EPA 600/R-09/089 | August 2011 | www.epa.gov/ord
United States
Environmental Protection
Agency
Development and Testing of
Methods to Decontaminate
a Building's Plumbing
System Impacted by a Water
Contamination Event:
DECONTAMINATION OF BACILLUS SPORES
Office of Research and Development
National Homeland Security Research Center
-------
EPA 600-R-09-089
August 2011
Development and Testing of
Methods to Decontaminate a
Building's Plumbing System
Impacted by a Water
Contamination Event:
Decontamination of
Bacillus Spores
United States Environmental Protection Agency
Cincinnati, Ohio 45268
National Institute of Standards and Technology
Gaithersburg, Maryland 20899
-------
Disclaimer
The U.S. Environmental Protection Agency (EPA) through its Office of Research and
Development partially funded and collaborated in the research described herein under
Interagency Agreement DW13-921677 to the Department of Commerce, National Institute of
Standards and Technology.. It has been reviewed by the Agency but does not necessarily reflect
the Agency's views. No official endorsement should be inferred. EPA does not endorse the
purchase or sale of any commercial products or services.
-------
DISCLAIMER 1
FOREWORD 4
ACRONYMS AND ABBREVIATIONS 5
ACKNOWLEDGEMENTS 6
LIST OF TABLES 7
LIST OF FIGURES 7
EXECUTIVE SUMMARY 8
1. 0 DECONTAMINATION OF BACILLUS SPORES ASSOCIATED WITH WATER
SYSTEM BIOFILMS 10
Introduction 10
Materials and Methods 11
Spore Preparations and Growth Conditions 11
Pipe Material Preparation 11
Biofilm Colonization of Pipe Materials 12
Pipe Loop Reactor Operation 12
CDC Biofilm Reactor (CBR) Operation 13
Preparation of Disinfectants 14
Spore Suspension Disinfection 14
Pipe Surface Disinfection 15
Estimation ofCt Value 15
Fluorescent In Situ Hybridization of Biofilm Organisms 15
Statistical Comparisons 16
Results 16
Solution Phase Spore Disinfection 16
Biofilm Accumulation 17
Spore Association with Biofilm-ConditionedPipe Materials 17
Decontamination of Surface Associated Spores 18
Biofilm Disinfection 20
Discussion of Disinfection of Spores in Solution and in Contact With Biofilms 22
-------
2.0 ENHANCED DECONTAMINATION OF BACILLUS SPORES IN A
SIMULATED DRINKING WATER SYSTEM 25
Introduction 25
Materials and Methods 26
Spore Preparations and Growth Conditions 26
Preparation of Disinfectants 27
Coupon Preparation and Biofilm Colonization of Pipe Materials 27
CDC Biofilm Reactor Operation 27
Statistical Comparisons 28
Decontamination Results 28
Spore Suspension Disinfection 29
Results of Spore Association with Biofilm-Conditioned Pipe Materials 30
Germinant Addition Impact on Biofilm 32
Discussion of Results 36
3.0 THE EFFECT OF HIGH FLOW ON THE ADHESION AND DISINFECTION
OF BT SPORES IN PIPE LOOP EXPERIMENTS 38
Introduction 38
Materials and Methods 38
Pipe Section Loop Reaction Protocol 38
BTSpore Inactivation 39
Results and Discussion 40
Conclusions 41
4.0 GAPS IN RESEARCH ON BIOLOGICAL THREATS TO BE INVESTIGATED
IN ADDITIONAL STUDIES 43
REFERENCES 45
APPENDIX A: PROTOCOLS FOR BIOLOGICAL THREAT
DECONTAMINATION 50
-------
Foreword
Following events of September 11, 2001, U.S. Environmental Protection Agency's (EPA's)
mission was expanded to account for critical needs related to homeland security. Presidential
Directives identified EPA as the primary federal agency responsible for the country's water
supplies and for decontamination following a chemical, biological, and/or radiological (CBR)
attack. To provide scientific and technical support to help EPA meet this expanded role, EPA's
National Homeland Security Research Center (NHSRC) was established. The NHSRC research
program is focused on conducting research and delivering products that improve the capability of
the Agency to carry out its homeland security responsibilities.
As part of this mission, NHSRC conducts research on contaminants that could be intentionally
injected into a community's water supply and water distribution system. The possibility of such
intentional contamination raised questions that had been heretofore largely unasked: what is the
fate of the contaminant in the water system? and how can the contaminant be removed? The
approach to answering these and other related questions depends greatly on where the
contaminant is located: the water treatment system, the water distribution system, or in a
building's plumbing system. For example, the remediation strategy for a large underground
water main that had been impacted by intentional contamination would most likely be different
than a remediation strategy for a similarly impacted plumbing system in a typical house.
The work summarized in this report addresses contamination of plumbing systems in houses and
other buildings. Specifically, the objective of the work was to develop and test of methods to
decontaminate a building's plumbing system impacted by a water contamination event. The
contaminants studied were Bacillus spores, which had proven challenging to decontaminate. A
follow on report will address other aspects of chemical and biological decontamination of
plumbing systems.
-------
Acronyms and Abbreviations
ANO V A
BA
BT
CBR
C/Co
CDC
CPU
cm
Ct
DAPI
DPA
DPD
EDTA
EPA
FISH
g
g
h
L
LB
jig
M
mg
mL
mm
mM
m/s
min
MQ
NaCl
NIST
PBS
PFA
PVC
RO
rpm
SDS
TOC
Tris
v/v
analy si s of vari ance
Bacillus anthracis Sterne
Bonide Thuricide™ Bacillus Thuringiensis (BT) Concentrate (Bonide Products,
Inc., Oriskany, New York)
CDC Biofilm Reactor (BioSurface Technologies, Bozeman, Montana)
concentration divided by initial concentration
Centers for Disease Control
colony forming unit
centimeter
concentration-time (concentration multiplied by time)
4',6-diamidino-2-phenylindole
dipicolinic acid
N,N-diethyl-p-phenylenediamine
ethylenediamine tetraacetic acid
U.S. Environmental Protection Agency
fluorescent in situ hybridization
grams
acceleration due to gravity, 9.81 m/s2
hour
liter
Luria-Bertani agar
microgram
molar (moles/liter)
milligram
milliliter
millimeter
millimolar
meters/second
minute
Milli-Q®
sodium chloride
National Institute of Standards and Technology
phospate buffered saline
paraformaldehyde
polyvinyl chloride
reverse osmosis
revolutions per minute
sodium dodecyl sulfate
Total Organic Carbon
tris (hydroxymethyl)aminomethane
volume to volume
-------
Acknowledgements
This document was prepared under an interagency agreement, DW 13-921677, between the US
EPA and the Department of Commerce, National Institute of Standards and Technology (NIST).
The contributing authors from NSIT were Dr. Kenneth Cole, Dr. Jayne Morrow and Dr. Stephen
Treado, Ms. Jamie Almeida, and Dr. Lisa Fitzgerald. The US EPA project officer was Mr.
Vicente Gallardo. The EPA review of the report was conducted by Mr. Gallardo, Dr. Jeff Szabo,
and Dr. Gene JAice.
-------
List of Tables
Table 1.1: Biofilm Surface Coverage in CFU/cm2 for the Different Reactor Types 17
Table 1.2: Initial Spore Surface Coverage in CFU/cm2 for the Different Reactor Types and
Contact Conditions 18
Table 2.1. Impact of Treatment Strategy onBT Spore Surface Coverage 33
Table 2.2. Impact of Treatment Strategy on BA Spore Surface Coverage 34
Table 2.3. Impact of Treatment Strategy on Biofilm Surface Coverage 35
List of Figures
Figure 1.1 Pipe Reactor Schematic 13
Figure 1.2 Solution Phase Disinfection of Spore Suspensions and Contact Time with
Disinfectants 16
Figure 1.3 Figure 1.3 Impact of Disinfectant and Contact Time on Spores Associated with
Biofilm-conditioned Cu or PVC Pipe Material Surfaces 19
Figure 1. 4. Biofilm Susceptibility to Disinfection 21
Figure 2.1 Flow Chart of Experimental Approach 29
Figure 2.2 Impact of Shear on BT Spore Association with Biofilm-Conditioned
Pipe Surfaces 30
Figure 2.3. Impact of Germination on Decontamination of Adhered BA Sterne Spores 31
Figure 3.1. The Effect of Water Flushing and Chlorine Disinfection Time
on the Reduction of BT Spores Associated with the Biofilm Conditioned Pipe
Sections 42
-------
Executive Summary
This report describes the work on decontamination of bacillus spores in building water systems
done at the National Institute of Standards and Technology (NIST) (Gaithersburg, Maryland).
Specifically Bacillus anthracis Sterne (BA) spores and a simulant, Bacillus thuringiensis (BT)
were the subject of this group of studies. Bacillus spores have proven to be very challenging
organisms to inactivate in a water environment. Compared to vegetative bacteria, the bacterial
endospores BA and BT are more resistant to disinfectants such as chlorine. In addition, spores
can adhere to water system biofilms which may provide protection from disinfectants. The work
with bacillus spores is summarized in this section, and divided into three separate topics:
1. Decontamination of Bacillus Spores Associated with Water System Biofilms
2. Enhanced Decontamination of Bacillus Spores in a Simulated Drinking Water System
3. The Effect of High Flow on the Adhesion and Disinfection of BT Spores in Pipe
Loop Experiments
In addition, gaps in research on biological threats were identified and studies have been
suggested for further investigation. These are given in Chapter 4 of this report.
Decontamination of Bacillus Spores Associated with Water System Biofilms
The objective of this work was to elucidate the disinfectant susceptibility and fate of BA and a
commercial preparation of BT spores associated with (i.e., attached to) a water system biofilm.
A native water system biofilm was accumulated on copper and polyvinyl chloride (PVC) pipe
material surfaces in a low-flow pipe loop and in a uniformly mixed tank reactor.
Spores were exposed to two commonly used disinfectants (free chlorine and monochloramine) in
planktonic phase (i.e., spores unattached to the biofilm) and after association with biofilm-
conditioned pipe materials. Biofilm associated spores required 5 to 10 fold higher disinfectant
concentrations to observe the same 2- to 4-logio reduction of viable spores as observed in the
planktonic phase. In a synthetic tap water, Ct values (where C is the concentration of
disinfectant in mg/L and t is exposure time in minutes) for a 2-logio reduction of BA and a
commercial preparation of BT (planktonic phase) were 290 mg-min/L and 620 mg-min/L,
respectively, for chlorine and 1200 mg-min/L and 880 mg-min/L, respectively, for
monochloramine. Shear during spore contact with biofilm-conditioned surfaces had a profound
effect on spore attachment. Application of a distributed, low-shear (60 revolutions per minute
(rpm), CDC biofilm reactor (CBR)) during spore contact with the biofilm-conditioned surfaces
resulted in a 1.0 and 1.6 logic increase in the number of BT spores associating with copper and
PVC surfaces, respectively, compared to uniform mixing alone (i.e., no applied shear). High
disinfectant concentrations (103 mg/L free chlorine and 49 mg/L monochloramine) and contact
time (60 minutes) yielded less than a 2-logio reduction of viable BT and BA spores associated
with the biofilm under low shear. Both spore preparations showed a similar susceptibility to the
disinfectants when associated with the biofilm-conditioned surfaces.
Enhanced Decontamination of Bacillus Spores in a Simulated Drinking Water System
Contact with germinant solutions, followed by commonly used disinfectants was investigated as
a way to enhance decontamination of BA and BT. PVC and copper pipe materials were used in
a continuously stirred tank reactor that permitted for controlled shear. Simulated water system
biofilms were accumulated on pipe material surfaces with synthetic tap water containing humic
-------
acids as a carbon source. Once the biofilms were established, BT subspecies kurstaki or BA
spores were added to the water system. Pipe surfaces were examined for biofilm accumulation,
spore adhesion, and disinfectant (chlorine and monochloramine) susceptibility before and after
germination with 1 mM inosine and 8 mM L-alanine. Biofilm associated spores required 5 to 10
fold higher disinfectant concentrations to observe the same 1 logio to 4 logio reduction of viable
spores as observed in the planktonic phase. Applied shear increased spore attachment (0.4 logio
to 1.6 logio increase) to biofilm-conditioned pipe surfaces. High disinfectant concentrations (103
mg/L free chlorine and 49 mg/L monochloramine) yielded less than a 2 logio reduction in
biofilm-associated viable spores after 60 minutes. A 4 logio reduction in the associated spores
was observed when coupons were in contact with germinants (24 hours) prior to sampling.
When germinant contact was followed by heat (50 °C, 20 minutes) or disinfectant contact, a
greater than 4 logio reduction in the associated viable spores was observed. Contact with
germinants appeared to dramatically enhance the susceptibility of surface-associated spores to
elevated water temperature and disinfectants.
The Effect of High Flow on the Adhesion and Disinfection of BT Spores in Pipe Loop
Experiments
In this study, a pipe loop with intermittent high flow rate was used to measure the effects of shear flow
rate on the biofilm growth, the adhesion of BT spores, and the disinfection process. It was seen that the
high flow rate had a large effect on the disinfection process by chlorine on BT spores. For high flow
conditions, moderate concentrations of chlorine (approximately 10 mg/L) effectively disinfected the
spores associated with the biofilm-conditioned pipe loops. This is in contrast to the results we obtained
with the static experiments in the laboratory in which much higher levels of chlorine (100 mg/L) were
required to achieve significant levels of disinfection.
-------
1. 0 Decontamination of Bacillus Spores Associated with Water
System Biofilms
Introduction
The fate and persistence of potential
bioterrorist agents such as Bacillus spores in
water distribution systems is a current
concern for homeland security. Several
recent studies have defined Ct (disinfectant
concentration multiplied by exposure time)
values for bioterrorist agents, including
Bacillus spores, by studying disinfectant
efficacy in simple buffer solutions (Rice, et
al. 2005; Rose, et al. 2005; Rose, et al.
2007). Bacillus spores were found to be
more resistant to commonly used
disinfectants (monochloramine and free
chlorine) than vegetative bacteria (Rice, et
al. 2005; Rose, et al. 2005; Rose, et al.
2007). However, little work has focused on
disinfectant efficacy for Bacillus spores in
water systems (Gibbs, et al. 2004; Szabo, et
al. 2007).
Contaminant organisms delivered to water
distributions systems under varied shear
conditions encounter complex solution
chemistries (Rose, et al. 2005; Dow, et al.
2006), as well as different native biofilms on
numerous pipe material surfaces (Ridgway
and Olson, 1981; LeChevallier, et al. 1993;
Manz, et al. 1993; Schwartz, et al. 2003).
Water quality parameters such as pH,
dissolved organic carbon and temperature
result in different Ct values for contaminant
inactivation (Dow, et al. 2006). Association
with biofilm matrices further increases
bacterial resistance to disinfection
(LeChevallier, et al. 1988; Cochran, et al.
2000; Donlan, et al. 2002; Gibbs, et al.
2004; Szabo, et al. 2006; Szabo, et al. 2007).
Substratum material and integrity can
impact biofilm formation and contaminant
survival in water systems (Domek, et al.
1984; LeChevallier, et al. 1987; Gagnon, et
al. 2004; Szabo, et al. 2006; Szabo, et al.
2007). Microorganisms have been found
associated with corrosion-induced pits and
tubercles where there are localized
differences in chemistry and hydrodynamic
conditions (LeChevallier, et al. 1988; De
Beer, etal. 1994). Hydrodynamic
conditions vary greatly in complex water
distribution systems, yet few studies have
addressed the role of fluid shear on bacterial
biofilm populations and pathogen
association with substrata (Rickard, et al.
2004; Azevedo, et al. 2006).
Disinfectant efficacy on biofilms and
contaminant organisms harbored by biofilms
is largely dependent on the chemical
reactivity and the ability of the disinfectant
to penetrate the biofilm matrix. Highly
reactive oxidants such as free chlorine, as
well as the less reactive chloramines, have
been shown to be consumed by the top
layers of the biofilm matrix, resulting in a
retarded penetration of the biofilm (De Beer,
et al. 1994; Huang, et al. 1995; Chen and
Stewart, 1996; Stewart, et al. 2001). Due to
its slower reaction kinetics, monochloramine
has been shown to be more effective at
penetrating polysaccharide layers that make
up the biofilm (LeChevallier, et al. 1990;
Samrakandi, et al. 1997; Turetgen, 2004).
Additionally, strong oxidants are more
corrosive on metal pipe materials and are
more susceptible to reaction with corrosion
byproducts, limiting their disinfectant ability
when compared to less corrosive biocides
(e.g., monochloramine) (LeChevallier, et al.
1993). In another study, B. atrophaeus
spores associated with biofilms on corroded
iron pipe surfaces were resistant to high
levels of chlorine (10, 25 and 70 mg/L) for
extended periods of time (Szabo, et al.
2007).
10
-------
The objective of this work was to elucidate
the association and disinfection sensitivity
of B. anthracis Sterne (BA) and a
commercial preparation of B. thuringiensis
(BT) spores when contacted with biofilm-
conditioned pipe material surfaces in a
simulated treated water system. We
measured the ability of commonly used
disinfectants (free chlorine and
monochloramine) to inactivate B A and BT
spores in synthetic tap water and when
associated with biofilm-conditioned surfaces
(copper and polyvinyl chloride (PVC)).
Two different reactor systems - a low-flow
pipe loop and a uniformly mixed tank
reactor, the CDC biofilm reactor (CBR)
(BioSurface Technologies Corp., Bozeman,
Montana) - were utilized to grow biofilms
similar to those found in water distribution
systems. The impact of fluid shear on
spores associated with biofilm-conditioned
surfaces and on subsequent decontamination
was evaluated for two contact conditions:
the uniformly mixed pipe loop reactor and
the low shear, uniformly mixed tank reactor.
Materials and Methods
Spore Preparations and Growth Conditions
Pure suspensions of BA were compared to a
commercially available BT preparation
(Bonide Thuricide™ Bacillus Thuringiensis
(BT) Concentrate, Bonide Products, Inc.,
Oriskany, New York), for the ability to
associate with water system biofilms and to
determine disinfectant susceptibility. BT
was chosen as a simulant for BA due to the
similar exosporium composition (Matz, et
al. 1970). The commercial BT preparation
was washed to yield a concentrated spore
preparation (~lx!09 (colony forming units)
CFU/mL) by the following procedure:
aliquots of the BT spore suspension were
centrifuged at 16,000 x g for 6 minutes,
supernatant was removed and discarded and
the spores were re-suspended in phosphate
buffered saline (PBS) containing 0.01 %
(v/v) Triton™ X-100. PBS (0.01 % Triton™
X-100 ) was prepared by dissolving 8 g of
NaCl, 0.2 g of potassium phosphate, 1.15 g
of sodium phosphate and 0.2 g of potassium
chloride in 1 L of MQ water (Milli-Q® ,
Millipore Corp., Billerica, Massachusetts),
resistivity 18 MQ), pH 7.4, and autoclaved
for 15 minutes, 121 °C. To minimize spore
aggregation, 1 mL of 10 % (v/v) Triton™ X-
100 was added to the PBS after it had cooled
to room temperature. The spore pellet was
washed 4 times in PBS (0.01 % (v/v)
Triton™ X-100) by centrifugation at 16 000
x g for 6 minutes, pipetting and vortexing to
re-suspend the sample, then rinsed one
additional time and stored in 20% (v/v)
ethanol at 4 °C. BA spore suspensions from
the U.S. Army's Dugway Proving Ground,
Dugway Proving Ground, Utah,
characterized and noted as "Lot 3" in a
previous publication (Almeida, et al. 2008)
(3xl08 CFU/ml) were stored in sterile Milli-
Q® (MQ) water at 4 °C. Spore
concentrations were determined by the
spread plate method on Luria-Bertani (LB)
agar and incubating at 35 °C (see Appendix
A, Protocol 8). PBS (0.01 % Triton™ X-
100) was used as a dilution buffer for
biofilm and spore enumeration to enhance
dispersal of biofilm bacteria and biofilm
associated spores. Spore preparations
contained > 95 % phase bright spores as
determined by phase microscopy.
