REVIEW OF COLIPHAGES AS POSSIBLE 

INDICATORS OF FECAL CONTAMINATION 

FOR AMBIENT WATER QUALITY 

820-R-15-098

EPA Office of Water 

Office of Science and Technology 

Health and Ecological Criteria Division 

April 17, 2015 
i 

NOTICES 

This document has been drafted and approved for publication by the Health and Ecological 

Criteria Division, Office of Science and Technology, United States (U.S.) Environmental 

Protection Agency (EPA), and is approved for publication. Mention of trade names or 

commercial products does not constitute endorsement or recommendation for use.
ii

ACKNOWLEDGMENTS 

The development of this criteria document was made possible through an effort led by Sharon 

Nappier, EPA Project Manager, EPA, Office of Science and Technology, Office of Water. EPA 

acknowledges the valuable contributions of EPA Internal Technical Reviewers who reviewed 

this document: Jamie Strong and Elizabeth Doyle. 

The project described here was managed by the Office of Science and Technology, Office of 

Water, EPA under EPA Contract EP-C-11-005 to ICF International. EPA also wishes to thank 

Audrey Ichida, Kirsten Jaglo, Jeffrey Soller, Arun Varghese, Alexandria Boehm, Kara Nelson, 

Margaret Nellor, and Kaedra Jones for their contributions and invaluable support. 

The primary contact regarding questions or comments to this document is: 

Sharon Nappier 

U.S. EPA Headquarters 

Office of Science and Technology, Office of Water 

1200 Pennsylvania Ave., NW 

Washington, DC 20460 

Mail Code 4304T 

Phone: (202) 566-0740 

Email: nappier.sharon@epa.gov 
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EXTERNAL PEER REVIEW WORKGROUP

The External Peer Review was managed by the Office of Science and Technology, Office of 

Water, EPA under EPA Contract No. EP-C-13-010 to Versar, Inc. The following professionals 

were part of the External Peer Review Workgroup that provided excellent technical and 

scientific review on the Draft regarding the content and technical approach in response to EPA 

Charge to the Peer Reviewers: 

Valerie J. Harwood 

University of South Florida 

Tampa, Florida 33620 

Sunny Jiang 

University of California, Irvine 

Irvine, California 92697 

Mark D. Sobsey 

University of North Carolina, Chapel Hill 

Chapel Hill, NC 27599 

EPA reviewed and incorporated their comments, where appropriate, to develop this literature 

review. 

Potential areas for conflict of interest were investigated with the Peer Reviewers, including a 

review of their current affiliations. No conflicts of interest were identified. 
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TABLE OF CONTENTS 

Notices ............................................................................................................................................. i

Acknowledgments........................................................................................................................... ii

External Peer Review Workgroup .................................................................................................iii

Table of Contents ........................................................................................................................... iv

Tables ............................................................................................................................................. vi

Figures........................................................................................................................................... vii

1. Introduction ............................................................................................................................. 1

1.1. Context and Purpose......................................................................................................... 1

1.2. Background ...................................................................................................................... 1

1.3. General Attributes of an Ideal Indicator of Enteric Viral Degradation ............................ 3

2. Bacteriophage Characteristics ................................................................................................. 5

2.1. Origin and Replication ..................................................................................................... 5

2.2. Morphology ...................................................................................................................... 7

2.2.1 Morphological Properties Affecting Persistence ........................................................... 9

2.3. Detection Methods ......................................................................................................... 10

2.3.1 Culture-Based Methods ............................................................................................... 10

2.3.2 Rapid Methods ............................................................................................................. 12

3. Epidemiological Relationships ............................................................................................. 16

3.1 Von Schirnding et al. (1992) ............................................................................................... 17

3.2 Lee et al. (1997) .................................................................................................................. 18

3.3 Medema et al. (1995) and Van Asperen et al. (1998) ......................................................... 19

3.4 Wiedenmann et al. (2006) ................................................................................................... 20

3.5 Colford et al. (2005, 2007) .................................................................................................. 21

3.6 Wade et al. (2010) ............................................................................................................... 22

3.7 Abdelzaher et al. (2011) ...................................................................................................... 23

3.8 Griffith et al. (personal communication, 2015) .................................................................. 24

3.9 General Conclusions from Epidemiological Studies .......................................................... 24

4. Occurrence in the Environment ............................................................................................ 27

4.1. Associations between Coliphages and Viruses .............................................................. 27

4.1.1 Coliphage – Virus Associations in Freshwater ............................................................ 31

4.1.2 Coliphage-Virus Associations in Saline or Brackish Water ........................................ 31

5. Environmental Factors and Fate ........................................................................................... 37

5.1. Temperature ................................................................................................................... 37

5.2. Sunlight .......................................................................................................................... 39
v

5.3. Salinity ........................................................................................................................... 44

5.4. Predation and Enzymatic Degradation ........................................................................... 47

5.5. Organic and Inorganic Matter ........................................................................................ 49

5.6. Environmental Factors Impacts Summary ..................................................................... 51

6. Wastewater Treatment .......................................................................................................... 56

6.1. Primary Treatment.......................................................................................................... 60

6.2. Secondary Treatment...................................................................................................... 61

6.3. Wastewater Treatment Ponds ......................................................................................... 63

6.4. Tertiary Treatment and Advanced Treatment ................................................................ 63

6.5. Disinfection .................................................................................................................... 66

6.5.1 Free Chlorine ............................................................................................................... 66

6.5.2 Combined Chlorine ...................................................................................................... 68

6.5.3 Ozone ........................................................................................................................... 71

6.5.4 UVC ............................................................................................................................. 72

6.5.5 UVA and UVB ............................................................................................................. 75

7. Conclusions ........................................................................................................................... 78

8. References ............................................................................................................................. 81

APPENDIX A: Literature Search Strategy and Summary of Literature Search Results ............ 110

APPENDIX B: Coliphage and NoV Densities during Wastewater Treatment .......................... 114
vi 

Tables 

Table 1. Morphology of a subsection of bacteriophages. ............................................................... 8

Table 2. Morphology of human enteric viruses that may be transmitted in aquatic 

environments. ................................................................................................................... 9

Table 3. Advantages and disadvantages of methods to detect coliphages. ................................... 14

Table 4. Summary of epidemiological studies. ............................................................................. 26

Table 5. Number of cases and outcome of the logistical regression analysis of the 

association between coliphages and pathogens in water. ............................................... 28

Table 6. Logistic regression of the association between indicators and different pathogens 

in water. .......................................................................................................................... 29

Table 7. Comparison of common methods for the detection of pathogenic human enteric 

viruses from environmental sources. .............................................................................. 30

Table 8. Summary table of coliphages – virus correlations in ambient water. ............................. 33

Table 9. Comparison of mean exponential decay rates of coliphages, fecal indicators and 

human viruses in different media at different temperatures. .......................................... 40

Table 10. Mean exponential decay rates of coliphages and fecal indicators in fresh river 

water contaminated with raw sewage or effluent under different light conditions. ....... 42

Table 11. Comparison of mean exponential decay rates of coliphages and human viruses 

under different light conditions. ..................................................................................... 43

Table 12. Comparison of mean exponential decay rates of coliphages and MNV at 

different concentrations of salt and at different temperatures. ....................................... 47

Table 13. Summary of environmental factors influencing viral inactivation in aquatic 

environments. ................................................................................................................. 52

Table 14. Log10 removals of enteric viruses and indicator organisms. ......................................... 60

Table 15. Virus densities in secondary treated wastewater samples from five Australian 

WWTPs. ......................................................................................................................... 62

Table 16. Average (and percent positive) microorganism effluent densities in a WWTP 

with free chlorine treatment of nitrified and filtered secondary wastewater. ................. 67

Table 17. Log10 reduction of FIB, enteric virus, and F-specific coliphages in sewage 

matrix due to chlorine (adapted from Tree et al., 2003). ................................................ 69

Table 18. Log10 reduction of bacteriophages and enteroviruses in spiked secondary 

effluent after chlorination with 20 mg/L of chlorine.* ................................................... 70

Table 19. Inactivation of FIB and F-specific RNA coliphage in ozone disinfected water.* ........ 72

Table 20. Average (and percent positive) microorganism densities in a WWTP with UV 

treatment of filtered secondary effluent (n=5). ............................................................... 73

Table 21. UVC inactivation of F-specific RNA coliphage MS2. ................................................. 74

Table 22. Log10 reduction in coliphages and enteric viruses in secondary effluent after 

lagooning in sunlight or UVC treatment. ....................................................................... 76

Table 23. Attributes of fecal contamination indicators. ................................................................ 80

Table A. Coliphage and NoV densities during wastewater treatment. ....................................... 114
vii 

Figures

Figure 1. Comparison of coliphage and enteric viruses in raw sewage and secondary 

effluent. ........................................................................................................................... 59
viii 

Acronyms and Abbreviations 

AOP advanced oxidation processes 

AOR adjusted odds ratio 

BOD biochemical oxygen demand 

C Celsius 

CDOM colored dissolved organic matter 

CFU colony forming unit 

CI confidence interval 

CLAT culture latex agglutination and typing 

CT disinfectant concentration multiplied by contact time 

cm centimeter 

DNA deoxyribonucleic acid 

ds double-stranded 

EPA Environmental Protection Agency 

E. coli Escherichia coli 

Famp E. coli resistant to streptomycin and ampicillin (host) 

FCV feline calicivirus 

FIB fecal indicator bacteria 

F-specific male-specific or F+ coliphage (“F” refers to the genetic fertility factor that is 

required for bacteria to produce a sex pilus necessary for conjugation) 

GE gastroenteritis 

GI genogroup I 

GII genogroup II 

GIII genogroup III 

GIV genogroup IV 

HCGI highly credible gastrointestinal illness 

ICC integrated cell culture 

ISO International Standards Organization 

kDa kilodalton 

L liter 

mg milligram 

MJ megajoule 

mJ millijoule 

mL milliliter 

mm millimeter 

mM millimolar 

MNV murine norovirus 

MPN most probable number 

MST microbial source tracking 

μm micrometer 

NEEAR National Epidemiological and Environmental Assessment of Recreational Water 

nm nanometer 

NOAEL no observed adverse effect level 

NoV norovirus

NTU nephelometric turbidity unit 
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OR odds ratio 

PCR polymerase chain reaction 

PFU plaque forming units 

QA/QC quality assurance/quality control 

QMRA quantitative microbial risk assessments 

qPCR quantitative polymerase chain reaction 

RNA ribonucleic acid 

RR relative risk 

RT-PCR reverse transcriptase polymerase chain reaction 

RT-qPCR reverse transcriptase quantitative polymerase chain reaction 

SAL single agar layer 

SD standard deviation 

ss single-stranded 

U.S. United States 

UV ultraviolet 

W Watt 

WERF Water Environment Research Foundation 

WRF Water Research Foundation 

WWTP wastewater treatment plant 
1

1. Introduction

1.1. Context and Purpose 

The United States (U.S.) Environmental Protection Agency (EPA) recommended the use of the 

fecal indicator bacteria (FIB) Escherichia coli (E. coli) and enterococci to determine the level of 

fecal contamination present in environmental waters and to establish the 2012 Recreational 

Water Quality Criteria (RWQC), which protect the designated use of primary contact recreation 

(U.S. EPA, 2012). The purpose of this review is to summarize the scientific literature on 

coliphage properties to assess their suitability as indicators of fecal contamination in ambient 

water. This review covers background information on coliphage characteristics and enumeration 

methods (Section 2); their relationship with human health risks in epidemiological studies 

(Section 3); their occurrence and associations with pathogens in the environment (Section 4); and 

the fate and transport of coliphages in the environment (Section 5) and during wastewater 

treatment (Section 6). Appendix A describes the literature search strategy and summarizes the 

results of literature search. 

At this time, EPA is considering the use of F-specific and somatic coliphages, as possible viral 

indicators of fecal contamination in ambient water. Coliphages are a subset of bacteriophages 

that infect E. coli. Other types of bacteriophages [i.e., those that infect Enterococcus and various 

Bacteroides species (spp.)] have also been evaluated for their use as indicators of fecal 

contamination. While some information on other types of bacteriophages is presented, this 

review primarily focuses on coliphages because there is more literature available on their 

occurrence, fate, and epidemiological relationships. Additionally, two standardized enumeration 

methods published by EPA are available for both coliphages. 

1.2. Background 

For over a century, FIB (i.e., total coliforms, fecal coliforms, E. coli, fecal streptococci, and 

enterococci) have been used to detect sewage contamination in water in order to protect the 

public from waterborne pathogens associated with fecal material (e.g., bacteria, protozoa, and 

viruses) (Kehr et al., 1941; NRC, 2004). The use of FIB as indicators of sewage contamination 

facilitated tremendous gains in public health protection, particularly by indicating the likely 

presence of bacterial pathogens such as Vibrio cholerae (which causes cholera) and Salmonella 

typhi (which causes typhoid fever). Although advances in wastewater treatment over the last half 

century have facilitated gains in public health, it has been suggested that viral pathogens are the 

leading causative agents of recreational waterborne illnesses (Jiang et al., 2007; Sinclair et al., 

2009). Unfortunately, because bacteria respond to water treatment processes and environmental 

degradation processes differently than viruses, traditional FIB may not be the best indicators of 

viral pathogens associated with fecal contamination. This review considers coliphages as 

possible indicators of fecal contamination in ambient water. Because FIB have long been used 

for managing water quality, much of this review compares coliphages to other commonly used 

FIB, such as E. coli and enterococci.

EPA conducted a series of prospective cohort epidemiological studies at multiple locations from 

1972 to 1979 to better understand the relationship between FIB and swimming-associated 

illnesses (Cabelli et al., 1982; Dufour, 1984). Symptoms of the swimming-associated illnesses 
2

included acute, self-limiting gastroenteritis (GE) with a short incubation period and duration. 

From their studies, Cabelli et al. (1982) concluded that human noroviruses (NoV) or rotaviruses 

were the most likely cause of the symptoms.1 Soller et al. (2010) reached similar conclusions 

regarding the causative agent of the illnesses observed in the EPA National Epidemiological and 

Environmental Assessment of Recreational Water (NEEAR) study. Additionally, numerous 

studies have identified the presence of viruses in wastewater treatment effluent, often when 

traditional FIB are non-detectable (Kageyama et al., 2003; da Silva et al., 2007; Haramoto et al., 

2007; Kitajima et al., 2009; Kuo et al., 2010; Simmons et al., 2011). 

The human viruses most frequently associated with recreational waterborne illnesses are NoV, 

adenoviruses, human enteroviruses, rotaviruses, astroviruses, and hepatitis E, with NoV 

responsible for the large majority of viral-based gastrointestinal illnesses (U.S. EPA, 2009a). For 

example, Sinclair et al. (2009) found that 18 of the 27 (67%) reported viral outbreaks in ambient 

recreational water (does not include pools) from 1951 to 2006 were due to NoV, and 2 (7%) 

were due to adenovirus. As viruses are an important cause of recreational waterborne illness, it 

has been suggested they may also be appropriate indicators of fecal contamination in water. 

Currently, there are limitations associated with using individual pathogenic viruses as indicators. 

For one, the measurement of densities of individual pathogenic viruses in water is expensive and

time consuming, as culture-based techniques to propagate them can take over a week. Secondly, 

human NoV has only recently been cultured and methods for culture-based quantification of 

environmental water samples have not been developed yet (Papafragkou et al., 2013; Jones et al., 

2014; Thorne and Goodfellow, 2014). NoV viral ribonucleic acid (RNA) can be detected and 

amplified through reverse transcriptase polymerase chain reaction (RT-PCR) (a semiquantitative method) and RT-quantitative PCR (RT-qPCR, a quantitative method) (Kageyama et 

al., 2003; Trujillo et al., 2006; Atmar et al., 2008; Tajiri-Utagawa et al., 2009; Cashdollar et al., 

2013). However, polymerase chain reaction (PCR) methods do not differentiate between 

infective and non-infective viruses. Human enteric adenoviruses are also difficult and slow to 

culture, and therefore are frequently detected using integrated cell culture (ICC) RT-PCR assays, 

which are semi-quantitative. Methods for these assays are improving, but they are still 

technically difficult and relatively slow to produce results (i.e., days) (Rodríguez et al., 2013; 

Polston et al., 2014). Therefore, numerous authors have proposed using bacteriophages (viruses 

that infect bacteria) as an indicator of human enteric viruses in water impacted by fecal 

contamination (Hilton and Stotzky, 1973; Gerba, 1987; Havelaar, 1987; Sobsey et al., 1995; 

Chung et al., 1998; Contreras-Coll et al., 2002; Hot et al., 2003; Skraber et al., 2004a, b; MocéLlivina et al., 2005). In particular, coliphages, or viruses that infect E. coli, have been the most 

thoroughly investigated for this purpose. 

Coliphages, particularly F-specific (also known as “male-specific” or “F+ phage”) and somatic 

coliphages, have been proposed as more reliable indicators of human viral pathogens associated 

with fecal contamination than FIB (Gerba, 1987; Palmateer et al., 1991; Havelaar et al., 1993; 

1

 Cabelli et al. (1982) suggested that “human rotavirus and/or the parvo-like viruses” were the etiological agents. In 

the 1970s the virus now called NoV was described morphologically as "picorna or parvovirus-like” (Kapikian et al., 

1972). 
3

Duran et al., 2002; Rose et al., 2004; Skraber et al., 2004a, b).2 This is based on their greater 

similarity to human enteric viruses in their physical structure, composition, and morphology, 

survivability in the environment, and persistence in treatment processes compared to FIB 

(Funderburg and Sorber, 1985; Havelaar et al., 1993; Gantzer et al., 1998; Grabow, 2001; 

Nappier et al., 2006). For example, F-specific RNA coliphages are morphologically similar to 

enteroviruses, caliciviruses, astroviruses, and hepatitis A and E viruses, and some somatic 

coliphages are similar to adenovirus (King et al., 2011). Coliphages can also be detected and 

quantified by simple, inexpensive, rapid, and reliable methods (Gerba, 1987; Havelaar, 1987). 

Although they are abundant in domestic wastewater, raw sewage sludge, and polluted waters 

(Havelaar et al., 1990; Debartolomeis and Cabelli, 1991; Leclerc et al., 2000; Mandilara et al., 

2006), coliphages are present at lower densities in fresh feces than in wastewater (Dhillon et al., 

1976; Osawa et al., 1981; Calci et al., 1998; Gantzer et al., 2002; Long et al., 2005). They 

originate almost exclusively from the feces of humans and other warm-blooded animals and can 

undergo limited multiplication in sewage under some conditions (i.e., high densities of 

coliphages and susceptible host E. coli at permissive temperatures) (Sobsey et al., 1995; Grabow, 

2001). Coliphages (detected by EPA Method 1601, 1602, or approved equivalent methods) are 

one of the fecal indicator organisms that can be selected for microbial monitoring of groundwater 

systems (U.S. EPA, 2006). 

1.3. General Attributes of an Ideal Indicator of Enteric Viral Degradation 

Methodological constraints limit reliable enumeration of individual pathogens in water (see 

Section 2). Because pathogen enumeration methods are not advanced enough at this time for use 

in routine water quality monitoring, FIB have been used to detect the presence of fecal 

contamination. Important attributes of an ideal indicator for fecal contamination include the 

following (NRC, 2004; Bitton, 2005): 

 the indicator should be a member of the intestinal microflora of warm-blooded animals 

(see Section 2); 

 the indicator should be present when pathogens are present and absent in uncontaminated 

samples (see Section 4); 

 the indicator should be present in greater numbers than the pathogen (see Sections 4 

and 6); 

 the indicator should be at least as resistant as the pathogen to environmental factors (see 

Section 5) and to disinfection in water and wastewater treatment plants (WWTPs) (see 

Section 6); 

 the indicator should not multiply in the environment (see Section 2); 

 the indicator should be detectable by easy, rapid, and inexpensive methods (see 

Section 2); 

 the indicator should be nonpathogenic (see Section 2); 

 the indicator should be correlated to health risk (see Section 3); and 

 the indicator should be specific to a fecal source or identifiable as to source of origin 

(microbial source tracking [MST] is not included in this review). 

 

2

 Coliphages are broken into two groups, F-specific (also referred to as male-specific or F+) RNA or DNA 

coliphages and somatic coliphages. Both infect E. coli; somatic coliphages infect E. coli cells through their outer 

membrane and F-specific coliphages infect E. coli via the pilus appendage, found on the surface of some types of 

bacteria for conjugative or motile functions. 
4

While there is no true “ideal” indicator that fits all of the criteria above, coliphages exhibit most 

of these attributes, including the following: 

 they are part of the intestinal microflora of warm-blooded animals (Sobsey et al., 1995; 

Grabow, 2001); 

 they are present in greater numbers than pathogens (Havelaar et al., 1990; Debartolomeis 

and Cabelli, 1991; Leclerc et al., 2000); 

 they are detectable by easy and rapid (1 day or less) methods (Wentsel et al., 1982; 

Gerba, 1987; Havelaar, 1987); and 

 they are nonpathogenic (Grabow, 2001; Pillai, 2006; Jończyk et al., 2011). 

Coliphages partially meet some of the other criteria, including the following: 

 they co-occur with pathogens in water in some studies (for example, Havelaar et al., 

1993; Jiang et al., 2001; Ballester et al., 2005; Pillai, 2006; Wu et al., 2011); 

 they are at least equally resistant as some viral pathogens to environmental factors and to 

disinfection in water and WWTPs (Havelaar, 1987, 1990; Yahya and Yanko, 1992; 

Nasser et al., 1993; Gantzer et al., 1998; Sinton et al., 2002; Hot et al., 2003; Bitton, 

2005; Lodder and de Roda Husman, 2005; Pillai et al., 2006; Charles et al., 2009; 

Bertrand et al., 2012; Seo et al., 2012; Silverman et al., 2013); 

 they undergo very limited to no multiplication in the environment (Grabow et al., 1980; 

Grabow, 2001; Luther and Fujioka, 2004; Muniesa and Jofre, 2004; Jofre, 2009; Jończyk 

et al., 2011); and 

 they have been shown to correlate with health risk in some studies (Lee et al., 1997; 

Colford et al., 2005, 2007; Wiedenmann et al., 2006; Abdelzaher et al., 2011). 

Additionally, while not the focus of this review, assays for bacteriophages have been developed 

to identify some sources of origin (Pina et al., 1998; Brion et al., 2002; Schaper et al., 2002a; 

Cole et al., 2003; Noble et al., 2003; Payan et al., 2005; Savichtcheva and Okabe, 2006; StewartPullaro et al., 2006; Ebdon et al., 2007, 2012; Lee et al., 2009; Wolf et al., 2010; Gómez-Doñate 

et al., 2011; Jofre et al., 2011; Lee et al., 2011; Nnane et al., 2011; Boehm et al., 2013).

This review evaluates the potential for coliphages to be useful as viral indicators of fecal 

contamination. The above attributes are considered in more detail throughout the review. 
5

2. Bacteriophage Characteristics

Bacteriophages (commonly referred to as phages) are viruses that infect bacteria. Phages were 

first described as a component of the human microbiome in the early 1900s and are 

nonpathogenic. They exist for all known bacterial species, and a wide variety have been isolated. 

Based on their size and morphology, bacteriophages are classified into 13 different phylogenetic 

families (Pillai, 2006). Generically, the bacteriophage virion (entire virus particle) consists of 

either double-stranded (ds) or single-stranded (ss) RNA or deoxyribonucleic acid (DNA), a 

protein capsid, and in some cases, a lipid membrane envelope (Pillai, 2006). Bacteriophages 

under evaluation as indicators of fecal contamination are nonenveloped, like many viral 

pathogens of interest. See Tables 1 and 2 below for more details. 

In recent years, bacteriophages that infect E. coli, Enterococcus, and various Bacteroides spp. 

have been considered as possible indicators of fecal contamination (Chung and Sobsey, 1993; 

Grabow et al., 1995; Jofre et al., 1995; Bradley et al., 1998; Gantzer et al., 1998; ISO, 1999; 

Duran et al., 2002; Lucena et al., 2003; Mandilara et al., 2006; Bonilla et al., 2010; SantiagoRodriguez et al., 2010; Vijayavel et al., 2010; Purnell et al., 2011). The majority of research on 

using bacteriophages as fecal indicators has been conducted on coliphages, which are 

bacteriophages that infect E. coli (U.S. EPA, 2001a). Coliphages, specifically F-specific and 

somatic coliphages, are the primary focus of this document and are described in detail below. 

2.1. Origin and Replication 

Bacteriophages are considered the most abundant form of “life” on earth and can be found in all 

environments where bacteria grow, including in soil, water, and inside other larger organisms 

(e.g., humans) harboring host bacteria (e.g., E. coli) (Clokie et al., 2011; Dutilh et al., 2014; 

Díaz-Muñoz and Koskella, 2014). However, these viruses only reproduce inside metabolizing 

bacterial hosts and are thus considered obligate intracellular parasites that cannot multiply 

independently in any environment outside of the host bacterial cell (Grabow, 2001; Brüssow et 

al., 2004; Jończyk et al., 2011). For replication to occur in a given environment, such as in 

recreational waters, their host must be both viable in that environment (Grabow, 2001; Bitton, 

2005; Jofre, 2009) and susceptible to bacteriophage infection (Wiggins and Alexander, 1985; 

Woody and Cliver, 1995, 1997). Bacteriophages use the host cell’s ribosomes, proteinsynthesizing machinery, amino acids, and energy generating systems to replicate. Some 

bacteriophage species possess fewer than 10 genes and use essentially all of the host’s cellular 

functions to replicate. In contrast, other bacteriophages have 30 to 100 genes and are less 

dependent on the host. For example, larger bacteriophage may not require host genes for DNA 

replication because their own genomes contain the necessary genes (Grabow, 2001). 

Bacteriophage replication includes the following steps: 

1) adsorption: the virion attaches itself to a host cell;

2) penetration: the genome enters the host cell;

3) viral synthesis: the host cell manufactures viral components;

4) maturation: the components are assembled into intact new virions; and

5) release: virus particles leave the infected cell (Goldman and Green, 2009).
6

The host-specificity of bacteriophages is determined by protein molecules that serve as receptor 

sites on the surface of potential host bacteria. Only specific bacteriophages will recognize and 

attach to these specific bacterial receptor sites. Attachment leads to infection of the bacterium 

host as described above (Grabow, 2001). 

Coliphages are generally found in the gut and are excreted in the feces of humans and other 

warm-blooded animals. Coliphages are present in large numbers in sewage (approximately 107

plaque forming units [PFU] per milliliter [mL]) (Ewert and Paynter, 1980; Lucena et al., 2004; 

Lodder and de Roda Husman, 2005). They have been investigated for years as possible viral 

indicators of fecal contamination (Simkova and Cervenka, 1981; Gerba, 1987; Havelaar et al., 

1993; Sobsey et al., 1995; Chung et al., 1998; Contreras-Coll et al., 2002; Cole et al., 2003; Hot 

et al., 2003; Lucena et al., 2003; Mocé-Llivina et al., 2005; Brezina and Baldini, 2008; Wu et al.,

2011). Coliphages can be divided into seven major morphological groups, or families; four of 

which contain somatic coliphages and three of which contain F-specific coliphages (Cole et al., 

2003; Mesquita et al., 2010). Somatic coliphages infect E. coli cells through their outer 

membrane; F-specific coliphages infect E. coli via the pilus appendage, found on the surface of 

many types of bacteria. Studies indicate that somatic coliphages are excreted at higher levels 

than F-specific RNA coliphages and that somatic coliphages are likely to be more persistent in 

water than F-specific RNA coliphages (Grabow, 2001; Schaper et al., 2002a; Lee and Sobsey, 

2011). Both F-specific and somatic coliphages, including their taxonomy, are described below.

Somatic coliphages are an abundant group of bacteriophages in feces and encompass DNA 

bacteriophages that infect coliform bacteria, including E. coli, via the outer membrane. The 

bacteriophage families Myoviridae, Siphoviridae, Podoviridae, and Microviridae have somatic 

coliphage representatives (Hayes, 1968; Grabow, 2001). E. coli strains that are used for 

propagating somatic coliphages include E. coli CN13 and E. coli WG5 (Muniesa et al., 2003). 

Coliphage strain ΦX174 from the Microviridae family is a model somatic coliphage that is 

widely used in laboratory settings. Coliphages in the Microviridae family have circular ds DNA. 

Coliphages in the Myoviridae, Siphoviridae, and Podoviridae families have linear ds DNA. For 

more information on coliphage families, see Section 2.2 (Table 1) below. 

Male-specific, or F-specific, coliphages are another broad group of coliphages that infect Gramnegative bacteria, including E. coli, which possess a plasmid coding for an F, or sex, pilus (Vinjé 

et al., 2004). F-specific coliphages are in the bacteriophage families Inoviridae, Leviviridae, and 

Tectiviridae (Cole et al., 2003; Lute et al., 2004; Ogorzaly et al., 2009; Mesquita et al., 2010). Fspecific coliphages in the Inoviridae family are filamentous, ssDNA phages, whereas F-specific 

coliphages in the Leviviridae family are small, icosahedral, ssRNA phages and F-specific 

coliphages in the Tectiviridae family are cubic capsid (icosahedral) with linear dsDNA and no 

tail (Cole et al., 2003; Mesquita et al., 2010). Based on serological cross-reactivity, replicase 

template activity, and phylogenetic analysis, the F-specific RNA coliphages in the Leviviridae 

family have been further broken down into genogroups GI, GII, GIII, and GIV (Vinjé et al., 

2004). In general, GII and GIII F-specific RNA coliphages are mainly found in environments 

associated with human waste, and GI and GIV F-specific RNA coliphages are mostly associated 

with animal waste, although these associations are not absolute (Schaper et al., 2002a; Cole et 

al., 2003; Vinjé et al., 2004). Several host strains of bacteria are used to enumerate F-specific 

coliphages in water samples, including E. coli resistant to streptomycin and ampicillin (Famp) and 
7

Salmonella enterica serovar Typhimurium WG49 (Stm WG49).3 Common laboratory strains of 

F-specific coliphages include MS2 (GI), GA (GII), Qβ (GIII), and SP (GIV) (Vinjé et al., 2004). 

Despite being frequently detected in the environment, data indicate that somatic and F-specific 

coliphages rarely, if ever, replicate in E. coli under environmental conditions (Contreras-Coll et 

al., 2002; Jofre, 2009). Lack of replication in the environment is partially because coliphages do 

not replicate below a bacterial host density of 104 colony-forming units per mL (Wiggins and 

Alexander, 1985; Woody and Cliver, 1997). Additionally, Woody and Cliver (1997) 

demonstrated that the F-specific RNA coliphage Qβ cannot replicate in E. coli in nutrient-poor 

environments, and Cornax et al. (1991) asserted that the low survivability of the E. coli bacterial 

host in marine environments does not support the replication of coliphages. 

F-specific coliphages have not been observed to multiply in E. coli suspended in water (Grabow, 

2001). As described above, F-specific coliphages require the presence of F-pili on the host 

bacteria for infection to occur. In addition to requiring high densities of bacterial hosts for 

replication, the F-pili production requires optimum temperatures between 30 and 37°Celsius (°C) 

with F-pili production decreasing rapidly below temperatures of 25°C (Franke et al., 2009). 

Additionally, most environmental isolates of E. coli have not been observed to produce pili even 

at elevated temperatures and are generally considered unsuitable hosts for F-specific RNA 

coliphages (Luther and Fujioka, 2004). F-specific coliphages can replicate in E. coli in certain 

water environments if fertility fimbriae are present and when the temperature is at least 30°C. 

However, replication under these conditions is unlikely as environmental conditions are not 

likely to support fertility fimbriae production (Grabow et al., 1980). However, some argue that Fspecific RNA coliphages may reproduce under environmental conditions according to the “mud 

puddle hypothesis.” This hypothesis argues that the presence of animal waste lagoons and 

stagnant small puddles in a watershed may provide an environment for the generation of 

coliform bacteria and F-specific and somatic coliphages (Jiang et al., 2007; Reyes and Jiang, 

2010). However, additional research to test whether the coliphages detected on environmental E. 

coli strains can also infect the E. coli strains used in laboratory assays is needed.

2.2. Morphology 

Bacteriophages are incredibly diverse in size, morphology, surface properties, and composition. 

Table 1 briefly describes the structure and morphology of the seven bacteriophage families that 

include coliphages.

 

3

 Stm WG49 contains an E. coli plasmid that codes for sex pili. 
8

Table 1. Morphology of a subsection of bacteriophages.

