United States EPA/600/R-10/149
Environmental Protection December 2010
Agency
A Study of the Various Parameters
that Affect the Performance of the
New Rapid U.S. Environmental
Protection Agency Quantitative
Polymerase Chain Reaction (qPCR)
Method for Enterococcus Detection
and Comparison with Other Methods
and Pathogens in Treated
Wastewater Mixed with Ambient
Water
RESEARCH AND DEVELOPMENT
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EPA/600/R-10/149
December 2010
A Study of the Various Parameters that Affect
the Performance of the New Rapid U. S.
Environmental Protection Agency Quantitative
Polymerase Chain Reaction (qPCR) Method for
Enterococcus Detection and Comparison with
Other Methods and Pathogens in Treated
Wastewater Mixed with Ambient Water
Kristen P. Brenner1, Kevin Oshima1, Ying Chu2, Larry J. Wymer1,
Richard A. Haugland1 and Eunice Chern1
1 U. S. Environmental Protection Agency
National Exposure Research Laboratory
Cincinnati, Ohio 45268
2 Dynamac Corporation
c/o U. S. Environmental Protection Agency
Cincinnati, Ohio 45268
Contract EP-D-06-096
National Exposure Research Laboratory
Office of Research and Development
U.S. Environmental Protection Agency
Cincinnati, OH 45268
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Disclaimer
Although this work was reviewed by U.S. EPA and approved for publication, it may not
necessarily reflect official Agency policy. Mention of trade names or commercial products does
not constitute endorsement or recommendation for use.
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Executive Summary
The U. S. Environmental Protection Agency's (U.S. EPA's) current recommended criteria for
recreational water quality are based upon culture measurements of Enterococcus fecal indicator
bacteria (FIB). However, a rapid method for monitoring water quality is needed to warn bathers
when FIB densities at public bathing beaches exceed recommended criteria levels. Currently,
warnings to swimmers are being delayed by the time needed to culture the FIB.
Rapid quantitative polymerase chain reaction (qPCR) methods for fecal indicator bacteria are
being considered by the U.S. EPA for beach monitoring and other uses to support new water
quality criteria. Beach epidemiological studies conducted by the U.S. EPA have shown a direct
relationship between densities of Enterococcus determined by qPCR and gastrointestinal illness
rates for both fresh water and marine beaches. However, there is a need for information on how
the fate of the qPCR signal compares with other more traditional culture-based methods that are
currently being used to support water quality criteria. These comparisons are particularly needed
within wastewater treatment plants (WTPs) and when treated effluent mixes with ambient water.
Understanding how well molecular and cultural method results mimic each other and how
pathogens decay in the environment is important in evaluating the applicability of these methods
for establishing water quality criteria under section 304(a)(9) of the Beach Act of 2000 and
addressing all Clean Water Act purposes. For example, further comparative data are needed on
the fates of fecal indicators (by molecular and cultural assessments) and pathogen densities
during wastewater treatment and after treated effluents are mixed with ambient waters. A
comparison of the decay of FIB densities determined by qPCR-based and culture-based methods
in WTPs is of particular interest in the present study because of differences in response to
disinfection between qPCR and culture. Understanding the behavior of qPCR and culture
assessments in the treatment process at WTPs is important, because WTPs represent a significant
source of fecal indicators and pathogens. An understanding of the similarities and differences
between the decay of the qPCR and culture-based signals in WTPs will help determine the
feasibility of using rapid molecular methods to measure treatment efficacy and the impact of
these molecular and culture-based targets of FIB at beaches.
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The purpose of this project was to evaluate and compare the reduction of the Enterococcus qPCR
signal and culture-based FIB during the wastewater treatment process. These same relationships
were also studied in mesocosms where treated effluent was mixed with surface water. The effect
of chlorine and ultraviolet light during disinfection and seasonal effects were also studied. A
small number of pathogens were also studied in the wastewater treatment study and in the
mesocosm studies.
Results from the wastewater treatment component of this project indicated that the reduction of
Enterococcus densities measured by qPCR and culture were similar during primary and
secondary treatment, but were significantly different (p=0.05) during disinfection using either
UV light disinfection or chlorination. The reduction of Enterococcus densities by culture was
significantly greater than the reduction of the qPCR method during disinfection and also during
the complete treatment processes. Similar patterns were observed between the Enterococcus
qPCR and Escherichia coli culture methods. The differences were less pronounced for
Enterococcus qPCR comparisons with F+ male-specific coliphage, Bacteroides, and Clostridium
perfringens culture methods. The effects of UV light and chlorination disinfection processes on
reductions of Enterococcus densities, as determined by qPCR, were similar. No association
between the degradation of Enteroviruses and fecal indicators could be determined, in part,
because of the very low concentrations of Enteroviruses that were detected in the treated
wastewater. Differences in the densities of Giardia cysts and Cryptosporidium oocysts could not
be detected between secondary and disinfected, secondary treated wastewater samples because of
the very low concentrations of both organisms.
Results from the holding studies indicated that, in general, greater reductions of fecal indicator
densities were observed by culture than by Enterococcus qPCR assays in effluents. Reductions
of fecal indicator densities observed by culture and by Enterococcus qPCR were generally more
consistent when holding effluents in the presence of ambient surface waters than when holding
effluents alone. For all holding studies, the initial densities of Enterococcus determined by qPCR
were generally several orders of magnitude higher than the corresponding densities of culturable
Enterococcus, E. coli, and F+ male-specific coliphages, except in the winter samples. For all of
in
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the holding studies, reductions of fecal indicator densities were lowest in the winter. Reductions
of spiked, attenuated polioviruses in wastewater effluent from Ohio River holding studies were
similar to those of Enterococcus determined by both the qPCR and culture methods.
IV
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Table of Contents
Disclaimer i
Executive Summary ii
Table of Contents v
Introduction 1
Material and Methods 4
Water Sample Locations 4
Mesocosm Studies 4
Study Design 5
Dry Run 5
Design of the Main Study 5
Part A 6
PartB 6
PartC 7
PartD 7
Sample Collection 9
Quality Assurance/Quality Control 11
Analytical Methods 12
Standard membrane filter method for Enterococci 12
Standard membrane filter method for Escherichia coli 13
Quantitative polymerase chain reaction (qPCR) method 13
Standard method for male-specific (F+) coliphage 14
Enumeration of Bacteroides and Clostridium perfringens by membrane filtration 15
Enterovirus plaque assay 15
Enterovirus cytopathic effect (CPE) most-probable-number (MPN) assay combined with
a reverse transcriptase polymerase chain reaction (RT-PCR) assay 16
Cryptosporidium and Giardia detection by U.S. EPA Method 1623 17
Cryptosporidium oocyst infectivity culture method 18
Ancillary measurements 18
Photographic data 18
Data Analysis 18
Results 19
Comparison of Enterococcus Densities Measured by qPCR and the Densities of Fecal
Indicators Measured by Cultural Methods through the Wastewater Treatment Process 19
Densities of Enterovirus, Cryptosporidium and Giardia Before and After Disinfection 21
Effluent Holding Studies 22
5% Disinfected Secondary Effluent-Ohio River Mesocosm Studies 22
20% Disinfected Secondary Effluent-Diluted Ohio River Mesocosm Studies 23
5% Disinfected Secondary Effluent-Ohio River Mesocosm Studies with Spiked Attenuated
Poliovirus 23
Discussion 24
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Wastewater Treatment Studies 24
Indicator Persistence Studies 28
Conclusions 31
Wastewater Treatment 31
Holding Studies 32
References 32
Tables 43
Figures 49
Appendix A-l
VI
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Introduction
The U. S. Environmental Protection Agency's (U.S. EPA) current recommended criteria for
recreational water quality are culture-based measurements of Enterococcus fecal indicator
bacteria (FIB; U.S. EPA 1986). However, a rapid method for monitoring water quality is needed
to warn bathers when FIB densities at public bathing beaches exceed recommended criteria
levels. Currently, warnings to swimmers are being delayed by the time needed to culture the
FIB. These delays are 24-32 hr after the collection of the sample. Rapid Quantitative
Polymerase Chain Reaction (qPCR) methods for fecal indicator bacteria are being considered by
the U.S. EPA for beach monitoring and other uses to support new water quality criteria.
A considerable amount of effort has been made in developing and characterizing the
performance of qPCR-based methods to detect and quantify FIB in recreational waters
(Haugland et al. 2005, Siefring et al. 2008, Chern et al. 2009, U.S. EPA, 2010) and to determine
health relationships at bathing beaches. Furthermore, the U.S. EPA has conducted a series of
beach epidemiological studies to assess the relationship between densities of FIB as determined
by qPCR and illness rates (Wade et al. 2006, Wade et al. 2008, Wade et al., 2010). These studies
have shown a direct relationship between gastrointestinal illness rates and qPCR-based densities
of Enterococcus for both fresh water and marine beaches. However, there is a need for
information on how the fate of the qPCR signal compares with other more traditional culture-
based methods that are currently being used to support water quality criteria. These comparisons
are particularly needed within wastewater treatment plants (WTPs) and when treated effluent
mixes with ambient water.
This project was initiated (1) to examine the relationship between the qPCR method and the
traditional indicator methods at wastewater treatment plants and in ambient water mixed with
disinfected, secondary effluents and (2) to collect information so that the Office of Water can
determine if the qPCR method can be used for Total Maximum Daily Limits (TMDLs) and
National Pollutant Discharge Elimination System (NPDES) Permits under the Clean Water Act.
Currently, there is little direct evidence comparing qPCR-based and culture-based FIB
quantification results as predictors of pathogen levels in wastewater or ambient water.
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Understanding how well molecular and cultural method results mimic each other and the decay
of pathogens in the environment is important in evaluating the applicability of these methods for
establishing water quality criteria under section 304(a)(9) of the Beach Act of 2000 and whether
they can be used for all Clean Water Act purposes. For example, further comparative data are
needed on the fates of fecal indicators (by molecular and cultural assessments) and pathogen
densities during wastewater treatment and after treated effluents are mixed with ambient waters.
The decay of fecal indicators has been studied extensively using culture methods for FIB. It is
known that temperature, sunlight, disinfection, predation, and salinity can effect the detection of
culturable bacterial cells (Anderson et al. 2005, Arnone and Walling 2007, Craig et al. 2004,
Harwood et al. 2009, Maiga et al. 2009, Muela et al. 2000, Scheuerman et al. 1988, Sinton et al.
2007). Other factors, such as particulates, have been shown to enhance the survival of fecal
indicators (Arnone and Walling 2007, Craig et al. 2004, Garcia-Armisen and Servais 2009, Lee
et al. 2006, Pote et al. 2009). There is less known about the decay of molecular-based markers of
FIB compared with culture and pathogens, although some studies have been reported recently
(Dick et al. 2010, Lavender and Kinzelman 2009, Shannon et al. 2007, Wery et al. 2008). A
major difference between the qPCR method and cultural methods is the fact that the qPCR
method measures DNA from both live and dead cells, as well as extracellular DNA in water or
wastewater, and culture methods measure only viable cells. A comparison of the decay of FIB
densities determined by qPCR-based and culture-based methods in WTP is of particular interest
in the present study because of differences in response to disinfection between qPCR and culture.
Understanding the behavior of qPCR and culture assessments in the treatment process at WTPs
is important because WTPs represent a significant source of fecal indicators and pathogens. A
better understanding of the comparison of the decay of the qPCR and culture-based signals in
WTPs will help determine the feasibility of using rapid molecular methods to measure treatment
efficacy and the impact of molecular and culture-based targets of FIB at beaches.
The objectives of the study described here were the following:
1) Evaluate and compare the reduction of Enterococcus qPCR signal and culture-based FIB
during the wastewater treatment process.
a) Between seasons.
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b) Between wastewater effluents disinfected by ultraviolet light (UV) and chlorine.
c) Compare indicators with selected pathogens, such as enteroviruses, Giardia, and
Cryptosporidium.
2) Evaluate and compare the reduction of Enterococcus qPCR signal and culture-based FIB over
time in mesocosms of treated WTP effluents.
a) Between seasons.
b) Between wastewater effluent disinfected by UV light and chlorine.
3) Evaluate and compare the reduction of Enterococcus qPCR signal and culture-based FIB in
mesocosms of WTP effluent samples mixed with ambient water collected from the Ohio River.
a) Between seasons.
b) Between wastewater effluent disinfected by UV light or chlorine.
c) Compare indicators with selected pathogens, such as enteroviruses, Giardia, and
Cryptosporidium.
A number of FIB, b acted ophage, and selected pathogens were measured in order to compare
densities of organisms determined by qPCR and culture methods. There is little information of
this kind reported in the literature particularly within WTPs. The results from this study will
help interpret qPCR-based estimates of densities of FIB at beaches impacted by sewage
treatment plants by characterizing how treatment processes at these plants affect the densities of
these organisms during and after treatment, as well as the environmental persistence of these
organisms after they are mixed with ambient receiving waters.
This information will be used to help interpret the impact of measurements of molecular and
culturable indicators in wastewater effluents on recreational waters and to evaluate the
applicability of these alternative methods for establishing water quality criteria under 304(a)(9)
of the Beach Act of 2000 and whether they can be used for all Clean Water Act purposes.
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Material and Methods
Water Sample Locations
Ambient Ohio River water samples were collected at the Greater Cincinnati Water Works,
California pumping station sample intake east of Cincinnati, Ohio. Partially-treated drinking
water samples (Ohio River source water) were also collected at the Greater Cincinnati Water
Works after sand filtration, but before activated carbon treatment. Wastewater samples were
collected from the following Hamilton County Metropolitan Sewer District wastewater treatment
plants (WTP) in Cincinnati, Ohio: Mill Creek WTP (MC; 52,526 MGD capacity), activated
sludge treatment with chlorine disinfection; Little Miami WTP (LM; 9,742 MGD capacity),
activated sludge treatment with chlorine disinfection; Muddy Creek WTP (MD; 5,371 MGD
capacity), activated sludge treatment with UV disinfection; and Polk Run WTP (PR; 1,810 MGD
capacity), activated sludge treatment with UV disinfection except in winter. The wastewater raw
influent was primarily domestic sewage, but Mill Creek WTP had some industrial waste input as
well. Samples were collected at four different locations at each WTP:
• raw sewage influent;
• primary effluent;
• secondary effluent before disinfection; and
• secondary effluent after disinfection by either chlorination or UV disinfection, but before
the effluent was discharged to the receiving streams.
Schematic diagrams of each WTP showing the sample collection sites can be found in the
Appendix (Figures A1-A4).
Mesocosm Studies
Mesocosm studies were done to characterize the degradation of qPCR and culture-based
quantification of indicators and enterovirus over a 6-day period. The following mesocosms were
used:
• secondary effluent after disinfection but before discharge into ambient water;
• 5% secondary effluent after disinfection mixed with 95% Ohio River water; and
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• 20% secondary effluent after disinfection mixed with 60% partially-treated drinking
water and 20% Ohio River water.
Study Design
The field and lab studies were conducted during 2009 by the U.S. EPA contractor, TetraTech,
Fairfax, Virginia (U.S. EPA Contract Number EP-C-08-004, Task Order 2008-026) and their
sub-contractors: Tetra Tech-Clancy Environmental (formerly Clancy Environmental Inc), Saint
Albans, Vermont (Field sampling and most of the laboratory analyses); EMSL Analytical, Inc.,
Cinnaminson, New Jersey (qPCR analyses); BioVir Laboratories, Inc., Benicia, California
(Enterovirus analyses), and the laboratory of Dr. Kellogg Schwab at John Hopkins Bloomberg
School of Public Health, Baltimore, Maryland (Enterovirus reverse transcriptase-polymerase
chain reaction [RT-PCR] method).
Dry Run
A preliminary dry run was conducted on March 23-26, 2009. The "dry run" was a preliminary
sampling visit to two of the WTPs (one using chlorination and one using ultraviolet light
disinfection), instead of the four WTPs required in the rest of the study, to allow the contractor
sampling and laboratory personnel to go through the entire study procedure, observed by U.S.
EPA and Contractor management personnel, but with a reduced analytical load. The purpose of
the dry run was to answer questions (if any), observe all activities in detail, and see if changes in
procedure or improvements in logistics were needed before the major part of the study began.
No QA/QC issues were identified in the dry run analysis and, thus, the dry run data were
included in the final data set.
Design of the Main Study
The research study was divided into four parts (Parts A, B, C, and D), as shown in the schematic
in Figures 1-3. The spring, summer, and winter seasonal sampling visits were conducted on
May 27, 2009-June 5, 2009; September 8-16, 2009; and December 8-16, 2009, respectively.
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Part A
The purpose of Part A was to determine the die-off of the indicator microorganisms and the
decay of the qPCR signal, expressed in cell equivalents (CE; Haugland et al. 2005), through the
treatment processes of the WTPs. Part A also compared performance of multiple methods with
the two different types of disinfection (chlorination and UV disinfection). In Part A of the study
(see Figures 1-3), wastewater samples were collected at all four of the different locations at each
facility.
Chlorinated, secondary effluents for this part of the study were treated with sodium thiosulfate (1
ml of a sterile 10% solution per L of wastewater sample) after collection. The samples were
collected during a dry run at Mill Creek WTP and Muddy Creek WTP and during three different
visits to each of the four WTPs, corresponding to three different seasons: spring, summer, and
winter. Each of the samples in Part A was analyzed by six different methods:
• New, rapid Enterococcus Quantitative Polymerase Chain Reaction (qPCR) Method
(Haugland et al. 2005) and
• Five fecal indicator cultural methods: Enterococcus membrane filter (MF) method (U.S.
EPA 2002a), Escherichia coli MF method (U.S. EPA 2002b), E. coli F+ male-specific
coliphage method (U.S. EPA 2001a), Bacteroides MF method (Livingston et al. 1978),
and Clostridiumperfringens MF method (Fout et al. 1996).
Part B
Part B of the study extended the evaluations in Part A to several pathogens. For this part of the
study (see Figures 1-3), the secondary effluent samples collected before and after disinfection
were also analyzed using the following pathogen methods:
• Enterovirus plaque assay (U.S. EPA 1987) using the continuous Buffalo Green Monkey
Kidney (BGM) cell line (maintained at the BioVir Laboratory, Benicia, California);
• Enterovirus cytopathic effect (CPE) most-probable number (MPN) method (U.S. EPA
2001) combined with an Enterovirus RT-PCR method (Gregory et al. 2006);
• U.S. EPA Method 1623 for both Giardia and Cryptosporidium (U.S. EPA 2005b); and
• Cryptosporidium infective oocyst cultural method (Johnson et al. 2010).
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PartC
In Part C (see Figures 1-3), disinfected, secondary effluent die-off studies were conducted. An
extra 20-L portion of each disinfected, secondary effluent sample from two WTPs for the
preliminary dry run (Mill Creek WTP and Muddy Creek WTP) and from each of the four WTPs
during each seasonal visit was stored at the local analytical laboratory at the seasonal
temperatures at which they were collected for an additional six days after the initial tests in Parts
A, which were designated as Day 0, and analyzed on Days 1, 2, 4, and 6 by the first four
methods used in Part A (qPCR and U.S. EPA Methods 1600, 1602, and 1603). The seasonal
temperatures used for the spring (April and May), summer (June-September), and winter
(December-March) visits were 15-17 °C, 20-23 °C, and 4-8 °C, respectively. The months and
temperatures for each season were chosen based on the period of representative seasonal (spring,
summer, and winter) ambient water temperatures for the Ohio River that were collected by the
U.S. EPA microbiology laboratory in Cincinnati, Ohio over a period of several years.
The secondary effluent samples disinfected by chlorine (Mill Creek and Little Miami WTPs) did
not receive sodium thiosulfate during collection, but sodium thiosulfate was added when the
individual samples were removed from the carboy for analysis on days 0, 1,2, 4, and 6. Samples
were covered to keep them completely in the dark and mixed twice daily, once in the morning
and once in the late afternoon, to simulate samples with little or no exposure to UV light. This
part of the study was used to determine the die-off of the microorganisms and the decay of the
qPCR signal in the stored effluents after treatment, but before effluent discharge.
PartD
In Part D of the study (see Figures 1-3), two simulated recreational water sample die-off studies
were conducted. In the first study, 2.5 L of disinfected, secondary effluent from two WTPs
during the dry run (Mill Creek and Muddy Creek WTPs) and from each of the four WTPs during
each of the three seasonal visits was mixed with 47.5 L of an Ohio River sample, collected the
day the wastewater sample was collected, to produce a simulated recreational water sample with
a final concentration of 5% wastewater. Attenuated poliovirus was added [final concentration of
approximately 1000 Plaque-Forming-Units (PFU) per ml] to boost the virus concentrations in the
simulated recreational water samples to detectable levels in order to measure the die-off of the
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viruses. Each sample was stored at the local analytical laboratory at the seasonal temperatures at
which they were collected for an additional six days and analyzed on Days 0, 1,2, 4, and 6 by
the first four methods used in Part A (qPCR and U.S. EPA Methods 1600, 1602, and 1603) and
by the three Enterovirus methods of Part B. The seasonal temperatures used for the spring (April
and May), summer (June-September), and winter (December-March) visits were 15-17 °C, 20-
23 °C, and 4-8 °C, respectively.