Pipe Material Preparation
PVC and copper pipe surfaces were rinsed
with reverse osmosis (RO) water prior to
connecting to the reactor tubing. (PVC and
copper coupons were purchased from
BioSurface Technologies Corp., Bozeman,
Montana)). PVC coupons were rinsed
briefly in 10 % (v/v) bleach solution
followed by RO water prior to inserting into
the reactor holder. Copper coupons were
briefly buffed in a circular motion with a 3M
Scotch-Brite™ pad and then soaked in 5 %
11
-------
(v/v) nitric acid for 30 minutes and rinsed
with RO water prior to inserting into the
reactor holder. For the pipe loop reactor, all
pipe surfaces had been conditioned by
several years of laboratory use.
Biofilm Colonization of Pipe Materials
Simulated water system biofilms were accumulated by contacting pipe surfaces in synthetic tap
water with the following formulation: 1.2 mM NaHCO3, 0.54 mM MgSO4-7H2O, 0.2 mM
CaSO4-2H2O, 0.004 mM K2HPO4, 0.002 mM KH2PO4, 0.08 mM (NH4)2SO4, 0.17 mM NaCl,
36xlO'6 mM FeSO4-7H2O, 0.011 mM NaNO3, 0.2 mM CaCO3, pH = 8.2 ± 0.2
(LeChevallier, 2005, personal
communication providing aforementioned
synthetic tap water formulation.) made from
the laboratory RO water supply. Growth of
organisms indigenous to the RO water
supply system at the National Institute of
Standards and Technology (NIST) was
stimulated by adding 24 mg/L humic acid
sodium salt to the synthetic tap water for
two weeks. A final carbon deprivation
period of 3 to 5 days, achieved by omitting
humic acids, was utilized to produce viable
plate counts of biofilm organisms consistent
with literature reports for water distribution
systems (Schwartz, et al. 2003). Biofilm
organisms were enumerated by spread
plating on R2A agar and incubating at room
temperature for up to 7 days (Schwartz, et
al. 2003). Biofilms were grown using two
reactor system types: the pipe loop reactor
and the CBR.
Pipe Loop Reactor Operation
The pipe loop reactor system consisted of a
maximum of eighteen, two inch long (51
mm), % inch nominal diameter (19 mm),
alternating PVC schedule 80 pipe and
copper pipe sections connected by silicone
tubing (50 mm long) (Figure 1.1). Synthetic
tap water was delivered from a 20 L carboy
with Norprene® Food Process Tubing (6402-
14) tubing at 1 mL/min (a fluid velocity of
5.8 xlO"6 m/s and hydraulic retention time of
15 to 30 hours (depending on reactor length)
with a peristaltic pump. Midway through
the biofilm accumulation period (7 days),
the pipe sections were rotated from influent
to effluent to obtain uniform biofilm growth
along the length of the reactor. After 14
days, the reactor was switched to synthetic
tap water without humic acids, and flow was
continued for an additional 3 to 5 days. Pipe
sections were removed from the reactor and
placed in sterile synthetic tap water. Biofilm
accumulation was quantified by scraping the
inside surface of the pipe with a sterile cell
scraper (23 cm, Nunc™ ). The inside of the
pipe section was then rinsed several times
with 10 mL of dilution buffer (0.0425 g/L
KH2PO4, 0.405 g/L MgCl2 6H2O) solution
into a sterile, 50 mL conical tube. The tube
was vigorously vortexed three times at 10
second intervals.
Pipe sections were contacted with bacterial
spores by placing individual sections
vertically on a perforated, plastic rack in a 4
L beaker with 1.5 L of sterile synthetic tap
water with 5 x 106 CFU/mL spores. To
reduce settling of the spores during batch
contact, the spore solution was gently stirred
with a DuPont™ Teflon® coated stir bar
under the perforated rack. After a given
contact time, pipe sections were removed
from the spore solution with sterile forceps
and gently rinsed by placing in sterile
synthetic tap water for several minutes using
gentle agitation to wash away the unbound
spores. Rinsed pipe sections were then
either directly sampled by scraping to
12
-------
determine initial spore association or placed
in a sterile 250 mL beaker containing 150
mL of the disinfectant solution. Initial
association experiments (performed for two
to 48 hours spore contact times) indicated
that steady state spore association was
reached after 24 hours in batch contact
conditions (data not shown). All
inactivation studies were performed with 24
hours spore contact time.
0.2 urn filter vent
Figure 1.1. Pipe Reactor Schematic. Synthetic water was pumped at 1 mL/min
from a 20 L carboy with a peristaltic pump (P) through alternating PVC and
copper pipe sections connected by silicon tubing.
CDC Biofilm Reactor (CBR) Operation
The impact of distributed shear on spore
association with the biofilm surface was
determined with the CBR. The CDC
biofilm reactor was chosen due to its history
as a model potable water system (Donlan, et
al. 2002) and for the ability to control the
applied fluid shear flow during biofilm
development and spore contact conditions
(Goeres, et al. 2005). The reactor was
assembled and vented as directed by the
manufacturer.
Synthetic tap water was supplied from a 20
L feed carboy to the CBR by a peristaltic
pump using Masterflex® tubing (Norprene®
Food Tubing 6402-14). Prepared coupons
were inserted into the coupon holders and
the reactor was filled with 400 mL of
synthetic tap water and autoclaved (121 °C
for 15 minutes). Reactors were allowed to
cool and then placed on digital stir plates
and connected to the peristaltic pump. A
flow rate of 0.3 mL/min (hydraulic retention
time of 29 hours) supplied the reactors with
synthetic tap water containing 24 mg/L
humic acids for 14 days at room
temperature. After 7 days, mixing was
introduced by turning on the digital stir plate
to begin baffle rotation at 120 ± 5 rpm
(confirmed by a tachometer) while fluid
flow was continued. After 2 weeks of
synthetic tap water with humic acids,
reactors were switched to synthetic tap water
without humic acids, applied at the same
flow rate for 3 to 5 days.
Contact with spores was performed by
filling the reactor beaker to the 700 mL
mark with synthetic tap water and adding
the appropriate amount of a concentrated
spore stock suspension resulting in a
concentration of 1.4 (0.8) x 107 CFU/mL of
spores. Spores were contacted for 24 hours
under uniform mixing in batch (baffle was
removed using only the stir bar rotating at
13
-------
60 rpm) and with an applied shear (baffled
stir bar with 60 rpm rotation) in the reactor.
Steady state spore association with the
biofilm-conditioned surfaces was found to
occur after 24 hours when tested for
adhesion for 2, 12, 24, 36 and 48 hours (data
not shown). The coupon holders were
removed, rinsed in sterile synthetic tap water
and sampled or exposed to disinfectants.
The CBR was sampled by removing coupon
holders and placing them on a sterile piece
of aluminum foil. Coupon surfaces were
scraped in a circular fashion, collecting the
biofilm in the center of the coupon while
still positioned in the coupon holder (Zelver,
et al. 1999; Zelver, et al. 2001). The method
by Zelver, et al. 2001 was adapted by using
a sterile piece of DuPont™ Teflon® cut to
resemble a spatula. The method had
originally called for the use of a cotton
swab, but the aforementioned spatula was
seen as superior. Coupons were removed
from the holder with a pair of sterile forceps,
positioned over a 15 mL BD Falcon™
polypropylene centrifuge tube and rinsed by
dispensing 5 mL of sterile synthetic tap
water directly on the coupon surface. The
rinsing procedure was repeated 2 times
using the same 5 mL of synthetic tap water
to minimize sample volume. Biofilm
samples were vortexed for 30 seconds at the
highest speed to disperse the cells and
associated spores. Biofilm and biofilms
post-spore contact were diluted and
enumerated on R2A and LB agar,
respectively. R2A plates were enumerated
after 7 days at room temperature. Spore
samples were counted after storing the LB
plates at 35 °C for 16 hours. Since it was
relatively straightforward to distinguish
between BA or BT spores and the native
bacteria in the biofilm, it was not necessary
to heat shock the spore samples prior to
spread plating onto the LB agar. The
biofilm bacteria grew slowly (i.e., about 1
week before colonies were visible), whereas
the spores germinated and grew rapidly
overnight on the plates and could be easily
recognized by the morphologies of the
colonies.
Preparation of Disinfectants
Disinfectant solutions consisted of 150 mL
of synthetic tap water containing either free
chlorine at average concentrations of 11
mg/L (ranged from 10 mg/L to 12 mg/L)
and 103 mg/L (ranged from 98 mg/L to 108
mg/L) or monochloramine at average
concentrations of 13 mg/L and 49 mg/L
(ranged from 8 mg/L to 18 mg/L and 47
mg/L to 57 mg/L, respectively). Free
chlorine was derived from a stock solution
of sodium hypochlorite (Clorox®, bleach).
Free chlorine concentrations were
determined using N,N-diethyl-p-
phenylenediamine (DPD) reagent and
chlorine standards (Hach Method 8021,
Hach Company, Loveland, Colorado).
Monochloramine solutions were prepared as
described by Camper et. al. (Camper, et al.
2003). Monochloramine concentrations
were determined by DPD and the
indophenol method (Hach Methods 8167,
8021, and 10171, Hach Company, Loveland,
Colorado). Concentrations of all
disinfectant solutions were determined after
30 minutes of stirring the post-disinfectant
addition to synthetic tap water to eliminate
chlorine demand contributions to effective
disinfectant determinations.
Spore Suspension Disinfection
BA and BT spores were diluted to
approximately 1 x 106CFU/mL in sterile
glass vials that contained either a
disinfectant in sterile synthetic tap water or
sterile synthetic tap water as a control.
Gentle stirring with small stir bars was used
to prevent settling of spore suspensions
during contact. Viability after contact was
determined at various time points by
14
-------
sampling 100 jil from the vial and diluting
10-fold in 0.3 mM sodium thiosulfate in
PBS (0.01% (v/v) Triton™ X-100).
Pipe Surface Disinfection
Pipe surfaces (coupons in coupon holders or
pipe sections) were contacted with
disinfectant solutions (150 mL) in 250 mL
beakers of the disinfectant solution. A small
stir bar was used to uniformly mix the
disinfectant solution during batch contact of
the CBR coupons. Beakers with pipe
sections were gently swirled every 10
minutes of contact time to ensure adequate
mixing. After the given contact time, pipe
surfaces were immediately removed and
placed in 150 mL of sterile synthetic tap
water containing 7.5 mM sodium thiosulfate
to neutralize residual disinfectant prior to
sampling.
Estimation ofCt Value
Ct values were estimated using the Chick-
Watson Law assuming first-order kinetics
(Weavers and Wickramanayake, 2001).
Logic (C/Co) for a given concentration
multiplied by the contact time (Ct,
mg-min/L) of spore inactivation was plotted
and linear regression analysis was
performed to determine the Ct value for a 2-
logio reduction in the viable spore fraction
(Le Dantec, et al. 2002).
Fluorescent In Situ Hybridization ofBiofilm
Organisms
Probes and hybridization conditions for
fluorescent in situ hybridization (FISH)
were used as described by Manz, et. al.
(1993) for drinking water biofilm organism
identification. Oligonucleotide probes for
the P- and y-proteobacteria labeled with
fluorescent dyes were purchased from
Invitrogen Corp. (Carlsbad, California).
Alexa Fluor 546 and Alexa Fluor 647 were
added to the 5' end of the |3-42a and y-42a
probes, respectively, as defined by Manz, et
al. (1993). The hybridization protocol
described by Manz, et. al. (1993) was
modified to probe biofilm samples removed
from the pipe material surfaces as described
in the following sampling procedure: 1 mL
of the 5 mL sample collected was
centrifuged and re-suspended in 1 part PBS
buffer and 3 parts 4% paraformaldehyde
(PFA). The sample was vortexed and fixed
for 3 hours at 4°C. Biofilm samples were
washed twice with PBS and re-suspended in
1 part PBS and 1 part ethanol (95 %).
Twenty \\L samples were immobilized by
spotting on poly-L-lysine slides and allowed
to air-dry. The samples were dehydrated by
dipping the slides in an ethanol series (50 %,
80 %, 95 %; 3 minutes each) and the slides
were air dried in a vertical position.
Twenty-five \\L hybridization buffer (20
mM Tris-HCl pH 7.2, 0.9 M NaCl, 39%
formamide, and 0.01 % sodium dodecyl
sulfate (SDS)) containing 80 ng of probe
was added to the sample surface. Two
milliliters of hybridization buffer was
poured into a 50 mL centrifuge tube that
contained a Kimtech Science Kimwipes®
(Kimberly-Clark Corp., Dallas, Texas)
wiper. The slide was sealed in the 50 mL
tube and incubated in a horizontal position
at 50 °C for 16 hours. Slides were rinsed
with preheated (50 °C) washing buffer (20
mM Tris-HCl pH 7.2, 40 mM NaCl, 5 mM
ethylenediamine tetraacectic acid (EDTA),
and 0.01 % SDS) and incubated at 50 °C for
15 minutes. The slides were rinsed with
water, air-dried and imaged with an
epifluorescence microscope (Olympus®
AX70) with the suited filter sets. Image
analysis was performed with Image J
software, a public domain, open source
software (Rasband, 1997-2007)
15
-------
Statistical Comparisons
Dunnett's multiple comparison procedure
was used for treatments versus controls. A
one-way analysis of variance (ANOVA) was
performed to test the hypothesis that the
average mean value across categories of the
groups were equal. In the presence of
significance for the omnibus ANOVA test, a
Newman-Keuls multiple comparison test is
used to perform pairwise comparisons.
Statistical decisions were made at a =0.05.
Statistics were performed using WINKS
SDA Software (TexaSoft, Cedar Hill,
Texas.)
Results
Solution Phase Spore Disinfection
Significant reductions in the viable spore
count were observed for both spore
suspensions after 60 minutes of contact with
both free chlorine and monochloramine
(Figure 1.2). However, free chlorine was
more effective than monochloramine at
reducing the viable spore fraction for both
preparations. BA spores were more
susceptible to free chlorine than the
commercial preparation of BT were. A four
logic reduction in the viable fraction of BA
spores was noted after 60 minutes of contact
with approximately 11 mg/L of free
chlorine. A 2.5 logio reduction in viable
spores was observed after 60 minutes of
contact with the same concentration of free
chlorine for the commercial preparation of
BT spores. Ct values for a 2-logio reduction
in the viable spore fraction when contacted
-6
20 40 60 80 100
Contact Time (min) with Disinfectants
120
140
160
Figure 1.2. Solution Phase Disinfection of Spore Suspensions and Contact Time with
Disinfectants. BA spores, A, and the commercial preparation of BT, •, were contacted with
free chlorine (average concentration of 11 mg/L) shown as solid symbols, and monochloramine
(average concentration of 13 mg/L) shown as open symbols, in sterile synthetic water, pH =
8.1, at 23°C for the given contact time in minutes, error bars indicate the observed range of
duplicate experiments, linear regression analysis R2 values are shown next to the trend lines.
C/Co indicates final concentration divided by the initial concentration of spores in suspension.
16
-------
with free chlorine were 290 and 620
mg-min/L for BA and BT spores,
respectively. Monochloramine was less
effective at killing free spores in solution
yielding only a 1.0 to 1.4 logio reduction
after 60 minutes of contact with the B A and
the commercial preparation of BT,
respectively. Ct values for a 2-logio
reduction in the viable spore fraction due to
monochloramine contact were 1200 and 880
mg-min/L for BA and BT spores,
respectively.
Biofilm Accumulation
Heterotrophic plate counts on R2A media
were utilized to quantify the biofilm
community associated with the pipe material
surface. Typically, six colony morphologies
were apparent on R2A plates. Average
heterotrophic plate counts of the biofilm
bacteria were not significantly different for
the CDC reactor compared to the pipe
reactor (a = 0.05, p = 0.105 and 0.069, for
PVC and copper, respectively) (Table 1.1).
In all cases, our heterotrophic plate counts
were quantitatively consistent at 1.1-2.5 x
105 CFU/cm2 (when converted to similar
units) with those reported in the literature
for mature water distribution systems
(ranges of 0.23 to 8.4 x 105 CFU/cm2)
(Schwartz, et al. 2003).
Table 1.1: Biofilm Surface Coverage in CFU/cm2 for the Different Reactor Types
Material Pipe Reactor CDC Reactor
Biofilm
PVC
Cu
2.5 (1.7) x 105
1.0 (0.8) x 105
1.7 (0.8) x 105
1.8 (1.4) x 105
Values presented are averages with 1 standard deviation shown in parentheses, n > 5.
Epifluorescent microscopy was utilized to
determine biofilm morphology and the
results of FISH probing. Imaging of
biofilms indicated sparsely distributed
aggregates (10 to 70 um in diameter) of
bacteria with a more uniform layer of
extracellular polysaccharides. FISH results
indicated 91 % and 94 % of the total
bacterial population as stained by 4',6-
diamidino-2-phenylindole (DAPI) were
positive for either |3 or y-proteobacteria,
respectively. All culturable colony
morphologies were positive for either the
P or y-proteobacteria.
Spore Association with Biofilm-Conditioned
Pipe Materials
The number of BT spores associating with
the biofilm-conditioned surfaces was largely
dependent on spore contacting conditions.
Significantly more spores were associated
with the pipe material surfaces when the
reactor was uniformly mixed and a low
shear was uniformly applied over the
coupon surfaces (60 rpm with the baffle
present in the CBR) during the 24 hours
contact time compared to uniform mixing
alone (pipe reactor and no baffle in the CDC
reactor, Table 1.2). Uniformly distributing
the fluid shear over the coupon surfaces in
the CDC reactor with the baffle present
resulted in more spores on average
associating with the biofilm compared to
results when the baffle was removed. The
impact of uniform shear application was
consistent for the spore preparations; both
spore preparations had similar surface
coverage of associated spores on the
different pipe material surfaces (Table 1.2).
17
-------
Table 1.2: Initial Spore Surface Coverage in CFU/cm2 for the Different Reactor Types and
Contact Conditions
Pipe Reactor
CDC Reactor
Spore Association, CFTJ/cm2
BT
Uniformly mixed
(no baffle)
PVC 1.1 (l.O)xlO3**
Cu 0.8(0.6)xl03**
Uniformly mixed
(no baffle)
1.8(0.8)xl03*
3.2(1.4) xlO3
Uniformly mixed,
Low-shear (baffle)
1.7(1.3)xl04
2.5(1.6)xl04
BA
PVC
Cu
ND
ND
ND
ND
5.6(5.4)xl04
5.7(5.3)xl04
Values presented are averages with 1 standard deviation shown in parentheses, n > 3.