Type Family (Examples) Nucleic acid Structure

Somatic coliphages Myoviridae (T2, T4) Linear dsDNA Nonenveloped, contractile 

tail, consisting of a sheath 

and central tube

Somatic coliphages and

Bacteroides

bacteriophages

Siphoviridae (λ, T1, T5) Linear dsDNA Nonenveloped, long 

noncontractile tail

Somatic coliphages Podoviridae (T3, T7, P22) Linear dsDNA Nonenveloped, short 

noncontractile tail

Somatic coliphages Microviridae (ΦX174) Circular dsDNA Nonenveloped, isometric

F-specific DNA 

coliphages

Tectiviridae (PR772) Linear dsDNA Nonenveloped, cubic capsid 

(isosahedral), no tail

F-specific RNA 

coliphages (Genogroups I, 

II, III, IV)

Leviviridae (MS2, Qβ, F2) Linear ssRNA Nonenveloped, isometric

F-specific DNA 

coliphages

Inoviridae (M13) Circular ssDNA Nonenveloped, filamentous

Source: Pillai, 2006; Mesquita et al., 2010; Jończyk et al., 2011

Coliphages within families are sometimes grouped by similar traits or have unique 

characteristics, which is important because morphology affects the susceptibility of viruses to 

inactivation in the environment. Some specific structural characteristics, such as tails, large 

capsids, and lack of an envelope, can be associated with greater resistance to external factors, 

such as thermal degradation and degradation in water (Ackermann et al., 2004; Jończyk et al., 

2011). For example, most coliphages belong to the ‘T group’ and have a tail structure. ‘T even’ 

coliphages possess a contractile sheath. T1 and T5 coliphages have long tails without contractile 

sheaths, whereas T3 and T7 coliphages have very short tails (Pillai, 2006; Jończyk et al., 2011). 

While morphology is linked with viral family and type, differences in morphology within these 

classifications have also been observed. For more details on viral properties that affect 

inactivation and environmental persistence of coliphages, see Section 5.0.

As illustrated in Table 2, similar to coliphages (shown in Table 1), many waterborne human 

enteric viruses are nonenveloped and display a range of nucleic acid structures. The virion of 

both coliphages and human enteric viruses consists of either ds or ss RNA or DNA (Pillai, 2006; 

Jończyk et al., 2011; King et al., 2011). Specifically, F-specific RNA coliphages are 

morphologically similar to enteroviruses, caliciviruses, astroviruses, and hepatitis A and E 

viruses, and somatic coliphages are more similar to adenovirus (King et al., 2011).
9

Table 2. Morphology of human enteric viruses that may be transmitted in aquatic 

environments.

Genus &

Common name(s) Nucleic acid Structure

Astrovirus

Astrovirus

Spherical ssRNA Nonenveloped

Calicivirus

Norovirus

Spherical ssRNA Nonenveloped

Coronavirus

Coronavirus

Linear ssRNA Enveloped

Enterovirus

Poliovirus, 

Coxsackievirus A & B, 

Echovirus

Linear ssRNA Nonenveloped

Enterovirus

Hepatitis A

Spherical ssRNA Nonenveloped

Hepevirus

Hepatitis E

Spherical ssRNA Nonenveloped

Mastadenovirus

Adenovirus

Linear dsDNA Nonenveloped

Parvovirus

Parvovirus

Linear ssDNA Nonenveloped

Reovirus

Reovirus

Linear dsRNA (segmented) Nonenveloped

Rotavirus

Rotavirus

Spherical dsRNA (segmented) Nonenveloped

Torovirus

Torovirus

Linear ssRNA Enveloped

Source: Bosch, 1998; King et al., 2011

2.2.1 Morphological Properties Affecting Persistence 

Coliphages can be inactivated, or made noninfective by various environmental factors, including 

temperature (Feng et al., 2003), pH (Feng et al., 2003), salinity (Sinton et al., 2002), sunlight 

(Sinton et al., 1999), and ultraviolet (UV) light (Sang et al., 2007). Viral inactivation occurs 

when viral components (nucleic acids, proteins, lipids) are destroyed. Therefore, characteristics 

that influence survival include coliphage morphology, including size and surface properties 

(Jończyk et al., 2011). 

Of greatest importance, surface conformations, such as whether the virus is enveloped or 

nonenveloped, affects virus inactivation. Due to their nonenveloped nature, NoV, poliovirus, 

coxsackievirus, and echovirus are presumed to be highly resistant to environmental degradation 

and chemical inactivation (Bae and Schwab, 2008). The lipid content of a viral envelop renders 

the virus more sensitive to environmental stress including desiccation and heat, and are generally 

believed to be less resistant to inactivation than non-enveloped viruses (Rosenthal, 2009). 

Coliphages are nonenveloped and are resistant to environmental degradation and chemical 

inactivation similar to other enteric nonenveloped viruses (Havelaar, 1987; Havelaar et al., 1990; 

Yahya and Yanko, 1992; Nasser et al., 1993; Gantzer et al., 1998; Sinton et al., 2002; Hot et al., 

2003; Ackermann et al., 2004; Bitton, 2005; Lodder and de Roda Husman, 2005; Pillai et al., 

2006; Jończyk et al., 2011; Bertrand et al., 2012; Seo et al., 2012; Silverman et al., 2013). 
10

Of additional consideration, differences in tail structure as well as capsid size and structure affect 

bacteriophage survival. For example, Ackermann et al. (2004) found that tailed bacteriophages 

were the most stable in adverse conditions, but found no difference in stability among 

bacteriophages with contractile, noncontractile, or short tails. Bacteriophages with a large capsid 

(100 nanometers [nm] in diameter) were found to have better preservation rates than 

bacteriophages with a smaller capsid (60 nm in diameter) (Ackermann et al., 2004). Lee and 

Sobsey (2011) found small diameter Microviridae to be among the most persistent of several 

tested somatic coliphages in water. Romero et al. (2011) attributed differences in solar 

inactivation rates between MS2 and rotavirus to their differing protein capsid structure and 

genomes, which the authors conclude may be responsible for observed differences in reactivity 

with individual reactive oxygen species. Overall, it is difficult to make generalizations given the 

complexity of interactions between physical characteristics and factors that affect bacteriophage 

survival. 

While there are differences in survival among viruses of different families, there are also 

differences in survival among viruses within the same family (Sobsey and Meschke, 2003; 

Nappier et al., 2006). A study that estimated the survival of several virus families and genera, 

including adenovirus, poliovirus, and coxsackievirus, found that survival varied by virus type 

(Mahl and Sadler, 1975). Siphoviridae with flexible tails are the most persistent in freshwater 

environments under adverse conditions (Muniesa et al., 1999). Additionally, coliphages within 

the same family and with similar structural similarities do not necessarily share the same survival 

characteristics (Jończyk et al., 2011). For example, results from laboratory studies showed that 

different F-specific RNA coliphages differ in their survival in water (Brion et al., 2002; Schaper 

et al., 2002b; Long and Sobsey, 2004; Nappier et al., 2006). There is also demonstrated 

variability within taxonomic types (Brion et al., 2002). 

2.3. Detection Methods 

Currently a variety of methods are available to detect bacteriophages. These include culturebased methods and “rapid” methods (defined as one day or less) which include immunology- and

molecular-based methods. Each type of method has advantages and disadvantages (see Table 3). 

Plaque assays are a typical culture-based technique used for enumerating infectious virus 

particles (ISO, 1995, 2000, 2001; Grabow, 2001; U.S. EPA, 2001a, b; Eaton et al., 2005; 

Rodríguez et al., 2012a). Additionally, there are three bacteriophage methods published by the 

International Organization for Standardization (ISO) for F-specific RNA bacteriophages, somatic 

coliphages, and bacteriophages infecting Bacteroides fragilis (B. fragilis) (ISO, 1995, 2000, 

2001). Rapid methods include immunology based methods (i.e., culture latex agglutination and 

typing [CLAT]), molecular methods (multiple types of PCR), and Fast Phage (a modified rapid 

version of EPA Method 1601) (Brussaard, 2004, 2009; Fong and Lipp, 2005; Kirs and Smith, 

2007; Love and Sobsey, 2007; Gentilomi et al., 2008; Salter et al., 2010; Rodríguez et al., 

2012b). 

2.3.1 Culture-Based Methods 

Standardized culture-based methods are available in both the United States and the European 

Union for the detection of coliphages in water (ISO, 1995, 2000, 2001; U.S. EPA, 2001a, b; 
11

Eaton et al., 2005). The ISO methods have been optimized and tested through interlaboratory 

comparison (Mooijman et al., 2001, 2002, 2005; Muniesa and Jofre, 2007). The ISO Standard 

Method 9224A-F provides protocols for detecting or enumerating coliphages (Eaton et al., 

2005). Two methods for coliphage monitoring in groundwater were approved by EPA in 2001 

(U.S. EPA 2001a, b). These methods include EPA Method 1601 (two-step enrichment process) 

and EPA Method 1602 (single agar layer [SAL] method). EPA Methods 1601 and 1602 have 

undergone multi-laboratory validation (U.S. EPA 2003a, b). The results of these inter-laboratory 

comparisons support the use of these methods in the determination and enumeration of F-specific 

and somatic coliphages in groundwater (U.S. EPA, 2003a, b). These methods are approved in 40 

Code of Federal Regulations Part 136 and can be used for detection of coliphages in wastewater.4

These culture-based methods have been applied to rivers, estuaries, drinking water, surface 

water, storm water, and wastewater (Havelaar, 1987; Davies et al., 2003; Borchardt et al., 2004; 

Lucena et al., 2004; Sobsey et al., 2004; Ballester et al., 2005; Lodder and de Roda Husman, 

2005; Nappier et al., 2006; Stewart-Pullaro et al., 2006; Bonilla et al., 2007; Locas et al., 2007, 

2008; Gomila et al., 2008; Love et al., 2010; Francy et al., 2011; Rodríguez et al., 2012a). 

EPA Method 1601 describes a qualitative two-step enrichment procedure for coliphages and was 

developed to help determine if groundwater is affected by fecal contamination (U.S. EPA, 

2001a). However, this validated4procedure determines the presence or absence of F-specific and 

somatic coliphages in groundwater, surface water, and other waters (U.S. EPA, 2003a). Method 

1601 may be used as a quantitative assay of coliphage densities in a most probable number 

(MPN) format (spot-plating). The Method 1601 protocol directs that a 100 mL or 1 liter (L) 

groundwater sample be enriched with a log-phase host bacteria (E. coli Famp for F-specific 

coliphages and E. coli CN-13 for somatic coliphages) for coliphages. After an overnight 

incubation, samples are put on to a lawn of host bacteria specific for each type of coliphage, 

incubated, and examined for circular lysis zones, which indicate the presence of coliphages. For 

quality control purposes, both a coliphage positive reagent (enumerated sewage filtrate or pure 

cultures of F-specific RNA coliphage MS2 or somatic coliphage ΦX174) water sample and a 

negative reagent water sample (method blank) are analyzed for each type of coliphage with each 

sample batch. This method is considered more sensitive than EPA Method 1602, a SAL 

procedure discussed below (U.S. EPA, 2001a), due to the larger sample volumes used in 1601 

(100 mL to 1 L) compared to Method 1602 (100 mL). In total, EPA Method 1601 requires 28 to 

40 hours for a final result, depending on incubation times (Salter et al., 2010). 

The EPA Method 1602 SAL procedure can be used to quantify coliphages in a sample. The 

Method 1602 protocol directs that a 100 mL water sample may be assayed by adding the logphase host bacteria (E. coli Famp for F-specific coliphage and E. coli CN-13 for somatic 

coliphage) and 100 mL of double-strength molten tryptic soy agar to the sample. The sample is 

then thoroughly mixed and the total volume is poured into multiple plates. After an incubation of 

16 to 24 hours, circular lysis zones (plaques) are counted and summed for all plates from a single 

sample. The quantity of coliphages in a sample is expressed as PFU per 100 mL. For quality 

control purposes, both a coliphage-positive reagent (enumerated sewage filtrate or pure cultures 

of F-specific RNA coliphage MS2 or somatic coliphage ΦX174) water sample and a negative 

4 http://water.epa.gov/scitech/methods/cwa/methods_index.cfm 

Method validation is defined as a process that demonstrates the suitability of an analytic method for its intended purpose (U.S. 

EPA, 2009b). 
12

reagent water sample (method blank) are analyzed for each type of coliphage with each sample 

batch. In total, EPA Method 1602 typically requires an overnight incubation (18 to 24 hours) up 

to 3 days, but results can be obtained in as few as 8 to 10 hours (Salter et al., 2010). 

There are also methods for coliphage detection that use membrane filters to concentrate 

coliphages from a water sample (Sobsey et al., 1990; Sobsey et al., 2004; Eaton et al., 2005). 

Volumes of water of 100 mL and greater can be concentrated on a membrane filter after addition 

of salts and or pH adjustments. Coliphages can then be eluted off the filter and used in one of the 

standard assays above, or they can be enumerated directly on the membrane filter (Eaton et al., 

2005). For direct filter assays, a single assay dish is utilized for each coliphage-adsorbed filter. 

This significantly reduces the time and materials required. However, extraneous material on the 

filter can interfere with the plaque assay. Both 47-millimeter (mm) membrane and 90-mm 

membrane filters have been used and the membrane filtration method can be used to detect both 

F-specific and somatic coliphages. 

One study evaluated the use of a single E. coli host (Escherichia coli host strain CB390) for the 

simultaneous detection of both somatic and F-specific coliphages (Guzmán et al., 2008). This 

host could be useful for detecting total coliphages. However, more independent and multilaboratory validation of this method is needed. Rose et al. (2004) used E. coli C-3000 (ATCC 

#15597), which they report can host both somatic and F-specific coliphages. 

EPA is currently evaluating a membrane filtration culture method and may also evaluate an 

ultrafiltration culture method for use in coliphage enumeration. The intralaboratory (single 

laboratory) method validation study is underway. 

2.3.2 Rapid Methods 

Recently, multiple methods have been published that are faster than EPA Methods 1601 and 

1602. Each method has advantages and disadvantages in terms of speed, accuracy, form of 

results (i.e., quantitative, qualitative, infectivity of virus), and level of training and equipment 

required. The rapid methods are outlined below in more detail. 

Polymerase Chain Reaction Methods 

The most common type of molecular method used to detect coliphages is PCR. PCR is a method 

of amplifying nucleic acids and involves cycling the reaction mixture through temperatures that 

allow for denaturing, annealing, and extension of new DNA fragments or amplicons. With each 

cycle, specific DNA fragments targeted by primers are doubled. This exponential amplification 

of DNA fragments allows samples with very small numbers of target sequences to be amplified 

into an amount of DNA that can be visualized on an agarose gel (Innis et al., 1990). Depending 

on the type of information needed (quantitative, qualitative), different types of PCR are used and 

are described in more detail below. Currently, there are no universal primers for the detection of 

coliphages, but primers are available for individual coliphage strains. 

RT-PCR: RT-PCR is used to determine the presence of RNA or RNA viruses, such as F-specific 

RNA coliphages. The viral RNA is first reverse transcribed into complementary DNA, which is 

used as a template for the PCR reaction (Fong and Lipp, 2005; Kirs and Smith, 2007).
13

Quantitative (q) PCR and RT-qPCR: Both qPCR and RT-qPCR assays, which detect and 

quantify the amount of nucleic acid present, have been developed for the quantification of 

coliphages (Smith, 2006). These assays can determine the amount of a given coliphage present in 

a given sample (Yong et al., 2006; Kirs and Smith, 2007; U.S. EPA, 2007, 2010; Gentilomi et 

al., 2008). These PCR assays often detect only a subgroup of the total coliphages that would be 

quantified by plaque assays. Most recently, PCR has been performed on digital microfluidic 

platforms (Hua et al., 2010; Jebrail and Wheeler, 2010; Mark et al., 2010) and has been used to 

detect bacteriophages (Tadmor et al., 2011) and coliphages (Reitinger et al., 2012). Digital PCR 

on microfluidic chips promises to be a fast and accurate high-throughput technique to determine 

phage genome quantification.

Multiplex PCR: Multiplex PCR (also including multiplex qPCR, RT-PCR, and RT-qPCR) was 

developed to detect multiple target sequences in the same reaction tube. Thus, multiplex PCR 

can detect more than one type of phage in one sample (U.S. EPA, 2007, 2010). For example, RTqPCR only quantitatively detects one type of coliphage per tube (i.e., GII F-specific RNA 

coliphage) while multiplex RT-qPCR quantitatively detects multiple phage targets per tube (i.e.,

GI, GII, and GIII F-specific RNA coliphages) (Kirs and Smith, 2007). 

Culture Latex Agglutination and Typing

The CLAT method has been validated for the detection of coliphages from fecal contamination 

in beach waters (Griffith et al., 2009; Wade et al. 2010) and combines a two-step enrichment 

process and latex agglutination serotyping to monitor for the presence of coliphages (Love and 

Sobsey, 2007; Rodríguez et al., 2012a). This rapid antibody-based method detects F-specific 

coliphages in water samples in 5 to 24 hours. Samples are generally scored as positive based on 

formation of clumps visible on the agglutination card after 60 seconds. Absence of such clumps 

signifies negative samples (Love and Sobsey, 2007). The assay is relatively inexpensive as 

reagents can be stored at ambient temperatures for months, unlike the reagents used for PCRbased assays (Love and Sobsey, 2007). 

Fast Phage Modified Method 1601 

A modification to EPA Method 1601 called Fast Phage has been described by Salter et al. 

(2010). This modification incorporates the use of shelf-stable, ready-to-use reagents in a 

simplified format. Within the Fast Phage method, isopropyl-β-D-1-thiogalactopyranoside is used 

as an enrichment medium to induce transcription of the host E. coli lac operon. Lysis of E. coli 

by coliphages is coupled with lac operon expression. Therefore, a large amplification and a rapid 

extracellular beta-galactosidase enzyme release during coliphage-induced lysis of the infected 

host are reported in comparison to the growing, uninfected host (Salter et al., 2010). Fast Phage 

is approved under EPA’s Alternative Test Procedure program for detection of coliphages in 

groundwater (Salter and Durbin, 2012). 
14

Table 3. Advantages and disadvantages of methods to detect coliphages.

Method Advantages Disadvantages

Culture EPA Method 

1601

 Qualitative (presence/absence);

 Differentiates between F-specific and somatic

coliphages;

 Infectivity is determined;

 More sensitive than Method 1602 (depending

on sample volume: Method 1601 with >100

mL is more sensitive than Method 1602 and

Method 1601 with <100 mL is less sensitive

than Method 1602); and

 Inexpensive.

 Not validated as a quantitative assay;

and

 Requires 24 hours–3 days for results.

Culture EPA Method 

1602

 Both qualitative and quantitative (PFU);

 Differentiates between F-specific and somatic

coliphages;

 Infectivity is determined; and

 Inexpensive.

 Requires 16–24 hours for results; and

 May be less sensitive than Method

1601 (depending on sample volume:

Method 1601 with >100 mL is more

sensitive than Method 1602 and

Method 1601 with <100 mL is less

sensitive than Method 1602).

SM9224F Membrane 

Filtration

 Both qualitative and quantitative (PFU);

 Differentiates between F-specific and somatic

coliphages;

 Infectivity is determined;

 Similar material requirements to EPA Methods

1601 and 1602; and

 Greater than 100 mL volume samples can be

evaluated, which increases sensitivity in

ambient waters.

 Requires 16–24 hours for results; and

 May have recovery loss due to

filtration and elution steps; and

 Turbidity in the sample may interfere

with plaque identification.

PCR/(reversetranscriptase) RT-PCR

 Rapid (~2–10 hours);

 Increased sensitivity compared to culture

methods; and

 Can test for specific types of DNA (PCR) or

RNA (RT-PCR) phages.

 Qualitative only;

 Infectivity (live vs. dead) not

determined;

 Inhibitors may be present in the

environmental matrix; and

 Expensive (PCR equipment) and

quality assurance/quality control

(QA/QC) expertise required.

qPCR/RT-qPCR 

(quantitative)

 Rapid (~2–10 hours);

 Quantitative;

 Increased sensitivity compared to culture

methods; and

 Can test for specific types of DNA (qPCR) or

RNA (RT-qPCR) phages.

 Infectivity not determined;

 Inhibitors may be present in the

environmental matrix; and

 Expensive (qPCR equipment) and

QA/QC expertise required.

Multiplex qPCR/RTqPCR

 Rapid (~2–10 hours).

 Increased sensitivity compared to culture

methods; and

 Can quantitatively distinguish between Fspecific DNA (qPCR) and RNA (RT-qPCR)

subgroups in one reaction tube.

 Infectivity not determined;

 Inhibitors may be present in the

environmental matrix;

 Expensive (qPCR equipment) and

QA/QC expertise required; and

 Multiple sets of primers and probes in

the multiplex qPCR reactions may

cross-react, creating issues in method

specificity (Batra et al., 2013).
15

Method Advantages Disadvantages

CLAT  Same day results (2–24 hours);

 Detects F-specific coliphages and has been 

applied to some somatic coliphage groups 

(Lee, 2009);

 Can differentiate among F-specific 

genogroups;

 Detects infectious coliphages;

 Inexpensive;

 Field portable; and

 When used in an enrichment-CLAT format it 

is as sensitive as EPA Methods 1601 and 1602

(Love, 2007).

 Not quantitative unless implemented 

in an MPN format (Love and Sobsey, 

2007).

Culture Fast Phage  Results within 16–24 hours;

 Differentiates between F-specific and somatic 

coliphages; 

 Infectivity is determined; and

 Considered “equivalent” to EPA Method 1601 

for groundwater monitoring by EPA’s 

Alternate Test Procedures program.

 Qualitative only; and

 Requires laboratory equipment, 

reagents (Fast Phage kit), and 

training.

Note: CLAT and PCR can be field portable, but all the quantitative methods require laboratory facilities.

Sources: ISO, 1995, 2000, 2001; Grabow, 2001; U.S. EPA, 2001a, b, 2007, 2010; Brussaard, 2004; Fong and Lipp, 

2005; Kirs and Smith, 2007; Love and Sobsey, 2007; Gentilomi et al., 2008; Brussaard, 2009; Salter et al., 2010; 

Rodríguez, et al., 2012a, b; Salter and Durbin, 2012.
16

3. Epidemiological Relationships 

Since the 1950s, epidemiological studies have been performed to evaluate relationships between 

fecal indicators and recreational swimming-associated illnesses in surface waters. The incidence 

of symptoms associated with gastrointestinal, eye, ear, and respiratory illnesses has been found 

to be higher in swimmers than in nonswimmers in ambient waters (Prüss, 1998; Wade et al., 

2003; Zmirou et al., 2003). 

Over the past several decades, EPA has conducted numerous epidemiological studies in both 

marine and freshwaters to evaluate the relationship of water quality indicators and human health 

risks. The results of an epidemiological study conducted by Cabelli et al. (1982) found that 

densities of enterococci in marine and freshwaters correlated with incidences of swimmingassociated gastrointestinal illness, whereas densities of E. coli were correlated with swimmingassociated gastrointestinal illness only in freshwaters. EPA’s NEEAR study found that the 

occurrence of gastrointestinal illness in swimmers was positively associated with exposure to 

levels of enterococci enumerated by EPA’s Enterococcus qPCR Method 1611 in marine and 

freshwater (Wade et al., 2008, 2010; U.S. EPA, 2012). The correlation between gastrointestinal 

illness and culturable enterococci in the NEEAR studies was positive, but not as strong as the 

relationship between illness and enterococci enumerated by qPCR. The odds of gastrointestinal 

illness was higher among swimmers compared to non-swimmers on days were coliphages were 

detected, but the associations did not achieve statistical significance (Wade et al., 2010). The 

statistical power was limited due to the relatively few positive results. In addition, only data on 

coliphage presence or absence in 100 mL volume samples were used for the analysis even 

though quantitative data may be available. Thus, further analyses of these data may be needed to 

fully understand the results of the study. 

In 1982, Cabelli et al. suggested that viruses were a primary cause of gastrointestinal illness, in 

agreement with quantitative microbial risk assessment (QMRA) modeling that used data from 

the NEEAR freshwater study (Soller et al., 2015). QMRA modeling demonstrated that the 

illnesses reported during the NEEAR study were consistent with a virus that had an incubation 

period similar to NoV (Soller et al., 2015). However, NoV has not been confirmed as the cause 

of illness in these primary contact recreators. Interestingly, adenovirus (detected by qPCR) has 

been positively associated with gastrointestinal illness at a freshwater beach in Ohio (Lee, 2011).

A consistent association between FIB (E. coli and enterococci) and illness has not been reported 

at all beaches where epidemiological studies have been conducted (Colford et al., 2007). This 

may be due partially to the fact that FIB in surface waters can come from sources other than 

wastewater, such as rainfall, plants, runoff, animals, and human shedding. In some subtropical 

and temperate climates, bacteria, such as E. coli and enterococci, can multiply in the 

environment, giving a false impression of an increase in fecal pollution (Solo-Gabriele et al., 

2000; Yamahara et al., 2009). Additionally, compared to non-spore-forming FIB, human enteric 

viruses have been found to be more persistent in water environments and more resistant to 

physical antagonism, such as heat (55°C) (Lee and Sobsey, 2011). There are clear advantages to 

having alternative indicators (e.g., other than E. coli or enterococci) that have the following 

attributes compared to E. coli and enterococci: The alternative indicators are more closely 

associated with viral gastrointestinal illnesses (e.g., that are present in intestinal microflora of 
17

humans); they do not come from other, non-fecal related sources; they offer improved detection 

methods; they do not multiply in the aquatic environment; and they are more closely linked to 

the pathogens of concern (i.e., often present when viruses are present and absent in 

uncontaminated samples and as resistant to environmental factors as some viral pathogens). 

Numerous studies have been conducted to determine whether both somatic and F-specific 

coliphages are associated with fecal contamination (Chung and Sobsey, 1993; Mocé-Llivina et 

al., 2005; Love and Sobsey, 2007). However, only a limited number of epidemiological studies 

have evaluated the use of coliphages as an indicator of human fecal contamination in recreational 

water. These results are summarized below in chronological order. When available, data on E. 

coli and enterococci are also presented for comparative purposes. 

3.1 Von Schirnding et al. (1992) 

Von Schirnding et al. (1992) conducted a prospective cohort study at two marine beaches in 

South Africa with 733 participants (including adults and children). Beach 1 was described as 

moderately impacted by human sources of fecal contamination, including septic tank overflows, 

feces-contaminated river water, and stormwater runoff. Beach 2 was considered to be less 

impacted by known sources of fecal contamination. Participants were recruited at the two 

beaches. Those who entered the water above their waist were considered “swimmers” and those 

who entered the water up to their waist or who did not enter the water were designated as 

“nonswimmers.” A telephone follow-up call 3 to 4 days later recorded symptoms that developed 

after the beach visit. The symptoms were grouped as gastrointestinal (i.e., diarrhea, vomiting, 

stomachache, and nausea), respiratory (i.e., sore throat, cough, cold, runny/stuffy nose), and skin 

symptoms. 

Water samples were collected on study days at three locations at each beach, both before and 

during maximum swimming activity. The following indicators were evaluated using culturebased methods: fecal coliforms, enterococci, staphylococci, somatic coliphages, and F-specific 

coliphages. The density of fecal coliforms and enterococci was statistically significantly higher 

at Beach 1 than at Beach 2 (median levels of fecal coliforms: 76.5 colony forming units (CFU) 

per 100 mL at Beach 1 and 8.0 CFU per 100 mL at Beach 2; median levels of enterococci: 51.5 

CFU per 100 mL at Beach 1 and 2.0 CFU per 100 mL at Beach 2). Insignificant densities of 

staphylococci and coliphages were detected at both beaches. 

The rates for gastrointestinal, respiratory, and skin symptoms (but not other symptoms including 

wheezing, earache, rashes, allergy, headache, backache) were higher for swimmers than 

nonswimmers at Beach 1, but the differences were not statistically significant. The relative risks 

(RR) of symptoms when comparing swimmers and nonswimmers were 2.45 (95% confidence 

interval [CI]: 0.55–10.9) for gastrointestinal symptoms, 3.28 (95% CI: 0.76–15.26) for 

respiratory symptoms, and 4.06 (95% CI: 0.52–31.72) for skin symptoms. The differences were 

not statistically significant for children younger than 10 years of age or for adults. 

The authors suggested that a possible explanation for the higher rates of symptoms among 

swimmers than nonswimmers at Beach 1 was that the main sources of contamination were likely 

the bathers themselves or the sanitary facilities at the informal settlements close to the study 

beaches. This conclusion is supported by the known sources of contamination at Beach 1 and the 
18

fact that insignificant densities of F-specific coliphages were detected at Beach 1 (because 

coliphages are more closely associated with sewage and septage than direct human fecal inputs). 

At Beach 2, higher rates of respiratory symptoms were observed among nonswimmers than 

swimmers (but the differences were not statistically significant). The authors suggested that this 

apparent anomaly may reflect the presence of a respiratory outbreak in the community (and thus 

children perceived as sick were restricted from swimming by the parents), but because the 

numbers were low and not statistically significant, these findings should not be over-interpreted. 

Overall, the authors felt that the study findings suggested a relationship between swimmingassociated illness and water quality, but that larger study sizes (4,000 subjects) would be needed 

to detect statistically significant differences. 

3.2 Lee et al. (1997) 

Lee et al. (1997) studied the risk of gastrointestinal illness associated with white-water canoeing 

and rafting in a cohort study of 473 canoeists and rafters using an artificial white-water course 

fed by the River Trent in England. The River Trent is a lowland river that receives considerable 

volumes of treated sewage and, during heavy rainfall, untreated sewage from storm overflows. 

The study was conducted on 11 nonconsecutive days between March and December 1995. 

Participants were recruited on the day of the study and given a questionnaire about their 

activities on the course, previous use of the course, medical history, and food eaten in the 

previous week. A second questionnaire (to be returned by prepaid postage 1 to 2 weeks after the 

study) included questions on the range of symptoms (respiratory tract, gastrointestinal, ear and 

eye, and general symptoms), date of onset and duration, additional water sports conducted 

(including at same course), and food eaten in the week after visiting the course. Gastrointestinal 

illness was defined as either vomiting or diarrhea (four or more loose stools in 24 hours), or fever 

combined with nausea, stomach pain, or loose bowels. 

On each study day, water was tested hourly for levels of E. coli, enterococci (fecal streptococci), 

sulfite-reducing clostridia, F-specific RNA coliphages (using ISO method 10705-1), and 

culturable enteroviruses. 

The study found a statistically significant association between risk of gastrointestinal illness and 

density of F-specific RNA coliphages. When comparing the exposure ranges of 26 to 32 PFU per 

10 mL and 69 to 308 PFU per 10 mL to the reference levels of 1 to 3 PFU per 10 mL, the RR of 

gastrointestinal illnesses was 2.6 (95% CI: 1.3–5.2) and 2.8 (95% CI: 1.3–6.0), respectively. 

Other variables significantly associated with increased risk of gastrointestinal illness were 

ingestion of water (RR = 1.9, 95% CI: 1.0–3.6 for swallowing two or more times compared to 

none), accidentally swimming in slalom course (RR = 2.3, 95% CI: 1.2–4.3), and eating and 

drinking before changing clothes (RR = 2.1, 95% CI: 1.1–4.0). Being a regular user of the course 

was associated with a reduced risk of gastrointestinal illness (RR = 1.6, 95% CI: 0.8–3.3) for one 

to six times per year compared to none; and RR = 0.3 (95% CI: 0.1–0.7) for seven or more uses 

compared to none). 
19

The authors stated the observed association between fecal streptococci and E. coli levels and 

gastrointestinal illness risk was not seen after controlling for the stronger association seen with 

F-specific coliphages. The authors concluded that this study demonstrates the value of F-specific 

RNA coliphages as indicators of human fecal contamination associated with risk of 

gastrointestinal illness in recreational water. 

3.3 Medema et al. (1995) and Van Asperen et al. (1998) 

Medema et al. (1995) conducted a pilot study to determine the relationship between 

microbiological freshwater quality parameters and the occurrence of health complaints among 

triathletes (n = 314) using run-bike-runners as controls (n = 81). Information on the occurrence 

of health complaints during the competition and in the week thereafter was collected through a 

written questionnaire. The authors did not link reported illnesses to water quality, other than to 

report the water quality during the time of the triathlon. The geometric means of FIB were 170 E. 

coli CFU 100 per mL and 13 fecal streptococci CFU per 100 mL. F-specific RNA coliphages 

geometric mean was 5.6 PFU per 100 mL. Enteroviruses were present at densities of 0.1 PFU per 

L. Triathletes reported higher rates of symptoms than run-bike-runners: gastrointestinal (7.7 

versus 2.5%), respiratory (5.5 versus 3.7%), skin/mucosal (2.6 versus 1.2%), general (3.5 versus 

1.2%), and total symptoms (14.8 versus 7.4%) in the week after the event. Approximately 75% 

of triathletes reported ingesting water during the swim event. 