Chlorine in the disinfected, secondary effluent samples from the WTPs that used chlorination
(Mill Creek and Little Miami WTPs) was neutralized by sodium thiosulfate during the initial
collection procedure. Samples were covered to protect them from light and mixed twice daily,
once in the morning and once in the late afternoon, to simulate samples with little or no exposure
to ultraviolet light. One (1) L of each of the spiked simulated recreational water samples for
each analysis day was sent to the virus analytical laboratory for processing and analysis. The
purpose of this part of the study was to determine the fate of the microorganisms, the viruses,
and the qPCR signal in the ambient water after wastewater treatment and effluent discharge into
the receiving body of water, in this case, the Ohio River water.
In the second die-off and degradation study, a sample containing 20% disinfected, secondary
effluent (10 L), 20% Ohio River water (10 L), and 60% partially-treated drinking water (30 L;
source water was the Ohio River) was prepared and stored at the local analytical laboratory at the
seasonal temperatures at which they were collected for an additional six days and analyzed on
Days 0, 1, 2, 4, and 6 by the first four methods used in Part A (qPCR and U.S. EPA Methods
1600, 1602, and 1603). The seasonal temperatures used for the spring (April and May), summer
(June-September), and winter (December-March) visits were 15-17 °C, 20-23 °C, and 4-8 °C,
respectively.
Chlorine in the disinfected, secondary effluent samples from the WTPs that used chlorination
(Mill Creek and Little Miami WTPs) was neutralized by sodium thiosulfate during the initial
collection procedure. Samples were covered to protect them from light and mixed twice daily,
once in the morning and once in the late afternoon, to simulate samples with little or no exposure
to UV light. This allowed for the maximum survival of the microorganisms, thereby presenting
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a worst case scenario for microbial die-off and degradation of the qPCR signal (Arnone and
Walling 2007, Scheuerman et al. 1988, Sinton et al. 2007). The purpose of this part of the study
was to determine the fate of the microorganisms and the qPCR signal in a sample where the
majority of the DNA measured by the qPCR method came from the wastewater Enterococci,
while retaining some of the natural predators from the Ohio River in the sample.
In addition, a non-spiked portion of each Ohio River water sample was collected and stored at
the local analytical laboratory at the seasonal temperatures at which they were collected for an
additional six days and analyzed on Days 0, 1,2, 4, and 6 by the first four methods used in Part
A (qPCR and U.S. EPA Methods 1600, 1602, and 1603) and by the Enterovirus Plaque Assay on
Day 0 as a control. The seasonal temperatures used for the spring (April and May), summer
(June-September), and winter visits (December-March) were 15-17 °C, 20-23 °C, and 4-8 °C,
respectively. Samples were covered to protect them from light and mixed twice daily, once in
the morning and once in the late afternoon, to simulate samples with little or no exposure to UV
light. This allowed for the maximum survival of the microorganisms, thereby presenting a worst
case scenario for microbial die-off and degradation of the qPCR signal (Arnone and Walling
2007, Scheuerman et al. 1988, Sinton et al. 2007). Partially-treated drinking water samples were
also analyzed by the same methods on Day 0 as a control, and a virus titer of the added
poliovirus was made at the time the simulated recreational water was spiked.
Sample Collection
Each sample had a unique identification number that included the date (month, day, and year),
actual time of collection (in military time), the study part (A, B, C, or D), sample visit (dry run,
spring, summer, or winter), WTP (MC, MD, LM, and PR), sampling location within the WTP,
method(s) to be used for the sample, storage day of the holding studies, and volume or dilution
of sample analyzed. Microbiological analysis of water samples (Haugland et al. 2005, U.S. EPA
2001, U.S. EPA 2002a, 2002b, Livingston et al. 1978, Fout et al. 1996) for Enterococci (qPCR
and Method 1600), Escherichia coli, Bacteroides fragilis group, Clostridium perfringens, and F+
male-specific coliphage, respectively, began within six hours of collection, the holding time for
wastewater and recreational water, and the analyses were completed within eight hours of
sampling (Bordner et al. 1978, CFR 1999).
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Raw sewage influent and primary effluent samples for Part A were collected aseptically using
sterile, polypropylene 500-ml or 1000-ml bottles at the locations designated by the personnel at
each WTP. Pre-sterilized sample bottles were purchased for use in this study, and the sterility of
a few randomly-chosen bottles from each lot were tested before field use by adding sterile
Trypticase Soy Broth to the bottles, incubating for 48-72 hours at 35 °C, and observing the
bottles for bacterial growth (turbidity). Samples were taken from a faucet, when available; about
one foot (0.3 m) under the surface of the wastewater; at an indoor trough; or at an effluent
overflow, and the bottles were filled, allowing approximately one inch of head space for
subsequent mixing. Faucets and troughs were flushed for 3-5 minutes to remove water in the
lines or standing water, respectively, before collecting the samples. Pumps with sterile,
replaceable tubing or polypropylene dippers with sterile containers were used, where
appropriate.
Pumps, large mixing tanks, and specially-designed sampling manifolds were used to
simultaneously collect water samples and the large volumes of water needed for the virus and
parasite filters. River water, partially-treated drinking water, secondary effluent, and secondary,
disinfected effluent samples, were each collected by the detailed protocols found in the
Appendix. Chlorine residuals in the containers of disinfected, secondary effluents from Mill
Creek and Little Miami WTPs were determined using a HACK CN-66 chlorine test kit
(Loveland, Colorado). Sample chlorine, if present, was neutralized by adding 1 ml of sterile
10% sodium thiosulfate solution per L of sample as soon as possible thereafter (except for the
disinfected, secondary effluent samples for the effluent holding studies in Part C), and the
samples were mixed thoroughly. A second total chlorine determination was made on each
sample treated with sodium thiosulfate to confirm the absence of chlorine. All samples were
stored on ice after collection and during transit to the laboratory for logging, distribution, and/or
packing and shipping and at 1-4 °C until the time of analysis.
Cartridge filters for the Enterovirus, Giardia, and Cryptosporidium analyses were placed inside
two sterile plastic bags after field filtration and taken to or shipped to the appropriate analytical
laboratory on wet ice or cold packs. Temperature-tracking devices, iButtons (Maxim Integrated
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Products, Sunnyvale, California and Dallas Semiconductor iButton Product Group, Dallas, Texas
75244), were placed in small ziplock bags and included in all shipping packages (but not
directly next to the cold packs or ice) to ensure that the pathogen filters were not frozen or
exposed to high temperatures en route to the analytical laboratories. Samples for virus and
parasite analysis were shipped daily during the dry run and each seasonal visit. All frozen qPCR
filters for each sampling visit were sent by overnight delivery on dry ice in a single shipment to
the qPCR analytical laboratory. The date and time of arrival of various samples at the analytical
laboratories and the time the packages are opened were recorded at the laboratories.
Examination of the iButton records showed that none of the virus or parasite filters were frozen
or exposed to high temperatures during shipping and that all qPCR filters remained frozen during
transit to the analytical laboratory.
Quality Assurance/Quality Control
The contract laboratories used standard good laboratory practice and Quality Assurance/Quality
Control (QA/QC) procedures in this study, as described in the U.S. EPA Microbiology Methods
Manual, Part IV, C (Bordner et al. 1978); Section 9000 of the 20th edition of Standard Methods
(Clesceri et al. 1998); the QC section of the U.S. EP A's Manual for the Certification of
Laboratories Analyzing Drinking Water (U.S. EPA 2005a); U.S. EPA Manual of Methods for
Virology (U.S. EPA 1987); Quality Assurance/Quality Control Guidance for Laboratories
PerformingPCR Analyses on Environmental Samples (U.S. EPA 2004); the QA Project Plan
(QAPP), the individual method protocols, and the instructions and QA recommendations of the
instrument manufacturers. Appropriate field blanks; supplies, media, and reagent sterility tests;
positive and negative controls for each of the microbial methods; and matrix spikes were
performed and documented. Known concentrations of calibrator cells were added to the qPCR
tests to establish the number of cell equivalents in the qPCR signal detected by the test
(Haugland et al. 2005). ColorSeed™ C & G (BTF Pty Ltd., Sydney, Australia) for Giardia and
for Cryptosporidium internal standards were used according to the manufacturer's instructions as
a post-filtration positive control for both organisms (Francy et al. 2004). iButtons (Maxim
Integrated Products, Inc., Sunnyvale, CA) were included in the shipping containers to determine
whether the qPCR filters remained frozen and to find out whether the parasite cartridge filters
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were exposed to elevated or freezing temperatures during transit to the remote analytical
laboratories.
The procedures at the local laboratory in Cincinnati, Ohio and sample collectors in the field were
observed by U.S. EPA task order manager, Kristen Brenner, on several occasions, including the
preliminary dry run. The "dry run" was a preliminary sampling visit to two of the WTPs (one
using chlorination and one using ultraviolet light disinfection), instead of the four WTPs required
in the rest of the study, to allow the contractor sampling and laboratory personnel to go through
the entire study procedure, observed by U.S. EPA and Contractor management personnel, but
with a reduced analytical load. The purpose of the dry run was to answer questions (if any),
observe all activities in detail, and see if changes in procedure or improvements in logistics were
needed before the major part of the study began. No QA/QC issues were identified in the dry
run analysis and, thus, the dry run data were included in the final data set. In addition, the U.S.
EPA, NERL-Cincinnati Quality Assurance Officer, Margie Vazquez, conducted a formal QA
audit of the local Laboratory and field operations on September 8-10, 2009 and a formal QA
audit of the EMSL (Cinnaminson, NJ) qPCR laboratory was conducted on October 7, 2009 by
the U.S. EPA NHEERL-RTP Quality Assurance Officer, Michael Ray, and the contractor QA
Officer, Trisha Johnson, TetraTech-Clancy Environmental.
Analytical Methods
Standard membrane filter method for Enterococci
U.S. EPA Method 1600 (Messer and Dufour 1998, U.S. EPA 2000, U.S. EPA 2002a), the U.S.
EPA-approved culture method for monitoring wastewater and recreational water, was used to
determine the Enterococcus concentrations of each of the various water samples in this study.
One 500-ml water sample of each type of wastewater at each WTP for each seasonal visit,
including the dry run, was collected, filtered through cellulose nitrate or mixed cellulose ester,
0.45-um pore size MFs, and analyzed for Enterococci using 3-5 volumes (or dilutions in
phosphate-buffered dilution water) of each sample (Bordner et al. 1978, Clesceri et al. 1998).
Additional volumes or dilutions were performed during the dry run to determine the appropriate
volume and/or dilution range for each type of wastewater or water sample. Analysis of each
sample began within six hours of its collection, and processing (filtration and plating) was
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completed no later than eight hours after collection. Analysis start time and the time and date
incubation began and ended were recorded for all samples. Media were checked for sterility, and
for positive and negative reactions, and filter and buffer controls were performed. Verification
tests (U.S. EPA 2000, U.S. EPA 2002) were performed for all water samples (5 colonies/sample)
from the first trip to each WTP. Results for this method are expressed in colony-forming-units
per 100ml(CFU/100ml).
Standard membrane filter method for Escherlchla coll
U.S. EPA Method 1603 (U.S. EPA 2000, U.S. EPA 2002b), the U.S. EPA-approved culture
method for monitoring wastewater and recreational water, was used to determine the Escherichia
coli concentrations of the various water samples in this study. One 500-ml water sample of each
type of wastewater at each WTP for each seasonal visit, including the dry run, was collected,
filtered through cellulose nitrate or mixed cellulose ester, 0.45-um pore size MFs, and analyzed
for E. coli using 3-5 volumes (or dilutions in phosphate-buffered dilution water; Bordner et al.
1978, Clesceri et al. 1998) of each sample. Additional volumes or dilutions were performed
during the dry run to determine the appropriate volume and/or dilution range for each type of
wastewater or water sample. Analysis of each sample began within six hours of its collection,
and processing (filtration and plating) was completed no later than eight hours after collection.
Analysis start time and the time and date incubation began and ended were recorded for all
samples. Media were checked for sterility, and for positive and negative reactions, and filter and
buffer controls were performed. Verification tests (U.S. EPA 2000, U.S. EPA 2002) were
performed for all water samples (5 colonies/sample) from the first trip to each WTP. Results for
this method are expressed in Colony-Forming-Units per 100 ml (CFU/100 ml).
Quantitative polymerase chain reaction (qPCR) method
The qPCR method (Haugland et al. 2005) describes the procedures for the detection of
Enterococci in water samples based on the collection of these organisms on MFs, extraction of
their total DNA, and PCR amplification of a genus-specific DNA sequence using the TaqMan™
PCR product detection system (see Figure A5 in the Appendix). The reactions were performed
in a specially-designed thermal cycling instrument (Cepheid Smart Cycler) that automates the
detection and quantitative measurement of the fluorescent signals produced by probe degradation
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during each cycle of amplification. The analyst at the qPCR laboratory received training in the
laboratory of Dr. Richard Haugland, the developer of the method, at the U.S. EPA in Cincinnati,
Ohio. Results for this method are expressed in Calibrator Cell Equivalents per 100 ml (CE/100
ml; Haugland et al. 2005). Calibrator cell equivalents are a measure of the qPCR signal density,
based on the qPCR signal generated by added Enterococcus calibrator cells where the number of
cells is known (Haugland et al. 2005).
A 1-L water sample of each type of wastewater at each WTP for each seasonal visit, including
the dry run, was collected for use in this method. All collected samples were analyzed for
Enterococci using sample volumes of 100 ml for the filters, except in special circumstances, such
as high turbidity or total suspended solids (TSS), which could clog filters and require smaller
volumes of sample. Five (5) replicate filtrations on 0.4-um polycarbonate filters were performed
for each sample, and the filters were transferred to extraction tubes, as described in the protocol
(Haugland et al. 2005), and stored at -80 °C until shipped to the qPCR analytical laboratory.
Filtration of each sample was initiated within six hours of its collection, and the filters were
stored in the freezer within eight hours of collection. Only one of the filters (Haugland et al.
2005) was extracted and analyzed, while the remaining four filters were stored in the freezer at -
80 °C as backups or for other/later analyses. All of the sample qPCR filters from a seasonal visit
were shipped together by overnight express on dry ice to EMSL Analytical (Westmont, New
Jersey) for analysis. iButtons were included in the shipping container to determine whether the
filters remained frozen during shipping. Specific QC requirements can be found in the method
or the PCR Quality Assurance Manual (U.S. EPA 2004; www.epa.gov/nerlcwww/qa_qc_pcr 10
_04.pdf).
Standard method for male-specific (F*) coliphage
U.S. EPA Method 1602 (U.S. EPA 2001), was used to determine the concentrations of male-
specific (F+) coliphage in the samples. The somatic coliphage portion of this test was not used in
this study. One (1)-L water samples of each type of wastewater at each WTP for each seasonal
visit, including the dry run, were collected and analyzed by the single agar layer method for
coliphage using sample volumes of 100, 10, and 1 ml, and dilutions of the samples in phosphate-
buffered dilution water (Bordner et al. 1978, Clesceri et al. 1998) were analyzed by the double
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agar layer method when needed. Analysis of each sample was initiated within six hours of its
collection, and processing (filtration and plating) was completed no later than eight hours after
collection. Specific QC requirements can be found in the method (U.S. EPA 2001). Positive and
negative controls were analyzed with each group of samples. Results for this method are
expressed in plaque-forming-units per ml (PFU/ml).
Enumeration of Bacteroides and Clostridium perfringens by membrane filtration
Each of the four types of wastewater from each of the WTPs, collected during the dry run and
the three seasonal visits, were filtered through cellulose nitrate or mixed cellulose ester, 0.45-um
pore size MFs, and analyzed for the Bacteroides fragilis group (Livingston et al. 1978) and for
Clostridium perfringens (Fout et al. 1996) using Bacteroides Bile Esculin Agar (BBE),
supplemented with 0.1 g of gentamicin after autoclaving (HEVIedia Laboratories, LTD), and
mCP agar, respectively. After anaerobically incubating the Bacteroides filters in a GasPak
Chamber at 36 °C for 18-48 hours (Livingston et al. 1978), grayish colonies surrounded by
blackening of the medium were counted and recorded. Clostridium perfringens filters were
incubated anaerobically for 24 hours in a GasPak Chamber at 44.5 °C, and straw yellow colonies
were counted (only total counts were made) and recorded. Results for these methods are
expressed in Colony-Forming-Units per 100 ml (CFU/100 ml).
Enterovims plaque assay
Secondary effluent samples before and after disinfection, the virus spike titrations, the Ohio
River water control samples, and the 5% wastewater in Ohio River water die-off study samples
in Part D were analyzed for Enterovirus by the Plaque Assay (U.S. EPA 1987), using the
continuous Buffalo Green Monkey (BGM) kidney cell line (as maintained at the BioVir
Laboratory, Benicia, California). Volumes of 100 L each of (1) secondary effluent, (2)
disinfected, secondary effluent, and (3) the river water samples were concentrated on a CUNO 1
MDS filter by field-filtration (U.S. EPA 2001) at each WTP for each seasonal visit, including the
dry run. Sodium thiosulfate neutralization of chlorine was performed on the secondary,
disinfected Mill Creek WTP and Little Miami WTP samples at the time of collection. Half of
the eluted viruses from each of the collected samples were analyzed for total culturable viruses
by the Virus Plaque Assay (U.S. EPA 1987). The other half was saved in the freezer at -80 °C,
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and shipped by overnight delivery on dry ice to the U.S. EPA at the end of the study. Analysis
of each sample began as soon as possible upon arrival at the virus analytical laboratory. Specific
QC requirements can be found in the method. The virus laboratory followed the general
laboratory practices in the U.S. EPA Manual for Methods in Virology (www.epa.gov/nerlcwww/
about.htm), and analyzed positive and negative controls along with the samples. Results for this
method are usually expressed in plaque-forming units per ml (PFU/ml), which can be converted
to PFU/100 ml for comparison studies by multiplying by 100.
Enterovirus cytopathic effect (CPE) most-probable-number (MPN) assay combined with a
reverse transcriptase polymerase chain reaction (RT-PCR) assay
The Enterovirus cytopathic effect (CPE) most-probable-number (MPN) assay (U.S. EPA 2001,
Chapron et al. 2000), combined with a presence-absence reverse transcriptase polymerase chain
reaction (RT-PCR) assay (Gregory et al. 2006), was also used for the (1) secondary effluents, (2)
disinfected, secondary effluents, (3) the control river water samples, and (4) the 5% wastewater
in Ohio River water die-off study samples in part D for each seasonal visit, including the dry run.
Sodium thiosulfate neutralization of chlorine was performed on the secondary, disinfected
effluents of the Mill Creek and Little Miami WTPs. Volumes of 100 L each of (1) secondary
effluent, (2) disinfected, secondary effluent, and (3) river water samples were concentrated on a
CUNO 1-MDS filter by field-filtration (U.S. EPA 2001), at each WTP for each visit. Half of the
eluted viruses from each of the collected samples were analyzed for enteroviruses by the CPE
MPN method, and the other half was saved, if unused, in the freezer at -80 °C and shipped by
overnight delivery on dry ice to the U.S. EPA at the end of the study. Analysis of each sample
began as soon as possible upon arrival at the analytical laboratory. The virus laboratory followed
the general laboratory practices in the U.S. EPA Manual for Methods in Virology (U.S. EPA
1987; www.epa.gov/nerlcwww/about.htm), and analyzed appropriate positive and negative
controls along with the samples. Specific QC requirements can be found in the method or the
PCR Quality Assurance Manual (U.S. EPA 2004; www.epa.gov/nerlcwww/qa_qc_pcrlO
_04.pdf). Results were expressed in Most Probable Number per L (MPN/L), which can be
converted to PFU/100 ml for comparison studies by dividing by 10. Supernatants from the CPE
MPN Assay were combined by sample and shipped to the laboratory of Dr. Kellogg Schwab at
John Hopkins Bloomberg School of Public Health, Baltimore, Maryland where the presence or
absence of viral RNA was determined by the RT-PCR Assay (Gregory et al. 2006).
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Cryptosporidium and Giardia detection by U.S. EPA Method 1623
U.S. EPA Method 1623 (U.S. EPA 2005b) describes one of the two methods used for the
detection of Cryptosporidium in this study. In addition, samples were analyzed by this method
for Giardia as well. The procedure described for Cryptosporidium and Giardia can be used to
identify both organisms to the genus level, but not to the species level. Method 1623 utilizes (1)
field filtration to concentrate the target organisms from the water samples; (2) immunomagnetic
separation of oocysts (Cryptosporidium) and cysts (Giardia) from background material that is
also concentrated by the filtration procedure; and (3) enumeration of target organisms through
the use of an immunofluorescence assay (IF A), 4',6-diamidino-2-phenylindole (DAPI) staining,
and differential interference contrast (DIG) microscopy. Specific QC requirements can be found
in the method, and forms for recording results can be found on the Internet.
Method 1623 is a performance-based method that has been validated by U.S. EPA in one or
more national inter-laboratory studies. The sub-contractor laboratory, Tetra Tech-Clancy
Environmental (formerly Clancy Environmental Inc.), performing these analyses was chosen
from the list of U.S. EPA-approved LT2 Laboratories (U.S. EPA 2007) that can be found on the
Internet at the following address: www.epa.gov/ogwdw/disinfection/lt2/lab_aprvlabs.html.
Two 50-L portions of each secondary effluent sample and two 50-L portions of each disinfected,
secondary effluent sample were concentrated in the field according to U.S. EPA Method 1623
using Envirochek HV filters (U.S. EPA 2005b) and analyzed for Cryptosporidium and Giardia.