* indicates values are significantly lower than the CDC reactor with the baffle in place (a =
0.05, p< 0.04),
** indicates values are significantly lower than the CDC reactor with and without the baffle
(a = 0.05, p< 0.009),
ND = not determined
Decontamination of Surface Associated
Spores
Spores associated with pipe materials
conditioned with synthetic tap water system
biofilms were more difficult to disinfect
with chlorine and monochloramine than
spores in suspension. In order to see the
same 2 to 4 logic reductions observed for
free spores in solution, biofilm associated
spores required 5 times the monochloramine
concentration and nearly 10 times the free
chlorine concentration (Figure 1.3).
Contrary to what was observed in
suspension, monochloramine was more
effective than free chlorine at reducing the
viable spore fraction when the spores were
associated with the biofilm-conditioned pipe
materials. Disinfection of the commercial
preparation of BT spores was tested for both
reactor configurations with free chlorine and
monochloramine (shown in Figure 1.3).
Depending on the pipe material surface and
contact conditions, a 1 to 4 logic reduction
in the viable spore fraction was observed
after contact with monochloramine at -49
mg/L. Spores associated with low-shear
contact conditions were more difficult to
disinfect resulting in a slightly lower
reduction in the viable spore fraction for the
CDC reactor (baffle attached) than just
uniformly mixed contact conditions (pipe
reactor). There were no significant
differences in the reduction of BA or BT
spores associated with the biofilm-
conditioned pipe surfaces following
monochloramine contact.
18
-------
Monochloramine (49 mg/L)
0 *
-1 -
o
o
O)
o
!
o
CO
-3
-4
0
A. Cu
-1 -
-2
-3
-4
B. PVC
20
40 60 0 20
Free Chlorine (103 mg/L)
40
60
-4
20 40
Contact Time (min)
60
20 40
Contact Time (min)
60
Figure 1.3. Impact of Disinfectant and Contact Time on Spores Associated with Biofilm-
conditioned Cu or PVC Pipe Material Surfaces. Inactivation curves, presented as logic of the
fraction remaining on the surface are depicted for the commercial preparation of BT, •, and BA, A,
after contact with monochloramine (49 mg/L), panels A and B, and free chlorine (103 mg/L), panels
C and D. Contact conditions for spore association were uniformly mixed, pipe reactor (open
symbols), and uniformly mixed under low-shear, CDC reactor (solid symbols). Temperature was 22
°C to 23 °C and pH 8.1. Averages of duplicate experiments are shown, bars indicate the observed
range of data.
19
-------
Free chlorine was not as effective as
monochloramine at inactivating spores from
the biofilm-conditioned surfaces. Free
chlorine contact at an average concentration
of 103 mg/L was required to yield a 1 to 4
logic reduction in viable BT spores
associated with the biofilm-conditioned pipe
surfaces. When spores were continuously
stirred with a baffle present to distribute the
shear across the coupon surface (CDC
reactor data, Figure 1.3), significant tailing
was observed for both pipe material surfaces
after 15 minutes of contact with free
chlorine at -103 mg/L. The tailing resulted
in single logic reductions after 30 and 60
minutes of contact that were the same as
reductions observed after 15 minutes of
contact. Tailing behavior was consistent for
both spore preparations resulting in
insignificant differences in measured logic
reductions for BA and BT spores (p < 0.4)
after 30 minutes of contact with free
chlorine. Tailing was not due to depletion
of disinfectant over the course of the study.
Residual disinfectant concentrations, post
biofilm contact, were not significantly
reduced during contact with biofilms (up to
120 minutes).
Biofilm Disinfection
Water system biofilms accumulated on pipe
surfaces were tested for vulnerability to two
disinfectants, free chlorine and
monochloramine, in the CDC reactor. Both
free chlorine and monochloramine at the
concentrations tested resulted in a
significant reduction in biofilm surface
coverage (p = 0.02-0.04, a = 0.05). As with
the spores associated with biofilm-
conditioned pipe materials, the biofilm
organisms themselves were more effectively
removed when contacted with
monochloramine compared to free chlorine.
A lower number of viable bacteria remained
on the copper and PVC coupons after 30
minutes of contact with monocholoramine
(-13 mg/L) than chlorine at -11 and 103
mg/L (Figure 1.4). Interestingly, a near 10
fold increase in the free chlorine
concentration did not result in a significant
decrease in the viable cell fraction of the
biofilm for either substratum (p = 0.22 and
0.27 for PVC and copper, respectively, a =
0.05).
20
-------
0
-0.5
-u
3 <—» A
00-1
O O
"S ^ -1.5
°- o)
_o> o
-2
-2.5
-3
Free Chlorine
(10mg/L)
Free Chlorine
(103mg/L)
Monochloramine
(13mg/L)
Disinfectant and Concentration
Figure 1.4. Biofilm Susceptibility to Disinfection. Heterotrophic plate count logic reductions of
biofilm organisms accumulated on pipe material surfaces in the CDC reactor, copper (solid) and
PVC (open), after contact with average concentrations with 11 mg/L and 103 mg/L free chlorine
and 13 mg/L monochloramine for 30 minutes. Temperature was 22 °C to 23 °C and pH 8.1.
Experimental averages are shown (n > 2), experimental ranges are indicated by bars.
21
-------
Discussion of Disinfection of Spores in
Solution and in Contact With Biofilms
Biofilms have been implicated as reservoirs
for their ability to harbor and protect
bacterial contaminants in water distribution
systems (LeChevallier, et al. 1987; Storey
and Ashbolt, 2003). This work sought to
examine the role of biofilm conditioning of
pipe materials on the fate and disinfection
susceptibility of spores in water systems.
Free chlorine and monochloramine are
proven disinfectants for the removal of
planktonic bacteria from the water column.
The inherent resistance of spores to
disinfection is due to the complex nature of
the spore coat and the inability of biocides to
penetrate and inactivate the spore core
(Cortezzo, et al. 2004). Typical chlorine Ct
values are 2-3 orders of magnitude higher
for spores than vegetative pathogens (Rose,
et al. 2005) and Ct values vary by an order
of magnitude among spores from different
Bacillus species (Rice, et al. 2005). The Ct
value reported here for B A inactivation by
free chlorine in synthetic tap water (290
mg-min/L) is higher than literature reported
values of 60 mg-min/L and 127 mg-min/L
for a 2-logio reduction in the same strain
when disinfection was performed at
equivalent pH and temperatures (Rice, et al.
2005; Rose, et al. 2005). Unlike the results
reported here where synthetic tap water was
utilized to simulate a real water distribution
system, the studies by Rice et. al. (1) and
Rose et. al. (2) were performed in a simple
potassium phosphate buffer. Differences in
calculated Ct values observed between
various studies have been attributed to minor
differences in water quality parameters (pH,
temperature) and different environements
during the spore preparation process, (Rose,
et al. 2005; Dow, et al. 2006). Although
chlorine demand by water components was
accounted for by contacting the water with
free chlorine for 30 minutes prior to
determining the chlorine and
monochloramine concentrations, water
constituents may still contribute to an
enhanced spore stability and disinfectant
resistance. Oxidizing agents are known to
damage the inner membrane of spores
(Cortezzo, et al. 2004). Synthetic tap water
constituents such as magnesium and
phosphorus may enhance membrane
stability (Sowers and Gunsalus, 1988;
Yethon and Whitfield, 2001; Juhna, et al.
2007) and repair oxidative damage resulting
in larger Ct values. Additionally, strain
differences and spore preparation procedures
may be responsible for the increased
resistance of BT spores to free chlorine
observed here (Ct = 620 mg-min/L for a 2-
logio reduction) in comparison to the BA
Sterne preparation and a literature reported
value of 246 mg-min/L for another strain of
B. thuringiensis (Rice, et al. 2005).
Monochloramine was less effective than free
chlorine at disinfecting the spores in solution
resulting in lower logic reductions and
higher Ct values. As previously reported,
monochloramine was less effective than free
chlorine at removing planktonic cells from
the water column (Gagnon, et al. 2004) and
free spores in suspension (Rose, et al. 2007).
A Ct value of 1,442 mg-min/L was reported
to achieve a 2-logio reduction of B.
anthracis Sterne spores with
monochloramine at 25 °C and a pH of 8.0 in
a monopotassium phosphate solution (Rose,
et al. 2007), which is consistent with values
reported here (1200 and 880 mg-min/L, for
BA and BT respectively).
Association with biofilm-conditioned pipe
surfaces increased the resistance of spores to
the two disinfectants tested. Once
associated with the biofilm-conditioned
surfaces, nearly 5 to 10 times the
disinfectant was required to inactivate the
associated spores when compared to
22
-------
concentrations needed to see the same logio
reductions in solution (a 16 to 46 fold
increase in estimated Ct values). No
significant difference in disinfectant
susceptibility was noted for the different
spore preparations (BA and the commercial
preparation of BT spores). Ct values for
free chlorine disinfection (10,000 and
28,570 mg-min/L, for BT spores associated
with PVC and copper pipe surfaces,
respectively) were comparable to the Ct
value of 31,680 mg-min/L reported by Szabo
et. al for a 2-logio reduction of B. atrophaeus
spores associated with biofilms on corroded
iron coupons (Szabo, et al. 2007).
Disinfectant efficacy was largely dependent
on the spore contact conditions.
Significantly lower BT spore association
was observed when diffusion alone was the
means of transport compared to the number
attached when a low shear was applied (60
rpm in the CDC reactor, Figure 1.3) and the
number that associated by diffusion were
easier to decontaminate (Figure 1.3). The
increase in the number of spores associating
with the biofilm-conditioned pipes is likely
due to an increase in the number of spores
coming in contact with the biofilm in the
CDC reactor where uniform mixing was
employed. However, the increase in
resistance to disinfectants when mixing is
uniform may be due to an increase in the
ability of the spores to penetrate the biofilm
matrix or change in biofilm morphology as a
result of the enhanced shear environment.
The biofilm morphology may be
significantly altered when mixing is
increased during spore contact in the CDC
reactor (60 rpm with the baffle). Altered
biofilm surface morphology can occur when
fluid shear is increased, initiating biofilm
sloughing events (Stoodley, et al. 1999;
Telgmann, et al. 2004), and contact with
biocides has been shown to alter biofilm
stability (Daly, et al. 1998; Chaw, et al.
2005). Therefore, biofilm sloughing events
that occur during spore contact may stabilize
the biofilm remaining on the pipe material
surfaces, resulting in a decrease in
subsequent vulnerability to sloughing from
additional disinfectant contact. Szabo, et. al.
observed an initial drop with a subsequent
stabilization and persistence of surface
associated spores at large disinfectant
residuals (Szabo, et al. 2007). A stable
biofilm during disinfectant contact may
explain the decreased sensitivity to the
disinfectants observed here for spores
associated under uniform mixing conditions.
The ability of the disinfectant to penetrate
the biofilm may also explain the differences
in disinfection observed for biofilm and
biofilm associated spores by free chlorine
and monochloramine. Monochloramine was
more effective than chlorine at reducing the
viable cell fraction in biofilms and biofilm
associated spores, but less effective than free
chlorine in suspension. In both Figures 1.3
and 1.4, the significant advantage of using
monochloramine to reduce the viable
fraction of associated spores and biofilm
bacteria, respectively, over free chlorine is
demonstrated. The observed enhanced
effectiveness of monochloramine compared
to free chlorine at biofilm disinfection has
been previously reported for biofilm bacteria
(LeChevallier, etal. 1990; Samrakandi, etal.
1997; Turetgen 2004). Monochloramine is
more stable and has a lower reactivity
toward inorganic and organic constituents in
the biofilm matrix resulting in a greater
disinfection ability (LeChevallier, et al.
1990; Samrakandi, etal. 1997). Chlorine
penetration of the biofilm matrix is the
limiting factor in biofilm disinfection (Chen
and Stewart, 1996). Chlorine has been
shown to be less effective than
monochloramine at disinfecting biofilm
organisms where high quantities of
extracellular polysaccharide are found, and
23
-------
the organic and inorganic constituents of
biofilms with high polysaccharide levels
consume the more reactive chlorine species
(Samrakandi, et al. 1997; Stewart, et al.
2001). Association with extracellular
matrices in natural water distribution
systems provides a survival niche for
microorganisms when disinfectant is present
(Ridgway and Olson, 1981; LeChevallier, et
al. 1987). Biofilm bacteria and spores
associated with the pipe surfaces might be
encased in a polysaccharide matrix or
shielded from the disinfectants by
extracellular components including the
inorganic material from pipe corrosion.
Bacterial attachment to biofilm-conditioned
surfaces is known to drastically increase the
inherent resistance to disinfection
(LeChevallier, et al. 1988) and cells
associated with biofilms are more resistant
to disinfection than planktonic cells
(Cochran, et al. 2000). Therefore, it is not
surprising that the association of spores with
the biofilm-conditioned pipe surfaces
resulted in an increased resistance to
disinfection. However, the degree to which
the spores are resistant and the potential for
continued sloughing and leaching of biofilm
associated spores into the water column
makes traditional chemical disinfection with
monochloramine and chlorine a less than
ideal strategy for decontamination of
Bacillus spores from treated water systems.
More research is needed to determine
alternate disinfectants and disinfection
strategies for the reduction of spores in
water systems in the case of an accidental or
intentional contamination of a water
distribution facility. Additionally, future
work should include investigating enhanced
disinfection due to increases in fluid shear
and examining the role of localized
alterations in hydrodynamic and chemical
differences due to corrosion as both have
been found to contribute to disinfectant
efficacy (De Beer, et al. 1994). Finally, this
work demonstrates the use of simulant
organisms is justified for large-scale
disinfection studies as BT spore disinfection
was either more difficult or similar to BA
inactivation in the simulated water system
reported here.
24
-------
2.0 Enhanced Decontamination of Bacillus Spores
in a Simulated Drinking Water System
Introduction
The fate of potential bioterror agents (e.g.,
Bacillus anthracis spores) in water
distribution systems is a homeland security
concern. As discussed in the previous
chapter, as well as in several recent studies,
biofilm-associated spores are dramatically
more resistant to commonly used
disinfectants as compared to their planktonic
counterparts (Ryu and Beuchat, 2005;
Morrow, et al. 2008; Szabo, et al. 2007).
Bacillus spores are more resistant to
commonly used disinfectants
(monochloramine and free chlorine) than
vegetative bacteria (Rice, et al. 2005; Rose,
et al. 2005; Rose, et al. 2007). Disinfectant
concentrations 5 to 10 times the levels
needed to reduce the viable spore fraction in
solution are required when spores are
associated with biofilm-conditioned surfaces
(Morrow, et al. 2008 ) and the spores can
persist up to a month at high (70 mg/L)
chlorine concentrations (Szabo, et al. 2007).
Spore resistance to commonly used
decontamination strategies is a concern for
the disinfection of the public water systems
in the event of a bioterrorist attack or an
unintentional release.
Many species of Bacillus and Clostridium
bacteria form spores in response to external
stress including nutrient starvation and
desiccation. The spore has a cell wall that is
similar in structure to vegetative cells
(Popham, et al. 1996) but spores also have
an additional series of membranes and a
cortex that protect the spores internal
components from heat, desiccation, UV and
oxidative damage (Hashimoto and Conti,
1971). Spores outgrow to vegetative cells
under favorable conditions initiated by small
molecules (amino acids and nucleosides) in
a process termed germination (Hashimoto
and Conti, 1971). Germination results in
degradation of the spore coat thereby
exposing the peptidoglycan layer of the
germ cell wall (Hashimoto and Conti, 1971).
Outgrowth to the vegetative cell does not
occur, however, unless sufficient nutrients
are available for restoration of full metabolic
activity (Setlow, 2003). During germination
spore components are released and lost to
the surrounding media (Kort, et al. 2005).
Such components, including dipicolinic acid
(DPA) protect the spore DNA by facilitating
internal desiccation. Upon DPA release,
sensitivity to heat inactivation increases
(Kort, et al. 2005). Spore resistance to
oxidizing agents (such as chlorine) is
primarily due to a decreased permeability
and inherently low water content of the
spore, a property that increases the
resistance of spores to heat inactivation as
well (Setlow, 1995). Oxidizing agents such
as chlorine dioxide, hydrogen peroxide and
hypochlorite damage the inner membrane of
spores making them more vulnerable to heat
treatment (Cortezzo and Koziol-Dube,
2004).
Disinfectant efficacy on biofilms and
contaminant organisms harbored by biofilms
is largely dependent on the chemical
reactivity and the ability of the disinfectant
to penetrate the biofilm matrix. Highly
reactive oxidants such as free chlorine have
been shown to be consumed by the top
layers of the biofilm matrix resulting in a
retarded penetration of the biofilm (Huang,
et al. 1995; Stewart, et al. 2001; De Beer, et
al. 1994; Chen and Stewart, 1996).
Monochloramine has been shown to be more
effective at penetrating polysaccharide
layers due to its slower reaction kinetics and
thus longer half-life (Samrakandi, et al.
1997; Turetgen, 2004; LeChevallier, et al.
25
-------
1990). Additionally, strong oxidants are
more corrosive on metal pipe materials and
are more susceptible to reaction with
corrosion byproducts, limiting their
disinfectant ability compared to less
corrosive biocides (e.g., monochloramine)
(LeChevallier, etal. 1993). Spore
germination has been studied as a means to
increase spore susceptibility to heat
inactivation and other forms of disinfection
(Kort, et al. 2005; Hornstra, et al. 2007), but
has not been applied to spores in contact
with biofilms.
Due to the unique resistance properties of
spores, improved technologies must be
developed to improve decontamination
strategies (Whitney, et al. 2003). The
objective of this work was to elucidate the
potential of combining germinant addition
with disinfection to increase the sensitivity
of Bacillus anthracis Sterne (BA) and a
commercial preparation of B. thuringiensis
(BT) spores associated with biofilm-
conditioned pipe material surfaces to
decontamination procedures. We measured
the ability of commonly used disinfectants
(free chlorine and monochloramine),
elevating the contact temperature (50 °C)
and then evaluating the impact of
disinfection and heat in combination with
germinant exposure to inactivate and
remove BA and BT spores from biofilm-
conditioned surfaces (copper and PVC).
Materials and Methods
Spore Preparations and Growth Conditions
Pure suspensions of BA were compared to a
commercially available BT preparation, for
the ability to associate with water system
biofilms and to determine their disinfectant
susceptibility. The commercial BT
preparation was washed to yield a
concentrated spore preparation at -IxlO9
CFU/mL (colony forming units/mL) by the
following procedure: aliquots of the BT
spore suspension were centrifuged at 16 000
x g for 6 minutes, supernatant was removed
and discarded and the spores were re-
suspended in phosphate buffered saline
(PBS) containing 0.01% (v/v) Triton™ X-
100 by pipetting several times and
vortexing. PBS (0.01% (v/v) Triton™ X-
100) was prepared by dissolving 8 g of
NaCl, 0.2 g of potassium phosphate, 1.15 g
of sodium phosphate and 0.2 g of potassium
chloride in 1 L of water, pH = 7.4, and
autoclaved for 15 minutes, 121 °C. 1 mL of
10 % Triton™ X-100 was added after to the
PBS after it had cooled to room temperature.