Van Asperen et al. (1998) extended the Medema et al. (1995) study over two summers. In a 

prospective cohort design, they followed 827 triathletes and 773 run-bike-run controls. A mailed 

detailed questionnaire collected data about age, sex, general health, medical, and race history, 

exposure to surface freshwaters in the week before and after the race, and occurrence of 

gastrointestinal complaints 2 days before, during, and 6 days after the race. Triathletes were also 

asked about goggle and wetsuit use during the race and if they ingested water during the 

swimming portion of the race. Four different GE endpoints were defined as follows: 

 GE_UK: (diarrhea AND three or more bowel movements per day) OR vomiting OR 

(nausea AND fever); 

 GE_US: vomiting OR (diarrhea AND fever) OR (nausea AND fever) OR (stomach pains 

AND fever); 

 GE_NL-1: (diarrhea AND three or more loose stools movements per day) AND (at least 

two of fever OR nausea OR vomiting OR stomach pains); and 

 GE_NL-2: diarrhea OR nausea OR vomiting OR stomach pains. 

On each exposure day, water samples were collected along the swimming course. Samples were 

analyzed for densities of E. coli, thermotolerant coliforms, fecal streptococci, enteroviruses, and 

reoviruses, F-specific RNA coliphages, Salmonella, Campylobacter, Aeromonas, Plesiomonas 

shigelloides, Pseudomonas aeruginosa, and Staphylococcus aureus. The geometric mean 

(ranges) of the microorganisms during triathlons were: thermotolerant coliforms 78 CFU per 100 

mL (0.6 to 650 CFU per 100 mL), E. coli 204 CFU per 100 mL (11 to 2,600 CFU per 100 mL), 

fecal streptococci 16 CFU per 100 mL (0.2 to 1,800 CFU per 100 mL), enteroviruses 0.04 PFU 

per L (0.007 to 7 PFU per L), and F-specific RNA coliphages 0.7 PFU per L (0.01 to 13.6 PFU 

per L). 
20

Depending on the case definition, the attack rates of GE in the week after the event were 

observed to be higher among triathletes than among run-bike-runners, with odds ratios (ORs) 

ranging from 1.6 to 2.3. The adjusted risk of GE ranged from 2.9 to 4.7, depending on the case 

definition. All ORs were statistically significant. 

The study showed that both E. coli and thermotolerant coliforms were associated with risk of 

gastrointestinal illness after bathing in freshwaters. The authors noted that levels of E. coli were 

higher than the levels of thermotolerant coliforms, and that the densities of E. coli were much 

more closely correlated with illness rates than the densities of thermotolerant coliforms. A 

relationship between health and fecal streptococci, enteroviruses, and F-specific RNA coliphages 

was not observed (Van Asperen et al., 1998). 

3.4 Wiedenmann et al. (2006) 

Wiedenmann et al. (2006) conducted a randomized control epidemiological study at five 

freshwater bathing beaches in Germany. The probable or possible sources of fecal contamination 

at these sites varied, but included raw and treated municipal effluent, agricultural runoff, and 

contamination from water fowl. Only one of the five lakes had a known point-source of 

contamination. A cohort of 2,196 participants (including adults, children, and teenagers) was 

recruited from the local population and randomized into bathers and nonbathers. Two to three 

days prior to exposure, participants were interviewed and underwent a brief medical 

examination. Bathers were exposed to water for 10 minutes and were asked to immerse their 

heads at least three times. Nonbathers made no contact with the water. One week after exposure, 

all participants were interviewed and underwent medical inspection of the throat, eyes, and ears. 

Three different GE endpoints were defined as follows: 

 GE_UK: (diarrhea AND three or more bowel movements per day) OR vomiting OR 

(nausea AND fever) OR (indigestion AND fever); 

 GE_UK-wf: (GE_UK without consideration of stool frequency: diarrhea OR vomiting 

OR (nausea AND fever) OR (indigestion AND fever); and 

 GE_NL-2: diarrhea OR nausea OR vomiting OR stomach pains. 

Water samples were collected at 20-minute intervals from swimming and nonswimming zones 

during the study period. The following microbiological parameters were evaluated: E. coli, 

enterococci, Clostridium perfringens, aeromonads, pyocyanine-positive Pseudomonas 

aeruginosa, and somatic coliphages. The median densities (ranges) of the microorganisms were: 

20 somatic coliphages PFU per 100 mL (10 to 3,780 PFU per 100 mL; method ISO 10705-2); 

136 E. coli CFU per 100 mL (4.7 to 5,344 CFU per 100 mL), 37 intestinal enterococci CFU per 

100 mL (3.0 to 1,504 CFU per 100 mL), 15 Clostridium perfringens CFU per 100 mL (9 to 260

CFU per 100 mL), 8,200 aeromonads CFU per 100 mL (600 to 31,400 CFU per 100 mL), and 10 

Pseudomonas aeruginosa CFU per 100 mL (10 to 100 CFU per 100 mL). 

For somatic coliphages, the no observed adverse effect level (NOAEL) was 10 PFU per 100 mL 

for the two less stringent (broader) illness definitions (GE_UK-wf and GE_NL-2) and 150 PFU 

per 100 mL for the most stringent (most narrowly defined) illness definition (GE_UK). The RRs

of GE_NL-2, GE_UK-wf, and GE_UK when comparing bathing in waters with somatic 

coliphage levels above the NOAEL, with nonbathing were statistically significant and ranged 

from 1.8 (95% CI: 1.2–2.6), 2.5 (95% CI: 1.5–4.0), and 4.6 (95% CI: 2.1–10.1), respectively. For 
21

all three illness definitions, swallowing water with somatic coliphage levels above the NOAEL 

resulted in a significantly higher attributable risk of illness than not swallowing water. 

The authors concluded that reasonable estimates for NOAELs at an average bathing intensity site 

are 100 E. coli CFU per 100 mL, 25 enterococci CFU per 100 mL, and 10 somatic coliphages 

PFU per 100 mL. Wiedenmann et al. (2006) concluded that a NOAEL approach would be 

practical for setting recreational water standards. The authors suggested that somatic coliphages 

would be appropriate alternative fecal indicators that could be used to set standards for 

freshwater just as well as E. coli and enterococci, especially in tropical climates, where E. coli

and enterococci may be less reliable as indicator organisms. 

3.5 Colford et al. (2005, 2007) 

Colford et al. (2005, 2007) conducted a prospective cohort epidemiological study at six beaches 

near Mission Bay, California in 2003. The study cohort consisted of nearly 8,000 participants. 

The authors reported that MST evaluation at Mission Bay suggested that only a minor portion of 

fecal input was from human point sources during the study period. 

Water quality was monitored using traditional FIB enumeration methods (culturable enterococci, 

fecal coliforms, and total coliforms) and a subset of samples was also evaluated using: (1) new 

methods for measuring traditional FIB (chromogenic substrate or qPCR), (2) Bacteroides, (3) 

coliphages (somatic and F-specific coliphages), and (4) human enteric viruses (adenovirus and 

NoV). F-specific and somatic coliphages were detected and quantified in 1 L volumes of water 

by a modification of EPA Method 1601 for enrichment and spot plating that provides a MPN 

estimate of coliphage density. Roughly 68% of the samples had detectable levels of somatic 

coliphages and maximum densities were observed near 36 MPN per 100 mL. F-specific 

coliphages were detected in 11% of the samples and maximum densities reached only one MPN 

per 100 mL. No NoV was found and adenovirus was found only in one sample. The observed 

geometric means for enterococci (measured by qPCR) and fecal coliforms were 65 estimated 

number per 100 mL and 25 MPN per 100 mL, respectively. 

Interviewers recorded which water sampling site was closest to the location of the individual or 

family on the beach. Participants were asked to complete a questionnaire prior to their departure 

from the beach. The questionnaire assessed possible exposures at the beach, and exposures or 

illnesses experienced during the previous two to three days. A follow-up telephone interview was 

conducted 10 to 14 days after the study to gain information on health outcomes. Health outcomes 

measured in the investigation included gastrointestinal illness, respiratory symptoms, and skin 

ailments. Two definitions of highly credible gastrointestinal illness (HCGI) were measured. One 

(HCGI-1) was defined as (1) vomiting; (2) diarrhea and fever; or (3) cramps and fever. The 

second (HCGI-2) was defined as vomiting plus fever. Multivariate analysis was conducted to 

assess relationships between health outcomes and degree of water contact or levels of water 

quality indicators. These analyses were adjusted for confounding covariates such as age, gender, 

and ethnicity. 

Of the measured health outcomes, only skin rash and diarrhea were consistently significantly 

elevated in swimmers compared to nonswimmers. For diarrhea, this risk was strongest among 

children 5 to 12 years old. No correlation was found between increased risk of illness and levels 
22

of Bacteroides or Enterococcus, as detected using rapid methods (qPCR) or for somatic 

coliphages. A significant association was observed between the levels of F-specific coliphages 

and HCGI-1, HCGI-2, nausea, cough, and fever. The adjusted odds ratio (AOR) was 1.26 (95% 

CI: 1.06–1.48) for HCGI-1; 1.43 (95% CI: 1.13–1.82) for HCGI-2; 1.34 (95% CI: 1.16–1.55) for 

nausea; 1.22 (95% CI: 1.02–1.48) for cough; and 1.25 (95% CI: 1.09–1.44) for fever. Colford et 

al. (2005, 2007) suggested that these associations be interpreted cautiously because only a small 

number of participants were exposed to the water at times when F-specific coliphages were 

detected. 

3.6 Wade et al. (2010) 

Wade et al. (2010) enrolled 6,350 participants in prospective cohort epidemiological studies 

conducted at three marine beaches. These beaches were located in Mississippi (Edgewater 

Beach), Rhode Island (Goddard Beach), and Alabama (Fairhope Beach), and were known to be 

impacted by discharge from nearby WWTPs. The study in Mississippi was conducted in 2005 

and studies in Rhode Island and Alabama were conducted in 2007. Upon study enrollment, 

participants were interviewed to gather information on exposure and health status and completed 

a questionnaire prior to exiting the beach for the day. Based on their activities for the day, 

participants were divided into cohorts that included swimmers and nonswimmers. Swimming 

was defined as body immersion (i.e., immersion to the waist or higher). Nonswimmers were 

considered unexposed to recreational water. Health endpoints evaluated during the study 

included upper respiratory illness (defined as any two of the following: sore throat, runny nose, 

cough, cold, or fever), earache, eye irritation, rash, and gastrointestinal illness (defined as any of 

the following: (1) diarrhea (three or more loose stools in a 24-hour period); (2) vomiting; (3) 

nausea and stomachache; or (4) nausea or stomachache, and interference with regular activities 

(missed regular activities as a result of the illness). 

Water samples were collected in duplicate at three different time points along three transects 

perpendicular to the shoreline on each study day. A total of 1,242 water samples were collected. 

Water samples were tested for a variety of indicators, including a faster test for F-specific 

coliphages based on a CLAT assay, which also distinguishes F-specific RNA coliphages and 

F-specific DNA coliphages. F-specific coliphages were also evaluated using a modified version 

of EPA Method 1601, called the 24-hour SPOT assay. Samples were tested for Enterococcus

spp. using EPA Method 1600 (a culture-based method) and Enterococcus and Bacteroidales by 

qPCR. 

Wade et al. (2010) reported that 56% (100 of 222) of samples at Fairhope Beach and 65% (203 

of 425) of samples at Goddard Beach were positive for F-specific coliphages by the modified 

EPA Method 1601.

5 Fewer samples were positive for F-specific coliphages by the CLAT assay. 

At Fairhope Beach, 4% (8 of 228) and 6% (14 of 224) of samples were positive for F-specific 

RNA and F-specific DNA coliphages, respectively. At Goddard Beach, 7% (31 of 425) and 9% 

(37 of 423) of samples were positive for F-specific RNA and F-specific DNA coliphages, 

respectively. The AOR of gastrointestinal illness was significantly higher among swimmers 

compared to nonswimmers on days when F-specific RNA (AOR = 1.80, 95% CI: 1.22–2.66) or 

 

5 Edgewater Beach, data collection was stopped several days early due to the effects of Hurricane Katrina. 

Bacteriophage results were not reported for Edgewater Beach. 
23

F-specific DNA coliphages (AOR = 1.69, 95% CI: 1.16–2.47) were detected by the CLAT assay 

or F-specific coliphages were detected by a modified 1601 method (AOR = 1.70, 95% CI: 1.12–

2.57). An increased, but not statistically significant risk of gastrointestinal illness among 

swimmers was observed for a 1-log10 increase in each of the three F-specific coliphages 

measured. F-specific coliphages measured by the modified 1601 method were not associated 

with gastrointestinal illness among swimmers. Other illnesses (i.e., respiratory illness, earache) 

did not show a relationship with the presence of coliphages. 

The risks of both gastrointestinal illness and diarrhea were significantly associated with exposure 

to Enterococcus and Bacteroidales (enumerated using qPCR). F-specific coliphages, using the 

modified 1601 method, had a positive correlation with gastrointestinal illness in marine waters, 

but like culturable enterococci, the association was not significant over the full range of water 

quality (Wade et al., 2010). 

3.7 Abdelzaher et al. (2011) 

Abdelzaher et al. (2011) performed a randomized control exposure epidemiological study to 

evaluate water quality and daily cumulative health effects for bathers at a nonpoint source 

subtropical marine recreational beach in Miami, Florida. Study participants were randomly 

assigned to either the ‘bather’ or the ‘nonbather’ categories. Those assigned to the bather 

category were asked to spend 15 minutes in the water and nonbathers were asked to spend 15 

minutes on the beach only. The daily number of bathers varied over the course of the study, with 

a total of 652 bathers (daily average = 43, daily range = 29–55). Similarly, for nonbathers, the 

total number was 651 (daily average = 43, and daily range = 25–60). 

Health effects considered during the study included gastrointestinal illness, skin ailments, and 

respiratory illness. Gastrointestinal illness was defined as all cases of vomiting or diarrhea, or all 

reported cases of indigestion or nausea accompanied by a fever. ‘Diarrhea’ was defined as 

having three or more runny stools within a 24-hour period. 

Water samples were categorized as “daily composite samples,” which were combined water 

samples collected throughout each sampling day either by bathers or study staff. Bather-collected 

samples were analyzed for a variety of indicator organisms, including enterococci (using three 

detection methods: membrane filtration, chromogenic substrate, and qPCR), fecal coliform, E. 

coli, Clostridium perfringens (all measured by membrane filtration), somatic and F-specific 

coliphages (measure by EPA Method 1602), human- and dog-associated MST markers 

(Bacteroides thetaiotaomicron, BacHum-UCD, HF8, and DogBac), human polyomavirus, and 

the esp gene of Enterococcus faecium. Pathogens evaluated in the study included: 

Staphylococcus aureus, Vibrio vulnificus, the protozoa Cryptosporidium spp. and Giardia spp., 

NoV, enterovirus, and hepatitis A virus. Investigator-collected composite samples were used for 

pathogen analysis using traditional large-volume concentration methods. 

Average daily excess illness percentage rates (calculated by subtracting the daily illness rates for 

nonswimmers from that for swimmers) for gastrointestinal, skin, and acute febrile respiratory 

illness were 2.0% (standard deviation [SD] = 3.3), 5.6% (SD = 4.7), and 1.2% (SD = 2.9), 

respectively. No statistically significant correlations between health outcomes and any of the 

indicator organisms, including coliphages, were identified in this investigation. 
24

Somatic coliphages were detected (range 0.3 to 1.7 PFU per 100 mL) on four of 15 days tested. 

On three of five days where the greatest level of excess gastrointestinal illness occurred, somatic 

coliphages were detected. Although no statistically significant associations between water 

quality and illness were observed, the authors state that this overlap is suggestive of a potential 

correspondence between the presence of somatic coliphages and increased risk of gastrointestinal 

illness. Given the low number of positive samples and that F-specific coliphages were not 

detected in any of the samples, no apparent association between this potential indicator and 

health outcomes was observed in this study. The authors suggest that a possible reason F-specific 

coliphages were not detected may be due to the small volume of water for each sample (100 mL) 

compared to other studies, such as Colford et al. (2005, 2007) who used 1 L samples, thereby 

increasing detection limits. 

3.8 Griffith et al. (personal communication, 2015) 

The Southern California Coastal Water Research Project Authority conducted two prospective 

cohort studies at California beaches (Avalon Bay (Avalon), Doheny State Beach (Doheny)) in 

2007 and 2008. Both Avalon and Doheny were impacted by faulty sanitary sewer infrastructure, 

which allowed microbial contamination to reach the beach via groundwater. 

The studies enrolled 8,226 swimmers across the two beaches and each swimmer’s water 

exposure was recorded. Water samples were collected several times per day at multiple locations 

at each beach and analyzed for up to 30 target indicators using more than 50 different 

methodologies. Interviewers contacted participants by phone 10 to 14 days later and recorded 

symptoms of gastrointestinal illness occurring after their beach visit. Regression models were 

used to evaluate the association between water quality indicators and gastrointestinal illness 

among swimmers at each beach. 

In these two studies, F-specific coliphages measured by EPA Method 1602 had a stronger 

association with health outcomes than did culturable enterococci measured by EPA Method 1600 

at Doheny and Avalon beaches. When all environmental conditions were considered in aggregate 

at Doheny, the OR for F-specific coliphages was 1.9 and statistically greater than 1 (p<0.05), 

whereas the OR for enterococci was only 1.2 and not significant (p>0.05). At Avalon, the OR for 

F-specific coliphages was 1.9 compared to less than 1.1 for enterococci, though neither was 

significantly different than 1.0 (p>0.05). Under highrisk conditions, F-specific coliphages were 

significantly associated with gastrointestinal illness (p<0.05) and the estimated OR was more 

than double that for culturable enterococci at both Avalon and Doheny. Associations were also 

found between F-specific coliphages and adenovirus observed at Doheny Beach (Love et al. 

2014). The authors noted that when the contamination source is primarily human fecal material, 

indicators like F-specific coliphages are better predictors of the health risk. 

3.9 General Conclusions from Epidemiological Studies 

Eight epidemiological investigations have evaluated the relationship between swimmingassociated illness and presence of coliphages. Studies specifically evaluating the link between 

levels of somatic or F-specific coliphages and incidence of illness resulting from exposure to 

fresh and marine waters are summarized in Table 4. 
25

With the exception of one small study (Von Schirnding et al., 1992), all of the epidemiological 

investigations that evaluated coliphages detected somatic or F-specific coliphages. There is 

considerable heterogeneity in the results of these studies, even within similar designs. For 

example, some studies found a significant association between the levels of F-specific coliphages 

and two definitions of gastrointestinal illness, cough, fever, and nausea, but found no association 

between increased risk of illness and levels of somatic coliphages, Bacteroides, or Enterococcus 

(Colford et al., 2005, 2007). Similarly, when comparing swimmers to nonswimmers on days 

when F-specific RNA or DNA coliphages were detected, Wade et al. (2010) found statistically 

significant increases in risk of gastrointestinal illness. In these cases, F-specific coliphages were 

potentially useful indicators. 

On the other hand, at a marine recreational beach in Miami with no known point source of 

contamination, Abdelzaher et al. (2011) detected somatic coliphages (range 0.3 to 1.7 PFU per 

100 mL) on four of 15 days tested, three of which were on days characterized by the highest 

excess gastrointestinal illness. However, F-specific coliphages were not detected in any of the 

samples, and no statistically significant correlations between water quality and illness were 

found. In this case somatic coliphages may have been useful, but F-specific coliphages were not 

useful, due to being below the detection limits of methods. 

Overall, the epidemiological evidence is suggestive of a potential relationship between 

coliphages and human health. In more than half the studies (Lee et al., 1997; Colford et al., 2005, 

2007; Wiedenmann et al., 2006; Abdelzaher et al., 2011, Wade et al., 2010), the presence of 

coliphages was associated with swimming-associated gastrointestinal illness. Wade et al. (2010) 

found that the AOR of gastrointestinal illness was higher among swimmers compared to 

nonswimmers on days when F-specific RNA and DNA coliphages were detected. These studies 

suggest that somatic (Wiedenmann et al., 2006) and F-specific coliphages (Lee et al., 1997; 

Colford et al., 2005, 2007; Wade et al., 2010; Griffith et al., personal communication, 2015) hold 

potential as feasible alternative water quality indicators in marine and freshwaters, with and 

without point-source contamination (Lee et al., 1997; Colford et al., 2005, 2007; Wiedenmann et 

al., 2006; Wade et al., 2010; Abdelzaher et al., 2011). As mentioned in Section 1.3, a good 

indicator should be correlated to health risk. Evaluation of the results of these eight 

epidemiological studies suggests that overall the studies support coliphages as potential 

indicators of gastrointestinal illness from recreational exposures. 
26

Table 4. Summary of epidemiological studies.

Study,

Sample size,

Water type

Indicators evaluated Results

Supports 

coliphages as water 

quality indicator?

Von Schirnding et al. 

(1992), 

n = 733, 

marine

Somatic 

coliphages and Fspecific coliphages, 

fecal coliforms, 

enterococci

Low densities of coliphages were detected at 

both beaches. Rates for gastrointestinal,

respiratory, and skin symptoms were higher for 

swimmers than nonswimmers at Beach 1, but 

the results were not statistically significant.

Coliphage levels 

were too low to 

evaluate.

Lee et al. (1997), 

n = 473, 

fresh

F-specific RNA 

coliphages, E. coli, 

enterococci, culturable 

enteroviruses

Statistically significant association between risk 

of gastrointestinal illness and density of Fspecific RNA coliphages. The observed 

association between fecal streptococci and E. 

coli levels and risk of gastrointestinal illness 

was not seen after controlling for the stronger 

association seen with F-specific coliphages.

Yes; F-specific 

coliphages

van Asperen (1998), 

827 triathletes and 

773 run-bike-run 

controls, 

Fresh

F-specific coliphages,

E. coli, fecal

streptococci, 

thermotolerant 

coliforms, 

enteroviruses

Risk of gastrointestinal illness increased 

significantly at levels with thermotolerant 

coliforms (≥220 CFU per 100 mL) or E. coli

(≥355 CFU per 100 mL), compared to lower 

levels (≤ 120 CFU per 100 mL tolerant 

coliforms or ≤238 CFU per 100 mL for E. coli). 

No exposure-response relationship observed for 

F-specific coliphages, fecal streptococci, or 

enteroviruses.

No

Wiedenmann et al. 

(2006),

n = 2,196, 

fresh

Somatic coliphages, E. 

coli, enterococci

Significantly increased RR of gastroenteritis for 

bathing in waters with somatic coliphage levels 

above the NOAEL (10 PFU per 100 mL) versus 

nonbathing.

Yes; somatic 

coliphages

Colford et al. (2005, 

2007),

n = 8,000, 

marine

F-specific coliphages 

(qPCR), somatic 

coliphages, culturable 

enterococci, fecal 

coliforms, total 

coliforms, adenovirus,

and NoV

Significant association between the levels of Fspecific coliphages and HCGI-1, HCGI-2, 

nausea, cough, and fever.

Yes; F-specific 

coliphages

Wade et al. (2010), 

n = 6,350, 

marine

F-specific RNA 

coliphages (CLAT), Fspecific DNA 

coliphages (CLAT), Fspecific coliphages 

(modified EPA 

Method 1601), 

enterococci

Significantly higher risk of gastrointestinal 

illness comparing swimmers with nonswimmers 

on days when coliphages were present.

Yes; F-specific 

coliphages

Abdelzaher et al. 

(2011), 

n = 652, 

marine

Somatic coliphages, 

enterococci, fecal 

coliforms, E. coli

No statistically significant correlations between 

health outcomes and any indicator organisms, 

including somatic coliphages.

Somatic coliphages

detection overlaps 

with highest illness 

days.

Griffith et al. 

(personal 

communication, 

2015)

F-specific coliphages 

(Method 1602); 

enterococci, and 30 

target indicators with 

50 different 

methodologies.

F-specific coliphages (measured using EPA 

Method 1602) had a stronger association with 

health outcomes than EPA Method 1600 at the 

two beaches studied.

Yes; F-specific

coliphages
27

4. Occurrence in the Environment

Coliphages, including F-specific DNA, F-specific RNA coliphages, and somatic coliphages have 

been detected and proposed as indicators of fecal contamination in a variety of environments. 

Most studies investigating coliphages as fecal indicators focused on environments that may be 

contaminated with human or animal fecal matter, such as water entering or exiting sewage 

treatment facilities, stormwater, natural lakes, rivers, streams, groundwater, seawater, and beach 

sand (Zaiss 1981; Sogaard, 1983; Payment et al., 1988; Araujo et al., 1997; Paul et al., 1997; 

Gantzer et al., 1998; Davies et al., 2003; Bonilla et al., 2007; Charles et al., 2009; Haramoto et 

al., 2009, 2011; Payment and Locas, 2011; Wu et al., 2011). 

Studies investigating the presence of coliphages and viruses in different types of environmental 

waters are described below (Section 4.1). A review of the literature shows that generalizations 

across studies are difficult because the detection of microorganisms from fecal contamination, 

including viruses and coliphages are inconsistent and dependent on a number of important 

factors (WHO, 2001). Generally, when any two studies on coliphages and viruses are compared, 

there are differences between the type of detection method used – both for the coliphages and the 

pathogen. In addition to different detection methods, the differences between studies might 

include the following: type of coliphage tested (i.e., somatic, F-specific DNA, F-specific RNA); 

specific pathogens tested; number of samples taken; volume of sample taken; level of 

contamination; type of environment from which samples were taken; location of the 

environment; resistance of the coliphages and pathogens to environmental stressors and growth; 

transport characteristics of the coliphages and pathogens; carriage rates and shedding patterns of 

the coliphages and pathogens among host populations; presence of host populations; waste 

management practices; rainfall; time of year; and statistical analyses used (WHO, 2001; Bonilla 

et al., 2007; Wu et al., 2011). Given these differences along with the variable occurrence of 

viruses in fecal sources, it is not surprising that the presence of fecal indicators including 

coliphages and the presence of enteric viruses varies between studies. 

4.1. Associations between Coliphages and Viruses 

Some studies have reported an association between the presence of coliphages and human 

viruses (Havelaar et al., 1993; Jiang et al., 2001; Ballester et al., 2005), while other studies have 

found no association between their presence (Ibarluzea et al., 2007; Jiang et al., 2007; Boehm et 

al., 2009; Viau et al., 2011b). Meta-analyses of peer-reviewed studies looking at the occurrence 

of microbial indicators and pathogens, including coliphages and viruses, can give an overview of 

the field. 

In one recent study, Wu et al. (2011) analyzed a broad range of 540 indicator-pathogen pairs 

from studies conducted between 1970 and 2009 in a variety of water environments including: 

rivers, lakes, reservoirs, ponds, estuaries, costal and marine waters, and wastewater (Wu et al., 

2011). Groundwaters, treated drinking waters, and sand/sediments were not included in the 

study. The data were analyzed using a logistic regression model adjusted for indicator classes, 

pathogen classes, water types, pathogen sources, sample size, the number of samples with 

pathogens, the detection method, year of publication, and statistical method. The association is 

presented as an OR, where an OR greater than one signifies that the presence of the indicator is 

associated with the presence of the pathogen. Not surprisingly, no single indicator was 
28

significantly correlated with all the pathogens evaluated.6 Coliphages (F-specific and somatic 

together) and F-specific coliphage densities were more likely to be correlated with pathogens 

than the other traditional indicators (E. coli, enterococci, and fecal coliforms) (Wu et al., 2011). 

The associations between coliphages and pathogens were not statistically significant (OR = 1.29, 

p-value = 0.186 and OR = 1.27, p-value = 0.625, respectively). See Table 5 below for specific 

OR and p-values between different categories of coliphages or indicators and pathogens in water. 

Silva et al. (2010) also found that in water samples collected from 16 beaches along the 

Portuguese coast there was no relationship between viral detection (hepatitis A and NoV) and the 

European regulatory-based bacterial indicators total coliform, fecal coliform, E. coli, and fecal 

enterococci.

Table 5. Number of cases and outcome of the logistical regression analysis of the 

association between coliphages and pathogens in water.

Fecal indicator Number of casesaOR Value p-Value 95% Confidence limits Uncorrelated Correlated

Coliphagesb 45 40 1.29 0.186 0.82 2.05

F-specific 

coliphages

24 16 1.27 0.625 0.48 3.35

F-specific RNA 

coliphages

15 8 0.75 0.518 0.31 1.80

Somatic coliphages 20 10 0.70 0.364 0.32 1.52

E. coli 29 11 0.52 0.070 0.25 1.06

Enterococci 34 12 0.47 0.032 0.24 0.94

Fecal coliforms 78 48 0.84 0.405 0.56 1.27

Source: Based on Table 2 in Wu et al. (2011).

a An individual case of an indicator-pathogen pair represents a statistical analysis of a published dataset of one 

indicator type with one pathogen type where the methods of statistical analysis, correlation coefficients, and pvalues were reported.

b Includes F-specific and somatic coliphages.

OR values above 1 are in bold.

Studies have evaluated the association between pathogens and different subsets of coliphages 

(i.e., somatic, F-specific DNA and RNA) and report variable results which are influenced by the 

environments in which the studies are conducted (Ballester et al., 2005; Savichtcheva and Okabe, 

2006; Payment and Locas, 2011). For example, Wu et al. (2011) report that no indicatorpathogen pairs were significantly associated, except for F-specific coliphage-adenovirus pairs 

(OR = 25.5, p-value = 0.019) (see Table 6 below). Wu et al. (2011) also found that the 

association between indicators and pathogens is significantly stronger in brackish and saline 

water than in freshwater. Therefore, the papers in this chapter are separated into those studies 

conducted in freshwater and those conducted in saline or brackish water. Because Wu et al. 

(2011) conducted a meta-analysis, which is summarized above, that includes most of the studies 

comparing coliphages to human viruses, only a few of the illustrative studies that compare 

coliphages to human viruses in ambient water are summarized in Sections 4.1.1 and 4.1.2 below. 

 

6

 Individual articles evaluated different pathogens. Pathogens (and pathogen genes) paired with indicators included 

Giardia, Cryptosporidium, Campylobacter, Helicobacter pylori, Salmonella, shiga toxin genes, Pseudomonas

aeruginosa, Aeromonads, Vibrio, Staphylococcus aureus, hepatitis A virus, adenoviruses, astroviruses, NoVs, 

sapoviruses, enteroviruses, human enteric viruses, filamentous fungi, yeasts, and Candida albicans. 
29

Table 6. Logistic regression of the association between indicators and different pathogens 

in water.

Pathogens

Indicators

F-specific coliphages

OR value (p-value); 

[95% Confidence 

Limits]

Somatic coliphages

OR value (p-value); 

[95% Confidence 

Limits]

E. coli; 

[95% Confidence 

Limits]

Enterococci; 

[95% Confidence 

Limits]

Adenoviruses 25.5 (p = 0.019); 

[1.72, 377.92]

1.25 (p = 0.862); 

[0.10, 15.50]

NR NR

Cryptosporidium

spp.

NR 0.74 (p = 0.791); 

[0.08, 6.97]

NR 0.73 (0.700);

[0.14, 3.70]

Enteroviruses 1.2 (p = 0.810);

[0.27, 5.29]

NR 1.19 (0.869);

[0.16, 8.99]

0.87 (0.858);

[0.18, 4.23]

Giardia spp. NR 1.06 (p = 0.965);

[0.09, 12.42]

NR 1.06 (0.950);

[0.18, 6.36]

Note: Data are from Wu et al. (2011).

Numbers in the table are the OR values followed by the p-values in parentheses. OR values above 1 are in bold. 

NR (not reported) indicates that the data were not included in the paper. 

Pathogens and indicators are listed in alphabetical order. 

Effects of human virus detection methods on associations between fecal indicators and 

pathogens 

As briefly described above in Section 2.3, there are currently numerous methods to detect human 

viruses. These include culture methods, molecular methods, and a combination of the two (Fong 

and Lipp, 2005). Similar to coliphage detection methods, each method has advantages and 

disadvantages, which in turn affect the type of data collected, including both quantity and the 

type(s) of virus(es) detected. An overview of the strengths and weaknesses of each enteric virus 

detection method is shown below in Table 7. For example, according to Mocé-Llivina et al.

(2005), genomic techniques used to detect human enteroviruses and other human viruses have 

detection rates from 7 to 70% and are not always consistent with the values of other methods for 

enumerating the same organisms. Reasons for the variability between PCR and culture-based 

techniques are due in part to: (1) PCR does not distinguish between infectious and noninfectious 

viruses (i.e., live and dead viruses); (2) the high sensitivity of PCR may contribute to artifacts, 

which could result in false positives; and (3) natural inhibitors in the environment may reduce or 

block PCR amplification resulting in false negatives or under-representation of infectious viruses 

(Fong and Lipp, 2005; Mocé-Llivina et al., 2005). It is important to keep in mind that differences 

in enteric virus detection methods (see Table 7) combined with differences in coliphage 

detection methods (see Table 3) may greatly affect the presence, absence and/or strength of 

correlations found between coliphages and enteric viruses. 
30

Table 7. Comparison of common methods for the detection of pathogenic human enteric 

viruses from environmental sources.

Method Advantages Disadvantages

Cell culture  Provides quantitative data; and 

 Infectivity can be determined.

