Evaluation of Vacuum-based Sampling Devices

EPA 600/R-13/137 | August 2013 | www.epa.gov/ord
Evaluation of Vacuum-based
Sampling Devices for Collection of
Bacillus Spores from Environmental
Surfaces
Assessment and Evaluation Report
Office of Research and Development
National Homeland Security Research Center
EPA 600-R-13-137
Evaluation of Vacuum-based Sampling Devices for Collection of
Bacillus spores from Environmental Surfaces
Assessment and Evaluation Report
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
ii
Disclaimer
The United States Environmental Protection Agency (EPA), through its Office of Research and
Development’s National Homeland Security Research Center, managed this investigation through EP-C09-027 WA 2-10 and 3-10 with ARCADIS U.S., Inc. The effort was funded through an interagency
agreement (RW-75-92345701) with the US Centers for Disease Control and Prevention (CDC). This
report has been peer and administratively reviewed and has been approved for publication as an EPA
document. It does not necessarily reflect the views of the EPA. No official endorsement should be
inferred. This report includes photographs of commercially available products. The photographs are
included for purposes of illustration only and are not intended to imply that EPA approves or endorses the
product or its manufacturer. The EPA does not endorse the purchase or sale of any commercial products
or services.
Questions concerning this document or its application should be addressed to:
M. Worth Calfee, Ph.D.
Decontamination and Consequence Management Division
National Homeland Security Research Center
U.S. Environmental Protection Agency (MD-E343-06)
Office of Research and Development
109 T.W. Alexander Drive
Research Triangle Park, NC 27711
Phone: 919-541-7600
Fax: 919-541-0496
E-mail: calfee.worth@ epa.gov
iii
Acknowledgments
This effort was managed by the principal investigator from the National Homeland Security Research
Center (NHSRC) within EPA’s Office of Research and Development. Laura Rose (CDC’s Clinical and
Environmental Laboratory Branch, Division of Healthcare Quality Promotion (DHQP) within the National
Center for Emerging and Zoonotic Infectious Diseases (NCEZID) and Dino Mattorano (EPA’s Chemical,
Biological, Radiological, and Nuclear Consequence Management Advisory Team (CBRN CMAT) within
the Office of Emergency Management (OEM) provided critical inputs into this investigation. Additionally,
the efforts of Stephen Morse and Angela Weber (CDC’s Office of Environmental Microbiology, Division of
Foodborne, Waterborne, and Environmental Diseases (DFWED), within the NCEZID are gratefully
recognized.
The authors would like to thank the peer reviewers for their significant contributions. Specifically, the
efforts of Lisa Delaney (National Institute for Occupational Safety and Health (NIOSH)), Matt Arduino
(CDC), and Erin Silvestri (EPA) are recognized.
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Table of Contents
Disclaimer ............................................................................................................................................. iii
Acknowledgments ................................................................................................................................ iv
Table of Contents .................................................................................................................................. v
List of Appendices............................................................................................................................... vii
List of Tables ...................................................................................................................................... viii
List of Acronyms and Abbreviations .................................................................................................... x
Executive Summary............................................................................................................................. xii
1
Introduction .................................................................................................................................... 1
1.1
Process ..................................................................................................................................... 1
1.2
Project Objectives...................................................................................................................... 1
1.3
Experimental Approach ............................................................................................................. 2
1.3.1
Vacuum-based Sampling Device Evaluation Tests – Phase 1 ............................................ 2
1.3.2
HVAC Inoculation Tests – Phase 2 .................................................................................... 2
1.3.3
HVAC Sampling Tests – Phase 2....................................................................................... 2
1.3.4
Inoculum Variability and Level Tests – Phase 3.................................................................. 3
1.3.5
Linearity Recovery Tests and Laboratory Variability Tests – Phase 3 ................................. 3
1.4
2
Definition of Sampling Efficiency ................................................................................................ 3
Materials and Methods ................................................................................................................... 4
2.1
Test Materials and Deposition.................................................................................................... 4
2.1.1
Coupon Preparation........................................................................................................... 4
2.1.2
Bacillus Spore Preparation................................................................................................. 6
2.1.3
Coupon Inoculation ............................................................................................................ 6
2.2
Test Matrix ................................................................................................................................ 9
2.3
Sampling and Analytical Procedures ........................................................................................ 11
2.3.1
Swab Sampling................................................................................................................ 11
2.3.2
Polyester-Rayon Blend (PRB) Wipe Sampling ................................................................. 11
2.3.3
Sponge-wipe Sampling .................................................................................................... 12
2.3.4
Vacuum Sock Sampling and Analysis .............................................................................. 12
2.3.5
37 mm MCE and 37 mm PTFE Sampling ......................................................................... 13
2.3.6
Trace Evidence Filter Sampling ....................................................................................... 14
2.3.7
Operational Assessment .................................................................................................. 14
2.3.8
HVAC filter extraction....................................................................................................... 15
v
2.3.9
Quartz extraction ............................................................................................................. 15
2.3.10
Direct Extraction of Stainless Steel .................................................................................. 15
2.4
Sampling Strategy ................................................................................................................... 16
2.4.1
2.5
Preventing Cross-contamination during Coupon Preparation ............................................ 18
2.5.2
Preventing Cross-contamination during Sampling ............................................................ 18
2.5.3
Preventing Cross-contamination during Analysis .............................................................. 19
2.5.4
Sample Containers .......................................................................................................... 19
2.5.5
Sample Identification ....................................................................................................... 19
2.5.6
Information Recorded by Field Personnel......................................................................... 20
2.5.7
Sample Preservation ....................................................................................................... 20
2.5.8
Sample Holding Times ..................................................................................................... 20
2.5.9
Sample Custody .............................................................................................................. 20
2.5.10
Sample Archiving............................................................................................................. 21
Statistical Analysis Methods..................................................................................................... 21
Results and Discussion ............................................................................................................... 22
3.1
4
Sampling Handling and Custody .............................................................................................. 18
2.5.1
2.6
3
Sampling/Monitoring Points ............................................................................................. 16
Phase 1 ................................................................................................................................... 22
3.1.1
Results of Statistical Analyses ......................................................................................... 24
3.1.2
Operational Parameters ................................................................................................... 24
3.1.2.1
Sampling Time and Ease ......................................................................................... 24
3.1.2.2
Analysis Time and Ease ........................................................................................... 25
3.1.2.3
Cost ......................................................................................................................... 25
3.2
Phase 2 ................................................................................................................................... 26
3.3
Phase 3 ................................................................................................................................... 30
Quality Assurance ........................................................................................................................ 35
4.1
Sampling, Monitoring, and Analysis Equipment Calibration ...................................................... 35
4.2
Data Quality ............................................................................................................................ 36
4.3
QA/QC Checks ........................................................................................................................ 36
4.4
Acceptance Criteria for Critical Measurements ......................................................................... 38
4.5
Data Quality Audits .................................................................................................................. 39
4.6
QA/QC Reporting .................................................................................................................... 39
vi
5
Summary....................................................................................................................................... 40
5.1
Phase 1 ................................................................................................................................... 40
5.2
Phase 2 ................................................................................................................................... 40
5.3
Phase 3 ................................................................................................................................... 40
5.4
Lessons Learned and Application of Vacuum-based Methods to Field Use .............................. 40
5.5
Future Research ...................................................................................................................... 42
References ........................................................................................................................................... 43
List of Appendices
Appendix A
Miscellaneous Operating Procedures (MOPs) included as a separate document
vii
List of Figures
Figure 2-1.
Carpet Coupon .................................................................................................................. 4
Figure 2-2.
Upholstery Coupon ............................................................................................................ 5
Figure 2-3.
Concrete Coupon .............................................................................................................. 5
Figure 2-4.
Schematic of Deposition Apparatus for Inoculation under Flow Conditions ......................... 8
Figure 2-5.
Vacuum Sock Kit and Individual Sock. ............................................................................. 13
Figure 2-6.
Filter (37 mm) Cassette with Nozzle and Tubing .............................................................. 14
Figure 2-7.
3M® Trace Evidence Filter (TEF) Vacuum-based Sampling Device .................................. 15
Figure 3-1.
Recovery from Materials for each Vacuum Method. . ...................................................... 22
Figure 3-2.
Relative Recovery – Data presented as mean relative recovery (RR) ............................... 23
Figure 3-3.
Recovery from HVAC Filters Using 5 Sampling Methods – Test HI2................................. 27
Figure 3-4.
Recovery from Mechanical MERV 8 Filter – Test HS ....................................................... 28
Figure 3-5.
Recovery from Electrostatic HVAC Filter – Test HS.......................................................... 29
Figure 3-6.
Recovery from Stainless Coupons during INOC Series Tests........................................... 31
Figure 3-7.
Mean CFU/Sample from Linear Recovery (LR) Test......................................................... 32
Figure 3-8.
Laboratory Variability for Two Vacuum Methods - Test LV ............................................... 34
List of Tables
Table 2-1.
Phase 1Test Matrix ............................................................................................................ 9
Table 2-2.
Phase 2 Test Matrix ........................................................................................................ 10
Table 2-3.
Phase 3 Test Matrix ......................................................................................................... 10
Table 2-4.
Frequency of Sampling Monitoring Events ....................................................................... 17
Table 2-5.
Critical and Non-Critical Measurements ........................................................................... 17
Table 3-1.
Recovery from Materials for each Vacuum Method .......................................................... 22
Table 3-2.
Relative Recoveries from all Devices and Material Surface Types ................................... 23
Table 3-3.
Average Recovery from HVAC Filter Extraction – Test HI1 .............................................. 26
Table 3-4.
Recovery from HVAC filters – Test HI2 ............................................................................ 26
Table 3-5.
HVAC Vacuuming Recovery – Test HS............................................................................ 27
Table 3-6.
Student’s t-test values from Electrostatic filters ................................................................ 28
Table 3-7.
Mean Recovery from Stainless Steel Coupons (n = 3), Phase 2 Tests ............................. 30
Table 3-8.
ANOVA – Comparison of the Three Stainless Steel Recovery Methods within each
Phase 2 Test Run (n = 3)................................................................................................. 30
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Table 3-9.
Mean Recoveries from Stainless Steel using the Sponge-wipe Method - Test INOC
(n = 10)............................................................................................................................ 30
Table 3-10.
Mean Recoveries (CFU/sample) from Test LR (n=5)........................................................ 31
Table 3-11.
Percent Relative Recovery of Vacuum Devices (Test LR) (n = 5) ..................................... 32
Table 3-12.
Relative Recovery of Vacuum Methods to Sponge-Wipe – Test LR (n=5)......................... 33
Table 3-13.
Laboratory Variability for Two Vacuum Methods (Test LV) ............................................... 34
Table 4-1.
Sampling and Monitoring Equipment Calibration Frequency............................................. 35
Table 4-2.
QA/QC Sample Acceptance Criteria ................................................................................ 37
Table 4-3.
Critical Measurement Acceptance Criteria........................................................................ 38
Table 5-1.
Summary of Advantages and Disadvantages of Each Vacuum-Based Sampling
Method. ........................................................................................................................... 41
ix
List of Acronyms and Abbreviations
%Cv
ADA
ANOVA
APPCD
ATCC
CBRN
CDC
CFM
CFU
CM
2
cm
CMAT
COC
DCMD
DFWED
DHQP
DPG
DQI
DQO
ECBC
EPA
EtO
2
ft
HAZMAT
H2O2
HVAC
ID
in.
INL
INOC
ISO
L
-1
L min
LR
LV
MCE
MDI
MERV
MOP
NCEZID
NHSRC
NIST
OEM
PBS
Percent Coefficient of Variation
aerosol deposition apparatus
Analysis of Variance
Air Pollution Prevention and Control Division
American Type Culture Collection
Chemical, Biological, Radiological, and Nuclear
Centers for Disease Control and Prevention
cubic feet per minute
colony forming unit(s)
critical measurement
square centimeter
Consequence Management Advisory Team
chain of custody
Decontamination and Consequence Management Division
Division of Foodborne, Waterborne, and Environmental Diseases
Division of Healthcare Quality Promotion
Dugway Proving Ground
Data Quality Indicator
Data Quality Objective
Edgewood Chemical Biological Center
U. S. Environmental Protection Agency
ethylene oxide
square feet
Hazardous Materials
hydrogen peroxide
heating, ventilation, and air conditioning
Internal Diameter
inch(es)
Idaho National Laboratory
Inoculation
International Organization for Standardization
liter(s)
liters per minute
Linear Range
Laboratory Variability
Mixed Cellulose Ester
metered dose inhaler
Minimum Efficiency Reporting Value
Miscellaneous Operating Procedure
National Center for Emerging and Zoonotic Infectious Diseases
National Homeland Security Research Center
National Institute of Standards and Technology
Office of Emergency Management
Phosphate Buffered Saline
x
PBST
PRB
PTFE
PVC
rpm
RR%
RSD
QA
QAPP
QC
SCFM
SOP
STS
TEF
VHP
WAM
®
Phosphate Buffered Saline with 0.05% TWEEN 20
Polyester-Rayon Blend
Polytetrafluoroethylene
Polyvinylchloride
rotations per minute
Relative Recovery Percentage
Relative Standard Deviation
Quality Assurance
Quality Assurance Project Plan
Quality Control
standard cubic feet per minute
Standard Operating Procedure
Sodium Thiosulfate
Trace Evidence Filter
Vaporous Hydrogen Peroxide
Work Assignment Manager
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Executive Summary
The existing surface sampling strategy for a biological incident involving B. anthracis spores necessitates
the use of various methods depending on the surface type. Currently-recommended surface sample