The spore pellet was washed 4 times in PBS
(0.01 % (v/v) Triton™ X-100) by
centrifuging at 16,000 x g for 6 minutes,
pipetting and vortexing to re-suspend the
sample and rinsed one additional time then
stored in 20 % ethanol at 4 °C to prevent
aggregation from residual dispersant
materials. Concentrated BA spore
preparations (Dugway Proving Ground,
Dugway, Utah) at ~3xl O8 CFU/mL were
stored in sterile MQ water (Milli-Q® ,
Millipore Corp., Billerica, Massachusetts
resistivity 18 MQ) at 4 °C. Spore
suspensions were subsequently diluted in
synthetic tap water (LeChavillier, 2005) for
disinfection studies as described in the
following sections. Spore concentrations
were determined by plating on Luria-Bertani
(LB) agar and incubating at 35 °C. PBS
(0.01 % (v/v) Triton™ X-100) was used as a
dilution buffer for biofilm and spore
enumeration. Spore preparations contained
greater than 95 % phase bright spores as
determined by phase contrast microscopy.
Spore germination was detected by plating
on LB agar before and after heat treatment at
65 °C for 25 minutes. The number of
germinated spores (vulnerable to an elevated
water temperature) was determined by the
difference in plate counts before and after
heat contact.
26
-------
Preparation of Disinfectants
Disinfectant solutions consisted of 150 mL
of synthetic tap water containing either free
chlorine at average concentrations of 11
mg/L (ranged 10 mg/L to 12 mg/L) and 103
mg/L (ranged from 98 mg/L to 108 mg/L) or
monochloramine at average concentrations
of 13 mg/L and 49 mg/L (ranged from 8
mg/L to 18 mg/L and 47 mg/L to 57 mg/L,
respectively). Free chlorine was derived
from a stock solution of sodium
hypochlorite (bleach). Free chlorine
concentrations were determined using N,N-
diethyl-p-phenylenediamine (DPD) reagent
and chlorine standards (Method #8021,
Hach Company, Loveland, Colorado).
Monochloramine solutions were prepared as
described by Camper, et. al.2003.
Monochloramine concentrations were
determined by DPD and the indophenol
method (Methods #8167, #8021, and
#10171, Hach Company, Loveland,
Colorado). Concentrations of all
disinfectant solutions were determined after
30 minutes of stirring post-disinfectant
addition to synthetic tap water to eliminate
chlorine demand contributions.
Coupon Preparation andBiofilm
Colonization of Pipe Materials
Polyvinyl chloride (PVC) and copper pipe
sections were rinsed with RO water prior to
connecting to the reactor tubing. PVC and
copper coupons were purchased from
BioSurface Technologies Corp (Bozeman,
Montana). PVC coupons were rinsed in 10
% (v/v) bleach and then RO water prior to
inserting into the reactor holder. Copper
coupons were buffed in a circular motion
with a 3M Scotch-Brite™ pad and then
soaked in 5 % (v/v) nitric acid for 30
minutes and rinsed with RO water prior to
inserting into the reactor holder.
Simulated water system biofilms were
accumulated by contacting pipe surfaces in
synthetic tap water (1.2 mM NaHCO3, 0.54
mM MgSO4-7H2O, 0.2 mM CaSO4-2H2O,
0.004 mM K2HPO4, 0.002 mM KH2PO4,
0.08 mM (NH4)2SO4, 0.17 mM NaCl, 36 nM
FeSO4-7H2O, 0.011 mMNaNO3, 0.2 mM
CaCO3, pH = 8.2 ± 0.2 (LeChevallier, M.
Personal communication, 2005)) made from
the laboratory RO water supply. Growth of
organisms indigenous to the RO water
supply system of NIST was stimulated by
adding 24 mg/L humic acid sodium salt
(Aldrich H16752-100G) to the synthetic tap
water for two weeks. A final low-carbon
adaptation period of 3 d to 5 d, achieved by
omitting humic acids, was utilized to
produce viable plate counts of biofilm
organisms consistent with literature reports
for water distribution systems (Schwartz, et
al. 2003). Biofilm organisms were
enumerated by plating on R2A media and
incubating at room temperature for up to 7
days.
CDC Biofilm Reactor Operation
The impact of a distributed shear on spore
association with the biofilm surface was
determined with a biofilm reactor designed
by the Centers for Disease Control and
Prevention (CDC). The CDC biofilm
reactor was chosen due to its history as a
model potable water system (Donlan et al,
2002) and for the ability to control the
applied fluid shear during biofilm
development and spore contact conditions
(Goeres, et al. 2005). The reactor was
assembled and vented as directed by the
manufacturer (BioSurface Technologies
Corp., Bozeman, Montana) and synthetic tap
water was supplied from a 20 L feed carboy
to the CDC reactor by a peristaltic pump via
Masterflex® tubing (Norprene® Food
Tubing 6402-14). Prepared coupons were
inserted into the coupon holders and the
reactor was filled with 400 mL of synthetic
tap water and autoclaved (121 °C for 15
minutes). Reactors were allowed to cool
27
-------
and then placed on digital stir plates and
connected to the peristaltic pump. A flow
rate of 0.3 mL/min (hydraulic retention time
of 29 hours) supplied the reactors with
synthetic tap water containing 24 mg/L
humic acids for 14 days at room
temperature. After 7 days, mixing was
introduced by turning on the digital stir plate
to begin baffle rotation at 120 ± 5 rpm
(confirmed by a tachometer) while fluid
flow was continued. After 2 weeks of
synthetic tap water plus humic acids,
reactors were switched to synthetic tap water
without humic acids, applied at the same
flow rate for 3 to 5 days.
Contacting with spores was performed by
filling the reactor vessel to the 700 mL mark
with synthetic tap water and adding the
appropriate amount of a concentrated spore
stock resulting in a concentration of 1.4 ±
0.8 x 107 CFU/mL of spores. Spores were
contacted for 24 hours under uniform
mixing in batch with an applied shear
(baffled stir bar with 180 rpm rotation) in
the CDC reactor. After this 24 hour period,
coupons mounted in holders were removed,
rinsed in sterile synthetic tap water and
either sampled directly, contacted with
disinfectants or germinants and subsequently
contacted with disinfectants.
The CDC reactor was sampled by removing
coupon holders and placing them on a sterile
piece of aluminum foil. Coupon surfaces
were scraped in a circular fashion collecting
the biofilm in the center of the coupon while
still positioned in the coupon holder (Zelver
and Hamilton, 2001; Zelver and Hamilton
1999). The method by Zelver and Hamilton,
1999 was adapted by using a spatula (made
from a sterile piece of DuPont™ Teflon®
cut to resemble a spatula) rather than a
cotton swab for scraping. Coupons were
removed from the holder with a pair of
sterile forceps, positioned over a 15 mL BD
Falcon™ tube and rinsed by dispensing 5
mL of sterile synthetic tap water directly on
the coupon surface. The rinsing procedure
was repeated two times using the same 5 mL
of synthetic tap water to minimize sample
volume. Biofilm samples were vortexed for
30 s at the highest speed to disperse the cells
and associated spores and were diluted and
enumerated under the described conditions.
Statistical Comparisons
Dunnett's multiple comparison procedure
was used for treatments versus controls. A
one-way Analysis of Variance was
performed to test the hypothesis that the
average mean value across categories of the
groups were equal. In the presence of
significance for the omnibus ANOVA test, a
Newman-Keuls multiple comparison test is
used to perform pairwise comparisons.
Statistical decisions were made at a =0.05.
Statistics were performed using WINKS
SDA Software (TexaSoft, Cedar Hill,
Texas.)
Decontamination Results
Once the biofilm was accumulated and spore
contact had occurred, coupons were
subjected to several different
decontamination strategies as shown in
Figure 2.1. Decontamination methods
included direct disinfection, germinant
addition, disinfection post germinant
contact, and heat inactivation before and
after germinant contact. In the
decontamination studies, pipe material
coupons in coupon holders were contacted
with disinfectant solutions (150 mL) in 250
mL beakers of the disinfectant solution. A
small stir bar was used to uniformly mix the
disinfectant solution during batch contact of
the CDC reactor coupons. After 30 minutes
contact times, coupons were immediately
removed and placed in 150 mL of sterile
synthetic tap water containing 7.5 mM
sodium thiosulfate to neutralize residual
28
-------
disinfectant prior to sampling. Concentrated
stocks of 10 mM inosine and 80 mM L-
alanine were prepared by dissolving
germinants in MQ water and filtering
through 0.2 um filters and stored at 4 °C
prior to use. The germinant stock solutions
were diluted directly in synthetic tap water
(pH 8.1) to a final concentration of 1 mM
inosine and 8 mM L-alanine (Barlass and
Houston, et al. 2002). Germination was
performed at room temperature (22 °C to 23
°C) by contacting coupons in holders in the
CDC reactor (containing 500 mL of
germinant solution) for 24 hours with
mixing (60 rpm). Heat treatment consisted
of placing coupon holders in preheated,
synthetic tap water in 50 mL conical tubes
(sealed with Parafilm®) at 50 °C for 25
minutes, cooling at room temperature,
followed by sampling as described above.
Spore Suspension Disinfection
The sensitivity of BA and BT spores that
detached from the coupon surface during
contact with germinant solutions was
determined for both free chlorine and
monochloramine. Germinant contact
solutions containing detached spores were
added to glass beakers containing synthetic
water with disinfectants (final
concentrations of 2 mg/mL and 10 mg/mL).
Gentle stirring with small stir bars was used
to prevent settling of spore suspensions
during contact. Viability after 30 minutes
contact was determined by sampling 100 jil
from the beaker and serially diluting in 0.3
mM sodium thiosulfate in PBS (0.01%
Triton™ X-100). All two-sample
1. Accumulate
biofilm on coupons
in the CDC Reactor
2. Add spores,
contact for 24 h
4. Add germinants
to reactor and
contact for 24 h
I
8. Remove coupon and
contact with
disinfectant or heat
(50°C, 25 min) in
beaker
5. Sample coupon and analyze for
spores, calculate reduction due to
germinants.
3. Remove coupons,
sample and analyze for
spores. Results serve as
initial surface coverage
for calculating log
reductions
6. Remove
coupon and
contact with
disinfectant or
heat (50°C, 25
min) in beaker.
9. Sample coupon and
analyze for spores,
calculate reduction due
to the combined
treatments: germinants
with either disinfectant
or heat.
7. Sample coupons and
analyze for spores,
calculate reduction due to
disinfection or heat.
Figure 2.1. Flow Chart of Experimental Approach. The experimental procedure for processing
coupons is shown.
29
-------
900
LL
"b
X
(D
o
Q.
tt>
T3
"ro
'o
o
No Baffle Baffle (60 rpm) Baffle (180 rpm)
Figure 2.2. Impact of Shear on BT Spore Association with Biofilm-Conditioned Pipe
Surfaces. Data for PVC surfaces, solid bars, and Cu, hatched bars, n > 3, error bars = 1
standard deviation.
comparisons employed Student's t-test to
calculate the t statistics assuming unequal
variances at a = 0.05. The p-values for
falsely rejecting the null hypothesis of no
difference in compared means are reported.
Results of Spore Association with Biofilm-
Conditioned Pipe Materials
This work confirms our previous report
(Morrow, et al. 2008) that increasing the
fluid shear during spore contact increases
the number of spores that associate with the
biofilm-conditioned surfaces up to the
maximum value tested of 180 rpm.
Significantly more spores (p < 0.01) were
associated with the pipe material surfaces
when the reactor was uniformly mixed and a
low shear was uniformly applied over the
coupon surfaces (180 rpm with the baffle
present in the CDC reactor) during the 24
hours contact time when compared to
previously published results (Morrow, et al.
2008) with lower shear (60 rpm) or uniform
mixing alone (Figure 2.2).
Spontaneous germination of spores
associated with the biofilm was measured by
comparing the number of associated spores
before and after contact at 65 °C for 25
minutes. Significant spontaneous
germination as indicated by heat sensitivity
was not detected. Plate counts of the
associated spores were not significantly
different before and after heat inactivation
for either PVC (p = 0.36) or copper (p =
0.26) surfaces, respectively.
Decontamination of attached spores was
attempted using disinfection
(monochloramine and free chlorine) and
elevated water temperature (50 °C). Results
fo r for BT are given in Table 2.1; results
for BA are given in Table 2.2 and Figure
2.3. The effect of germination before heat
treatment or before treatment with the
disinfectants was evaluated. Reduction of
the viable spores associated with the
biofilm-conditioned surfaces was significant
when germinant addition preceded
disinfection or heat treatment. Germinant
30
-------
addition alone reduced the viable spore
fraction by 1.5 to 3.0 logs compared to the
number that were initially adhered to the
biofilm-conditioned pipe surfaces (Tables
2.1 and 2.2). Of the remaining spores,
99.9% bound to the copper surfaces were
heat sensitive (1.3 xlO1 CFU/cm2 were still
viable after heat treatment) and no viable
spores were detected on the PVC surfaces
after heat treatment. Germination in
combination with disinfection or heat
treatment resulted in an even more dramatic
reduction in the number of viable spores
associated with the biofilm-conditioned pipe
surface numbers. Free chlorine or
monochloramine contact post-germination
resulted in a 1.5 to 3.4 logic or 2.3 to 3.4
logio reduction of attached spores,
respectively (Tables 2.1 and 2.2). BothBA
and BT spores that had detached and
germinated in the germinant solution were
inactivated (below the limit of detection of
the plate count method) by both
disinfectants at 10 mg/L and were reduced
by 1.0 and 1.2 logic after 30 minutes of
contact with 2 mg/L monochloramine and
free chlorine, respectively.
CI10
MC10
H50
Germ
-1 --
-2
o
O
O 3
*•••* O
0>
o
-4 --
-5 --
Germ +
MC10
Germ +
H50
none
detected
Figure 2.3. Impact of Germination on Decontamination of Adhered BA Sterne Spores. Log
reductions in viable spores accumulated on biofilm-conditioned pipe material surfaces, copper (solid) and
PVC (hatched), before and after germination (Germ) and after contact with 10 mg/L free chlorine (Cl 10)
and monochloramine (MC 10) for 30 min or water temperature elevated to 50 °C (H 50). Experimental
averages are shown (n > 3), ranges are indicated by bars, logic Co values were 4.1 for copper and 4.6 for
PVC. See Table 2.2 for initial spore levels.
-------
Interestingly, the most effective means of
eliminating spores from biofilm-conditioned
surfaces was completing a germination step
and then elevating the contact temperature to
50°C for 25 minutes (Tables 2.1 and 2.2. and
Figure 2.3). No viable BA spores were
detected in the solution or on the biofilm-
conditioned surface post-contact at 50 °C.
Heat inactivation was more effective at the
removal of viable BT spores from copper
pipe surfaces compared with PVC (only a
2.6 log reduction was observed for PVC
compared to below detection limits for
copper). Additionally, it is important to note
that BT spores were slightly more resistant
to decontamination methods than BA,
suggesting BT is a good simulant as it poses
a greater challenge than BA Sterne to
decontamination.
Germinant Addition Impact on Biofilm
Contact with inosine and L-alanine resulted
in an insignificant (p < 0.02, a = 0.05)
change in the surface coverage of the
biofilm (Table 2.3). Synthetic tap water
biofilms accumulated on pipe surfaces were
tested for vulnerability to free chlorine and
monochloramine, prior to and after contact
with germinant solutions. Monochloramine
resulted in a significant reduction in biofilm
surface coverage on both copper and PVC (p
< 0.03) with and without germinant addition.
Chlorine at ~ 100 mg/L resulted in a
significant decrease in the biofilm
population pre and post germinant contact (p
< 0.03) on both substrata. Chlorine at ~ 10
mg/L resulted in a significant decrease in the
biofilm population only on copper surfaces
pre and post germinant contact (p = 0.02).
Biofilms on PVC surfaces were resistant to
-10 mg/L free chlorine prior to (p = 0.13)
and post (p = 0.12) germinant addition.
Elevated water temperature in the presence
of germinant resulted in a significant
reduction in the biofilm population on
copper surfaces but not PVC.
32
-------
Table 2.1. Impact of Treatment Strategy on BT Spore Surface Coverage
Treatment
Initial
Germ
Non-Germ
CM 00
Germ
CM 00
Non-Germ
CI10
Germ
CMO
Non-Germ
MC10
Germ
MC10
Non-Germ
H50
Germ
H50
Copper Pipe Surface
T, X102 364(221)
CFU/cm2
Total LOG Reduction
PVC Pipe Surface
0.53 (0.52)*
2.8
r, xicr
556(504) 16.9(21.2)*
CFU/cm"
Total LOG Reduction
1.5
93.0(108)
0.6
188(106)
0.5
ND
=4.6
0.40 (0.39)*
3.1
1.58(1.42)* 401(701) 0.40(0.01)* 414(269)
2.4
0.0
3.0
0.0
ND
=4.6
16.2(12.8)* 25.0(16.4)* 2.70(3.06)* 217(134) 1.45(1.14)*
1.5
1.3
2.3
0.4
2.6
Total LOG Reduction is the reduction of the initial spore surface coverage (Initial) due to the individual or combined treatments;
Treatment notation is Germ = 24 h contact with 1 mM inosine and 8 mM L-alanine, Non-Germ = no germinant contact, Cl = free
chlorine with concentration, at either 10 mg/L or 100 mg/L, MC = monochloramine with concentration of 10 mg/L, H = heating to
50 °C for 25 minutes, contact with free chlorine at 10 mg/L was not performed for Non-Germ samples due to observed limited
reduction at 100 mg/L;
Values presented are averages with 1 standard deviation shown in parentheses, n > 3;
ND = none detected;
Consecutive treatments are shown with germination first and second treatment below;
indicates significant reduction in surface coverage compared to initial value (determined using Dunnett's comparison test a = 0.05
described in Statistical Comparison section).
33
-------
Treatment
Initial Germ
Copper Pipe Surface
CFU/cm2 134(123) °-13(a20)*
Total LOG Reduction 3.0
PVC Pipe Surface
CFU/cm2 363(459) 1'32(Z28)*
Total LOG Reduction 2.5
Non-Germ Germ Non-Germ Germ Non-Germ
CI10 CI10 MC10 MC10 H50
241(191) 0.13(0.23)* 14.5(2.28)* 0.10(0.17)* 50.0(25.4)*
0.0 3.0 1.0 3.1 0.4
134(77.8) 0.13(0.23)* 18.4(4.56) 0.19(0.48) 77.6(25.7)
0.4 3.4 1.3 3.4 0.7
Germ
H50
ND
=4.1
ND
=4.6
Total LOG Reduction is the reduction of the initial spore surface coverage (Initial) due to the individual or combined
treatments; Treatment notation is Germ = 24 h contact with 1 mM inosine and 8 mM L-alanine, Non-Germ = no germinant
contact, Cl = free chlorine with concentration, at either 10 mg/L or 100 mg/L, MC = monochloramine with concentration of 10
mg/L, H = heating to 50 °C for 25 minutes, contact with free chlorine at 100 mg/L for both Germ and Non-Germ was not
performed for B A spores;
Values presented are averages with 1 standard deviation shown in parentheses, n > 3;
ND = none detected;
Consecutive treatments are shown with germination first and second treatment below;
indicates significant reduction in surface coverage compared to initial (determined using Dunnett's comparison test a = 0.05
described in Statistical Comparison section).