 Lengthy processing time (takes days to 

weeks); 

 Relatively more expensive than PCR;

and

 Not all viruses from environmental 

samples can grow in cell culture (e.g., 

NoV).

PCR (RT-PCR)  Rapid; 

 Can be quantitative (e.g., end point 

analysis); and

 Increased sensitivity and specificity 

compared to cell culture.

 Usually qualitative; 

 Inhibitors may be present in the 

environmental matrix; and

 Infectivity cannot be determineda.

Nested PCR 

(semi-/heminested)

 Increased sensitivity compared to 

conventional PCR; and

 Can replace PCR confirmation 

steps, such as hybridization.

 Qualitative only; 

 Inhibitors may be present in the 

environmental matrix; 

 Potential risk of carryover 

contamination when transferring PCR 

products; and

 Infectivity cannot be determineda.

Multiplex PCR and Multiplex 

RT-PCR

 Several types, groups, or species of 

viruses can be detected in a single 

reaction; and

 Saves time and cost compared to 

PCR.

 Difficult to achieve equal sensitivity 

for all targeted virus species, groups, or

types; 

 May produce nonspecific amplification 

in environmental samples); 

 Inhibitors may be present in the 

environmental matrix; and

 Infectivity cannot be determineda.

qPCR/RT-qPCR  Provides quantitative data; 

 Confirmation of PCR products is 

not required (saves time); and

 Can be done in a closed system, 

which reduces risk of 

contamination compared to nested 

PCR.

 The lower limit of quantification is 

higher than the lower limit of detection, 

so qPCR can be considered less 

sensitive than presence/absence PCR;

 Can be more affected by inhibitors 

present in the environmental matrix

than culture methods; and

 Infectivity cannot be determineda.

ICC-PCR and ICC-RT-PCR  Improves detection of infectious 

viral pathogens compared to 

conventional cell culture; 

 Detects viruses that do not produce 

cytoplasmic effects in cell culture; 

and

 Provides results in half the time 

required for conventional cell 

culture.

 Less time-efficient and more costly 

than direct PCR detection; 

 Carryover detection of DNA of 

inactivated viruses inoculated onto 

cultured cells is possible; and

 Cannot be used for viruses that cannot 

be cultured.

Note: Table modified from Table 2 in Fong and Lipp (2005).

a Can determine infectivity if conducted in combination with ICC. See row on ICC-PCR and ICC-RT-PCR in table 

for more details.
31

4.1.1 Coliphage – Virus Associations in Freshwater

In studies that evaluated the association between the occurrence of coliphages and viruses in 

freshwater, results have varied. For example, Espinosa et al. (2009) found a strong association 

between F-specific coliphages and enterovirus (p-value = 0.0182), but a weak relationship with 

coliphages and rotavirus (p-value = 0.1502) and astrovirus (p-value = 0.4587) in high-altitude 

surface water. 

In a four-year study of surface source waters using 10 testing locations in the Netherlands, 

Lodder et al. (2010) found a significant association between densities of coliphages (F-specific 

and somatic) and enteroviruses, but not between coliphages and other viruses (NoV, rotavirus, 

and reovirus) or between the other viruses (NoV, rotavirus, and reovirus). NoV and rotavirus 

were detected in 45% and 48% of the samples, respectively. Infectious enterovirus and reovirus 

were detected in approximately 80% of the tested samples. Somatic and F-specific coliphages 

were detected in 100% and 97% of the samples, respectively. In the two samples where no Fspecific coliphages could be detected, enteroviruses were present, and in one sample and

rotavirus and NoV was also detected. Lodder et al. (2010) concluded that their results do not 

support a role for coliphages as indicators of source water quality, however, they also conclude 

that coliphages may be useful for determining treatment efficiencies. 

Payment and Locas (2011) used 20 years of sampling data from their laboratory to examine the 

association between pathogens and multiple microbial indicators, including coliphages, in 

sewage, surface water, and groundwater. Although the authors review data for several water 

types, coliphage associations with pathogens were investigated in groundwater. Their analysis of 

242 samples from 25 municipal groundwater well sites indicated that somatic and F-specific 

RNA coliphages were not predictive of virus presence or absence. This was due in part to the 

low numbers of coliphages present in the samples and their infrequent detection (Payment and 

Locas, 2011). 

Viau et al. (2011b) found no significant association between the presence of F-specific 

coliphages and adenovirus, enterovirus, NoV GI, and NoV GII in tropical coastal streams.

Additionally, Hot et al. (2003) found no significant association between the density of somatic 

coliphages and the presence of viral pathogens (RT-PCR detection of the genome of hepatitis A 

virus, NoV GI and GII, astrovirus, rotavirus, and infectious enteroviruses) in concentrated 

surface river water samples. In the 68 samples taken over 12 months, genomic detection of 

human pathogenic viruses was not statistically associated with the levels of somatic coliphages 

in surface water (Hot et al., 2003). For more information on the detection methods used, see 

Table 8 below. 

4.1.2 Coliphage-Virus Associations in Saline or Brackish Water 

The associations between coliphages and viruses in saline or brackish waters are also varied. 

Jiang et al. (2001) found that in urban runoff- impacted coastal waters, the presence of human 

adenovirus was significantly associated with the presence of F-specific coliphages (Jiang et al., 

2001). Mocé-Llivina et al. (2005) found that in seawater samples at public beaches, somatic 

coliphages were the best indicators of enteroviruses out of all of the indicators tested (F-specific 

coliphages, total coliforms, fecal coliforms, and enterococci) as they were found in higher 
32

numbers than other fecal indicators, including F-specific coliphages, and their amounts were 

indicative of enterovirus levels (Mocé-Llivina et al., 2005). Similarly, Ballester et al. (2005) 

found that in samples of marine water, both F-specific and somatic coliphages were significantly 

associated with adenoviruses, and F-specific coliphages were also significantly associated with 

rotavirus and enterovirus. Neither type of coliphage was significantly associated with the 

presence of astroviruses (Ballester et al., 2005). The amounts of coliphages and viruses varied by 

season. From seasonal and proximity data, it appeared that coliphages were more associated with 

viral presence than E. coli and that F-specific coliphages had the highest association with viral 

presence (Ballester et al., 2005). 

In contrast, in a study of the occurrence and distribution of FIB (total coliform, fecal coliform, 

and Enterococcus), F-specific coliphages, human adenovirus, and enterovirus in freshwater 

streams and an estuary, Jiang et al. (2007) found a strong association between the occurrence of 

FIB and F-specific coliphages, but no association between the presence of F-specific coliphages 

and human adenovirus or enterovirus. Jiang et al. (2007) found that the detection of human 

viruses depends on a seasonal and freshwater-to-saltwater distribution pattern that was the 

opposite of that of FIB and coliphages. For more information on the detection methods used, see 

Table 8 below. Similarly, Boehm et al. (2009) did not find an association between the presence 

of coliphages, including somatic and F-specific DNA and F-specific RNA coliphages, and 

human enterovirus or adenovirus in marine waters in Avalon Beach, California (Boehm et al., 

2009). 

A summary of the above papers, detection methods, and quantitative data (when available) are 

presented below in Table 8. A systematic literature review was not conducted, so the studies 

shown in Table 8 are only a subset of the studies that likely exist. 
33

Table 8. Summary table of coliphages – virus correlations in ambient water.

Study Water type

(Location)

Coliphages detected

Coliphage 

detection 

method

Viruses detected Virus detection 

method Occurrence findings

Baggi et al. 

(2001)

Fresh 

(Switzerland)

(upstream of 

WWTP)

Somatic coliphages 

(means: 1.9 and 3-

log10 PFU per mL)

F-specific coliphages 

(range of means: 

1.5–3-log10 PFU per 

mL)

ISO 10705-1 Enteroviruses, rotaviruses, and 

hepatitis A (41–44% of samples 

positive)

RT-PCR plus 

nested PCR

Coliphages associated with 

viruses. FIB not associated with 

viruses.

Jiang et al.

(2001)

Marine

Coastal waters

impacted by 

urban run-off 

(Southern CA)

Somatic coliphages

(5.3–3,332 PFU per 

L)

F-specific

(5.5–300 PFU per L)

EPA Method 

1601

Adenovirus

(880–7,500 genomes per L)

Nested PCR The presence of human 

adenovirus was significantly 

associated with F-specific 

coliphages.

Hot et al.

(2003)

Fresh river

(France)

Somatic coliphages

(range of densities: 

4×102–1.6×105 PFU 

per L)

ISO 10705-2 Cell culture: total culturable 

enteroviruses (later determined 

to be poliovirus type3)

Molecular methods: hepatitis A 

virus (1 positive /68 total), 

astrovirus (2/68), NoV GI (0 

detects), NoV GII (1/68),

rotavirus (0 detects), and 

enterovirus (2/68).

Cell culture and 

RT-PCR followed 

by Southern Blot

No significant association was 

observed between the density 

of somatic coliphages and the 

presence of infectious 

enteroviruses, or enterovirus 

genomes.

Skraber et al. 

(2004b)

Fresh river 

(France)

Somatic coliphages

(Mean: 3.06-log10

PFU/100 mL)

ISO 10705-2 Enterovirus spp. and NoV GII

(34 samples out of 90 (38%) 

were positive for enterovirus 

(13%) and/or NoV GII (27%) 

genome)

Enterovirus spp.:

cell culture, ICCRT-PCR, and RTPCR 

NoV GII: RT-PCR

The number of samples positive 

for pathogenic viral genome 

increased with increasing 

densities of somatic coliphages.

Ballester et al.

(2005)

Marine

Coastal water

impacted by 

WWTP

discharge

(Boston, MA)

Somatic and Fspecific coliphagesEPA Method 

1602

Human astrovirus, 

enteroviruses, rotavirus, and 

adenovirus types 40 and 41

ICC-nPCR, ICCRT-nPCRThe presence of enteric viruses 

and adenovirus was 

significantly associated with 

the presence of F-specific 

coliphages and somatic 

coliphages. Only F-specific 

coliphages were significantly 

associated with the presence of 

rotavirus and enterovirus.
34

Study Water type

(Location)

Coliphages detected

Coliphage 

detection 

method

Viruses detected Virus detection 

method Occurrence findings

Betancourt and 

Rose (2005)

Wetland and 

proposed 

sources for 

wetland 

restoration 

(Florida, USA)

F-specific coliphages 

(5 PFU per 100 mL 

reported for one 

wetland lake sample)

Agar overlay 

method and 

enrichment 

protocol 

developed by B. 

Yanko

Enteric viruses (detected in 

14/28 samples)

Cell culture Not discussed by the authors 

but low levels of occurrence in 

the sample set indicate 

association is unlikely.

Mocé-Llivina 

et al. (2005)

Marine

coastal water 

impacted by 

urban run-off

(Barcelona, 

Spain)

Somatic coliphages

(9–12,240 PFU per 

100 mL)

F-specific coliphages

(0–84 PFU per 100 

mL)

ISO 10705–1 

10705-2

Culturable enteroviruses (0–158 

PFU per 10 L)

Cell culture 

methods: standard 

plaque assay, 

double-layer plaque 

assay, VIRADEN 

method, RT-PCR,

and RT-nPCR

Receiver operating 

characteristic curves of 

“numbers of enteroviruses in 10 

L of seawater” indicated that 

the numbers of somatic 

coliphages (and enterococci) 

most accurately predicted the 

numbers of cultivable 

enteroviruses.

Westrell et al. 

(2006)

Fresh

river impacted 

by WWTP

(The 

Netherlands)

F-specific coliphages

In 2001:

Range: 6–7400 PFU 

per L

In 2002–2003:

Peak: 5,100

Median: 1,300 PFU 

per L

ISO 10705-1 NoV

In 2001: 

January peak: 240 PCR 

detectable units per L

In 2002–2003:

Peak: 2,000–3,000 

Mean: 12–1,700 PCR detectable 

units per L

RT-PCR Peaks in NoV did not coincide 

with those of enteroviruses, Fspecific coliphages, or 

turbidity.

Jiang et al. 

(2007)

Marine and fresh

coastal estuary

(Newport Bay, 

CA)

F-specific coliphages EPA Method 

1601

Adenovirus, enterovirus RT-PCR 

(enterovirus), 

nested PCR 

(adenovirus)

The seasonal and freshwater-tosaltwater distribution pattern of 

human viruses is the opposite 

of FIB and coliphages. 

Boehm et al. 

(2009)

Marine

sewage impacted 

beach

(Avalon, CA)

F-specific DNA and 

RNAcoliphages, 

somatic coliphages

Membrane 

filtration

Adenovirus, enterovirus RT-PCR 

(enterovirus), 

nested PCR 

(adenovirus)

No association between 

coliphages and adenovirus or 

enterovirus.

Espinosa et al. 

(2009)

Fresh 

high-altitude 

surface water

(Mexico City, 

Mexico)

Not specified (but

likely F-specific

coliphages)

Double layer 

culture (K12 Hfr 

host)

Enterovirus, rotavirus, 

astrovirus

RT-PCR Coliphages showed strong 

association with enterovirus, 

but weak association with other 

enteric viruses. 
35

Study Water type

(Location)

Coliphages detected

Coliphage 

detection 

method

Viruses detected Virus detection 

method Occurrence findings

Lodder et al.

(2010)

Fresh 

rivers

(The 

Netherlands)

Somatic coliphages

(1.1 to 114,156 PFU 

per L),

F-specific coliphages

(0.12 to 14,403 PFU 

per L)

ISO 10705–1 

ISO 10705–2

Enterovirus (present in 75% of 

samples (range, 0.0033 to 5.2 

PFU per L)

Reovirus (83% of samples 

(0.0030 to 5.9 PFU per L),

Cell culture using 

RT-PCR and ICCRT-PCR

A significant association was 

observed between the densities 

of the two coliphages and 

enteroviruses. 

Payment and 

Locas (2011), 

using data 

taken from 

Locas et al. 

(2007, 2008)

Fresh 

groundwater

(Canada)

Somatic and Fspecific RNA 

coliphages

EPA Methods 

1601 and 1602

Cell culture and 

immunoperoxidase: total 

culturable human enteric viruses 

Molecular methods: NoV, 

adenovirus types 40 and 41, 

enteroviruses, and reoviruses 

types 1, 2, and 3 

Cell culture,

immunoperoxidase, 

ICC-PCR, ICC-RTPCR, and RT-PCR

Somatic and F-specific RNA 

coliphages were not predictive 

of virus presence or absence. 

Coliphages were present only 

in low numbers and less 

frequently than bacterial 

indicators.

Viau et al. 

(2011b), using 

data presented 

in Viau et al.

(2011a)

Fresh,brackish 

and marine

tropical coastal 

streams and 

estuaries

(Hawaii)

F-specific coliphages

(present in 85/88 

samples, log10 mean 

1.2 ± 0.8 per 100 

mL)

Membrane 

filtration and 

double agar 

layer

Adenovirus (present in 13/88 

samples, 0.8 to 4.2 gene copies 

per 100 mL) 

Enterovirus (5/88 samples, 0.4 

to 4.8 gene copies per 100 mL) 

NoV GI (19/88 samples, 1.2 to 

1,441 gene copies per 100 mL) 

NoV GII (11/88 samples, 0.9 to 

62.4 gene copies per 100 mL) 

qPCR, 

RT-qPCR

There were no associations 

between occurrence of viruses 

and fecal indicator densities 

(including coliphages).

Love et al. 

(2014)

Marine 

recreational 

beaches 

F-specific coliphages

(median 

concentrations at 

both beaches 0.3 

MPN per 100 mL)

Somatic coliphages

(median 

concentrations were 

4.9 and 3.1 MPN per 

100 mL)

Modified 

version of 

modified version 

of EPA Method 

1601

Adenovirus (25.5% of water 

samples at Doheny Beach

and in 9.3% at Avalon Beach

NoV (22.3% of water samples at 

Doheny Beach and 0.7% at 

Avalon Beach

Adenovirus: nested 

PCR

NoV: nested RTPCR

The presence of F-specific

coliphages was positively 

associated with the probability

of detecting adenovirus. NoV

was not significantly associated 

with either type of coliphages.
36

Study Water type

(Location)

Coliphages detected

Coliphage 

detection 

method

Viruses detected Virus detection 

method Occurrence findings

Rezaeinejad et 

al. (2014)

Urbanized 

catchment 

waters

(freshwater) in 

tropical 

Singapore

F-specific coliphages

(mean concentration 

= 1.1 x 102 PFU per 

100 mL)

Somatic coliphages

(mean concentration 

= 2.2 x 102 PFU per 

100 mL)

EPA Method 

1602

Adenovirus (mean = 9.4 x 101 

gene copies/L)

Astrovirus (mean = 2.9 x 102 

gene copies/L)

NoV GII (mean = 3.7 x 102 gene 

copies/L)

Rotavirus (mean = 2.5 x 102 

gene copies/L)

Adenovirus: real 

time PCR

Astrovirus, 

rotavirus, NoV G I 

and GII: real time 

RT-PCR

F-specific coliphages were 

positively associated with NoV

densities.

VIRADEN method = “virus adsorption enumeration” based on the direct enumeration of viruses adsorbed into nitrate-acetate cellulose membranes.

Note: Bacterial hosts for somatic coliphages include: WG5, CN13, E. coli 036; bacterial hosts for F-specific coliphages include: Stm WG49, E. coli Famp, K12 

Hfr.
37

5. Environmental Factors and Fate

The ability of coliphages (including different coliphage genogroups) and other enteric viruses to 

survive in environmental media varies widely (Callahan et al., 1995; Reyes and Jiang, 2010; 

Jończyk et al., 2011; Bertrand et al., 2012). As described previously, the effect of environmental 

factors on coliphage survival is associated with morphology, where some specific structural 

characteristics, such as tails, large capsids, and lack of an envelope have been shown to be 

associated with greater resistance to external factors (Ackermann et al., 2004; Jończyk et al., 

2011). Researchers have investigated the survival of coliphages and enteric viruses under a 

variety of environmental conditions. Studies have examined the effects of physical stress (e.g., 

temperatures and sunlight), biological antagonists (e.g., microbial predation and enzymatic 

degradation), and chemical antagonists (e.g., disinfection). This section focuses on physical and 

biological antagonists in natural aquatic environments, mechanisms of inactivation, and where 

data are available, compares inactivation rates of somatic, F-specific and Bacteroides

bacteriophages to inactivation of human enteric viruses. Chemical treatment and other 

disinfection methods are discussed in Section 6 (Wastewater Treatment). 

5.1. Temperature 

Temperature is an important factor in viral ecology as it plays a fundamental role in attachment, 

penetration, multiplication, occurrence, and viability (Sobsey and Meschke, 2003; Pradeep Ram 

et al., 2005; Jończyk et al., 2011). Many studies have examined the effect of temperature on the 

survival of different viruses in aquatic environments. Both enteric viruses and coliphages have 

been reported to survive longer and occur more frequently at lower temperatures in natural 

environments and decay more rapidly at higher temperatures (i.e., seawater, river, and 

groundwater) (Long and Sobsey, 2004; Fong and Lipp, 2005). Below is a brief summary of the 

evidence of the effects of temperature on human enteric virus and coliphage inactivation in 

aquatic systems.

Bertrand et al. (2012) conducted a meta-analysis of the effects of temperature on the inactivation 

of enteric viruses and bacteriophages in food and water. The study collected 658 data points from 

76 published studies and analyzed the effects of virus type, matrix (simple or complex), and 

temperature (<50 and ≥50°C) on virus survival. A simple matrix included: (1) synthetic media; 

(2) drinking water; and (3) groundwater. A complex matrix included: (1) freshwater; (2) natural 

seawater; (4) sewage; (4) soil; (5) dairy products; (6) food; and (7) urine (Bertrand et al., 2012). 

The study determined that, overall, virus inactivation was faster at temperatures ≥50°C than at 

temperatures <50°C and that virus inactivation was less sensitive to temperature change in 

complex matrices than in simple matrices (Bertrand et al., 2012). The somatic coliphage ΦX174 

was highly persistent under all temperatures and matrices tested. 

Studies reported differences in survival among different F-specific coliphage groups across 

temperature gradients. For example, Long and Sobsey (2004) reported that at 4°C, GI and GII 

F-specific RNA coliphages were detectable for over 100 days, GIII F-specific RNA coliphages 

were detectable for 3 weeks, and GIV F-specific RNA coliphages were reduced to the limit of 

detection after 10 days (Long and Sobsey, 2004). Of the F-specific DNA coliphages, all strains 

were detectable after 110 days at 4°C (Long and Sobsey, 2004). The authors also noted that the 

GI F-specific RNA coliphage MS2 and F-specific DNA coliphage M13 demonstrated a longer 
38

survival in environmental waters than other F-specific coliphage species (Long and Sobsey, 

2004). 

Temperature can also affect survival of somatic and Bacteroides bacteriophages in aquatic 

systems. Lee and Sobsey (2011) estimated the temperature inactivation of four types of somatic 

coliphages in laboratory tests using both reagent grade water and surface water. The authors 

found that T4 (Myoviridae family), ΦX174 (Microviridae family), and λ (Siphoviridae family), 

survived better than T1 (Siphoviridae family), and T7 (Podoviridae family), at low temperatures 

(4°C) and high temperatures (25°C). Chung and Sobsey (1993) found that B. fragilis coliphages 

survived comparable to or better than hepatitis A, poliovirus, and rotavirus (measured using cell 

culture) in seawater exposed to low (5°C) and high (25°C) temperatures. 

Reported comparisons between decay rates of F-specific RNA coliphages and human enteric 

viruses, or proxies to human enteric viruses, indicate that decay rates of both vary by temperature 

and water conditions. For example, in their two studies, Allwood et al. (2003, 2005) compared 

the survival of GI F-specific RNA coliphage MS2, feline calicivirus (FCV), and E. coli at 4°C, 

25°C, and 37°C in chlorinated and dechlorinated water. In dechlorinated water at 4°C and 25°C, 

MS2 survived three times longer than both E. coli and FCV, whereas they had similar survival 

rates at 37°C (Allwood et al., 2003). 

Similarly, Romero et al. (2011) found that porcine rotavirus and GI F-specific RNA coliphage 

MS2 had relatively low inactivation rate constants in the dark from 14 to 42°C, 10-fold increases 

in inactivation rates at 50°C and between 10- and 60-fold increases in inactivation rates at 60°C. 

In a similar experiment, Seo et al. (2012) compared the decay rates of murine NoV (MNV) and 

GI F-specific RNA coliphage MS2 over a temperature range of 24 to 85°C. They found that 

decay rate of MS2 was lower than MNV between 24°C and 60°C and that both were rapidly 

inactivated by temperatures >60°C (Seo et al., 2012). For more details on the decay rates at 

different temperatures, see Table 9 below. 

Synergistic effects between temperature and other environmental factors

The importance of temperature as a determinant of coliphage survival has been found to vary 

between freshwater and saltwater environments. For example, Reyes and Jiang (2010) noted that 

temperature is more important in influencing coliphage occurrence in freshwater environments 

than in saltwater environments (See Section 5.3 for more information on salinity). The 

importance of temperature as a determinant of virus survival is also dependent on the presence of 

sunlight. Romero et al. (2011) found that temperature played an important role in sunlightmediated inactivation. For example, degradation rates of both GI F-specific coliphage MS2 and 

porcine rotavirus were higher for the same temperatures under different light conditions (full 

solar spectrum and only UVA and visible light) as compared to in the dark (Romero et al., 2011) 

(See Section 5.2 for more information on sunlight). Hurst et al. (1989) showed that temperature 

effects on inactivation of enterovirus was dependent on the water sources used as the aqueous 

phase in experiments. 

Summary 
39

In summary, conclusions drawn in multiple studies indicate that the coliphages are equally 

persistent to, or more persistent than enteric viruses. Bertrand et al. (2012) found that somatic 

coliphage ΦX174 was highly persistent under all matrices and temperatures tested, and at higher 

temperatures, somatic and F-specific coliphages were classified as the most persistent as 

compared to enteric viruses. These data are consistent with the results of Allwood et al. (2003, 

2005) and Seo et al. (2012). Combined, these data indicate that coliphages may be conservative 

surrogates for the behavior of enteric viruses under a range of temperatures (meaning they 

persist as long or longer than human viruses). Table 9 presents the decay rates of different types 

of coliphages, other fecal indicators, and human viruses. 

5.2. Sunlight 

Sunlight is also an important factor leading to virus inactivation (Sobsey and Meschke, 2003; 

Fong and Lipp, 2005; Jończyk et al., 2011). Sunlight that reaches Earth’s surface is composed of 

medium and long wavelength UV light [UVB (280 to 320 nm); UVA (320 to 400 nm)], visible 

light (400 to 700 nm), and longer wavelengths (Love et al., 2010). There are three proposed 

types of virus inactivation caused by the UV wavelengths in light: endogenous direct, 

endogenous indirect, and exogenous indirect (Silverman et al., 2013). While UV radiation is 

utilized in wastewater treatment processes, this application uses primarily UVC wavelengths 

(which do not reach Earth’s surface due to the ozone layer) and will be discussed in Section 6 on 

wastewater treatment. This section will focus on inactivation of viruses due to natural or 

simulated sunlight. Below is a brief summary of the evidence of the effects of sunlight on human 

enteric viruses and coliphage inactivation in aquatic systems. 
40

Table 9. Comparison of mean exponential decay rates of coliphages, fecal indicators and human viruses in different media at 

different temperatures. 

Study Allwood et al.

(2003)

Allwood et al. 

(2005)

Nasser (1993)

Long and Sobsey 

(2004)

Lee and Sobsey 

(2011)

Seo et al. (2012)

Temperature 4°C 25°C 37°C 4°C 25°C 37°C 10°C 20°C 30°C 4°C 20°C 4°C 25°C 24°C 37°C

Organism Mean Exponential Decay Rate, k (d-1)

a

E. coli 0.30 0.40 0.77 0.23 0.20 0.14 0.03 0.18 0.20

FCV 0.32 0.44 1.15 0.15 0.58

GI F-specific RNA 

coliphage MS2 0.09 0.12 0.85 0.32 0.35 0.37 0.02 0.16 0.05 0.20

F-specific coliphages N/O 0.08 N/O

Human adenovirus 0.12 0.15 0.18

Poliovirus type 1 0.01 0.14 0.21

F-specific DNA 

coliphages

0.02–

0.23

0.13–

1.44

GI F-specific RNA 

coliphages

0.02–

0.03

0.16–

0.20

GII F-specific RNA 

coliphages 0.09 0.38

GIII F-specific RNA 

coliphages 0.55 1.59

GIV F-specific RNA 

coliphages 0.55 2.71

GV F-specific RNA 

coliphages 0.63 2.88

Somatic coliphage T1 0.10 1.15

Somatic coliphage T4 <0.01 0.07

Somatic coliphage T7 0.10 1.15

Somatic coliphage 

ΦX174 0.02 0.15

Somatic coliphage λ 0.01 0.12

MNV 0.82 2.43

Study Medium Dechlorinated tap 

water

Dechlorinated tap 

water

Raw wastewater, 30-

day incubation Lake water Lake water

Dulbecco’s 

Modified

Eagle’s Medium

N/0 = die off was not observed, empty cells = not reported; aThe mean exponential decay rate k may be used in the exponential decay equation: Nt = N0e

-

kt

.Alternatively, k may be used in the base 10 exponential decay equation as Nt = N010-kt/

ln(10)

.
41

Effects of sunlight on coliphage decay rates and decay rates of other fecal indicators 

Love et al. (2010) found a correlation between the size of the genome and the inactivation rate of 

environmental isolates of somatic coliphages in sunlight: Larger genomes were correlated with 

higher inactivation rates. They also found that F-specific RNA coliphages were significantly 

more resistant to sunlight inactivation than the F-specific DNA coliphages over an 8-hour period 

(Love et al., 2010). Overall, they found that under full-spectrum-simulated sunlight, inactivation 

rates varied more widely for ssDNA and dsDNA viruses than for ssRNA viruses, and that 

differences in virus inactivation rate were not just a function of nucleic acid type, but also 

genome length and morphology (Love et al., 2010). 

Sinton et al. (1999) studied the inactivation rates of sewage-isolated somatic coliphages, 

F-specific coliphages, B. fragilis bacteriophages, and fecal coliforms by solar radiation in 

sewage-seawater mixtures. Overall, their data showed that sunlight conditions resulted in faster 

decay rates of all indicators as compared to dark conditions and that, under all conditions,

somatic and F-specific coliphages had lower decay rates than B. fragilis bacteriophages and fecal 

coliforms (Sinton et al., 1999). The authors also found that colder water resulted in slower decay 

rates than warmer water under all light and dark conditions tested (Sinton et al., 1999). 

In their follow-up study, Sinton et al. (2002) investigated the inactivation rates of waste 

stabilization pond effluent isolated fecal coliforms, enterococci, E. coli, somatic coliphages, and 

F-specific RNA coliphages by solar radiation in freshwater (Table 10 below). Overall, their data 

showed that, for all indicators, sunlight conditions resulted in faster decay rates than dark 

conditions and that under both light and dark conditions, somatic and F-specific RNA coliphages 

had smaller decay rates than E. coli, enterococci, and fecal coliforms (Sinton et al., 2002). Sinton 

et al. (2002) also found that F-specific RNA coliphages were inactivated by a wide range of 

wavelengths, whereas somatic coliphages were mainly inactivated by UVB wavelengths (318 

nm). 
42

Table 10. Mean exponential decay rates of coliphages and fecal indicators in fresh river 

water contaminated with raw sewage or effluent under different light conditions. 

Organism Source of contamination

Dark Summer Winter Summer Winter

kD(/hour)

a ks(m2

/MJ)b kL(/hour)

a

Fecal coliforms

Wastewater effluent 0.02 0.09 0.08 0.26 0.14

Raw sewage 0.01 0.28 0.22 0.70 0.30

E. coli

Wastewater effluent 0.02 0.08 0.07 0.25 0.11

Raw sewage 0.02 0.29 0.24 0.70 0.33

Enterococci

Wastewater effluent 0.02 0.28 0.11 0.77 0.16

Raw sewage 0.01 0.14 0.14 0.36 0.18

Somatic coliphages

Wastewater effluent 0.01 0.08 0.05 0.20 0.08

Raw sewage 0.00 0.10 0.09 0.28 0.14

F-specific RNA 

coliphages

Wastewater effluent 0.01 0.07 0.05 0.17 0.08

Raw sewage 0.00 0.08 0.07 0.18 0.12

Source: Sinton et al. (2002)

a The mean exponential decay rate, kD and kL, may be used in the exponential decay equation: Nt = N0e-kt

.

Alternatively, kD (decay in the dark) and kL (decay in the light) may be used in the base 10 exponential decay 

equation as Nt = N010-kt/ln(10). Note that kL may be used only in equivalent solar insolation conditions as the study.

b The mean solar inactivation rate ks may be used in the exponential decay equation: Nt = N0es-ksIt, where I is the 

solar irradiance.

Effects of sunlight on decay rates of enteric viruses and coliphages 

Individual enteric viruses and coliphages also have different levels of resistance to sunlight. For 

example, Love et al. (2010) observed that in seawater under sunlit conditions, the decay rates of 

adenovirus 2 and GI, GII, GIII and GIV F-specific RNA coliphages were similar and slower than 

the decay rates of F-specific DNA coliphages, somatic coliphages, and poliovirus type 3 (Love et 

al., 2010). These results are consistent with field experiments under conditions of similar 

sunlight intensity (Love, et al., 2010). 

Romero et al. (2011) used both full spectrum sunlight and a combination of UVA and visible 

light to determine the decay rates of GI F-specific RNA coliphage MS2 and porcine rotavirus at 

temperatures ranging from 14 to 50°C (see Table 11 below). Under dark conditions, decay rates 

were not detected for either virus between 14 and 42°C whereas at 50°C, low decay rates were 

detected for both (Romero et al., 2011). Under full spectrum sunlight, the decay rates (Kobs) of 

both viruses increased and those for GI F-specific RNA coliphage MS2 were below those of 

porcine rotavirus (Romero et al., 2011). Under a combination of UVA and visible light, both 

viruses had low, approximately constant degradation rates between 14 and 42°C, whereas at 

50°C the rates increased slightly (Romero et al., 2011). The very low levels of degradation of 

both MS2 and porcine rotavirus in the absence of UVB were consistent with previous studies 

indicating that the majority of sunlight degradation of viruses in water is due to UVB light 

(Sinton et al., 2002; Romero et al., 2011). These results are consistent with the findings of Fisher 

et al. (2011) who found that in phosphate buffered saline, GI F-specific RNA coliphage MS2 

was resistant to UVA but highly sensitive to UVB wavelengths. 
43

Silverman et al. (2013) compared the inactivation rates of poliovirus type 3, adenovirus type 2,

and GI F-specific RNA coliphages (MS2 and PRD1) under dark and full simulated sunlight 

conditions in four different types of environmental water (seawater from two marine beaches, 

river estuary water, coastal wetland, and coastal wetland collected near cattail plants) and in 

phosphate-buffered saline (see Table 11). They found that all dark control inactivation rates were 

less than those obtained from experiments conducted under full-spectrum simulated sunlight for 

all three viruses in all five types of water (Silverman et al., 2013). Additionally, they found that 

decay rates of GI F-specific RNA coliphages under full-spectrum simulated sunlight were 

significantly below those of poliovirus type 3 in all five types of water and less than or equal to 

those of adenovirus type 2. The authors conclude that GI F-specific RNA coliphages are a 

conservative surrogate for predicting poliovirus type 3 and adenovirus type 2 decay in all five 

types of water tested (Silverman et al., 2013). 

