by 35 cycles of 94 ºC for 1 min, annealing of 58 ºC for 1 min and an

extension of 72 ºC for 1 min. int PCR parameters were: an initial denaturation of 95 ºC for 2 min,

followed by 35 cycles of 94 ºC for 45 s, annealing of 55 ºC for 45 s, followed by an extension of

72 ºC for 1 min and a final extension of 72 ºC for 7 min. To ensure that no bacterial DNA was

present in the viral DNA, amplification of the 16S rRNA gene was performed using the

following PCR conditions: an initial denaturation of 95 ºC for 3 min, followed by 30 cycles of 95

ºC for 30 s, annealing of 52 ºC for 30 s, followed by an extension of 72 ºC for 30 s and a final

extension of 72 ºC for 10 min. PCR products were visualized in 1.0 % agarose gels using GelStar

Nucleic Acid gel stain. Positive controls were included for tetM and both esp variants and

potential int products were sequenced using an ABI 3130xl Genetic Analyzer.


Fisher’s exact tests were used to determined differences in the prevalence of the mentioned genes

between sample types within the sample sites. The same analysis aimed to determine significant

differences in the prevalence of the esp variants between sample types. All analyses were

performed with the R statistical software (v.2.11.1) [38].

RESULTS

Bacterial fraction and enterococci isolates

Table 2 shows the prevalence of the tested genes in the bacterial fractions of the water and sand

samples. A significant difference was noted in the prevalence of the esp variants of water

samples collected from CBB (p = 0.030). Table 3 shows the prevalence of the esp variants and

int in the enterococci isolates. Table 4 shows the prevalence of tetracycline and vancomycin-

resistant enterococci and tetM, vanA and vanB. Not all enterococci exhibiting resistance to either

antibiotic harbored tetM or vanA and vanB. Interestingly, several of these isolates harbored two

of the genes of interest. In OP waters, 2 of the isolates were positive for both tetM and vanA, 8

harbored tetM and the esp variant present in E. faecalis and 1 harbored both vanA and the esp

variant found in E. faecalis. In wet sands collected from OP and CBB, 2 and 1 of the isolates

were positive for tetM and the esp variant present in E. faecium, respectively. In dry sands

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collected from CBB, 3 of the enterococci isolated were positive for tetM and the esp variant

present in E. faecium. Similarly, 1 of the isolates from LC was positive for the latter combination

of genes (data not shown). Significant differences were noted for the prevalence of tetM in CBB

waters and dry sand (p = 0.0015) and CBB wet and dry sands (p = 0.0038). The prevalence of

both esp variants in waters collected from OP was significantly different as well (p = 0.033).

OP

water

(n=35)

OP wet

sand

(n=16)

OP dry

sand

(n=15)

CBB

water

(n=13)

CBB wet

sand

(n=14)

CBB dry

sand

(n=11)

LC

(n=8)

RP

(n=10)

tetM 6 (17) 2 (13) 2 (13) 4 (31) 1 (7) 1 (9) 2 (25) 2 (20)

vanA 7 (20) 4 (25) 2 (13) ND* 1 (7) 2 (18) 2 (25) ND*

vanB ND* ND* ND* ND* ND* ND* ND* ND*

esp E. faecalis 2 (5) 2 (13) 2 (13) 7 (54) 7 (50) 4 (36) 2 (25) 6 (60)

esp E. faecium 6 (17) 4 (25) 5 (33) 1 (8) ND* ND* ND* ND*

int 4 (11) 1 (6) 1 (7) 2 (15) 1 (7) ND* 3 (38) 2 (20)

ND*=not detected

Table 2: Prevalence of antibiotic-resistance and virulence-encoding genes in the bacterial

fractions of tropical marine (OP, CBB) and fresh waters (LC, RP) and wet and dry sands (OP,

CBB). Genes included those conferring resistance to tetracycline (tetM) and vancomycin (vanA

and vanB), the enterococcal surface protein (esp) variants present in E. faecalis and E. faecium

and an integrase-encoding gene specific for E. faecalis phages (int). Percents are presented in

parenthesis.