34
-------
Table 2.3. Impact of Treatment Strategy on Biofilm Surface Coverage
Treatment
Initial Germ Non-Germ Germ Non-Germ Germ Non-Germ Germ Non-Germ Germ
CM 00 CM 00 CI10 CI10 MC 10 MC 10 H 50 H 50
Copper Pipe Surface
r, xi cr
387(217) 347(235)
CFU/cnT
Total LOG Reduction
PVC Pipe Surface
0.1
r,
715 (425) 560 (454)
CFU/cmz
Total LOG Reduction
0.1
88(120)*
0.7
183(162)*
0.6
165 (113)*
0.4
256 (87)*
0.5
116(45)* 145(48)* 15(20)*
0.5
0.4
1.4
299(184) 273(48) 22(23)*
0.4
0.4
1.5
35 (44)*
1.0
73 (79)*
1.0
109 (9)*
0.6
271 (160)
0.4
Total LOG Reduction is the reduction of the initial spore surface coverage (Initial) due to the individual or combined
treatments;
Treatment notation is Germ = 24 h contact with 1 mM inosine and 8 mM L-alanine, Non-Germ = no germinant contact, Cl =
free chlorine with concentration, at either 10 mg/L or 100 mg/L, MC = monochloramine with concentration of 10 mg/L, H =
heating to 50 °C for 25 minutes, heat contact with Non-Germ samples was not performed;
Values presented are averages with 1 standard deviation shown in parentheses, n > 3;
ND = none detected;
Consecutive treatments are shown with germination first and second treatment below;
indicates significant reduction in surface coverage compared to initial value (determined using Dunnett's comparison test a =
0.05 described in Statistical Comparison section).
35
-------
Discussion of Results
Biofilms have been implicated as reservoirs
for their ability to harbor and protect
bacterial contaminants in water distribution
systems (Morrow, et al. 2008; Szabo et al.
2007; LeChevallier, et al. 1987; Szabo, et al.
2006) and matrix materials may protect
contaminants from the disinfectants. This
work confirms previous reports (Morrow, et
al. 2008; Szabo et al. 2007) that indicate
spores are exceptionally resistant to high
levels of disinfectant when associated with
biofilm matrices and examines the role of
spore germination as a means to enhance
disinfectant efficacy.
Germinant addition resulted in a significant
reduction in the number of viable spores
associated with the biofilm-conditioned
surfaces, disinfectants and elevated water
temperatures (Table 2.1 and Figure 2.3).
The exact mechanism of the success of
germinant addition is unknown; however,
the observed vulnerability of germinated
spores is believed to be due to a combination
of physiological changes in the cell form
and the inability to compete with the native
biofilm bacteria for nutrients. Germinant
addition resulted in a significant reduction in
the viable spores associated with the
biofilm-conditioned pipe surfaces.
Ultrastructural changes occurring during
germination result in a more vulnerable cell
form as degradation of the spore coat
exposes the underlying peptidoglycan layer
of the germ cell wall to detrimental
environmental effects (Hashimoto and
Conti, 1971). Spore coat degradation and
subsequent hydrolysis of DNA and cell
proteins occurring during germination
further enhance the spore's susceptibility to
oxidants and heat that yield cell lethality
through damage of those cell components
(Setlow, 1995). Additionally, the depleted
nutrient levels in the simulated water system
are believed to limit the ability of
germinated spores to outgrow and develop
into vegetative cells as germination alone
resulted in a significant decrease in the
surface-bound viable spore fraction and
spontaneous germination was not observed.
Due in part to the depleted nutrient levels, it
is likely that the germinated spores did not
develop into fully vegetative cells, but that
the spores began the germination process
and became more susceptible to disinfection.
In addition, any germinated spores
remaining associated with the biofilm or in
the water column are vulnerable to
competition with the bacterial biofilm
organisms for available nutrients.
The impact of germinant addition on the
biofilm bacterial population was of concern
as drinking water biofilm bacteria can
readily utilize simple amino acids and
chlorine demand has been shown to increase
when biofilms are grown on amino acids
(Butterfield, et al. 2002). Interestingly,
germinant addition did not result in a
significant increase in the biofilm bacterial
population as biofilm surface coverage
values were similar before and after
germinant addition (Table 2.1).
Furthermore, disinfectant efficacy on
biofilm control was not significantly altered
for either chlorine or monochloramine.
The disinfectant efficacy was dependent on
the chemistry of the disinfectant when
germinants were not added. The higher
logio reduction of biofilm-associated spores
by monochloramine compared to free
chlorine is consistent with previous reports
that indicate its stability and lower reactivity
toward inorganic and organic constituents
results in a greater disinfection ability
toward biofilms (Samrakandi, et al. 1997;
LeChevallier, et al. 1990). Monochloramine
has been shown to be more effective at
reducing the viable cell fraction and adhered
spores in some biofilms (Morrow, et al.
36
-------
2008; (Samrakandi, etal. 1997;
LeChevallier, etal. 1990)). However,
similar log reductions in the viable spore
fraction were observed for both disinfectants
for surface bound spores when germinants
were added (Table 2.1). Finally, 2 mg/L of
both free chlorine and monochloramine was
effective at reducing the viable spore
fraction that detached in the germinant
solution by 1.0 to 1.2 logic, respectively, and
no viable spores were detected in the
germinant solution after contact with -10
mg/L of disinfectant. Disinfectant efficacy
for germinated spores in the water column is
consistent with values reported for the
inactivation of other bioterrorist agents and
vegetative cell forms of Bacillus (Rose et al.
2007; Rose et al. 2005; Beuchat, et al.
2005).
Contact at an elevated water temperature
resulted in a significant decrease in the
viable spores associated with the biofilm-
conditioned surfaces. Heat sensitivity was
enhanced with germinant addition resulting
in no detectable viable spores on the
biofilm-conditioned surfaces after contact at
an elevated water temperature. The
resistance of spores to heat is related to the
dehydration state of the cell due to the
presence of DP A (Kort, et al. 2005; Setlow,
1995). Spores are known to undergo a
phase transition from glassy-like to a
rubbery-like phase between 50 °C and 60 °C
corresponding to the release of DP A
(Alimova, et al. 2006). Such a physiological
change may yield the observed vulnerability
to the elevated water temperature (0.4 to 0.7
logio reductions) observed here and DPA
release is enhanced when germinants are
added, stimulating spore coat degradation.
Furthermore, oxidizing agents such as
chlorine dioxide, hydrogen peroxide and
hypochlorite damage the inner membrane of
spores making them more vulnerable to
sublethal heat levels and demonstrate an
increased germination rate (Cortezzo, et al.
2004).
Attachment to biofilm-conditioned surfaces
is known to increase the inherent resistance
of spores to disinfection. This work
demonstrates that adding germinants in
combination with either a disinfectant or
elevated water temperature drastically
reduces or eliminates the need for high
concentrations of reactive disinfectants to
treat spore contamination of a treated water
system.
37
-------
3.0 The Effect of High Flow on the Adhesion and
Disinfection of BT Spores in Pipe Loop Experiments
Introduction
The aim of this study is to evaluate the
effect of a high flow rate on the disinfection
step of Bacillus spores using chlorine in a
simulated drinking water system. In a
previous study (Morrow, et al. 2008), we
measured the disinfection of B. thuringiensis
(BT) and B. anthracis (B A) spores in a
simulated drinking water system using a
commercial CDC biofilm reactor and a pipe-
section loop bioreactor. Both of these
reactors have been described in previous
sections of this report. The reactors were
used to establish a water system biofilm by
contacting the plumbing materials with a
synthetic formulation of tap water
containing humic acids. The humic acids
stimulated the growth of water system
bacteria in a short period of time (three
weeks) to levels similar to those found in
mature water distribution systems
(Schwartz, et al. 2003). A stirred paddle was
used to establish a fluid shear in the CDC
biofilm reactor; and a creeping flow
(Reynolds number < 1) was used to grow
the biofilm organisms in the pipe section
reactor. An important feature of the
experimental design in the earlier study was
that the pipe section or coupons (CDC
biofilm reactor) were removed from the
bioreactors and the disinfection step was
done by contacting the pipe materials with
the chlorine solution without flow. We
found that the spores adhered to the biofilm
materials were resistant to disinfection to
chlorine solutions when using these
conditions (Morrow, et al. 2008).
In this study BT was chosen as a safe
simulant for B A because the species are
genetically closely related (Radnedge, et al.
2003) and the exosporium (outermost spore
layer) composition of the two species is also
similar (Matz and Beaman, et al. 2001). The
disinfection of Bacillus spores by chlorine
solutions is determined by the solution
conditions used and the characteristics of the
spore sample (Dychdala, 2001). Studies
have indicated that the inactivation kinetics
of BA and BT are similar (Rice, et al. 2005;
Rose, et al. 2005) validating the use of BT
as a simulant for BA, for chlorine
disinfection studies. We also observed
similar spore adhesion and disinfection
kinetics for BT and BA (Sterne strain) in a
previous study (Morrow, et al. 2008).
In this study, we focused on the effect of a
relatively high flow (Reynolds number =
2,800) on the critical disinfection step using
a pipe section bioreactor in a loop format
using local tap water supplemented with
humic acids to stimulate the biofilm growth.
A high flow rate was used in all phases of
the experiments including biofilm growth,
spore contacting, flushing, and disinfection
with chlorine. These changes were done
with the pipe section reactor to better
simulate the conditions that would be found
in a building water system.
Materials and Methods
Pipe Section Loop Reaction Protocol
Water from a cold water faucet was
collected for these experiments at the NIST
facility in Gaithersburg, Maryland. The
water obtained did not contain a significant
amount of active chlorine (less than 0.04
mg/L) and was used without further
treatment for the experiments. The pipe
section bioreactor consisted of alternating
PVC and copper pipe sections (19 mm inner
38
-------
diameter and 51 mm long) connected by
silicone tubing in a loop with a 20 L water
container vented to the atmosphere. Silicone
tubing (inside diameter 9.7 mm) and a
peristaltic pump (operated at approximately
500 rpm) was used to circulate the water
from the 20 L container through the pipe
section loop and back to the container at a
flow rate of 2.5 L/min (14.7 crn/s). The
volume of the loop section containing the
pipe sections was approximately 1 L with 36
pipe sections. During the entire biofilm
accumulation stage, the peristaltic pump had
a duty cycle of 2 hours of flow and 2 hours
no flow, established by means of an
electronic timer. Biofilm was established by
contacting the pipe sections with 20 L of tap
water containing humic acids (24 mg/L,
sodium acid for a total of 21 days. The tap
water was drained from the reactor and
renewed with a fresh solution after 7 and 14
days (for a total of 21 days of biofilm
growth with humic acids). The pipe loop
was then flushed for 1 day with 20 L of
fresh tap water (without humic acids) before
contacting with the BT spores.
The pipe section biofilm reactor was then
drained and contacted with 2 L of BT spores
(in tap water containing a total of 2 x 109
CPU) for 24 hours with flow of 2.5 L/min
(duty cycle 2 hours flow and 2 hours no
flow). A commercial BT preparation was
prepared as previously described (1). After
spore contacting, the pipe-section bioreactor
was drained, rinsed, and flushed for 2 hours
with 2 L of fresh tap water. A 10 mg/L
solution of free chlorine was prepared using
a stock solution of sodium hypochlorite
(commercial bleach) diluted in tap water.
Free and total chlorine concentrations were
measured using N, N-diethyl-p-
phenylenediamine (DPD) reagent and
chlorine standards (Method numbers #8167
and 8021, Hach Company, Loveland,
Colorado). The chlorine concentration was
measured on a spectrophotometer
(Shimadzu UV160U) after 30 minutes of
stirring. Chlorine disinfection was
performed in the entire pipe section pipe
loop at 2.5 L/min. The initial disinfection
step involved introducing 2 L of chlorine
solution (10 mg/L), which was replaced
after 30 minutes by a fresh 2 L solution of
chlorine (10 mg/L). After the disinfection
steps duplicate pipe sections of both copper
and PVC were removed and rinsed with 150
mL of tap water containing 7.5 mM
thiosulfate. The biofilm was scraped off the
pipe surfaces using sterile cell scrapers and
rinsed with phosphate buffered saline (PBS,
0.01 M phosphate, 0.138MNaCl, 0.0027 M
KC1, pH 7.4 ) containing 0.01% (vol/vol)
Triton™ X-100 and sampled as previously
described (Morrow, et al. 2008). Spore
counts were determined by plating on Luria-
Bertani agar and biofilm organisms were
counted on R2A agar plates (Morrow, et al.
2008). The stock spore samples and samples
that had been exposed to disinfectant were
concentrated by filtering half the sample (5
mL) on to a 0.45 micron filter (Nalgene
#145-2045, Rochester, NY) that was applied
directly to the nutrient plates for growth.
BT Spore Inactivation
To measure the disinfection of the BT
spores used in the experiments we measured
the inactivation of the spores in solution
with a solution of chlorine. A solution of
active chlorine (10 mg/L) was prepared in
phosphate buffer (0.1 M pH 7.8). BT spores
were suspended in the chlorine-phosphate
buffer solution at a concentration of
approximately 5 x 105 CFU/mL in a
borosilicate glass vial. The glass vial was
placed on a rocker to mix the contents
during the inactivation process. Samples
were taken at 10 minutes time intervals by
vortexing the vial (5 sec) and removing a
sample. The concentration of viable spores
39
-------
was measured as described in the previous
section.
Results and Discussion
The heterotrophic plate count of the biofilm
organisms grown on the copper and PVC
surfaces were 1.5 (1.8) x 109 CFU/m2and 9.0
(6.5) x 108 CFU/m2, respectively (means
from 4 independent experiments, values in
parenthesis are a standard error). These
numbers were consistent with values in the
literature for water distribution systems
which range from 0.23 to 8.4 x 109 CFU/m2
(Schwartz, et al. 2003).
The number of BT spores associated with
the biofilm-conditioned pipes was 5.5 (6.0)
x 108 CFU/m2 and 2.4 (3.2) x 108 CFU/m2
for copper and PVC pipe sections,
respectively (means and standard deviations
from 4 independent experiments). These
values are comparable with the previous
study using BT spores in the CDC
bioreactor with a low shear contacting
method that had 2.5 (1.6) x 108 CFU/m2 and
1.7 (1.3) x 108 CFU/m2 and for copper and
PVC pipe sections, respectively (Morrow, et
al. 2008). In this present study we used a
flow rate of 2.5 L/min in 19 mm diameter
pipe sections, resulting in a Reynolds
number of approximately 2800, indicating
the flow is transitioning to turbulent. The
resulting flow velocity was 14.7 cm/s. The
effect of a 2 hours flush with tap water on
the removal of spores was reduction by
approximately half a log (Figure 3.1).
Chlorine disinfection of the pipe loop was
done in two stages because of depletion of
the active chlorine by the large surface area
of biofilm on the pipes and tubing of the
reactor. The first chorine solution (2 L with
concentration of 10 mg/L) was depleted to
approximately half of the initial value after
30 minutes. A fresh chlorine solution was
then used for the rest of the disinfection
process. The second disinfection solution (2
L with concentration of 10 mg/L) was also
reduced to approximately half of the initial
value by the end of the second disinfection
step (150 minutes). Figure 3.1 shows the
effect of disinfection on BT spores
associated with the biofilm in the pipe
section loop. The first 30 minutes
disinfection step resulted in a reduction of
approximately 1.5 logs of the BT spores
associated with the surface. The second
disinfection step resulted in a further
reduction of approximately 1.5 logs. The
overall total reduction of BT spores was
approximately 3 to 4 logs in the pipe section
loop. Similar reductions were seen in either
PVC or copper pipe sections.
Figure 3.1 also shows the effect of the
chlorine disinfection on the removal of the
biofilm organisms. The data shows that
there is a significant decrease in the level of
biofilm organisms that correlates with the
removal of the BT spores. The biofilm
organisms on copper and PVC pipe sections
were reduced by approximately 2.5 logs
using 10 mg/L chlorine at high flow rates,
while in the previous study the reduction
was less than 1 log reduction for copper and
less than 0.5 log for PVC using 10 mg/L
chlorine for 30 minutes without flow
(Morrow, et al. 2008).
To make comparison to the disinfection
effectiveness of chlorine, it is common to
measure a CT value (the product of chlorine
concentration times the exposure time)
required to achieve a 2 or 3 log reduction in
viability. The CT value to achieve a 2 log
inactivation of BT spores in 0.1 M sodium
phosphate buffer (pH 7.8) using 10 mg/L
chlorine was 150 (60) min-mg/L (mean of 3
determinations and 1 standard deviation in
parenthesis). This value for BT inactivation
is lower than the value previously measured
in the synthetic water formulation (Morrow,
40
-------
et al. 2008), and comparable to values
measured in a potassium phosphate buffer
(pH 8) for BT (Rice, et al. 2005). The
chlorine disinfection solutions used in pipe
loop of this study were tap water with a pH
of 8.0.
Conclusions
This study shows the importance of a high
flow rate to increase the efficiency in
disinfection of spores associated with
biofilms on pipe surfaces. The high flow
likely resulted in improved penetration of
the disinfectant solution into the biofilm
layer, which resulted in better removal of the
biofilm layer and the entrapped spores. The
calculated Reynolds number (value of 2800
based on velocity of 14.7 cm/s and!9 mm
diameter pipe) for the flow rate indicated
transition to turbulent flow conditions in the
pipe section reactor. The pipe section reactor
was assembled with a number (up to 36) of
short (51 mm) pipe sections joined by
silicone tubing and several bends to achieve
a loop configuration resulting in a number of
changes in the diameter of the flow that
would make laminar flow unlikely.
Therefore, the flow was probably closer to
turbulent conditions than the calculation
suggested. The presence of turbulent flow
would be expected to increase the delivery
of fresh chlorine solution to the surface of
the biofilm layer resulting in an improved
mass transfer of active chlorine and a more
effective disinfection process. Turbulent
flow would also be expected to increase the
sloughing of the biofilm also resulting in a
more effective removal of the biofilm and
the adhered spores.
We showed that an efficient reduction
(approximately 3 logs) of spores from the
pipe surfaces can be achieved in
approximately 5 hours and using a moderate
volume of disinfectant solution
(approximately 5 volumes of the loop).
Additional experiments are needed to
determine the extent of reduction that was
due to physical removal of the spores
adhered to the biofilm due to the flushing
and the extent due to inactivation of the
spores adhered to the biofilm surfaces. An
important consideration is that the chlorine
will be consumed by the biomass in a short
period of time under recirculation
conditions, so it is important to monitor the
active chlorine concentration and replenish
it as necessary.
41
-------
.Chlorine Disinfection Time-
initial Water
Contact flush 2 h 15 min 30 min 45 min 60 min 90 min 180min
Oc
- I .U
-— «. -1 C
~3 -1.0 -
O
o 90
o
•ti o ^
D
J Q n
D)
° 35
4n
4c
c; n -
i
T
1
\
DBT Copper
D BT PVC
D Biofilm Copper
• Biofilm PVC
••
1
••
••
-•
1
-•
-r
I
1
-L
.
J
T
-•
Figure 3.1. The Effect of Water Flushing and Chlorine Disinfection Time on the Reduction of
BT Spores Associated with the Biofilm Conditioned Pipe Sections. Copper or PVC pipe
sections were sampled at the indicated stage and the log (base 10) reduction calculated by
dividing the measured BT concentration (C) by the BT concentration at the initial contact
time (Co). The conditions of the experiments were done in tap water pH 8.0 and 22 °C to 23
°C. The BT values are the means of 4 independent experiments (bars are 1 standard deviation
of the mean) and the biofilm values (for 15, 45, and 90 minutes) are the means from two
independent experiments.