One filter of each type of sample was used for Method 1623, and the other was used for the
Cryptosporidium live oocyst culture method (Johnson et al. 2010). Sodium thiosulfate
neutralization of chlorine was performed for the secondary, disinfected effluents of the Mill
Creek and Little Miami WTPs in the field. ColorSeed™ C & G (BTF Pty Ltd., Sydney,
Australia) for Giardia and for Cryptosporidium internal standards were used according to the
manufacturer's instructions as a post-filtration positive control for both organisms (Francy et al.
2004). Results for these methods were expressed as oocysts (Cryptosporidium) or cysts
(Giardia) per L.
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Cryptosporidium oocyst infectivity culture method
The second method for the detection of Cryptosporidium was an oocyst infectivity culture
method (Johnson et al. 2010, Di Giovanni et al. 2006). The method was used to analyze all
secondary effluent samples before and after disinfection at each WTP for each seasonal visit,
including the dry run. Sodium thiosulfate neutralization of chlorine was used for the secondary,
disinfected effluents from the Mill Creek and Little Miami WTPs. The sub-contractor
laboratory, Tetra Tech-Clancy Environmental (formerly Clancy Environmental Inc.), performing
these analyses was chosen from the list of U.S. EPA-approved LT2 Laboratories (U.S. EPA
2007) that can be found on the Internet at the following address: www.epa.gov/ogwdw/
di sinfecti on/lt2/l ab_aprvl ab s. html.
Ancillary measurements
Additional sample information collected and ancillary measurements made are shown in Figure
A6 in the Appendix. The first five items in Figure A6 (date and time, air and water temperature,
rainfall, and cloud cover) and the last five items (GPS, pH, turbidity, conductivity, and total
suspended solids) applied to all samples at the WTPs and the Ohio River. The remaining items
with an asterisk apply only to the Ohio River samples. UV light readings were taken whenever
possible. In addition, treatment parameters at the WTP were recorded, including whether
disinfection was being used when the samples were collected. Ancillary measurements listed in
Figure A6 were collected by a variety of means, some by simple observation, while others
involved the use of equipment, such as pH meters, wind gauges, and rain gauges, etc.
Photographic data
Digital photographs were taken at all sample locations during the dry run, the three seasonal
sampling trips at the WTPs, and at the Ohio River sample intake.
Data Analysis
The SAS MIXED procedure (v. 9, SAS Institute, Cary, NC; Littell et al. 2006) was used to
model the effects of the treatment processes and the holding studies and to assess the differences
between methods. To compute the logarithms, the value of one was added to all of the data.
Values preceded by a greater than (>) or less than sign (<) were plotted at the value with the sign
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removed and marked with an asterisk in the figures. The fixed effects were Time (dry run,
spring, summer, and winter), Treatment (raw influent, primary, secondary, and disinfected
secondary), and Method. Wastewater treatment plants were treated as random effects. Separate
analyses were performed for each type of disinfection. The logic reductions were calculated for
each organism. The water summary table shows cumulative (versus raw influent) and stepwise
(versus the previous degree of treatment) logic reductions by type of disinfection (chlorination or
UV disinfection) for each type of FIB and virus, as measured by qPCR and the existing U.S.
EPA-approved methods or cited methods for that organism. The p-values (a=0.05) were
determined for the difference between qPCR and the U.S. EPA-approved or other FIB methods
with respect to stepwise log™ reductions.
Results
Comparison of Enterococcus Densities Measured by qPCR and the Densities of Fecal
Indicators Measured by Cultural Methods through the Wastewater Treatment Process
Mean fecal indicator densities in samples from the four WTPs, as determined by each of the five
culture-based methods including Enterococcus (Method 1600), Escherichia coll (Method 1603),
F+ male-specific coliphage (Method 1602), Bacteroides, and Clostridiumperfringens, were
compared with the estimated mean densities of Enterococcus calibrator cell equivalents, as
determined by the qPCR method, at each stage of treatment for each of the seasonal visits in
Figures 4-13. The mean logic reduction values for all seasons at each stage of treatment and
over the entire treatment processes are shown in Tables 1-5. The relationships between overall
logio reductions determined by these methods in treatment processes with UV light disinfection
as the final stage can be summarized as follows: Escherichia coll Method 1603 (-4.49) >
Enterococci Method 1600 (-4.33) > Bacteraides (-3.78) > F+ coliphage Method 1602 (-3.63) >
Enterococcus qPCR (-2.77) > Clostridiumperfringens (-1.85). The same relationships for the
treatment processes with chlorine disinfection as the final stage can be expressed as: Enterococci
Method 1600 (-3.50) >Escherichia coli Method 1603 (-3.30) > F+ coliphage Method 1602 (-
3.10) >Bacteroides (-2.23) > Enterococcus qPCR (-2.06) > Clostridiumperfringens (-0.89).
All of the methods, including Enterococcus qPCR, showed a decrease in the densities of their
respective target organisms as the wastewater progressed through the WTPs (raw sewage
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influent to primary effluent to secondary effluent to disinfected, secondary effluent), but the
overall decreases in Enterococcus qPCR cell equivalents were less than the reductions in the
culturable densities of all fecal indicators except Clostridium perfringens. These differences in
overall reductions across seasons and facilities were statistically significant in comparisons
between results for Enterococcus qPCR and the culture methods for Enterococcus, F+ male-
specific coliphage and E. coli, (p=<0.0001-0.0002). The logic reductions seen for these three
culture methods were also generally greater than those for the two anaerobic bacteria methods
(Bacteroides and Clostridium). Of the two anaerobic indicator methods, only the results of the
Clostridium method for the WTPs disinfected by chlorine showed significant differences
(p=0.0014) with those of the Enterococcus qPCR method and, in this instance as mentioned
above, the difference was associated with a smaller reduction in indicator densities as determined
by the culture method. Additional graphs showing individual results for each of the four WTPs
are shown in Figures A7-A46 in the Appendix.
The changes in Enterococcus densities determined by the qPCR method going from raw influent
to primary effluent were small and similar to those seen by all of the culture methods.
Substantial reductions in indicator densities were shown by all of the methods including
Enterococcus qPCR going from primary to secondary effluents. Reductions during secondary
treatment were similar and relatively consistent across WTP facilities and seasons as determined
by the Enterococcus qPCR method and the Enterococcus., E. coli, and Bacteroides culture
methods. The qPCR results were less similar to those of the F+ specific coliphage and
Clostridium methods, with the former method tending to show greater reductions than qPCR and
the latter method generally showing smaller reductions during this stage of treatment.
The most pronounced differences between the qPCR and culture methods were observed during
the disinfection processes (i.e., from secondary effluent to either chlorine- or UV- disinfected,
secondary effluent). The decreases in Enterococcus densities determined by qPCR were
significantly lower (p=<0.0001-0.0417) than those of all the culture methods except for
Clostridium, regardless of the disinfection method, except for F+ coliphage where the difference
was for chlorination only. The relatively consistent differences between the qPCR and the
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culture method results in response to disinfection strongly influenced the overall differences seen
between these methods in terms of total indicator reductions in the complete treatment processes.
UV irradiation caused larger reductions in indicator densities than chlorination as determined by
four of the five culture-base methods, as well as by the Enterococcus qPCR method (Tables 1-
5). Differences between the two disinfection methods in logic reductions (UV disinfection minus
chlorination) were as follows: Bacteroides (0.52); Escherichia coli Method 1603 (0.36);
Clostridium perfringens (0.46); Enterococci Method 1600 (-0.09); Enterococcus qPCR (0.21);
F+ coliphage Method 1602 (0.45).
Densities of Enterovirus, Cryptosporidium and Giardia Before and After Disinfection
The results from analyses of the secondary and disinfected, secondary effluent samples by the
two Enterovirus methods (Plaque Assay and the Cytopathic Effect MPN Method), as well as by
Method 1623 for Giardia and Cryptosporidium and by the Cryptosporidium live oocyst method,
are shown in Table 6. Enteroviruses were detected by both the plaque and CPE MPN assays in
very few samples. The majority of the detections with the plaque assay occurred in winter.
Detection of enteroviruses by the CPE MPN method was sporadic, and detection of Enterovirus
RNA by the RT-PCR test in the supernatants from the samples used for the CPE MPN tests did
not always match the CPE MPN positives. Like the plaque assay, more positives were found in
winter than in other seasons with the RT-PCR method.
Detection of Giardia and Cryptosporidium, using Method 1623, occurred more frequently than
the detection of enteroviruses. Giardia were recovered in all of the secondary and disinfected,
secondary effluents at Mill Creek WTP during each of the visits, and at Polk Run WTP, except
for the spring disinfected, secondary effluent. Recoveries of these pathogens at the other two
WTPs were sporadic. However, the secondary and/or disinfected, secondary effluents at all four
WTPs were all positive for Giardia cysts in winter, and almost all of them were also positive for
Cryptosporidium oocysts in winter, as well. Since the mean recoveries of Giardia cysts and
Cryptosporidium oocysts using Colorseed as a post-filtration control ranged from 2-150% (mean
=43%) and from 0-62% (mean=20.3%), respectively, the actual numbers were probably higher.
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The number of positive detections with the Cryptosporidium live oocyst culture method was few
in number and sporadic in occurrence.
Effluent Holding Studies
Results of the disinfected, secondary effluent holding studies for all four WTPs during the dry
run and all three seasonal visits are shown in Figures 14-19. Compared to the culture-based
methods for Enterococcus, F+ male-specific coliphage andE. coli (Methods 1600, 1602, and
1603), the Enterococcus densities determined by qPCR were usually 1-3 logic higher when
sampling almost immediately after the disinfection process (Day 0). These three culture-based
indicator methods each showed reductions of several logs in their target organism densities over
the 6-day holding period down to <10 or <100 per 100 ml, except in winter, while the
Enterococcus densities determined by qPCR remained relatively stable with little reduction in
the winter and dry run studies, but larger decreases in the spring and summer studies. Decreases
in the culture method indicator densities were lower during winter compared to the other seasons
and more nearly approximated the decreases in the Enterococcus qPCR results. However, the
densities of these culturable indicators in the winter effluents were still at least one order of
magnitude lower than those determined by Enterococcus qPCR.
5% Disinfected Secondary Effluent-Ohio River Mesocosm Studies
Results of the seasonal holding time studies using mesocosms containing 5% disinfected,
secondary effluents from each of the four WTPs in 95% Ohio River water are shown in Figures
20-25. As in the results of the effluent holding studies, the initial densities of Enterococcus cell
equivalents determined by qPCR were generally several orders of magnitude higher than the
corresponding initial densities of Enterococcus, E. coli, and F+ male-specific coliphage
determined by the culture methods. Most of the culturable indicator densities decreased to
values <10 per 100 ml of sample over the 6-day holding period, except in winter when the levels
remained relatively stable. Enterococcus qPCR densities generally changed in a similar manner
to the culture results over time, but remained well above the method detection limit in all
instances. Effluents from three of the four treatment plants used for spiking were disinfected in
each of the seasonal studies. Although the Polk Run WTP discontinued their UV light treatment
of secondary effluents for the winter, the mesocosms containing these effluents showed the same
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seasonal trends as those containing the other WTP effluents. Graphs in Figures A47-A62, found
in the Appendix, show the seasonal variations for mesocosms containing effluents from each of
the four individual WTPs.
20% Disinfected Secondary Effluent-Diluted Ohio River Mesocosm Studies
The overall results of the seasonal holding studies using mesocosms containing 20% disinfected,
secondary effluent from each of the four WTPs, 20% Ohio River water, and 60% partially-
treated Ohio River drinking water (source water was Ohio river) as diluents are shown in Figures
26-31. These mesocosms were designed to increase the relative amounts of indicator organisms
originating from the disinfected, secondary effluents compared to untreated, naturally-occurring
indicator organisms in the river water to more directly assess the persistence of the treated
wastewater FIB, while maintaining some natural predators in the mesocosms. Despite the higher
ratios of indicator organisms originating from the treated effluents in these mesocosms, neither
the relative nor absolute changes in their densities over time, as determined by the culture and
qPCR methods, were appreciably different from those observed in the 5% effluent mesocosms.
As in the other holding time studies, the indicator densities determined by the three culture-based
methods (Methods 1600, 1602, and 1603) generally decreased to <10 CFU or PFU per 100 ml by
Day 6, while the Enterococcus densities, determined by the qPCR method, remained several logs
higher over the entire holding period.
5% Disinfected Secondary Effluent-Ohio River Mesocosm Studies with Spiked Attenuated
Poliovirus
Enterovirus levels were usually very low or below the detection limit in the secondary and
disinfected, secondary WTP effluents examined in this study (Table 6), and Ohio River control
samples were negative by the plaque assay and the RT-PCR method. Consequently, to compare
the persistence of these viruses with Enterococcus determined by the qPCR method and fecal
indicators determined by culture methods, 5% effluent mesocosms were spiked with attenuated
poliovirus to initial densities of approximately 1000 PFU/ml. Seasonal holding time results for
the Enterococcus qPCR method, Enterococcus Method 1600, and for the two Enterovirus
methods from these mesocosms are shown in Figures 32 and 33. All of the RT-PCR tests,
performed on the pooled CPE-MPN supernatants from each sample collected from the
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poliovirus-spiked mesocosm, tested positive for Enterovirus RNA. The Enterovirus plaque
method consistently recovered more viruses than the CPE - MPN method when the
concentrations were adjusted to similar volumes of sample. Poliovirus densities, determined by
both of the Enterovirus analytical methods, decreased from initial values by -1-2 logs over the
6-day holding period in each of the seasonal mesocosms, with the exception of the winter study.
The virus densities showed either similar or up to ~1 log greater reductions compared to
Enterococcus qPCR densities over the 6-day holding period in each of the seasonal mesocosms
with the lowest reductions occurring for both methods in winter. Analysis results by four
methods (Enterococcus qPCR, Enterovirus plaque assay, Enterovirus CPE-MPN method, and
Enterococcus Method 1600) from mesocosms containing effluents from each of the four
individual WTPs are shown in Figure A63-A66 of the Appendix.
Discussion
Wastewater Treatment Studies
Samples analyzed from the four WTPs in this study indicated that Enterococcus densities,
determined by the qPCR method, were generally reduced in a similar manner to the culturable
indicator densities during the primary and secondary treatment processes, with potentially
noteworthy differences observed with F+ male-specific coliphage and Clostridium perfringens
results. In contrast, larger and often significant differences were observed between the results of
the qPCR method and the majority of the culture methods in response to the disinfection
processes. Enterococcus densities, determined by the qPCR method, were only slightly affected
by either of the two disinfection methods examined in this study, whereas indicator densities
determined by the two currently approved cultural methods for Enterococcus (Method 1600) and
Escherichia coli (Method 1603) were reduced by greater than one log after disinfection,
regardless of which method of disinfection was used. These results are consistent with previous
reports suggesting that qPCR methods may not respond in the same way as culture-based
methods when used to estimate the effectiveness of the disinfection processes in inactivating FIB
and bacterial pathogens (He and Jiang 2005, Stapleton et. al 2009, Varma et al. 2009) and
suggest that the qPCR method cannot be substituted for cultural methods in determining the
effects of wastewater treatment. Disinfection, a widely- used step in WTP treatment processes,
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is often critical for facilities to meet the requirements of National Pollutant Discharge
Elimination System (NPDES) permits, as well as total maximum daily loads (TMDLs).
An alternative to traditional qPCR methods that do not distinguish between viable and non-
viable cells is an approach that utilizes propidium monoazide (PMA) in the qPCR assay to allow
of the quantification of intact and, presumably, viable cells (Bae and Wuertz 2009a, 2009b,
Nocker et al. 2006, Varma et al. 2009). These studies have determined that lower densities of
Enterococcus and Bacteroidales were detected when PMA-qPCR was used, compared to
traditional qPCR indicating the presence of viable cells. In future studies, the inclusion of
comparisons between the PMA-qPCR, qPCR, and culture methods may indicate whether the
PMA-qPCR is a suitable alternative that responds to disinfection in a similar manner to the
culture methods.
It is potentially noteworthy, however, that the disinfection processes also had smaller effects on
reductions of F+ male-specific coliphage, and particularly Clostridium perfringens densities,
compared to the approved indicator bacteria as determined by their respective culture methods.
These observations were consistent with published reports indicating that both of these less
conventional culturable indicator groups are relatively resistant to wastewater disinfection
(Chauret et al. 1999). Coliphages (Chauret et al. 1999, Skraber et al. 2002) and have been
suggested as potentially superior indicators of the effectiveness of wastewater treatment
processes on viral pathogens, whereas Clostridium spores have been argued as potentially
superior surrogates of protozoan pathogens (Chauret et al. 1999). Reductions of Enterococcus
densities determined by the qPCR method differed, in some instances significantly, from those of
these two alternate culturable indicator groups in response to disinfection, as well as in response
to the overall treatment processes. However, the intermediate position of the qPCR results
between the F+ coliphage and Clostridium perfringens methods in the disinfection rankings and
the more conservative reduction of qPCR signals compared to all of the FIB methods
demonstrating reductions in this study suggest that the qPCR method may have more value as a
general predictor of treatment efficacy for all non-bacterial pathogens. Research in which non-
bacterial pathogens have been shown to be more resistant to disinfection than the general
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culturable indicators also supports this hypothesis (Blatchely et al. 2007, Bonadonna et al. 2002,
Crockett 2007, Tree et al. 2003, Varma et al. 2009).
It is also of potential interest that, of all alternative methods examined in this study, the one that
showed the greatest similarity to Enterococcus qPCR in terms of demonstrated indicator
reductions through the entire treatment processes was the culture method for Bacteroides. While
the significance of this observation is currently unclear, it is noteworthy that genetic markers
from Bacteroides species are becoming increasingly popular targets in microbial fecal source-
tracking investigations, and some studies have provided evidence that the persistence of
Bacteroides genetic markers in the environment may mimic certain pathogens (Walters et al.
2009). In the future, the analysis of qPCR methods for other FIB will be compared to their
corresponding culture methods to determine if similar patterns emerge.
The results of this study and others (He and Jiang 2005, Stapleton et al. 2009) suggest that levels
of fecal indicators determined by qPCR are generally less affected by overall treatment than
when analyzed by culture. However, other studies have indicated that qPCR and culture-based
methods are more closely correlated (Lavender and Kinzelman 2009, Varma et al. 2009).
Previous studies have suggested that the culturable FIB may overestimate the effectiveness of
wastewater treatment compared to viral and protozoan pathogens (Blatchely et al. 2007,
Bonadonna et al. 2002, Crockett 2007, Tree et al. 2003, Varma et al. 2009). In this study,
physical removal, the major process in primary and secondary treatment for the removal of fecal
indicators, has been found to have similar effects on the densities of fecal indicators by culture
and qPCR. Significant differences between FIB density estimates by qPCR and culture occur
when disinfection processes were used. During the disinfection process, inactivated cells are not
physically removed from the treated effluent, and, thus, are still present to be detected by qPCR
even though the inactivated cells can no longer be cultured.
Many factors were considered while deciding which pathogens to include in the study. The
biology of the pathogens, ease of detection, and the availability of culture and non-culture
methods for a given pathogen were considered. In addition, it is known that protozoa and viruses
are believed to be a leading cause of waterborne illness (Henrickson et al. 2001). Norovirus is an
26
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important pathogen, but no culture method is available; so it was not included. Enterovirus was
selected because well-characterized culture and molecular methods were available.
The most important function of any fecal indicator method with respect to monitoring
wastewater treatment processes is to accurately predict the efficacy of different processes in
removing and/or inactivating pathogens. To examine this question with respect to the different
indicator methods used in this study, Enteroviruses, Giardia, and Cryptosporidium pathogen
densities in the secondary and disinfected, secondary effluents of each of the WTPs were also
determined.
Except in winter, the levels of all three pathogen groups were often below the detection limits of
the analytical methods, making any comparisons of treatment effects on the pathogens and the
indicators difficult. In the winter sampling visit, where Giardia and Cryptosporidium were
detected in the majority of either secondary and/or disinfected, secondary effluents, no clear
reductions in densities of these organisms were observed in response to the disinfection
processes at the different facilities, although Giardia levels were slightly higher than those for
Cryptosporidium. The reduced effectiveness of the disinfection process appeared to be more
consistent with the results of the Enterococcus qPCR and Clostridium culture methods. Because
the Enterovirus densities were usually below the detection limits, the relationship of the
treatment effectiveness of Enterovirus with the fecal indicators could not be quantified.
Increased sensitivity (larger volumes and more efficient recoveries) in the detection of
Enterovirus may be needed in order to document detectable levels of virus through the
disinfection process. As indicated above in this report and discussed previously by others
(Harwood et al. 2005), the diversity of different types of pathogens that may occur in
wastewaters presents challenges in using any single indicator method to predict the efficacy of
different treatment processes in reducing overall pathogen content.
The results from this study did not provide conclusive comparisons between changes in
Enterovirus, Cryptosporidium, and Giardia densities during wastewater treatment and changes in
fecal indictor densities, determined by either culture or qPCR methods, because of the very low
levels of the pathogens that were detected and quantified from the wastewater samples.
27
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However, the logic reduction values of Enterococcus densities determined by qPCR in this study
(-2.8 log reduction for Enterococcus qPCR compared to -4.3 log reduction for Enterococcus
culture) appeared to be similar to logio reduction values that have been reported for pathogens
that are more resistant to disinfection in other studies (i.e., -1.5-3.5 log reduction for
Cryptosporidium oocysts and Giardia cysts and -2.5-3.0 log reduction for enteric viruses;
Chauret et al. 1999, Rose et al. 2004, Varma et al. 2009). There have been a number of studies
that have examined the effects of wastewater treatment on fecal indicators and pathogens, and
the majority of these studies have shown that, when culture-based methods are used, FIB
densities are greatly reduced while pathogen densities are not as efficiently removed (Baggi et al.