OP

water

(n=99)

OP wet

sand

(n=55)

OP dry

sand

(n=46)

CBB

water

(n=25)

CBB wet

sand

(n=21)

CBB dry

sand

(n=25)

LC

(n=19)

RP

(n=66)

esp E.

faecalis

10 (10) 1 (2) 1 (2) 2 (8) 1 (5) ND* ND* 6 (9)

esp E.

faecium

2 (2) 6 (11) 3 (7) 2 (8) 2 (10) 5 (20) ND* 5 (8)

int 3 (3) ND* ND* ND* ND* ND* 3 (16) ND*

ND*=not detected

153

Table 3: Prevalence of the esp variants present in E. faecalis and E. faecium and int in tropical

marine (OP, CBB) and fresh waters (LC, RP) and wet and dry beach sands in the Enterococcus

isolates. Percents are presented in parenthesis.

ND*=not detected

Table 4: Enterococci isolates tested for tetracycline (16 μg/mL) and vancomycin (20 μg/mL)

resistance isolated from tropical samples. The presence of tetracycline (tetM) and vancomycin

(vanA and vanB) resistance genes was also tested in the isolates.

Viral fraction and induced phages

None of the antibiotic-resistance or virulence genes were detected in the viral DNA fractions.

None of the samples were positive for the 16S rRNA gene and only int was detected in one of

the filtrates. Similarly, the antibiotic-resistance and virulence genes tested were not detected in

the induced enterococci phages. Not all induced enterococci phages were able to infect most of

the Enterococcus or S. aureus type strains in this study, but those that formed viral plaques were

able to replicate at 22 and 37ºC in E. faecalis. In addition, enterococci phages induced from an

enterococci isolate exhibited two different viral plaque morphologies (Figure 2A). Some plaques

were turbid on the edges and translucent in the center, while other plaques were completely

translucent, and these phages were not positive for the integrase-encoding gene tested in this

study. On the other hand, one of the induced enterococci phages that did not form viral plaques

using the mentioned enterococci type strains was positive for int. Sequencing of int showed a 99

% similarity (e value=4 e-81) with int of E. faecalis phages phiFL1A, 1B, 1C, 2A, 2B, 3A and

Antibiotic

Gene

Sample Tested Tetracycline Vancomycin Tested tetM vanA vanB

OP water 188 26(14) 18(10) 99 13(13) 2 ND*

OP wet sand 75 14(19) 10(13) 55 9(16) ND* ND*

OP dry sand 136 31(23) 31(23) 46 9(20) ND* ND*

CBB water 65 5(8) 3(5) 25 2(8) ND* ND*

CBB wet sand 75 23(31) 8(11) 21 2(10) ND* ND*

CBB dry sand 130 74(57) 16(12) 25 13(52) ND* ND*

LC 173 50(29) 70(40) 19 3(16) ND* ND*

RP 87 22(29) 24(28) 66 3(5) ND* ND*

154

3B. In terms of the morphology of one of the induced enterococci phage, this exhibited an

icosahedral capsid and a flexible tail of approximately 80 and 200nm, respectively (Figure 2B).

(A) (B)

Figure 2: Characterization of induced enterococci phages. Panel (A) shows the viral plaque

morphologies of an enterococci isolate. Panel (B) shows the morphology of an induced

enterococci phage (Bar=200 nm). Phage exhibited an icosahedral capsid of approximately 80 nm

and a flexible tail of approximately 200 nm.