42
-------
4.0 Gaps in Research on Biological Threats to be
Investigated in Additional Studies
This chapter summarizes research gaps that
were identified during the course of this
study and recommends additional research
to help bridge these gaps. The objective of
this research was to study the adherence and
decontamination of biological contaminants.
As the research progressed, knowledge gaps
were identified that presented challenges in
the development of decontamination
strategies.
Recommended Research
The following is a summary of
recommended research for bridging these
knowledge gaps:
1. Long term persistence of bacteria
and spores should be investigated
in water systems. These
experiments are technically
difficult to do and will require
significant investment in time and
effort. One approach would be to
investigate the fate of biological
threats in water system using the
fluorescent-tagged bacteria and
spores that were developed in the
third year of the project. The fate of
Bacillus spores and E. coll in
biofilms on a microscopic level
using fluorescence confocal
microscopy could be a subject of
investigation.
2. Adhesion of other biological threats
and disinfection of surfaces
contaminated by them should be
measured. Additional threat
possibilities include Yersiniapestis,
the bacterium that causes plague, and
the single-stranded RNA virus, male-
specific bacteriophage, MS2. The
MS2 phage is a commonly used as a
viral simulant in water systems.
3. Adhesion of biological threats and
simulants to additional plumbing
materials (e.g., stainless steel,
rubber, and plastic surfaces) should
be measured along with surface
decontamination. The inclusion of
used material with mineral deposits
would be valuable addition to the
existing data.
4. Transport and removal of bacteria
and spores in a large-scale pipe
system should be studied. The large-
scale experiments will require
significant planning and coordination
to carry out the experiments in order
to obtain meaningful results. The
results of the bench level
experiments completed in the current
project and discussed in this report
could be used as the basis of these
experiments. The system would
include additional water system
appliances, and complex
architecture. Experiments would
involve injection of a sample of the
simulant and measurement of the
transport in the pipe system by
sampling the water and coupons at
various points. Development of
sensors to detect the fluorescence
signatures of the simulants would
also be carried out. Classical
microbiological techniques would be
used to measure the bacterial and
identification would be confirmed by
the presence of the fluorescent tags.
5. Disinfection studies should be
carried out on large-scale systems
that have been contacted with the
bacteria. The most promising
conditions for disinfection of the
biological stimulants (determined in
43
-------
the previous years of the project)
would be used on the large scale
system to determine the efficiency of
disinfection in a near to real life
system. As in the contacting
experiments, the sampling and
analysis of these experiments would
require significant planning and
coordination of effort because of the
large number and nature of the
samples. Data on the efficiency of
the disinfection process in the large
scale systems would be valuable
because of the relevance to real life
systems.
44
-------
References
1. Alimova, A., Katz, A., Gottlieb, P., and Alfano, R. R. (2006). "Proteins and dipicolinic
acid released during heat shock activation of Bacillus subtilis spores probed by optical
spectroscopy." Applied Optics, 45(3), 445-450.
2. Almeida, J. L., Harper, B., and Cole, K. D. (2008). "Bacillus anthracis spore suspensions:
Determination of stability and comparison of enumeration techniques." Journal of
Applied Microbiology, 104(5), 1442-1448.
3. Azevedo, N. F., Pinto, A. R., Reis, N. M., Vieira, M. J., and Keevil, C. W. (2006). "Shear
stress, temperature, and inoculation concentration influence the adhesion of water-
stressed Helicobacter pylori to stainless steel 304 and polypropylene." Applied and
Environmental Microbiology, 72(4), 2936-2941.
4. Barlass, P. J., Houston, C. W., Clements, M. O., and Moir, A. (2002). "Germination of
Bacillus cereus spores in response to L-alanine and to inosine: The roles of gerL and
gerQ operons" Microbiology, 148(7), 2089-2095.
5. Beuchat, L. R., Pettigrew, C. A., Tremblay, M. E., Roselle, B. J., and Scouten, A. J.
(2005). "Lethality of chlorine, chlorine dioxide, and a commercial fruit and vegetable
sanitizer to vegetative cells and spores of Bacillus cereus and spores of Bacillus
thuringiensis." Journal of Industrial Microbiology and Biotechnology, 32(7), 301-308.
6. Butterfield, P. W., Camper, A. K., Ellis, B. D., and Jones, W. L. (2002). "Chlorination of
model drinking water biofilm: Implications for growth and organic carbon removal."
Water Research, 36(17), 4391-4405.
7. Camper, A. K., Brastrup, K., Sandvig, A., Clement, J., Spencer, C., and Capuzzi, A. J.
(2003). "Effect of distribution system materials on bacterial regrowth." Journal/
American Water Works Association, 95(7), 107-121+112.
8. Chaw, K. C., Manimaran, M., and Tay, F. E. H. (2005). "Role of silver ions in
destabilization of intermolecular adhesion forces measured by atomic force microscopy
in Staphylococcus epidermidis biofilms." Antimicrobial Agents and Chemotherapy,
49(12), 4853-4859.
9. Chen, X., and Stewart, P. S. (1996). "Chlorine penetration into artificial biofilm is limited
by a reaction-diffusion interaction." Environmental Science and Technology, 30(6), 2078-
2083.
10. Cochran, W. L., McFeters, G. A., and Stewart, P. S. (2000). "Reduced susceptibility of
thin Pseudomonas aeruginosa biofilms to hydrogen peroxide and monochloramine."
Journal of Applied Microbiology, 88(1), 22-30.
11. Cortezzo, D. E., Koziol-Dube, K., Setlow, B., and Setlow, P. (2004). "Treatment with
oxidizing agents damages the inner membrane of spores of Bacillus subtilis and sensitizes
spores to subsequent stress." Journal of Applied Microbiology, 97(4), 838-852.
12. Daly, B., Betts, W. B., Brown, A. P., and O'Neill, J. G. (1998). "Bacterial loss from
biofilms exposed to free chlorine." Microbios, 96(383), 7-21.
13. De Beer, D., Srinivasan, R., and Stewart, P. S. (1994). "Direct measurement of chlorine
penetration into biofilms during disinfection." Applied and Environmental Microbiology,
60(12), 4339-4344.
45
-------
14. Domek, M. 1, LeChevallier, M. W., Cameron, S. C., and McFeters, G. A. (1984).
"Evidence for the role of copper in the injury process of coliform bacteria in drinking
water." Applied and Environmental Microbiology, 48(2), 289-293.
15. Donlan, R. M., R.; Carpenter, I; Brown, E.; Besser, R.; Fields, B. (2002).
"Monochloramine Disinfection of Biofilm-Associated Legionella pneumophila in a
Potable Water Model System." In: Legionella., R. Marre, Kwaik, Y. A., Bartlett, C.,
Cianciotto, N. P., Fields, B. S., Frosch, M., Hacker, I, Luck, P. C., ed., ASM Press,
Washington DC, 406-410.
16. Dow, S. M., Barbeau, B., Von Gunten, U., Chandrakanth, M., Amy, G., and Hernandez,
M. (2006). "The impact of selected water quality parameters on the inactivation of
Bacillus subtilis spores by monochloramine and ozone." Water Research, 40(2), 373-382.
17. Dychdala, G. R. (2001). "Chlorine and chlorine compounds." In: Disinfection,
Sterilization, and Preservation, S. S. Block, ed., Lippincott Williams & Wilkins,
Philadelphia, 135-157.
18. Gagnon, G. A., O'Leary, K. C., Volk, C. I, Chauret, C., Stover, L., and Andrews, R. C.
(2004). "Comparative analysis of chlorine dioxide, free chlorine and chloramines on
bacterial water quality in model distribution systems." Journal of Environmental
Engineering, 130(11), 1269-1279.
19. Gibbs, S. G., Meckes, M. C., Green, C. F., and Scarpino, P. V. (2007). "Evaluation of the
ability of chlorine to inactivate selected organisms from the biofilm of a drinking water
distribution system simulator following a long-term wastewater cross-connection."
Journal of Environmental Engineering and Science, 3(2), 97-105.
20. Goeres, D. M., Loetterle, L. R., Hamilton, M. A., Murga, R., Kirby, D. W., and Donlan,
R. M. (2005). "Statistical assessment of a laboratory method for growing biofilms."
Microbiology, 151(3), 757-762.
21. Hashimoto, T., and Conti, S. F. (1971). "Ultrastructural changes associated with
activation and germination of Bacillus cereus T spores." Journal of Bacteriology, 105(1),
361-368.
22. Hornstra, L. M., de Leeuw, P. L. A., Moezelaar, R., Wolbert, E. I, de Vries, Y. P., de
Vos, W. M., and Abee, T. (2007). "Germination of Bacillus cereus spores adhered to
stainless steel." International Journal of Food Microbiology, 116(3), 367-371.
23. Huang, C. T., Yu, F. P., McFeters, G. A., and Stewart, P. S. (1995). "Nonuniform spatial
patterns of respiratory activity within biofilms during disinfection." Applied and
Environmental Microbiology, 61(6), 2252-2256.
24. Juhna, T., Birzniece, D., and Rubulis, J. (2007). "Effect of phosphorus on survival of
Escherichia coli in drinking water biofilms." Applied and Environmental Microbiology,
73(11), 3755-3758.
25. Kort, R., O'Brien, A. C., Van Stokkum, I. H. M., Oomes, S. J. C. M., Crielaard, W.,
Hellingwerf, K. J., andBrul, S. (2005). "Assessment of heat resistance of bacterial spores
from food product isolates by fluorescence monitoring of dipicolinic acid release."
Applied and Environmental Microbiology, 71(7), 3556-3564.
26. Le Dantec, C., Duguet, J. P., Montiel, A., Dumoutier, N., Dubrou, S., and Vincent, V.
(2002). "Chlorine disinfection of atypical mycobacteria isolated from a water distribution
system." Applied and Environmental Microbiology, 68(3), 1025-1032.
46
-------
27. LeChevallier, M. W., Babcock, T. M., and Lee, R. G. (1987). "Examination and
characterization of distribution system biofilms." Applied and Environmental
Microbiology, 53(12), 2714-2724.
28. LeChevallier, M. W., Cawthon, C. D., and Lee, R. G. (1988). "Factors promoting
survival of bacteria in chlorinated water supplies." Applied and Environmental
Microbiology, 54(3), 649-654.
29. LeChevallier, M. W., Lowry, C. D., and Lee, R. G. (1990). "Disinfecting biofilms in a
model distribution system." Journal/American Water Works Association, 82(7), 87-99.
30. LeChevallier, M. W., Lowry, C. D., Lee, R. G., and Gibbon, D. L. (1993). "Examining
the relationship between iron corrosion and the disinfection of biofilm bacteria." Journal
/American Water Works Association, 85(7), 111-123.
31. LeChevallier, M.W., personal communication, 2005, with Kenneth Cole providing
synthetic tap water formulation. American Water Works Service Company, Inc.:
Voorhees, NJ.
32. Manz, W., Szewzyk, U., Ericsson, P., Amann, R., Schleifer, K. H., and Stenstrom, T. A.
(1993). "In situ identification of bacteria in drinking water and adjoining biofilms by
hybridization with 16S and 23S rRNA-directed fluorescent oligonucleotide probes."
Applied and Environmental Microbiology, 59(7), 2293-2298.
33. Matz, L. L., Beaman, T. C., and Gerhardt, P. (1970). "Chemical composition of
exosporium from spores of Bacillus cereus." Journal of Bacteriology, 101(1), 196-201.
34. Morrow, J. B., Almeida, J. L., Fitzgerald, L. A., and Cole, K. D. (2008). "Association and
decontamination of Bacillus spores in a simulated drinking water system." Water
Research, 42(20), 5011-5021.
35. Popham, D. L., Helin, J., Costello, C. E., and Setlow, P. (1996). "Analysis of the
peptidoglycan structure of Bacillus subtilis endospores." Journal of Bacteriology,
178(22), 6451-6458.
36. Radnedge, L., Agron, P. G., Hill, K. K., Jackson, P. J., Ticknor, L. O., Keim, P., and
Andersen, G. L. (2003). "Genome differences that distinguish Bacillus anthracis from
Bacillus cereus and Bacillus thuringiensis." Applied and Environmental Microbiology,
69(5), 2755-2764.
37. Rasband, W. S. (1997-2007). "ImageJ." U.S. National Institutes of Health, Bethesda,
MD. http://rsb.info.nih.gov/ij/
38. Rice, E. W., Adcock, N. J., Sivaganesan, M., and Rose, L. J. (2005). "Inactivation of
spores of Bacillus anthracis sterne, Bacillus cereus, and Bacillus thuringiensis subsp.
israelensis by chlorination." Applied and Environmental Microbiology, 71(9), 5587-5589.
39. Rickard, A. H., McBain, A. J., Stead, A. T., and Gilbert, P. (2004). "Shear rate moderates
community diversity in freshwater biofilms." Applied and Environmental Microbiology,
70(12), 7426-7435.
40. Ridgway, H. F., and Olson, B. H. (1981). "Scanning electron microscope evidence for
bacterial colonization of a drinking-water distribution system." Applied and
Environmental Microbiology, 41(1), 274-287.
41. Rose, L. J., Rice, E. W., Hodges, L., Peterson, A., and Arduino, M. J. (2007).
"Monochloramine inactivation of bacterial select agents." Applied and Environmental
Microbiology, 73(10), 3437-3439.
47
-------
42. Rose, L. J., Rice, E. W., Jensen, B., Murga, R., Peterson, A., Donlan, R. M., and Arduino,
M. J. (2005). "Chlorine inactivation of bacterial bioterrorism agents." Applied and
Environmental Microbiology, 71(1), 566-568.
43. Ryu, J. H., and Beuchat, L. R. (2005). "Biofilm formation and sporulation by Bacillus
cereus on a stainless steel surface and subsequent resistance of vegetative cells and spores
to chlorine, chlorine dioxide, and a peroxyacetic acid-based sanitizer." Journal of Food
Protection, 68(12), 2614-2622.
44. Samrakandi, M. M., Roques, C., and Michel, G. (1997). "Influence of trophic conditions
on exopolysaccharide production: Bacterial biofilm susceptibility to chlorine and
monochloramine." Canadian Journal of Microbiology, 43(8), 751-758.
45. Schwartz, T., Hoffmann, S., and Obst, U. (2003). "Formation of natural biofilms during
chlorine dioxide and u.v. disinfection in a public drinking water distribution system."
Journal of Applied Microbiology, 95(3), 591-601.
46. Setlow, P. (1995). "Mechanisms for the prevention of damage to DNA in spores of
Bacillus species." Annual Review of Microbiology, 49, 29-54.
47. Setlow, P. (2003). "Spore germination." Current Opinion in Microbiology, 6(6), 550-556.
48. Sowers, K. R., and Gunsalus, R. P. (1988). "Adaptation for growth at various saline
concentrations by the archaebacterium Methanosarcina thermophila." Journal of
Bacteriology, 170(2), 998-1002.
49. Spotts Whitney, E. A., Beatty, M. E., Taylor Jr, T. H., Weyant, R., Sobel, J., Arduino, M.
J., and Ashford, D. A. (2003). "Inactivation of Bacillus anthracis spores." Emerging
Infectious Diseases, 9(6), 623-627.
50. Stewart, P. S., Rayner, J., Roe, F., and Rees, W. M. (2001). "Biofilm penetration and
disinfection efficacy of alkaline hypochlorite and chlorosulfamates." Journal of Applied
Microbiology, 91(3), 525-532.
51. Stoodley, P., Lewandowski, Z., Boyle, J. D., and Lappin-Scott, H. M. (1999). "Structural
deformation of bacterial biofilms caused by short-term fluctuations in fluid shear: An in
situ investigation of biofilm rheology." Biotechnology and Bioengineering, 65(1), 83-92.
52. Storey, M. V., and Ashbolt, N. J. (2003). "Enteric virions and microbial biofilms - A
secondary source of public health concern?" In: Water Science and Technology, 97-104.
53. Szabo, J. G., Rice, E. W., and Bishop, P. L. (2006). "Persistence of Klebsiella
pneumoniae on simulated biofilm in a model drinking water system." Environmental
Science and Technology, 40(16), 4996-5002.
54. Szabo, J. G., Rice, E. W., and Bishop, P. L. (2007). "Persistence and decontamination of
Bacillus atrophaeus subsp. globigii spores on corroded iron in a model drinking water
system." Applied and Environmental Microbiology, 73(8), 2451-2457.
55. Telgmann, U., Horn, H., and Morgenroth, E. (2004). "Influence of growth history on
sloughing and erosion from biofilms." Water Research, 38(17), 3671-3684.
56. Turetgen, I. (2004). "Comparison of the efficacy of free residual chlorine and
monochloramine against biofilms in model and full scale cooling towers." Biofouling,
20(2), 81-85.
57. Weavers, L. K. W., G. B. (2001). "Kinetics of the Inactivation of Microorganisms." In:
Disinfection, Sterilization, and Preservation, S. S. Block, ed., Lippincott Williams &
Wilkins, Philadelphia, 65-78.
48
-------
58. Yethon, J. A., and Whitfield, C. (2001). "Purification and Characterization of WaaP from
Escherichia coli, a Lipopolysaccharide Kinase Essential for Outer Membrane Stability."
Journal of Biological Chemistry, 276(8), 5498-5504.
59. Zelver, N., Hamilton, M., Goeres, D., and Heersink, J. (2001). "Development of a
standardized antibiofilm test." In: Methods inEnzymology, 363-376.
60. Zelver, N., Hamilton, M., Pitts, B., Goeres, D., Walker, D., Sturman, P., and Heersink, J.
(1999). "Measuring antimicrobial effects on biofilm bacteria: From laboratory to field."
In: Methods in Enzymology, 608-628.
49
-------
Appendix A: Protocols for Biological Threat Decontamination
List of Protocols
1. Protocol for Biosafety Level 2 Laboratory
2. Protocol for Titration of Bacillus anthracis Sterne
3. Protocol for Fluorescence In Situ Hybridization (FISH)
4. Protocol for CDC Biofilm Reactor Operation
5. Protocol for Sampling CDC Biofilm Reactor
6. Protocol for Contacting Pathogen Simulant in CDC Biofilm Reactor
7. Procedure for Free and Total Chlorine Determination
8. Protocol for Titration of Bacillus Spores
9. Protocol for Preparation of Bacillus thuringiensis spores
10. Preparation of Standard Solution of Monochloramine and Monochloramine
Measurement
11. Procedure for Disinfection of Pipe Materials Using the Contact Method
12. Protocol for Preparation of Synthetic Water
13. Protocol for Pipe Section CDC Biofilm Reactor Operation
50
-------
1. Protocol for Biosafety Level 2 Laboratory
Purpose: To use proper safety techniques when in the biosafety level 2 laboratory .
Materials:
Laboratory coat
Latex gloves
Safety glasses
Class II, Type A2 laminar flow biological safety cabinet
Clorox® Bleach, diluted 1:10 (equivalent oto 0.5 - 0.6% sodium hypochlorite)
70% ethanol
Paper towels
NuAire Operation and Maintenance Manual for Class II Biological Safety Cabinet
Methods:
1. Enter key code on the key pad to enter room A212.
2. Gloves, safety glasses, and a laboratory coat should be worn at all times.
3. Turn on the blower in the biological safety cabinet. Spray the hood surface with 70%
ethanol and wipe dry.