collection methods include pre-moistened wipes (for smooth nonporous surfaces), vacuuming (for rough
and porous surfaces), and wet swabs (for small and/or hard-to-sample areas such as keyboards). The
currently-used vacuum-based method utilizes woven collection socks attached to a cardboard nozzle.
Some criticisms of the current method are that the vacuum socks often come from the manufacturer with
visible holes in the sock seams, the method is vulnerable to cross-contamination between samples, the
socks are constructed of materials with large pore-sizes (> 1µm), and the filters are cumbersome for
laboratory handling and extraction during analysis. This project comparatively evaluated the vacuum sock
and two additional vacuum-based collection devices (37 mm filter cassette and 3M™ Trace Evidence
Filter) for their sampling efficacy. The 37 mm filter cassette was evaluated with mixed cellulose ester
(MCE) filters or polytetrafluoroethylene (PTFE) filters installed, each was considered a unique device.
These data were generated so appropriate sampling devices could be selected following a B. anthracis
incident.
A known quantity of Bacillus atrophaeus (B. anthracis surrogate) spores was aerosolized and deposited
2
onto large coupons (1 square foot (ft )) of various materials common to the built environment, including
carpet, upholstery, unpainted (smooth finish) concrete, and two types (electrostatic and mechanical) of
heating, ventilation and air conditioning (HVAC) filters. Coupons were then subjected to vacuum-based
sampling and sample analysis according to protocols developed jointly by the US Centers for Disease
Control and Prevention (CDC) and US Environmental Protection Agency (EPA). Recovery was
determined for each sampling method according to culture-based microbiological assays following
physical extraction methods developed by the CDC.
Phase 1 of this study included the evaluation of vacuum-based sampling devices. These tests were
conducted with four different devices and one sampling variation (fast or slow sampling rate at which the
device passed over the sampled surface) for the vacuum sock. Samples were collected from three porous
surface types for each device, each with ten replicate samples. Vacuum and wipe samples were also
collected from stainless steel coupons as a method to standardize results collected on different days and
with different inoculation devices.
For carpet, the slow vacuum sock method had the highest spore recovery. For concrete, the 37 mm MCE
had the highest spore recovery. For upholstery, the 37 mm MCE had the highest spore recovery. The
vacuum sock method afforded more ease of use and may therefore be more desirable for larger sample
areas. However, on concrete and upholstery, the 37 mm MCE filter method demonstrated higher
recoveries per unit area than did the vacuum sock device. The significance of these differences was
dependent upon the statistical test performed. There are advantages and disadvantages of each device.
Changing the speed at which the vacuum sock sampling device passed over the surface did not
necessarily improve the recovery of spores from the porous surfaces tested.
During Phase 2, collection of B. atrophaeus spores from contaminated HVAC filters with two vacuumbased devices (devices with the highest recoveries from Phase 1, vacuum sock (slow rate) and 37 mm
MCE cassette) was compared to sampling by extractive methods (removing a portion of the filter and
extracting spores from the filter matrix). Contamination of HVAC filters during a biological incident, unlike
xii
most surfaces, is expected to occur under flow conditions, with HVAC blowers pulling spores to the
interior of the filter. A method was developed which successfully deposited spores on (and within) HVAC
filters under flow conditions.
Two types of clean (new) HVAC filters (electrostatic and mechanical) were inoculated similarly to Phase
1, yet under flow conditions. The contaminated surfaces were then evaluated using three methods: direct
extraction from excised sections of the HVAC filter, vacuum sock sampling, and 37 mm MCE filter
vacuum sampling. In addition, the magnitude and variability of the inoculating metered dose inhaler (MDI)
was evaluated with three methods: sponge-wipe sampling of stainless steel coupons, pre-moistened
(polyester-rayon blend, PRB) wipe sampling of stainless steel coupons, and direct extraction of stainless
steel coupons.
Three methods (two vacuum methods and one extractive) of sampling HVAC filters performed reliably on
mechanical and electrostatic filters. The data suggest that extractive methods are more efficient than
vacuum-based recovery methods for electrostatic filters, but not for mechanical filters. When comparing
the two vacuum-based methods, the vacuum sock method provided higher recoveries than the 37 mm
MCE method from electrostatic filters, but there was no statistical difference in recoveries from the
electrostatic filters. Vacuum-based methods may be more applicable to HVAC filter media that are not
easily sectioned.
Phase 2 data also suggested there was no statistical difference between direct extraction, sponge-wipes,
or pre-moistened PRB wipes for recovery (inoculum magnitude) from stainless steel coupons. Mean
variability (%Cv) was also similar across all methods, at 28%, 22%, and 20% for the extraction-based
method, sponge-wipe method, and PRB wipe method.
During Phase 3, Inoculum Variability and Level Tests (INOCs) were conducted to determine the
repeatability and magnitude of inoculation from the MDI/ aerosol deposition apparatus (ADA) dosing
4
6
method. Three inoculum levels (corresponding to three concentrations of spores within MDIs - 10 , 10 ,
7
10 spores per coupon) were used to dose up to ten replicate stainless steel coupons. The MDI
inoculation method was demonstrated over a broad range of surface contamination concentrations, from
4
2
1 to 1 × 10 CFU/square centimeter (cm ).
Also during Phase 3, Linearity Recovery (LR) Tests and Laboratory Variability (LV) Tests were conducted
to determine: (1) if vacuum-based method recoveries were linear over a range of concentrations; and (2)
if recoveries varied significantly between two different technicians processing (extracting and plating) the
samples. During these tests, carpet coupons were sampled using two chosen vacuum sampling
methods: vacuum sock (slow) and 37 mm MCE. Following collection, all samples were extracted and
plated by two technicians. All samples for both devices and all three inoculum levels were collected on a
single day so that comparisons could be made across devices, inoculum levels, and technicians. One
technician operated the vacuum-based collection device for all samples. Recoveries were compared
between the two technicians to determine if technician-induced variability was significant. The linearity of
recoveries for each device was determined graphically by plotting recovery versus the targeted inoculum.
The two vacuum methods were demonstrated effective for all inoculum concentrations. Compared to
sponge-wipes (from stainless steel), the vacuum sock method recovered between 3% and 22% CFU,
while the 37 mm MCE method recovered between 4% and 140% CFU from carpet samples. Relative
recoveries greater than 100% were possible because these values were determined by dividing vacuum
method recovery by wipe-based recovery from stainless steel. Greater than 100% relative recovery
xiii
indicated that the vacuum method out-performed the wipe method. Finally, the extraction procedures for
both vacuum methods were evaluated by two independent laboratory technicians with no statistical
difference in the two recoveries.
These data were collected to aid in sampling device and strategy selection following a biological
contamination incident.
xiv
1
Introduction
The U.S. Environmental Protection Agency (EPA) conducted a study, in collaboration with the Centers for
Disease Control and Prevention (CDC), to evaluate several vacuum-based sampling devices for
collection of biological agent from environmental surfaces. Methods for detection and characterization of
biological agent on surfaces following a bioterrorism incident include the use of swabs, wipes, and
vacuum. Vacuum-based methods are preferred when sampling porous surfaces. Currently, there are no
vacuum-based methods validated for collection of Bacillus anthracis spores. Further, multiple vacuumbased devices, sample collection methods, and sample extraction methods were utilized to characterize
the extent of contamination following the 2001 anthrax incidents. Recently, some work has been
conducted to characterize the performance of vacuum-based surface sampling devices when used to
collect Bacillus spores [1-3]. However, significant gaps remain in our understanding of vacuum-based
sampler performance, efficiencies, ease of use, and applicability to various material surface types. The
current study was conducted to generate data that could be used to inform selection of appropriate
sampling methodologies following a B. anthracis incident. Scientifically tested sampling methods will
provide increased confidence in the ability to characterize contamination following such an incident.
1.1
Process
Consistent with previous sampling studies [1], spores of Bacillus atrophaeus (formerly known as Bacillus
globigii or Bacillus subtilis var. niger) served as surrogates for Bacillus anthracis spores. Collection of B.
atrophaeus spores from multiple surface types was evaluated with four vacuum-based sampling devices.
In addition, collection of biological agent from contaminated heating, ventilation, and air conditioning
(HVAC) filters with the two top-performing vacuum-based devices was compared to sampling by
destructive methods (removing a portion of the filter and extracting spores directly from the filter matrix).
A known quantity of aerosolized dry B.atrophaeus spores was gravitationally deposited onto large
coupons (35.6 cm x 35.6 cm) of various materials common to the built environment, which included
carpet, upholstery, unpainted (smooth finish) concrete, and two types of HVAC filters. The coupons were
then subjected to vacuum-based sampling according to protocols developed jointly by the CDC and EPA.
Recovery was determined for each sampling method according to culture-based microbiological assays
following physical extraction methods developed by the CDC. All test parameters, such as coupon
materials and sizes, sampling methods, and methods of extraction and analysis were determined by
agreement among participating experts from EPA and the CDC.
1.2
Project Objectives
The objective of this project was to evaluate four currently-available vacuum-based devices for their
ability to recover Bacillus spores from environmental surfaces. Performance (relative recovery) of the
devices was compared to recoveries from pre-moistened gauze wipes, used to sample reference
stainless steel coupons that were inoculated at the same time as the test coupons. Evaluation of
operational parameters included time required for sample collection, the physical impact on the sampling
team during collection, time required for sample analysis, and the cost of media and analysis equipment
and supplies. Another objective of this study was to evaluate vacuum-based sampling of contaminated
HVAC filters to that of extraction-based methods.
1
1.3
Experimental Approach
The experimental approaches that were used to meet the objectives of this project are:
•
Use of controlled chambers, standardized sections and spore inoculums.
•
Inoculation of material coupons via aerosol deposition of bacterial spores.
•
Quantitative assessment of spore recoveries with each device, by sampling representative
sections of materials and then extraction of sampling media to recover collected spores
(enumerated and reported as colony forming units (CFU)).
•
Use of reference stainless steel coupons to quantify and standardize results for cross-comparison
of tests.
•
Documentation of operational considerations (e.g., cross-contamination, procedural time, impacts
on materials and personnel).
All testing was conducted at EPA’s Research Triangle Park, NC, campus.
1.3.1 Vacuum-based Sampling Device Evaluation Tests – Phase 1
The Vacuum-based Sampling Device Evaluation Tests were conducted with four different device types
and one sampling variation (sampling speed) for one device type (Table 2-1). Samples were collected
from three surface types (carpet, upholstery, and concrete) for each device with ten replicate samples.
Pre-moistened polyester rayon blend (PRB) wipe samples were also collected from reference stainless
steel coupons as a method to standardize results collected on different days and with different inoculation
devices. The foundation for this test matrix was described in the Quality Assurance Project Plan (QAPP)
entitled, “Evaluation of Vacuum-based Sampling Devices for Collection of Biological Agent
(Decontamination and Consequence Management Division (DCMD) 3.60)” (available upon request).
1.3.2 HVAC Inoculation Tests – Phase 2
HVAC Inoculation (HI) Tests were conducted to determine the best inoculation method for HVAC filters.
Contamination of HVAC filters during a biological incident, unlike most surfaces, is expected to occur
under flow conditions, with HVAC blowers pulling spores to the interior of the filter. Also, the porous
nature of the substrate presents challenges to even the settling-based aerosol inoculation. For this
reason, a modified version of the method developed by Calfee et al., 2013, for depositing aerosolized
spores on material surfaces, was utilized for this substrate. These tests were conducted to verify that the
inoculation target (quantity of spores) could be achieved without excessive variability.
1.3.3 HVAC Sampling Tests – Phase 2
HVAC Sampling (HS) Tests were conducted to determine spore recovery from HVAC filters. Two types of
HVAC filters (electrostatic and mechanical) were inoculated under flow conditions. The contaminated
surfaces were then evaluated using three methods: direct extraction, vacuum sock sampling, and 37 mm
MCE filter vacuum sampling. In addition, the variability of the inoculating method was evaluated with two
2
sampling methods: sponge-wipe sampling of stainless steel coupons and direct extraction of stainless
steel coupons.
1.3.4 Inoculum Variability and Level Tests – Phase 3
Inoculum Variability and Level (INOC) Tests were conducted to determine the repeatability and
3
magnitude of inoculation from metered dose inhaler (MDI) dosing method. Three inoculum levels (10 ,
6
7
10 , 10 spores per coupon) were deposited onto as many as ten replicate stainless steel coupons.
Coupons were sampled by sponge-wipe following the required 18-h deposition time.
1.3.5 Linearity Recovery Tests and Laboratory Variability Tests – Phase 3
Linearity Recovery (LR) Tests and Laboratory Variability (LV) Tests were conducted to determine: (1) if
vacuum-based recoveries were linear over the three inoculation levels demonstrated in the INOC tests;
and (2) if recoveries varied significantly between two different technicians processing (extracting and
plating) the samples. During these tests, carpet coupons were sampled using two chosen vacuum
sampling methods: vacuum sock and 37 mm MCE filters. Following collection, all samples were extracted
and plated by two laboratory technicians. All samples for both devices and all three inoculum levels were
collected on a single day so that comparisons could be made across devices, inoculum levels, and
laboratory technicians. Recoveries were compared between the two laboratory technicians to determine
if technician-induced variability was significant. The linearity of recoveries for each device was determined
graphically by plotting recovery versus the targeted inoculum.
1.4
Definition of Sampling Efficiency
The recovery from vacuum sampling methods was compared to recovery from wipe-based methods (PRB
wipes or sponge-wipes, or both, dependent upon the test) used to sample stainless steel surfaces with
identical inoculums. Surface samples from stainless steel were considered the best estimate of the
number of spores inoculated by the aerosol method due to the high repeatability and high recovery
efficiencies when wipe sampling from this surface type. When side-by-side comparisons could not be
made (from separate test days due to the large number of vacuum samples prescribed by the test matrix),
results were normalized to the stainless steel surface samples.
3
2
2.1
Materials and Methods
Test Materials and Deposition
2.1.1 Coupon Preparation
Carpet coupons (Figure 2-1) were 30.5 cm x 30.5 cm (12” x 12”) carpet tiles (Shaw Living Berber sand
loop, Home Depot, Model # 3W05300100) attached to a 35.6 cm x 35.6 cm (14” x 14”) square of 1.9 cm
(¾”)-thick plywood. The tiles were attached to the plywood using their self-adhesive backs plus three
staples on each side.
Figure 2-1.
Carpet Coupon
Upholstery coupons (Figure 2-2) were constructed from a 61.0 cm x 61.0 cm (24” x 24”) piece of fabric
(Indoor/Outdoor Modern Houndstooth Red Fabric, www.fabric.com, Part# UJ-849) covering a 30.5 cm x
30.5 cm (12” x 12”) piece of upholstery padding (432 Poly Foam, 2.5 cm by 61.0 cm (1” x 24”),
OnlineFabricStore, Item# 1243310), placed in the center of a 35.6 cm x 35.6 cm (14” by 14”) square of
1.9 cm (¾”)-thick plywood. The fabric square was stretched over the foam and wood, and excess fabric
was folded underneath and stapled to the back side of the plywood backing.
4
Figure 2-2.
Upholstery Coupon
Concrete coupons (Figure 2-3) were formed from Sakrete Sand Mix poured into custom 35.6 cm x 35.6
cm (14” by 14”) forms. The sand mix was prepared according to indications on the package using a
trough and garden hose for the water supply. The coupon was smoothed with a trowel and allowed to set
and dry overnight. The coupons were allowed to cure under humid conditions for at least five days.
Figure 2-3.
Concrete Coupon
Mechanical MERV 8 HVAC filters were 35.6 cm x 35.6 cm x 2.5 cm (14” x 14” x 1”) Purafilter 2000 Blue
series (Purafilter 2000, Las Vegas, NV; http://purafilter2000.com/products.php). Electrostatic MERV 8
filters were 35.6 cm x 35.6 cm x 2.5 cm (14” x 14” x 1”) Eco-Aire MERV 8 High Cap (Con-Air Industried,
Inc., Orlando, FL). Stainless steel coupons were Grade 316, cut to 35.6 cm x 35.6 cm (14” by 14”) or 10.2
cm x 15.2 cm (4” x 6”). Only the centermost 30.5 cm x 30.5 cm (12” x 12”) of each 35.6 cm x 35.6 cm
(14” by 14”) coupon was sampled.
5
All coupons were placed in sterilization bags and sterilized prior to use. Carpet and upholstery coupons
were sterilized by a minimum 250 ppmv Vaporous Hydrogen Peroxide (VHP) cycle for four hours using a
STERIS ED1000. Concrete and stainless steel coupons were sterilized prior to use by steam autoclave
utilizing a gravity cycle program consistent with the EPA NHSRC Microbiology Laboratory Miscellaneous
Operating Procedure (MOP) 6570 (see Appendix A for all associated MOPs). HVAC filters were sterilized
with ethylene oxide. Sterility was evaluated by swab sampling one coupon from each sterilization batch
according to MOP 3135. Prior to use, the coupons treated with VHP were incubated at 30-35 °C for a
minimum of two days to force off-gassing of residual hydrogen peroxide (H2O2) from material coupons as
suggested by Baron et al. [4], so that biocidal activity was prevented.
2.1.2 Bacillus Spore Preparation
The test organism for this work was a powdered spore preparation of B. atrophaeus (American Type
Culture Collection (ATCC) 9372) and silicon dioxide particles. The preparation was obtained from the
U.S. Army Dugway Proving Ground (DPG) Life Science Division, in Dugway Utah. The preparation
procedure is reported in Brown et al. [5]. Briefly, after 80 – 90 percent sporulation, the suspension was
centrifuged to generate a preparation of approximately 20 percent solids. A preparation resulting in a