Table 11. Comparison of mean exponential decay rates of coliphages and human viruses 

under different light conditions. 

Medium/

Conditions

Romero et al. (2011)

Silverman 

et al. (2013)

b

20 mg C/L of riverine natural organic material (full-spectrum sunlight)

a

Fullspectrum 

sunlight

Microorganism 14°C 23–26°C 34°C 42°C 50°C Temperature 

not provided

kD(/h) kL(/h) kD(/h) kL(/h) kD(/h) kL(/h) kD(/h) kL(/h) kD(/h) kL(/h) kD(/h) kL(/h)

MS2 ND 4.00 ND 4.23 ND 4.49 ND 5.00 0.50 NS 0.10

Rotavirus ND 7.31 ND 8.58 ND 8.63 ND 9.42 0.33

Poliovirus (Type 3) NS 0.08

Adenovirus (Type2) NS 0.11

ND = Nondetect

NS = Not significantly different from zero

empty cells = not reported

mg = milligrams

a The mean exponential decay rate kD and kL may be used in the exponential decay equation: Nt = N0e-kt

.

Alternatively, kD (decay in the dark) and kL (decay in the light) may be used in the base 10 exponential decay 

equation as Nt = N010-kt/ln(10). Note that kL may be used only in equivalent solar insolation conditions as the study.

b The decay rates reported in Silverman et al. (2013) are from water collected from Tijuana River estuary (Imperial 

Beach, California) at the end of the ebb tide.

Synergistic effects between sunlight and other environmental factors 

Several studies have found synergy between sunlight and other environmental factors in the 

inactivation rates of viruses, such as the presence of organic matter or particulate matter, 

sunlight, and salinity. For example, inactivation of viruses may be greater in waters with organic 

matter that produces reactive oxygen species (Kohn et al., 2007; Love et al., 2010; Romero et al., 

2011). However, the presence of flora, fauna, and dissolved and particulate matter may also 

increase viral survival by blocking or absorbing photons from passing through water (Bitton et 

al., 1979; Romero et al., 2011). Please refer to Sections 5.4 and 5.5 for more information on 

microbial activity and organics, respectively. The synergy between sunlight and temperature 

appears to play a role in the inactivation of viruses. For example, Romero et al. (2011) concluded 

that temperature is a critical factor in the sunlight-mediated inactivation of GI F-specific 
44

coliphage MS2. Please refer to Section 5.1 for more information on temperature. Differential 

inactivation of coliphages by sunlight can also occur in saltwater versus freshwater. For example, 

Sinton et al. (1999, 2002) found that salinity had a synergistic effect with sunlight. Specifically,

sunlight inactivation increased with increasing salinity. For more information on salinity please 

see Section 5.3. 

Summary 

In summary, data indicated that human enteric viruses and coliphages have faster decay rates 

under conditions of full sunlight as compared to in the dark (Sinton et al., 1999, 2002; Romero et 

al., 2011). Reported decay rates varied by virus, amount and wavelengths of light (UVA, UVB),

temperature, and aquatic conditions (salt or freshwater), however, several studies indicated that 

coliphage decay rate is generally lower than enteric virus or FIB decay rate in various sunlight 

conditions (Sinton et al., 2002; Love et al., 2010; Romero et al., 2011; Silverman et al., 2013). 

Thus, coliphages may be a conservative surrogate for predicting virus decay due to sunlight.

5.3. Salinity 

The types and concentrations of salts found in natural waters differ depending on the type of 

water. Generally, seawater is considered to be 35 parts per thousand salt. Chloride (Na) and 

sodium (Cl) are the most prevalent ions and account for more than 85% of the salt content by 

mass (Murray, 2004). Concentrations of these ions (Na and Cl) are significantly lower in 

freshwaters, and vary depending on type and source of water (Murray, 2004). 

Salts, or salinity, can influence viral survival in aquatic environments. Salinity can either 

increase or decrease degradation rates of viruses depending on the type and concentration of salt, 

the temperature, and the specific virus (Hurst and Gerba, 1980; Gutierrez et al., 2010; Mylon et 

al., 2010; da Silva et al., 2011; Nguyen et al., 2011; Seo et al., 2012). It has been hypothesized 

that monovalent salts provide strong steric and electrosteric stabilization of GI F-specific 

coliphage MS2, whereas divalent salts have been found to cause MS2 aggregation (Mylon et al., 

2010; Nguyen et al., 2011). Similar results have been shown for rotavirus and NoV G1.1 

(Gutierrez et al., 2010; da Silva et al., 2011). Aggregation of viruses can make it difficult to 

measure their infectivity, as plaque assays result in underestimates (e.g., a single PFU may be 

comprised of clumps of virus particles). Additionally, osmotic shock through rapid changes in 

osmotic pressure can trigger inactivation of coliphages via direct oxidization, which can cause 

capsid degradation and dispersion, tail fragmentation, and release of viral nucleic acids into the 

aquatic environment (Jończyk et al., 2011). This section will describe the effects of salinity on 

viral degradation. Below is a brief summary of the evidence of the effects of salinity on human 

enteric viruses and coliphage inactivation in aquatic environments. 

Effects of salinity on decay rates of coliphages 

Sinton et al. (1999, 2002) found that salt water affected the decay rates of F-specific and somatic 

coliphages under both dark and sunlight exposed conditions. Sinton et al. (1999) studied the 

inactivation rates of sewage-isolated somatic coliphages and F-specific DNA and RNA 

coliphages in sewage-seawater mixtures. Sinton et al. (2002) studied the inactivation rates of 

somatic and F-specific RNA coliphages isolated from waste-stabilization pond effluent under 
45

both dark and sunlight exposed conditions in river water, simulated estuarine water (50% river 

water, 50% seawater), and seawater. Under dark conditions, both somatic and F-specific 

coliphages had lower decay rates in river water (somatic coliphages (kD = 0.008 h-1), F-specific 

RNA coliphages (kD = 0.014 h-1)) than in sea water (somatic coliphages (kD = 0.044 h-1), Fspecific RNA coliphages (kD = 0.044 h-1) (Sinton et al., 1999, 2002). Degradation rates of 

somatic coliphages increased 5.5 fold in salt water compared to river water under dark conditions 

whereas F-specific RNA coliphages rates increased 3.1 fold under the same conditions. These 

data indicate that somatic coliphages are less stable in seawater than F-specific RNA coliphages 

under the tested conditions. 

Somatic coliphages were more sensitive to salt water under sunlight conditions as well. For 

example, Sinton et al. (2002) determined the degradation rates of somatic coliphages and Fspecific RNA coliphages isolated from waste-stabilization pond effluent under full sunlight 

conditions in freshwater and 50:50 water and seawater. For somatic coliphages degradation rates 

were kS = 0.079 m2 megajoules (MJ)

-1 in river water, kS = 0.129 m2 MJ-1 in 50:50 water and kS = 

0.184 m2 MJ-1 in sea water. Similarly, F-specific RNA coliphages rates were: kS = 0.086 m2 MJ-1

in river water, kS = 0.092 m2 MJ-1 in 50:50 water and kS = 0.123 m2 MJ-1 in sea water (Sinton et 

al., 2002). Degradation rates of somatic coliphages increased 2.3 fold in salt water compared to 

river water whereas F-specific RNA coliphages rates increased 1.4 fold under the same 

conditions. These data indicate that somatic coliphages are more sensitive to salt water than Fspecific RNA coliphages under these conditions. 

Overall, the authors concluded that as salinity increases, inactivation of coliphages increases as 

well (Sinton et al., 2002). In particular, inactivation of F-specific RNA coliphages obtained from 

sewage increased with salinity, but the trend in F-specific RNA coliphages obtained from 

stabilization ponds was less pronounced (Sinton et al., 2002). These conclusions are in 

agreement with those of Savichtcheva and Okabe (2006) who found that F-specific RNA 

coliphages were more sensitive to sunlight inactivation at high salinity. 

Seo et al. (2012) investigated the differences in tolerance of MNV and GI F-specific RNA 

coliphage MS2 to different concentrations of NaCl (0.3, 1.3, 3.3, and 6.3% NaCl) at three 

different temperatures, 24°C, 37°C, and 50°C. Their results show that there are complex 

interactions between salt concentration and temperature for both of the viruses, with several 

differences between the two. They found that MS2 was more resistant to NaCl than MNV at all 

concentrations of NaCl and temperatures tested (Seo et al., 2012). At 24°C, MS2 did not show 

any reduction in infectivity at any of the NaCl concentrations and at higher temperatures, NaCl 

seem to have a protective effect (Table 12; Seo et al., 2012). 

Hurst and Gerba (1980) compared the decay of poliovirus, echovirus, coxsackievirus and simian 

rotavirus in estuarine and freshwater during two different years. Decay was quicker in estuarine 

water relative to freshwater in one year, and decay was similar in the waters in the second years 

suggesting factors other than salinity may have been contributing to viral decay. 
46

Synergistic effects between salinity and other environmental factors 

Several studies have found synergy between salinity and other environmental factors in the 

inactivation rates of viruses. For example, Seo et al. (2012) found an interaction between 

temperature and salt concentration. Depending on the specific virus, incubation in high 

concentrations of NaCl at high temperatures could either reduce virus infectivity (MNV) or 

increase virus infectivity (GI F-specific RNA coliphage MS2) as compared to lower 

concentrations of salt at the same temperature. The susceptibility of MNV to all concentrations 

of NaCl increased rapidly at 37°C and 50°C, whereas at the same temperatures, GI F-specific 

RNA coliphage MS2 was more stable at higher NaCl concentrations (1.3 to 6.3% NaCl) than at 

low concentrations (0.3% NaCl) (Seo et al., 2012). The authors hypothesized that the high NaCl 

concentration may “protect against thermally induced capsid opening or stabilize the viral 

protein-RNA complex” (Seo et al., 2012). For more information on the effects of temperature on 

virus degradation, see Section 5.1. Other studies have reported synergistic effects between salt 

and natural organic and inorganic matter (Mylon et al., 2010). Mylon et al. (2010) found that GI 

F-specific RNA coliphage MS2 aggregated at lower concentrations of Ca2+ in the presence of 10 

mg/L Suwannee River organic matter (100 millimolar (mM) Ca2+) as compared to just Ca2+ (160 

mM Ca2+). Lukasik et al. (2000) observed that mono-, di-, and trivalent salts (NaCl, MgCl2, and 

AlCl3) either promoted or interfered with adsorption of GI F-specific RNA coliphage MS2, 

somatic coliphage ΦX174, and poliovirus type 1 to different types of filters at different pH 

levels. For more information on adsorption to organic and inorganic matter, please see Section 

5.5 below. 

Summary

In summary, both enteric viruses and coliphages are affected by salinity, the specific effects of 

which vary depending on the type of virus and the type and concentration of salt, as well as 

temperature. In terms of aggregation, multiple studies have found that monovalent cations are 

either ineffective at, or are less effective at causing aggregation of coliphages (F-specific RNA 

coliphage MS2) and enteric viruses (NoV GI.1 and rotavirus) than divalent cations (Gutierrez et 

al., 2010; Mylon et al., 2010; da Silva et al., 2011; Nguyen et al., 2011) and aggregation can 

affect the number of PFUs measured in a sample. In terms of decay rates, Seo et al. (2012) found 

that MS2 had lower decay rates than MNV at all NaCl concentrations tested (0.3 to 6.3%) at 

three different temperatures (24°C, 37°C, and 50°C). Table 12 below shows the decay rates from 

Seo et al. (2012). 
47

Table 12. Comparison of mean exponential decay rates of coliphages and MNV at different 

concentrations of salt and at different temperatures. 

°C NaCl%

Organism

MNV GI F-specific coliphage MS2

k(d-1)

24

0.3 1.17 0.05

1.3 3.03 0.05

3.3 2.96 0.05

6.3 2.73 0.05

37

0.3 2.54 0.19

1.3 4.44 0.18

3.3 4.11 0.21

6.3 4.11 0.11

50

0.3 61.40 10.80

1.3 72.08 3.15

3.3 122.80 5.54

6.3 118.42 4.09

Source: Seo et al. (2012)

Higher k = faster decay

5.4. Predation and Enzymatic Degradation 

Inactivation of viruses can occur via predation or release of virucidal agents from endogenous 

microorganisms in environmental waters (Sobsey and Cooper, 1973; Fujioka et al., 1980; Ward 

et al., 1986). Many bacteria produce proteolytic enzymes that are capable of inactivating viruses, 

including human enteric viruses, by degradation of protein capsids (Bae and Schwab, 2008). In 

seawater, virioplankton are postulated to be inactivated in part by enzymatic attack and predation 

(Finiguerra et al., 2011). One study found that the presence/absence of microorganisms is a more 

important factor than temperature on virus survival in groundwater (Wetz et al., 2004). Other 

studies have shown that association with biofilms can also affect the inactivation of enteric 

viruses and coliphages. Helmi et al. (2011) found that poliovirus, GI F-specific RNA coliphage 

MS2, and somatic coliphage ΦX174 densities in drinking water biofilms decreased after 6 days 

due to inactivation and detachment, but previous research has found that biofilms protect viruses 

from inactivation (Skraber et al., 2007). While the effects of microbial antagonism and 

enzymatic degradation on coliphages are not as well studied as the effects on human enteric 

viruses, these processes are thought to inactivate them as well. For example, studies examining 

coliphages in waste stabilization ponds have shown that while sunlight is the major cause of 

inactivation, predation may also play a role (da Silva et al., 2008). Below is a brief summary of 

the evidence of human enteric virus and coliphage predation- and enzymatic degradationmediated inactivation in aquatic systems. 

Effects of predation and enzymatic degradation on decay rates of coliphages 

In a study examining the role of aquatic plants in freshwater and salt water wetlands on the 

survival of waterborne coliphages, Karim et al. (2008) found that the presence of wetland 

vegetation significantly increased the inactivation of GI F-specific RNA coliphage MS2. The 

authors hypothesized that the presence of aquatic plants may enhance rhizosphere bacterial 
48

populations, which increase coliphage inactivation due to the presence of metabolites or the 

presence of proteolytic substances released by microbes or plants (Karim et al., 2008). 

Finiguerra et al. (2011) investigated the light-independent mechanisms of inactivation of somatic 

coliphage T4 (marine host: PWH3a) and coliphage P1 (enteric host: E. coli B) in seawater. They 

found that decay rates of both coliphages were reduced in particle-free seawater (<2 micrometers 

[μm]) as compared to seawater containing nanoplankton (<10 μm) and the lowest decay rates 

were found in ultra-filtered seawater (<10 kilodaltons [kDa]). The authors concluded that 

inactivation of these coliphages is accelerated by naturally occurring particles, which include 

living organisms and heat-labile colloids and macromolecules >10 kDa (Finiguerra et al., 2011). 

Effects of predation and enzymatic degradation on decay rates of human viruses 

A number of studies have examined the effect of microbial activity on enteric virus survival in 

aquatic systems. Direct predation of enteric viruses can occur via engulfment or ingestion by 

bacteria, protozoa, helminthes, and other aquatic organisms (Sobsey and Meschke, 2003). 

Fujioka et al. (1980) demonstrated that inactivation of enteric viruses (poliovirus type 1, 

coxsackievirus B4, and echovirus 7) in marine and estuarine waters is associated with the natural 

microbial community. Microbial activity has also been shown to decrease persistence of 

rotaviruses in raw and treated freshwaters (Raphael et al., 1985) and hepatitis A in mixed septic 

tank effluent (Deng and Cliver, 1995). Toranzo et al. (1982) confirmed the ability of bacteria to 

release virucidal agents by isolating marine bacteria that had marked activity against poliovirus 

(net 2-log10 inactivation or greater within 6 to 8 days), coxsackievirus B-5, and echovirus 6. 

Sobsey and Cooper (1973) showed that microbial activity in waste stabilization pond water 

contributed to poliovirus inactivation. Similarly, Herrmann et al. (1974) showed that 

enteroviruses decayed more quickly in lake water compared to sterilized lake water. Ward et al. 

(1986) also showed that proteolytic bacterial enzymes were responsible for echovirus 

inactivation in freshwater. 

Wetz et al. (2004) studied the inactivation rate of poliovirus in filtered natural seawater, 

unfiltered natural seawater, artificial seawater, and deionized water at 22 and 30°C. They found 

that the highest rates of virus inactivation occurred in unfiltered natural seawater at both 

temperatures tested. Prior to spiking they exposed all of the water in their experiments to >14 

hours of UV light (to kill the indigenous microorganisms). Because the indigenous 

microorganisms were killed, the authors hypothesized that direct microbial inactivation of the 

viruses was highly unlikely and degradation was likely caused by a release of cellular proteases, 

nucleases, and other enzymes (Wetz et al., 2004). 

Synergistic effects between predation and enzymatic degradation and other environmental 

factors 

Several studies have identified synergy between predation and enzymatic degradation and other 

environmental factors in the inactivation rates of viruses. There is some evidence that when 

viruses, including enteric viruses and coliphages, adsorb to particles, the associated particle may 

offer them some protection from predation (Fong and Lipp, 2005; Weaver and Sinton, 2009; 

Finiguerra et al., 2011). For more information on adsorption to organic and inorganic matter, see 
49

Section 5.5 below. There are also synergistic effects between enzymatic degradation and 

temperature. For example, Wetz et al. (2004) found a synergistic effect between temperature and 

enzymatic degradation of poliovirus in natural seawater, as degradation rates were higher at 30°C 

than at 22°C. For more information on effects of temperature, see Section 5.1 above. 

Summary

Microbial predation and enzymatic degradation are both important mechanisms of virus 

inactivation in natural waters. Both predation and enzymatic degradation have been shown to 

increase human virus degradation rates in freshwater, salt water, treated water and septic system 

effluent (Fujioka et al., 1980; Toranzo et al., 1982; Raphael et al., 1985; Deng and Cliver, 1995; 

Wetz et al., 2004). While there are fewer data for coliphages, there is some evidence that 

microbial predation and enzymatic degradation do contribute to virus inactivation in natural 

waters. Due to lack of data, it is not currently possible to compare degradation rates of enteric 

viruses and coliphages by microbial predation or enzymatic degradation in natural waters. 

5.5. Organic and Inorganic Matter 

Aquatic environments contain both organic and inorganic matter. Inorganic matter consists of 

materials made from nonbiological sources and do not contain carbon (except for CO2 and CH4).

These include metals, chemicals, sand, clay, salts, and ions. Natural organic matter consists of 

materials that are made from biological sources and contain carbon. These include exudates from 

organisms and the materials that are produced from their decay. Organic matter in water is a 

diverse mixture of organic compounds ranging from macromolecules to low molecular-weight 

compounds (USGS, 2013). Organic matter is capable of both attenuating light (thus decreasing 

photoactivation rates) and producing reactive oxygen species (thus increasing photoactivation 

rates) (Silverman et al., 2013). Depending on the absolute amount of sunlight that reaches the 

virus and the amount of reactive oxygen species produced, the overall effect of organic matter 

can either result in decreased or increased viral photoinactivation rates. 

Viruses in the environment are often associated with particulate matter, which has a major effect 

on persistence and transport in the environment (Gerba, 1984). For example, clay surface 

exchange capacity and particle size and shape affect the virus-adsorption activity of a clay 

(Carlson et al., 1968). Laboratory-based predictions suggest that as many as 99% of viruses in 

coastal waters should be adsorbed to naturally occurring colloids and particles (Finiguerra et al., 

2011). If the resultant aggregate is dense and large, it can settle out of the water column 

(Characklis et al., 2005; Shen et al., 2008). If the aggregate is less dense, viruses may remain 

more mobile in the environment (Characklis et al., 2005). 

The isoelectric point of the virus dictates its overall charge at a given pH, ionic strength, and 

water chemistry and thus affects virus adsorption. For example, reoviruses adsorb primarily to 

negatively charged sites on clay, while T1 and T7 coliphages adsorb to positively charged sites at 

environmentally relevant pHs (Gerba, 1984). Stotsky et al. (1980) found that adsorption to clay 

by reovirus (the family to which rotavirus belongs) and somatic coliphages (T1 and T7) 

increased the persistence of the viruses in lake water (Stotsky et al., 1980, as cited in Sobsey and 

Meschke, 2003). 
50

This section will focus on inactivation of viruses due to interactions with organic and inorganic 

matter. Below is a brief summary of the evidence of the effects of organic and inorganic matter 

on human enteric viruses and coliphage inactivation in aquatic systems.

Effect of organic and inorganic matter on decay rates of coliphages 

Finiguerra et al. (2011) investigated the effects of particulate, dissolved, and colloidal organic 

and inorganic material in seawater on the inactivation rate of somatic coliphage T4. They 

determined that a significant fraction of viral inactivation (39–65%) can be attributed to passive 

sorption to living and inert planktonic particles (sterile debris was produced from cultivated 

phytoplankton species; 0.2 to 10 μm) (Finiguerra et al., 2011). The lowest decay rates were in 

oxidized filtrate from a 10 kDa tangential filtration system. The authors identified virucidal 

material between 10 kDa and 0.2 μm in size that is resistant to autoclaving. They concluded that 

inorganic solutes may be the primary inactivating mechanism in the dissolved fraction 

(Finiguerra et al., 2011). 

Effects of organic and inorganic matter on decay rates of enteric viruses and coliphages 

LaBelle and Gerba (1980) found that adsorption to marine sediment increased the time required 

for 99% inactivation from 1 hour to greater than 4 days for poliovirus and from 1.4 days to 

greater than 6 days for echovirus. Another study found that enteroviruses associated with marine 

solids are infectious for longer (19 days) than unassociated enteroviruses in the water column (9 

days) (Griffin et al., 2003). Shen et al. (2008) estimated somatic coliphage P22 inactivation rates 

to be in the range 0.27 to 0.57 per day (0.12 to 0.25 log10 per day) with the highest inactivation 

rate found in samples with high suspended solids concentration, relatively low dissolved organic 

carbon content, and sediment with high clay content. 

Chung and Sobsey (1993) found both temperature- and sediment-dependent differences between 

the decay rates of the five viruses tested: F-specific coliphages, B. fragilis phages, hepatitis A, 

poliovirus, and rotavirus. The effect of sediment differed among the viruses. Sediment protected 

poliovirus and human adenovirus at 5°C and 25°C and F-specific coliphages at 25°C, whereas it 

accelerated inactivation of rotavirus at both temperatures. B. fragilis phage survival was not 

affected by sediment at either temperature (Chung and Sobsey, 1993). Interestingly, at 5°C, all of 

the viruses had increasing levels of association with the sediment fraction over a 60-day period, 

except for hepatitis A, which had approximately constant rates over the entire period. 

Association with the sediment did not correlate with inactivation rates (Chung and Sobsey, 

1993). All five of the viruses tested had faster decay rates at 25°C than at 5°C (Chung and 

Sobsey, 1993). Under the conditions tested, F-specific coliphages had similar decay rates to 

poliovirus in sediment at 25°C and in seawater at 5°C, and rotavirus in sediment at 5°C (Chung 

and Sobsey, 1993).

Silverman et al. (2013) found that the presence of photosensitizers (presumably colored 

dissolved organic matter [CDOM]) in five different natural waters, had different effects on 

human virus and bacteriophage photoinactivation in different waters exposed to full spectrum, 

simulated sunlight. In four of the five natural waters, the inactivation rate of poliovirus type 3 

was significantly slowed relative to a clear, buffered control. In one of the five natural waters, 
51

the inactivation of PRD-1 (which infects Salmonella LT2) was significantly slowed. In three of 

five waters, adenovirus and GI F-specific coliphage MS2 inactivation was significantly faster 

than the clear control buffer. The authors also examined inactivation rates in UVB-blocked 

simulated sunlight to gain insight into the mechanisms of photoinactivation of the different 

viruses. The authors concluded that exogenous mechanisms (reaction reactive species formed by 

photosensitizers in the water column) contributed significantly to inactivation of the viruses other 

than poliovirus type 3 for which endogenous processes are likely dominant. 

Synergistic effects between organic and inorganic matter and other environmental factors 

Several studies have found synergy between organic and inorganic matter and other 

environmental factors in the inactivation rates of viruses. Sunlight has been shown to have 

synergistic effects with CDOM present in the water matrix, the effects of which vary depending 

on the type of virus, the amount of UVB attenuated by the CDOM, and the number and 

concentration of damaging radicals produced (Silverman et al., 2013). Please see Section 5.1, 

5.2, and 5.3 for more information sunlight, temperature, and salinity. Viral adsorption to 

biofilms, sediment and organic matter can protect viruses from inactivation or expose viruses to 

detrimental microbial activity. Please see Section 5.4 for more information on biofilms and 

predation and degradation by microbes. 

Summary

In summary, depending on the specific environmental conditions coliphages may be a 

conservative surrogate for the inactivation of human enteric viruses. The presence of organic and 

inorganic matter affects inactivation of enteric viruses and coliphages in aquatic systems. Both 

organic and inorganic matter have been shown to either increase or decrease degradation rates, 

depending on the type of the virus and the nature of the organic matter (Chung and Sobsey, 

1993; Silverman et al., 2013). For example, several groups found that poliovirus, echovirus, and 

enterovirus adsorption to sediment or solids decreased inactivation of the viruses (LaBelle and 

Gerba, 1980; Griffin et al., 2003), whereas others have found that inactivation rates increased in 

samples with high suspended solids and sediment with high clay content (Shen et al., 2008). 

While it is impossible to compare coliphages with all human enteric viruses under all conditions, 

Silverman et al. (2013) found that GI F-specific coliphage MS2 was a conservative surrogate for 

poliovirus type 3 and human adenovirus type 2 (i.e., GI F-specific coliphage MS2 had a slower 

decay rate than the human viruses) in five environmental waters with varying levels of 

photosensitizing molecules both in the dark and in full sunlight. 

5.6. Environmental Factors Impacts Summary 

Some studies have found that coliphages are more resistant to environmental stressors than 

human viruses, but such findings are highly contextual and dictated by a host of local 

environmental conditions. The inactivation kinetics of coliphages is also relative. In general, 

temperature, pH, sunlight, CDOM and the association with solids are some of the most important 

factors influencing survival of coliphages (Schaper et al., 2002b). Table 13 summarizes these 

environmental factors and their mechanisms of inactivation for human enteric viruses and 

coliphages. 
 52

Table 13. Summary of environmental factors influencing viral inactivation in aquatic environments.

Effects

References

Factor Coliphages

NoV and other

human enteric viruses Conclusions

Physical

Temperature  Variable decay rates among 

strains; wild isolates more stable 

than laboratory strains. 

 F-specific RNA coliphages are 

more resistant to decay at low 

temperatures than high 

temperatures. 

 Somatic coliphage ΦX174 is 

highly persistent under all 

temperatures from 0°C–100°C in 

a variety of matrices tested. 

Somatic and F-specific 

coliphages highly persistent at

higher temperatures. 

 At 4°C, GI and GII F-specific 

RNA coliphages are detectable 

for over 100 days, GIII Fspecific RNA coliphages

detectable for 3 weeks and GIV 

F-specific RNA coliphages 

reduced to the limit of detection 

after 10 days. 

 Different viruses have different 

decay rates at the same 

temperature.

 MNV more stable than human 

adenovirus and human rotavirus 

at 0°C and 50°C in a variety of 

media. 

 Viruses decay at faster rates at higher 

temperatures. More stable at 4°C, than 

at 20°C, and at <50°C more stable than 

at ≥50°C in a variety of media. 

 Salinity and sunlight have synergistic 

effects at temperatures ranging from 

0°C to 100°C (but in general, coliphage 

ΦX174 has lower decay rates than 

rotavirus, adenovirus, and MNV)

 Somatic and F-specific coliphages are 

classified as the most persistent of the 

viruses at higher temperatures.

 At 4°C and 25°C GI F-specific RNA 

coliphage MS2 has lower decay rates 

than FCV and similar survival rates at 

37°C in dechlorinated water. GI Fspecific RNA coliphage MS2 has fourfold lower decay rates than MNV at 

24°C and 37°C and three-fold lower 

decay rates at 50°C and 60°C.

Hurst et al., 1980; 

Chung and Sobsey, 

1993; Nasser et al., 

1993; Skraber et al., 

2002; Allwood et al., 

2003, 2005; 

Savichtcheva and 

Okabe, 2006; Lee 

and Sobsey, 2011; 

Bertrand et al., 2012; 

Seo et al., 2012
 53

Sunlight  Different viruses have different 

decay rates under the same 

sunlight conditions. Genome size 

of somatic coliphages is 

correlated with decay rate.

 F-specific RNA coliphages are

more resistant to sunlight than Fspecific DNA coliphages in clear 

seawater. 

 Inactivation rates vary more 

widely for ssDNA and dsDNA 

viruses than for ssRNA viruses 

based on nucleic acid type, 

genome length, and morphology. 

 Different viruses have different 

decay rates under the same 

sunlight conditions. 

 Poliovirus type 3 has faster 

decay rates than human 

adenovirus type 2 under full 

sunlight in four different 

environmental waters. 

 Virus inactivation rates are higher in 

sunlight conditions than in the dark. 

 UVB wavelengths are the most 

damaging. 

 Synergistic effects with temperature, 

salinity, organic, and inorganic matter. 

 Direct damage to protein capsid and 

genetic material and indirect 

inactivation due to reactive oxygen 

species and other free radicals. 

 In full sunlight in seawater, the decay 

rates of human adenovirus type 2 and 

F-specific coliphages (MS2, Fi, Qβ, 

and Sp) are similar.

 The decay rates of F-specific DNA 

coliphage M13 and poliovirus type 3 

are also similar.

 Decay rates for porcine rotavirus are 

two- to three-fold higher than decay 

rates for GI F-specific RNA coliphage 

MS2 when tested between 14°C and 

42°C. 

 GI F-specific RNA coliphage MS2 is a 

conservative surrogate for decay of 

poliovirus type 3 and human 

adenovirus type 2 in four types of

environmental water.

Bitton et al.,1979; 

Sinton et al., 1999, 

2002; Sobsey and 

Meschke, 2003; 

Duizer et al., 2004; 

Fong and Lipp, 2005; 

Love et al., 2010; 

Jończyk et al., 2011; 

Lee and Sobsey, 

2011; Romero et al., 

2011; Silverman et 

al., 2013
 54

Chemical

Salinity  GI F-specific RNA coliphage 

MS2 does not aggregate in high 

concentrations of monovalent 

cations but does aggregate in

high concentrations of divalent 

cations.

 Concentrations of 1.3 to 6.3%

NaCl were protective of GI Fspecific RNA coliphage MS2 at 

37°C and 50°C. 

 Salt water compared to 

freshwater affects the decay rate

of F-specific coliphages and 

somatic coliphages under both 

dark and sunlight-exposed 

conditions. 

 F-specific coliphages are more 

tolerant in salt water than in 

freshwater in the dark.

 Somatic coliphages are more 

tolerant of salt water than 

freshwater under sunlight 

conditions. 

 NoV GI.1 aggregates with both 

mono- and divalent cations, 

rotavirus aggregates with 

divalent cations.

 In seawater, FCV has an initial 

reduction (due to salt content), 

but retains infectivity over a 

month. 

 Salinity either increases or decreases

degradation rates of viruses based on 

type and concentration of salt and 

specific virus. 

 Salinity can affect viral adsorption to 

organic and inorganic matter.

 There are synergistic effects with 

salinity and temperature and organic 

and inorganic matter. 

 At 24°C, 37°C, and 50°C, GI F-specific 

RNA coliphage MS2 is more resistant 

to 0.3–6.3% NaCl concentrations than 

MNV.

Slomka and 

Appleton, 1998; 

Gutierrez et al., 2010; 

Mylon et al., 2010; 

da Silva et al., 2011; 

Nguyen et al., 2011; 

Seo et al., 2012
 55

Organic and 

Inorganic 

Matter

 39 to 65% of viral inactivation of 

coliphage T4 due to living and 

inert planktonic particles 0.2 

to10 μm.

 Both somatic and F-specific 

coliphages adsorb to particles 

<5 μm in size. 

 Somatic coliphages attach 

preferentially to particles <2 μm 

in size. 

 Sediment protects F-specific 

coliphages at 25°C, but not at 

higher temperatures. 

 Under full sunlight in 

environmental waters containing 

CDOM, GI F-specific RNA 

coliphage MS2 degradation is 

dominated by exogenous 

mechanisms.

 Adsorption to clay increases

persistence in lake water for 

reovirus (the family to which 

rotavirus belongs).