2001, Bonnadonna et al. 2002, Chauret et al. 1999, Rose et al. 2004, Tyrrell et al. 1995, Varma et
al. 2009). Fewer studies have examined the relationships between pathogens and FIB levels
determined by qPCR (He and Jiang 2005, Shannon et al. 2007, Stapleton et al. 2009, Varma et
al. 2009).
Indicator Persistence Studies
The purpose of the effluent holding portion of the study was to determine if holding disinfected,
secondary effluents for up to six days would reduce the Enterococcus qPCR values to levels that
are similar to fecal indicator levels determined by culture. Such an outcome might be predicted
if inactivating the indicator organisms by disinfection has a delayed effect on the stability of their
nucleic acids and could potentially provide support for a hypothesis that indicator densities from
treated effluents will reach similar levels after some period of time in ambient waters, as
determined by either culture or qPCR methods. Fecal indicator densities determined by the three
culture-based methods (Methods 1600, 1602, and 1603) decreased to very low levels over the 6-
day holding time, except in winter. The greater persistence of these culturable organisms in
winter was expected, as cold temperatures generally favor survival of microorganisms (Dick et
al. 2010, Arnone and Walling 2007, Okabe and Shimizu 2007, Seurinck et al. 2005, Terzieva and
McFeters 1991). Persistence particularly in the winter months were less resolved because of the
slower decay compared to the other seasons.
In contrast, Enterococcus densities, as determined by the qPCR method, in the disinfected
effluents generally declined at a slower rate. The qPCR-determined indicator densities were also
28
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generally several orders of magnitude higher than the corresponding indicator densities
determined by the three culture-based indicator methods. These results contradict the hypothesis
that fecal indicator densities from treated WTP effluents will reach comparable levels, as
determined by culture and Enterococcus qPCR methods, after a reasonable amount of time in the
absence of other environmental factors that may affect the fate of these organisms in ambient
receiving waters.
The mesocosm holding time studies, containing 5% disinfected, secondary effluents from each of
the four WTPs in Ohio River water, were designed to compare the persistence of indicators from
treated wastewater, as determined by the qPCR method and by three culture-based FIB methods
(Methods 1600, 1602, and 1603), in the presence of ambient receiving waters. As in the effluent
holding studies, the initial Enterococcus densities determined by the qPCR method were
generally several orders of magnitude higher than the corresponding indicator densities
determined by the three culture-based indicator methods in these seeded mesocosms. In contrast
to the effluent holding studies, however, the densities of Enterococcus qPCR cell equivalents and
the culture-based fecal indicators in these mesocosms showed fairly similar patterns and overall
levels of reduction during the holding period. These results suggest that the factors that affect
the persistence these organisms in ambient receiving waters may affect both their viability and
the stability of their nucleic acids in a relatively similar manner.
It is important to note that the mesocosm studies were conducted in the dark and measured
persistence without the influence of sunlight. The results, therefore, represent conditions that
may increase the persistence of culture and molecular targets compared to conditions that include
the effects of sunlight. These simulated recreational water samples with little to no exposure to
UV light during storage at seasonal temperatures are representative of a worst case situation
when disinfected, secondary effluents are discharged into their receiving waters. These
conditions favor organism survival and greater persistence in their detection.
Results from several studies that have compared the persistence of culture and molecular targets
have indicated that sunlight increases the decay of cultured indicators, but has less of an effect on
DNA targets in fresh and marine water (Bae and Wuertz 2009b, Sinton et al. 1999, Walters and
29
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Field 2009, Walters et al. 2009, Dick et al. 2010, Muela et al. 2000, Green et al., Personal
Communication). The mesocosms in this study were done in the dark. As stated above for
qPCR, studies done in the dark may provide a relatively good predictor for persistence in
sunlight. However, a recent study by Walters et al. (2009) found that light did have an influence
on decay rates for Bacteroidales as measured by qPCR. However, the results from other studies
have generally found that the densities of FIB measured by qPCR are not as affected by light as
the culture method. Turbidity may play a role on the effect of sunlight on the persistence of
culture and qPCR targets (Cantwell and Hofmann 2008, Dick et al. 2010). The results from the
present study also clearly indicate that UV disinfection has a differential effect on culture and
molecular targets with significantly greater impacts on culture-based measurements. A steeper
decay of culture-based indicators might have been observed if the mesocosms were exposed to
sunlight. Other studies have observed an inverse relationship between densities of culturable
bacterial fecal indicators and increasing sunlight (Fujioka et al. 1981, Lessard et al. 1983,
Kapuscinski and Mitchell 1981).
In this study, there were differences in the relative starting concentrations of the culturable and
non-culturable indicators. Relatively high ratios of qPCR-detectable to culturable indicators
were used in this study as a result of seeding the mesocosms with disinfected effluents, as
opposed to raw sewage. Parallel analyses of mesocosms containing only Ohio River water in
this study indicated that the FIB levels determined by all methods were at least one log lower
than those in the seeded macrocosms (results not shown) and, therefore, had little impact on the
interpretation of results, except in winter months when ambient densities of fecal indicators were
higher.
Mesocosms seeded with 20% disinfected effluents in a sample containing 20% Ohio River water
and 60% partially-treated drinking water to increase the relative amounts of indicator organisms
originating from the effluents gave results similar to those of the mesocosms containing only 5%
disinfected effluents. A conclusion from these studies is that ambient waters that are consistently
impacted by large amounts of treated wastewater effluents should be expected to show relatively
high ratios of indicator densities as determined by qPCR compared to culture methods, despite
the fact that their decay rates may be similar.
30
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As is the case with monitoring wastewater treatment processes, the most important question for
any fecal indicator method with respect to monitoring of ambient waters for microbial-related
health risks is how accurately the method predicts the occurrence of pathogens. To accurately
predict pathogen occurrence, the persistence of the fecal indicator and pathogen must be similar
in ambient waters. To examine this question with respect to the different indicator methods used
in this study, the 5% effluent mesocosms were spiked with attenuated poliovirus to initial
densities of approximately 1000 PFU per ml. While laboratory-grown viruses, may behave
differently than naturally-occurring viruses in terms of exhibiting lower persistence (Tree et al.
2003), this procedure was necessitated by the very low Enterovirus levels that were found to
occur in all secondary WTP effluents in this study. Results from this study showed similar
patterns of persistence of the spiked polioviruses compared to Enterococcus determined by the
qPCR and culture method. It should be noted, however, that the viruses that cause the bulk of
recreational water disease are not enteroviruses, and these viruses may persist much longer than
enteroviruses.
Conclusions
Wastewater Treatment
1. The reduction of Enterococcus densities measured by qPCR and culture were similar during
primary and secondary treatment, but were significantly different (p=0.05) during
disinfection using either UV light disinfection or chlorination. The reduction of
Enterococcus densities by culture were significantly greater than the reduction of the qPCR
method during disinfection and also during the complete treatment processes. Similar
patterns were observed between the Enterococcus qPCR and E. coli culture methods.
2. The differences were less pronounced for Enterococcus qPCR comparisons with F+ male-
specific coliphage, Bacteroides and Clostridium perfringens culture methods.
3. The effects of UV light and chlorination disinfection processes on reductions of
Enterococcus densities, as determined by qPCR, were similar.
4. No association between the degradation of enteroviruses and fecal indicators could be
determined, in part because of the very low concentrations of enteroviruses that were
31
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detected in the treated wastewater. Differences in the densities of Giardia cysts and
Cryptosporidium oocysts could not be detected between secondary and disinfected,
secondary treated wastewater samples because of the very low concentrations of both
organisms.
Holding Studies
1. In general, greater reductions of fecal indicator densities were observed by culture than by
Enterococcus qPCR assays in effluent holding studies.
2. Reductions of fecal indicator densities observed by culture and by Enterococcus qPCR were
generally more consistent when holding effluents in the presence of ambient surface waters
than when holding effluents alone.
3. For all holding studies, the initial densities of Enterococcus determined by qPCR were
generally several orders of magnitude higher than the corresponding densities of culturable
Enterococcus, E. coli, and F+male-specific coliphages except in the winter samples.
4. For all of the holding studies, reductions of all fecal indicators densities were lowest in the
winter.
5. Reductions of spiked, attenuated polioviruses in wastewater effluent-Ohio River holding
studies were similar to those of Enterococcus determined by both the qPCR and culture
methods.
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viruses by positive charge 1 MDS cartridge filters and organic flocculation. In: U.S. EPA
Manual of Methods for Virology. Publication EPA-600/4-84/013 (N14), Office of Research and
Development, Washington, DC. Available at: www.epa.gov/microbes.
U. S. Environmental Protection Agency. 2001d. Total culturable virus quantal assay. In: U.S.
EPA Manual of Methods for Virology. Publication EPA-600/4-84/013 (N15), Office of Research
and Development, Washington, DC. Available at: www.epa.gov/microbes.
U. S. Environmental Protection Agency. 2002a. EPA Method 1600: Enterococci in water by
membrane filtration using membrane-Enterococcus Indoxyl-p-D-Glucoside Agar (mEI Agar)
(September, 2002), Publication EPA 821-R-02-022, Office of Water, Washington, DC.
Available at: www.epa.gov/microbes.
U. S. Environmental Protection Agency. 2002b. EPA Method 1603: Escherichia coll (E.colf) in
water by membrane filtration using modified membrane-thermotolerant Escherichia coll agar
(Modified mTEC agar) (September, 2002), Publication EPA821-R-02-023, Office of Water,
Washington, DC. Available at: www.epa.gov/microbes.
40
-------
U. S. Environmental Protection Agency. 2004. Quality assurance/quality control guidance for
laboratories performing PCR analyses on environmental samples. EPA 815-B-04-001, Office of
Water, U.S. Environmental Protection Agency, Washington, DC.
U. S. Environmental Protection Agency. 2005a. Manual for the certification of laboratories
analyzing drinking water: Criteria and procedures, quality assurance, 5th edition. Publication
EPA 815-R-05-004, Office of Water, Office of Ground Water and Drinking Water, Technical
Support Division, Cincinnati, OH.
U. S. Environmental Protection Agency. 2005b. EPA Method 1623: Cryptosporidium and
Giardia in water by filtration/IMS/FA (December, 2005), Publication EPA 815-R-05-002, Office
of Water, Washington, DC. Available at: www.epa.gov/microbes.
U. S. Environmental Protection Agency. 2007. Laboratories approved for the analysis of
Cryptosporidium under the Safe Drinking Water Act; Long Term 2 Enhanced Surface
Water Treatment Rule (LT2). Available at: www.epa.gov/ogwdw/disinfection/lt2/lab_
aprvlabs.html.
Wade, T. I, R. L. Calderon, E. Sams, M. Beach, K. P. Brenner, A. H. Williams, and A. P.
Dufour. 2006. Rapidly measured indicators of recreational water quality are predictive of
swimming-associated gastrointestinal illness. Environ Health Persp. 114(l):24-28.
Wade, T. I, R. L. Calderon, K. P. Brenner, E. Sams, M. Beach, R. Haugland, L. Wymer, and A.
P. Dufour. 2008. High sensitivity of children to swimming-associated gastrointestinal illness:
results sing a rapid assay of recreational water quality. Epidemiology. 19(3):375-383.
Wade, T.J., E. Sams, K. P. Brenner, R. Haugland, E. Chern, M. Beach, L. Wymer, C. C. Rankin,
D. Love, Q. Li, R. Noble, and A. P. Dufour. 2010. Rapidly measured indicators of recreational
water quality and swimming-associated illness at marine beaches. Submitted.
41
-------
Walters, S. P., and K. G. Field. 2009. Survival and persistence of human and ruminant-specific
faecal Bacteroidales in freshwater microcosms. Environ. Microbiol. 11:1410-1421.
Walters, S. P., K. M. Yamahara, and A. B. Boehm. 2009. Persistence of nucleic acid markers of
health-relevant organisms in seawater microcosms: implications for their use in assessing risk in
recreational waters. Water Res. 43:4929-4939.
Varma, M., R. Field, M. Stinson, B. Rukovetsc, L. Wymer, and R. Haugland. 2009. Quantitative
real-time PCR analysis of total and propidium monoazide-resistant fecal indicator bacteria in
wastewater. Water Res. 43:4790-4801.
42
-------
Table 1. Enterococcus qPCR-CE Compared To Method 1600 Enterococcus CPUs Logio Reduction
Through The Treatment Stages: Combined Results Over All Seasons
Disinfection
Method Treatment Step
CI2 Raw -> Primary
Primary -> Secondary
Secondary -> Disinfection
Raw-> Disinfection
UV Raw -> Primary
Primary -> Secondary
Secondary -> Disinfection
Raw-> Disinfection
Log10 Reduction (-) or Increase (+)
qPCR(Log10 Range)
-0.3680 (-0.8858 to -0.0560)
-1.5730 (-2.7801 to -0.2653)
-0.1 150 (-0.5368 to 1.0287)
-2.0560 (-2.6719 to -1.6331)
-0.1200 (-0.4757 to 0.3814)
-2.3440 (-3.2218 to -1.9121)
-0.3033 (-0.8246 to 0.1780)
-2.7670 (-3.1048 to -2.1479)
CPU (Log 10 Range)
-0.1 250 (-0.6830 to 0.1 453)
-1.6980 (-2.7778 to -1.0279)
-1.6780 (-2.91 16 to -0.5309)
-3.5020 (-4.5932 to -2.0464)
-0.1720 (-0.7773 to 0.0582)
-2.5650 (-2.8586 to -2.0830)
-1.5890(-2.2952 to 0.1 808)
-4.3260(-4.9028 to -2. 6794)
Difference in Log10
Reduction
qPCR-CFU
-0.2430
0.1250
1.5630
1.4460
0.0520
0.2210
1.2857
1.5590
P-valuefor
Difference
0.4471
0.6936
<0.0001"
<0.0001"
0.7782
0.2300
<0.0001"
<0.0001"
* Bold values are statistically significant (p=0.05).
-------
Table 2. Enterococcus qPCR-CE Compared To Method 1602 F+ Coliphage PFUs Log™ Reduction
Through The Treatment Stages: Combined Results Over All Seasons
Disinfection Treatment step
Method
CI2 Raw -> Primary
Primary -> Secondary
Secondary -> Disinfection
Raw -> Disinfection
UV Raw -> Primary
Primary -> Secondary
Secondary -> Disinfection
Raw -> Disinfection
Log10Reduction(-)
qPCR( Log 10 Range)
-0.3680 (-0.8858 to -0.0560)
-1.5730 (-2.7801 to -0.2653)
-0.1 150 (-0.5368 to 1.0287)
-2.0560 (-2.6719 to -1.6331)
-0.1 200 (-0.4757 to 0.3814)
-2.3440 (-3.2218 to -1.9121)
-0.3033 (-0.8246 to 0.1780)
-2.7670 (-3.1048 to -2.1479)
or Increase (+)
PFU(Log10 Range)
-0.1 112 (-0.4155 to 0.4006)
-2.5195 (-4.6415 to -1.9314)
-0.4712 (-2.3782 to 0.2526)
-3. 1019 (-4.5953 to -2.2565)
-0.1047 (-1.2186 to 0.2116)
-2.6089 (-3.0212 to -2.0159)
-0.9178 (-1.8839 to 0.3428)
-3.6314 (-5.2967 to -2.5925)
Difference in Log10
Reduction
qPCR-PFU
-0.2568
0.9465
0.3562
1.0459
-0.0153
0.2649
0.6144
0.8643
P-value for
Difference
0.2728
0.0001"
0.1299
<0.0001*
0.9407
0.2040
0.0060"
0.0002"
* Bold values are statistically significant (p=0.05).
-------
Table 3. Enterococcus qPCR-CE Compared To Method 1603 E, coli CPUs Log-|0 Reduction
Through The Treatment Stages: Combined Results Over All Seasons
Disinfection Treatment step
Method
CI2 Raw -> Primary
Primary -> Secondary
Secondary -> Disinfection
Raw -> Disinfection
UV Raw -> Primary
Primary -> Secondary
Secondary -> Disinfection
Raw-> Disinfection
Log10 Reduction {-)
qPCR( Log10 Range)
-0.3680 (-0.8858 to -0.0560)
-1.5730 (-2.7801 to -0.2653)
-0.1150 (-0.5368 to 1.0287)
-2.0560 (-2.6719 to -1.6331)
-0.1200 (-0.4757 to 0.3814)
-2. 3440 (-3. 22 18 to-1.9121)
-0.3033 (-0.8246 to 0.1780)
-2.7670 (-3.1048 to -2. 1479)
or Increase (+)
CPU (Log 10 Range)
0.0809 (-0.2685 to 0.4434)
-1.9135 (-2.6049 to -1.2250)
-1.4643 (-3.0562 to -0.2609)
-3.2969 (-4.6233 to -2.6271)
0.0602 (-0.1042 to 0.3391)
-2.7238 (-3.1997 to -2.4009)
-1.8218 (-2.6055 to 0.0509)
•4.4854 (-4.9441 to -2.7871)
Difference in Log10
Reduction
qPCR-CFU
-0.4489
0.3405
1.3493
1.2409
-0.1802
0.3798
1.5185
1.7184
P-value for
Difference
0.1253
0.2421
<0.0001"
<0.0001a
0.2722
0.0230"
<0.0001"
<0.0001"
* Bold values are statistically significant (p=0.05).
-------
Table 4. Enterococcus qPCR-CE Compared To Bacteroides fragilis CPUs Log10 Reduction Through
The Treatment Stages: Combined Results Over All Seasons
Disinfection Treatment step
Method
CI2 Raw -> Primary
Primary -> Secondary
Secondary -> Disinfection
Raw -> Disinfection
UV Raw -> Primary
Primary -> Secondary
Second ary-> Disinfection
Raw -> Disinfection
Log 10 Reduction (-)
qPCR( Log 10 Range)
-0.3680 (-0.8858 to -0.0560)
-1.5730 (-2.7801 to -0.2653)
-0.1150 (-0.5368 to 1.0287)
-2.0560 (-2.6719 to -1.6331)
-0.1200 (-0.4757 to 0.3814)
-2.3440 (-3.2218 to -1.9121)
-0.3033 (-0.8246 to 0.1780)
-2. 7670 (-3. 1048 to -2, 1479)
or Increase (+)
CPU (Log10 Range)
-0.0352 (-0.5071 to 0.5128)
-1.4560 (-2.8789 to -0.4697)
-0.7437 (-1.7086 to -0.0988)
-2.2349 (-2.8361 to -1.2004)
-0.1033 (-0.5484 to 2.2235)
-2.4126 (-3.9219 to -1.7194)
-1.2660 (-1.7899 to 0.0934)
-3.7818 (-5.4116 to -0.5024)
Difference in Log10
Reduction
qPCR-CFU
-0.3328
-0.1170
0.6287
0.1789
-0.0167
0.0686
0.9627
1.0148
P-value for
Difference
0.2532
0.6862
0.0344"
0.5464
0.3945
0.8750
0.0417"
0.1633
VO
* Bold values are statistically significant (p=0.05).
-------
Table 5. Enterococcus qPCR-CE Compared To Clostridium perfringens CPUs Logio Reduction
Through The Treatment Stages: Combined Results Over All Seasons
Disinfection
Method
CI2
uv
Treatment Step
Raw -> Primary
Primary -> Secondary
Secondary -> Disinfection
Raw -> Disinfection
Raw -> Primary
Primary -> Secondary
Secondary -> Disinfection
Raw -> Disinfection
Log10 Reduction i
qPCR(Log10 Range)
-0.3680 (-0.8858 to -0.0560)
-1.5730 (-2.7801 to -0.2653)
-0.1150 (-0.5368 to 1.0287)
-2.0560 (-2.6719 to -1.6331)
-0.1200 (-0.4757 to 0.3814)
-2.3440 (-3.2218 to -1.9121)
-0.3033 (-0.8246 to 0.1780)
-2.7670 (-3.1048 to -2.1479)
(-) or Increase (+)
CPU (Log 10 Range)
-0.2610 (-1.2000 to 0.5139)
-0.6894 (-1.4418 to 0.6811)
0.0613 (-0.1909 to 0.8969)
-0.8892 (-1.7869 to 0.3780)
-0.1234 (-1.1629 to 0.9720)
-1.3248 (-2.7635 to 0.6926)
-0.4036 (-0.6423 to 0.1896)
-1.8517 (-3.0426 to -0.0432)
Difference in Log 10
Reduction
qPCR-CFU
-0.1070
-0.8836
-0.1763
-1.1669
0.0034
-1.0192
0.1003
-0.9153
P-value for
Difference
0.7600
0.0136"
0.6126
0.0014"
0.9958
0.0279"
0.8290
0.0528
" Bold values are statistically significant (p=0.05).