DISCUSSION

Many studies focusing on the prevalence of tet determinants in the environment have been done

in waters associated with swine production facilities and livestock [27, 39, 40]. However, few

studies that focus on the prevalence of tetracycline-resistance genes in tropical recreational

waters are available. In the present study, tetM was detected in both the bacterial fractions and

enterococci isolated from waters and wet and dry sands. Interestingly, significant differences

were noted in the prevalence of tetM in waters and wet and dry sands collected from CBB. This

may suggest that the prevalence of tetM may be independent of the sample type, but this was not

the case for all the sampled sites. The detection of tetracycline resistant bacteria in the sites

tested may suggest that these are being introduced via human fecal material (as there are no

animal husbandry facilities in proximity). Another possibility is that the presence of tetracycline

in waters is selecting for resistant bacteria. This is reasonable since it has been shown that

tetracycline can be introduced via feces and urine and detected in concentrations of

approximately 0.10 μg/L [41, 42]. The horizontal transfer of genes conferring tetracycline-

155

resistance via mobile elements between autochthonous and exogenous bacteria is likely, but in

tropical recreational waters it may be difficult to identify enterococci from a fecal source and

those from the environment. Interestingly, enterococci that exhibited resistance to tetracycline

but were negative for tetM suggests that these may harbor other tet determinants.

This is also one of the few reports on the detection of vanA and vanB in tropical recreational

waters. Results suggest that several bacterial species in tropical environments may harbor vanA

or vanA-like genes (e.g. S. aureus) [43]. The present study is comparable with previous results in

which enterococci harboring vanA have been isolated from marine waters [5, 44]. In contrast,

vanB was not detected in any of the samples tested, but this is similar to results presented

elsewhere [45]. Although the reasons for this remain to be determined, it is possible that

harboring vanA, instead of vanB, may be more beneficial since harboring vanA confers resistance

to vancomycin and teicoplanin [46]. It should be noted that most of the enterococci isolates in

the present study that exhibited resistance to vancomycin lacked vanA or vanB. It is possible that

the enterococci isolates tested harbored other genes conferring resistance to vancomycin (e.g.

vanC, vanD and vanE); although it is also feasible that genes conferring resistance to other

antibiotics may also be conferring resistance to vancomycin. Similar outcomes have been

reported in Europe in which vancomycin-resistant enterococci have been associated with the use

of avoparcin [47]. The source of vanA in tropical waters and sand remains to be determined,

although the input of fecal matter remains a possibility since vancomycin-resistant bacteria have

been detected in sewage effluents [44]. As with tetracycline, future studies need to determine if

vancomycin is being introduced into the waters tested and if it is a selective force in tropical

environments.

Detection of both esp variants in the bacterial fractions and enterococci isolates of several of the

samples may suggest an input of human and animal fecal material [13]. The absence of the E.

faecium variant in the bacterial fractions and in enterococci of several of the samples evaluated

was not expected since these sites are visited throughout the year and may not be free from fecal

contamination from non-point sources. This may suggest that the E. faecium variant may not

necessarily indicate human fecal pollution in all water types, especially in the tropics, in which

enterococci are naturally found in many water bodies [48]. Only the bacterial fraction of waters

156

collected from CBB and the enterococci isolated from OP water exhibited a difference in the

prevalence of both esp variants. This may suggest that, in tropical environments, similar results

are obtained when detecting either esp variant.

In the present study, none of the tested genes were detected in the viral DNA fraction or the

induced enterococci phages. However, the presence of other antibiotic-resistance and virulence-

encoding genes in bacteriophages should not be ignored. In terms of int, although it was detected

in the enterococci isolates and one of the induced phages, it is possible that other Enterococcus

phages may harbor int sequences different from the variant tested in the present study. Phages

infecting enterococci have exhibited differences in their lysogeny modules and int sequences [24,

49]. The present study also suggests that Enterococcus isolates may harbor more than one

inducible prophage, but the function of various prophages in the genomes of enterococci remains

to be determined [50, 51].

CONCLUSIONS

This is one of the first reports which aimed to identify a battery of antibiotic-resistance and

virulence genes in both the bacterial and viral fractions of tropical recreational waters and sand.