 Inactivation of poliovirus and 

echovirus decreases with 

adsorption to marine sediment. 

 Enteroviruses are associated 

with particulate matter protected 

from degradation. 

 Sediment protects poliovirus and 

human adenovirus at 5°C and 

25°C. 

 Sediment accelerates 

inactivation of rotavirus at 5°C

and 25°C.

 Under full sunlight, in 

environmental waters containing 

CDOM, poliovirus type 3 

degradation is dominated by 

endogenous mechanisms, human 

adenovirus type 2 degradation 

dominated by exogenous 

mechanisms.

 Organic and inorganic material impact 

viral degradation rates. 

 Organic matter either decreases or 

increases viral deactivation rates. 

 Viruses adsorb to suspended particulate 

matter. 

 Synergistic effects among sunlight, 

temperature, pH, and salinity. 

 F-specific coliphage decay rates are 

similar to poliovirus in sediment at 

25°C and in seawater at 5°C, and 

rotavirus in sediment at 5°C.

 MS2 is a conservative surrogate for 

decay of poliovirus type 3 and human 

adenovirus type 2 in different types of 

environmental waters. 

Chung and Sobsey, 

1993; Griffin et al., 

2003; Sobsey and 

Meschke, 2003; 

Characklis et al., 

2005; Kohn et al., 

2007; Shen et al., 

2008; Finiguerra et 

al., 2011; Romero et 

al., 2011; Silverman 

et al., 2013

Biological

Predation 

and 

Enzymatic 

Degradation 

(including 

biofilms)

 Wetland vegetation increases

inactivation of GI F-specific 

RNA coliphage MS2. 

 Somatic coliphage T4 

inactivation in seawater is 

accelerated by naturally 

occurring particles >10 kDa.

 Biofilms protect viruses from 

inactivation: somatic coliphage 

ΦX174 and GI F-specific RNA 

coliphage MS2 densities in 

drinking water biofilms decrease 

after 6 days. 

 The presence of microbial 

activity in raw and treated 

freshwaters decrease rotavirus 

persistence.

 Highest poliovirus inactivation 

rates in unfiltered natural 

seawater compared to filtered 

natural seawater, artificial 

seawater, and deionized water. 

 NoV is more persistent in 

biofilm samples and viral 

genomes persist longer in 

biofilm than in wastewater.

 Both predation and enzymatic 

degradation increase virus degradation 

rates in different types of water. 

 Virus adsorption to biofilms is 

protective, but microbial activity in 

biofilms causes virus inactivation and 

degradation. 

 Synergistic effects with temperature 

and the presence of organic and 

inorganic matter. 

 No data on direct comparisons of 

predation effects on human enteric 

viruses and coliphages.

Fujioka et al., 1980; 

Toranzo et al., 1982; 

Raphael et al., 1985; 

Deng and Cliver, 

1995; Wetz et al., 

2004; Skraber et al., 

2007; Karim et al., 

2008; Shen et al., 

2008; Skraber et al., 

2009; Weaver and 

Sinton, 2009; 

Finiguerra et al., 

2011; Helmi et al., 

2011
56

6. Wastewater Treatment 

Treated wastewater is a source of viruses in ambient water (Kageyama et al., 2003; da Silva et 

al., 2007; Haramoto et al., 2007; Kitajima et al., 2009; Kuo et al., 2010; Simmons et al., 2011). 

This section provides a broad overview of how coliphages, human enteric viruses, and FIB 

behave during various wastewater treatment processes. This section does not evaluate 

engineering technologies or provide specifics on treatment processes. Rather, the overall context 

is to evaluate whether coliphages could be better than traditional FIB at indicating removal or 

inactivation of human enteric viruses during wastewater treatment. 

Coliphages have been considered useful microorganisms for evaluating wastewater treatment 

efficacy (Duran et al., 2003; Lucena et al., 2004; Bitton, 2005). Because coliphages and human 

enteric viruses have similar morphological and structural characteristics (see Section 2.2), often 

co-occur in feces, and often share fate and transport characteristics (see Section 5.0), the 

reduction of human enteric viruses and coliphages may follow similar patterns during wastewater 

treatment depending on the method of pathogen removal (Havelaar et al., 1993; Turner and 

Lewis, 1995; Rose et al., 2004). These shared attributes of viruses also suggest that coliphages 

would be better indicators for human enteric viruses than traditional FIB in wastewater. This 

section discusses coliphage behavior during wastewater treatment and compares it to other 

enteric viruses (with a focus on NoV) and FIB. 

Somatic coliphages have been reported to outnumber F-specific coliphages in both treated and 

untreated wastewater sources (Grabow et al., 1993; Gantzer et al., 1998; Grabow, 2001; Aw and 

Gin, 2010). The lower density of indigenous F-specific coliphages is a potential limitation of 

their use as an indicator. The range of coliphage densities, specifically the lower end found in 

influent is highly variable. For example, within the influent for six WWTPs, Rose et al. (2004) 

found somatic and F-specific coliphages from 103to 106 PFU per 100 mL (host strain ATTC 

15597), and F-specific coliphages at 102to 108 PFU per 100 mL (host strain ATTC 700891). In a 

study of eight WWTPs in Canada that serve 20,000 to 60,000 people, Payment and Locas (2011) 

found F-specific coliphages in influent in a range of 102to 106 PFU per 100 mL. Because 

bacteria in biological treatment systems are not in logarithmic growth, it is unlikely that Fspecific coliphages replicate during treatment (Rose et al., 2004). 

Wastewater treatment processes are often categorized as primary, secondary, tertiary and 

advanced treatment, and disinfection. There are a variety of different secondary treatment unit 

processes that can produce different qualities of water. Tertiary treatment has different purposes 

and definitions depending on the State, and outside the United States. Definitions can vary 

widely. In addition, natural treatment systems, such as waste stabilization ponds, are commonly 

used to provide treatment that is roughly similar to primary and secondary treatment. There are a 

wide variety of technologies available for wastewater treatment, and almost all WWTPs in the 

United States include secondary treatment and some type of disinfection. This section focuses on 

the removal or inactivation of coliphages and enteric viruses during the various steps of 

wastewater treatment. 

It is important to understand that treatment efficacy depends on the quality of the effluent prior to 

disinfection (particularly turbidity or UV transmittance), pH, temperature, the type of 
57

chlorination (free or combined), chlorine dose and contact time, UV dose, and a number of other 

factors (Asano et al., 2007). Given the importance of the specifics of the treatment processes, it is 

difficult to draw generalized conclusions from studies that do not provide adequate information 

on the treatment specifics. For example, efficacy of chlorination depends on the type of 

chlorination, the contact time, and the specific nature of the secondary treated water. Coliphages 

can be resistant to some chlorination practices (Havelaar, 1987; Sobsey, 1989; Havelaar et al., 

1990; Yahya and Yanko, 1992; Nasser et al., 1993; Gantzer et al., 1998; Bitton, 2005; Harwood 

et al., 2005), but sequential chlorination (the free chlorine portion) can provide up to 6-log10

removal (LACSD, 2013). 

Ideally, to examine the question of how coliphages, FIB, and enteric viruses compare during 

wastewater treatment, a study would include the following design attributes: 

 enumeration of indigenous somatic and F-specific coliphages, FIB, and one or more 

human viruses (not addition of a laboratory generated stock of virus); 

 density in influent and effluent; 

 calculated log10 reduction values; 

 detailed information on the treatment processes that were applied including information 

on discharge requirements that would impact level of treatment; and 

 full-scale wastewater treatment facilities (not pilot and bench scale studies). 

Although most of the literature found for this review did not include all of the above attributes, 

the studies that provided the most relevant information are discussed in more detail (Rose et al., 

2004; Harwood et al., 2005; Aw and Gin, 2010; Keegan et al., 2012). It is beyond the scope of 

this review to conduct a meta-analysis for synthesizing data from the different studies, so each 

study is discussed individually. Given that this is a broad, high-level review and treatment details 

are lacking in most of the studies, the nuances of wastewater treatment diversity are not 

discussed. 

The Water Environment Research Foundation (WERF) conducted an evaluation of the reduction 

of pathogens, FIB, and alternative indicators (including somatic and F-specific coliphages) at six 

WWTPs that produced tertiary recycled water by collecting samples five times (approximately 

once every 2 months) over the course of a year (Rose et al., 2004). Samples were obtained from 

the WWTPs at various stages of the treatment process and the microorganism density was 

evaluated by culture dependent methods. A comparison of the log10 reductions of all coliphages 

and enteroviruses across all facilities indicated that the combination of primary and biological 

secondary treatment results in a ~2-log10 reduction of coliphages and enteroviruses, filtration 

results in a ~0.5-log10 reduction of both coliphages and enteric viruses, and disinfection results in 

a ~0.5-log10 reduction of coliphages, and little to no reduction of enteroviruses (Rose et al., 

2004). The lower average reduction of enteric viruses from disinfection at all six plants was 

concluded to be partially due to the fact that enteroviruses were below detection limits in 69% of 

the samples, and samples with no detection were recorded as being at the detection limit (Rose et 

al., 2004). Coliphages were closer in log10 reductions to enteroviruses than traditional FIB. 

Whereas coliphages and enteroviruses both had a cumulative reduction of ~3 to 4 log10, FIB had 

a cumulative reduction of ~5 to 6- log10.
58

The study found that the densities of coliphages and viruses in the influent samples from the 

different WWTPs were not significantly different; whereas the densities in final effluent were 

significantly different among WWTP (Rose et al., 2004). These data indicate that differences in 

effluent densities were related to the treatment processes employed in each WWTP. Although no 

correlation of the density of coliphages and enteroviruses was found, the authors suggest that it is 

possible to predict the absence of enteroviruses based on coliphage levels. Levels less than 10 

coliphage PFU per 100 mL (either F-specific coliphages, or F-specific combined with somatic 

coliphages) were indicative of effluents with no detectable cultivatable enteroviruses (Rose et al., 

2004). While Rose et al. (2004) reported log10 reductions, they did not provide detailed 

information on the treatment processes. 

Harwood et al. (2005) evaluated the same data reported in Rose et al. (2004). F-specific 

coliphages were detected in 100% of the influent samples at densities ranging from 103 PFU per 

100 mL to 108 PFU per 100 mL. Although enteroviruses were above detection limits in 31% of 

the disinfected effluent samples, coliphages and enteroviruses co-occurred in only 13% of the 

disinfected effluent samples. The authors reported a weakly significant relationship between the 

presence or absence of enteroviruses and coliphages in disinfected wastewater effluent (Harwood 

et al., 2005). 

Aw and Gin (2010) reported that, when comparing raw sewage to secondary effluent at a plant in 

Singapore (where wastewater is treated using activated sludge processes), on average, somatic 

and F-specific coliphage densities were reduced by 2.4-log10 and NoV GI and GII were reduced 

by ~2-log10. Specifically, somatic coliphages were reduced from 1.8 × 105to 102 PFU per 100 

mL, F-specific coliphages from 4.3 × 104to 102 PFU per 100 mL, NoV GI from 3.2 × 105to 7.1 

× 103 gene copies per 100 mL and NoV GII from 2.3 × 105

to 5.2 × 103gene copies per 100 mL. 

Coliphages were quantified by infectivity assays and NoV were quantified by qPCR 

amplification. PCR amplification can amplify both infectious and noninfectious virus particles 

and may therefore overestimate the number of infectious NoV particles. The authors found 

significant correlation between levels of somatic coliphages and adenoviruses, and between Fspecific coliphages and NoV GII in raw sewage samples (Aw and Gin, 2010).

Figure 1 shows example reductions for three WWTPs in Singapore (secondary effluent - 

activated sludge). Somatic and F-specific coliphages had on average 2.4-log10 reduction, and 

were reduced at a similar rate as enteric viruses, adenovirus, and astrovirus. NoV reductions 

were less, but assays were based on qPCR results evaluating both viable and nonviable NoV (Aw 

and Gin, 2010). 
59

Figure 1. Comparison of coliphages and enteric viruses in raw sewage 

and secondary effluent. 

Somatic and F-specific coliphages (PFU per 100 mL) and enteric viruses 

(gene copy number per 100 mL) isolated from raw sewage (n = 18) and 

secondary effluent (n = 18). The box represents 50% of the data values. The 

line across the inside of the box represents the median value, and the lines 

extending from the box represent the 95% CIs. Outliers are represented by 

circles. Hashed boxes are raw sewage, and open boxes are secondary effluent 

(adapted from Aw and Gin, 2010). 

Keegan et al. (2012) investigated the required chlorine and chloramine contact times for 

inactivating enterovirus (Coxsackie B5) and adenovirus 2. Enterovirus and adenovirus 2 were 

cultured and added separately to wastewater with varying turbidity levels (0.2, 2, 5, and 20 

nephelometric turbidity unit [NTU]) and pH (7, 8, and 9) at 10°C. The spiked samples were 

exposed to different chlorine/chloramine concentrations and contact times to determine the 

contact times for up to 4-log10 virus inactivation. Results demonstrated that increasing contact 

times are needed with increased turbidity and increased pH. For both viruses, a 4-log10

inactivation was possible even at the highest turbidity tested (20 NTU). The authors indicated

that the results of the study will be used in the development of new wastewater disinfection 

guidelines for Australia. 

Regulatory agencies in Australia use coliphages as indicators for wastewater treatment efficacy. 

When evaluating a WWTP, the South Australian and Victorian Departments of Health use 

minimum removal values as defaults for each treatment process, unless it has been demonstrated 

that a greater inactivation is achievable in the system (Keegan et al., 2012). Table 14 shows the 

log10 reductions for wastewater treatments used by the South Australian and Victorian 
60

Departments of Health. Note that coliphage removals are more similar to human virus removal 

than E. coli or bacterial pathogen removal for many treatments. 

Table 14. Log10 removals of enteric viruses and indicator organisms.

Treatment

Indicative Log10 Removalsa

Viruses (including 

adenoviruses, rotaviruses 

and enteroviruses)

Coliphages E. coli Bacterial 

pathogens

Primary treatment 0–0.1 N/A 0–0.5 0–0.5

Secondary treatment 0.5–2.0 0.5–2.5 1.0–3.0 1.0–3.0

Dual media filtration with 

coagulation

0.5–3.0 1.0–4.0 0–1.0 0–1.0

Membrane filtration 2.5–>6.0 3.0–>6.0 3.5–>6.0 3.5–>6.0

Reverse osmosis >6.0 >6.0 >6.0 >6.0

Lagoon storage 1.0–4.0 1.0–4.0 1.0–5.0 1.0–5.0

Chlorination 1.0–3.0 0–2.5 2.0–6.0 2.0–6.0

Ozonation 3.0–6.0 2.0–6.0 2.0–6.0 2.0–6.0

UVC light >1.0 adenovirus

>3.0 enterovirus, hepatitis 

A virus

3.0–6.0 2.0–>4.0 2.0–>4.0

Sources: Australian Guidelines for Water Recycling (2008) and Keegan et al. (2012)

a Reductions depend on specific features of the process, including detention times, pore size, filter depths, and 

disinfectant. The default values are accumulated across the treatment train processes. Each row shows only the 

reduction for that treatment step.

6.1. Primary Treatment 

Primary treatment of wastewater involves settling of solids in settling tanks and results in 

different reduction rates of different microbe groups. Viruses are too small to settle and are only 

removed during primary treatment if they are attached to larger particles. The settling velocities 

of individual bacteria and protozoan cysts are low compared to the retention time of 

sedimentation tanks; thus, their removal is also enhanced by attachment to larger particles. As a 

result, the removal efficiencies of microorganisms is a function of their association with 

wastewater particles. Asano et al. (2007) report that typical removal is <0.1- to 0.3-log10 for fecal 

coliforms, 0.1- to 1.0-log10 for Cryptosporidium, <1-log10 for Giardia, and <0.1-log10 for enteric 

viruses. Additionally, Lucena et al. (2004) determined the density of bacterial indicators (e.g., 

fecal coliforms, enterococci, and sulfate-reducing bacteria) and bacteriophages (e.g., somatic 

coliphages, F-specific coliphages, and B. fragilis-specific bacteriophages) that are present in 

incoming wastewater and effluent after primary treatment (for secondary treatment see below) 

from treatment plants in Argentina, Columbia, France, and Spain. The average reductions for the 

various indicators during primary settling ranged from 0.3- to 0.5-log10 units. Irrespective of the 

geographical location, no significant difference in the reduction of any of the indicator 

microorganisms was observed (Lucena et al., 2004). In the same study, the addition of lime had a 

significant effect on F-specific RNA coliphage removal, which approached 2-log10 units, but not 

on somatic coliphage removal (Lucena et al., 2004). At another WWTP in Ireland, after primary 

treatment the mean reduction of F-specific RNA coliphages was 0.32-log10 (SD ± 0.55-log10) 

(Flannery et al., 2012). Finally, Ottoson (2005) investigated the reduction of microorganisms and 

indicators at multiple WWTPs in Sweden, and found that during primary treatment, somatic 
61

coliphages were reduced by 0.8-log10 (SD = 0.4) and F-specific coliphages were reduced by 1.3-

log10 (SD = 0.7) (Ottoson, 2005). 

6.2. Secondary Treatment 

Secondary treatment of wastewater involves the use of a natural population of bacteria, such as 

the mixed liquor flocs in activated sludge treatment or the biofilm on trickling filters, to decrease 

biochemical oxygen demand (BOD), organic material, and in some cases nutrients (depending on 

the design). In activated sludge treatment, aeration is necessary to support the growth of the 

aerobic heterotrophic bacteria that consume the soluble organic material in the wastewater. 

Although secondary treatment is not designed to remove pathogens, removal of indicator 

organisms and pathogens often occurs.

Secondary treatment results in different log10 reduction values for different microorganisms and 

depends on the specifics of the secondary treatment. In a widely used general resource book 

(Water Reuse), Asano et al. (2007) report that the typical range of removal is 0 to 2-log10 for 

fecal coliforms, 1-log10 for Cryptosporidium, 2-log10 for Giardia, and 0- to 2-log10 for enteric 

viruses. In addition, Asano et al. (2007) report that secondary treatment using activated sludge 

results in a mean reduction of 1.83-log10 for GI F-specific RNA coliphage MS2. The Australian 

Guidelines for Water Recycling report log10 reduction ranges of 1- to 3-log10 for E. coli, 0.5- to 

2.5-log10 for coliphages, and 0.5- to 2-log10 for enteric viruses (NRMMC-EPHC-NHMRC, 

2008). A study of WWTPs in Argentina, Colombia, France, and Spain found that secondary 

treatment reduced somatic coliphages, Bacteroides fragilis bacteriophages, and F-specific 

coliphages between 1.0- to 1.6-log10 units (Lucena et al., 2004). In a study of WWTPs in 

Switzerland, Baggi et al. (2001) found that three WWTPs with mechanical, biological, and 

chemical processes provided 0.6- to 0.8-log10 reductions for F-specific and somatic coliphages. 

A fourth WWTP with mechanical, biological, and chemical processes, plus sand filtration 

provided 1- to 4.4-log10 reductions for F-specific and somatic coliphages (Baggi et al., 2001). 

While some coliphages and human virus removal occurs during secondary treatment, they are 

still typically detectable in non-disinfected secondary effluent. Aw and Gin (2010) detected 

somatic coliphages and F-specific coliphages along with adenoviruses, astroviruses, and NoVs in 

100% of the secondary effluent samples tested (Figure 1). Somatic coliphages and F-specific 

coliphages were present in secondary effluent at 100 PFU per 100 mL (Aw and Gin, 2010). In 

six WWTPs secondary effluents, Rose et al. (2004) found that somatic and F-specific coliphages 

ranged from 10 to 105 PFU per 100 mL, enterococci from 103to 105 CFU per 100 mL, and 

enteroviruses from 10 to 102 MPN per 100 mL. However, in 27% of the secondary effluent 

samples, enteroviruses were below the detection limits. In five Australian WWTPs, Keegan et al. 

(2012) found coliphages, adenoviruses, rotaviruses, reoviruses, NoV, and enteroviruses in 

secondary effluent (Table 15). 
62

Table 15. Virus densities in secondary treated wastewater samples from five Australian 

WWTPs. 

Sample 

location

Microorganism

Adenovirus

genome/L

Enterovirus

genome/L

Reovirus

genome/L

NoV

genome/L

Rotavirus

genome/L

F-specific 

RNA 

coliphages

PFU/L

Bolivar 1.59 x 105 0 3.7 x 108 2.7 x 105 1.63 x 104 5.0 x 105

Bolivar 2 9.3 x 106 0 0 2.0 x 103 >6.0 x 103 3.7 x 104

Glenelg 2.8 x 105 0 1.16 x 108 2.3 x 104 1.05 x 104 5.0 x 103

Cairns 8.1 x 105 ND ND ND ND ND

Brisbane 1.7 x 106 ND ND ND ND ND

ACT ND ND ND 4.0 x 103 ND 21

Source: Keegan et al. (2012)

Performed in triplicates with mean results shown in the table. 

ND = not detected.

Some other studies also measured log10 reduction values for coliphages and enteric viruses, but 

did not provide enough information on treatment design and operations to understand how these 

reductions might apply in other WWTPs. For example, Lodder and de Roda Husman (2005) 

found that secondary treatment resulted in the reduction of 1.8-log10 for NoV, 1.6-log10 for Fspecific coliphages, and 1.1-log10 for somatic coliphages. Ottoson et al. (2005) found that 

secondary treatment mean reductions from multiple WWTPs in Sweden were 1.73-log10

(SD = 0.6) for F-specific coliphages and 1.04-log10 (SD = 0.3) for somatic coliphages, which 

were similar to reductions of enteroviruses 1.3-log10 (SD = 0.7), and NoVs 0.89-log10 (SD = 0.3). 

FIB had higher log10 reduction values; enterococci was reduced 2- log10 (SD = 0.5) and E. coli

was reduced 2.3- log10 (SD = 0.6) (Ottoson et al., 2005). Flannery et al. (2012) measured the 

densities of FIB, F-specific coliphages, and NoV GI and GII in both influent and final effluent at

a WWTP. Treatment included preliminary processing by screening and grit removal followed by 

treatment with a conventional activated sludge system, including primary sedimentation, 

aeration, and secondary clarification, but no further treatment details were provided. A 

comparison of influent to secondary effluent found that mean culturable F-specific coliphage 

densities were reduced by 2.13-log10, NoV GI gene copy densities were reduced by 0.8-log10,

NoV GII gene copy densities were reduced by 0.92-log10, and E. coli densities were reduced by 

1.49-log10 (Flannery et al., 2012). 

Appendix B is a compilation of studies that investigated coliphage and NoV densities before, 

during, and/or after wastewater treatment. It includes mostly non-disinfected secondary effluent, 

but some disinfected effluents are also included as noted in the table. The Appendix B 

information is focused on NoV compared to coliphages, because of NoV’s importance as an 

enteric pathogen. NoV is the leading etiological agent of gastrointestinal illness in the United 

States, and of an estimated 36.4 million cases of domestically acquired gastrointestinal illness, 

NoV causes an estimated average of 20.8 million cases annually (Scallan et al., 2011). 

Some of the studies reviewed in this section evaluated correlations between coliphages and 

enteric viruses to determine the usefulness of coliphages as surrogates for human viral presence 

in non-disinfected secondary effluent. Gantzer et al. (1998) showed a significant correlation 

between the density of coliphages and infectious enteroviruses in secondary effluent and the 
63

correlation between the density of somatic coliphages and the presence of the enterovirus 

genomes (p-value <0.0001). No enteroviruses were isolated in secondary effluent (without 

disinfection) when the somatic coliphage density was between 100 and 10,000 PFU per L 

(Gantzer et al., 1998). Although the treatment specifics were different, these results are similar to 

those in Rose et al. (2004), who found that coliphage levels less than 10 PFU per 100 mL in final 

disinfected effluent contained no detectable cultivatable enteric viruses (Rose et al., 2004). The 

threshold level in the WERF study is based on tertiary disinfected effluent and not nondisinfected secondary effluent (Rose et al., 2004). Ottoson et al. (2006) found there was no 

significant correlation between the reduction of coliphages or FIB compared to viruses 

(enteroviruses and NoV) in secondary treated wastewater. Flannery et al. (2012) also found no 

correlation between the densities of E. coli and F-specific coliphages with either NoV GI or NoV 

GII levels in effluent wastewater (r < 0.07 in all instances). 

6.3. Wastewater Treatment Ponds 

Wastewater treatment ponds, also known as waste stabilization ponds or lagoons, are shallow 

synthetic basins that treat sewage in a single or series of anaerobic, facultative or maturation 

ponds. Aeration and encouragement of aquatic life are other possible features of wastewater 

treatment ponds. Verbyla and Mihelcic (2015) analyzed virus removal data from 71 different 

systems. They found weak to moderate correlation between virus removal and hydraulic 

retention time. For each log10 reduction of viruses a geometric mean of 14.5 days of retention 

(95th percentile was 54 days of retention) was required. GI F-specific RNA coliphage MS2 

coliphage is considered to be the best surrogate for studying sunlight disinfection in wastewater 

treatment ponds. Inactivation of coliphages by solar radiation in lagoons and ponding systems 

tends to be seasonal, with the most effective inactivation occurring in summer months (DaviesColley et al., 2005; Blatchley et al., 2007). Sunlight inactivation of viruses is discussed in 

Section 5.2 and is compared to UVC inactivation in Section 6.5.4.

The open water wetland is similar to a maturation pond, but instead of having planktonic algae, 

the algae are part of a biomat on the bottom of the pond. Silverman et al. (2015) found that 

removals of F-specific and somatic coliphages were similar in a pilot-scale system. Based on 

laboratory and modeling work, they determined that GI F-specific RNA coliphage MS2 was 

inactivated more slowly than poliovirus under summer conditions, but more rapidly under winter 

conditions. More research is needed to determine how the relative inactivation rates of 

indigenous coliphages (F-specific and somatic coliphages) and other enteric viruses change 

seasonally in open water wetlands.

6.4. Tertiary Treatment and Advanced Treatment 

Tertiary treatment typically refers to particle removal processes (e.g., granular media filtration, 

cloth filtration, or membrane filtration) that are employed before final disinfection. The amount 

by which viruses (and other pathogens) are reduced by filtration varies depending on filter 

characteristics, operating practices, microbial properties, including size, surface properties, and 

degree of association with other microorganisms or particles, and water quality variables (Levine 

et al., 2008). Tertiary treatment may also refer to chemical or biological nutrient removal 

processes (e.g., targeting nitrogen and/or phosphorus), although these processes are sometimes 

considered part of secondary treatment. Literature reports for treatment plants employing nutrient 
64

removal were included in Section 6.2. Advanced treatment trains, which can be applied to 

filtered tertiary effluents, can be used to further purify water for indirect or direct potable reuse. 

Advanced treatment typically involves advanced oxidation processes (AOPs) and dense 

membranes (nanofiltration and reverse osmosis) that target the removal of pathogens and trace 

organic contaminants (Leverenz et al., 2011; NRC, 2011; Gerrity et al., 2013). Membrane 

processes are reviewed here. Disinfection processes, including UV, ozone, free chlorine, 

combined chlorine, and AOPs are described in detail in Section 6.5.

Depth filtration involves the use of granular media (e.g., sand, anthracite, garnet, or activated 

carbon) in single (mono-media) or layered configurations (multi-media). Microorganism removal 

differs based on a variety of factors, including water quality, the type and size of granular media,

the filtration velocity, and the use of coagulant and/or polymer. Typical removals from depth 

filtration are reported to be 0 to 1-log10 for fecal coliforms, 0 to 3-log10 for Cryptosporidium, 0- 

to 3-log10 for Giardia, 0- to 1-log10 for enteric viruses and ~0.14- to 2-log10 for coliphages 

(Rajala et al., 2003; Hijnen et al., 2004; Zanetti et al., 2006; Asano et al., 2007; Hijnen and 

Medema, 2007). Asano et al. (2007) report that tertiary treatment using depth filtration results in 

a mean reduction of 0.29-log10 for GI F-specific RNA coliphage MS2, and Zanetti et al. (2006) 

found that tertiary sand filtration resulted in a mean reduction of 0.31-log10 for E. coli and 0.14-

log10 for somatic coliphages. 

Rajala et al. (2003) conducted both laboratory and pilot-scale experiments on rapid sand 

filtration of wastewater effluent from WWTPs in Finland. In the laboratory experiment, the rapid 

sand filtration reduced coliphages by 0.15- to 0.26-log10 (30–46%) at a hydraulic load of 5 

meters per hour and 0.13- to 0.27-log10 (23–38%) at a hydraulic load of 10 meters per hour. In 

the pilot experiments (hydraulic loads range 7.7 to 10 meters per hour), coliphages were reduced 

by 0.66- to 1.5-log10 (7–97%), depending on the plant (Rajala et al., 2003). Based on pilot-scale 

filter studies on rapid depth filtration, Williams et al. (2007) found that the removal efficiency of 

GI F-specific RNA coliphage MS2 (seeded into secondary effluent) was similar to that of E. coli

and total coliforms (~ 0.8 log10 at a loading rate of 12.2 meters per hour). The removal efficiency 

of MS2 was more sensitive to the coagulant dose, compared to the indicator bacteria. In an 

experimental rapid sand filtration setup, virus size (based on ΦX174, MS2, and T4 coliphages) 

was the only factor that influenced retention and the larger the virus, the greater the retention 

(Aronino et al., 2009). 

Levine et al. (2008) conducted experiments to examine pathogen reduction from sand filtration 

of secondary effluent at five full-scale water reclamation facilities in the United States (three 

plants using monomedium and two plants using dual media) at peak usage over the course of a 

year. These are the same facilities that are reported in Rose et al. (2004). The average reductions 

for all five plants ranged from 0.1- to 4.2-log10 for fecal coliforms, 0.3- to 1.1-log10 for infectious 

Cryptosporidium, 0.7- to 1.5-log10 for Giardia, 0.3- to 1.2-log10 for culturable enteroviruses, 0.3- 

to 1.3-log10 for F-specific coliphages, and 0.2- to 0.8-log10 for somatic and F-specific coliphages 

(Levine et al., 2008). The authors found that the differences in average reduction rates between 

plants were likely due to a combination of loading rates, chemical addition practices (chlorine 

and coagulant), backwashing and post backwashing operating strategies, and the effectiveness of 

upstream biological treatment and sedimentation (Levine et al., 2008). In general, log10

reductions of indicator bacteria (coliforms, enterococci, and Clostridium) was 2-to 9-fold greater 
65

than the log10 reduction of pathogens, suggesting that monitoring with bacterial indicators may 

over predict pathogen reductions. Rose et al. (2004) noted that shallow sand filters were more 

effective than deep-bed dual-media or monomedia filters for removal of coliphages and viruses. 

However, this result was affected by the fact that pre-disinfection (pre-chlorination) was used for 

the shallow sand filter tests but not for the deep-bed filters. 

Surface Filtration includes mechanical sieving of secondary effluent, through cloth, metal or 

synthetic woven materials with a pore size of ~10 to 30 μm. In comparative testing for 15- to

30-μm particles, surface filtration removed more particles than granular filtration over all particle 

sizes tested (Olivier et al., 2003). Asano et al. (2007) reported average reductions for surface 

filtration of 0- to 1-log10 for coliform bacteria and 0- to 0.5-log10 for enteric viruses. These 

results are consistent with Levine et al. (2008), who found that cloth filtration of secondary 

effluent at a full-scale water reclamation facility at peak usage over the course of a year resulted 

in average reductions of 3-log10 (range: 1.9 to 4.3) for fecal coliforms, 0.5-log10 (range: 0.3 to 

0.7) for infectious Cryptosporidium, 0.5-log10 (range: -0.4 to 1.3) for Giardia, 0.5-log10 (range: 

0.3 to 0.8) for culturable enteric viruses, 0.6-log10 (range: -0.1 to 1.8) for F-specific coliphages,

and 0.4-log10 (range: -0.1 to 1) for somatic and F-specific coliphages. 

Membrane filtration, a type of advanced treatment, involves forcing wastewater through a thin 

membrane filtering under pressure. Membranes with different sized pores can be used, including 

microfilters (>50 nm), ultrafilters (2 to 50 nm), nanofilters (<2 nm), and reverse osmosis 

(polymer matrix without discrete pores; particles are excluded and uncharged molecules pass 

through membrane by diffusion). In general, the smaller the pore size used, the greater the 

reduction of pathogens and the higher the operating pressure (Asano et al., 2007). For example, 

Asano et al. (2007) reported that typical removal of pathogens from microfiltration are 1- to 4-

log10 for fecal coliforms, 1- to 4-log10 for Cryptosporidium, 2- to 6-log10 for Giardia, and 0- to 2-

log10 for enteric viruses. Ultrafiltration results in removal of 3- to 6-log10 for fecal coliforms, >6-

log10 for protozoa, and 2- to 7-log10 for viruses. Nanofiltration results in removal of 3- to 6-log10

for all types of bacteria, >6-log10 for protozoa, and 3- to 5-log10 for viruses. Reverse osmosis 

results in reductions of 4- to 7-log10 for fecal coliforms, 4- to 7-log10 for Cryptosporidium, >7-

log10 for Giardia, and 4- to 7-log10 for enteric viruses (Asano et al., 2007). 