powdered matrix containing approximately 1 x 1011 viable spores per gram was prepared by dry blending
and jet milling the dried spores with fumed silica particles (Deguss, Frankfurt am Main, Germany). The
powdered preparation was loaded into MDIs by the U.S. Army Edgewood Chemical Biological Center
(ECBC) or by ARCADIS according to a proprietary protocol. Control checks for each MDI were included in
the batches of coupons contaminated with a single MDI.
2.1.3 Coupon Inoculation
Coupons were inoculated with spores of B. atrophaeus from an MDI using the procedure described by
Calfee et al. [6] and detailed in MOPs 3161-LD and 3161-HD for low dose (≤2 x 104) and high dose (>2 x
4
10 ) concentrations. Briefly, each coupon was inoculated independently by being placed into a separate
aerosol dosing apparatus (ADA) designed to fit one 35.6 cm x 35.6 cm (14” by 14”) coupon of any
thickness. In accordance with MOP 3161-LD or -HD, the MDI was discharged into the ADA a single time
2
for most concentrations, but three discharges were administered when the 2 x 10 colony forming units
2
(CFU) per dose MDI was used, for a total dose of ~6 x 10 CFU per coupon for these inoculations. The
spores were allowed to settle onto the coupon surfaces for a minimum period of 18 h. When porous
coupons were used (HVAC filters), coupons were placed on double-coated carpet tape (Polyken Model
105C) to prevent re-entrainment of spores during handling and sampling. After the minimum 18-h period,
the coupons were then removed from the ADA and sampled. The ADAs were removed from only those
coupons required for a single sample at a time (some samples were comprised of three coupons, others
a single coupon). After use, the coupons were placed in a bin of soapy water before disposal. The
handling of the contaminated coupons was done in a way to minimize or control spore dispersal. One
person was tasked with removing the clamps holding the ADA to the coupon and the removal of the ADA
and gasket from the coupon. A second person, wearing new gloves for each coupon, was then tasked
with placing the sampling template atop the coupon. A third person executed the actual sample collection
procedure (e.g., vacuum-device usage, wipe-based sample collection, etc). The first person then
removed the coupon for neutralization. All personnel conducted the same job throughout the entirety of a
test operation.
6
The MDIs are claimed to provide 200 discharges per MDI. The number of discharges per MDI was
tracked so that use did not exceed this value. For any inoculation event for all three materials, a new MDI
had to be used to avoid exceeding 200 discharges per MDI. Additionally, in accordance with MOP 3161LD and -HD, the weight of each MDI was determined after completion of the inoculation of each coupon.
For quality control of the MDIs, on each day of testing at least three reference stainless steel inoculation
control coupons were inoculated simultaneously and interspersed with test coupons. The reference
coupons were inoculated as the first, middle, and last coupons within a single group of coupons, all
inoculated by the same MDI within a single test. These inoculation control (reference) coupons were a
stainless steel coupon (35.6 cm by 35.6 cm) inoculated in accordance with MOP 3161-LD or -HD,
sampled and analyzed in accordance with Section 2.3.2 or Section 2.3.3. If the results from the
inoculation controls were outside the acceptance criteria (Table 4-2), the results were discussed
immediately to determine the corrective action.
A log was maintained for each set of coupons that was dosed via the method of MOP 3161-LD or -HD.
Each record contained the unique coupon identifier, the MDI unique identifier, the date, the operator, the
weight of the MDI before dissemination into the coupon dosing device, the weight of the MDI after
dissemination, and the difference between these two weights. The coupon codes were pre-printed on the
log sheet prior to the start of coupon inoculation (dosing).
A second method was used to inoculate HVAC filters under air flow (~1000 cubic feet per minute, CFM).
To provide the flow, the back of the filter was connected to a blower from a high-volume air sampler
(Thermo Scientific High Volume Air Sampler VFC-PM10.for FRM RFPS-1287-063
(https://www.thermo.com/eThermo/CMA/PDFs/Various/File_52267.pdf)). The outlet of the blower was
directed to the intake of the H130 enclosure exhaust system to control contamination from breakthrough.
On top of the filter was an ADA assembled exactly as described in MOP 3161-LD or -HD. Positioned
between the HVAC filter and the blower, a sterile quartz filter was used to collect any spores that escaped
capture by the HVAC filter (see Figure 2-4). The quartz filter was installed as outlined in MOP 3168. The
blower was operated for 15 sec, then the MDI was activated, and the blower operated for another 15 sec
before deactivation. Once the blower ceased, the filter and ADA were lifted from the blower assembly and
placed on carpet tape as the filters inoculated using the first method. The quartz filter was collected as
outlined in MOP 3168. The blower adapter was cleaned and decontaminated between coupons with
Dispatch™ disinfectant wipes (Clorox Corp, CA), followed by 3% sodium thiosulfate (STS) and ethanol
wipes. This method is described in more detail in MOP 3161-F.
7
35.6 cm x 35.6 cm
ADA
ADAdapter
35.6 cm x 35.6
cm HVAC filter
Hi-Vol Blower
assembly
20.3 cm x 25.4 cm
Quartz filter
Flow
Figure 2-4.
Schematic of Deposition Apparatus for Inoculation under Flow Conditions
Additionally, after a coupon was dosed via the above procedure, the coupon was labeled with the unique
identifier described in Section 2.5.5. The identification (ID) was written on the ADA with a Sharpie or
equivalent permanent marker.
8
2.2
Test Matrix
Testing was conducted in three phases. Data from the first phase of testing were used to compare
recoveries from the various vacuum devices. The second phase was designed to perform side-by-side
comparisons of the two devices with the highest recoveries (from Phase 1) when used to collect spores from
HVAC filters. The third phase was designed to better understand the variability between different
laboratories in performing the inoculation, recovery, and extraction activities. The Phase 1 test matrix is
shown in Table 2-1.
Table 2-1. Phase 1Test Matrix
Replicate Test
Samples (Total
number of
coupons)
Negative Controls
Samples
Total
Coupons
Vacuum Device
Material
Target
Inoculum
(CFU/coupon)
Method 1
(Vacuum sock –
fast sampling)
Carpet
10
7
10 (30 coupons)
1 (3 coupons)
33
10
7
10 (30 coupons)
1 (3 coupons)
33
Concrete
10
7
10 (30 coupons)
1 (3 coupons)
33
Carpet
10
7
10 (30 coupons)
1 (3 coupons)
33
10
7
10 (30 coupons)
1 (3 coupons)
33
Concrete
10
7
10 (30 coupons)
1 (3 coupons)
33
Carpet
107
10 coupons
1 coupon
11
10
7
10 coupons
1 coupon
11
Concrete
10
7
10 coupons
1 coupon
11
Carpet
107
10 (30 coupons)
1 (3 coupons)
33
Upholstery
107
10 (30 coupons)
1 (3 coupons)
33
7
10 (30 coupons)
1 (3 coupons)
33
Method 1
(Vacuum sock –
slow sampling)
Method 2
(37 mm MCE)
Method 3
(Trace Evidence
Filter)
Method 4
(37 mm PTFE)
Upholstery
Upholstery
Upholstery
Concrete
10
Carpet
107
10 coupons
1 coupon
11
10
7
10 coupons
1 coupon
11
10
7
10 coupons
1 coupon
11
Upholstery
Concrete
MCE = Mixed Cellulose Ester, PTFE = Polytetrafluoroethylene
Vacuum methods applicable to large areas were challenged with three coupons per sample (Methods 1
and 3), while methods requiring a longer sample time per area were challenged with only one coupon per
sample (Methods 2 and 4).
The Phase 2 test matrix is shown in Table 2-2.
9
Table 2-2.
Phase 2 Test Matrix
Test Series
Sampling
Methods
Surface Type
Deposition
Method(s)
Purpose
HI1
Clean electrostatic
HVAC filters
Extraction
Settling vs. under flow,
8
10 spores/actuation
MDI used for
inoculation
HI2
Clean electrostatic
HVAC filters
Vacuum sock, 37
mm MCE, and 37
mm PTFE vs.
extraction
Under flow, 10
spores/actuation MDI
used for inoculation
HS
Clean electrostatic
or mechanical
HVAC filters
Vacuum sock, 37
mm MCE, and 37
mm PTFE vs.
extraction
Under flow, 10
spores/actuation MDI
used for inoculation
Measure the recovery
based on extraction
and test the validity of
the inoculation
procedure
8
Compare recovery of
vacuum methods to
extraction method
8
Compare recovery of
vacuum methods to
extraction method
One test from the HS series was performed twice (Test HS1 and Test HS1b). Test HS1b included PRB wipe
sampling of stainless steel coupons in addition to sponge-wipe sampling.
The Phase 3 test matrix is shown in Table 2-3.
Table 2-3. Phase 3 Test Matrix
Test Series
(Test
Name)
Target
Inoculum
Level
(CFU/coupon)
Inoculation
10
(INOC)
106
3
103
Stainless
steel
Stainless
steel
Stainless
steel
Carpet
106
Carpet
7
Carpet
107
Carpet
107
Carpet
107
Linearity
Recovery
Material
Type
(LR)
10
Laboratory
Variability
(LV)
Sample
Method
Sample Type
Reps
Sponge-wipe
Experimental
10
Sponge-wipe
Experimental
10
Sponge-wipe
Experimental
10
Vacuum sock
and 37 mm
Vacuum sock
and 37 mm
Vacuum sock
and 37 mm
Vacuum sock
and 37 mm
Vacuum sock
and 37 mm
MCE
Experimental
5
Experimental
5
Experimental
5
Experimental
to Laboratory
Experimental
to Laboratory
B
10
each
10
each
10
Purpose
To determine the
repeatability and
magnitude of
inoculation from
the MDI/ADA
dosing method
To determine
linearity of
recovery as a
function of
inoculation dose
To determine
variability
between
laboratory
technicians
(laboratories)
Each sample analysis day required laboratory blank samples analyzed in parallel with test samples.
2.3
Sampling and Analytical Procedures
Within a single test, surface sampling was first completed for all negative control coupons before
sampling of any inoculated coupon was performed. Surface sampling was conducted either by PRB wipe
sampling, sponge-wipe sampling, or vacuum sampling in accordance with the protocols documented
below. These methods encompass those currently used by the EPA and CDC during biological
contamination sampling events.
Laboratory surfaces were covered with new bench liner each day immediately prior to testing.
Paper sampling templates were sterilized with ethylene oxide (EtO) before each use.
Prior to the sampling event, all materials needed for sampling were prepared using aseptic techniques.
The materials specific to each protocol are indicated in the relevant sections below. In addition, general
sampling supplies were also needed. A sampling material bin was stocked for each sampling event. The
bin contained enough wipe sampling and vacuum sampling kits to accommodate all required samples for
the specific test. Additional kits of each type were also included for backup. A sample collection bin was
used to transport samples to the NHSRC Microbiology Laboratory after collection. The exterior of the
transport container was decontaminated by wiping all surfaces with a bleach wipe or towelette moistened
with a 5000 ppm hypochlorite solution prior to transport from the sampling location to the NHSRC
Microbiology Laboratory.
2.3.1 Swab Sampling
Swab sampling was used to verify sterility of coupons prior to inoculation. One coupon per sterilization
2
batch, or one coupon per 10 coupons in large sterilization batches, was swab-sampled (approx. 25 cm )
according to MOP 3135 and analyzed according to MOP 6563.
2.3.2 Polyester-Rayon Blend (PRB) Wipe Sampling
The centermost 30.5 cm x 30.5 cm area of a coupon was delineated by a sterile paper template and
sampled with a pre-moistened PRB wipe. Wipe sampling is typically used for small sample areas (i.e., 1
2
ft ) and is effective on nonporous, smooth surfaces such as ceramics, vinyl, metals, painted surfaces, and
plastics [7]. Wipe sampling was used for stainless steel control samples only and conducted according to
MOP 3144. The general approach is that a moistened sterile nonwoven PRB pad is used to wipe a
specified area to recover bacteria, viruses, and biological toxins. The protocol that was used in this
project is described in MOP 3144 and has been adapted from that provided by Busher et al. [7], Brown et
al. [5], and documented in the Idaho National Laboratory (INL) 2008 Evaluation Protocols . Wipe samples
®
were extracted in 20 mL Phosphate Buffered Saline (PBS) with 0.05% TWEEN 20 (PBST) according to
MOP 6567 and subjected to serial 10-fold dilution and spread-plating onto Tryptic Soy Agar (TSA, BD,
Franklin Lakes, NJ) according to MOP 6535a.
11
2.3.3 Sponge-wipe Sampling
The centermost 30.5 cm x 30.5 cm area of the coupon was delineated by a sterile paper template and
sampled with a pre-moistened sponge. Sponge-wipe samples, described in MOP 3165, were collected
using the following five patterns: (1) using the flat side of the sponge-wipe, the surface was sampled
using horizontal S-strokes, covering the entire template area; (2) the sponge-wipe was then flipped over
to the opposite side to sample the surface in a vertical pattern, covering the entire template area; (3)
using the narrow edges of the sponge-wipe, the surface was sampled using the same S-strokes but
applied diagonally across the template, (4) rotating the sponge to use the opposite side starting at the
midway point of the coupon; and (5) the tip of the sponge-wipe was then used to sample the perimeter of
the sampling area. The sampling method is described in detail in the study Rose et al. [8] and are
consistent with the CDC-developed field collection procedures (www.cdc.gov/niosh/topics/emres/surfacesampling-bacillus-anthracis.html). Sponge-wipe samples were extracted in 90 mL PBST as described in
MOP 6580 and subjected to 10-fold serial dilution and spread-plating according to MOP 6535a.
2.3.4 Vacuum Sock Sampling and Analysis
Vacuum sock sampling was conducted in accordance with MOP 3145 with the following modification:
three coupons were vacuumed per sample for some Phase 1 tests. A single coupon was vacuumed for
Phase 2 and 3 tests. The centermost 30.5 cm x 30.5 cm (12” x 12”) square area was vacuumed on each
coupon. For Phase 1, vacuum sock nozzles were moved across the coupon at two speeds, approximately
1 sec per 30.5 cm pass (fast) and approximately 3 sec per 30.5 cm pass (slow). For example, on one
square foot of surface area (30.5 cm x 30.5 cm), sampling with the vacuum sock (slow) or vacuum sock
(fast) required approximately 60 passes each (30 passes in one direction, then 30 additional passes in a
direction oriented 90 degrees to the first), and approximately 240 or 90 sec, respectively. For the three
coupon sample, the collection time required was 720 or 270 sec for the slow or fast method, respectively.
Each speed was considered a different vacuum method. All vacuum samples were collected using an
-1
OmegaVac (Atrix, Int.; Burnsville, MN), which supplied airflow of approximately 2000 L min . This
equipment was powered by alternating current (120 V) supplied by a wall receptacle. Phases 2 and 3
used only the slow speed technique. Vacuum socks were extracted in 20 mL PBST according to MOP
6572 and subjected to 10-fold serial dilution and spread-plating according to MOP 6535a. Figure 2-5
shows an assembled vacuum sock kit with cardboard nozzle and connection tubes, and the vacuum sock
removed from the tubes.
12
Figure 2-5.
Vacuum Sock Kit and Individual Sock.
2.3.5 37 mm MCE and 37 mm PTFE Sampling
Filter samples (37 mm MCE and PTFE) were collected according to Section A of MOP 3164, based on
methods developed jointly by EPA and CDC. In short, a vacuum pump (Vac-U-Go, SKC, Inc., Eighty
-1
Four, PA) at the back end of the filter pulled 20 L min of air through the filter. This pump was powered by
120 V alternating current supplied by a wall receptacle. A section of Tygon (~3 cm ) tubing was cut to an
angle of 45° on one end, the non-angled terminus was attached to the cassette via a polyvinyl chloride
(PVC) adapter (SKC, Inc., P/N 225-132A), and the angled end was used as a nozzle. The nozzle and
-1
filter were moved along the coupon at roughly ten cm sec (three seconds per 30.5 cm pass) in both
directions (i.e., horizontally and vertically). For example, on one square foot of surface area (30.5 cm x
30.5 cm), sampling with the 37 mm cassette (MCE or PTFE) required approximately 100 total passes and
approximately 400 sec. A single coupon was vacuumed when sampling with either of these two devices.
The nozzle was extracted separately (described in MOP 6579), the nozzle extract was then combined
with the filter extraction vessel, and filter extraction commenced. The combined resulting extract was
subjected to 10-fold serial dilution and spread-plating according to MOP 6535a.
Figure 2-6 shows the 37 mm cassette with nozzle and tubing.
13
Figure 2-6.
Filter (37 mm) Cassette with Nozzle and Tubing
2.3.6 Trace Evidence Filter Sampling
Trace Evidence Filter (TEF) samples were collected according to Section B of MOP 3164, based on
methods developed jointly by EPA and CDC. In short, a vacuum (Atrix OmegaVac, operating at 50%
voltage to prevent overcoming filter housing) pulled 790 L min-1 (28 standard cubic feet per minute
(SCFM)) of air through the filter. The nozzle and filter were moved along the coupon at roughly ten cm
-1
sec (three seconds per 30.5 cm pass) in both directions (i.e., horizontally and vertically). On one square
foot of surface area (30.5 cm x 30.5 cm), sampling with the vacuum TEF required approximately 30 total
passes and approximately 120 sec for sample collection. Three coupons per sample were vacuumed
when sampling with this device. TEF filters were extracted in 90 mL PBST as described in MOP 6582 and
subjected to 10-fold serial dilution and spread-plating according to MOP 6535a. Figure 2-7 shows the
capped TEF cassette.
2.3.7 Operational Assessment
Operational data, such as time required for sample collection, ease of sample collection, ease of device
use, malfunction of device, and ease of laboratory analytical procedures, were collected for each
sampling method. These data were used to qualitatively compare methods based on their ease of use.
14
Figure 2-7.
3M ® Trace Evidence Filter (TEF) Vacuum-based Sampling Device
2.3.8 HVAC filter extraction
2
2
Sections (930 cm , 1 ft ) of HVAC filters were excised and further cut into half or quarter sections (15.3
cm x 30.5 cm (6” x 12”) or 15.3 cm x 15.3 cm (6” x 6”)) and using sterile scissors, folded, and placed in a
sterile 1 L container (one section per container). The size chosen for filter sub-sections was based upon
optimization experiments conducted with the same 1 L containers at CDC (data not shown). Excision and
sectioning of the filter typically required 90 sec (each 30.5 cm x 30.5 cm (12” x 12”) section) once cutting
was initiated. HVAC filters were then extracted in PBST for 30 min using an orbital shaker (300 rpm).
Quarter-sections of the filters were extracted in 500 mL PBST, while half-sections of the filters were
extracted in 700 mL PBST. Details on the method are provided in MOP 6593.
2.3.9 Quartz extraction
Quartz filters were extracted in 100 mL of PBST as described in MOP 6586.
2.3.10 Direct Extraction of Stainless Steel
Six 15.3 cm x 30.5 cm (4” x 6”) stainless steel coupon parts were aseptically transferred straight from the
ADA to a sterile 10L beaker. Two sterile glass rods, bent symmetrically at a 90 degree angle, were placed
in the bottom of the beaker, followed by one sextant of the inoculated coupon, followed by another two
sterile glass rods. The process of stacking the glass rods and coupons continued until there were no
more inoculated coupon pieces. Each coupon was placed in the beaker so the inoculated side was
upright. The glass beaker was sealed (sterile aluminum foil; the same foil with which the beaker was
autoclaved) and transported to the NHSRC Microbiology Laboratory for extraction.
15