-------
Table 6, Pathogens in Secondary and Disinfected Secondary Effluents
SEASONAL CONCENTRATIONS3
Organism
Enterovirus
Giardia
Ciyptosporidium
Method
Used
Plaque
(PFU per ml)
CPE MPN
(MPN per L)
RT-PCR
(Presence-absence)
Method 1623
(Cysts per L)
Method 1623
(Oocysts per L)
Culture
(Live oocysts
per mL)
Preliminary
WTP
2°
MC <0.1
MD <0.1
LM
PR
MC <0.07
MD <0.07
LM
PR
MC -/-
MD -/-
LM
PR
MC 0.35
MD 0.36
LM
PR
MC 0.25
MD 1.12
LM
PR
MC b
MD b
LM
PR
Disin-
fected
2°
<0.1
<0.1
<0.07
<0,07
'-'-
0.067
0
0.4
0
b
b
Spring
2°
<0,1
<0.1
<0.1
<0.1
0.067
<0.067
<0.067
0.067
;
0.033
0
0
0.02
0.033
0
0
0
0.05
0.05
0
0
Disin-
fected
2°
<0.1
<0.1
<0.067
<0.067
<0.067
<0.067
;
0.15
0
0
0
0
0
0
0
0
0
0
0.1
Summer
2°
<0.1
<0.1
<0.1
0.1
<0.067
<0.067
0.067
0.067
:i:
0.63
0
0
1.35
0
0
0
0.15
0
0
0
0
Disin-
fected
2°
0.1
<0.1
<0.1
<0.1
<0.067
<0.067
<0.067
<0.067
->-
0.3
0
0
1.8
0
0
0
0
0
0
0
0
Winter
2°
0.2
0.2
0.4
0.1
<0.067
<0.067
<0.067
<0.067
;:
0.1
0.02
0.325
1.68
0
0.06
0.025
0.04
0
0
0.05
0
Disin-
fected
2°
<0.1
<0.1
<0.1
<0.067
<0.067
<0.067
0.07
t
0.417
0.125
3.175
0.84
0.33
0.075
0.05
0.02
0
0
0
0
oo
a Bold values are above the detection limit of the method.
b Lost Monolayers
-------
METHOD ANALYTE
Method 1600 Enterococei
Wastewater Study (Day 0)
DISINFECTED
WTP INFLUENT PRIMARY SECONDARY SECONDARY
LM XXX
Method 1603 Escherichia coli
QPCR
Enterococei
Method 1602 Coliphage
mCP Agar Clostridium
perrringens
BBE Agar Bacteroides
ttagilis
M8
MC
MD
PR
LM
XXX
xxx
XXX
XXX
W
Dieoff Study
Seconda
of Disinfected
ry Effluent
Dieoff Study of Virus-Spiked
Disinfected 5% Wastewater in River Water
DAY 1 DAY 2 DAY 4 DAY 6 DAY 0 DAY 1 DAY 2 DAY 4 DAY 6
xxx xxx
xxx xxx
XXX XXX
XXX XXX
XXX XXX
XXX XXX
XXX XXX XXX XXX
XXX XXX XXX XXX
XXX XXX XXX XXX
XXX XXX XXX XXX
CONTROLS
Dieoff Study of Non-Spiked Virus
River Water Spike
DAY 0 DAY 1 DAY 2 DAY 4 DAY 6 DAY 0
XXX XXX XXX XXX XXX
XXX XXX XXX XXX XXX
XXX XXX XXX XXX XXX
XXX XXX XXX XXX XXX
XXX XXX XXX XXX XXX
XXX XXX XXX XXX XXX
XXX XXX XXX XXX XXX
XXX XXX XXX XXX XXX
CPE + RT-PCR Enterovrrus
PFU
Enterovirus
Method 1623 Cryptosporidium
and Giardia
Oocysts Cryptosporidium
XXX
XXX
XXX
XXX
XXX
XXX
XXX
NOTE: Disinfected Secondary Wastewater Samples in Parts A
and B are also the Wastewater Controls in the Part D Study.
NOTE: Each X stands for the sample(s)
taken during each trip/visit.
* Consecutive Grab Samples
# All samples obtained from a larger sample
collected and mixed in a 100-gallon tank.
" This sample is DAY 0 for Part C.
COLOR KEY:
D Part A
D PartB
n PartC
Q PartD
Wastewater Treatment Plant, WTP Disinfection
MC
MD
Mill Creek WTP
_
Little Miami WTP
CL2
UV
CL2
FIGURE 1. Wastewater Study - Water Sample Analysis
-------
CONTROLS
Dleoff Study of Disinfected Dleoff Study of Virus-Spiked
Wastewater Study (Day 0) Secondary Effluent Disinfected 5% Wastewater Dieoff Study of Non-Spiked
DISINFECTED in River Water River Water
DAY 4 DAY 6 DAY 0 DAY 1 DAY 2 DAY 4 DAY 6
METHOD
Method 1600
Method 1603
QPCR
Method 1602
mCP Agar
BBE Agar
CPE + RT-PCR
PFU
Method 1623
Oocysts
ANALYTE
Enterococci
Escnerichia coli
Enterococci
Coliphage
Clostridium
perfringens
Bacteroides
fragilis
Enterovirus
Enterovirus
Cryptosporidium
and Giardia
Cryptosporidium
WTP
MC
MD
MS
MS
MS
MB
MC
MD
MC
MD
MC
MD
MC
MD
MB
INFLUENT PRIMARY
X
X
X
X
X
X
x
X
X
X
X
X
X
X
X
X
X
X
X
X
X
SECONDARY
X
X
X
X
X
X
8
X
X
X
X
X
X
X
8
X
SECONDARY
X
X
X
X
X
8
x D
X
X
X
X
x n
DAY 1 DAY 2
X
X
X
X
X
£
II
I
X
X
x
X
X
X
DAY 4
X
X
X
X
X
X
X
X
DAY 6
X
X
X
X
X
X
X
X
H
DAYO
X
X
X
X
x
X
X
X
DAY1
X
X
X
X
X
X
X
X
s
DAY 2
X
X
X
X
X
X
X
X
X
X
X
DAY 4
X
X
X
X
X
X
X
X
X
X
DAY
X
X
X
X
X
X
X
x
X
X
Virus
Spike
DAYO
NOTE: Each X stands for the sample(s)
taken during each trip/visit.
'Consecutive Grab Samples
#AII samples obtained from a larger sample
collected and mixed in a 100-gallon tank.
" This sample is DAY 0 in Part C.
COLOR KEY:
o Part A
d PartB
dPartC
a Part D
Wastewater Treatment Plant, WTP Disinfection
MC Mill Creek WTP
MD Muddy Creek WTP
CL2
UV
FIGURE 2. Wastewater Study - Water Sample Analysis - Dry Run Samples
-------
METHOD ANALYTE
Method 1600 Enterococci
Method 1603 Escherichia coli
QPCR Enterococci
Method 1602 Coliphage
mCP Agar Clostridtum
perfringens
BBE Agar Bacteroides
fragilis
CPE + RT-PCREnterovirus
PFU
Enterovirus
Method 1623 Cryptosporidium
ana Giardia
Oocysts Cryptosporidium
WTP
Dieoff Study of Filtered River Water Spiked
With 20 % Wastewater and 20 % Partially Treated DW
DAYO DAY1 DAY 2 DAY 4 DAYS
MC
MD
PR
LM
MC
MD
PR
LM
MC
MD
PR
LM
MC
MD
PR
LM
MC
MD
PR
LM
MC
MD
PR
LM
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
NOTE
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
Each X stands for the sample(s)
taken during each trip/visit.
# All samples obtained
MC
MD
PR
LM
MC
MD
PR
LM
MC
MD
PR
LM
MC
MO
PR
LM
from a larger sample
collected and mixed in a 100-gaTlon tank.
Partially
Treated
DW Control
Day 0
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
XXX
Dry Run Dieoff Study of Filtered River Water Spiked
With 20 % Wastewater and 20 % Partially Treated DW
DAYO
DAY1
DAY 2
DAY 4
DAY 6
Dry Run
Partially
Treated
DW Control
Day 0
X
X
X
X
D
COLOR KEY:
a Part A
a Part B
DPartC
npartD
Wastewater Treatment Plant, WTP Disinfection
MC
MD
PR
LM
Mill Creek WTP
Muddy Creek WTP
Polk Run WTP
Little Miami WTP
CL2
UV
UV
CL2
FIGURE 3. Wastewater Study - Water Sample Analysis, Including the Dry Run
-------
E
o
o
Is
(1) O
§1
O o
107-
106-
io5-
O 10"
103-
ill 102
o
1 -
10
10°
(N
Method 1600 CPU
qPCR-CE
"i 1 1 1 r~
•& ^
-i 1 —i r~
*
Preliminary
Spring
Summer
Winter
Figure 4. Overall Comparison of EPA Method 1600 Enterococcus CPUs (o) and Quantitative
Polymerase Chain Reaction (qPCR) Cell Equivalents (•) Through the Wastewater Treatment Process
(These values are based on the use of 56 samples for each method, 16 samples each for Mill Creek
and Muddy Creek, and 12 samples each for Little Miami and Polk Run)
-------
E
o
o
.§ =>
CtJ
Q_
(D O
11
O o
107
106
1Q5-
104-
103-
o io2-
10°
Method 1602PFU
qPCR-CE
jff
Preliminary
Spring
Summer
Winter
Figure 5. Overall Comparison of EPA Method 1602 F+ Coliphage PFUs (O) and Quantitative
Polymerase Chain Reaction (qPCR) Cell Equivalents (•) Through the Wastewater Treatment Process
(These values are based on the use of 56 samples for each method, 16 samples each for Mill Creek
and Muddy Creek, and 12 samples each for Little Miami and Polk Run)
-------
E
o
o
g ID
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|o
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§1
O o
LU
O
107,
106
105
104
103
102
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qPCR-CE
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-------
E
o
o
c ;:
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11
O o
lil
O
107 i
106
105
104
102
10°
S. frap/7/sCFU
qPCR-CE
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1
o 1Q5
o =>
2o io4-
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O —
o § 103
O o
S 102
1 -
10
10°
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0)
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0 -
-0.5 -
-1 '
O)
o
O
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(D CO
-3.5 -
-4
-4.5
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Preliminary
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Spring
Summer
-------
Preliminary
Spring
Summer
Winter
o>
8
_I CD
c *
.0 o
£0 CJJ
•££
CD CD
C ®
O £
O
oo
Method 1602PFU
qPCR-CE
Figure 10. Overall Comparison of the Cumulative Log,0 Reduction in EPA Method 1602
F+ Coliphage PFUs (O) and Quantitative Polymerase Chain Reaction (qPCR) Cell
Equivalents (•) Through the Wastewater Treatment Process (These values are based on
the use of 56 samples for each method, 16 samples each for Mill Creek and Muddy Creek,
and 12 samples each for Little Miami and Polk Run)
-------
Preliminary
Spring
Summer
Winter
-O—Method 1603 CPU
qPCR-CE
Figure 11 .Overall Comparison of the Cumulative Log10 Reduction in EPA Method
1603 E. co//CPUs (O) and Quantitative Polymerase Chain Reaction (qPCR)
Cell Equivalents («) Through the Wastewater Treatment Process (These values are
based on the use of 56 samples for each method, 16 samples each for Mill Creek
and Muddy Creek, and 12 samples each for Little Miami and Polk Run)
-------
Preliminary
Spring
Summer
Winter
3?
/ /
—O-B. frailisCFU
o
VO
Figure 12. Overall Comparison of the Cumulative Log10 Reduction in the Bacteroides fragilis
Method CPUs (O) and Quantitative Polymerase Chain Reaction (qPCR) Cell Equivalents
(CE) (•) Through the Wastewater Treatment Process (These values are based on the use
of 5Q samples for each method, 16 samples each for Mill Creek and Muddy Creek, and 12
samples each for Little Miami and Polk Run)
-------
Preliminary
Spring
Summer
Winter
CD
2)'5r
1 c?
^ CD
SCO
D) >„
5 1
C ^-
.2 0
«+-* -a-*
J5 CD
"c •—
® "co
c "®
o C£
O
0.5 -i
0 -
-0.5 -
-1 -
-1.5 -
-2
-2.5 -
-3 J
• C/. perfringens CPU
•qPCR-CE
Figure 13. Overall Comparison of the Cumulative Log10 Reduction in Clostridium perfringens
Method CPUs (O) and Quantitative Polymerase Chain Reaction (qPCR) Cell Equivalents (CE)
(•) Through the Wastewater Treatment Process (These values are based on the use of 56
samples for each method, 16 samples each for Mill Creek and Muddy Creek, and 12 samples
each for Little Miami and Polk Run)
-------
10s
^ to4
8
c ^
| 2 103
"c ,_
-------
E
o
o
O LU
'
§8
o ^
CD
Q.
LU
O
106i
104-
103
102-
V*-
| !
0
b-
"S,
1 1 1 1
1246
Preliminary
o
01246
Spring
-O— Method 1602 PFU
-•— qPCR-CE
01246 0124
Summer Winter
6
10°
Number of Days the Sample was
Held at Seasonal Temperatures
Figure 15. Comparison of EPA Method 1602 F+ Coliphage PFUs (O) and Quantitative
Polymerase Chain Reaction (qPCR) Cell Equivalents (•) From Disinfected, Secondary
Effluent Holding Studies
vo
-------
O
0)
O —v
'•§ u- 103
^_» c_?
0) O
Ji102
O o
d> in1 -I
Q_
LJJ
o,
10°
-o- Method 1603
-•— qPCR-CE
i i i i i
01246
Preliminary
i
CFU
i i i i i
01246
Spring
—i— i i i i i i
01246
Summer
i i i i
0124
Winter
i
6
Number of Days the Sample was
Held at Seasonal Temperatures
Figure 16. Overall Comparison of EPA Method 1603 E. coli CPUs (o) and Quantitative
Polymerase Chain Reaction (qPCR) Cell Equivalents (• ) From Disinfected, Secondary
Effluent Holding Studies
-------
Number of Days the Sample was
Held at Seasonal Temperatures
0.5 T
Preliminary
01246
Spring
1246
Summer
1246
Winter
01246
Method 1600 CPU
qPCR-CE
VO
-2 -1
Figure 17. Overall Cumulative Log10 Reduction of Method 1600 Enterococcus CPUs (O) and
QPCR Cell Equivalents (•) in the Effluent Holding Study
-------
Number of Days the Sample was
Held at Seasonal Temperatures
0.5
-2
Preliminary
01246
Spring
01246
Summer
01246
A
•Method 1602PFU
qPCR-CE
Figure 18. Overall Cumulative Log10 Reduction of Method 1602 F+ Coliphage
PFUs (o) and qPCR Cell Equivalents (•) in the Effluent Holding Study
-------
Preliminary
0.2 i 0 1 2 4 6
Number of Days the Sample was
Held at Seasonal Temperatures
Spring
01246
Summer
01246
Winter
0124
Method 1603 CPU
qPCR-CE
Figure 19. Overall Cumulative Log10 Reduction of Method 1603 £ coli
CFUs (o) and qPCR Cell Equivalents (•) in the Effluent Holding Study
VO
-------
SJ
LLJ
O
108-
107-|
1
8
.1 B
2 o 1°5
1Q3-
102
10°
Method 1600 CFU
qPCR-CE
oo
VO
01246
Preliminary
—i 1 1 1 r
01246
Spring
-i 1 1 r
01246
Summer
0124
Winter
Number of Days the Sample was
Held at Seasonal Temperatures
Figure 20. Overall Comparison of EPA Method 1600 Enterococcus CPUs (o) and Quantitative
Polymerase Chain Reaction (qPCR) Cell Equivalents (•) From Simulated Recreational Water
Holding Studies Using 5% Wastewater Effluent in Ohio River Water
-------
108i
1Q7
106-
o
o
.2
ft o
§1
O o
105
104
LU
O 102
10°
• Method 1602 PFU
•qPCR-CE
01246
Preliminary
01246
Spring
01246
Summer
01246
Winter
Number of Days the Sample was
Held at Seasonal Temperatures
Figure 21. Overall Comparison of EPA Method 1602 F+ Coliphage PFUs (O) and
Quantitative Polymerase Chain Reaction (qPCR) Cell Equivalents (•) From Simulated
Recreational Water Holding Studies Using 5% Wastewater Effluent in Ohio River Water
-------
I
.1
i o
§1
O o
LU
o
108
107
106
105
104
103
10' 1
10M
•Method 1603 CPU
•qPCR-CE
01246
Preliminary
01246
Spring
01246
Summer
1246
Winter
Number of Days the Sample was
Held at Seasonal Temperatures
Figure 22. Overall Comparison of EPA Method 1603 E. co//CPUs (O) and Quantitative
Polymerase Chain Reaction (qPCR) Cell Equivalents (•) From Simulated
Recreational Water Holding Studies Using 5% Wastewater Effluent in Ohio River Water
-------
0)
O)
c
CD
D) Q
II
2 ji
"c <»
1.5T
1 --
0.5--
o
O
-0.5 -
.1 --
-1.5-
Preliminary
01246
Number of Days the Sample was
Held at Seasonal Temperatures
Spring
01246
Summer
01246
Method 1600 CPU
qPCR-CE
Winter
01246
Figure 23. Overall Cumulative Log10 Reduction of Method 1600 Enterococcus
CPUs (o) and qPCR Cell Equivalents (•) in the 5% Wastewater Holding Study
-------
-1.5
Preliminary
01246
Number of Days the Sample was
Held at Seasonal Temperatures
Spring
01246
Summer
01246
Winter
01246
-0
(N
•Method 1602PFU
•qPCR-CE
Figure 24. Overall Cumulative Log10 Reduction of Method 1602 F+ Coliphage
PFUs (o) and qPCR Cell Equivalents ( •) in the 5% Wastewater Holding Study
-------
1.5 n
-1.5 J
Preliminary
01246
Number of Days the Sample was
Held at Seasonal Temperatures
Spring
01246
Summer
01246
•Method 1603 CPU
•qPCR-CE
Winter
01246
Figure 25. Overall Cumulative Log10 Reduction of Method 1603 £ coll
CPUs (O) and qPCR Cell Equivalents («)in the 5% Wastewater Holding Study
-------
105n
fo
0) O
§1
O o
LLI
O
,4.
10
E
o
o
o ;D 103
102-
10°
•Method 1600 CPU
•qPCR-CE
01246
Preliminary
01246
Spring
01246
Summer
01246
Winter
Number of Days the Sample was
Held at Seasonal Temperatures
Figure 26. Overall Comparison of EPA Method 1600 Enterococcus CPUs (O) and
Quantitative Polymerase Chain Reaction (qPCR) Cell Equivalents (•) From Simulated
Recreational Water Holding Studies Using 20% Wastewater Effluent in a Sample
Containing 20% Ohio River Water and 60% Partially-Treated Drinking Water
-------
E
o
o
.2 Z>
o
-------
E
o
o
-------
Number of Days the Sample was
Held at Seasonal Temperatures
-------
Number of Days the Sample was
Held at Seasonal Temperatures
1 n
0.5-
-1.5 J
Preliminary
01246
Spring
01246
Summer
01246
Winter
01246
Method 1 602 PFU
qPCR-CE
oo
Figure 30. Overall Cumulative Log10 Reduction of Method 1602 F+ Coliphage
PFUs (o) and qPCR Cell Equivalents (•) in the 20% Wastewater Holding Study
-------
Number of Days the Sample was
Held at Seasonal Temperatures
0.5 i
Preliminary
Spring
01246
Summer
01246
Winter
01246
Method 1603 CPU
qPCR-CE
_2 J
Figure 31. Overall Cumulative Log10 Reduction of Method 1603 £ co//CFUs (O) and
qPCR Cell Equivalents (•) in the 20% Wastewater Holding Study
-------
o
oo
—O-PLAQUE
—a— MPN
A qPCR
——1600
01246
Preliminary
01246
Spring
01246
Summer
0124
Winter
Number of Days the Sample was
Held at Seasonal Temperatures
Figure 32. Overall Comparison of the Enterococcus Quantitative Polymerase Chain Reaction
(qPCR) Method Cell Equivalents (CE) (A), the Enterococcus Culture Method CPUs (Method 1600)
(-) and the Concentrations of Two Enterovirus Methods in a Simulated Recreational Water Con-
taining 5 % Wastewater in Ohio River Water - The two Enterovirus Methods were the
Plaque Assay (PFU) (Q and the CPE-MPN Method (MPN) (a).
-------
Number of Days the Sample was
Held at Seasonal Temperatures
0)
O)
to
O) Q
1.5
Preliminary
01246
§
-------
EPA/600/R-10/149
December 2010
APPENDIX
Table of Contents
Figure Al. Schematic of Mill Creek WTP A-2
Figure A2. Schematic of Muddy Creek WTP A-3
Figure A3. Schematic of Little Miami WTP A-4
Figure A4. Schematic of Polk Run WTP A-5
Sample Collection Protocols A-6
Figure A5. Quantitative Polymerase Chain Reaction (qPCR) Schematic A-29
Figure A6. Ancillary Measurements A-30
Figures A7-A66. Comparison of Results atFour WTPs A-32
A-l
-------
1
tail. CREEK TOEATOEOT PUNT
SHEET 1 OF
Fig. A1. Schematic of Mill Creek WTP, an activated sludge treatment plant in Cincinnati, Ohio that uses chlorination.
-------
RIVER ROAD
aiMMi
SJE:°
MUDDY CREEK
WASTEWATER TREATMENT PLANT
SCALE.
1* - 40"
SHE LI NO.
1 OF 1
Fig. A2. Schematic of Muddy Creek WTP, an activated sludge treatment plant in Cincinnati, Ohio that uses ultraviolet light disinfection.
-------
I I I 111
unit MIAMI
WASTEWATER TREATMENT PUNT
SITE PLAN
SHEET NO.