The presence of antibiotic-resistance and virulence genes in the environmental microbiota of

tropical samples suggests that the transmission of these genes is not restricted to strains of the

clinical setting. The exact mechanisms of horizontal transfer of antibiotic-resistance and

virulence genes under these environmental conditions need to be determined, as do the role of

environmental microbiota as reservoirs of these genes. By detecting int in lysogenic phages,

further studies could determine the presence of antibiotic-resistance and virulence factors in their

genomes. Our data also point to the need to take into consideration lysogenic phages when

determining the presence or absence of bacterial viruses in the environment.

Most epidemiological studies on recreational waters have focused on gastrointestinal illness;

though a few recent ones have also included and eye, skin and ear infections. The present study

showed a relatively high prevalence of Enterococcus spp. exhibiting resistance to antibiotics

which also harbored virulence genes. It remains to be determined if these bacteria represent a

risk to bathers and if these are specifically involved in skin infections. However, it is worrying

157

that opportunistic pathogens involved in skin infections and harboring antibiotic resistance are

present in recreational waters and sands, where bathers may be exposed to them.

ACKNOWLEDGEMENTS

We thank M.G. Dominguez-Bello, N. Shankar and M.N. Byappanahalli for sharing the positive

controls used in this study; Miguel Urdaneta for the phage microscopy and Rita Patricio, Jean M.

Carrasquillo, Roberto Lorenzini, Gabriel Vázquez and Gabriela Tirado for sample collection and

processing. This research was partially supported by MBRS-RISE (NIH Grant Number

2R25GM061151-09).

158

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CHAPTER 7

General conclusions and future directions

GENERAL CONCLUSIONS

A culture method was developed for the detection of Enterococcus faecalis phages

(enterophages) in recreational waters, sewage, sand and fecal samples. Our method is the most

recent approach to infer the virological quality of water sources. This method can also be tested

for the detection of enterophages in other sample types such as soils and food. It should be noted

that if the enterophage method is to be tested in other laboratories, a consistency should be kept

(reagents and the bacterial host) in order to compare results. The method enabled us to isolate

these viruses to further characterize them ecologically, morphologically and genetically.

Enterophages possess many of the characteristics of an indicator of fecal pollution and

specifically that from a human source. Enterophages seem to be restricted to human fecal

material, as these were not detected in pig, chicken, cow or dog feces. The present study

determined the inactivation rates and survival times of enterophages under various conditions.

Results were compared to those of human enteric viruses reported elsewhere under similar

conditions. Moreover, the survival of enterophages in beach sand was determined, and their short

survival rates, compared to coliphages, suggest recent fecal pollution. Enterophages could be

considered surrogates of certain human enteric viruses since their inactivation rates and survival

times are similar to those reported. Enterophages exhibit resistances to removal and inactivation

methods, similarly to human enteric pathogens.

Enterophages were tested as indicators of fecal pollution in tropical inland waters as few studies

have evaluated the microbial quality of these waters. One problem is the need to develop

appropriate water quality standards for tropical waters since those used in other geographical

regions may not be applicable. We tested various Microbial Source Tracking (MST) methods

(enerococci, thermotolerant coliforms, enterophages, coliphages, human- and cattle-specific

Bacteroides, chicken-specific Bacteroides and Clostridium, and enterococci by qPCR) to assess

the microbial quality of a tropical watershed in Puerto Rico. Enumeration of both coliphages and

enterophages may be a more appropriate to infer fecal contamination, compared to the

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bacteriological method. The reason is that these bacteriophages are not ubiquitous in tropical

waters. The human Bacteroides seemed a reliable means of inferring human fecal contamination

in tropical waters, unlike the cattle and chicken-specific Bacteroides and Clostridium markers.

Interestingly, positive correlations between the culture and molecular methods for the detection

of enterococci should be considered prior to implementing qPCR as a rapid method for inferring

fecal pollution in the tropics. The qPCR method for enterococci may not apply to all water

bodies, specifically to tropical inland lakes. qPCR assays targeting enterococci in tropical waters

may overestimate the numbers of enterococci as many molecular methods may also amplify

naked DNA.