Sterile PBST (1.5 liters) was aseptically added to the 10L beaker containing the sample and then resealed with the aluminum foil. The entire beaker was placed into an ultrasonic cleaner (Branson model
8510, Danbury, CT) and the sample was sonicated (40 kHz) for 15 min. Immediately following sonication,
1L of extraction liquid was removed and transferred to a 1L specimen container. The sample was then
homogenized by manual agitation/swirling before being 10-fold serially diluted and plated according to
MOP 6535a.
2.4
Sampling Strategy
2.4.1 Sampling/Monitoring Points
The experimental samples are listed in Section 2.2. For each inoculation event, additional samples
collected from stainless steel surfaces were used as control samples. These control samples included
wipe samples (used as inoculation controls or reference coupons) and vacuum samples (to compare
collection efficacy among vacuum sampling methods). Wipe and vacuum samples were collected by
sampling within a 30.5 cm x 30.5cm (12” x 12”) sampling template (SKC, Inc., P/N 225-2416) centered on
the coupons. Direct extraction techniques were also used to quantify inoculation levels on stainless steel
and HVAC filters. Each coupon was sampled only once.
The time required to collect and analyze vacuum samples, both singly and in the aggregate, was logged
in the laboratory notebook.
Table 2-4 lists the samples collected for each test.
16
Table 2-4. Frequency of Sampling Monitoring Events
Sample Type
Sample Number
Purpose
Vacuum sample
3 to 10 per test condition per
coupon material
To determine the number of viable
spores recovered via the vacuum method
Negative coupon
sample (vacuum
sample of sterile
coupon)
1 per vacuum method per test
To determine extent of crosscontamination from coupon handling and
sampling
Field blank sample (1
minute sample of
laboratory air)
1 per vacuum method per test
To determine extent of crosscontamination from laboratory air
Positive control
coupons sampled
either by extraction,
wipe, or sponge
1 set of 3 stainless steel coupons
per sampling method inoculated
at the beginning, middle, and end
of test coupon inoculations
To provide the best estimate of the
number of viable spores deposited onto
the material test coupons
Swab samples
1 per material or equipment per
sterilization batch
To demonstrate sterility of coupons and
inoculation materials
Microbiology
Laboratory material
blanks
3 per material
To demonstrate sterility of extraction and
plating materials
Table 2-5 lists the critical and noncritical measurements for each sample.
Table 2-5. Critical and Non-Critical Measurements
Sample Type
Critical Measurements
Non-critical Measurement
Vacuum sample
Plated volume, incubation
temperature, extracted volume, CFU,
collection area
Storage time, storage temperature,
collection time, extraction time
Negative coupon sample
(vacuum sample of sterile
coupon)
Plated volume, incubation
temperature, extracted volume, CFU,
collection area
Storage time, storage temperature,
collection time, extraction time
Field blank sample (1 minute
sample of laboratory air)
Plated volume, incubation
temperature, extracted volume, CFU
Storage time, storage temperature,
collection time, extraction time
Positive control coupons
sampled either by extraction,
wipe, or sponge
Plated volume, incubation
temperature, extracted volume, CFU
Storage time, storage temperature,
collection time, extraction time
Swab samples
Plated volume, incubation
temperature, extracted volume, CFU
Storage time, storage temperature
Microbiology Laboratory
material blanks
Plated volume, incubation
temperature, extracted volume, CFU
Storage time, storage temperature
17
2.5
Sampling Handling and Custody
2.5.1 Preventing Cross-contamination during Coupon Preparation
Coupon preparation included the activities performed on each pre-fabricated material coupon and
procedural blank coupon prior to the inoculation procedure. Sterilization methods depended on the
sample type: concrete coupons were sterilized using the gravity cycle of the autoclave; carpet and
upholstery coupons were sterilized using VHP. Swab sampling of coupons from sterilization batches
were used to confirm sterility of the materials after sterilization. The sterilization procedure was repeated
if results were positive for the target organism. Swabs showing foreign contamination could be cause for
repeating the sterilization procedure or taking other corrective action. The blank coupon sampling
occurred before sampling of any inoculated coupons.
2.5.2 Preventing Cross-contamination during Sampling
Sampling poses a significant opportunity for cross-contamination of samples. In an effort to minimize the
potential for cross-contamination, several management controls were followed.
•
In accordance with aseptic technique, a sampling team made up of a “sampler,” a “support
person,” and a “sample handler” was utilized.
•
The sample handler was the only person to handle deposition pyramids (ADAs) or material
coupons during the sampling event. The support person had the responsibility of handing sterile
templates to the sampler.
•
The sampler handled only the sampling media and the support person handled all other supplies.
The sampler sampled the surface according to the appropriate procedure as described in Section
2.3.
•
The collection medium was then placed into a sample container that was opened, held and
closed by the support person.
•
The sealed sample was handled only by the support person.
•
All of the following actions were performed only by the support person, using aseptic technique:
•
o
The sealed bag with the sample was placed into another sterile plastic bag that was then
sealed; that bag was then decontaminated using a bleach wipe.
o
The double-bagged sample was then placed into a third sterile bag that was sealed and
then placed into a sterile sample container for transport.
o
The exterior of the transport container was decontaminated by wiping all surfaces with a
bleach wipe or towelette moistened with a solution of 5000 ppm hypochlorite prior to
transport from the sampling location to the NHSRC Microbiology Laboratory.
After the sample was placed into the container for transport, the sample handling team placed the
sampled coupon in soapy water for eventual disposal.
The sampling crew then changed their gloves in preparation for working with the next sample.
Additionally, and equally important, the order of sampling was as follows: (1) first field blank; (2) all blank
coupons; (3) second field blank (when required by test plan); (4) inoculated coupons; and (5) last field
18
blank (when required by test plan). This order ensured that test coupons were handled in an order from
least level of contamination to the most, and field blank control samples provided evidence that samples
were handled properly, without cross-contamination.
2.5.3 Preventing Cross-contamination during Analysis
General aseptic laboratory technique was followed and was embedded in the standard operating
procedures (SOPs) and MOPs used by the NHSRC Microbiology Laboratory to recover and plate
samples. The SOPs and MOPs document the aseptic technique employed to prevent crosscontamination. Additionally, the order of analysis was (1) all blank coupons, then (2) all inoculated
coupons.
2.5.4 Sample Containers
For each PRB wipe sample, the primary containment was an individual sterile 50 mL conical tube.
Conical tubes (15mL) were primary containment for swab samples. Secondary containment for swab and
PRB wipe samples was sterile sampling bags. The sponge-wipe primary containment was the stomacher
bag used for extraction, and secondary containment was sterile sampling bags. The primary containment
of the vacuum sock was a sterile sampling bag. A four inch cable tie was also used to close the open end
of the newly-collected vacuum sock sample. The secondary containment of each vacuum method
sample was separate sterile sampling bags. All biological samples from a single test were then placed in
a sterilized container. After samples were placed in the container for storage and transport to the NHSRC
Microbiology Laboratory, the container was wiped with a towelette saturated with a ≥5000 ppm
hypochlorite solution. A single container was used for storage of materials during sampling and for
transport of samples to the NHSRC Microbiology Laboratory.
2.5.5 Sample Identification
Each coupon or sample was identified by a unique sample ID. The sampling team maintained an explicit
laboratory log which included records of each unique sample ID and its associated test number, inoculum
level, sampling method, and the date sampled. Each coupon was marked with only the material
descriptor and unique code number. Sample IDs included descriptors, where necessary, for project
number (WA 10), test ID, coupon material type, vacuum or other sample type, inoculation type, sample
purpose (test, control, field blank, etc.) and replicate number. The sample codes eased written
identification. A typical sample ID was 10-HS1-F-V-4, which identified the sample as from WA 10, Test
HS1, inoculated under Flow, sampled by Vacuum sock, replicate 4. Once samples were transferred to the
NHSRC Microbiology Laboratory for microbiological analysis, each sample (plate) was additionally
identified by replicate number and dilution. The NHSRC Microbiology Laboratory also included on each
plate the date it was placed in the incubator.
The samples from blank coupons had a two-letter material code, with the first letter being “X”.
The sequence number was added to the test number to distinguish control samples in the case where all
materials were not inoculated at the same time for a single vacuum method. For instance, a stainless
steel PRB wipe sample control coupon for Test 1 for carpet and upholstery may be labeled 10-1.1-S-W-1,
while for the concrete, inoculated on a different day, the sample would be labeled 10-1.2-S-W-1.
19
2.5.6 Information Recorded by Field Personnel
The sampling team members’ names, date, run number, and all sample codes with corresponding
coupon codes were recorded in the laboratory notebook, along with sample times and durations. Any
deviations from sampling protocols were documented in the laboratory notebook, along with any
observations.
Digital video was collected during sampling of each material using each vacuum method in order to
document the process.
2.5.7 Sample Preservation
Following transfer to the NHSRC Microbiology Laboratory, all samples were stored at 4 ± 2 °C until
analyzed. All samples were allowed to equilibrate at room temperature for one hour prior to analysis.
2.5.8 Sample Holding Times
After sample collection for a single test was complete, all biological samples were transported to the
NHSRC Microbiology Laboratory immediately, with appropriate chain of custody (COC) form(s). Liquid
samples were stored no longer than 24 h prior to analysis. Samples of other matrices were stored no
longer than five days before the primary analysis. Typical hold times, prior to analyses, for most
biological samples was ≤ two days.
2.5.9 Sample Custody
Careful coordination with the NHSRC Microbiology Laboratory was required to achieve successful
transfer of uncompromised samples in a timely manner for analysis. Test schedules were confirmed with
the Microbiology Laboratory prior to the start of each test. To ensure the integrity of samples and to
maintain a timely and traceable transfer of samples, an established and proven chain of custody or
possession is mandatory. Accurate records were maintained whenever samples were created,
transferred, stored, analyzed, or destroyed. The primary objective of these procedures was to create an
accurate written record that could be used to trace the possession of the sample from the moment of its
creation through the reporting of the results. A sample was in custody in any one of the following states:
•
•
•
•
•
In actual physical possession
In view, after being in physical possession
In physical possession and locked up so that no one could tamper with it
In a secured area, restricted except to authorized personnel
In transit.
Laboratory test team members received copies of the test plans prior to each test. Pre-study briefings
were held to apprise all participants of the objectives, test protocols, and COC procedures to be followed.
These protocols were required to be consistent with any protocols established by EPA. In the transfer of
custody, each custodian signed, recorded, and dated the transfer on the COC. Sample transfer could be
on a sample-by-sample basis or on a bulk basis. The following protocol was followed for all samples as
they were collected and prepared for distribution:
•
A COC record accompanied the samples. When turning over possession of samples, the transferor
and recipient signed, dated, and noted the time on the record sheet. This record sheet allowed
20
transfer of custody of a group of samples from the sample collection laboratory to the NHSRC
Microbiology Laboratory.
•
If the custodian had not been assigned, the laboratory operator had the responsibility of packaging
the samples for transport. Samples were carefully packed and hand-carried between on-site
laboratories. The COC record showing the identity of the contents accompanied all packages.
2.5.10 Sample Archiving
All samples and diluted samples were archived for a minimum of two weeks following completion of
analysis. This time allowed for review of the data to determine if any re-plating of selected samples was
required. Samples were archived by maintaining the primary extract at 4 ± 2 °C in a sealed extraction
vessel.
2.6
Statistical Analysis Methods
Relative recovery data were analyzed using a one-way analysis of variance (ANOVA). A three-way
ANOVA model with full interactions between Device, Material, and Experiment/Control was used to
analyze raw recovery data. For the three-way ANOVA, mean log10 reduction was then computed as a
linear contrast of model coefficients to assess recovery rate. Mean differences in log10 reduction between
devices were also computed as linear contrasts. Statistical significance (p ≤ 0.05) of differences between
devices was then assessed via t-tests on the contrasts, and two sets of p-values were computed. First, pvalues were computed without adjusting for multiple comparisons. Subsequently, p-values which
accounted for multiple comparisons were computed, based on the multivariate t-distribution of the test
statistics.
Both statistical methods were deemed valid by a statistical contractor (Neptune, Inc.). The advantage of
the one-way ANOVA is that it compared relative recovery data that were normalized across experiments
(normalized by PRB wipe recoveries). The disadvantage is that one-way ANOVA is not robust with
regards to multiple sources of variation (i.e., variation in treatment groups and variation in control
recoveries). The three-way ANOVA compared recoveries of the sampling methods using raw recovery
values, and included recovery data from PRB wipe samples as a treatment group. While this method
accounted for multiple sources of variation and interaction, the method, as used, was less robust for
direct comparisons of sampling methods across tests.
The results of both statistical approaches are presented and discussed.
21
3
3.1
Results and Discussion
Phase 1
The results of the vacuum method scoping tests comparing recovery of five techniques from three
materials are summarized in Table 3-1 and Figure 3-1.
Table 3-1. Recovery from Materials for each Vacuum Method
Mean CFU/cm2 (n = 10) (Standard Deviation in parentheses)
Vacuum Sock
– Fast Speed
Vacuum Sock
– Slow Speed
Carpet
2.01 x 104
3
(9.33 x 10 )
2.66 x 104
3
(8.49 x 10 )
2.83 x 104
4
(1.19 x 10 )
1.28 x 104
3
(2.21 x 10 )
2.18 x 104
4
(1.05 x 10 )
Concrete
1.44 x 104
3
(3.55 x 10 )
3.18 x 104
3
(5.55 x 10 )
7.41 x 104
4
(3.88 x 10 )
3.47 x 104
4
(1.06 x 10 )
2.57 x 104
4
(1.49 x 10 )
Upholstery
1.86 x 104
3
(4.16 x 10 )
7.29 x 103
3
(2.73 x 10 )
2.08 x 104
3
(5.64 x 10 )
3.76 x 103
3
(3.78 x 10 )
1.39 x 104
3
(8.60 x 10 )
Material
37 mm MCE
TEF
37 mm PTFE
Recovery (CFU cm-2)
1.00E+06
1.00E+05
vacuum sock - fast
vacuum sock - slow
37 mm MCE
TEF
1.00E+04
37 mm PTFE
1.00E+03
Carpet
Figure 3-1.
Concrete
Upholstery
Recovery from Materials for each Vacuum Method. Data are plotted on a log-scale, as
mean ± standard deviation.
These data are standardized for number of coupons sampled, with three coupons per sample for all
methods except the 37 mm MCE and PTFE methods, which used only one coupon. Even without this
standardization, the 37 mm MCE vacuum method shows higher recoveries from concrete (i.e., more
22
spores recovered by this method even though surface area sampled was one-third that of the vacuum
sock and TEF methods).
To account for differing inoculation levels achieved across numerous test days and MDIs, recovery was
further standardized by normalizing vacuum recoveries to PRB wipe samples collected from stainless
steel coupons inoculated by the same MDI and collected on the same test day as the test samples.
Figure 3-2 and Table 3-2 summarize these results. Interestingly, the 37 mm MCE method demonstrated
higher recovery on concrete than the wipe-based method on stainless steel, resulting in a relative
recovery ≥100%. One explanation is that the 37 mm MCE device efficiently collected fine dust or debris
particles from the concrete surface. Spores bound to these debris particles may have been collected
more efficiently as a consequence, because the ease of particle resuspension from surfaces increases
proportionately with cross-sectional area (i.e., larger particles are more easily resuspended than smaller
particles).
140%
Relative Recovery (%)
120%
100%
Vacuum sock - fast
80%
Vacuum sock - slow
37 mm MCE
60%
TEF
40%
37 mm PTFE
20%
0%
Concrete
Upholstery
Carpet
Relative Recovery – Data presented as mean relative recovery (RR). RR calculated as
percent of wipe recovery from stainless steel surface.
Figure 3-2.
Relative Recoveries from all Devices and Material Surface Types
Table 3-2.
Average CFU/cm2 vacuum method / Average CFU/cm2 stainless steel wipe
Vacuum Sock
– Fast Speed
Vacuum Sock
– Slow Speed
37 mm MCE
TEF
37 mm PTFE
Concrete
30%
26%
124%
41%
24%
Upholstery
23%
11%
35%
3%
13%
Carpet
39%
64%
47%
12%
20%
23
The null hypothesis that recoveries obtained during slow operation of the vacuum sock nozzle were not
significantly different (p > 0.05) from the recoveries obtained during fast operation was not disproven by
these tests. Recovery using the slow vacuum sock method was higher from carpet coupons but lower
(than the fast vacuum sock method) from upholstery and concrete coupons, possibly due to the rapid
motion of the nozzle agitating the coupon surface. Operation of the vacuum sock nozzle is expected to be
highly variable between individual operators. Based on these data, operational variability may not greatly
affect recoveries.
Preliminary research (based upon direct spiking, conducted at the CDC) suggested that recovery from
PTFE was higher than recovery from MCE for the 37 mm filter media. The current data suggest that
overall recovery (collection from the surface, retention on the filter, then extraction from the filter) with
MCE filters was higher. The TEF vacuum method performed poorly for upholstery and carpet coupons,
compared to the other methods. The sock (slow) method performed poorly on upholstery.
3.1.1 Results of Statistical Analyses
For carpet, the slow vacuum sock method has the highest recovery rate (one-way ANOVA, p ≤ 0.001).
When analyzed by three-way ANOVA without correcting for multiple comparisons, the vacuum sock had
significantly higher recoveries than the 37mm PTFE (p = 0.004) and TEF (p = 0.003) methods. On
carpet, none of the method comparisons were statistically significant after correcting for multiple
comparisons (three-way ANOVA, p > 0.05).
For concrete, the 37mm MCE method demonstrated the highest recovery (one-way ANOVA, p ≤ 0.001).