1 OF 1
Fig. A3. Schematic of Little Miami WTP, an activated sludge treatment plant in Cincinnati. Ohio that uses chlorination.
-------
POLK RUN
WASTEWATER TREATMENT PUNT
SITE PLAN
<
SHEET NO
1 OF I
Fig. A4. Schematic of Polk Run WTP, an activated sludge treatment plant in Cincinnati, Ohio that uses ultraviolet light disinfection.
-------
Sample Collection Protocols
1. Raw Water
Sampling procedure will be the same for all 4 wastewater treatment plants (WWTPs) (Mill Creek, Muddy
Creek, Little Miami, and Polk Run). See Figure 1.
1. Check in at site, where necessary.
2. Go to Raw sample location.
3, Record ambient site observations), including:
a. Date/time
b. Air Temp (°C)
c. Cloud cover (S, MS, C, MC, O)*
d. Rainfall (current conditions, site rain gauge measurement, if available)
e. Photograph sample site
*S, MS, C, MC, O: Sunny; Mostly Sunny (20-50% cloud cover); Cloudy (50-70% cloud cover); Mostly
Cloudy (70-99% cloud cover); Overcast
4, Prepare waste bag.
5. Set up dip sampler with sterile 1 L bottle. (Exception: Mill Creek samples may be collected
directly from sample trough in building).
6. Spread out ground cloth if ground surface is wet or dirty. Lay out bottles conveniently near
sample location.
7. Collect samples by dipping sterile 1 L bottle, pouring off into sample bottles:
Raw Water sample summary:
Volume Number Purpose
500 mL 2 TSS, field parameters (wide-mouth)
1L 2 micro parameters
8, From the wide mouth 500 mL (field parameter) bottle, measure field parameters: pH,
conductivity, water temperature, and turbidity. (See instrument instructions in sampling
documents.)
9. Record field parameters on field data sheet. Pour remaining field sample back into waste
stream and discard empty bottle into the waste bag.
10, Clean outer surface of remaining 500-mL and 1-L sample bottles with sanitizing wipes and place
in a Ziploc bags.
11. Place bagged bottles in cooler labeled "Raw-Primary samples". Add an ice bag from ICE cooler.
12. Remove dipping bottle (if used) from pole and discard in waste bag.
13. Clean dip sampler with sanitizing wipe and rinse with tap water from spray bottle.
14. Complete chain-of-custody form.
15. Check field sheets for completeness before leaving the site.
PageB-2ofB-189
A-6
-------
September 7, 2009 version
2. Primary Effluent
Sampling procedure will be the same for all 4 WWTPs (Mill Creek, Muddy Creek, Little Miami, and Polk
Run). See Figure 1.
I. Go to Primary effluent sample location.
2, REFERENCE RAW FIELD SHEET, OR: Record ambient site observations), including;
a. Date/time
b. Air temp (°C)
c. Cloud cover (S» MS, C, MC, Oj*
d. Rainfall (current conditions, site rain gauge measurement, if available)
e. Photograph sample site
*S, MS, C, MC, O: Sunny; Mostly Sunny (20-50% cloud cover); Cloudy (50-70% cloud cover); Mostly
Cloudy (70-99% cloud cover); Overcast
3. Set up dip sampler with sterile 1 L bottle by securing to pole with cable tie and tape.
4. Spread out ground cloth if ground surface is wet or dirty. Assemble sample bottles conveniently
near sample location.
5. Collect samples by dipping sterile 1 L bottle, pouring off into sample bottles:
Primary Effluent sample summary:
Volume Number Purpose
500 ml 2 TSS, field parameters (wide-mouth)
1L 3 micro parameters, EPA
6. From the wide mouth 500 mL (field parameter) bottle, measure field parameters: pH,
conductivity, water temperature, and turbidity. (See instrument instructions in sampling
documents.)
7. Record field parameters on field data sheet. Pour remaining field sample back into waste
stream and discard empty bottle into the waste bag,
8. Clean outer surface of remaining 500-mL and 1-L sample bottles with sanitizing wipes and place
in a Ziploc bags.
9. Place bagged bottles in cooler labeled "Raw-Primary samples". Add more ice bags or loose ice
from ICE cooler, if needed.
10. Remove bottle from dipping pole and discard in waste bag. Clean dip sampler with sanitizing
wipe and rinse with tap water from spray bottle.
11. Designate one team member to proceed to secondary effluent sites to photograph locations.
12. Complete chain-of-custody form.
13. Check field sheets for completeness before leaving the site.
PageB-3of 6-189
A-7
-------
September 7, 2009 version
Figure 1. Raw and Primary Effluent Sampling
Raw or Primary Effluent Stream
PageB-4ofB-189
A-8
-------
September 7, 2009 version
3. Secondary Effluent Pre Disinfection
Sampling procedure will be the same for all 4 WWTPs (Mill Creek, Muddy Creek, Little Miami, and Poik
Run}.
1. Check in at site, where necessary.
2. Go to Secondary Effluent, Pre-Dtsinfectiort sampling location,
3. Record ambient site observations), including:
a. Date/time
b. Air temp (*C)
c. Cloud cover (S, MS, C, MC, O}»
d. Rainfall (current conditions, site rain gauge measurement, if available)
e. Photograph sample site
•S, MS, C, MC, O: Sunny; Mostly Sunny (20-50% cloud cover); Cloudy fSO-70% cloud cover); Mostly
Ctoudy (70-99% cloud cover); Overcast
4, Spread out ground cloth if ground surface is wet or dirty. Assemble sample bottles, filters,
containers conveniently near sample location,
5. Label and ready sample bottles, filters, containers.
6. Unpack pump feed tubing (reinforced, with foot valve) and recirculating (recirc)/flush tubing.
7. See Figure 2A. Secure tubing (to railing and/or pump) and lower feed tubing into sample
stream, away from channel walls.
8, Pump has been primed. Tip pump up to remove cap from suction port, then connect feed tubing
to pump inlet. Run electric power to pump from nearest outlet (in or on disinfection buildings).
Extension cord may be required.
9. Connect Recirc/Flush line to small ball valve on pump TEE.
10. Crack open recirc/flush valve (small ball valve on pump Tee). Close manifold valve (large ball
valve). Turn on pump, slowly opening flush valve, and directing discharge downstream of feed
tubing/foot valve. *IF PUMP HAS LOST PRIME, ADD DISTILLED WATER SUPPLIED WITH
SAMPLING MATERIALS. Add through sterile plug cap or directly into pump discharge outlet.
11. See Figure 28. Unpack manifold, set up on a waste container for support, and assemble
remaining 4 waste containers around manifold.
12. Plumb manifold to pump discharge, using washing machine hose supplied.
13. Install discharge tubing on all manifold ports, directed back to waste stream, downstream of
sample collection site. Use barbed connectors in place of filters during sample port flushing.
14. Turn on pump and set flows through the manifold to rate appropriate to sample collection;
• 2 CUNO 1MDS Virus filter branches: 4 L/min ea. Sample volume: 100 L
each
• 3 Pall Envirochek HV filter branches: 2 L/min ea. Sample volume: 50 L
each
• Bulk 10 L carboy: 0.4 L/min
Page B-5 of B-189
A-9
-------
September 7, 2009 version
15. Allow all manifold ports to flush 4 min, directing discharges to one of the waste containers.
After 4 min, collect two 5QO-mt samples (for T5S and field parameters} from one of the sample
discharge lines,
16. After flushing, empty the container well downstream of sample intake line.
17. Add household bleach to the four large volume-measuring waste containers: Add 100 ml
bleach to each large (100 L| volume-measuring container downstream of cartridge filter pairs,
and 50 ml bleach to each low container (50 L) capturing capsule discharges.
18. Turn off pump. Install filters on appropriate manifold ports. PLACE ALL FILTER END CAPS IN
CLEAN BAG AND KEEP HANDY.
19. Direct filter discharge lines back to volume containers.
20. Attach 54" tubing to carboy cap fitting. Keep barbed fitting cap handy.
21. Loosen carboy cap to allow venting during sampling.
22. Turn on pump, adjust flows as necessary.
23, While sampling, measure field parameters: pH, conductivity, water temperature, and turbidity
from the wide mouth Q.S-L field parameter bottle {See instrument instructions in sampling
documents).
24. Record field parameters on field data sheet. Pour remaining field sample back into waste
stream and discard empty bottle into the waste bag.
25. Collect samples to volumes required. Shut off pump.
26. Record volumes filtered on sample sheet.
27. Separate virus filter pairs. Turn upside down to drain. Cap filter housings. Pack the 2 pre-filters
and 2 virus filters in one shipping cooler. Add layer of bubble pack, and place four ice packs
from ICE cooler on top.
28, Prepare carboy for delivery to project laboratory: place in one of the SO L bins and add one ice
bag from cooler.
29. Pack HV filters and 500 mL TSS sample in ice cooler with remaining ice bag for delivery to project
laboratory.
30. Fill out chain of custody forms.
31. Place sample tube/foot valve in 25 gallon container of chlorinated filtered sample water
remaining in sample filter discharge containers. Turn on pump and pump water through sample
assembly, from foot valve through pump, all manifold branches and discharge lines.
32, Pump out remaining containers through manifold or directly to waste using bypass line, or
empty into secondary effluent channel.
33. Cap pump, and pack manifold and other equipment.
34. Check field sheets for completeness before leaving the site.
35. Proceed to Post disinfection site.
PageB-€ofB-189
A-10
-------
September 7, 2009 version
Figure 2A. Secondary Effluent Pre-Disinfection Pumping and Polk Run POST UV
To Manifold
(WASHING MACHINE HOSE)
BAGD
Flush line
(3/4" TUBING)
(BagE)
PUMP
DISCHARGE
TEE
BAGC
PUMP
Pump Feed {3/4 reinforced tubing)
(BAG B)
channel
I
Foot valve
PageB-7ofB-i89
A-ll
-------
September 7,2009 version
Figure 28. Secondary Effluent Pre-Disinfection Sampling
Secondary PRE
Disinfection
It systems needed:
All 4 plants secondary pre-disinfection
HV fitters
80
Collect 0.5 LTSS
and 0.5 L field parameter samples
after manifold flush
Secondary waste
stream
Pre Disinfection
CF—
Booster pump
1/4" tubing
BAGG
1/4" tubing
0.4 L/mm
10 L carboy
bottom dispensing
8
Target
Flow rate Volume Waste tubing
(L/min) (Liters)
50 L 1/2" plastic
2.0 SOL
2.0 SOL
.0 100 L
1/2" plastic
1/2" plastic
1/2" plastic
Virus Cartridge filters (2 pair)
100 L
1/2" plastic
Bag F
Page B-8 of B-189
A-12
-------
September 7, 2009 version
4. Secondary Effluent
4.1.A Post UV Disinfection: Muddy Creek
See Figure 3A.
1. Check in at site, where necessary.
2. Go to Secondary Effluent, Post-UV sampling location.
3. Record ambient site observations, including:
a. Date/time
b. Air temp (8C)
c. Cloud cover (S, MS, C, MC, O) *
d. Rainfall (current conditions, site rain gauge measurement, if available)
e. Photograph sample site (or verify that other team member(s) photographed the site)
*S, MS, C, MC, O: Sunny; Mostly Sunny (20-50% cloud cover); Cloudy (50-70% cloud cover); Mostly
Cloudy (70-99% cloud cover); Overcast
4. Label and ready sample bottles, filters, containers.
5. Unpack washing machine hose and manifold.
6. Install sample equipment on outlet of ball valve at sampling location. Open this valve (sample
valve) to flush sample line. Throttle main flow pipe to provide more flow to sample line, but be
sure flow to site instruments is maintained. Shut sample vatve.
7. Install discharge tubing on all manifold ports, directed back to sample trough. Use barbed
unions in place of filters during flushing.
8, Open sample ball valve and set flows through the manifold to rate appropriate to sample
collection:
Type Flow Rate Sample Volume
2 CUNO 1MOS Virus
filter pairs
2 Pall Envlrocheck HV
capsules
Bulk 20 L carboy
4 L/min ea.
2 L/min ea.
0.8 L/min
100 L each
50 L each
201
9, Allow all manifold ports to flush 4 min. If flow is not sufficient, throttle large ball valve below
and to the right of the sample trough. Maintain some flow to site instruments.
10. After 4 min, collect two 0.5 L samples (for TSS and field parameters) from one of the sample
discharge lines.
11. Add household bleach to the four large volume-measuring waste containers: Add 100 mL
bleach to each large (100 L) volume-measuring container downstream of cartridge filter pairs,
and 50 mL bleach to each low container (501) capturing capsule discharges.
12. Turn sample valve (and pump, if used) off.
13. Install filter housings/filters on appropriate manifold ports. PLACE ALL FILTER END CAPS IN
CLEAN BAG AND KEEP HANDY.
PaieB-9ofB-189
A-13
-------
September 7, 2009 version
14. Direct filter discharge lines back to volume containers
15, Attach %" tubing to carboy cap fitting. Keep barbed fitting cap handy.
16. Loosen carboy cap to allow venting during sampling.
17. Turn sample valve on and adjust flows as necessary. If any filter leg flow rates decrease to the
point the target rate cannot be maintained by opening the flowmeter valve(s), decrease flows
to all legs proportionally (so that simultaneous sampling of all sample legs is achieved).
18. While sampling, measure field parameters: pH, conductivity, water temperature, and turbidity
from the wide mouth 500 ml field parameter bottle (See instrument instructions in sampling
documents).
19. Record field parameters on field data sheet. Pour remaining field sample back into waste
stream and discard empty bottle into the waste bag.
20. Collect samples to volumes required. Shut off sample valve.
21, Separate virus filter pairs. Turn upside down to drain. Cap filter housings. Pack the 2 pre-fllters
and 2 virus filters in one shipping cooler. Add layer of bubble pack, and place four ice packs
from ICE cooler on top.
22. Prepare carboy for delivery to project laboratory: place in a 50 L bin and add one ice bag from
cooler.
23. Pack HV filters and 500 ml TSS sample in Ice cooler with remaining ice bag for delivery to project
laboratory.
24. Fill out chain of custody forms.
25. Empty waste containers to secondary stream by siphon or dumping,
26. Pack equipment in labeled containers for storage.
27. Check field sheets for completeness before leaving the site.
10
PageB-IOofB-189
A-14
-------
September 7, 2009 version
Figure 3A. Secondary Effluent Post UV Sampling at MUDDY CREEK
POST UV
Secondary effluent lamping
PostUV
from sampling trough
In secondary building
Hook up wrtn tubing
clamped to 1.2" pipe
80
Collect 0.5 L TSS
and 0.5 L field parameter samples
after manifold flush
Flow rate (Uminj
M
2.0 SO L total volume
-•>
HV Men 121
20 50 L total volume
-*•
4[ 100 L total volume
— Virus Cartridge Filers (Z pair)
4.0 100 L lotal volume
0.8
L carboy
for holding ttudy, EPA.
and all micro samples
11
PageB-11 ofB-189
A-15
-------
September 7, 2009 version
4.IB Post UV Disinfection POLK Run
S@e Figures 2A 38,
1. Check in at site, where necessary.
2. Go to Secondary Effluent, Post-UV sampling location.
3. Record ambient site observations, including;
a. Date/time
b. Air temp (*C)
c. Cloud cover (S, MS, C, MC, O)*
d. Rainfall (current conditions, site rain gauge measurement, if available)
e. Photograph sample site (or verify that other team member(s) photographed the site)
*S, MS, C, MC, O: Sunny; Mostly Sunny {20-50% cloud cover); Cloudy (50-70% cioyd cover); Mostly
Cloudy (70-99% cloud cover); Overcast
4. Label and ready sample bottles, filters, containers.
5. Unpack washing machine hose and manifold.
i. Install sample equipment on outlet of ball valve at sampling location. Open this valve (sample
valve) to flush sample line. Throttle main flow pipe to provide more flow to sample line, but be
sure flow to site instruments is maintained. Shut sample valve,
7. Install discharge tubing on all manifold ports, directed back to sample trough. Use barbed
unions in place of filters during flushing.
8. Open sample ball valve and set flows through the manifold to rate appropriate to sample
collection:
Tvoe FlojMRajte Sample Volume
2 CUNO1MDS Virus 4L/mlnea. 100 Leach
filter pairs
2 Pall Envirocheck HV 2 L/min ea. 50 L each
capsules
Bulk 20 L carboy 0.8 L/min 20 L
9. Allow all manifold ports to flush 4 min. If flow is not sufficient, throttle large ball valve below
and to the right of the sample trough. Maintain some flow to site instruments.
10. After 4 min, collect two 0.5 L samples (for TSS and field parameters) from one of the sample
discharge lines.
11. Add household bleach to the four large volume-measuring waste containers: Add 100 mL
bleach to each large {100 L) volume-measuring container downstream of cartridge filter pairs,
and 50 mL bleach to each low container (50 L) capturing capsule discharges.
12. Turn sample valve (and pump, if used} off.
13, Install filter housings/fitters on appropriate manifold ports, PLACE ALL FILTER END CAPS IN
CLEAN BAG AND KEEP HANDY.
14, Direct filter discharge lines back to volume containers
12
Page B-l2of 8-189
A-16
-------
September 7, 2009 version
15. Attach K" tubing to carboy cap fitting. Keep barbed fitting cap handy.
16. Loosen carboy cap to allow venting during sampling.
17. Turn sample valve on and adjust flows as necessary, if any filter leg flow rates decrease to the
point the target rate cannot be maintained by opening the flowmeter valve{s)» decrease flows
to all legs proportionally (so that simultaneous sampling of all sample legs is achieved).
18. While sampling, measure field parameters: pH, conductivity, water temperature, and turbidity
from the wide mouth 500 ml field parameter bottle (See instrument instructions in sampling
documents).
19. Record field parameters on field data sheet Pour remaining field sample back into waste
stream and discard empty bottle into the waste bag.
20. Collect samples to volumes required. Shut off sample valve.
21. Separate virus fitter pairs. Turn upside down to drain. Cap filter housings. Pack the 2 pre-fitters
and 2 virus filters in one shipping cooler. Add layer of bubble pack, and place four ice packs
from ICE cooler on top.
22, Prepare carboy for delivery to project laboratory: place in a 50 L bin and add one Ice bag from
cooler.
23. Pack HV filters and 500 mL TSS sample in ice cooler with remaining ice bag for delivery to project
laboratory.
24. Fill out chain of custody forms.
25. Empty waste containers to secondary stream by siphon or dumping.
26. Pack equipment in labeled containers for storage.
27. Check field sheets for completeness before leaving the site.
13
PageB-13of B-189
A-17
-------
September 7, 2009 version
Figure 38. Secondary Effluent Post UV Sampling at POLK RUN
Secondary Post UV
Polk Run Only Target
Flow rate Volume Waste tubing
(Urn in) (Liters)
HV filtere
n«2
80
Collect 0.5 L TSS
and O.S L field parameter samples
after manifold flush
-*•
Secondary wasU
stream
• •
Booster pump
1/4" tubing
2.0 SOL
-*•
2.0 50 L 1/2- tubing in Bag F
4.0 100 L
Virus Cartridge fitters (2 pair)
4.0 100 L
1/4" tubing in Bag G
0.4 L/min
10 L carboy
bottom dispensing
14
PageB-14ofB-l89
A-18
-------
September 7, 2009 version
4.2. Post Chlorine Disinfection: Mill Creek and Little Miami
See Figure 4A and Figure 4B.
1. Go to Secondary Post Chlorine sample location,
2. Record ambient site observations), including:
a. Date/time
b. Air temp (*C)
c. Cloud cover (S, MS, C, MC. 0}*
d. Rainfall (current conditions, site rain gauge measurement, if available)
e. Photograph sample site (or verify that other team member(s) photographed site
*S, MS, C, MC, O: Sunny; Mostly Sunny (20-50% cloud cover); Cloudy (50-70% cloud cover); Mostly
Cloudy (70-99% cloud cover); Overcast
3. Set 100 gallon tank near sample port in Secondary treatment building (Mill Creek) or in hall
outside sample room (Little Miami), Plumb hose to sample port and flush sample plumbing to
drain for 1 minute.
4. Direct sample hose from sample port to 100 gallon tank.
5, Open valve to fill tank.
6. As tank fills, set up sample/recirculation loop consisting of tubing, pump and manifold,
7. Plumb discharge lines to drain manifold. Direct drain manifold discharge back to tank.
8. Begin recirculation with recirc loop and all legs of manifold open and returning to tank.
9, When tank is full, close sample valve.
10. Collect the samples not requiring dechlorination:
f. Collect the 500 ml TSS sample. Note on field data sheet and CEC Chain of Custody form
g. Collect 500 mL field sample (wide-mouth bottle) and measure field parameters; total
chlorine, pH, conductivity, water temperature, and turbidity.
h. Record field parameters on field data sheet. Pour remaining field sample back into
waste stream and discard empty bottle into the waste bag.
11. Add 100 mL sodium thiosulfate solution to tank.
12. Allow dechlorinated water to recirc/mix for S minutes. Measure total chlorine to confirm no
residual present at pump/manifold outlet.
13. Do not waste water from tank as It contains just more than will be needed for all sample
volumes.
14. Add household bleach to the four large volume-measuring waste containers: Add 500 ml
bleach to each large (100 L) volume-measuring container downstream of cartridge filter pairs,
and 250 mL bleach to each low container (50 L) capturing capsule discharges,
15, Valve off manifold, and disconnect tubing from drain manifold.
16. Attach M" tubing to carboy cap fitting. Keep barbed fitting cap handy.
17, Loosen carboy cap to allow venting during sampling.