Perfectly intact nanofiltration and reverse osmosis membranes should not allow passage of any 

bacteria or viruses; however, leaks in seals, and membrane imperfections or damage to the 

membranes could allow their passage. Thus, monitoring the integrity of membranes is critical to 

ensuring high removal of microorganisms. 

Juby (2003) found that microfiltration of screened primary effluent at a demonstration plant in 

California resulted in typical reductions of 4.7-log10 for fecal coliforms and 1.7-log10 for 

coliphages. Gomez et al. (2006) used secondary effluent from a WWTP in Spain to determine 

pathogen reduction rates from microfiltration and ultrafiltration. They found that microfiltration 

resulted in a 2.7-log10 reduction in fecal coliform, the removal of E. coli below detectable limits,

and a 1.3-log10 reduction of coliphages, whereas ultrafiltration resulted in a 4.7-log10 reduction of 

fecal coliform, the removal of E. coli below detectable limits, and a 3.5-log10 reduction of 

coliphages (Gomez et al., 2006). 
66

Membrane Bioreactors are a relatively new technology that combine an activated sludge 

bioreactor with membrane filtration, which replaces both secondary and tertiary treatment.

Zhang and Farahbakhsh (2007) found that membrane bioreactor pilot plants achieved 5.8-log10

removal of coliphages. Whereas conventional activated sludge process followed by advanced 

tertiary treatment (nitrifying rotating biological contactors, sand filtration, and chlorination), 

achieved 5.5-log10 removal of coliphages. For membrane systems, coliphages appear to be better 

indicators of microbial removal efficacy (especially viral removal) probably because the pore 

size of most microfiltration and ultrafiltration membranes exclude fecal coliforms, but some 

coliphages can still pass through the membrane pores (Zhang and Farahbakhsh, 2007). In a fullscale membrane bioreactor study, Purnell et al. (2015) reported a 5.3-log10 reduction in 

indigenous somatic coliphages. Indigenous F-specific coliphages were less abundant and 

demonstrated a 3.5-log10 reduction. In ‘spiking’ experiments, suspended GI F-specific RNA 

coliphage MS2 demonstrated a 2.25-log10 reduction (Purnell et al., 2015). 

6.5. Disinfection 

Disinfection of secondary effluent can be achieved using physical (UVC radiation) and chemical 

(chlorine, chloramines, and ozone) treatments. This section focuses on the physical and chemical 

treatments of secondary effluent and the effects of these treatments on coliphages, FIB, and 

enteric virus inactivation. Although solar radiation can also play a role in further disinfection of 

secondary effluent by lagooning (Gomila et al., 2008), lagooning is not typically considered a 

disinfection treatment (see Section 6.3). It is important to note that if ammonium levels are not 

reported, it cannot be determined whether free chlorine or combined chlorine was present during 

the disinfection step. This is important because many studies report on water samples from 

secondary disinfected effluent, but there is wide variation in what secondary disinfected effluent 

includes. 

As mentioned previously, studies with water samples collected at full-scale WWTPs are 

preferred. However, pilot scale and bench-scale studies are also included when full-scale data 

were not available. 

6.5.1 Free Chlorine 

Chlorine (Cl-) is the most widely used wastewater disinfectant (Asano et al., 2007). Chlorine is 

an efficient disinfectant for most enteric bacteria, but is generally less efficient against viruses, 

protozoan cysts, and bacterial spores (Keegan et al., 2012). The effectiveness of chlorine is 

impacted by disinfection dose, contact time, temperature, and water quality variables (pH, 

turbidity, presence of ammonia and oxidant demand) (Rose et al., 2004; Asano et al., 2007). For 

example, above pH 7, a 10 mg per L residual chlorine resulted in 4-log10 reduction of F2 

coliphages, whereas at low pH, as little as one-fifth of this chlorine achieved the same reduction 

(Hajenian and Butler, 1980). 

Due to the highly reactive chemical nature of free chlorine (HOCl and OCl-

), in secondary 

effluents containing ammonium, it rapidly combines with ammonium to form chloramines, a 

form of combined chlorine (U.S. EPA, 2002; Tree et al., 2003; Asano et al., 2007). Disinfection 

with free chlorine can be achieved in ammonium-containing effluents if “breakpoint” 

chlorination is practiced, in which sufficient free chlorine is added to convert all ammonium to 
67

nitrogen. Free chlorine is a much stronger oxidant than combined chlorine, and is more effective 

at inactivating pathogenic bacteria and coliphages than combined chlorine (Tyrrell et al., 1995; 

Duran et al., 2003; Tree et al., 2003; Keegan et al., 2012). Combined chlorine will be discussed 

in more detail in Section 6.5.2. It is important to note that both free chlorine and combined 

chlorine can result in formation of hazardous disinfection byproducts such as trihalomethanes, 

haloacetic acids, chlorite, and other hazardous compounds. 

In a bench-scale study, Shin and Sobsey (2008) studied the inactivation of NoV by free chlorine 

using molecular methods and compared its inactivation to F-specific RNA coliphage MS2 and 

poliovirus type 1. F-specific RNA coliphage MS2 and poliovirus type 1 were measured using 

both culture-based and molecular methods. The authors conducted experiments using 1 and 5 mg 

per L free chlorine. Inactivation of NoV was similar to F-specific RNA coliphage MS2 and faster 

than poliovirus type 1 when densities were measured using molecular methods. They also 

showed that the CT (disinfectant concentration times contact time) required for NoV inactivation 

was not significantly different from other viruses even though the molecular methods likely 

overestimate the CT needed. Thus, the study authors concluded that chlorine is a useful 

disinfectant for NoV. 

If a study evaluates disinfected effluent, but does not report ammonium levels, it cannot be 

determined whether free chlorine or combined chlorine was present. Of the six WWTPs 

evaluated in Rose et al. (2004), four used chlorine disinfection, but only one of the WWTP had 

ammonium levels that allowed for free chlorine. Rose et al. (2004) combined the data from the 

four WWTPs and found that on average, 300 minutes of contact time with chlorine (or combined 

chlorine) in secondary effluent resulted in a 3-log10 reduction of enterococci, whereas 500 

minutes of contact time were required for a 3-log10 reduction of enteroviruses. Data from the 

WWTP with free chlorine disinfection of nitrified and filtered effluent is show in Table 16. More 

studies that compare chlorine inactivation of FIB, indigenous F-specific and somatic coliphages, 

and enteric viruses in nitrified effluent are needed. 

Table 16. Average (and percent positive) microorganism effluent densities in a WWTP with 

free chlorine treatment of nitrified and filtered secondary wastewater.

Treatment

Microorganism

Total coliform

CFU/100 mL

Enterococci

CFU/100 mL

Somatic and Fspecific 

coliphages

PFU/100 mL

F-specific

coliphages

PFU/100 mL

Enterovirus

MPN/100 mL

Influent 3.41 x 107

(100%)

7.36 x 105

(100%)

2.84 x 105

(100%)

3.14 x 105

(100%)

1.52 x 104

(100%)

Filtered and 

disinfected with 

free chlorine

11.3

(60%)

0.2 detection 

limit

(0%)

10.4

(80%)

10.4

(80%)

0.3

(80%)

Source: Rose et al. (2004)

Over a 13-month period water samples were taken at Easterly WWTP in Vacaville, California 

and evaluated for FIB, F-specific coliphages, NoV, and other pathogens (Olivieri et al., 2012). 

This WWTP has bar screens, grit removal, primary clarification, secondary treatment with 

activated sludge, secondary clarification, nitrification, and chlorine disinfection and de-
68

chlorination. Log10 removal of fecal coliform through secondary disinfection was a median of 

6.8-log10 (range 6.1- to 8.4-log10; all non-detects were set at the detection limit). In finished 

effluent, fecal coliforms were not detected in 50 of the 55 samples (<2 MPN per 100 mL 

detection limit), and the remaining 5 were at the limit of detection. For enterococci, log removals 

through secondary disinfection was a median of 5.8-log10 (range 3.2- to 6.2-log10; all non-detects 

were set at the detection limit). In finished effluent, enterococci were not detected in 22 of 32 

samples (<1 MPN per 100 mL detection limit). The 10 detectable enterococci samples ranged 

from 1 to 648.8 MPN per 100 mL. F-specific coliphages were detected in all 32 influent samples. 

Densities ranged from 60 PFU per 100 mL to 13,000 PFU per mL, with a median of 2,750 PFU 

per 100 mL. F-specific coliphages in the final disinfected and dechlorinated effluent were below 

the detection limit (<1 PFU/100 mL) in all but two samples. One sample was at the detection 

limit (1 PFU/100 mL), and the other sample was 2 PFU per 100 mL. The log removal of Fspecific coliphages had a median of 3.4-log10 (range 1.8- to 4.1-log10). NoV were present in the 

ten WWTP influent samples and not detected in eleven final disinfected and dechlorinated 

effluent samples (Olivieri et al., 2012). This demonstrates that free chlorine applied to nitrified 

effluent is quite effective at inactivating F-specific coliphages and NoV. 

6.5.2 Combined Chlorine 

As stated above, studies have shown that in secondary effluent with ammonium, the free chlorine 

rapidly combines with the ammonium to form chloramines (Asano et al., 2007). Combined 

chlorine compounds are less effective at inactivating microorganisms than free chlorine (Tyrrell 

et al., 1995; Duran et al., 2003; Tree et al., 2003; Keegan et al., 2012). 

Tree et al. (2003) studied the chlorine-mediated inactivation of both seeded (E. coli,

Enterococcus faecalis, GI F-specific RNA coliphage MS2, and poliovirus – all measured using 

culture-based methods) and naturally occurring (E. coli, enterococci, F-specific coliphages, and 

enterovirus – also using culture-based methods) bacterial and viral indicators in primary sewage 

effluent. The inactivation rates of three applied doses of free chlorine (8, 16, and 30 mg per L) 

were investigated in both seeded sterilized primary effluent and unsterilized primary effluent. 

Although free chlorine was applied, Tree et al. (2003) found that the amount of free chlorine 

available in effluent decreased rapidly within the first 5 minutes and then remained 

approximately constant for the duration of the experiments (30 minutes). In both experiments, 

the authors found that FIB (E. coli and enterococci) were inactivated more rapidly and at lower 

doses than the viruses (F-specific coliphages, poliovirus, and enterovirus) and that chlorine dose 

and time of exposure had significant effects on survival of all organisms tested in both 

experiments (Table 17). Both E. coli (laboratory-cultured and indigenous) and enterococci had 

linear degradation rates and were completely inactivated over the course of the experiment at all 

chlorine applications tested. In contrast, enteroviruses had biphasic degradation rates, with a 

rapid initial rate, followed by a slower inactivation rate. F-specific RNA coliphages (both 

laboratory-cultured and naturally occurring) only showed degradation for the first 5 minutes of 

exposure after which no further degradation occurred. The authors suggest that the slower rate of 

degradation for enteroviruses and lack of degradation of F-specific RNA coliphages after 5

minutes is likely due to the weaker effect of combined chlorine on viruses. The authors conclude 

that F-specific RNA coliphages are a useful and conservative model surrogate for chlorine 

inactivation of viruses in sewage (Tree et al., 2005). 
69

Table 17. Log10 reduction of FIB, enteric virus, and F-specific coliphages in sewage matrix 

due to chlorine (adapted from Tree et al., 2003).

Matrix Microorganism Applied chlorine concentration

8 mg/L 16 mg/L 30 mg/L

Seeded into sterilized primary 

sewage matrix

E. coli >5

(<5 min.)

>5

(<5 min.)

>5

(5 min.)

Enterococcus 1

(<5 min.)

<1

(<15 min.)

>5

(5 min.)

Poliovirus 1

(after 30 min.)

<2

(after 30 min.)

4

(after 30 min.)

F-specific RNA 

coliphage MS2

<1

(after 30 min.)

<1

(after 30 min.)

1

(after 30 min.)

Naturally occurring in raw 

sewage (after primary 

treatment)

Indigenous E. coli 4

(5 min.)

4-5

(5 min.)

5

(5 min.)

Indigenous

Enterococcus

>3

(15 min.)

>3

(5 min.)

>3

(5 min.)

Indigenous

Enteroviruses

<1

(after 30 min.)

>1

(after 30 min.)*

>1

(after 5)*

Indigenous Fspecific RNA 

coliphages

<1

(after 30 min.)

<1

(after 30 min.)

<1

(after 30 min.)

* Because these were naturally occurring level (so lower density than seeded samples), the detection limit was

reached after about a 1-log10 reduction. 

Note: Times in parenthesis indicate the duration of the chlorine treatment. Log reductions were estimated based on 

graphical information. 

Duran et al. (2003) determined chlorine-mediated inactivation rates of both spiked and naturally 

occurring FIB, bacteriophages, and enteroviruses in secondary effluents. The authors found that 

after secondary effluent was exposed to 10 mg per L chlorine at the WWTP, naturally occurring 

FIB (E. coli and enterococci) were reduced at significantly higher rates than were naturally 

occurring viruses (F-specific RNA coliphages, somatic coliphages, B. fragilis bacteriophages, 

and enteroviruses). Specifically, mean reductions of naturally occurring microorganisms in 

chlorinated secondary effluent were: 2.9-log10 (SD = 2.5) for fecal coliforms, 2.0-log10 (SD = 0.7) 

for enterococci, 1.6-log10 (SD = 0.6) for somatic coliphages, 0.6-log10 (SD = 0.5) for F- specific 

RNA coliphages, 0.3-log10 (SD = 0.3) for B. fragilis bacteriophages, and 0.4-log10 for 

enteroviruses (no SD was given due the low number of positive samples) (Duran et al., 2003). 

Chlorination of secondary effluent in the laboratory resulted in similar inactivation rates as those 

found from chlorination at the WWTP (Duran et al., 2003). Both F-specific and somatic 

coliphages were inactivated more efficiently than enteroviruses. However, coliphages were 

closer to enterovirus log10 reductions than FIB. 

To determine if different types of viruses have different levels of resistance to chlorine, Duran et 

al. (2003) spiked secondary effluent with several bacteriophages, the vaccine strain of poliovirus 

type 1 Lsc 2a, and an enterovirus isolated from the environment, AR51101-1. The log10 reduction 

of bacteriophages and enteroviruses are presented in Table 18.
70

Table 18. Log10 reduction of bacteriophages and enteroviruses in spiked secondary effluent 

after chlorination with 20 mg/L of chlorine.*

Phage or Virus 20 Min 30 Min

Enteroviruses AR51101-1 3.1 3.6

Poliovirus type 1 1.3 1.6

Somatic coliphages ΦX174 2.8 3.3

MY2 2.3 2.8

SS13 0.2 0.4

SR51 2.0 2.5

SC12 1.0 1.7

F-specific RNA coliphage MS2 1.3 2.1

Bacteriophages specific to B. 

fragilis HSP40

B40-8 0.1 0.4

Source: Duran et al. (2003)

*Concentration of free chlorine was 0 after 20 minutes.

Haramoto et al. (2006) determined the densities of NoV GI and GII and F-specific coliphages in 

the raw sewage and final effluent at a WWTP that used conventional activated sludge process 

followed by chlorination. When comparing raw sewage to chlorinated secondary effluent, on 

average, NoV GII genome copies were reduced by 3.69-log10, E. coli densities were reduced by 

3.37-log10, total coliform densities were reduced by 3.05-log10, F-specific coliphage densities 

were reduced by 2.81-log10, and NoV GI genome copies were reduced by 2.27-log10 (Haramoto 

et al., 2006). 

Other studies have also demonstrated that FIB are more sensitive to chlorine than viruses. Tyrrell 

et al. (1995) found that combined chlorine was ineffective at inactivating F-specific coliphages 

(such as F-specific RNA coliphages), but was effective at eliminating vegetative bacteria from 

secondary effluent. Dee and Fogleman (1992) evaluated coliphage removal in a Denver 

treatment facility, because routine monitoring of the plant showed that coliphages were escaping 

the wastewater treatment processes. Monochloramine residuals were monitored in the effluent,

and the contact times that coliphages had with monochloramines were measured. In the WWTP 

studied monochloramines alone were not capable of a 2-log10 reduction in coliphage density 

(Dee and Fogleman, 1992). Average coliphage densities in effluent were 0.36 ± 1.03 PFU per 

100 mL in the summer and 0.08 ± 0.19 PFU per 100 mL for the rest of the year (Dee and 

Fogleman, 1992). 

In a bench-scale study, Sobsey et al. (1988) compared the inactivation of hepatitis A virus, 

coxsackievirus B5, F-specific RNA coliphage MS2, and somatic coliphage ΦX174 when 

exposed to 0.5 mg free chlorine (pH 6-10) per L and 10 mg monochloramine per L in phosphate 

buffer. Hepatitis A virus was inactivated quickly by free chlorine, but was relatively resistant to 

monochloramine. Coxsackievirus was relatively resistant to both. Somatic coliphage ΦX174 was 

most sensitive of the viruses to free chlorine at all pHs, except pH 10, and was the virus most 

sensitive to monochloramine. Inactivation of F-specific RNA coliphage MS2 by free chlorine 

was quicker than hepatitis A at low pH, but less rapid at higher pHs. In this study F-specific 

RNA coliphage MS2 was the virus most resistant to monochloromine. 

Log10 reduction values for all the human enteric viruses compared to coliphages do not seem to 

be available. Given treatment diversity, it might be impossible to rank all the enteric viruses in 
71

order of resistance. However, it should be noted that adenoviruses seem to be more resistant to 

combined chlorine than enterovirus (Irving and Smith, 1981; Cromeans et al., 2010). 

Adenovirus-2 is one of the most resistant viruses to chloramines and adenovirus-2 has similar 

resistance as adenovirus 40 and 41 (Keegan et al., 2012). Reovirus may be even more difficult to 

remove than adenovirus and enteroviruses through secondary treatment processes (Irving and 

Smith, 1981). Future studies that compare the behavior of coliphages to adenovirus would be 

helpful for evaluating the utility of coliphages as indicators of the presence of viruses in 

wastewater.

6.5.3 Ozone 

Ozone is a highly reactive chemical that damages microorganisms and reacts with water to 

produce hydroxyl radicals (OH-) that oxidize organic pollutants (Paraskeva and Graham, 2002). 

Like other disinfection processes, high doses of ozone can result in hazardous disinfection 

byproduct formation such as bromate. Additionally ozone decomposition occurs faster at higher 

temperatures and higher pH, which can alter disinfection efficacy (U.S. EPA, 1999). Viruses as,

a group, are the most sensitive microorganisms to ozone of all the microorganisms listed on 

EPA’s Contaminant Candidate List (U.S. EPA, 1998; Gerba et al., 2003). 

Several studies have shown that in secondary effluent, ozone is more effective at inactivating 

coliphages than FIB (Tyrrell et al., 1995; Gehr et al., 2003; Tanner et al., 2004). For example, 

Gehr et al. (2003) found that a transferred ozone dose of 17 mg per L resulted in a 3-log10

reduction of F-specific RNA coliphages, whereas a transferred ozone dose of 30 to 50 mg per L

was required for a 2-log10 reduction in fecal coliforms and resulted in less than a 1-log10

reduction in Clostridium perfringens (Gehr et al., 2003). Tyrrell et al. (1995) found that 

secondary sewage treated with a pulse of ozone [mean residual ozone concentrations of 0.30 

ppm (SD = 0.08)] resulted in approximate mean reductions of 2.5-log10 for F-specific coliphages, 

2.25-log10 for somatic coliphages, 1.3-log10 for fecal coliforms, 1.2-log10 for enterococci, and 

0.2-log10 for C. perfringens. Lazarova et al. (1998) found that 5 minutes of contact time of a 5-

mg per L dose of ozone resulted in a 5-log10 removal of F-specific RNA coliphage MS2. 

Tanner et al. (2004) investigated the effects of ozone on poliovirus, F-specific RNA coliphage 

MS2, Klebsiella terrigena, E. coli, heterotrophic plate count bacteria, fecal coliforms, and total 

coliforms, either in secondary effluent or reverse osmosis treated water. In secondary effluent, 

continuous ozone treatment for 1 minute resulted in an average inactivation of 2.5-log10 for 

coliphages, and <1.5-log10 reductions for total coliforms, fecal coliforms, and heterotrophic plate 

count bacteria. In demand-free reverse osmosis treated water, the authors found that 1 minute in 

0.2 mg ozone per L resulted in a >3-log10 inactivation of poliovirus and 1 minute in 0.25 mg 

ozone per L resulted in a 6-log10 inactivation of F-specific RNA coliphage MS2. Tanner et al. 

(2004) also found that increasing the concentration of ozone reverse osmosis treated water 

resulted in increased inactivation of all indicator organisms tested (F-specific RNA coliphage 

MS2, Klebsiella terrigena, and E. coli). Table 19 presents the highest log10 reductions of each 

indicator organism at a given ozone concentration. 
72

Table 19. Inactivation of FIB and F-specific RNA coliphage in ozone disinfected water.*

Indicator Log10 inactivation Ozone concentration (mg ozone/L)

F-specific RNA coliphage MS2 ≥5.41 0.22

Klebsiella terrigena 4.71 0.25

E. coli 4.15 0.25

poliovirus >3 0.2

Source: Tanner et al. (2004)

*Water was spiked after receiving RO treatment.

Finch and Fairbairn (1991) found that in demand-free phosphate buffer, 1.6-log10 units more 

inactivation was observed with GI F-specific RNA coliphage MS2 than with poliovirus type 3. 

The authors conclude that use of MS2 coliphage as a surrogate organism for studies of enteric 

virus with ozone disinfection systems can overestimate the inactivation of enteric viruses. In 

contrast, Shin and Sobsey (2003) documented the inactivation of MS2, Norwalk virus, and 

poliovirus type 1 in the presence of ozone using infectivity assays (for MS2 and poliovirus type 

1) and RT-PCR (all three viruses). Using a 0.37-mg per L dose of ozone at pH 7 and 5°C, the

authors found that the three viruses were inactivated approximately 3-log10 within 5 minutes and 

hence had similar inactivation behavior when detected using molecular methods. Inactivation 

measured by infectivity of F-specific RNA coliphage MS2 and poliovirus type 1 agreed well 

with their inactivation using molecular methods. Hall and Sobsey (1993) studied the decay of Fspecific RNA coliphage MS2 and hepatitis A virus in buffered water when exposed to ozone, as 

well as ozone and hydrogen peroxide in series. They found that both F-specific RNA coliphage 

MS2 and hepatitis A virus behaved similarly and were rapidly inactivated (up to 6-log10 in 5 

seconds) by two types of treatments. 

6.5.4 UVC 

In contrast to oxidative disinfection processes with chemicals like free chlorine and ozone, the 

efficacy of UVC (hereafter referred to as UV) disinfection is not affected by conditions like 

temperature, pH, and the presence and concentration of reactive organic matter (UV absorbance 

by organic and inorganic matter can shield microorganisms from UV, but the reactive properties 

of the organic matter don’t affect the UV, as happens with chemical disinfectants) (Oppenheimer 

et al., 1993; Hijnen et al., 2006). UV light is primarily absorbed by nucleic acids of 

microorganisms, causing harmful photoproducts such as thymine dimers on the same nucleic 

acid strand. If the damage is not repaired, replication is blocked, leading to subsequent 

inactivation of microorganisms (Ko et al., 2005). UV inactivation of microorganisms, including 

coliphages, is proportional to the UV fluence or dose, the product of the UV intensity and 

exposure time. Unlike free and combined chlorine and ozone, UV does not produce harmful 

disinfection byproducts (Oppenheimer et al., 1993). UVC has a shorter wavelength than UVA 

and UVB. UVC is filtered by the atmosphere, so does not occur in sunlight that reaches the 

surface of the earth. UVC is the most biologically damaging of the three UV wavelength classes 

and can be created artificially with UVC bulbs for treatment of water. UVC wavelengths (100 to 

280 nm), also called short-wave or germicidal UV, have been shown to result in a 1.09- to 2-

log10 reduction of indicator bacteria and coliphages in secondary effluent (Rose et al., 2004). 
73

In general, coliphages have been found to be more resistant to UVC light than FIB. For example, 

Gehr et al. (2003) demonstrated that GI F-specific RNA coliphage MS2 is more resistant to UV 

inactivation than fecal coliforms in effluent. Tree et al. (2005) found that F-specific RNA 

coliphage MS2 was more resistant to UV reduction than E. coli (4-log10 reduction required 62.5 

mJ/cm2for MS2 and 5.32 mJ/cm2for E. coli) in seeded, sterilized secondary effluent. Wilson et 

al. (1992) found that the viruses GI F-specific coliphage MS2, rotavirus, poliovirus, and hepatitis 

A were at least 7.1 times more resistant than the bacteria Klebsiella terrigena, Legionella 

pneumophila, Salmonella typhi, Aeromonas hydrophila, E. coli, Campylobacter jejuni, Yersina 

enterocolitica, Shigella dysenteriae, and Vibrio cholerae. In general, the bacteria tested were 

three to ten times more susceptible to UV irradiation than the viruses (Wilson et al., 1992). 

One of the six WWTPs studied in Rose et al. (2004) used UV disinfection. Table 20 shows the 

densities of microorganisms in the filtered secondary effluent compared to the filtered secondary 

effluent after UV disinfection. 

Table 20. Average (and percent positive) microorganism densities in a WWTP with UV 

treatment of filtered secondary effluent (n=5).

Treatment

Microorganism

Total coliform

CFU/100 mL

Enterococci

CFU/100 mL

Somatic and Fspecific 

coliphages

PFU/100 mL

F-specific 

coliphages

PFU/100 mL

Enterovirus

MPN/100 mL

Filtered 

secondary 

effluent

1.79 x 104

(100%)

5.8 x 102

(100%)

1.14 x 103

(100%)

1.41 x 102

(100%)

0.7

(40%)

Filtered 

secondary 

effluent 

disinfected with 

UV with free 

chlorine

11.9

(80%)

4.38

(20%)

10 detection 

limit

(0%)

10 detection 

limit

(0%)

0.5 detection 

limit

(0%)

Source: Rose et al. (2004).

The National Water Research Institute (NWRI) and Water Research Foundation (WRF) 

Ultraviolet Disinfection Guidelines for Drinking Water and Water Reuse provided information 

on log10 inactivation of viruses and FIB. They indicated that water is essentially “pathogen-free” 

if a 5-log10 poliovirus reduction and a 7-day median total coliform density of 2.2 MPN per 100 

mL is achieved. When media filtration is employed, effluent quality can vary, and particulate 

matter may shield pathogens from UV light to various degrees. In these cases, a reduction 

equivalent dose of 100 mJ (millijoules) per square centimeter (cm2) is typically adequate to 

inactivate total coliform to less than 2.2 MPN per 100 mL. The report also indicated a 5-log10

reduction of poliovirus can be achieved with a UV dose of 50 mJ/cm2based on laboratory 

studies, however the 100 mJ per cm2dose is recommended to account for effluent variability. 

When using membrane filtration or ultrafiltration prior to UV treatment, the impact of particles is 

normally eliminated. In this situation, the Ultraviolet Disinfection Guidelines for Drinking Water 

and Water Reuse noted that a 5-log10 reduction in poliovirus can be achieved with a UV dose of 

50 mJ/cm2, and a design UV dose of 80 mJ per cm2is suggested to account for variability in the 
74

effluent quality for membrane filtration or ultrafiltration. When using reverse osmosis for 

filtration, a reduction of at least 2-log10 for viruses can be achieved through the reverse osmosis 

process, and the additional 3-log10 reduction required for poliovirus can be achieved with a UV 

dose of about 30 mJ per cm2. Therefore, to account for variability in the effluent quality for 

reverse osmosis the design UV dose of 50 mJ per cm2 is recommended (NWRI-WRF, 2012). 

UV disinfection efficiency of secondary effluent is influenced by hydraulic properties of the 

reactor and wastewater characteristics, such as initial density of microbes, UV absorbance, and 

the concentration and characteristics of suspended solids (Koivunen, 2007; NWRI-WRF, 2012). 

For example, organic humic acid floc particles shielded viral surrogates (F-specific RNA 

coliphage MS2 and somatic coliphage T4) from UV light to a greater degree than inorganic 

kaolin clay floc particles of similar size (diameters <2 mm) (Templeton et al., 2005). However, 

humic acid floc particles also caused a greater reduction in log10 inactivation virus than larger 

activated sludge particles, which suggests that particulate chemical composition (e.g., UV 

absorbing content) and size may be important factors in the survival of particle-associated 

viruses during UV disinfection (Templeton et al., 2005). Because the study did not include 

human viruses, more data are needed to know whether these results extend to human viruses. 

Table 21 presents the estimated rate constants from a study on UV inactivation of F-specific 

RNA coliphage MS2 (NWRI-WRF 2012). 

Table 21. UVC inactivation of F-specific RNA coliphage MS2. 

Dose

(mJ/cm2)

Surviving density

(PFU/mL)

Log survival

(log PFU/mL)

Log10 inactivation

(log PFU/mL)

0 1.00 x 107 7.0 0.0

20 1.12 x 106 6.05 0.95

40 6.76 x 104 4.83 2.17

60 1.95 x 104 4.29 2.71

80 4.37 x 103 3.64 3.36

100 1.20 x 103 3.08 3.92

120 7.08 x 101 1.85 5.15

140 1.48 x 101 1.17 5.83

Source: NWRI-WRF (2012) 

Different types of somatic coliphages have different levels of resistance to UV light. For 

example, Lee and Sobsey (2011) estimated the UV inactivation of five types of somatic 

coliphages (T1, T4, T7 ΦX174, λ) representing the four families (Microviridae (ΦX174), 

Myoviridae (T4), Podoviridae (t7) and Siphoviridae (λ, T1)) in laboratory tests using both 

reagent-grade water and surface water. Using regression analysis, the authors predicted the UV 

doses (mJ per cm2) for a 4-log10 inactivation of each of the somatic coliphages to be (in order of 

most to least resistant): 24 for λ, 18 for ΦX174, 12 for T7, 11 for T1, and 4 for T4. Note that 

these doses are lower than what is recommended for treatment plants. Based on these results, the 

authors concluded that different somatic coliphage families can have very different inactivation 

rates and that ΦX174 and λ are the most resistant to UV radiation. In addition, different 

wavelengths have different efficacy at coliphage attenuation. For example, GI F-specific RNA 

coliphage MS2 was three times more sensitive to wavelengths near 214-nm compared to the 

254-nm output of low-pressure lamps in simulated drinking water (Mamane-Gravetz et al., 
75

2005). There is also evidence that laboratory propagated F-specific RNA coliphage MS2 is 

inactivated by UV at a rate that is twice that of indigenous F-specific coliphages (Oppenheimer 

et al., 1993). This highlights the importance of data evaluating indigenous coliphages. 

Nuanualsuwan et al. (2002) evaluated coliphages and other virus inactivation in a phosphatebuffered solution and UV light treatment. The UV dose (mJ/cm2

) required for 1-log10

inactivation was 47.85 for FCV, 36.50 for hepatitis A virus, 24.10 for poliovirus type 1, 23.04 

for GI F-specific RNA coliphage MS2, and 15.48 for somatic coliphage ΦX174 (Nuanualsuwan 

et al., 2002). The coliphages were slightly more sensitive to UV compared to the human viruses. 

In contrast, other studies have found that coliphages are more resistant to UV than human 

viruses. For example, Wiedenmann et al. (1993) found that in a sodium chloride solution, to 

achieve 4-log10 inactivation, a three-times higher UV dose was required for GI F-specific RNA 

coliphage MS2 compared to hepatitis A virus. Havelaar (1987) found that, in secondary effluent, 

F-specific coliphages are more resistant to UV treatment than coxsackievirus, rotavirus, and 

poliovirus. Similarly, Tree et al. (2005) found that UV reduction of F-specific RNA coliphage 

MS2 was less than that of poliovirus and FCV (a surrogate for NoV) in seeded, sterilized 

secondary effluent. To achieve a 4-log10 reduction, doses (mJ per cm2) of 62.5 for GI F-specific 

RNA coliphage MS2, 27.51 for poliovirus, and 19.04 for FCV were required. In bench-scale 

experiments Wilson et al. (1992) found that GI F-specific coliphage MS2 was 1.9 times more 

resistant to UV irradiation that the viruses, rotavirus, poliovirus, and hepatitis A. 

Human adenoviruses are more resistant to UV light than other waterborne (enteric) viruses with 

single and ds RNA genomes. The human adenovirus genome is comprised of dsDNA, which 

affords the virus the ability to use host cell repair enzymes to repair damage in the DNA caused 

by UV light (Hijnen et al., 2006). In contrast, viral genomes that are single stranded DNA cannot 

be repaired in host cells because there is no second strand to serve as a template for replication of 

the nucleic acid. Viral genomes made of RNA are not repaired efficiently because mammalian 

hosts do not have sufficient repair mechanisms for RNA (Eischeid et al., 2011). Thompson et al. 