Contrasts of the methods were also significant when analyzed by three-way ANOVA, before adjusting for
multiple comparisons (all p ≤ 0.02). After adjusting for multiple comparisons, the 37mm MCE recoveries
were statistically significant only when contrasted with recoveries of the 37 mm PTFE method (p = 0.03).
For upholstery, the 37 mm MCE method demonstrated the highest recovery (one-way ANOVA, p ≤
0.001). When analyzed by three-way ANOVA, the recoveries using the 37 mm MCE method were
significantly higher than those of the 37 mm PTFE, TEF, and slow vacuum sock methods. After adjusting
for multiple comparisons, was the 37 mm MCE method recoveries were only statistically higher than
those using the TEF method. The TEF method demonstrated the lowest recovery for upholstery
compared to all other devices. This result was statistically significant even after adjusting for multiple
comparisons (three-way ANOVA, all p ≤ 0.001).
3.1.2 Operational Parameters
3.1.2.1
Sampling Time and Ease
As discussed above, two sampling durations were used for vacuum sock sampling. Based on those data,
there was conflicting effects of sampling time on vacuum sock samples.
For the 37 mm devices, sampling is very time consuming due to the small surface area of the vacuum
nozzle. A sample time of five min ft-2 limits the number of samples that can be collected per person per
day, especially within the confines of Hazardous Materials (HAZMAT) operations. The TEF method
sampling time was comparable to that of the vacuum sock.
24
The vacuum sock and 37 mm vacuums are both commercially-available items. The recommended
vacuum for the TEF filters caused the filters to warp within the housing, which caused loss of sample. For
these tests, the recommended vacuum had to be operated at 50% power to prevent filter collapse.
Defining the operation of the TEF vacuum was thus more complicated and may be more difficult to
implement in the case of a large response.
3.1.2.2
Analysis Time and Ease
Vacuum Socks
Sample kit preparation, as well as sampling and extraction procedures, is straightforward. In the current
study, the extraction was completed in 20 mL of PBST. Increasing the volume of PBST may be
beneficial. With the 20 mL, the entire white (sample) portion of the sock cannot be wetted during the filter
segmentation steps without allowing the nonsterile blue portion of the sock to touch the inside of the
extraction cup. Durable, ergonomic scissors that are resilient to repeated sterilizations are strongly
recommended. Custom-made racks for the orbital shaker were needed to accommodate the extraction
vessels. Vacuum sock samples required many plating iterations (each iteration was conducted in
triplicate) to meet QA goals for variability. This variability may be due to the presence of debris in the
vacuum sock extraction fluid. Average processing time (all unpackaging and extraction procedures, not
plating and analysis) for a typical sample batch (12 sock samples) was 90 min, or 7.5 min per sample.
Filter (37 mm)
Assembly of the 37 mm sampling kits and extracting the samples was time-consuming. Transferring liquid
from the filter cartridge can be difficult, especially when large amounts of debris are present. The nozzle
was extracted separately and usually contained large fragments of carpet or other debris. The debris from
the concrete, when mixed with the PBST, created a paste-like substance that was difficult to transfer via
pipet. The carpet debris also was difficult to pipet, as it frequently obstructed pipet tips. Average
processing time (all unpackaging and extraction procedures, not plating and analysis) for a typical sample
batch (12 sock samples) was 120 min, or 10 min per sample.
TEF
Aseptically removing the TEF filter from the filter housing proved difficult as it was impossible to predict
which half of the two-part housing the filter would align itself with upon opening. Rinsing the filter and
cartridge was also time consuming and difficult, especially with samples collected from concrete. The
dust and debris from the concrete samples generated a paste during sample extraction and was difficult
to manipulate. Occasionally the filters punctured the stomacher bag, resulting in a compromised sample.
Sample throughput was limited by the stomacher. Average processing time (all unpackaging and
extraction procedures, not plating and analysis) for a typical sample batch (10 sock samples) was 180
min, or 18 min per sample.
3.1.2.3
Cost
While the cost of all three methods was tracked, it is difficult to determine if any method was significantly
more expensive than any another. The capital costs of each method would vary as well from laboratory to
laboratory, depending on the availability of equipment. The following extraction equipment is
recommended for each laboratory handling 50 samples daily:
Vacuum Sock method: Orbital shaker with appropriate rack for extraction vessels;
37 mm filter samples: Ultrasonic water bath (sonicator);
25
TEF filter samples: Stomacher (preferably two), bench-top centrifuge.
3.2
Phase 2
The results from Test HI1 are summarized in Table 3-3 and suggest that MOP 3161-F, the proposed
method for inoculating HVAC filters under flow, worked as expected and yielded repeatable results (the
6
data quality objective was to achieve 1.0 x 10 recovered spores (CFU) and less than 100% RSD). During
8
inoculation, an average 1.1 x 10 spores (CFU) were captured on the quartz filter having passed
completely through the HVAC MERV 8 filter. However, the amounts recovered from HVAC filters
inoculated under flow were similar to the HVAC filters inoculated with settling conditions only. Because
similar recoveries were obtained from both methods and the flow-based method is likely to be more
representative of HVAC filter contamination in the field, MOP 3161-F (flow-based method) was used on
all subsequent HVAC filter inoculations.
Table 3-3. Average Recovery from HVAC Filter Extraction – Test HI1
Inoculation Method
Flow
Settling
Mean Recovery (CFU) from HVAC Filter (n=3)
RSD* (%)
3.96 x 106
54%
6
39%
9.47 x 10
*RSD – Relative Standard Deviation, i.e., Coefficient of Variation as a percentage
Test HI2 was a scoping test to evaluate the feasibility of sampling flow-inoculated filters using vacuum
methods. Table 3-4 and Figure 3-3 show the average recovery (CFU) from HVAC filters using four
vacuum methods, as well as direct extraction. The method for direct extraction can be used on the small,
thin filters chosen for this project but is not expected to be feasible for larger or thicker HVAC filters.
Table 3-4. Recovery from HVAC filters – Test HI2
Vacuum Method
37 mm PTFE
TEF
37 mm MCE
Vacuum sock (slow)
Extraction
Mean Recovery (CFU) (n=3)
RSD* (%)
1.83 x 10
8
15%
1.17 x 10
8
98%
5.41 x 10
7
33%
1.37 x 10
8
51%
1.63 x 10
8
21%
*RSD – Relative Standard Deviation, i.e., Coefficient of Variation as a percentage
26
Recovery from HVAC Filters
Recovery (CFU)
1.00E+09
1.00E+08
1.00E+07
1.00E+06
37 mm PTFE
Figure 3-3.
37 mm MCE
TEF
Vacuum sock
Extraction
Recovery from HVAC Filters Using 5 Sampling Methods – Test HI2. Data are plotted
on a log-scale, as mean ± standard deviation.
Results from Tests HS1 and HS2 are shown in Table 3-5. All tested methods performed well, with some
higher variability in the TEF recovery. Though in Phase 2 the 37 mm PTFE filters provided better recovery
than the 37 mm MCE filters (Figure 3-3), the MCE filters were chosen for side-by-side evaluation with
vacuum socks based on the results from Phase 1 (Table 3-2).
Table 3-5.
HVAC Vacuuming Recovery – Test HS
Mechanical MERV 8 Filter
Comparison
Mean
to Extraction
CFU/Sample RSD
(T-test p(n=5)
(%)
value)
Sample Type
4.90 x 106
Extraction
Vacuum sock
a
37 mm MCE
a
50%
6
43%
1.58 x 106
40%
1.34 x 10
0.029
0.036
Electrostatic MERV 8 filter
Comparison
Mean
to Extraction
CFU/Sample
RSD
(T-test p(n=5)
(%)
value)
4.60 x 106
84%
6
43%
3.36 x 105
40%
1.39 x 10
0.26
0.16
One electrostatic filter sample was lost during extraction. n = 4.
The single factor ANOVA p-value of 4.4 x 10-3 suggests a difference between all three recovery methods
from Minimum Efficiency Reporting Value (MERV) 8 mechanical filters and the heteroscedastic Student’s
t-test values between the vacuum methods and extraction recovery indicate neither vacuum method
provided recoveries as high as extraction from the mechanical MERV 8 filter. Figure 3-4 shows recovery
data from the mechanical MERV 8 filter with the three collection methods. Neither vacuum method
provided a statistically significant difference in recovery (heteroscedastic Student’s t-test, p = 0.56).
27
Recovery from Mechanical MERV 8 HVAC Filters
Recovery (CFU)
1.00E+07
1.00E+06
1.00E+05
1.00E+04
Vacuum sock
Figure 3-4.
HVAC Extraction
37 mm MCE
Recovery from Mechanical MERV 8 Filter – Test HS. Data are plotted on a log-scale, as
mean ± standard deviation.
Unlike the case for mechanical type filters, the single factor ANOVA suggests no statistical difference
between all three recovery methods from electrostatic MERV 8 filters (p = 0.16) (Figure 3-5). However,
reanalysis using log10-transformed recovery, thus stabilizing variability, indicates there is a statistically
significant difference between the three methods (ANOVA, p = 0.002). This difference is indicative in part
to a high uncertainty in the number of spores deposited. Table 3-6 shows the heteroscedastic Student’s ttest between method pairs for both raw recovery and log-transformed recovery.
Table 3-6. Student’s t-test values from Electrostatic filters
Heteroscedastic Student’s T-test p-values
Method Pairs
Recovery
Log10 Recovery
vacuum sock vs. 37 mm
0.036147
0.005965
vacuum sock vs. extraction
0.262986
0.187705
37 mm vs. extraction
0.157613
0.007319
Table 3-6 suggests a statistically significant difference between vacuum sock and 37 mm MCE vacuum
methods, indicating higher recovery from vacuum sock. Table 3-6 also suggests a potential difference
between 37 mm and extraction methods.
28
Recovery from Electrostatic MERV 8 Filters
1.00E+08
Recovery (CFU)
1.00E+07
1.00E+06
1.00E+05
1.00E+04
Vacuum sock
Figure 3-5.
HVAC Extraction
37 mm MCE
Recovery from Electrostatic HVAC Filter – Test HS. Data are plotted on a log scale, as
mean ± standard deviation.
The vacuum sock method was much easier to perform on HVAC filters than the operation of the 37 mm
filters. However, the ease of performance may be true only for certain types of HVAC filters. Filter types
with very deep crevices may require a sampling method such as the 37 mm filter method, which uses a
more narrow nozzle.
The Phase 2 test matrix also included comparisons of recovery methods from stainless steel deposition.
While these depositions were in part included for quality control − as a measure of the stability of the
inoculation MDI − they also provide information on spore recovery from nonporous surfaces. These data
are not directly comparable to the HVAC data, due to the spores that passed completely through the filter
as discussed above (Test HI1). Tables 3-7 and 3-8 summarize the recovery results from stainless steel
surface samples, collected and analyzed during Phase 2 tests. The data are segregated by test because
different MDIs were used for each. But taken collectively, the three recovery methods were equivalent,
with no method producing statistically significant higher recovery.
29
Table 3-7. Mean Recovery from Stainless Steel Coupons (n = 3), Phase 2 Tests
Log10 CFU (± Standard Deviation)
Sampling Method
HS2
HS1b
7
6
HS1
7
6
extraction
2.2 x 10 (6.61 x 10 )
1.75 x 10 (5.23 x 10 )
1.11 x 108 (2.68 x 107)
sponge-wipe
2.63 x 107 (8.8 x 106)
1.90 x 107 (6.61 x 106)
7.52 x 107 (2.10 x 107)
PRB wipes
1.93 x 107 (3.02 x 106)
2.05 x 107 (5.11 x 106)
NA
Note: Standard deviation is in parentheses.
Table 3-8. ANOVA – Comparison of the Three Stainless Steel Recovery Methods within each
Phase 2 Test Run (n = 3)
Source of
Variation
Test
a
df
F
P-value
F crit
HS2
Between Groupsa
2
0.87
0.46
5.14
HS1b
Between Groups
2
0.20
0.82
5.14
HS1
Between Groups
2
3.21
0.15
7.71
Groups: Direct extraction, sponge-wipe, and PRB wipe of stainless steel.
3.3
Phase 3
The results from the INOC test series are summarized in Table 3-9 and graphically displayed in Figure 36.
Table 3-9. Mean Recoveries from Stainless Steel using the Sponge-wipe Method - Test INOC (n =
10)
MDI (Target Dose)
Mean Recovery (CFU)
RSD
103
104
107
1.27 x 103
1.48 x 104
1.39 x 107
37%
31%
52%
30
Recovery from Stainless Coupons
Recovery (CFU)
1.00E+08
1.00E+07
1.00E+06
1.00E+05
E3
1.00E+04
E4
E7
1.00E+03
1.00E+02
1
2
3
4
5
6
7
8
9
10
Replicate
Figure 3-6.
Recovery from Stainless Coupons during INOC Series Tests.
These results demonstrate the ability of both the deposition method and the sponge-wipe recovery
method over a range of spore concentrations. This approach was then applied to carpet coupons so that
vacuum-based methods could be evaluated over a range of spore concentrations. The mean recoveries
(CFU) for each vacuum-based method are shown in Table 3-10 and Figure 3-7. These data were
gathered to evaluate the linearity of recoveries by each method when challenged with inocula spanning
several orders of magnitude.
Table 3-10. Mean Recoveries (CFU/sample) from Test LR (n=5)
Inoculation Level
104
Vacuum Sock
(Carpet)
1.06 x 103
Mean Recoveries (CFU/sample)
37 mm MCE
Extraction
(Carpet)
(Stainless Steel)
2.48 x 103
1.60 x 104
Sponge-wipe
(Stainless Steel)
1.27 x 104
106
2.02 x 105
5.07 x 105
1.93 x 106
1.81 x 106
107
2.45 x 106
8.65 x 106
2.27 x 107
1.69 x 107
31
Linear Recovery Test
1.00E+08
Recover (CFU)
1.00E+07
1.00E+06
1.00E+05
1.00E+04
Vacuum sock
1.00E+03
37 mm MCE
1.00E+02
1.00E+01
1.00E+00
E4
Figure 3-7.
E6
E7
Mean CFU/Sample from Linear Recovery (LR) Test. Data are plotted on a log-scale as
mean ± standard deviation.
Table 3-11 shows the relative recovery (percentage) based on stainless steel extraction method.
Table 3-11. Percent Relative Recovery of Vacuum Devices (Test LR) (n = 5)
Target
Inoculum
104
10
6
10
7
Relative Recovery
(percent of stainless steel
extraction recovery)
Vacuum Sock
37 mm MCE
4%
14%
3%
7%
6%
11%
7%
16%
9%
10%
12%
11%
16%
9%
6%
30%
12%
3%
11%
21%
21%
15%
19%
51%
25%
35%
104%
20%
9%
22%
Mean Relative Recovery
(percent of stainless steel extraction
recovery)
32
Vacuum Sock
37 mm MCE
7%
15%
10%
26%
11%
38%
The relative recovery (RR%) consists of the mean recovery (CFU) from each vacuum sample divided by
the mean recovery (CFU) from sponge-wipe sampling stainless steel (Table 3-12). These data suggest a
slight advantage of the 37 mm MCE vacuum method over the vacuum sock method, though the Student
t-test returned p-values ≥ 0.05 for each comparison.
Table 3-12. Relative Recovery of Vacuum Methods to Sponge-Wipe – Test LR (n=5).
Target
Inoculum
10
4
106
107
1
Relative Recovery
(percent of sponge-wipe recovery)
Vacuum Sock
37 mm MCE
5%
17%
3%
8%
8%
12%
7%
17%
9%
11%
17%
14%
22%
12%
8%
38%
15%
4%
14%
27%
22%
16%
21%
54%
27%
47%
140%1
27%
12%
30%
Mean Relative Recovery
(percent of sponge-wipe recovery)
Vacuum Sock
37 mm MCE
8.3%
19.6%
11.1%
28.0%
14.5%
51.2%
– Data point is within two standard deviations about the mean and the value was therefore not considered an outlier.
ANOVA analysis of CR% over the three inoculums suggested that there was no difference in the
recovery over the entire range for either vacuum method. (i.e., no statistical difference in CR% between
devices at each inoculum)
Results from Test LV showing variation in laboratory and personnel for two vacuum methods are shown
in Figure 3-8 and Table 3-13.
33
Laboratory Variability
7.10E+06
Recovery (CFU)
6.10E+06
5.10E+06
4.10E+06
Lab 1
3.10E+06
Lab 2
2.10E+06
1.10E+06
1.00E+05
37 mm MCE
Figure 3-8.
Vacuum Sock
Laboratory Variability for Two Vacuum Methods - Test LV
Table 3-13. Laboratory Variability for Two Vacuum Methods (Test LV)
Sample Type
n
% Recovery of Extraction Controls
Average CFU/Sample
Lab 1
Lab 2
37 mm MCE
10
3.58 x 10
6
Vacuum Sock
10
2.01 x 10
6
Lab 1
Lab 2
4.12 x 10
6
12.4 %
13.7 %
2.47 x 10
6
6.8%
8.4%
Standard Deviation
Sample Type
n
Lab 1
Standard Deviation
Lab 2
37 mm MCE
10
1.34 x 10
6
Vacuum Sock
10
1.06 x 10
6
Lab 1
Lab 2
1.25 x 10
6
4.6 %
4.6%
4.86 x 10
5
3.7 %
1.7%
CV%
CV%
Sample Type
n
Lab 1
Lab 2
Lab 1
Lab 2
37 mm MCE
10
37%
34%
37%
34%
Vacuum Sock
10
53%
20%
53%
20%
T-Test Between Laboratories
T-Test Between Laboratories
Sample Type
n
37 mm MCE
10
0.522
0.522
Vacuum Sock
10
0.234
0.234
There was no statistically significant difference between the recoveries of the two laboratories. Pooling
the results of the two laboratories, however, does show a significant difference between the results of the
-5
two vacuum methods, with a Student’s t-test value of 4.9 x 10 . With this large number of replicates, the
37 mm MCE vacuum method did provide better recovery than the vacuum sock method.
34
4
Quality Assurance
This project was performed under an approved Category III QAPP titled Evaluation of Vacuum-based
Sampling Devices for Collection of Biological Agent (DCMD 3.60) (October 2011).
4.1
Sampling, Monitoring, and Analysis Equipment Calibration
There were SOPs for the maintenance and calibration of all laboratory and NHSRC Microbiology
Laboratory equipment. All equipment was verified as being certified calibrated or having the calibration
validated by EPA’s APPCD on-site (RTP, NC) Metrology Laboratory at the time of use. Standard
laboratory equipment such as balances, pH meters, biological safety cabinets and incubators were
routinely monitored for proper performance. Calibration of instruments was done at the frequency shown
in Table 4-1. Any deficiencies were noted. The instrument was adjusted to meet calibration tolerances
and recalibrated within 24 h. If tolerances were not met after recalibration, additional corrective action was
taken, possibly including recalibration or/and replacement of the equipment.
Table 4-1. Sampling and Monitoring Equipment Calibration Frequency
Equipment
Calibration/Certification
Expected Tolerance
Thermometer
Compare to independent NIST thermometer (this is
a thermometer that is recertified annually by either
NIST or an International Organization for
Standardization (ISO)-17025 facility) value once per
quarter
±1°C
Stopwatch
Compare against NIST Official U.S. time at
http://nist.time.gov/timezone.cgi?Eastern/d/-5/java
once every 30 days.
±1 min/30 days
Clock
Compare to office U.S. Time @ time.gov every 30
days.
±1 min/30 days
Micropipets
All micropipets will be certified as calibrated at time
of use. Pipettes are recalibrated by gravimetric
evaluation of pipette performance to manufacturer's
specifications every year.
±5%
Scale
Compare reading to Class S weights
±1%
35
4.2
Data Quality
The primary objective of this project was to evaluate up to four currently-available vacuum-based devices
for biological sampling efficiency. Performance (recovery) of devices was compared to the currentlypreferred method, the "Vacuum Sock" (Midwest Filtration; Cincinnati, OH). Evaluation of operational
parameters included time required for sample collection, the physical impact on the sampling team during
collection, time required for sample analysis, and the cost of media and analysis equipment and supplies.
This section discusses the Quality Assurance/Quality Control (QA/QC) checks (Section 4.3) and
Acceptance Criteria for Critical Measurements (Section 4.4) considered critical to accomplishing the
project objectives.
4.3
QA/QC Checks
Uniformity of the test materials was a critical attribute to ensuring reliable test results. Uniformity was
maintained by obtaining a large enough quantity of material so that multiple material sections and
coupons could be constructed with presumably uniform characteristics. Samples and test chemicals were
maintained to ensure their integrity. Samples were stored away from standards or other samples which
could cross-contaminate them.
Supplies and consumables were acquired from reputable sources and were National Institute of