15
PageB-lSofB-189
A-19
-------
September 7, 2009 version
18. Install 2 HVs and 2 virus filters on manifold legs.
19. Direct HV drain lines to low containers (SO L).
20, Direct virus drain lines to large containers (100 L)
21. Begin flow through filters and to carboy. Direct discharge of filters to volume-measuring
containers.
22. Throttle redrc/rrtix loop if necessary to boost pressure to filters.
Type How Rate Sample Volume
2CUN01MDSVirus 4L/minea. 100 Leach
filter pairs
2 Pal! Envirocheck HVs 2 L/min ea. 50 L each
Bulk 20 L carboy 0.8 L/min 20 L
23. Turn off pump when appropriate volumes have been collected.
24. Separate vims filter pairs. Turn upside down to drain. Cap filter housings. Pack the 2 pre-fitters
and 2 virus filters in one shipping cooler. Add layer of bubble pack, and place four ice packs
from ICE cooler on top.
25. Prepare carboy for delivery to project laboratory: place in a 50 L bin and add one ice bag from
cooler.
26. Pack HV filters and SQQ mL TSS sample in ice cooler with remaining ice bag for delivery to project
laboratory.
27. Fill out chain of custody forms.
28. Place pump feed tubing and foot valve in volume-measuring container (garbage can) of 100 L
filtered virus filtrate.
29. Begin pumping and allow chlorinated water to flow through manifold assembly. Cap pump,
manifold, and pack equipment in labeled containers for storage.
30. Pump out remaining containers through manifold or directly to waste using recirc line.
31. Remove tank liner and place in field trash bag. Discard the entire field trash bag in one of the
covered red bins marked as "haiardous waste* at the project laboratory.
32. Check field sheets for completeness before leaving the site.
16
Page B-18 of B-189
A-20
-------
September 7, 2009 version
Virus Sampling Directions
a) After adjusting the flow rates, shut off flow.
b) Remove brass union from between manifold branch and flow meter,
c) Remove the foil from each end of the prefilter module and connect the prefilter module to the
manifold, noting flow direction arrow,
d) Remove the foil from the female x female adapter and connect to the discharge of the prefilter.
e) Remove foil from each end of the virus fitter module and connect it to the prefilter with the
previously installed adapter,
f) Connect flow meter to discharge of virus filter.
g) Direct discharge line to volume—measuring waste container,
h) Label capsules with sharpie pen.
i) Slowly turn on water and establish desired flow rate (4 L/min).
j) Monitor flow rate during sampling and adjust as needed.
k) Sample until volume in waste containers equals the target volume (100 L),
I) Turn off sample pump, (ff target volume is not attained in 1 hour, shut off pump to end
sampling and note volume sampled on field sheet.)
m) Record volume sampled on field sheet.
n) Loosen the swivel connections and remove both modules from manifold.
o) Turn both housings ypside down and allow excess drain water to flow out as waste water.
p) Turn housings upright.
q) Remove endcaps from foil wrapping and cap both ends of both housings.
r) Place in cooler.
17
PafeB-17 of 8-188
A-21
-------
September 7, 2009 version
Envirochek HV Sampling Directions
1. Open package and remove capsules. Leave end-caps in package and set aside.
2. Label capsules [or apply pre-pnnted label] with sharpie pen. Use same 10 as on Field Sheet.
3. Identify inlet and outlet ends.
4. Insert Inlet end into M* tubing on 2 L/min flowmeter.
5. Insert outlet end into ¥t" tubing directed to waste container.
6. When flow begins, set flowmeter to 2 L/min. Monitor and adjust during filtration to maintain 2
L/min flow rate.
7. After 50 L have been filtered, shut off pump and close flowmeter vaive,
8. Remove capsule from tubing without allowing water to exit inlet port. Place end-caps on inlet
and outlet ports.
9. Record on the filter the exact volume filtered.
10. Place capsule In ZipLoc bag and place in cooler,
11. Do not allow ice packs in cooler to contact capsule bags.
18
PageB-iaofB-189
A-22
-------
September 7, 2009 version
Figure 4A: Tank Sample Plumbing for Post Chlorine at Mill Creek and Little Miami.
SITE SAMPLE PIPE
PVC
Sample line
Hook tip at
3/4 MPT below
Ibali valve
r
MILL CREEK
POST CHLORINE
SAMPLE SET UP
To Manifold
(WASHING MACHINE HOSE)
BAG"D"
3/4" tubing
Tubing bag
"A"
First:
Flush to drain
then to tank
BagE
REC1RC LINE
, (3/4" TUBING)
PUMP
DISCHARGE
TEE
BAGC
PUMP
FEED
from tank
(3M- tubing, Bag "B")
I
Foot valve
19
Page B-19 of B-189
A-23
-------
Sept 7, 2009 version
Figure 48. Secondary Effluent Post Chlorine Disinfection at Mill Creek and Little Miami.
POST Chlorine
Recirc/Mixing/Sampling Method
Recirc/mix loop
Secondary waUe stream
(Post disinfection)
N-
378 Liter
100 gallon
tank
i
i
•ancillary samples pulled pre-dechlor
•- ---
pIQ'TSSand
LJ Ufield paramet
'"N.
i
Waste tubing return (during manifold flush only)
•------+-
{^) Flow rate Target
(Umin) Volume
irs (Liters)
HV filters 1 1/2" plastic
1 IK 1 I BagF
— ^ f^j * 2 Umin SO L
KT~|~^I ^ 4 Umin 100 L
IXlTvl 1 1/2" plastic
\J NJ | BagG
Virus C jpture filters
1 IT~1 K/j j 4 Umin 100 L
r*"'* 0.8
*t Umin 20 L
"^k carboy
1/4" tubing in Bag H
(N
<
micro samples, holding study, EPA
20
PageB-20ofB-189
-------
Sept 7, 2009 version
5. Ohio River at Greater Cincinnati Water Works Pumping
Station
See Figure 5.
1. Check in at Entrance Gate.
2. Go to Pumping station sample location.
3. Record ambient site observations), including;
a. Date/time
b. Air temp TO
c. Cloud cover (S, MS, C, MC, 0)*
d. Rainfall (current conditions, site rain gauge measurement, if available)
e. Photograph sample site
*S, MS, C, MC, O: Sunny; Mostly Sunny (20-50% cloud cover); Cloudy (50-70% cloud cover); Mostly
Cloudy (70-99% cloud cover); Overcast
4. Flush sample tap through extra hose to drain sink on sample panel.
5. Unpack sample hose and manifold.
6, install sample equipment on outlet of ball valve at sampling location.
7, Install discharge tubing on ail manifold ports, directed back to sample trough. Use barbed
unions in place of filters during flushing.
8, Open sample ball valve and set flows through the manifold to rate appropriate to sample
collection:
Type Flow Rate Sample Volume
2CUN01MDSVirus 4L/minea. 100 Leach
filter pairs
Bulk 20 L carboy #1 0.8 I/mm 20 L {blending study #1)
Bulk 20 L carboy #2 0,8 L/min 20 L (blending study #2)
Bulk 20 L carboy #3 0.8 L/min 20 L (EPA sample)
*This is onty needed on one day of each season or dry run)
Bulk 20 L carboy 0.8 L/min 20 L (Bulk micro samples)
9, Allow all manifold ports to flush 4 min,
10. After 4 min flush, collect TSS and field parameter samples in 500-mL bottles. Note turbidity at
on-site meter, and record on field sheet.
11. Close sample valve.
12. Measure chemical parameters out of 500-mL wide mouth field parameter bottle: pH,
conductivity, and water temperature.
13. Record field parameters on field data sheet. Pour remaining field sample back into waste
stream and discard empty bottle into the waste bag.
14. Install filter housings/filters on appropriate manifold ports.
21
PageB-21 of 8-189
A-25
-------
Sept 7, 2009 version
15. Direct discharge lines back to stream, downstream of 'Sample uptake,
16. Direct other ports to containers,
17. Open sample valve and adjust flows as necessary.
18. Collect samples to ¥olumes required. Shut off pump.
19. Again note turbidity and record.
20. Pack virus filters and 500-mL TSS sample in cooler.
21. Prepare carboy samples for delivery to project laboratory.
22. Cap manifold, and pack equipment in labeled containers for storage.
23. Check field sheets for completeness before leaving the site.
22
PageB.22ofB-189
A-26
-------
Sept 7, 2009 version
Figure 5. Ohio River Sampling Location at GCWW Pump Station.
River intake sampling
at GCWW
to EPA*
Collect at
GCWW Intake
Building
Collect 2 each day
Rec study (spike) Rec study
WTP1 WTP 2
0.8 L/min
20 L
carboy*
Reclrculating mix loop
'[ I K/l ^4.0 100 L total volume
\ *M l/min
Virus Capture filters
L4.0 100 L total volume
tanln
(N
<
0.8
0.8 L/|nin L/min
EC—
-------
Sept 7, 2009 version
6. Treated Ohio River at Greater Cincinnati Water Works
Richard Miller Treatment Plant
Sampling will be performed by River intake sampling personnel.
I, Record ambient site observations), including:
« Date/time
« Air temp (*C)
• Cloud cover (S, MS, C, MC, 0)*
• Rainfall (current conditions, site rain gauge measurement, if available)
• Photograph sample site
*S, MS, C, MC, O: Sunny; Mostly Sunny (20-50% cloud cover); Cloudy (50-70% cloud cover); Mostly
Cloudy (70-99% cloud cover); Overcast
2, At sample location, open sample tap and allow it to flush,
3, After 4 minutes, collect:
Volume Number Purpose
Treated River at GCWW Bulk samples: 500 mL 2 TSS, field
10 L carboys 3 (Bulk micro, blending studies)
4. Measure chemical parameters: pH, conductivity, and water temperature from wide-mouth
field parameter bottle (turbidity read from on-site instrument).
5. Record field parameters on field data sheet. Pour remaining field sample back into waste
stream and discard empty bottle into the waste bag,
6. Prepare samples for delivery to project laboratory.
7. Fill out chain-of-custody form.
8. Check field sheets for completeness before leaving the site.
24
PsgeB-24ofB-189
A-28
-------
Figure A5. Quantitative Polymerase Chain Reaction (qPCR) Schematic
Hut
1 copy of DNA
Separate/denature strands
Attach 2 primers and probe.
Taq polymerase attaches
» DNA replicates, and probe is
| | cleaved and degraded.
Instrument detects free
fluorescent reporter molecule,
and DNA elongates
2 copies of ONA
Key | Attached Fluorescent Reporter Molecule
I Fluorescence Quencher (when attached)
• Free Fluorescent Reporter Molecule
A-29
-------
Figure A6. Ancillary Measurements
-
Measurement
I>*seription
twits/Format MQOs
Dale and Time iDaic anJ I (me of day |mm dd'vy:
!hh:rnm
Air temperature Measured hy thermometer at a fixed location °C
ocrv v i>it
Water temperature
Cloud Cover
Rainfall
Wind speed
Wind direction
Current Direct ion
Boats
Animals''Birds
Debris
UV Light Reading
Measured by thermometer for ambient water at a C'C
fixtd samp! ing location at appropriate depth lor
the thermometer on every \i\it, measured Iitr
waste-water using a container oilier than the
sample bottles to maintain itcrilil} ol the
samples
Sunny. Mostly Sunn> f 20-50 'a cloud covert. S. MS, C, MC. 0
Cloud> t50-70*ot:o\eri Mo-!l> Cloudy |70-W.j
cover). Overcast
Measured by rain gauge near wastewalei Rain in inches,
sampling area, collected each da> Jt time tit Other
sampling and any lime tarn is knows; to have observations
occurred since the last meastrerneni \\as taken noted in
Current conditions, such a& ram. lightning, hail, comments field
etc. noted
Sustained speed measured a? sample collection
site by wind gauge; gusts indicated in comments
fields.
Compass direction to nearest seminjuadMn!
leeward measured on wind gauge.
Described in relation to shoreline while facing
the water.
Number/approximate number of boats in the
water, within approximately 500 M of sampling
area.
Animals and birds potentially affecting the water
(within approximately 20 M of the sampling area
in the water or laterally within 20 M of the
sample location!; also includes number of fowl or
other birds in !he air near the sampling area.
Description of any debris floating in the water or
washed on shore near the sampling location.
Miles per hour.
N KF E SE S
SW. \\.orNW
Descriptive
(onshore, right,
etc.)
Categorical,
None, 1-5, 5-10,
10-20,20-30,
etc.. etc.
Types of
Animals.
Numbers of
Animal Types on
beach and in
water
Categorical;
"None," "Very
Little." "Little,"
"Lots;" describe
types
Measured by ultraviolet i\VTm"
A-30
-------
Measurement
Position
pH
Turbidity
Total Suspended
Solids (TSS)
Conductivity
Description
Coordinates shall be taken where each sample is
collected at each WTP on each visit or trip,
including the dry run,.
Each sample measured ifter microbiological
analysis processing, per "Standard Methods" (3) or
equivalent.
Each simple measured by nephlometer ifter
microbiological analysis processing, per "Standard
Methods" (3) or equivalent ,
Measured by "Standard Methods" (3) from the
samples taken for Enterococci or £ co// (EPA
Methods 1600 and 1603, respectively), after those
analyses are complete.
Each sample measured after microbiological
analysis processing, per "Standard Methods" (3) or
equivalent.
Units/Format
Latitude/Longitude
pH units
Nephlornetric
Turbidity Units
(NTUs)
mg/1
Micro-Siemens or
Midi-Siemens, as
appropriate.
MQOs
Field Person or
Team Consensus.
i 0.2 unils
Range
dependent; see
Standard
Methods 2 1 30 13.
Field Person.
Field Person. i
A-31
-------
1 -Mill CreekWTP
E
o
o
-------
2 - Little Miami WTP
o 3
ffe
cu o
Si
O o
107
106
o
O 1Q5
104-
103
g 102
101.
10°
-O~ Method 1600 CPU
-»— qPCR-CE
' I 1 T I 1 III
cfAA^ c? ^ ^ d>
eg® <& -Jo »«' (ff5
Spring
Summer
Winter
Figure A8. Comparison of the Effect of the Wastewater Treatment Processes at
Little Miami WTP on the Enterococcus Concentrations, Determined Using EPA Method
1600 (o), and the Quantitative Polymerase Chain Reaction (qPCR) Method Cell Equivalents (•)
-------
107
106-
8 10B
c
.0 ID
(U O
§1
O o
1Q4-
103i
UJ 10a
1 .
10
10°
3 - Muddy Creek WTP
Method 1600 CFU
qPCR-CE
/
Preliminary
Spring
Summer
G$
Winter
Figure A9, Comparison of the Effect of the Wastewater Treatment Processes at
Muddy Creek WTP on the Enterococcus Concentrations, Determined Using EPA
Method 1600 (O), and the Quantitative Polymerase Chain Reaction (qPCR) Method
Cell Equivalents (•)
-------
o
o
II
(U O
§1
O o
LU
O
108-
106
10s
104
103
102
10°
/
4 - Polk Run WTP
—O— Method 1800 CPU
-*-qPCR-CE
/ /
4?
<0
Spring
Summer
Winter
Figure A10, Comparison of the Effect of the Wastewater Treatment Processes at Polk Run WTP
on the Enterococcus Concentrations, Determined Using EPA Method 1600 (o), and the
Quantitative Polymerase Chain Reaction (qPCR) Method Cell Equivalents {«)
-------
1 - Mill Creek WTP
o
IS Q
c o
108,
107-
108-
105-
104-
103J
10°
• Method 1602 PFU
qPCR-CE
#
ts° v •Jo «s*
/^ ^./
J
VO
Preliminary
Spring
Summer
Winter
Figure A11. Comparison of the Effect of the Wastewater Treatment Processes at Mill Creek WTP
on the F+ Coliphage Concentrations, Determined Using EPA Method 1602 (o), and the
Quantitative Polymerase Chain Reaction (qPCR) Method Cell Equivalents (•)
-------
2 - Little Miami WTP
o
o
106-
106
S
•43 rr
ca O
^° 1°4
Q "g
S | w
t 10'
1 .
10
10°
—O— Method 1602 PFU
— •— qPCR-CE
Spring
Summer
Winter
Figure A12. Comparison of the Effect of the Wastewater Treatment Processes at Little Miami WTP
on the F+ Coliphage Concentrations, Determined Using EPA Method 1602 (O), and the Quantitative
Polymerase Chain Reaction (qPCR) Method Cell Equivalents (•)
-------
3 - Muddy Creek WTP
Spring
& <
/ /
Summer
oo
Method 1602 PFU
-qPCR-CE
Winter
Figure A13. Comparison of the Effect of the Wastewater Treatment Processes at Muddy Creek WTP
on the F+ Coliphage Concentrations, Determined Using EPA Method 1602 (o), and the Quantitative
Polymerase Chain Reaction (qPCR) Method Cell Equivalents (•)
-------
4 - Polk Run WTP
o
o
!§
£o
9 E
u_
a.
108-,
107-
106-
105-
104
103
10°
V-i
® r\ /\ ^ ^* A
-O— Method 1602PFU
-*-qPCR-CE
>^ ^ <#
/ / /
^ ^ u
-------
1 - Mill Creek WTP
o
o
c *-
8|
§1
O o
LU
O
108!
107
1Q6
10s
104-
103-
Preliminary
Spring
1 f/
Summer
1Q2'
io1-
4 no
lU -f
^
-0— Method 1603 CPU
-•— qPCR-CE
i i i i i i i i i i i i i i i i i i i
$ & & & cf^1^^ ^^^^ <$ <^ & &
-
-------
2 - Little Miami WTP
107
106
8 10*
C
0
-------
3 - Muddy Creek WTP
107
106
I
8 10s
J 3
|0
(U O
o •=
a
10°
(N
Method 1603 CPU
qPCR-CE
* £
Preliminary
Spring
Summer
Winter
Figure A17. Comparison of the Effect of the Wastewater Treatment Processes at Muddy Creek WTP
on the E.co// Concentrations (o), Determined Using EPA Method 1603, and the Quantitative
Polymerase Chain Reaction (qPCR) Method Cell Equivalents (•)
-------
4 - Polk Run WTP
108
107
I 10s
O
LLJ
O
10=
•p« —
C fe
8 | 10*
gj
O o
^ 103
102
10°
6
<^ & £* *i*
?P -^ fP .of1 .jT .^ /P -w1
b (F & -^
-------
o
o
.ii
Ife
c «-
h
Q o
LJJ
O
108 -
107-
106
10s
104
103
102
10°
1 - Mill Creek WTP
-O—a fragHisCFU
-•— qPCrf-CE
Preliminary
Spring Summer
Number of Days the Sample was
Held at Seasonal Temperatures
Winter
Figure A19. Comparison of the Effect of the Wastewater Treatment Processes at Mill Creek WTP
on the Bacteroides fragilis Concentrations (o) and the Quantitative Polymerase Chain Reaction
(qPCR) Method Cell Equivalents (CE) (•)
-------
o
o
O I3
0) O
si
O o
LU
107-
106-
10s"
1Q4.
103-
1Q2.
2 - Little Miami WTP
—O— fl- frag/7/sCFU
-^-qPCR-CE
05
Spring
&?