4. Place items to be used for the experiment (plates, dilution tubes, etc.) in the hood.
5. Allow the blower to run for 15 minutes before beginning work.
6. When working in the hood, do not make quick movements with your arms. This disrupts
the air flow. Move arms slowly to shift items within the hood.
7. Do not place any items on the front grill or against the back wall of the hood as this
disrupts air flow.
8. If a biological spill occurs, absorb the liquid using a paper towel and discard paper towel
and gloves into the biohazardous waste. Replace gloves and apply 10% bleach to the
location of the spill for 10 minutes. Wipe up the bleach with paper towel and place into
biohazardous waste. Spray surface with 70% ethanol to remove any residual bleach and
wipe clean.
9. A sterile working environment within the hood is located in the center of the hood
(approximately 6" from the front grill). It is suggested to designate a "clean" and "dirty"
side of the hood to avoid contamination.
10. Sharps are prohibited in the room (razor blades, syringe needles, scissors, etc.).
11. Design experiments to minimize aerosols whenever possible.
12. When work in the hood is completed, spray the hood surface with 70% ethanol and wipe
dry with a paper towel. Discard paper towel in biohazardous waste.
13. Pull the sliding window all the way down and turn on the UV light for 15 minutes.
14. Turn off the UV light.
15. Remove laboratory coat and place gloves in biohazardous waste.
16. Wash hands thoroughly with soap and water before exiting the laboratory.
51
-------
2 Protocol for Titration of Bacillus anthracis Sterne
Purpose: To determine concentration of Bacillus anthracis spores.
Materials:
Bacillus anthracis Sterne (suspension in sterile distilled water, stored at 4 °C)
Sterile water
Dilution tubes
Luria-Bertani (LB) agar (Fisher Chemicals #BP1425-500)
Petri dishes
Class II Type A2 laminar flow biosafety cabinet
0.5-0.6% Clorox®bleach, diluted 1:10
70% ethanol
Latex gloves
Laboratory coat
Safety glasses
Ethanol lamp
Plate spinner
Bacterial spreaders
1 mL Pipetman® (RAININ #P-1000)
200|il Pipetman® (RAININ #P-200)
Aerosol barrier pipet tips - 200|il (RAININ #GP-200F)
Aerosol barrier pipet tips - 1000|il (RAININ #GP-1000F)
Methods:
1. Refer to SOP# A212-1 for the procedures used in the biosafety level 2 laboratory.
2. In the hood, label dilution tubes and add appropriate volume of diluent (sterile water,
etc.) for dilutions.
3. Vortex vial containing Bacillus anthracis Sterne to evenly distribute the microbes.
4. Perform serial dilutions making sure to vortex between each dilution.
5. Light the ethanol lamp.
6. Add appropriate volume of dilution to LB agar plates (usually between 50-200|il).
7. Dip the end of the bacterial spreader in 70% ethanol and flame the end to burn the
ethanol off for sterilization.
8. Spread the dilution around the plate using the sterile bacterial spreader (and plate spinner
if needed).
9. Extinguish the flame in the ethanol lamp.
10. Allow plates to dry for 10 minutes.
11. Turn plates upside down and incubate at 37°C for 16 hours or until colonies are visible.
12. Count and record the number of colonies.
13. Place agar plates in the biohazardous waste.
14. Autoclave waste when it has sufficiently accumulated using a dry cycle of 40 minutes at
15-20 psi.
52
-------
3 Protocol for Fluorescence In Situ Hybridization (FISH)
Materials:
Fluorescent labeled oligonucleotide probes labeled (Alexa Fluor 546 and Alexa Fluor 647) on 5'
ends. Probe sequences as described in Manz, et al. (1993).
Beta 42a oligonucleotide probe labeled with Alexa Fluor 546 (GCC TTC CCA CTT CGT TT )
Gamma 42a probe labeled with Alexa Fluor 647 (GCC TTC CCA CAT CGT TT)
Pseudomonas probe labeled with Alexa Fluor 546 or 647 (GAT CCG GAC TAC GAT CGG
TTT)
Phosphate buffered saline (0.01 M sodium phosphate, 0.138 M NaCl, 0.0027 M KC1, pH 7.4).
95% ethanol
Microscope slides
4',6-diamidino-2-phenylindole (DAPI)
Fluorescence microscope with appropriate filters
Hybridization buffer (39% formamide, 0.9 M NaCl, 0.01 % sodium dodecyl sulfate, 0.02 M
tris(hydroxymethyl)aminomethane (Tris) pH 7.2)
Washing buffer (0.01% SDS, 0.04 M NaCl, 0.005 M (EDTA), 0.02 M Tris pH 7.2)
Methods:
Fixation for preparation of bacteria
1. Add a colony to 20|jL IX PBS, resuspend, and vortex 30 sec on high
2. Spin down cells, 16,100 x g, for 0.5 minutes; remove supernatant
3. Add 60|jL 4% PFA (PFA was at room temperature), vortex high for 30 seconds
4. Fix for 4 hours 5 minutes at 4°C
5. Spin down cells, 16,100 xg, for 0.5 minutes
6. Remove supernatant and wash with 80|jL IX PBS; vortex 15 sec
7. Wash twice with 1 X PBS
8. End fixation and storage in PBS/ethanol at -20°C
a. 10 (jL 1 X PBS + 10|jL ethanol; vortex 5 seconds
Immobilization procedure
1. Spot 3 |j,L of sample on slide and air dry
2. Dehydrate samples by dipping the slides in an ethanol series (50-80-95%; 3 minutes
each), let air dry in vertical position
Probe Procedure
53
-------
1. Add 10|jL hybridization buffer plus probe and mix carefully by pipetting
2. Pour remaining 2 mL hybridization buffer in 50 mL centrifuge tube
3. Place slide in prepared 50 ml tube, close the tube and incubate in horizontal position
at 50°C for 16 hrs.
4. Preheat wash buffer at 50°C
5. Open hybridization tube in hood, rinse with 1 mL wash buffer and place slide in wash
buffer and put the tube back at 50°C for 15 minutes
6. Rinse carefully with sterile water, let air dry
D API-staining:
- Per well, apply 10 |jL DAPI, stain for 10 minutes on ice, rinse with water, let air dry
54
-------
4. Protocol for CDC Biofilm Reactor Operation
Purpose: To grow and accumulate biofilm samples on pipe material coupons using the
CDC biofilm reactor.
Materials:
CDC biofilm reactor (CBR)
Pipe surface coupons (RD128-CU-A101 - ASTM designation of C10100 and RD128-PVC)
20 L Nalgene® Carboy (Fisher Scientific # 2210-005)
Masterflex® tubing (6424-18 Precision C-Flex® andNorprene® Food Tubing 6402-14)
Cole-Parmer Masterflex® L/S® digital console pump (Model 7524-50)
Millipore Millex®-FG vent filter (Fisher Scientific # SLFG05010)
Corning® PC-410D stirrer
Latex gloves
Laboratory coat
Safety glasses
95 % ethanol
10% bleach solution
3M Scotch-Brite™ pads (cat. 7448)
15% nitric acid
Small hex head screw driver
20 L synthetic water with and without humic acids at 24 mg/L (SOP)
Method:
1. Assemble the CBR as directed by the manufacturer with the 20 L Nalgene Carboy
with the Masterflex® tubing (Norprene® Food Tubing 6402-14) leading through the
Masterflex® peristaltic pump to the CBR influent drip.
2. Vent the reactor using the Masterflex® tubing (6424-18) and the Millipore Millex®-
FG vent filter.
3. Connect the Masterflex® tubing (6424-18) to the reactor effluent.
4. Pretreat coupons by removing any large corrosion products or debris on the copper
coupons by sanding in a circular motion with 3M Scotch-Brite™ pads, to remove
debris and large machine marks then soaked in 15% nitric acid for 30 minutes prior to
inserting into the reactor holder. PVC coupons are rinsed briefly in 10 % bleach in-
between experiments to remove any organic contaminants collected in the previous
run. Note: copper and PVC coupons are both used continuously and age with the
experimentation.
5. Insert the coupons into the coupon holder and tighten the holder screws with the small
hex head screw driver.
6. Fill the reactor to the 400 mL mark with synthetic water without humic acids, add the
coupons to the coupon holders, place them in the reactor submerged in the synthetic
water and autoclave the whole system minus the feed carboy at 121 °C for 15
minutes.
55
-------
7. Remove from the autoclave, allow liquid and reactor to cool then place the reactors
on the Corning PC-410D stir plate and connect the tubing to the Masterflex® pump.
8. Connect the tubing to the feed carboy and begin pumping synthetic water plus humics
at 1 mL/min to each reactor for 5 days at room temperature.
9. After 5 days of shear free incubation, start shear by rotating baffle at 120 ± 5 rpm by
adjusting the stir plate and confirming speed with the tachometer.
10. Continuously apply synthetic water plus humic acids for an additional 9 days (a total
of 2 weeks with humic acid application).
11. After 2 weeks of synthetic water plus humic acids, switch to synthetic water without
humic acids and apply at the same flow rate for 4 ± 1 days at room temperature.
12. Contact and sample according to appropriate SOPs.
56
-------
5. Protocol for Sampling CDC Biofilm Reactor
Purpose: To obtain biofilm samples from pipe material coupons contacted in the CBR .
Materials:
CBR grown biofilms
Latex gloves
Laboratory coat
Safety glasses
Ethanol lamp
Plate spinner
Bacterial spreaders
70 % ethanol
95 % ethanol
10% Bleach Solution
3 test tube racks
Drummond electronic pipette
ImL Pipetman® (RAININ P-1000)
200 |il Pipetman® (RAININ P-200)
Aluminum foil
Small hex screw driver provided with CDC reactor to loosen coupon holder screws.
Vortex Genie
Sterile Items (autoclave at 121 °C or purchase sterile):
10 L Sterile synthetic water (SOP KC-1)
Nutrient agar plates (R2A (VWR #EM Science 1.00416.0500) for biofilm and LB (Luria-Bertani
Miller agar plates, Fisher Scientific #BP1425-500) or selective media for pathogen simulants)
Dilution buffer (PBS, 0.01% Triton™ X-100)
1.7 mL Microcentrifuge tubes
15 mL BD Falcon™ polypropylene centrifuge tubes
5 mL Serological pipettes
Aerosol barrier pipette tips - 200|il
Aerosol barrier pipette tips - lOOOjil
DuPont™ Teflon® scrapers
Kelly Forceps
Method:
1. Refer to SOP JBM-1 for the procedures used to grow and maintain the simulated drinking
water biofilm with the CBR..
2. Prepare dilution buffer (1 x PBS with 0.01 % Triton™ X-100) by dissolving 8 grams of
sodium chloride, 0.2 grams of potassium phosphate, 1.15 grams of sodium phosphate and
0.2 grams of potassium chloride in 1 L of reverse osmosis (RO) water, test pH is at 7.4,
autoclave for 15 minutes, 121 °C. Add 1 mL of 10 % Triton™ X-100 in RO water and
mix.
3. Dispense 900 |jL of dilution buffer into 1.7 mL microfuge tubes as needed.
57
-------
4. Dispense 5 mL synthetic water into 15 mL BD Falcon™ tubes with 5 mL serological
pipette.
5. Remove the coupon holder from the CBR with gloved hands.
6. Place coupon holder on a 8 by 18 inch piece of aluminum foil wiped with 95 % ethanol.
7. Using the sterile DuPont™ Teflon® scraper, scrape the surface of the first of three
coupons with the scraper in a circular motion collecting the biofilm in the center of the
coupon.
8. Repeat the process to make sure all biofilm is loosened in the center of the coupon.
9. Fill the 5 mL serological pipette with sterile synthetic water from one BD Falcon™ tube.
10. Ethanol and flame the small hex screwdriver then loosen the coupon holder screw and
using the forceps; gently apply pressure to the coupon to pop it out of the holder.
11. Pick up the coupon with the forceps and position over the BD Falcon™ tube and position
the 5 mL pipette over the coupon and dispense the liquid directly on the coupon surface,
rinsing the biofilm from the surface.
12. Repeat the rinsing procedure by pipetting the biofilm laden synthetic water and
dispensing it directly on the coupon surface. Gently tap any residual liquid from the
coupon surface into the BD Falcon™ tube.
13. Cap the BD Falcon™ tube and vortex the tube for 30 seconds at the highest speed.
14. Dilute biofilm samples to 10"2 dilution in dilution buffer.
15. Place lOOjil of diluted biofilm sample on the surface of a labeled nutrient agar (R2A for
biofilm heterotrophic plate count) plate. With flame sterilized bacterial spreaders (soak
in the bleach solution for 10 minutes and blot dry prior to ethanol if working with
spores), careful spin and distribute the liquid over the surface of the nutrient agar plate.
Repeat according to diluted sample number.
16. Stack spread plates, then bag and store at room temperature for 7 days for biofilm and 37
°C for 16 hours for pathogen simulants.
17. Count all visible colonies after incubation period.
58
-------
6. Protocol for Contacting Pathogen Simulant in CDC Biofilm Reactor
Purpose: To monitor the association of a pathogen simulant to synthetic drinking water
system biofilms in the biofilm reactor.
Materials:
CDC Reactor (BioSurface Technolgies, Corp. Bozeman, Montana)
run according to SOP
stir plate
Latex gloves
Laboratory coat
Safety glasses
95 % ethanol
Drummond electronic pipette
ImL Pipetman® (RAININ P-1000)
200 |il Pipetman® (RAININ P-200)
10% bleach solution
Tubing clamp
Sterile Items (autoclave at 121 °C or purchase sterile):
5 mL serological pipettes
Aerosol barrier pipette tips - 200|il
Aerosol barrier pipette tips - lOOOjil
200 mL Nalgene low-form polypropylene beakers
3 L synthetic water without humic acids
Nutrient agar plates
Dilution buffer (PBS, 0.01% X-100 ™)
1.7 mL microcentrifuge tubes
1 L Kimax tall form beaker
1 L Kimax tall form beaker with a stir bar
LB, Miller agar plates (Fisher Scientific #BP1425-500)
LB, Lennox broth (Fisher Scientific #BP1427-500)
Organisms:
Escherichia coli O157:H7 , nontoxigenic, biosafety level 1
Bacillus thuringiensis kurstaki, Thuricide, Bonide Products Inc. Oriskany, New York, washed as
described in Protocol 9 of this appendix, " Protocol for Preparation of Bacillus thuringiensis
spores "
Method:
1. Accumulate the synthetic drinking water system biofilm and sample any background
biofilm controls needed as described in SOP JBM-1 prior to starting the pathogen
contact.
2. Use the tubing clamp to clamp off the reactor effluent tube.
3. Fill the reactor beaker to the 700 mL mark with synthetic water at a flow rate of 3
mL/min.
59
-------
4. Prepare dilution buffer (1 x PBS with 0.01 % Triton™ X-100) by dissolving 8 grams of
sodium chloride, 0.2 grams of potassium phosphate, 1.15 grams of sodium phosphate and
0.2 grams of potassium chloride in 1 L of RO water, test pH is at 7.4, autoclave for 15
minutes, 121 °C. Add 1 mL of 10 % Triton™ X-100 in RO water and mix.
5. Dispense 900 |jL of dilution buffer into 1.7 mL microfuge tubes as needed.
6. Add the appropriate amount of the pathogen simulant to the beaker to get a final
concentration of 3.2 ± 1.6 x 107 cells/mL for E. coli O157:H7 and 1.4 ± 0.8 x 107
CFU/mL for B. thuringiensis spores.
7. Mix at 125 rpm using the CBR baffled stir bar for 30 minutes then remove 1 mL from the
supernatant, dilute and plate to determine initial concentration of simulant.
8. Start the stir plate set at either a high shear (180 rpm) or low shear (60 rpm) by adjusting
stir plate digital setting.
9. Serially dilute the supernatant sample by transferring 100 \\L of the sample into the 900
\\L of dilution buffer in a 1.7 mL microfuge tube, vortexing and repeating in the next
dilution tube.
10. Spread plate the dilutions by placing lOOjil of diluted sample on the surface of a labeled
LB agar plate. With flame sterilized bacterial spreaders (soak in the bleach solution for
10 minutes and blot dry prior to ethanol if working with spores), careful spin and
distribute the liquid over the surface of the LB agar plate. Repeat according to diluted
sample number.
11. Stack spread plates, then bag and store at 37 °C for 16 hours for pathogen simulants.
12. Contact the CBR with the pathogen simulant for 24 hours at room temperature.
13. Sample the supernatant liquid as needed by removing a desired volume of liquid from the
reactor with the 5 mL serological pipette.
14. Add 150 mL of the sterile synthetic water to a 200 mL Nalgene low-form polypropylene
beakers; prepare two beakers per coupon holder rod.
15. After the contact time has ceased, turn off the stir plate and remove the coupon holders,
placing them directly in the first 150 mL of sterile synthetic water.
16. Transfer the coupon holder to a new Nalgene low-form polypropylene beaker of sterile
synthetic water.
17. Sample the coupons as described in SOP.
18. Once the samples are collected in the 15 mL BD Falcon™ tubes, spread plate as
described in steps 9 and 10 on appropriate nutrient agar. For Bacillus spore contacted
samples, place dilution tubes at 65 °C for 25 minutes to inactivate any vegetative cells
providing an estimate of Bacillus spore concentration, then plate on nutrient agar.
60
-------
7. Procedure for Free and Total Chlorine Determination
Purpose: To determine the amount of free and total chlorine in a sample.
Quality Assurance:
1. Run a chlorine standard with each set of measurements, prepared as described below.
2. Run a standard curve as described below every month when doing measurements or
repeat if standard is different than standard curve by more than 5%.
Materials:
SwifTest Bulk Dispenser, with DPD Total Chlorine Refill (Hach #2802400)
Total chlorine refill (Hach #2105660)
SwifTest Bulk Dispenser, with DPD Free Chlorine Refill (Hach #2802300)
Free Chlorine refill (Hach #2105560)
Chlorine Standard Solution, 50-75 mg/L as C12, pk/16 10 mL Ampules (Hach #1426810)
15mL conical tubes
ImL cuvettes
Methods:
Free chlorine
1. Place 10 mL of sample in tube or appropriate dilution in a 10 mL volume. Add 1
addition of DPD from Swiftest.
2. Cap tube and mix tube by inversion to dissolve powder. Start a timer for 1 minute.
3. Place sample in cuvette and read absorbance at 515 nm at 1 minutes.
Total chlorine
1. Place 10 mL of sample in tube or appropriate dilution in a 10 mL volume. Add 1
addition of DPD from Swiftest.
2. Cap tube and mix tube by inversion to dissolve powder. Start a timer for 3 minutes.
3. Place sample in cuvette and read absorbance at 515 nm at 3 minutes.
Preparation of Chlorine Standards by Serial Dilutions:
Should be linear from 0.02 to 2 mg/mL (maybe 4)
Stock solution is 66 mg/L.