Standards and Technology (NIST)-traceable when possible. Supplies and consumables were examined
for evidence of tampering or damage upon receipt and prior to use, as appropriate. Supplies and
consumables showing evidence of tampering or damage were not used. All examinations were
documented and supplies were appropriately labeled. Project personnel checked supplies and
consumables prior to use to verify that they met specified task quality objectives and did not exceed
expiration dates.
Quantitative standards do not exist for biological agents. Quantitative determinations of organisms in this
investigation did not involve the use of analytical measurement devices. Rather, the CFU were
enumerated manually and recorded. QC checks for critical measurements/parameters are shown in Table
4-2. These checks also served as DQI goals. The acceptance criteria were set at the most stringent level
that could be routinely achieved and are consistent with the DQOs described in Section 4.4. Positive
controls and procedural blanks were included along with the test samples in the experiments so that wellcontrolled quantitative values were obtained. Verification of the sterility of samples prior to inoculation and
other background checks were also included as part of the standard protocol. Replicate coupons were
included for each set of test conditions. MOPs using qualified, trained and experienced personnel were
used to ensure data collection consistency. The confirmation procedure, controls, blanks, and method
validation efforts were the basis of support for biological investigation results. If necessary, training
sessions were conducted by knowledgeable parties, and in-house practice runs were used to gain
expertise and proficiency prior to initiating the research.
36
Table 4-2. QA/QC Sample Acceptance Criteria
Corrective
Actions
Sample Type
Purpose
Frequency
Acceptance Criteria
Negative control
(coupon without
biological agent)
Controls for sterility
of materials and
methods used in
the sampling
procedure.
1 per test
No observed CFU
Identify and
remove source
of
contamination.
Consult WAM.
Wipe Control
(wipe sample from
stainless steel
coupon
inoculation with
biological agent)
Verify inoculation
level on the
coupons and to
Demonstrate
plate’s ability to
support growth.
3 replicates per MDI
use
For high inoculation,
7
target loading of 10 CFU
per sample with a
standard deviation of
6
< 0.5 log10. (5 x 10 – 5 x
7
10 CFU/sample);
Grubbs outlier test (or
equivalent).
Outside target
range: discuss
potential impact
on results with
EPA WAM;
correct loading
procedure for
next test and
repeat,
depending on
decided impact.
Outlier: evaluate
stability of
MDIs.
Blank plating of
microbiological
supplies
Controls for sterility
of supplies used in
dilution plating,
includes beads,
PBST, dilution
tubes, and could
include other
supplies if filter
plates are needed.
3 of each supply per
plating event
No observed growth
following incubation
Sterilize or
dispose of
source of
contamination.
Re-plate
samples.
Blank Tryptic Soy
Agar Sterility
Control
(plate incubated,
but not inoculated)
Controls for sterility
of plates.
Each plate is incubated
at least 18 but fewer
than 24 h
No observed growth
following incubation.
All plates are
incubated prior
to use; all
contaminated
plates will be
discarded.
Field Blank
Samples
(Sample matrices
handled in
sampling area
without contact
with surfaces)
The level of
contamination
present during
sampling
3 per sampling event
No observed growth
following incubation
Clean up
environment.
Sterilize
sampling
materials before
use.
Tests with conditions falling outside these criteria were rejected and repeated. Decisions to accept or
reject tests were based upon engineering judgment used to assess the likely impact of the parameter on
the conclusions drawn from the data. For the current study, no tests required repeating.
Potential confounding organisms were excluded or controlled by sterilization of the materials and use of
aseptic technique, procedural blank controls, and a pure initial culture. Aseptic technique was used to
ensure that the culture remained pure. Blank controls were set up and sampled in parallel with the
37
contaminated materials. Infrequently, colonies were observed from negative control and procedural blank
samples. The magnitude of the recoveries from these samples was always at least 2 orders of magnitude
lower than test samples, and therefore did not significantly affect results.
4.4
Acceptance Criteria for Critical Measurements
The DQOs are used to determine the CMs needed to address the stated objectives and specify tolerable
levels of potential errors associated with simulating the prescribed decontamination environments. The
following measurements were deemed to be critical to accomplish part or all of the project objectives:
•
•
Enumeration of spores recovered from the surface of the coupons.
Total number of coupons vacuumed per sample.
The DQIs listed in Table 4-3 are specific criteria used to quantify how well the collected data met the
DQOs. Failure to provide a measurement method or device that meets these goals results in the rejection
of results derived from the CM. For instance, if the plated volume of a sample is not known (i.e., is not
100% complete), then that sample is invalid.
Table 4-3. Critical Measurement Acceptance Criteria
Critical
Measurement
Measurement
Device
Accuracy
Precision
Detection
Limit
Completeness
Pipet
±2%
±1%
NA
100%
CFU/Plate
Enumeration
by visual
inspection
±10% (between
2 counters)
±5
1 CFU
100%
Number of
coupons per
plate
Enumeration
by visual
inspection
±1%
±0.5
1
100%
Plated Volume
Plated volume critical measurement goals were met. All pipets are calibrated yearly by an outside
contractor (Calibrate, Inc., Carborro, NC) and verified gravimetrically at the conclusion of testing.
Plates were analyzed quantitatively (CFU/plate) using a visual inspection-based counting method. For
each set of results (per test), a second enumeration was performed on 25 percent of the plates within the
desirable range (30-300 CFU per plate). All second counts were found to be within 10 percent of the
original count.
There are many QA/QC checks used to validate microbiological measurements. These checks include
samples that demonstrate the ability of the NHSRC Microbiology Laboratory to culture the test organism,
as well as to demonstrate that materials used in this effort do not themselves contain spores. The checks
include:
38
•
Field blank coupons: sterile coupons sampled at the same time as inoculated coupons.
•
Field blank sample: vacuum-based sampling media attached to vacuum device, and device was
activated for one minute. Unfiltered laboratory air was collected, no surfaces were sampled.
•
Laboratory material coupons: includes all materials, individually, used by the NHSRC Microbiology
Laboratory in sample analysis.
•
Inoculation control coupons: stainless steel coupons inoculated at beginning, middle, and end of each
inoculation campaign. After 18 – 24 h, surfaces were sampled with PRB wipes and analyzed to
assess the precision and accuracy of the MDI during the inoculation operation.
4.5
Data Quality Audits
This project was assigned QA Category III and did not require technical systems or performance
evaluation audits.
4.6
QA/QC Reporting
QA/QC procedures were performed in accordance with the QAPP for this investigation.
39
5
Summary
5.1
Phase 1
Four vacuum methods (vacuum sock, 37 mm MCE filter, 37 mm PTFE filter, and TEF), were evaluated for
sampling porous materials (concrete, carpet, and upholstery). The vacuum sock method was tested
additionally at two sampling speeds. The vacuum sock and TEF method afforded more ease of use to
those collecting the samples and may therefore be more desirable for larger sample areas. However, the
37 MCE filter method demonstrated higher recoveries per unit area than did the vacuum sock device. The
TEF and vacuum sock (slow) method performed poorly on upholstery. The TEF also performed poorly on
carpet. There are advantages and disadvantages for each device.
Some conclusions can be reached based on Phase 1 recovery:
•
the speed of vacuum sock sampling (rate of speed the vacuum device traversed the coupon
surface) does not necessarily impact collection; and
•
overall recovery (including collection from the surface, retention on the filter, then extraction from
the filter) of spores using the 37 mm MCE vacuum method was often higher than the recovery
achieved from the other devices.
5.2
Phase 2
A method was developed which successfully deposited spores on (and through) HVAC filters under flow
conditions. Three methods of sampling HVAC filters (two vacuum methods and one extractive) performed
reliably on mechanical and electrostatic filters. The data suggest extractive methods may be more
efficient than vacuum-based recovery methods, depending on filter type. Vacuum sock sampling provided
higher recovery from mechanical filters, but both sock and 37 mm cassettes performed similarly when
sampling from electrostatic filters. Vacuum-based methods may be more applicable to HVAC filter media
that are not easily sectioned.
Phase 2 data also suggested there was no statistical difference between extraction, sponge-wipes, or
PRB wipes for recovery from stainless steel coupons.
5.3
Phase 3
The MDI inoculation method and the sponge-wipe recovery method were demonstrated to be effective
4
2
3
7
over a broad range of concentrations, from 1 to 1 × 10 CFU/cm (1 x 10 to 1x10 CFU/sample). Two
vacuum methods were also demonstrated effective over the same range. Compared to sponge-wipes
(from stainless steel), the vacuum sock method demonstrated recoveries between 3% and 22%, while the
37 mm MCE filter method demonstrated recoveries between 4% and 140% from carpet samples. The
extraction procedures for both vacuum methods were evaluated by two independent laboratory
technicians with no statistical difference in the number of spores recovered.
5.4
Lessons Learned and Application of Vacuum-based Methods to Field Use
Based on the data generated during this study, the 37 mm MCE and the vacuum sock sampling methods
were most efficient among the combination of materials and vacuum-based methods evaluated. When
40
looking at these methods from an operational perspective, the vacuum sock and trace evidence filter
methods offer advantages in that they allow a greater amount of surface area to be sampled in a given
amount of time. Increasing the amount of area sampled increases the representativeness of the sample.
Collecting samples more rapidly decreases the sampler’s time down range (in the contaminated area),
resulting in decreasing health and safety-related risks. However, each method has a unique set of
advantages and disadvantages. Table 5-1 below summarizes these advantages and disadvantages.
Table 5-1. Summary of Advantages and Disadvantages of Each Vacuum-Based Sampling Method.
37 mm (MCE or PTFE)
Advantages
Disadvantages
Vacuum Sock
Advantages
Disadvantages
Trace Evidence Filter
Advantages
Disadvantages
High collection
AC power required
Larger surface
AC power required;
Larger surface
AC power required;
efficiency for
for the Vac-U-Go
area collected
however, newer Omega
area collected
however, newer
some surfaces
SKC sampling pump
vacuum (Global
Omega vacuum
(i.e., concrete
(SKC, Inc.)
Industrial, Port
comes with battery
Washington, NY) comes
option
and upholstery)
with battery option
Sample is
Long sample
Short sample
Heavy equipment,
Short sample
Volumetric flow
easily contained
collection time, >5
collection time:
cumbersome in the field
collection time:
above 790 L min
and packaged
min per 1 ft
2 min per 1 ft2
will cause filter to
2
1.5 to 4 min per 1
2
following
-1
unseat from base,
ft
allowing air to
collection
bypass filter
Relatively
Increases health and
Vacuum housing
Poor quality control in
Vacuum
Expensive (~$20
simple
safety concerns due
can be
device construction
housing can be
per sample device)
extraction
to longer sampling
decontaminated
(some have visible holes
decontaminate
procedure
duration required for
by wiping
in the seams)
d by wiping
a given number of
samples
Shortest time
Sample is
Longest time
requirement of
easily
requirement of
the methods
contained and
methods evaluated
evaluated for
packaged
for laboratory
laboratory
following
extraction
extraction
sample
collection
In summary, all three methods are available and could be utilized during a large-scale event. The results
of the current study suggest the 37 mm and vacuum sock methods perform well and with less variability
than the TEF. When selecting a sampling method for nonporous surfaces, the method should be based
on sampling efficiency, portability, sampling time, surface area sampled, and health and safety of
sampling personnel.
41
5.5
Future Research
Future research efforts may focus on improving the 37 mm filter cassette sampling method. These
improvements could involve increasing the sampling speed of the 37 mm MCE collection to determine the
effect of sampling speed. Alternatively, the width of the 37 mm nozzle could be increased while
maintaining the cross-sectional area of the nozzle. Further testing would need to be conducted to
determine the effect of this nozzle alteration.
More testing should be conducted on a variety of HVAC filters to vet the methods more thoroughly. In
addition, further study is needed to investigate the effect of surface grime or conditions on recovery.
42
References
1.
Brown, G.S., R.G. Betty, J.E. Brockmann, D.A. Lucero, C.A. Souza, K.S. Walsh, R.M. Boucher, M.S. Tezak,
and M.C. Wilson, Evaluation of vacuum filter sock surface sample collection method for Bacillus spores from
porous and non-porous surfaces. Journal of Environmental Monitoring, 2007. 9(7): p. 666-671.
2.
Frawley, D.A., M.N. Samaan, R.L. Bull, J.M. Robertson, A.J. Mateczun, and P.C.B. Turnbull, Recovery
efficiencies of anthrax spores and ricin from nonporous or nonabsorbent and porous or absorbent surfaces
by a variety of sampling methods. Journal of Forensic Sciences, 2008. 53(5): p. 1102-1107.
3.
Estill, C.F., P.A. Baron, J.K. Beard, M.J. Hein, L.D. Larsen, L. Rose, F.W. Schaefer, 3rd, J. Noble-Wang, L.
Hodges, H.D. Lindquist, G.J. Deye, and M.J. Arduino, Recovery efficiency and limit of detection of
aerosolized Bacillus anthracis Sterne from environmental surface samples. Applied and Environmental
Microbiology, 2009. 75(13): p. 4297-306.
4.
Baron, P.A., C.F. Estill, J.K. Beard, M.J. Hein, and L. Larsen, Bacterial endospore inactivation caused by
outgassing of vapourous hydrogen peroxide from polymethyl methacrylate (Plexiglas®). Letters in Applied
Microbiology, 2007. 45(5): p. 485-490.
5.
Brown, G.S., R.G. Betty, J.E. Brockmann, D.A. Lucero, C.A. Souza, K.S. Walsh, R.M. Boucher, M. Tezak,
M.C. Wilson, and T. Rudolph, Evaluation of a wipe surface sample method for collection of bacillus spores
from nonporous surfaces. Applied and Environmental Microbiology, 2007. 73(3): p. 706-710.
6.
Calfee, M.W., S.D. Lee, and S.P. Ryan, A rapid and repeatable method to deposit bioaerosols on material
surfaces. Journal of Microbiological Methods, 2013. 92(3): p. 375-80.
7.
Busher, A., J. Noble-Wang, and L. Rose, Sampling for Biological Agents in the Environment, P. Emanual,
Roos, J.W.,Niyogi,K., Editor. 2008, ASM Press: Washington, DC,. p. pp 95-131.
8.
Rose, L.J., L. Hodges, H. O'Connell, and J. Noble-Wang, National validation study of a cellulose sponge
wipe-processing method for use after sampling Bacillus anthracis spores from surfaces. Applied
Environmental Microbiology, 2011. 77(23): p. 8355-9.
43
Appendix A: Miscellaneous Operating Procedures
MOP 3135
Procedure for Sample Collection using BactiSwabTM Collection and Transport Systems
MOP 3144
Procedure for Wipe Sampling of Coupons
MOP 3145
Procedure for Vacuum Sock Sampling of Large and Small Coupons
MOP 3161-F
Aerosol Deposition of Spores on HVAC Filter Under Flow
MOP 3161-HD
Aerosol Deposition of Spores onto Material Coupon Surfaces Using The Aerosol
Deposition Apparatus (ADA) – High Dosing
MOP 3161-LD
Aerosol Deposition of Spores onto Material Coupon Surfaces Using The Aerosol
Deposition Apparatus (ADA) – Low Dosing
MOP 3164
Procedure for 37MM Cassette and Trace Evidence Filter Vacuum Sampling of Large
and Small Coupons
MOP 3165
Sponge Sample Collection Protocol
MOP 3168
Aggressive Air Sampling (AAS) for WA 3-28: Phase 1 Sampling Approach
MOP 6535a
Serial Dilution: Spread Plate Procedure to Quantify Viable Bacterial Spore
MOP 6555:
Petri Dish Media Inoculation Using Beads
MOP 6562
Preparing Pre-Measured Tubes with Aliquotted Amounts of Phosphate Buffered Saline
with Tween 20 (PBST)
MOP 6563
Swab Streak Sampling and Analysis
MOP 6565
Filtration and Plating of Bacteria from Liquid Extracts
MOP 6567
Recovery of Bacillus Spores from Wipe Samples
MOP 6570
Use of STERIS Amsco Century SV 120 Scientific Pre-vacuum Sterilizer
MOP 6572
Recovery of Spores from Vacuum Sock Samples
MOP 6579
Recovery of Bacillus Spores from 37MM Filter Cassettes
MOP 6580
Recovery of Bacillus Spores from 3M Sponge-Stick™ Samples
MOP 6582
Recovery of Bacillus Spores from Trace Evidence Filters
MOP 6586
Recovery of Bacillus Spores from Quartz Filters
MOP 6593
Recovery of Spores from HVAC Filters
44
Appendix B: Miscellaneous Operating Procedures
TM
MOP 3135
Procedure for Sample Collection using BactiSwab
Collection and Transport Systems
MOP 3144
Procedure for Wipe Sampling of Coupons
MOP 3145
Procedure for Vacuum Sock Sampling of Large and Small Coupons
MOP 3161-F
Aerosol Deposition of Spores on HVAC Filter Under Flow
MOP 3161-HD
Aerosol Deposition of Spores onto Material Coupon Surfaces Using The Aerosol
Deposition Apparatus (ADA) – High Dosing
MOP 3161-LD
Aerosol Deposition of Spores onto Material Coupon Surfaces Using The Aerosol
Deposition Apparatus (ADA) – Low Dosing
MOP 3164
Procedure for 37MM Cassette and Trace Evidence Filter Vacuum Sampling of Large
and Small Coupons
MOP 3165
Sponge Sample Collection Protocol
MOP 3168
Aggressive Air Sampling (AAS) for WA 3-28: Phase I Sampling Approach
MOP 6535a
Serial Dilution: Spread Plate Procedure to Quantify Viable Bacterial Spore
MOP 6555:
Petri Dish Media Inoculation Using Beads
MOP 6562
Preparing Pre-Measured Tubes with Aliquoted Amounts of Phosphate Buffered Saline
with Tween 20 (PBST)
MOP 6563
Swab Streak Sampling and Analysis
MOP 6565
Filtration and Plating of Bacteria from Liquid Extracts
MOP 6567
Recovery of Bacillus Spores from Wipe Samples
MOP 6570
Use of STERIS Amsco Century SV 120 Scientific Prevacuum Sterilizer
MOP 6572
Recovery of Spores from Vacuum Sock Samples
MOP 6579
Recovery of Bacillus Spores from 37MM Filter Casettes
MOP 6580
Recovery of Bacillus Spores from 3M Sponge-Stick™ Samples
MOP 6582
Recovery of Bacillus Spores from Trace Evidence Filters
MOP 6586
Recovery of Bacillus Spores from Quartz Filters
MOP 6593
Recovery of Spores from HVAC Filters
MOP-3135
Revision 2
November 2012
Page 1 of 3
Miscellaneous Operating Procedure (MOP) 3135:
Procedure for Sample Collection Using BactiSwabTM
Collection and Transport Systems
Prepared by: __________________________________________
Date: 11/15/2012
Stella McDonald, ARCADIS Work Assignment Leader
Reviewed by: __________________________________________
Dahman Touati, ARCADIS Project Manager
Date: 11/15/2012
Approved by: __________________________________________
Worth Calfee, EPA Work Assignment Manager
Date: 11/15/2012
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
MOP-3135
Revision 2
November 2012
Page 2 of 3
MOP 3135
TITLE:
Procedure for Sample Collection using BactiSwabTM Collection and Transport
Systems
SCOPE:
This MOP describes the procedure for collecting swab samples for Low Tech
Decontamination Technique Testing
PURPOSE:
The purpose if this MOP is to ensure all swab sampling is performed in a
consistent manner.
Equipment/Reagents
1.0