Summer
/
«? <#
Winter
Number of Days the Sample was
Held at Seasonal Temperatures
Figure A20. Comparison of the Effect of the Wastewater Treatment Processes at Little Miami
WTP on the Bacteroides fragilis Concentrations (O) and the Quantitative Polymerase Chain
Reaction (qPCR) Method Cell Equivalents (CE) (•)
-------
3 - Muddy Creek WTP
o
108
107.
o 106
o
|g 105
So
C £ 1Q4
O —
58 103
10'J
10°
B. fragilis CPU
qPCR-CE
VO
Preliminary
/ /
/ /<
J *
Spring
<
«
Summer
Winter
Figure A21, Comparison of the Effect of the Wastewater Treatment Processes at Muddy Creek
WTP on the Bacteroides fragilis Concentrations (O) and the Quantitative Polymerase Chain
Reaction (qPCR) Method Cell Equivalents (CE) (•}
-------
4 - Polk Run WTP
o
o
108
107-
106 -
105-
o B
So
g £ 104
8 I
O 8 1°3
UJ 1Q2
10°
B. frag///s CPU
•qPCR-CE
Spring
& ^ &
a& % -J&
^ ^
-------
108,
107-
0 10N
8
!& 1°5
I | 10*
§1
O o
LU
102-
10°
1 - Mill Creek WTP
oo
—O— Cl, perfringerts CPU
-•—qPCR-CE
Preliminary
Spring
Summer
Winter
Figure A23. Comparison of the Effect of the Wastewater Treatment Processes at Mill Creek WTP
on the Clostridium perfringens Concentrations (o) and the Quantitative Polymerase Chain
Reaction (qPCR) Method Cell Equivalents (CE) (»)
-------
2 - Little Miami WTP
10
7 -,
o
o
105-
•g w 104
fc o
o —
o S 103
O o
o
10°
-o—a
CL perfringens CPU
qPCR-CE
Spring
f
^
-------
107~
108-
o
2 105
iV
Winter
Figure A25. Comparison of the Effect of the Wastewater Treatment Processes at Muddy Creek
WTP on the Clostridium perfringens Concentrations (o) and the Quantitative Polymerase Chain
Reaction (qPCR) Method Cell Equivalents (CE) (•}
-------
4 - Polk Run WTP
rf—H
I
O
0
ji
"c fc-
-------
1 - Mill Creek WTP
Preliminary
Spring
Summer
Winter
0.5 -
0
Q>
c ® -0-5 -
g5 $ -1.5-
I I -" H
0) 05
gl -3H
O £
0 -3.5 H
_4 -
-4.5 -
Method 1600 CPU
qPCR-CE
/
(N
Figure A27. Comparison of the Effect of the Wastewater Treatment Processes at Mill Creek
WTP on the Cumulative Log10 Reduction in Enterococcus Concentrations, Determined
Using EPA Method 1600 (o), and the Quantitative Polymerase Chain Reaction (qPCR)
Method Cell Equivalents (•)
-------
2 - Little Miami WTP
Spring
Summer
Winter
—O—Method 1600 CPU
qPCR-CE
Figure A28. Comparison of the Effect of the Wastewater Treatment Processes at Little Miami
WTP on the Cumulative Log10 Reduction in Enterococcus Concentrations, Determined Using
EPA Method 1600 (o), and the Quantitative Polymerase Chain Reaction (qPCR) Method
Cell Equivalents (•}
-------
3 - Muddy Creek WTP
Preliminary
Spring
Summer
Winter
0
Method 1600 CPU
qPCR-CE
Figure A29. Comparison of the Effect of the Wastewater Treatment Processes at Muddy Creek
WTP on the Cumulative Log10 Reduction in Enterococcus Concentrations, Determined Using
EPA Method 1600 (O), and the Quantitative Polymerase Chain Reaction (qPCR) Method
Cell Equivalents (•)
-------
4 - Polk Run WTP
Spring
Summer
Winter
QJ
O)
Method 1600 CPU
qPCR-CE
-5 J
Figure A30. Comparison of the Effect of the Wastewater Treatment Processes at Polk Run
WTP on the Cumulative Log10 Reduction in Enterococcus Concentrations, Determined Using
EPA Method 1600 (o), and the Quantitative Polymerase Chain Reaction (qPCR) Method
Cell Equivalents (•)
-------
1 - Mill Creek WTP
Spring Summer
Winter
cs *
*/ /
•* £^ ff v
-------
2 - Little Miami WTP
Spring
Summer
Winter
Method 1602PFU
qPCR-CE
Figure A32. Comparison of the Effect of the Wastewater Treatment Processes at Little Miami WTP on the
Cumulative Log16 Reduction in F+ Coiiphage Concentrations, Determined Using EPA Method 1602 (o),
and the Quantitative Polymerase Chain Reaction (qPCR) Method Cell Equivalents (•)
-------
3 - Muddy Creek WTP
Preliminary
Summer
Winter
oo
Method 1602 PFU
qPCR-CE
Figure A33. Comparison of the Effect of the Wastewater Treatment Processes at Muddy Creek WTP
on the Cumulative Log10 Reduction in F+ Coliphage Concentrations, Determined Using EPA Method 1602 (o),
and the Quantitative Polymerase Chain Reaction (qPCR) Method Cell Equivalents (•)
-------
4 - Polk Run WTP
Spring
Summer
Winter
, T , r HQ
Method 1602PFU
qPCR-CE
Figure A34. Comparison of the Effect of the Wastewater Treatment Processes at Polk Run WTP
on the Cumulative Log10 Reduction in F+ Coliphage Concentrations, Determined Using EPA Method 1602 (o),
and the Quantitative Pofymerase Chain Reaction (qPCR) Method Cell Equivalents (»)
-------
1 - Mill Creek WTP
Preliminary
Spring
Summer
Winter
§>•«
I?
0.5-
0
S1-0.5J
W > '1 '
-2 -
CM ^
O >
_J CO
c Of -1.5
.2 o
•43 *-
2 ®
c •-
0 16 -2.5
c ®
o S -3
O
-3.5 J
^A #
,e» ^ -Q
** ./
-------
2 - Little Miami WTP
Spring
Summer
Winter
I I
-O—Method 1603CFU
-•—qPCR-CE
Figure A36. Comparison of the Effect of the Wastewater Treatment Processes at Little Miami
WTP on the Cumulative Logio Reduction in E. coll Concentrations, Determined Using EPA
Method 1603 (o), and the Quantitative Polymerase Chain Reaction (qPCR) Method
Cell Equivalents (•)
-------
3 - Muddy Creek WTP
01
$
c -«p
(N
—O- Method 1603 CPU
-»~qPCR-CE
Figure A37, Comparison of the Effect of the Wastewater Treatment Processes at Muddy Creek
WTP on the Cumulative Logio Reduction in £ coli Concentrations, Determined Using EPA
Method 1603 (o)» and the Quantitative Polymerase Chain Reaction
(qPCR) Method Cell Equivalents (•)
-------
4 - Polk Run WTP
Spring
Summer
Winter
Method 1603 CPU
qPCR-CE
Figure A38. Comparison of the Effect of the Wastewater Treatment Processes at Polk Run
WTP on the Cumulative Log 10 Reduction in £ coli Concentrations, Determined Using EPA
Method 1603 (o), and the Quantitative Polymerase Chain Reaction (qPCR) Method Cell
Equivalents (•)
vo
-------
1 - Mil) Creek WTP
Preliminary
Spring
Summer
Winter
—O— fi. fragilis CPU
qPCR-CE
Figure A39. Comparison of the Effect of the Wastewater Treatment Processes at Mill Creek WTP
on the Log,0 Reduction in Bacteroides fragilis Concentrations (o) and the Quantitative Polymerase Chain
Reaction (qPCR) Method Cell Equivalents (•)
-------
Spring
2 - Little Miami WTP
Summer
^
Winter
—O— B. fragilis CPU
-*— qPCR-CE
-3,5
Figure A40. Comparison of the Effect of the Wastewater Treatment Processes at Little Miami WTP on the Log10
Reduction in Bacteroides fragilis Concentrations (o) and the Quantitative Pofymerase Chain Reaction (qPCR) Method
Cell Equivalents (•)
-------
O
Preliminary
3 - Muddy Creek WTP
Spring Summer
Jj * v jSf ,p
o ».© Si *-\ o t©
-N ^K &? rs~* r+ jS.
Winter
/
/•
-O—B. fragf/feCFU
-»— qPCR-CE
-6 J
Figure A41, Comparison of the Effect of the Wastewater Treatment Processes at Muddy Creek WTP
on the Log10 Reduction in Bacteroides fragilis Concentrations (O) and the Quantitative Polymerase Chain
Reaction (qPCR) Method Cell Equivalents (•)
-------
4 - Polk Run WTP
Spring
Summer
Winter
—O— B. fragitis CPU
—»— qPCFT-CE
Figure A42. Comparison of the Effect of the Wastewater Treatment Processes at Polk Run WTP
on the Log10 Reduction in Bacteroides fragilis Concentrations (O) and the Quantitative Polymerase Chain
Reaction (qPCR) Method Cell Equivalents (•}
-------
1 - Mill Creek WTP
Preliminary
Spring
Summer
Winter
5?
Cl, perfringens CPU
qPCR-CE
oo
VO
Figure A43. Comparison of the Effect of the Wastewater Treatment Processes at Mill Creek
WTP on the Cumulative Log10 Reduction in Clostridium perfringens Concentrations (o) and the
Quantitative Polymerase Chain Reaction (qPCR) Method Cell Equivalents (CE) (•)
-------
2 - Little Miami WTP
Spring
Summer
Winter
CL perfringens CPU
qPCR-CE
Figure A44. Comparison of the Effect of the Wastewater Treatment Processes at Little Miami
WTP on the Cumulative Log10 Reduction in Clostridium perfringens Concentrations (O) and the
Quantitative Polymerase Chain Reaction (qPCR) Method Cell Equivalents (CE) (•}
-------
3 - Muddy Creek WTP
Preliminary
Spring
Summer
Winter
» ^ g
•Jtf O
/^ :£>
g? ff
& s& c
C/, perfringens CPU
qPCR-CE
-3,5 J
Figure A45. Comparison of the Effect of the Wastewater Treatment Processes at Muddy Creek
WTP on the Cumulative Logto Reduction in Clostridium perfringens Concentrations (O) and the
Quantitative Polymerase Chain Reaction (qPCR) Method Cell Equivalents (CE) (•)
-------
4 - Polk Run WTP
Spring
Summer
Winter
i=£' 0)
I 81
CO
O
-.85
2 g>
c --
I— 4_l
O
Cl. perfringens CPU
qPCR-CE
Figure A46. Comparison of the Effect of the Wastewater Treatment Processes at Polk Run WTP
on the Cumulative Log10 Reduction in Clostridium perfringens Concentrations (O) and the
Quantitative Polymerase Chain Reaction (qPCR) Method Cell Equivalents (CE) (•)
-------
1 - Mill Creek WTP
105-
ff
C T-
10°
(N
01246
Preliminary
01246 01246
Spring Summer
Number of Days the Sample was
Held at Seasonal Temperatures
01246
Winter
Figure A47. Comparison of Three Seasonal Die-Off Studies of Method 1600 Enterococcus
Concentrations Using Wastewater from Mill Creek WTP - EH, Disinfected, Secondary Effluent
Holding Study; 5%, 5% Wastewater in Ohio River Water Holding Study; 20%, Holding Study
with 20% Wastewater in a Sample Containing 20% Ohio River Water and 60% Partially-Treated
Drinking Water.
-------
2 - Little Miami WTP
10S
I
0
Spring
2 4
Winter
Number of Days the Sample was
Held at Seasonal Temperatures
Figure A48, Comparison of Three Seasonal Die-Off Studies of Method 1600 Enterococcus
Concentrations Using Wastewater from Little Miami WTP - EH, Disinfected, Secondary Effluent
Holding Study; 5%, 5% Wastewater in Ohio River Water Holding Study; 20%, Holding Study
with 20% Wastewater in a Sample Containing 20% Ohio River Water and 60% Partially-Treated
Drinking Water.
-------
3 - Muddy Creek WTP
I""*
O "7=
"-S £
m o
4= o
c T-
o p
c ii
o o
O —
10s -,
104~
103-
102-
101 -
10°
01246
Preliminary
0 1
i i r
246
Spring
01246
Summer
i i r
246
Winter
Number of Days the Sample was
Held at Seasonal Temperatures
Figure A49. Comparison of Three Seasonal Die-Off Studies of Method 1600 Enterococcus
Concentrations Using Wastewater from Muddy Creek WTP - EH, Disinfected, Secondary Effluent
Holding Study; 5%, 5% Wastewater in Ohio River Water Holding Study; 20%, Holding Study
with 20% Wastewater in a Sample Containing 20% Ohio River Water and 60% Partially-Treated
Drinking Water.
-------
4 - Polk Run WTP
£ 1Q3
5 o
is o
c •«-
102-
Winter
10°
1 2 4
Summer
Number of Days the Sample was
Held at Seasonal Temperatures
Figure A50. Comparison of Three Seasonal Die-Off Studies of Method 1600 Enterococcus
Concentrations Using Wastewater from Polk Run WTP - EH, Disinfected, Secondary Effluent
Holding Study; 5%, 5% Wastewater in Ohio River Water Holding Study; 20%, Holding Study
with 20% Wastewater in a Sample Containing 20% Ohio River Water and 60% Partially-Treated
Drinking Water.
-------
1 - Mill Creek WTP
10"
o •=
'•g E
2 o
.to o
0)
103
102
10
1 .
10°
VO
01246
Preliminary
1246
Spring
0124
Summer
1246
Winter
Number of Days the Sample was
Held at Seasonal Temperatures
Figure A51. Comparison of Three Seasonal Die-Off Studies of Method 1602 F+ Coliphage
Concentrations Using Wastewater from Mill Creek WTP- EH, Disinfected, Secondary Effluent
Holding Study; 5%, 5% Wastewater in Ohio River Water Holding Study; 20%, Holding Study
with 20% Wastewater in a Sample Containing 20% Ohio River Water and 60% Partially-Treated
Drinking Water.
-------
2 - Little Miami WTP
f—
,0 •=
CO o
J= O
® 5
o ID
c u_
3^
10°
Number of Days the Sample was
Held at Seasonal Temperatures
Figure A52, Comparison of Three Seasonal Die-Off Studies of Method 1602 F+ Coliphage
Concentrations Using Wastewater from Little Miami WTP - EH, Disinfected, Secondary Effluent
Holding Study; 5%, 5% Wastewater in Ohio River Water Holding Study; 20%, Holding Study
with 20% Wastewater in a Sample Containing 20% Ohio River Water and 60% Partially-Treated
Drinking Water.
-------
•Il
|8
CD S
M2
O Q_
Q —
3 - Muddy Creek WTP
103i
102
10
1 -
10°
oo
01246
Preliminary
01246
Spring
0124
Summer
1 2 4
Winter
Number of Days the Sample was
Held at Seasonal Temperatures
Figure A53. Comparison of Three Seasonal Die-Off Studies of Method 1602 F+ Coliphage
Concentrations Using Wastewater from Muddy Creek WTP - EH, Disinfected, Secondary Effluent
Holding Study; 5%, 5% Wastewater in Ohio River Water Holding Study; 20%, Holding Study
with 20% Wastewater in a Sample Containing 20% Ohio River Water and 60% Partially-Treated
Drinking Water,
-------
4 - Polk Run WTP
103i
0
2 4
Winter
6
1 2 4
Summer
Number of Days the Sample was
Held at Seasonal Temperatures
Figure A54. Comparison of Three Seasonal Die-Off Studies of Method 1602 F+ Coliphage
Concentrations Using Wastewater from Polk Run WTP - EH, Disinfected, Secondary Effluent
Holding Study; 5%, 5% Wastewater in Ohio River Water Holding Study; 20%, Holding Study
with 20% Wastewater in a Sample Containing 20% Ohio River Water and 60% Partially-Treated
Drinking Water.
-------
1 - Mill Creek WTP
10S
c s~-.
O "F
'•5 c
ss
103
10'
10°
O
oo
01248
Preliminary
01246
Spring
01246
Summer
1246
Winter
Number of Days the Sample was
Held at Seasonal Temperatures
Figure A55. Comparison of Three Seasonal Die-Off Studies of Method 1603 E. colt
Concentrations Using Wastewater from Mil! Creek WTP - EH, Disinfected, Secondary Effluent
Holding Study; 5%, 5% Wastewater in Ohio River Water Holding Study; 20%, Holding Study
with 20% Wastewater in a Sample Containing 20% Ohio River Water and 60% Partially-Treated
Drinking Water.
-------
10"
2 - Little Miami WTP
O
O
102
10°
oo
0124
Spring
_i—
6
01246
Summer
Number of Days the Sample was
Held at Seasonal Temperatures
01246
Winter
Figure A56. Comparison of Three Seasonal Die-Off Studies of Method 1603 E coli
Concentrations Using Wastewater from Little Miami WTP - EH, Disinfected, Secondary Effluent
Holding Study; 5%, 5% Wastewater in Ohio River Water Holding Study; 20%, Holding Study
with 20% Wastewater in a Sample Containing 20% Ohio River Water and 60% Partially-Treated
Drinking Water.
-------
3 - Muddy Creek WTP
103!
102
.2 "c
4= C
OJ O
i= O
C <-
0) 2-
o p
c u.
o o
O —
10
1'
10°
(N
oo
01246
Preliminary
01246
Spring
01246
Summer
01246
Winter
Number of Days the Sample was
Held at Seasonal Temperatures
Figure A57. Comparison of Three Seasonal Die-Off Studies of Method 1603 £ coli Concentrations
Using Wastewater from Muddy Creek WTP - EH, Disinfected, Secondary Effluent Holding Study;
5%, 5% Wastewater in Ohio River Water Holding Study; 20%, Holding Study with 20% Wastewater
in a Sample Containing 20% Ohio River Water and 60% Partially-Treated Drinking Water.
-------
4 - Polk Run WTP
1041
J f:
CO O
-b o
C T-
0 St
O P
C LL
o O
0 ^=^
103
102
10°
oo
-0-EH
^j-5%
^-20%
III!
1 I
T t
01246
Spring
0124
Summer
0
1246
Winter
Number of Days the Sample was
Held at Seasonal Temperatures
Figure A58. Comparison of Three Seasonal Die-Off Studies of Method 1603 £ coli Concentrations
Using Wastewater from Polk Run WTP - EH, Disinfected, Secondary Effluent Holding Study;
5%, 5% Wastewater in Ohio River Water Holding Study; 20%( Holding Study with 20% Wastewater
in a Sample Containing 20% Ohio River Water and 60% Partially-Treated Drinking Water.
-------
1 - Mill Creek WTP Method
5 E
'•5 °
CO o
CD
-------
2 - Little Miami WTP
ll
II
^ UJ
o O
O ^
10s-
1Q4-
102-
101 -
10°
oo
-A- 20%
i i i
0 1 2
i i i
4 6
i
0
i
1
i
2
i
4
i
6
i i
0
i
1
i
2
4 6
Spring
Summer
Number of Days the Sample was
Held at Seasonal Temperatures
Winter
Figure A60. Comparison of Three Seasonal Die-Off Studies of the Entemcoccus Quantitative
Polymerase Chain Reaction (qPCR) Method Cell Equivalents Using Wastewater from Little Miami
WTP - EH, Disinfected, Secondary Effluent Holding Study; 5%, 5% Wastewater in Ohio River
Water Holding Study; 20%, Holding Study with 20% Wastewater in a Sample Containing 20%
Ohio River Water and 60% Partially-Treated Drinking Water.
-------
3 - Muddy Creek WTP
c
O ZZ"
"S3 C
03 j=
Jb °
I?
§8
o
10s
104
103
102
101
10°
VO
oo
01246
Preliminary
01246
Spring
01246
Summer
01246
Winter
Number of Days the Sample was
Held at Seasonal Temperatures
Figure A61. Comparison of Three Seasonal Die-Off Studies of the Enterococcus Quantitative
Polymerase Chain Reaction (qPCR) Method Cell Equivalents Using Wastewater from Muddy
Creek WTP - EH, Disinfected, Secondary Effluent Holding Study; 5%, 5% Wastewater in Ohio
River Water Holding Study; 20%, Holding Study with 20% Wastewater in a Sample Containing
20% Ohio River Water and 60% Partially-Treated Drinking Water.
-------
4 - Polk Run WTP
••*=* c
to fc
f** CJ
o u3
O v
iu -
108-
107-
106-
105-
104-
103-
-o-EH
-o-5%
-A- 20%
fc^
101-l
10°
oo
0124
Spring
i
0
2 •
Summer
0124
Winter
Number of Days the Sample was
Held at Seasonal Temperatures
Figure A62. Comparison of Three Seasonal Die-Off Studies of the Enterococcus Quantitative
Polymerase Chain Reaction (qPCR) Method Cell Equivalents Using Wastewater from Polk Run
WTP - EH, Disinfected, Secondary Effluent Holding Study; 5%, 5% Wastewater in Ohio River
Water Holding Study; 20%, Holding Study with 20% Wastewater in a Sample Containing 20%
Ohio River Water and 60% Partially-Treated Drinking Water,
-------
- Mill Creek WTP
o
o
OJ
'•g Q-
OJ «e
** ?
Q} O
ll
u_
o
LU
O
\7 1
oo
oo
PLAQUE
MPN
qPCR
1600
01246
Preliminary
0124
Spring
01246
Summer
01246
Winter
Number of Days the Sample was
Held at Seasonal Temperatures
Figure A63. Seasonal Die-Off Study of the Enterococcus Quantitative Polymerase Chain Reaction
(qPCR) Method Cell Equivalents (CE) (A), the Enterococcus Culture Method CFUs (Method 1600) (- ),
and the Concentrations of Two Enterovirus Methods in a Simulated Recreational Water Containing
5% Wastewater from Mill Creek WTP in Ohio River Water - The two Enterovirus Methods were the
Plaque Assay (PFU) (o) and the CPE-MPN Method (MPN) (a).
-------
2 - Little Miami WTP
o
o
&
(D O
o
_
O
yJ
o
oo
-O—PLAQUE
—a—MPN
-A—qPCR
1600
01246
Spring
01246
Summer
Number of Days the Sample was
Held at Seasonal Temperatures
0124
Winter
Figure A64. Seasonal Die-Off Study of the Enterococcus Quantitative Polymerase Chain Reaction
(qPCR) Method Gel! Equivalents (CE) (A), the Enterococcus Culture Method CPUs (Method 1600) (-),
and the Concentrations of Two Enterovirus Methods in a Simulated Recreational Water Containing
5% Wastewater from Little Miami WTP in Ohio River Water - The two Enterovirus Methods were the
Plaque Assay (PFU) (O) and the CPE-MPN Method (MPN) (n).
-------
3 - Muddy Creek WTP
E
o
o
VITT1
i—
CD
c Qt
•"§1
43 ^
C ±-
-------
4 - Polk Run WTP
01246
Preliminary
0124
Spring
-O— PLAQUl
—o—MPN
—A—qPCR
—— ?6QO
01246
Summer
01246
Winter
Number of Days the Sample was
Held at Seasonal Temperatures
Figure A66. Seasonal Die-Off Study of the Enterococcus Quantitative Polymerase Chain Reaction
(qPCR) Method Cell Equivalents (CE) (A), the Enterococcus Culture Method CFUs (Method 1600) (.),
and the Concentrations of Two Entemvirus Methods in a Simulated Recreational Water Containing
5% Wastewater from Polk Run WTP in Ohio River Water - The two Entemvirus Methods were the
Plaque Assay (PFU) (O) and the CPE-MPN Method (MPN) (a).
------- |