(2003) conducted a pilot-scale study to examine the effects of UV disinfection on viruses in 

wastewater. In seeded tertiary treated wastewater, 4-log10 inactivation of poliovirus type 1 

required 35 mJ per cm2, GI F-specific RNA coliphage MS2 required 100 mJ per cm2

, and human 

adenovirus (types 15 and 2) required 170 mJ per cm2. In a buffered demand-free water 4-log10

inactivation of FCV required 36 mJ per cm2, GI F-specific RNA coliphage MS2 required 119 mJ 

per cm2, and human adenovirus-40 would have required 226 mJ per cm2(extrapolated value, 4-

log10 reduction was not achieved) (Thurston-Enriquez et al., 2003).

6.5.5 UVA and UVB 

Solar radiation, which consists of UVA/UVB in addition to longer wavelengths, can also be used 

as a disinfection method. Wastewater treatment ponds (see Section 6.3) can be used to treat 

sewage or to further treat secondary effluent. For example, Davies-Colley et al. (2005) 

constructed an outdoor (exposed to solar radiation) advanced pond system to determine the solar 
76

inactivation of somatic and F-specific RNA coliphages and E. coli in secondary effluent.7 Tests 

were conducted in both the summer and winter to determine effects of seasonal variation in solar 

radiation intensity. Somatic coliphages showed a 2.2-log10 reduction in summer and a 0.45-log10

reduction in winter, whereas E. coli had a greater than 4-log10 removal in both seasons.

Reductions of F-specific RNA coliphages were not reported due to low native coliphage 

densities in the influent. The authors found that solar radiation within the UVB range 

(represented by measurements at 311 nm) was responsible for somatic coliphage inactivation, 

whereas F-specific RNA coliphages were hypothesized to be inactivated by both UVA and UVB 

(Davies-Colley et al., 2005). These results are consistent with Sinton et al. (2002), who found 

that under a variety of conditions, F-specific RNA coliphages were inactivated by a wide range 

of wavelengths, whereas somatic coliphages were mainly inactivated by UVB wavelengths (less 

than 318 nm). Davies-Colley et al. (2005) concluded that the advanced pond system is efficient 

at removing coliphages mainly during the summer (or in the tropics), but not in the winter due to 

decreased intensity of solar radiation. 

Gomila et al. (2008) compared inactivation of coliphages and enteric viruses in secondary 

effluent that was treated by UVC radiation (laminar flow through four banks of eight lamps of 

87.5 Watts [W] each) or treated in a sunlit aerobic lagooning system with a residence time of 60 

days. Inactivation of somatic coliphages, F-specific coliphages, and enteroviruses was greater in 

a lagooning system compared to UVC treatment, as shown in Table 22. Using either a UVC 

radiation step in a treatment facility or solar radiation in a lagooning system yielded a greater or 

equal inactivation of coliphages as compared to enteric viruses. Costán-Longares et al. (2008) 

investigated the log10 inactivation of enteroviruses between secondary effluent and different 

types of tertiary treatment at WWTPs in Spain. The authors found a greater than 2-log10

reduction in enteric virus density from secondary treatment after lagooning (Costán-Longares et 

al., 2008). 

Table 22. Log10 reduction in coliphages and enteric viruses in secondary 

effluent after lagooning in sunlight or UVC treatment. 

Microorganism Log10 reduction (lagooning) Log10 reduction (UVC treatment)

Somatic coliphages 0.8 0.5

F-specific coliphages 1.6 0.5

Enteroviruses 0.7 0.5

Source: Gomila et al. (2008)

To compare the effects of UV wavelengths present in sunlight on both coliphages and enteric 

viruses, Lee and Ko (2013) exposed F-specific RNA coliphages MS2 and MNV, to UVA and 

UVB lamps. For all experiments, viruses were suspended in either saline solution or groundwater 

and viral densities were measured by EPA Method 1602 (MS2) or plaque assays (MNV). UVA 

irradiation resulted in a negligible effect on both F-specific RNA coliphages MS2 and MNV 

across the dose range tested (0 to 1500 mJ per cm2). In contrast, MNV was found to be 

significantly more susceptible to UVB than MS2; exposure to 376 mJ per cm2 UVB resulted in a 

7

 Sewage from the Ruakura Research Centre, near Hamilton, New Zealand fed into the advanced pond system. The 

F-specific coliphage level was consistently low in the Ruakura sewage, so primary treated sewage from Hamilton 

City was “spiked” into the pond system. 
77

4-log10 reduction of MNV whereas 909 mJ per cm2 UVB was required for the same reduction of 

F-specific RNA coliphage MS2 (Lee and Ko, 2013). Duizer et al. (2004) found that caliciviruses 

(enteric canine calicivirus no. 48 and respiratory FCV F9) were more susceptible to UVB than 

coliphages as <50 mJ per cm2 UVB resulted in a 4-log10 reduction of both viruses (as determined 

by cell culture) when suspended in buffer. 
78

7. Conclusions 

This review provides background information relevant to the use of coliphages, specifically 

somatic and F-specific coliphages, as indicators of fecal contamination. The following is a 

summary of the major conclusions of this review. 

Methods. Coliphage enumeration methods are adequate for water quality monitoring. EPA 

Method 1601 may be the most useful. Rapid methods are possible and MST methods for 

coliphage are under development. The ability to measure both somatic and F-specific coliphages 

on a single host may be useful, but needs validation (Rose et al., 2004; Guzmán et al., 2008). 

EPA is currently evaluating a membrane filtration culture method and may also evaluate an 

ultrafiltration culture method for use in coliphage enumeration. The intralaboratory (single 

laboratory) method validation study is underway. 

Epidemiological studies. This review summarizes eight epidemiological studies that evaluated 

the relationship of coliphages and gastrointestinal illness from exposure to recreational water. 

Five of the eight studies found a statistically significant relationship between F-specific 

coliphages and illness levels (Lee et al., 1997; Colford et al., 2005, 2007; Wiedenmann et al., 

2006; Wade et al., 2010; Griffith et al., personal communication, 2015). One of the studies found 

a statistically significant increase in RR of GE in bathers when somatic coliphage levels were 

above the NOAEL of 10 PFU per 100 mL (Wiedenmann et al., 2006). 

Occurrence in the Environment. Some studies have reported an association between the 

presence of coliphages and human viruses, while other studies have found no association 

between their presence in environmental waters. The results are strongly influenced by the 

environments in which the studies are conducted. For example, an association between indicators 

and pathogens has more often been reported for brackish and saline water than for freshwater. 

There is evidence that coliphage and F-specific coliphage densities are more strongly associated 

with pathogens than other traditional indicators (E. coli, enterococci, and fecal coliforms). 

Environmental Fate. Coliphages might be reasonable surrogates for enteric viruses in the 

environment. Human viruses and coliphages both decay faster at temperatures above 50°C 

compared to lower temperatures. Human viruses and coliphages both decay faster in sunlight 

than in the dark and are most stable near neutral pH (~6 to 9), but can also survive in lower pH 

environments (i.e., in the gastrointestinal tract of warm-blooded animals). The effect of salinity 

is equivocal and some studies have shown increased and others decreased inactivation in higher 

salinity waters. In fresh, treated, septic, and salt water, predation and environmental factors (e.g., 

temperature, sunlight, pH) have been shown to increase degradation of both coliphages and 

enteric viruses. Organic and inorganic matter affect inactivation—both have been shown to 

increase or decrease decay rates, depending on the virus and the nature of the composition of the 

organic or inorganic matter. There are synergistic or antagonistic interactions between all these 

environmental stressors. 

Wastewater treatment. For primary and secondary treatment, the removal efficiencies of FIB, 

F-specific coliphages, somatic coliphages, and enteric viruses are not substantially different. 

Disinfection is the key step in wastewater treatment for microbial inactivation. Although 
79

disinfection efficacies vary depending on the details of the treatment process, and the 

characteristics of the incoming water at each step, some general conclusions are possible. UVC 

and ozone are most effective at virus inactivation, followed by free chlorine. Combined chlorine 

is not as effective at virus inactivation as the other disinfection treatments. For free chlorine 

(chlorination of nitrified effluents), there are insufficient data to draw conclusions about the 

relative inactivation efficiencies of FIB, indigenous F-specific coliphages, indigenous somatic 

coliphages, and enteric viruses. For combined chlorine (non-breakpoint chlorination of unnitrified effluents), F-specific coliphages and enteric viruses are more resistant to inactivation 

than FIB. In laboratory studies of UVC disinfection, coliphages and enteric viruses are generally 

more resistant to inactivation compared to FIB. However, the inactivation rates of individual 

strains of F-specific and somatic coliphages, as well as enteric viruses, are variable. For example,

adenovirus is highly resistant to UVC. With the exception of ozone, F-specific and somatic 

coliphages overall are likely to be more conservative indicators than FIB in water treated by 

most disinfectants. 

Overall. Table 23 compares coliphage attributes against the currently recommended indicators 

of fecal contamination, E. coli and enterococci. Each indicator/method combination is

summarized and compared against indicator attributes described in Section 1.3. While some of 

the same limitations exist, coliphages are likely a better indicator of viruses in fecal 

contamination than the current FIB (i.e., enterococci and E. coli).
80

Table 23. Attributes of fecal contamination indicators. 

Indicator 

Attribute

Enterococci 

(e.g. EPA Method 1600)

E. coli

(e.g. EPA Method 1603)

Coliphages

(e.g. EPA Method 1602)

Intestinal microflora of warmblooded animalsYes Yes Yes

Present when pathogens are 

present and absent in 

uncontaminated samples

Present when fecal 

pathogens are present, but 

may also be present in 

nonfecally contaminated 

ambient water.

Present when fecal 

pathogens are present, but 

may also be present in 

nonfecally contaminated 

ambient water.

Present when fecal pathogens are 

present, but is likely absent in 

nonfecally contaminated ambient 

water.

Not indicative of viruses in 

WWTP effluent.

Not indicative of viruses 

in WWTP effluent.

Better surrogate for viruses than 

enterococci or E. coli in WWTP 

effluent.

Present in greater numbers 

than the pathogen (in this case,

human viruses)

Depends on sourcea Depends on sourceaIn most cases

Equally resistant as pathogens 

(in this case viruses) to 

environmental factors 

Not as resistant as viruses Not as resistant as viruses Under most conditions

Equally resistant as pathogens 

(in this case viruses) to 

disinfection in water and 

WWTPs

Not as resistant as viruses 

(except for ozone).

Not as resistant as viruses 

(except for ozone).

Under most conditions.

However, adenovirus is more 

resistant than coliphages and 

other enteric viruses to UV 

inactivation.

Should not multiply in the 

environment

Can multiply in the 

environment

Can multiply in the 

environment

Not likely enough to affect 

criteria levels

Detectable by means of easy, 

rapid, and inexpensive 

methods

Yes, but need EPA Method 

1611 for rapid 

enumeration. Other easy 

and rapid methods are 

available.

Yes, but EPA method is 

not considered rapid 

(requires overnight 

incubation). Other easy 

and rapid methods are 

available.

Yes, but Method 1601 needs 

validation for quantification.

Other easy and rapid methods are 

available.

Indicator organism should be 

nonpathogenic

Generally nonpathogenicb Generally nonpathogenic.

c Nonpathogenic

Demonstrated association with 

illness from epidemiological

studies

Yes Yes Yes

Specific to a fecal source or 

identifiable as to source of 

origin

Not EPA Method 1600, but 

MST methods being 

developed.

Not EPA Method 1603, 

but MST methods being 

developed.

Not EPA Method 1602, but MST 

methods being developed.

a

In raw sewage FIB are present in greater numbers than pathogens. Viruses are less vulnerable to treatment processes 

than bacteria, so could survive treatment in greater numbers than bacteria.

b Enterococci can be pathogenic or antibiotic resistant in some settings, like hospitals, but generally not in ambient water.

c Enterohemorrhagic E. coli, specifically O157:H7, grows poorly at 44°C and is often negative for beta-glucuronidase, so 

is not detected by Method 1603 (Degnan and Standridge, 2006). Other pathogenic strains could be detected by EPA

Method 1603.
81

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110

APPENDIX A: Literature Search Strategy and Summary of Literature Search Results 

The literature search strategy consisted of a number of combined approaches. The following 

‘synopsis of information’ and search terms were used to search online databases, including 

PubMed. To supplement these searches, individual authors used free search engines on the 

internet to find articles pertaining to specific information needed. The titles of literature cited in 

specific reports, books, review articles, and conference proceedings were evaluated for 

relevance. 

Synopsis of information gathered during the 2012 literature search: 

 Evaluate the use of bacteriophage as indicators of fecal contamination or wastewater 

treatment efficacy 

 Determine the sources and persistence of bacteriophage in the environment 

 Evaluate the correlation between bacteriophage occurrence and pathogens and traditional 

fecal indicator bacteria (FIB) in wastewater 

 Evaluate properties that affect fate and transport of viruses and bacteriophage 

 Describe the different methods used for detection and analysis of bacteriophage 

concentrations, particularly in recreational waters 

 Evaluate the environmental factors (e.g., organics, temperature, pH, UV/sunlight, 

predation, salinity, porosity, etc.) affecting viral and coliphage degradation 

 Compare degradation in WWTP for bacteriophage and enteric viruses (i.e., primary, 

secondary, and tertiary disinfection) 

Initial Literature Search Strategy Conducted by Professional Librarian 

Database: PubMed 

Dates: 1985-present (Search conducted on July 5, 2012) 

Language: No restrictions 

Retrieve: Titles and year 

Results Format: Microsoft Word; EndNote 

Interested in international and domestic journals and government reports. 

Search terms: 

Set 1: bacteriophage OR coliphage 

AND

Set 2: Water OR illness OR health OR risk 

Search Results from PubMed 

The PubMed search resulted in approximately 2,400 records after removing duplicates. These 

titles were reviewed for relevance based on the outline for the literature review and the synopsis 

above. From the database of titles, 391 articles were sorted as “yes” and 81 were sorted as 

“maybe.” Because this number of titles was still large, 125 “top” articles were selected. The 125 

articles were retrieved by the EPA librarian and sent to the contractor (ICF International). The 
111

125 articles were the starting place for the literature review. Additional targeted literature 

searches were required to obtain more complete information on specific topics. 

HECD Resources 

HECD has developed a robust library of references on waterborne pathogens. These resources 

were included in the resources for this project. Primary authors had access to the HECD library 

of PDFs. 

Supplemental Searches by Primary Authors 

Primary Author 1: 

Search Engine: Google Scholar (http://scholar.google.com/) 

Search terms: Number of Records Considered: 

Bacteriophage AND temperature 20

Bacteriophage AND pH 20

Bacteriophage AND sunlight 20

Bacteriophage AND UVA 10

Bacteriophage AND UVB 10

Bacteriophage AND organic matter 20

Bacteriophage AND sediment 20

Bacteriophage AND predation 20

Bacteriophage AND degradation 20

Bacteriophage AND biofilms 15

Bacteriophage AND morphology AND survival 25

Enterovirus AND temperature 20

Enterovirus AND pH 20

Enterovirus AND sunlight 20

Enterovirus AND UVA 10

Enterovirus AND UVB 10

Enterovirus AND organic matter 20

Enterovirus AND sediment 20

Enterovirus AND predation 20

Enterovirus AND degradation 20

Enterovirus AND biofilms 15

Primary Author 2: 

Date Search 

Engine

Search Terms # of Titles 

Reviewed

# of Articles 

retrieved

9-20-12 PubMed reviews for fecal source tracking 20 2

9-20-12 PubMed reviews for coliphages as viral 

indicators

20 2

9-20-12 PubMed reviews for microbial indicators and 

pathogens

20 2
112

9-20-12 Google coliphage environment fecal 

contamination

20 2

9-20-12 Google coliphage indicator environment 20 2

9-25-12 PubMed Policies and practices for beach 

monitoring

40 2

9-25-12 Google 

Scholar

coliphage detection in fresh water 20 2

9-25-12 Google 

Scholar

coliphage presence in environment 

review

20 2

9-25-12 Google 

Scholar

alternative indicators of fecal 

pollution

20 2

9-26-12 Google 

Scholar

Bacteriophage in the environment 20 4

9-27-12 Google 

Scholar

environmental detection coliphage 40 4

9-27-12 Google 

Scholar

coliphages environment 40 4

9-27-12 Google 

Scholar

fecal source tracking 40 4

10-1-12 PubMed coliphage source tracking 30 2

The document was undergoing internal EPA review and external peer review throughout 2013-

2014. Additional supplemental searches were conducted to address EPA internal and external 

peer reviewer comments. Ultimately, over 2,500 titles were reviewed for inclusion in the 

literature review. There are 342 citations in the final document. 

The Quality Assurance Project Plan includes Information Decision Criteria for selection of cited 

references. The relevant Information Decision Criteria for this project from the Quality 

Assurance Project Plan includes: 

1. More recent references were preferred over older references, unless the older reference 

was particularly notable, important, or widely cited. 

2. Accessibility – References needed to be obtained within project time and budget 

constraints. 

3. English language was required. 

4. Scientific, peer-reviewed publications were preferred, along with others references that 

presented a balanced and objective tone. Government publications such as federal 

regulations, state regulations, standards, permits, guidance documents, and other 

government publications were acceptable. 

5. The reference related to the scope of the information sought. In this case a document 

outline was available. 

6. Geographic relevance – Data collected in the U.S was preferred, but data from other 

countries was also relevant. 
113

7. For a given topic, a literature review citation may have been used in lieu of listing 

numerous primary research articles cited in the literature review. This was done when 

the additional detail provided by the primary citations was not needed. 

8. If a particular point was already in the document and a citation was already provided, 

additional citations backing up this same point were not added. Redundant articles were 

not necessarily cited. 

9. Information provided through personal communication (phone, email) was used only 

when another more widely obtainable source was not available for the same 

information. Information obtained through personal communication needed to be 

highly relevant. 

10. Books citations were acceptable, but sources that could be more easily obtained were 

preferred. Book citations were preferred when the book is an important resource in the 

field. 

11. Newspaper articles were not searched or cited. 

12. Websites were not cited as primary sources of information. URLs are provided for 

some of the references. 

13. Information and references that presented alternative points of view or conclusions to 

the mainstream view were given equal considerations to consensus or majority 

viewpoints. Both alternative and majority viewpoints and conclusions had to provide 

justification based on facts, employ accepted methodologies, and be grounded in the 

scientific method. 
114

APPENDIX B: Coliphage and NoV Densities during Wastewater Treatment 

Table A illustrates how coliphage reduction compares to NoV reduction during wastewater treatment. To be included in Table A, the 

study has to include quantitative norovirus data, quantitative coliphage data, and water samples from raw sewage or effluent. The 

studies are listed in chronological order by year of publication. 

Table A. Coliphage and NoV densities during wastewater treatment. 

Organism Influent (raw) Effluent

Log 

reduction

(log10)

Treatment Location Reference

NoV

(RT-qPCR)

Mean 2 × 105

PCR detectable 

unit per L

NRa 1.8 Non-disinfected activated 

sludge secondary 

treatment (preceded by 

primary and phosphorus 

removal);

No design/operational 

information provided to 

understand secondary 

treatment or performance.

Netherlands Lodder and de 

Roda 

Husman, 2005

F-specific 

bacteriophages

(ISO 10705-1)

Mean 106

PFU per L

NRa 1.6

Somatic 

coliphages

(ISO 10705-2)

Mean 106

PFU per L

NRa 1.1

NoV GI

(RT-qPCR)

0.17–260 copies 

per mL (range)

NR 2.27 ± 0.67 Activated sludge 

secondary treatment and 

chlorination;

No design/operational 

information provided to 

understand secondary 

treatment or performance; 

no information if this is 

free chlorine, combined 

chlorine, ammonia 

concentration.

Tokyo, Japan Haramoto et 

al., 2006

NoV GII

(RT-qPCR)

2.4–1900 copies 

per mL (range)

NR 3.69 ± 1.21

F-specific 

coliphages

(ISO 10705-1)

NR NR 2.81 ± 0.77

NoV

(RT-qPCR)

Mean 3.29 ± 0.26 

(<2.9–3.65) log10 

MPN PCR units 

per L

NR 0.89 ± 0.26 

(0.39->1.3)

Non-disinfected chemical 

precipitation and activated 

sludge (1 plant filtered the 

effluent and 1 plant 

provided additional 

nitrogen removal);

Sweden Ottoson et al., 

2006

F-specific 

coliphages

(ISO 10705-1)

NR NR 1.73 ± 0.59 

(0.74-2.63)
115

Organism Influent (raw) Effluent

Log 

reduction

(log10)

Treatment Location Reference

Somatic 

coliphages

(ISO 10705-2)

NR NR 1.04 ± 0.32 

(0.61-1.86)

No design/operational 

information provided to 

understand secondary 

treatment or performance.

Somatic 

coliphages

(ISO 10705-2)

Mean 2.9 × 106 

(± 2 × 106) PFU 

per mL

Mean 2.5 × 10

4 (± 2.9 × 

104) PFU per mL

Mean 2.16 ± 

0.42

Activated sludge 

secondary treatment and 

chlorination;

No design/operational 

information provided to 

understand secondary 

treatment or performance; 

no information if this is 

free chlorine, combined 

chlorine, ammonia 

concentration.

Italy Carducci et 

al., 2009

Somatic 

coliphages

(EPA Method 

1602)

Mean 1.8 × 105

PFU per 100 mL

Mean 102 PFU per 100 

mL

2.4 Non-disinfected activated 

sludge secondary 

treatment;

No design/operational 

information provided to 

understand secondary 

treatment or performance.

Singapore Aw and Gin, 

2010

F-specific 

coliphages

(EPA Method 

1602)

Mean 4.3 × 104

PFU per mL

Mean 102 PFU per 100 

mL

2.4

NoV GI

(RT- qPCR)

Mean 3.2 × 105

copies per 100 

mL

Mean 7.1 × 103copies 

per 100 mL

~2b

NoV GII

(RT-qPCR)

Mean 2.3 × 105

copies per 100 

mL

Mean 5.2 × 103copies 

per 100 mL

~2b

Somatic 

coliphages (EPA 

Method 1602)

Range 9.1 × 104 

to 4.5 × 105 PFU 

per 100 mL

Range 3 to 63 PFU per 

100 mL

NR Conventional secondary 

treatment with chlorine 

disinfection. No 

design/operational 

information provided to 

understand secondary 

treatment or performance.

Ohio, United 

States

Francy et al. 

2011

F-specific 

coliphages (EPA 

Method 1602)

Range 3.1 × 105 

to 2.2 × 107 PFU 

per 100 mL

Range <1 to 37 PFU 

per 100 mL

NR

NoV GI (qRTPCR)Range 230 cto 

2.2 × 103 GC per 

L

Range <2.7 to 1.8 c GC 

per L

NR
116

Organism Influent (raw) Effluent

Log 

reduction

(log10)

Treatment Location Reference

Somatic 

coliphages (EPA 

Method 1602)

Range 2.4 × 104 

to 3.0 × 106 PFU 

per 100 mL

All <1 PFU per 100 mL NR Conventional tertiary 

treatment (sand filtration) 

with UV disinfection. No 

design/operational 

information provided to 

understand secondary 

treatment or performance.

F-specific 

coliphages (EPA 

Method 1602)

Range 3.8 × 104 

to 2.1 × 105 PFU 

per 100 mL

All <1 PFU per 100 mL NR

NoV GI (qRTPCR)Range <560 to 

<8.3 × 103 GC 

per L

Range <36 to <67 GC 

per L

NR

Somatic 

coliphages (EPA 

Method 1602)

Range 2.6 × 104 

to 2.2 × 106 PFU 

per 100 mL

Range <1 to 1.1 × 103 

PFU per 100 mL

NR Membrane bioreactor 

(Kubota® Membrane 

Systems by Ovivo MBR) 

with UV disinfection. No 

design/operational 

information provided to 

understand secondary 

treatment or performance.

F-specific 

coliphages (EPA 

Method 1602)

Range 3.8 × 104 

to 1.9 × 106 PFU 

per 100 mL

Range <1 to 19 PFU 

per 100 mL

NR

NoV GI (qRTPCR)Range 49 cto 1.8

× 104 GC per L

Range <1.5 to <130 GC 

per L

NR

Somatic 

coliphages

(EPA Method 

1602)

Range 2.6 × 104

to 2.2 × 106 PFU 

per 100 mL

Range <1 to 8 PFU per 

100 mL

NR Two medium-sized 

KubotaTM (Osaka, Japan) 

system microfiltration 

membrane bioreactors 

with UV disinfection. No 

design/operational 

information provided to 

understand secondary 

treatment or performance.

Ohio, United 

States

Francy et al., 

2012

F-specific 

coliphages

(EPA Method 

1602)

Range 3.8 × 104

to 1.9 × 106 PFU 

per 100 mL

Range <1 to 2 PFU per 

100 mL

NR

NoV (qPCR, 

qRT-PCR)

Range 49 cto 1.8 

× 104 GC per L

Range <1.5 to <53 GC 

per L

NR

Somatic 

coliphages

(EPA Method 

1602)

Range 2.4 × 104

to 3.0 × 106 PFU 

per 100 mL

<1 PFU per 100 mL NR One small-sized 

conventional secondary 

plant with tertiary 

treatment and UV 

disinfection. No 

design/operational 

information provided to 

F-specific 

coliphages

(EPA Method 

1602)

Range 1.1× 104

to 2.1 × 105 PFU 

per 100 mL

<1 PFU per 100 mL NR
117

Organism Influent (raw) Effluent

Log 

reduction

(log10)

Treatment Location Reference

NoV (qPCR, 

qRT-PCR)

Range <5.6 × 102

to <8.3 × 103 GC 

per L

Range <36 to <67 GC 

per L

NR understand secondary 

treatment or performance.

Somatic 

coliphages

(EPA Method 

1602)

Range 9.1 × 104

to 4.5 × 105 PFU 

per 100 mL

Range 3 to 63 PFU per 

100 mL

NR One medium-sized 

conventional secondary 

plant with chlorine 

disinfection. No 

design/operational 

information provided to 

understand secondary 

treatment or performance.

F-specific 

coliphages

(EPA Method 

1602)

Range 3.1 × 105

to 2.2 × 107 PFU 

per 100 mL

Range <1 to 37 PFU 

per 100 mL

NR

NoV (qPCR, 

qRT-PCR)

Range 2.3 × 102c

to 1.5 × 103 GC 

per L

Range <1.8 cto <2.7 

GC per L

NR

F-specific RNA 

coliphages

(double agar 

layer)

NR Means 5 × 103to 5 × 

105 PFU per L

NR Secondary treated 

wastewater (2 WWTPs)d

Australia Keegan et al., 

2012

Somatic 

coliphages

(double agar 

layer)

NR Means 9 × 104

to 1.67 × 

105 PFU per L NR

NoV

(RT-qPCR) NR Means ND to 2.7 × 105genomes per L NR Secondary treated wastewater (5 WWTPs)

NoV GI

(RT-qPCR)

Mean 3.32 ± 0.64 

(range 2.05–4.76) 

log10 density

Mean 2.53 ± 0.57 

(range 1.26–4.06) log10 

density

0.80 ± 0.49 Non-disinfected activated 

sludge secondary 

treatment;

No design/operational 

information provided to 

understand secondary 

treatment or performance.

Ireland Flannery et 

al., 2012

NoV GII

(RT-qPCR)

Mean 3.55 ± 0.89 

(range 1.81–5.34) 

log10 density

Mean 2.63 ± 0.71 

(range 1.51–4.08) log10 

density

0.92 ± 0.76

F-specific 

coliphages

(ISO 10705-1)

Mean 5.54 ± 0.51 

(range 3.87–6.82) 

log10 density

Mean 3.41 ± 0.77 

(range 2.00–5.84) log10 

density

2.13 ± 0.76

Somatic 

coliphages (ISO

10705-2)

Mean 7.10 ± 

0.40-log10 PFU 

per L

Mean 4.99 ± 0.53-log10

PFU per L

2.11 ± 0.40 Rainy days: Activated 

sludge, chlorination. No 

design/operational 

Italy Carducci and 

Verani, 2013
118

Organism Influent (raw) Effluent

Log 

reduction

(log10)

Treatment Location Reference

NoV

(RT-qPCR)

Mean 5.83 ± 

2.87-log10 GC 

per L

Mean 5.80 ± 2.75-log10

GC per L

0.02 ± 0.61 information provided to 

understand secondary 

treatment or performance.

Somatic 

coliphages (ISO 

10705-2)

Mean 7.22 ± 

0.40-log10 PFU 

per L

Mean 5.00 ± 0.56-log10

PFU per L

2.21 ± 0.46 Dry days: Activated 

sludge, chlorination. No 

design/operational 

information provided to 

understand secondary 

treatment or performance.

NoV

(RT-qPCR)

Mean 5.92 ± 

2.86-log10 GC 

per L

Mean 6.04 ± 2.94-log10

GC per L

−0.11 ± 0.34

F-specific RNA

coliphages (ISO 

10705-1)

Mean 5.26-log10

PFU per 100 mL

Mean 2.96-log10 PFU 

per 100 mL

NR Screening and grit 

removal, phosphate 

removal through ferric 

sulfate, secondary 

treatment, UV 

disinfection. No 

design/operational 

information provided to 

understand secondary 

treatment or performance.

United States Flannery et al. 

2013

F-specific RNA

coliphage (RTqPCR)

Mean 5.11-log10

GC per 100 mL

Mean 4.57-log10 GC 

per 100 mL

NR

NoV GII (RTqPCR)Mean 3.87-log10

GC per 100 mL

Mean 3.61-log10 GC 

per 100 mL

NR

F-specific RNA

coliphages (ISO 

10705-1)

Range 5.9 × 104

to 7.5 × 105 PFU 

per L

Range 1.5 × 103 to 2.5 × 

104 PFU per L

NR Primary sedimentation. 

Influent data includes 

samples taken when 

treatment was interrupted. 

No design/operational 

information provided to 

understand secondary 

treatment or performance.

Fjellfoten, 

Norway

GrøndahlRosado et al. 

2014

NoV GI (qPCR) Range 2.9 × 103

to 1.4 × 106 GC 

per L

Range 5.6 × 103to 9.2 

× 104 GC per L

NR

NoV GII (qPCR) Range 6.0 × 104

to 1.4 × 107 GC 

per L

Range ND to 2.0 × 105

GC per L

NR

Somatic

coliphages (ISO 

10705-2.2)

Mean 4.9 × 103

PFU per mL

Range <0.01 to 2.5 × 

104 PFU per mL

Range of 

Mean 

Reduction 

0.6 ± 0.6 to 

5.2 ± 1.2

Raw municipal postscreen wastewater influent 

and effluent from three 

pilot-scale sand filters 

with different filter 

material and grain size 

designs and one with a 

separate phosphorous 

removal unit. No 

design/operational 

Kuopio, 

Finland

Kauppinen et 

al. 2014

F-specific

coliphages (ISO 

10705-1)

Mean 3.6 × 103

PFU per mL

Range <0.01 to 1.8 × 

104 PFU per mL

Range of 

Mean 

Reduction 

0.7 ± 0.6 to 

5.3 ± 1.3
119

Organism Influent (raw) Effluent

Log 

reduction

(log10)

Treatment Location Reference

NoV GI (RTPCR)Mean 80 GC per 

mL

Range <0.5 to 1.5 × 103

GC per mL

Range of 

Mean 

Reduction 

0.6 ± 0.6 to 

2.2 ± 0.8

information provided to 

understand treatment or 

performance.

NoV GII (RTPCR)Mean 2.3 × 104

GC per mL

Range <0.4 to 1.1 × 104

GC per mL

Range of 

Mean 

Reduction 

0.5 ± 0.2 to 

4.0 ± 0.6

Density units are as reported in the cited reference.

NR = not reported (in some cases effluent or influent densities were not reported, but log10 reduction was reported); Stm WG49 = Salmonella 

enterica serovar Typhimurium WG49 (host); ND = Not detected; GC = genome copies

a PFU reported in graphical format for influent, digester, high rate pond, algal settling pond, and maturation pond

b Estimated from figure

c Reported as estimated value extrapolated at the low end. PCR threshold cycles were past the upper limit of the standard curve.

d

Information from Table 3.6 and 3.7 in Keegan et al. (2012). The Bolivar WWTP consists of primary treatment (screening, grit removal, 

sedimentation), activated sludge, lagoon (16 day retention), chlorination (or dissolved air flotation and chlorination); collected undisinfected 

samples from the lagoon influent (thus secondary effluent), lagoon effluent, and post dissolved air flotation prior to chlorination. Two Melbourne 

WWTPs were tested: MW1 – primary settling, ASP (not defined); MW2 – anaerobic digestion (not defined), ASP (not defined), and lagoon 

polishing (26 days). This study was done for a very specific focus related to water recycling.