Disposable lab coat

Nitrile examination gloves

P95 Respirator

Shoe covers

Bouffant cap

Safety glasses

BactiSwabTM Collection and Transport System
PROCEDURE
1. Before starting the swabbing procedure, make sure you are wearing the appropriate, projectspecific PPE (at a minimum gloves, lab coat, and safety glasses).
2. Through the sleeve, crush the BactiSwabTM ampule at midpoint.
3. Hold BactiSwabTM tip end up for at least five seconds to allow the medium to wet the swab.
4. Open the package and remove the BactiSwabTM.
5. Label the plastic tube appropriately using the following scheme:
X-Y-N where,
X is the test number,
Y is the material abbreviation, and
N is the material number
MOP-3135
Revision 2
November 2012
Page 3 of 3
6. Remove the cap-swab from the plastic tube.
7. Swab the surface while spinning the cap-swab between the thumb and index fingers.
Swabbing should be conducted by following the recommend guidelines for each material as
detailed in the project documentation (usually the QAPP).
8. Return cap-swab to tube.
9. Date and initial each sample tube. Enter this information into the lab notebook.
10. Complete the chain of custody form and relinquish the samples to the BioLab.
MOP-3144
Revision 2
November 2012
Page 1 of 7
Miscellaneous Operating Procedure (MOP) 3144:
Procedures for Wipe Sampling of Coupons
Prepared by: __________________________________________
Date: 11/15/2012
Stella McDonald, ARCADIS Work Assignment Leader
Reviewed by: __________________________________________
Dahman Touati, ARCADIS Project Manager
Date: 11/15/2012
Approved by: __________________________________________
Worth Calfee, EPA Work Assignment Manager
Date: 11/15/2012
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
MOP-3144
Revision 2
November 2012
Page 2 of 7
MOP-3144
TITLE:
PROCEDURE FOR WIPE SAMPLING OF COUPONS
SCOPE:
This MOP describes the procedure for wipe sampling both small and large
coupons.
PURPOSE:
The purpose of this MOP is to ensure consistent and representative sampling of
such coupons.
EQUIPMENT (quantities are per sampling kit)
1.0

Sterile sampling bag (10” x 14”) – outer bag

Sterile sampling bag (5.5” x 9 “) – inner “sample collection sterile sampling bag”

Two sterile 50 mL Falcon Blue-Max™ Polypropylene Conical Tubes

Sterile Kendall (ref. # 8402) 4-ply all-purpose sponge

Sterile phosphate buffered saline with 0.005% TWEEN®-20, prepared according to
MOP -6562

Pipette or other method for aseptic dispensing of 5 mL liquid

Sterile Posi-grip© forceps

P-95 Particulate Respirators – to prevent contamination and for respiratory protection.
(Specific projects may require additional respiratory protection and will be addressed in
the project Quality Assurance Project Plan (QAPP), e.g, SAR)

Powder-free Nitrile gloves (support person) and Kimtech Pure G3 Sterile Nitrile gloves
(sampler)

Dispatch® bleach wipes
PREPARATION
1. All materials needed for collection of each sample will be prepared in advance using aseptic
technique. A sample kit for a single wipe sample will be prepared as follows:
a. Two sterile sampling bags (10” x 14”, 5.5” x 9 “) and a 50 mL conical tube, capped,
will be uniquely labeled as specified in the project QAPP. These bags and conical
tube will have the same label. The 5.5” x 9” labeled sterile sampling bag will be
referred to as the sample collection sampling bag.
b. A sterile all-purpose sponge will be placed in an unlabeled sterile 50 mL conical tube
using sterile forceps and aseptic technique. The all-purpose sponge will be moistened
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by adding 2.5 mL of sterile phosphate buffered saline with 0.005% TWEEN®-20.
The tube will then be capped.
c. The labeled 50 mL conical tube (capped), the unlabeled conical tube containing the
pre-moistened all-purpose sponge, and the 5.5” x 9” labeled sampling bag will be
placed into the 10” x 14” labeled sampling bag. Hence, each labeled sampling bag
will contain a labeled 50 mL conical tube (capped), an unlabeled capped conical tube
containing a pre-moistened all-purpose sponge, and an empty labeled sampling bag.
d. Each prepared bag is one sampling kit.
2.0
SAMPLING PROCEDURE FOR SMALL 14”x14” COUPONS
1. A three person team will be used, employing aseptic technique throughout. The team will
consist of a sampler, sample handler, and support person.
2. Throughout the procedure, the support person will log anything they deem to be significant
into the laboratory notebook.
3. In general, the team works from the least contaminated sample set (i.e., control blanks)
towards the most contaminated sample set (i.e., positive controls).
4. The sampling team will each don a pair of sampling gloves (a new pair per sample, nonsterile, as they will only be handling non-sterile items); the sampler’s gloves shall be sterile
sampling gloves as they are the only member of the team in contact with the sample. All
members shall wear dust masks to further minimize potential contamination of the samples.
Depending on the situation, respiratory protection beyond a dust mask may be required to
protect the sampling team (e.g., SAR; this will be specified in the project QAPP). New
disposable lab coats are required for the sample handler when changing between different
types of materials or when direct contact between the coupon and lab coat occurs.
5. The sample handler will remove the coupon from the appropriate cabinet and place it on the
sampling area, being careful to handle the coupon only around the edges.
6. The support person will record the coupon code on the sampling log sheet.
7. The support person will remove a template from the bag and aseptically unwrap it such that
the sampler may grab it wearing sterile gloves.
8. The sampler will place the template onto the coupon surface and align it such that the edges
of the coupon are visible through the holes on the template.
9. The support person will remove a sample kit from the sampling bin and record the sample
tube number on the sampling log sheet next to the corresponding coupon code just recorded.
10. The sampler and support person will verify the sample code and ensure that the correct
coupon and location are being sampled.
11. The support person will:
a. Open the outer sampling bag touching the outside of the bag.
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b. Touching only the outside of the (10” x 14”) bag, remove and open the unlabeled
conical tube and pour the pre-moistened all-purpose sponge onto the sample or into
the sampler’s hands.
c. Discard the unlabeled conical tube.
d. Remove the sample collection sample bag (5.5” x 9”), being careful to not touch the
inside of the outer sampling bag, and open it touching only the outside.
e. Maneuver the labeled 50 mL conical tube to the end of the outer sterile sampling bag
and loosen the cap.
f. Remove the cap from 50 mL conical tube immediately preceding the introduction of
the sample into the tube.
12. The sampler will:
a. Wipe the surface of the sample horizontally using S-strokes to cover the entire sample
area of the coupon using a consistent amount of pressure.
b. Fold the all-purpose sponge concealing the exposed side and then wipe the same
surface vertically using the same technique.
c. Fold the all-purpose sponge over again and roll up the folded sponge to fit into the
conical tube.
d. Carefully place the all-purpose sponge into the 50 mL conical tube that the support
person is holding, being careful not to touch the surface of the 50 mL conical tube or
plastic sterile sampling bag.
13. The support person will then immediately close and tighten the cap to the 50 mL conical
tube and slide the tube back into the sample collection sampling bag and seal it.
14. The support person will then wipe the sample collection sampling bag with a Dispatch®
bleach wipe and place it into the outer sampling bag.
15. The support person will then seal the outer sample collection bag now containing the capped
50 mL conical tube (containing the all-purpose sponge) inside a sealed 5.5” x 9” sample
collection bag.
16. The support person will then decontaminate the outer sample bag by wiping it with a
Dispatch® bleach wipe.
17. The support person will then place the triply contained sample into the sample collection
bin.
18. All members of the sampling team will remove and discard their gloves.
19. Steps 2 – 18 will be repeated for each sample to be collected.
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3.0
SAMPLING METHOD FOR LARGE (4’x4’ or larger) COUPONS
3.1
Sample Layout
The sampling of large coupons is carried out using a sample grid to divide the large coupons into
representative sections. These sections are then numbered and selected to be sampled at different
times during the course of the experiment as a blank, a control, or an experimental group sample.
This selection grid is pre-determined and the Project Quality Assurance Project Plan (QAPP)
may overrule the template shown in Figure 1 if otherwise specified.
As in the example below, the first cell is sampled as a Blank before contamination. Starting in
cell 3, every third cell is sampled as a positive Control. This sample is to be taken postcontamination and before decontamination. Every cell directly following a Control cell is
sampled as Experimental and is taken following decontamination. The sample kit labeling will
be based on this grid and the sampling team must ensure to correctly sample the coupons based
on this template.
1
Blank
2
3
Control
4
Experimental
5
6
Control
7
Experimental
8
9
Control
10
Experimental
11
12
Control
13
Experimental
14
15
Control
16
Experimental
Figure 1. 4’ x 4’ Material Section Template and Sample Grid
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3.2
Sampling Procedure
1. A two-person team will be used, employing aseptic technique throughout. The team will
consist of a sampler and a support person.
2. Throughout the procedure, the support person will log anything they deem to be significant
into the laboratory notebook.
3. The sampling team will each don a pair of sampling gloves (a new pair per sample, nonsterile, as they will only be handling non-sterile items); the sampler’s gloves shall be sterile
sampling gloves as they are the only member of the team in contact with the sample. All
members shall wear dust masks to further minimize potential contamination of the samples.
Depending on the situation, respiratory protection beyond a dust mask may be required to
protect the sampling team (e.g., SAR; this will be specified in the project QAPP).
4. The support person will record the coupon code on the sampling log sheet.
5. The sampler will place the template onto the coupon surface (using clamps as necessary).
6. The support person will remove a sample kit from the sampling bin and record the sample
tube number on the sampling log sheet next to the corresponding coupon code just recorded.
7. The sampler and support person will verify the sample code and ensure that the correct
coupon and location (cell) is being sampled.
8. The support person will:
a. Open the outer sampling bag touching the outside of the bag.
b. Touching only the outside of the (10” x 14”) bag, remove and open the unlabeled
conical tube and pour the pre-moistened all-purpose sponge onto the sample or into
the sampler’s hands.
c. The unlabeled conical tube is retained for Step 9.
d. Remove the sample collection sample bag (5.5” x 9”) being careful to not touch the
inside of the outer sampling bag and open it touching only the outside.
e. Maneuver the labeled 50 mL conical tube to the end of the outer sterile sampling bag
and loosen the cap.
f. Remove the cap from 50 mL conical tube immediately preceding the introduction of
the sample into the tube.
9. The sampler will:
a. For a vertical coupon, the sampler will squeeze excess moisture from the sampling
sponge to prevent dripping down the sampling surface. The excess moisture is caught
in the unlabeled conical tube from Step 8c, and is then discarded.
b. Wipe the surface of the sample using S-strokes to cover the entire sample area of the
coupon (inside the grid) using a consistent amount of pressure.
c. Fold the all-purpose sponge concealing the exposed side and then wipe the same
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surface vertically using the same technique.
d. Fold the all-purpose sponge over again and roll up the folded sponge to fit into the
conical tube.
e. Carefully place the all-purpose sponge into the 50 mL conical tube that the support
person is holding being careful not to touch the surface of the 50 mL conical tube or
plastic sterile sampling bag.
10. The support person will then immediately close and tighten the cap to the 50 mL conical
tube and slide the tube into the sample collection sampling bag and seal it.
11. The support person will then wipe the sample collection sampling bag with a Dispatch®
bleach wipe and place it into the outer sampling bag.
12. The support person will then seal the outer sample collection bag now containing the capped
50 mL conical tube (containing the all-purpose sponge) inside a sealed 5.5” x 9” sample
collection bag.
13. The support person will then decontaminate the outer sample bag by wiping it with a
Dispatch® bleach wipe.
14. The support person will then place the triply contained sample into the sample collection
bin.
15. All members of the sampling team will remove and discard their gloves.
16. Steps 2 – 15 will be repeated for each sample to be collected.
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Miscellaneous Operating Procedure (MOP) 3145:
Procedure for Vacuum Sock Sampling of Large and Small Coupons
Prepared by: __________________________________________
Date: 11/15/2012
Stella McDonald, ARCADIS Work Assignment Leader
Reviewed by: __________________________________________
Dahman Touati, ARCADIS Project Manager
Date: 11/15/2012
Approved by: __________________________________________
Date: 11/15/2012
Worth Calfee, EPA Work Assignment Manager
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
MOP-3145
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November 2012
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MOP-3145
TITLE:
PROCEDURE FOR VACUUM SOCK SAMPLING OF LARGE AND SMALL
COUPONS
SCOPE:
This MOP describes the procedure for vacuum sampling of porous areas.
PURPOSE:
The purpose of this MOP is to ensure consistent and representative sampling of
such areas.
EQUIPMENT (quantities are per sampling kit)
•
1 – Gamma irradiated vacuum sock filtration kit
•
2 - Fisherbrand bags with round wire enclosure, 5.5” x 15” (Fisher Scientific, p/n 14-955181)
•
1 - Fisherbrand bag with round wire enclosure, 10” x 14” (Fisher Scientific, p/n 01-00253)
•
1 – 7.5” “cable-tie” (McMaster Car, Item# 7130K59)
•
Dispatch wipes
•
Fisher Scientific CLEAN_WIPES (Catalog No. 06-664-14) prepared with 400 mL of 3%
Sodium Thiosulfate solution (90mL of 1N STS in 310mL DI water)
•
Fisher Scientific CLEAN_WIPES (Catalog No. 06-664-24) premoistened with 70%
isopropyl alcohol
•
Permanent marker
•
Nitrile gloves
•
OMEGA HEPA vacuum
1.0 PREPARATION
All materials needed for each sample to be collected will be prepared in advance. A sample kit
for a single vacuum sock sample will be prepared using the procedure in MOP-3141A,
Procedure for Assembling Irradiated Vacuum Sock Sampling Kits.
2.0 VACUUM SAMPLING OF SMALL (14” by 14”) COUPONS
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The following procedure will be used in this study for vacuum sock sampling of each coupon
surface:
1. A two person team will be used, employing aseptic technique. The team will consist of a
sampler and a support person.
2. Both members of the sampling team will each don a pair of sampling gloves (a new pair per
sample); the sampler’s gloves shall be sterile sampling gloves if he/she is placing a template
onto the sample. Both members shall wear dust masks to further minimize potential
contamination of the samples. Further respiratory protection beyond a dust mask may be
required to protect the sampling team (e.g., SAR; this will be specified in the project QAPP).
3. The sampler will plug in the vacuum power cord and then don his/her sterile gloves.
4. The vacuum will be powered through a foot switch to allow the sampler to turn the vacuum
on and off. In general, care should be taken to direct the vacuum exhaust away from samples.
5. The support person will aseptically unwrap a template (if used) from the bag and present it to
the sampler, taking care to not touch the template.
6. The sampler will place the template onto the coupon surface.
7. The sampler will wipe the hose connection end (that receives the vacuum sock) first with a
fresh dispatch wipe, followed with a fresh CLEAN-WIPE containing 3% STS, followed by a
fresh CLEAN-WIPE containing the 70% isopropyl alcohol.
8. The sampler will hold the vacuum nozzle for the support person to place the vacuum sock
assembly onto the nozzle.
9. The support person will open the sampling supply bin and remove the vacuum sock sample
kit from the bin.
10. The support person will record the sample collection bag ID number on the sampling log
sheet or in the laboratory notebook.
11. The sampler and support person will ensure that the correct sample coupon has been selected,
referencing the coupon code on the sampling bag.
12. The support person will record the coupon code on the sampling log sheet next to the
corresponding vacuum sock collection bag number that was just recorded.
13. The support person will:
a. Open the vacuum sock sample kit outer bag and remove the unlabelled vacuum sock
assembly bag.
b. Tear open the bag containing the vacuum sock assembly and, working from the
outside of the bag, maneuver the assembly from the bottom to expose the cardboard
applicator tube opening.
c. Firmly place the vacuum sock assembly onto the nozzle of the vacuum tube, using the
bag to handle the vacuum sock assembly, while the sampler holds the vacuum nozzle.
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14. The sampler will:
a. Ensure that the sock is correctly placed on the nozzle and adjust, if necessary.
Care must be taken to not puncture or tear the sock.
b. Turn on the vacuum.
c. Vacuum “horizontally” using S-strokes to cover the entire area of the material
surface not covered by the template, while keeping the vacuum nozzle angled so
that the tapered opening of the vacuum sock is flush with the sample surface.
d. Vacuum the same area “vertically” using the same technique.
e. Turn off the vacuum when sampling is completed.
15. The sampler will remove the vacuum sock assembly from the nozzle, taking care to only
touch the cardboard and blue sections of the sock.
a. Remove the sock assembly from the vacuum hose. Take care not to touch the
sock inside the tube (Figure 1a shows the sock assembly).
b. Loosen the nozzle by pulling it free of the longer tube, while holding onto the
blue sock filter, and lightly replacing it in the longer tube.
c. Unfold the blue portion of the sock filter so that it is folded over the nozzle tubing
(Figure 1b).
d. While holding both section of cardboard tubing (one in each hand), partially
remove the angled section of tubing.
e. Present the sock to the support person, who will cinch the sock closed using a
plastic “cable tie” at the blue portion between the cardboard tubes (Figures 1c and
1d).
f. Reconnect the two cardboard tubes (Figure 1e).
g. Place nozzle end up in a labeled 5.5” x 15” bag held by the support person.
a.
b.
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c.
d.
e.
Figure 1. (a) Vacuum sock assembly, (b) blue portion of sock over nozzle tubing, (c,d)
cinching the sock with the “cable tie”, and (e) the two cardboard tubes
reconnected.
16. The support person will then seal the outer sterile sampling bag and wipe it with a
Dispatch® wipe.
17. The support person will then place this into the labeled 10” x 14” sample bag now
containing the outer and inner bags, the inner containing the vacuum sock assembly. The
outermost bag will then be wiped with a Dispatch® wipe.
18. The sampler will wipe down the nozzle (inside and out) and end of the tubing first with a
Dispatch® wipe, next with a wipe pre-moistened with 3% STS, next with a wipe premoistened with 70 % ethanol.
19. The support person will then place the triply contained sample into the sample collection
bin.
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20. All members of the sampling team will remove and discard their gloves.
21. Steps 2 – 20 will be repeated for each sample to be collected.
MOP-3161-F
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Miscellaneous Operating Procedure (MOP) 3161-F:
Aerosol Deposition of Spores on HVAC Filter Under Flow
Prepared by:
_________________________________________
Date: 4/29/2013
Matt Clayton, ARCADIS Work Assignment Leader
Reviewed by:
_________________________________________
Date: 4/29/2013
Dahman Touati, ARCADIS Project Manager
Approved by:
__________________________________________
Worth Calfee, EPA Work Assignment Manager
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
Date: 4/29/2013
MOP-3161-F
Revision 1
April 2013
Page 2 of 7
MOP 3161-F
TITLE:
AEROSOL DEPOSITION OF SPORES ON HVAC FILTER UNDER
FLOW
SCOPE:
This MOP outlines the procedure for the deposition of a spore loaded aerosol onto
an HVAC filter under high flow conditions.
PURPOSE:
To provide a procedure for the consistent deposition of spores onto HVAC filters
under flow.
MATERIALS