The latter study is one of the few (if not the only) that has considered the effect of rainfall events

in the detection of a battery of microbial indicators, coliphages and enterophages in tropical

waters. Rainfall is often not considered when determining the prevalence of microbial indicators

in surface waters. Rainfall events may result in the resuspension of sediments into surface

waters. Sediments often harbor enteric microbes, specifically those that can replicate and/or

persist for long periods in the environment. This represents a concern to public health since many

waterborne outbreaks occur after precipitation events, but can also represent a disadvantage for

traditional indicators. The reason is that if the numbers of a microbial indicator in surface waters

increase after rainfall events may suggest that their input may not be recent. This is turn may

suggest that these microbes have replicated under environmental conditions.

The presence of enterophages in chicken, cattle, dog and pig feces, as well as their prevalence in

domestic sewage were determined using E. faecalis, E. faecium, E. gallinarum, E. hirae, E.

durans, E. dispar, E. casseliflavus and E. pseudoavium as the bacterial hosts. Results showed

that enterophages infect E. faecalis exclusively, and that both humans and animals harbor

Enterococcus phages. Interestingly, phages infecting E. faecium, E. casseliflavus and E.

pseudoavium replicating at 37 °C may be used to infer the presence of chicken-fecal matter in

fresh water sources.

The present project determined the prevalence of tetracycline (tetM) and vancomycin-resistance

(vanA and vanB) and the enterococcal surface protein genes (esp) of E. faecalis and E. faecium in

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both the bacterial and viral fractions of tropical environmental samples. Detection of the

mentioned genes in the environmental microbiota of tropical samples suggests that the

transmission of these genes is not restricted to strains of the clinical setting. The present study

showed a relatively high prevalence of Enterococcus spp. exhibiting resistance to antibiotics,

which also harbor the respective antibiotic-resistance genes and esp. Although phages did not

harbor the genes tested in the present study, the presence of other genes should not be ignored, as

well as their role as possible vectors. Detection of an integrase-encoding gene in lysogenic

phages will enable the detection of specific antibiotic-resistance and virulence-encoding genes in

their genomes. Importantly, results suggest that lysogenic phages should be considered when

determining the prevalence of bacteriophages in the environment.

FUTURE DIRECTIONS

Future studies need to simultaneously determine the prevalence of enterophages and enteric

viruses in various water types. This will enable direct correlations between indicators and enteric

viruses, and thus could further support the potential of enterophages as surrogates of these

pathogens. In addition, there is still the need to correlate the presence of enterophages with

gastrointestinal illnesses, eye, ear and skin infections.

The detection of enterophages by molecular methods needs to be developed and optimized. For

this reason, the complete genomes of lytic and lysogenic phages infecting E. faecalis are

currently being sequenced in collaboration with the Human Microbiome Project. This will not

only enable to develop the molecular techniques for the detection of these phages in the

environment, but will also represent a means to understand the genomes of lytic and lysogenic

phages infecting E. faecalis. The role of bacteriophages in biogeochemical processes, bacterial

pathogenicity and evolution has been a subject of increasing interest [1]. Many studies have

focused on the effects of strictly lytic phages on bacterial populations and as a possible tool to

treat bacterial infections [2, 3]. Lytic phages may be less genetically diverse (compared to

lysogenic phages) since DNA exchange seems to be more restricted due to their strictly virulent

nature [4]. Lysogenic phages, on the other hand, are the main focus of many genomic studies

since these may help shape bacterial genomes [5, 6]. Lysogenic phages are important vehicles for

DNA segments, which can make the bacterial host more competitive in a current ecosystem or

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allow it to adapt to a new one [2, 7, 8]. Sequencing lytic and lysogenic E. faecalis phages from

humans may open up the opportunity to further study their possible roles and how these may

affect Enterococcus populations in the intestine. Although many recent studies have focused on

lysogenic phages, understanding the genomic structures of both strictly virulent and lysogenic

phages is important since both phage types are involved in shaping bacterial populations.

167

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