1. Add 0.606 mL of stock to 9.3 94 mL of water (4 mg/mL)
2. Add 0.303 mL of stock to 9.697 mL of water (2 mg/mL)
3. Add 0.152 mL of stock to 9.848 mL of water (1 mg/mL)
4. Add 0.076 mL of stock to 9.924 mL of water (0.5 mg/mL)
5. Add 0.015 mL of stock to 9.985 mL of water (0.1 mg/mL)
Plus blank (no stock)
Example of Values from Standard Curve
61
-------
E
c
m
S>
Q
O
Standard Curve for Total and Free Chlorine
1 9
-l
I
n R
0 fi
.4 -
00
n
y = 0.2546x + 0.01 44 j±
R2 = 0.9986 ^
y = 0.2609X - 0.0028 ^^"^
R2 = 0.9988 ^Jjf^^
^.^''^ Afree chl
i^^^ • total chl
B^*
012345
Chlorine (mg/L)
62
-------
8. Protocol for Titration of Bacillus Spores
Purpose: To determine concentration of Bacillus spores.
Materials:
Bacillus anthracis
Sterile water
Dilution tubes
Luria-Bertani agar (Fisher Chemicals #BP1425-500)
Petri dishes
Class II laminar flow biosafety cabinet
0.5-0.6% Bleach, diluted 1:10
70% Ethanol
Latex gloves
Lab coat
Safety glasses
Ethanol lamp
Plate spinner
Bacterial spreaders
ImL Pipetman® (RAININ P-1000)
200 |il Pipetman® (RAININ P-200)
Aerosol barrier pipet tips - 200|il (RAININ #GP-200F)
Aerosol barrier pipet tips - 1000|il (RAININ #GP-1000F)
Methods:
1. Refer to SOP for the procedures used in the biosafety level 2 lab.
2. In the hood, label dilution tubes and add appropriate volume of diluent (sterile water,
etc.) for dilutions.
3. Vortex vial containing Bacillus spores to evenly distribute the microbe.
4. Perform serial dilutions making sure to vortex between each dilution.
5. Light the ethanol lamp.
6. Add appropriate volume of dilution to agar plates (usually between 50-200|il).
7. Dip the end of the bacterial spreader in 70% ethanol and flame the end to burn the
ethanol off for sterilization.
8. Spread the dilution around the plate using the sterile bacterial spreader (and plate
spinner if needed).
9. Extinguish the flame in the ethanol lamp.
10. Allow plates to dry for 10 minutes.
11. Turn plates upside down and incubate at 37°C for 16 hours or until colonies are
visible.
12. Count and record the number of colonies.
13. Place agar plates in the biohazardous waste.
14. Autoclave waste when it has sufficiently accumulated using a dry cycle of 40 minutes
at 15-20psi.
63
-------
9. Protocol for preparation of Bacillus thuringiensis spores
Purpose: To prepare a working stock of Bacillus thuringiensis spores.
Materials:
Thuricide, Bonide Products Inc. (contains B. thuringiensis kurstaki spores,)
Sterile PBS/0.01%Triton™ X-100
Sterile water
Sterile 1.5mL microfuge tubes
Luria-Bertani agar (Fisher Chemicals #BP1425-500)
Petri dishes
Microfuge (capable of 16,000xg-)
20% Ethanol
Sterile 50 mL conical tube
Latex gloves
Lab coat
Safety glasses
ImL Pipetman® (RAININ #P-1000)
Aerosol barrier pipet tips - lOOOjil (RAININ #GP-1000F)
Methods:
1. Shake the Bonide Thuricide bottle vigorously to mix the solution.
2. Aliquot ImL of Bonide solution into ten microfuge tubes.
3. Spin down spores in microfuge tubes for 6 minutes at 16k xg.
4. Remove the supernatant.
5. Wash the spore pellet by resuspending the pellet in ImL of sterile water by pipetting
up and down.
6. Vortex all the tubes.
7. Repeat step 3.
8. Wash the pellet once more with water (repeat steps 4-7).
9. Wash pellet twice with PBS/0.01% Triton™ X-100.
10. Resuspend pellet in 20% ethanol and pool all suspensions into a sterile 50mL conical
tube.
11. Store at 4°C.
12. Quantify the spore stock.
64
-------
10. Preparation of Standard Solution ofMonochloramine and Monochloramine Measurement
Purpose: To prepare a concentrated solution of monochloramine and measurement of
mochloramine in solutions.
Quality Assurance:
1. Run a monochloramine with each set of measurements, prepared as described below.
2. Run a standard curve as described below every month when doing measurements or
repeat if standard is different than standard curve by more than 5%.
Materials:
Potassium phosphate dibasic (Sigma #P8281)
Ammonium chloride (Riedel-de Haen #11209)
Bleach (Clorox®)
Buffer Powder Pillow, pH 8.3 (Hach #89868)
Chlorine Standard Solution, 50-75 mg/L as C12, pk/16 10 mL ampules (Hach #1426810)
monochloramine F pillows (Hach #28022-46)
Nitrogen, Ammonia Standard Solution, 100 mg/L (Hach #2406549)
Glass beakers, volumetric flasks, balance, pipettors, stir bars
Method:
To prepare 100 mL of monochloramine solution (approximately 300-250 mg/L)
1. Add 0.5g potassium phosphate to 1L RO water in a sterile glass beaker with stir bar. Stir
solution to mix. Check the pH of the solution (should be in the range of 8.9-9.2). Adjust
pH if necessary.
2. Add 0.1 Ig ammonium chloride to 100 mL of phosphate solution in a glass beaker and stir
using a stir bar.
3. Prepare a 4% bleach solution.
4. In a chemical hood, add 200|il per minute of the 4% bleach solution to the ammonium
chloride solution for 5 minutes (total volume added is ImL) while the solution is mixing.
NOTE: Solution may get hot - do not add more than 200|il of the bleach solution per
minute. Chlorine gas is released; solution must be made in the hood.
5. Allow solution to stir for 30 minutes in the hood. If making several preparations of 100
mL of monochloramine, after the 30 minutes stirring in the hood, the solutions can all be
combined at this point before determining the concentration.
65
-------
Procedure
1. Prepare the monochloramine standard fresh before use.
2. Add the contents of one Buffer Powder Pillow, pH 8.3 to about 50-mL of
organic-free water in a clean 100-mL Class A volumetric flask. Swirl to
dissolve the powder.
3. Using a Class A volumetric pipet, transfer 2.00 mL of Nitrogen, Ammonia
Standard Solution, 100 mg/L as NH3-N into the flask.
4. Dilute to volume with organic-free water, cap and mix thoroughly. This is a
2.00 mg/L buffered ammonia standard.
5. Pipet 50.00 mL of the buffered ammonia standard into a clean 100-mL beaker.
Add a stir bar.
6. Obtain a recent lot of Chlorine Solution Ampules, 50-70 mg/L, and note the
actual free chlorine concentration for this lot.
7. Calculate the amount of Chlorine Solution to be added to the ammonia
standard using the following equation:
example
mL chlorine solution required = 4557 (free chlorine concentration)
our stock is 66 mg/L so 455/66 = 6.89 mL
8. Open an ampoule and, using a glass Mohr pipet, add the calculated amount of
Chlorine Solution slowly to the ammonia standard, while mixing at medium
speed on a stir-plate.
9. Allow the monochloramine solution to mix for 1 minute after all Chlorine
Solution is added.
10. Quantitatively transfer the monochloramine solution to a clean 100-mL
Class A volumetric flask. Dilute to the mark with organic-free water, cap, and
mix thoroughly. This is a nominal 4.5 mg/L (as Cl2) monochloramine
standard. USE STANDARD WITHIN ONE HOUR OF PREPARATION
Indophenol Method of Monochloramine Assay (HACK 10172)
1. Turn on spectrophotometer to warm up and change wavelength to 655 nm.
2. Pipette 2 mL sample (or diluted sample range up to 10 mg/L) in cuvette place in
instrument and press auto zero to blank.
3. Add contents of 1 pillow of monochlor F (Hach #28022-46) and seal cuvette with
Parafilm®, start timer, and mix contents for about 20 seconds to dissolve.
4. After 5 minutes place in spectrophotometer and read absorbance.
5. Construct standard curve using standard
66
-------
11. Procedure for disinfection of pipe materials using the contact method
Purpose: To disinfect pipes or pipe materials that have biofilms and other
biologicals associated with them using various disinfectants.
Materials:
Monochloramine solution
Chlorine solution (dilute Clorox®)
0.1M Sodium thiosulfate (Sigma #87026)
Potassium phosphate monobasic (Mallinckrodt #7100)
Magnesium chloride hexahydrate (Fluka #63068)
250 mL plastic beakers
Synthetic water
Stainless steel forceps
0.45|im bacterial plate filter (Nalgene #145-2045)
LB agar (Fisher Chemicals #BP1425-500)
R2A agar (EM Science #1.00416.0500)
Cell scraper (Nunc™ #179693)
DuPont™ Teflon® scraper
Petri dish
70% ethanol
10% bleach
Method:
1. Prepare the dilution buffered solution (0.0425g/L KH2PO4, 0.405g/L MgCl2-6H2O).
Autoclave the solution.
2. Prepare the disinfectants of a certain concentration.
3. Prepare 0.0075M sodium thiosulfate.
4. Transfer the pipe/pipe material to a beaker containing 150 mL of disinfectant (either
monochloramine or chlorine).
5. Allow the pipe material to contact the disinfectant for a specified amount of time.
6. Remove the pipe material using sterilized forceps (70% ethanol and flame) and place it
into a new 250 mL beaker containing 150 mL of synthetic water and 0.0075 M sodium
thiosulfate (this concentration will stop the chlorine at 100 mg/L concentrations of MC
and free chlorine).
7. Add 11.2 mL of 0.1M sodium thiosulfate to the 150 mL of disinfectant (0.0075 M
concentration of sodium thiosulfate) that the pipe material was being contacted in to stop
further action of the chlorine/chloramine. Filter the 150 mL using a Nalgene 0.45|im
filter and place the filter onto a LB agar plate or a R2A agar plate using sterilized forceps.
8. Sample the pipe/pipe material by scraping the surface either with a DuPont™ Teflon®
scraper that has been sterilized using 70% ethanol and a flame, or by using a sterile cell
scraper, and rinse the scraping into a 50 mL conical tube with 10 mL of dilution buffered
solution.
67
-------
12. Protocol for Preparation of Synthetic Water
Purpose: To obtain water for biofilm growth.
Materials:
RO water
Sodium bicarbonate (Sigma #8-7277)
Magnesium sulfate heptahydrate (Sigma #M5921)
Calcium sulfate dehydrate (Sigma #C3771)
Potassium phosphate, dibasic (Sigma #P8281)
Potassium phosphate, monobasic (Mallinckrodt #7100)
Ammonium sulfate (Sigma #A4418)
Sodium chloride (J.T. Baker #3624-01)
Iron (II) sulfate heptahydrate (Sigma #F7002)
Humic acid sodium salt (Aldrich #H16752)
Sodium nitrate (Sigma #8-5022)
Calcium carbonate (Sigma #C6763)
Methods:
SYNTHETIC SOLUTION PREPARATION
Preparation of Synthetic Solution—10L are prepared at a time
1
2
O
4
5
6
7
8
9
10
11
Chemicals needed
NaHCO3
MgSO4.7H2O
CaSO4-2H2O (corrected from
anhydrous)
K2HPO4 (dibasic)
KH2PO4 (monobasic)
(NH4)2S04
NaCl
FeSO4- 72H2O (corrected from
anhydrous)
Humic acid sodium salt
NaNO3
CaCO3
F.W.
84.0
246.5
172
174.2
136.1
132.1
58.4
278.0
84.99
100.1
Cone mM
1.2
0.054
0.2
0.004
0.002
.00008
.00017
.000036
0.011
0.2
mg/lOL
1000
134
342
7.0
3.0
0.1*
0.1*
0.01**
80-240
10
200
* Solutions prepared at 1 mg/mL add 0.1 mL to 10 L
68
-------
** Solution prepared at 1.82 mg/mL add 0.01 mL to 10 L
Add powders to 10 L water and stir. Check pH and conductivity
This synthetic water, theoretically, should yield 2.0mg/L total organic carbon (TOC). If
4.0mg/L TOC is desired, then 16mg/L humic acid sodium salt will have to be added. The
addition of humic acid to the synthetic water causes a visible color to form.
Synthetic water is prepared with 20mg/L CaCO3. If lOOmg/L CaCO3 is required, add an
additional 80mg/L CaCO3 to synthetic water.
Measure synthetic water for the following:
pH 7.6-7.8
Conductivity +/- 10% of nominal (TBD)
Turbidity < 0.1 NTU (prior to addition of humic acid)
Chlorine < 0.05 ppm
TOC +/- 10% of nominal (2mg/L or 4mg/L)
The pH adjustment is done with IN NaOH to get 8.5 pH and IN HC1 to lower pH to 7.2.
The prepared synthetic water does not have any storage conditions, no refrigeration
necessary.
Temperature: room temperature, 4°C
pH: 7.2, 8.5
Hardness: lOOmg/L NaHCO3/20 mg/L CaCO3, lOOmg/L NaHCO3/100mg/L CaCO3
TOC: 2mg/L, 4mg/L (8mg/L humic acid =2mg/L TOC, 16mg/L humic acid=4mg/L)
Incubation period: 7 days
Formula taken from: AwwaRF 2981, Standard Operating Procedures for Decontamination of
Water Infrastructure, Phase la Report by Mark LeChevallier and Stacey Spangler
69
-------
13. Protocol for Pipe Section CBR Operation
Purpose: The operation of a bioreactor designed to grow a biofilm on the inner surface of a
pipe section. The methods for contacting the pipe sections with a biological agent, sampling the
inner surface, and sampling the disinfectant solution are also included.
Materials:
Safety gloves
Lab coat
Safety glasses
Pipe sections made of standard plumbing materials (typically 0.5 inch to 0.75 inch in
diameter by 2 inches long)
Silicone tubing (inside diameter to be slightly smaller than outside diameter of pipe sections
being used)
Plastic connectors and diameter reducers
Plastic cell scrapers (Nunc™ #179693)
R2A agar plates (EM Science #1.00416.0500)
Synthetic tap water (referrer to SOP#KC-1 for formulation)
Humic acids sodium salt (technical grade Aldrich Chemical #H16752).
Sterile dilution buffer solution (0.405 g/L MgCl-6H2O, 0.0425 g/L KH2PO4)
Sterile dilution tubes (plastic)
Rubber stoppers
Automatic pipette devices
Plastic pipits
Peristaltic pump with silicone tubing (flow rates range of 0.5 mL/min to 2 mL/min)
Plastic 20 L carboys
Volumetric cylinders of various sizes
Stopwatch
Monochloramine solution (refer to SOP# JLA-6)
Chlorine solution (dilute Clorox")
Sodium thiosulfate (Sigma #87026)
Potassium phosphate monobasic (Mallinckrodt #7100)
Magnesium chloride hexahydrate (Fluka #63068)
250 mL plastic beakers
Synthetic water
Stainless steel forceps
0.45|im bacterial plate filter (Nalgene #145-2045)
LB agar Fisher Chemicals #BP1425-500)
R2A agar (EM Science #1.00416.0500)
Cell scraper (Nunc™ #179693)
DuPont™ Teflon® scraper
Petri dish
70% Ethanol
10% Bleach
70
-------
Methods:
1. The synthetic tap water was prepared in large 20 L plastic carboys. The water used was
for the formulation of the synthetic tap water was reverse osmosis prepared in house
prepared and the resulting solutions were not sterilized. The carboys have lids with a
sterile vent filter (0.22 micron) that were connected to the peristaltic pump.
2. The pipe sections were assembled by the use of 5 cm lengths of silicone tubing. The
silicone tubing inner diameter should be a snug fit around the outer diameter of the pipe
section. It should be sufficiently snug to prevent leakage of the solution. Pipe sections
were linked in chains up to 18 sections per fluid path. Different pipe section materials can
be alternated in the chain. Additional chains can be placed in parallel using another fluid
paths to obtain more pipe sections for large experiments.
3. Flow rate was established by pumping synthetic water containing humic acids (24 mg/L)
for a period of 14 days. The fluid paths are kept free of bubbles by slightly raising the
dripping end of the pipe section chain. Flow rate was determined by collection the fluid
dripping out the end of a chain into a volumetric cylinder and measuring the time with a
stopwatch. Typical flow rates were 1 mL/min for 0.75 inch inside diameter pipe sections.
4. At 7 days, midway through the growth period, the pump was stopped and the pipe
sections were disconnected from the pump, rubber stoppers inserted into the ends, and the
chain of pipe sections were reversed. The stoppers were removed and the pipe section
chains were reconnected to the pump, so that the section that was nearest the pump is
now furthest away from the pump. The pump was turned back on and the growth stage
continued for another 7 days.
5. At the end of the 14-day growth period using synthetic tap water with humic acids, the
flow was stopped. The plastic 20 L carboy is switched to a new carboy containing only
synthetic tap water and the flow started again. This period of starvation was continued for
3 days.
6. At the end of this period, the flow was stopped and pipe sections were disassembled. Pipe
sections were placed in sterile synthetic tap water until used for contacting with a
biological threat agent or used to determine the level of biofilm bacteria on the pipe
section.
7. To contact a pipe section with a biological agent (such as bacterial spores or vegetative
cells) the pipe sections were placed in beaker containing the contacting solution. The
solution was gently stirred by use of a DuPont™ Teflon® coated stir bar to prevent
settling of the spores or bacterial cells. After contacting the pipe section was removed
from the contacting solution and gently rinsed in sterile synthetic water for five minutes
with gentle agitation of the solution to wash away the unbound biological agent.
8. The contacted pipe sections can be treated with a disinfectant solution to inactivate the
biological agent. The rinsed pipe section was placed into a sterile 250 mL beaker
containing 150 mL of the disinfectant solution (typically chlorine or monochloramine).
After the desired time, the pipe section was removed using sterilized forceps and placed
into a fresh 250 mL beaker containing 150 mL of synthetic water (for chlorine and
monochloramine solutions the solution contained 7.5 mM sodium thiosulfate to stop the
action of the chlorine and monochloramine). The surface of the pipe section may then be
sampled.
71
-------
9. To sample for the presence of viable biological agents in the disinfectant solution the
disinfectant must be neutralized. A solution of thiosulfate (11.2 mL of 0.1M) was added
to neutralize 150 mL solutions of chlorine or monochloramine (up to 100 mg active
chlorine/L). A portion or the entire volume was vacuum filtered through a sterile 0.45|im
filter. The filter was then placed on top of a nutrient agar plate (LB agar for bacteria and
spores or a R2A agar for biofilm bacteria) using sterilized forceps. The plates with the
filters were placed into an incubator (30 °C or 37 °C) for up to 6 days and checked daily
for growth of bacterial colonies.
10. The pipe sections were sampled for biofilm bacteria or biological agent by material by
scraping the surface either with a sterile cell scraper. The inside of the pipe section was
rinsed with 10 mL of the dilution buffer solution into a 50 mL conical tube. The tube was
vigorously vortexed several times for 10 sec. Dilutions were made and the solution was
spread on nutrient agar plates (LB agar for bacteria and spores or a R2A agar for biofilm
bacteria) in triplicate. The plates were placed into an incubator (30 °C or 37 °C) for up to
6 days and checked daily for growth of bacterial colonies.
72
-------
United States
Environmental Protection
Agency
PRESORTED STANDARD
POSTAGE & FEES PAID
EPA
PERMIT NO. G-35
Office of Research and Development (8101R)
Washington, DC 20460
Official Business
Penalty for Private Use
$300
------- |