One 14” x 14” Aerosol Deposition Apparatus (ADA) (per inoculation)

One 14” x 14” gasket (per inoculation)

ADAdapter (see Figure 2)

Dispatch® wipes

STS Wipes

Isopropyl Alcohol Wipes

High Volume Sampler (Thermo Andersen 100042-00)

MOP 3161 ( and associated materials)

MOP 3168

Sterile forceps – Posi-Grip® Ref # 7190

Quartz filter sampling kits (prepared by BioLab)

Sterile filter sampling bag (VHP bag sterilized with EtO)

Enclosure aeration duct

Fisherbrand ® labeling tape – Catalog # - 15-951

Foot power switch – McMaster Item #96618

Sterilized Quartz filter (1 per inoculation) - WHATMAN® G653 Cat#-18209932

Sterilized 14” x 14” HVAC filter (1 per inoculation) - Purafilter 2000®

Medium binder clips – Staples ® Model#-10668-CC

Large binder clips - Staples ® Model#-10669

Blue spring clamps – Workforce 573-682

12” Double-sided sticky tape – Polyken 105C Carpet Tape
MOP-3161-F
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1.0
PREPARATION AND ASSEMBLY
1. Sterilize one 14” x 14” ADA, one 14” x 14” gasket, and an ADAdapter at least 3 days prior
to allow for off-gassing.
2. Sterilize one VHP bag per coupon using ethylene oxide (EtO) at least 3 days prior to
testing.
3. Set up the High Volume Sampler in the enclosure vented to the enclosure aeration system.
NOTE: Refer to Figure 1 to ensure assembly is correct during Steps 3-8. Figure 2 shows
the completed blower and ADA assembly.
4. The filter base on the blower assembly must be lifted above the top face of the sampler using
angle iron (See Figure 1a) to allow the seal between the base and ADAdapter to be clamped.
5. A two-person team is employed to aseptically install the quartz filter on top of the blower
assembly base as in MOP 3168, except there is no need for a third person to hold the bracket
as it is not required with this assembly (see Figure 1b).
6. Install the ADAdapter on top of the filter ensuring contact with the gasket. It is important that
the filter is installed such that all edges are covered by the gasket on the bottom of the
ADAdapter to ensure all airflow passes through it. Clamp the ADAdapter to the assembly
using small binder clips (see Figure 1c).
7. Once the ADAdapter is securely in place, aseptically install the 14” x14” HVAC filter.
Remove the filter from the bag using sterile gloves and place on top of the ADAdapter.
Ensure the filter is installed in the orientation pictured in Figure 1d.
8. Finally, Install the 14” x 14” gasket and ADA on top of the filter and secure the seal with the
spring clamps. Ensure the ADA is equipped with 4 filters and the top port is sealed with the
included mechanism (see Figure 1e).
9. Before proceeding to Section 2.0, prepare adequate space for the number of filters to be
puffed on cleaned tables. Use double-sided tape to create 16” x 16” settling areas, one for
each puffed filter. It is necessary to clean the table surfaces with Dispatch wipes, water and
alcohol. Do not use paper coverings due to using double-sided tape.
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a.
b.
c.
d.
e.
Figure 1. (a) Blower assembly base, (b) Quartz filter installed, (c) ADAdapter installed and clamped,
(d) HVAC filter installed, and (e) Completed assembly.
MOP-3161-F
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14x14 ADA
ADAdapter
Blower Assembly
base
Figure 2. Blower and ADA assembly
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2.0
PUFFING PROCEDURE
1. Determine the weight of the MDI canister using a balance. Record the MDI ID number and
the weight (to the nearest 0.01g) in lab notebook. In addition, keep a record of the total
number of ‘puffs’ dispensed for each MDI canister.
NOTE: The MDI canister full is approximately 15 grams, an empty canister is approx 9.5
grams. To ensure the canister contains adequate spore suspension for dosing,
canisters should be retired from use when their weight falls below 10.5 grams.
2. Next, assemble the MDI and actuator by inserting the MDI into the actuator, taking care not to
activate the MDI.
3. Vortex the MDI/actuator assembly for 30 seconds (the MDI canister should be in direct contact
with the vortex mixer).
4. Holding the MDI/actuator assembly upright, in a swift, firm motion, dispense three test ‘puffs’
into the aerosol trap to prime the MDI. It is important to vortex the assembly 10 seconds
before every puff (except 30 seconds prior to the initial puff of the experiment).
5. Vortex the assembly 10 seconds, and attach it to the ADA lid by mating the ADA adaptor to
the hole in the ADA lid. Loosen the lid screws enough to allow the lid to be slid into the
‘open’ position. Secure the lid in the open position by tightening the lid screws.
NOTE: The ‘open’ position is achieved when the hole in the lid aligns with the hole in the top
of the ADA.
6. Using the foot switch, start the blower and allow the unit to flow for 15 seconds.
7. At 15 seconds, dispense the spores by activating the MDI with a swift, firm motion. Hold the
MDI in the activated position for 3 seconds before releasing. Activation is best achieved by
grasping the MDI/actuator with two hands, and using a thumb to press the bottom of the MDI
canister.
8. After 30 seconds total, follow the reverse order of the lid opening procedure to close the ADA
lid, and turn off the blower using the foot switch.
9. Determine the weight of the actuator-MDI using a balance, and record the weight in lab
notebook.
NOTE: If the dosing puff is faulty, return to Step 5 and attempt a second puff on the current
filter. Do not proceed to the next filter until a ‘successful’ puff has been delivered. A
‘successful’ puff is achieved when the weight of the actuator- MDI assembly has a
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0.04gram-0.07gram loss. Familiarity and professional judgment will be needed to
determine the success of a puff.
10. Once the filter is successfully puffed, remove the ADA , gasket, and filter assembly. Clamp the
filter to the assembly using large binder clips, and place on the double sided tape.
11. Remove the ADAdapter and place on a clean surface.
12. Aseptically recover the quartz filter as in MOP 3168.
13. Clean the ADAdapter with Dispatch® wipes, STS wipes, and isopropyl alcohol wipes before
proceeding to Step 14.
14. For each additional filter, proceed to apparatus assembly, using a new sterile ADA and gasket
for each inoculation (Step 5, Section 1).
15. Repeat Steps 1.5 to 2.14 until all filters have been puffed (see test plan for number of samples).
16. Once all coupons have been puffed, remove the MDI from the actuator and weigh. Record the
final weight and total number of puffs.
17. Allow spores to settle onto the coupon surface for at least 18 hours. Settling time should not
exceed 26 hours.
3.0
RECOVERY
1. After 18 hours settling time, carefully remove binder clips and remove ADA and gasket from
coupon surface, taking care not to disturb the surface of the coupon.
2. Don sterile gloves and transfer the filter to sterilized VHP bag. Carefully tape the bag closed,
ensuring an air tight seal.
3. The VHP bag should be placed in tertiary containment before transport to the APPCD
Biocontaminant Laboratory for analysis.
MOP 3161-HD
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November 2012
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Miscellaneous Operating Procedure (MOP) 3161-HD:
Aerosol Deposition of Spores onto Material Coupon Surfaces using
the Aerosol Deposition Apparatus (ADA) – High Dosing
Prepared by: __________________________________________
Date: 11/15/2012
Stella McDonald, ARCADIS Work Assignment Leader
Reviewed by: __________________________________________
Dahman Touati, ARCADIS Project Manager
Date: 11/15/2012
Approved by: __________________________________________
Worth Calfee, EPA Work Assignment Manager
Date: 11/15/2012
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
MOP 3161-HD
Revision 3
November 2012
Page 2 of 7
MOP 3161-HD
TITLE:
AEROSOL DEPOSITION OF SPORES ONTO MATERIAL COUPON
SURFACES USING THE AEROSOL DEPOSITION APPARATUS (ADA) –
HIGH DOSING
SCOPE:
This MOP outlines the procedure for assembly and usage of the Aerosol
Deposition Apparatus (ADA).
PURPOSE:
Precise and highly repeatable aerosol deposition of bacterial spores onto material
surfaces for detection, sampling, and/or decontamination studies.
Materials:

Aerosol Deposition Apparatus (ADA) (shown in Figure 1)

Metered Dose Inhaler (MDI) preloaded with a bacterial spore suspension of known
concentration (i.e., 1 x 109 spores per puff)

Vertical MDI Actuator (shown in Figure 2)

Material coupon (with dimensions at least that of the ADA)

ADA-coupon gasket (1 per ADA) (see Figure 1)

Clamping devices (i.e., medium-size steel binder clips, C-clamps (8 per ADA))

Vortex mixer (shown in Figure 4)

Aerosol trap (described in Appendix A and shown in Figure 4)

Personal Protective Equipment (PPE) (gloves, lab coat, safety goggles)

pH-adjusted bleach (pAB) (MOP 3128-A)

0.22µm pore-size syringe filters (shown in Figure 1)

PVC tubing (3/8” OD, 1/4” ID)

Mass balance (with 0.01 gram accuracy)

Bench liner
1.0
STERILIZATION OF MATERIALS
Prior to the start of any experiment, all components must be sterilized and stored in a sterile
environment until usage. Sterilization is not necessary for binder clips, MDI, vortex, or the
aerosol trap.
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November 2012
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ADAs can be sterilized by autoclave, VHP, or by wiping with pH-adjusted bleach (pAB) with
subsequent deionized (DI) water and ethanol rinse/wipes. The ADA lid should be attached and
in the closed position during the sterilization.
ADA with lid in
closed position
2.2um
syringe filter
Gasket
ADA
Figure 1. ADA apparatus
The MDI actuator, with attached MDI adaptor, can be wiped with pAB then rinsed with DI
water.
MDI
Vertical
Actuator
Figure 2. MDI and vertical actuator
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Sterilization requirements for coupons vary by material. Regardless of the sterilization method,
quality control (QC) checks (typically by collecting a swab sample per MOP 3135) should be
administered to ensure the effectiveness of the sterilization method.
Gasket sterilization may also vary by material. Care should be taken to thoroughly degas gaskets
if sterilized via fumigation.
2.0
PROCEDURE
1. Begin by donning PPE (gloves, lab coat, and protective eyewear).
2. Clean the workspace by wiping with pAB, next with DI water , and lastly with a 70-90%
solution of denatured ethanol. Alternatively, new, clean bench liner may be placed on the
work surfaces. Make sure the workspace is clean and free of debris.
3. Discard gloves and replace with fresh pair.
4. Using aseptic techniques (when possible) assemble the coupon/ADA by first placing the
sterilized material coupon onto the clean lab bench or workspace, next place the sterilized
gasket on top of the coupon, and lastly seat the ADA on the coupon + gasket. Orient each
component so that it fits squarely with the previously placed item. Take care not to touch the
inside of the ADA or the coupon surface. Secure these components by attaching mediumsize binder clips, one at each corner, and one at the midpoint of each of the four sides of the
ADA. The binders should firmly secure the coupon to the ADA, and apply sufficient
pressure to the gasket to seal the union. If material coupons are too large to use binder clips
other methods may be used to secure the coupon and gasket to the ADA (i.e., larger clamps,
weight added to the ADA, etc.). Lastly, attach 0.2 um syringe filters to each vent tube on all
ADAs (4 per ADA). Syringe filters can be attached using PVC tubing (3/8” OD, 1/4” ID).
5. Determine the weight of the MDI canister using a balance. Record the MDI ID number and
the weight (to the nearest 0.01g) in lab notebook. In addition, keep a record of the total
number of ‘puffs’ dispensed for each MDI canister.
NOTE: The MDI canister full is approximately 15 g, an empty canister is approx 9.5 g. To
ensure the canister contains adequate spore suspension for dosing, canisters should
be retired from use when their weight falls below 10.5 g.
6. Next, assemble the MDI and actuator by inserting the MDI into the actuator, taking care not
to activate the MDI.
7. Vortex the MDI/actuator assembly for 30 seconds (the MDI canister should be in direct
contact with the vortex mixer).
8. Holding the MDI/actuator assembly upright (Figure 3), with a swift, firm motion, dispense
three test ‘puffs’ into the aerosol trap to prime the MDI. It is important to vortex the
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assembly 10 seconds before every puff (the exception being 30 seconds prior to the initial
puff of the experiment, as prescribed in Step 7).
Figure 3. MDI orientation while dispensing test puffs into the aerosol trap.
9. Vortex the assembly for 10 seconds and then attach to the ADA lid by mating the ADA
adaptor to the hole in the ADA lid. Loosen the lid screws enough to allow the lid to be slid
into the ‘open’ position. Secure the lid in the open position by tightening the lid screws.
NOTE: The ‘open’ position is achieved when the hole in the lid aligns with the hole in the
top of the ADA.
10. With a swift, firm motion, dispense the spores by activating the MDI. Hold the MDI in the
activated position for 3 seconds before releasing. Activation is best achieved by grasping the
MDI/actuator with two hands, and using a thumb to press the bottom of the MDI canister.
11. Follow the reverse order of the lid opening procedure to close the ADA lid.
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12. Determine the weight of the actuator-MDI using a balance, and record the weight in lab
notebook.
NOTE: If the dosing puff is faulty, return to Step 9 and attempt a second puff on the current
coupon. Do not proceed to the next coupon until a ‘successful’ puff has been
delivered. A ‘successful’ puff is achieved when the weight of the actuator-MDI
assembly has a 0.04 g to 0.07 g loss. Familiarity and professional judgment will be
needed to determine the success of a puff.
13. Vortex the assembly for 10 seconds, then proceed to dosing the next coupon (Step 9).
14. Repeat Steps 9 through 13 until all coupons have been dosed.
15. Once all coupons have been puffed, remove the MDI from the actuator and weigh. Record
the final weight and total number of puffs.
16. Allow spores to settle onto the coupon surface for at least 18 hours. Settling time should not
exceed 26 hours.
17. Carefully remove binder clips (or other attachment device), and remove ADA and gasket
from coupon surface, taking care not to disturb the surface of the coupon.
18. Test coupon is now ready for use.
19. Decontaminate the ADA and associated components with the same procedures utilized
during the initial sterilization.
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APPENDIX A - Aerosol Trap
Purpose: This device allows test puffs of the MDI to be deployed without contamination of the
surrounding area. Spores are pulled into the trap, contained, and inactivated.
This device consists of a suction source, a trap (containing pAB), and an inlet funnel.
Aerosolized spores are pulled into the funnel, and forced into the trap. The spores are collected
and inactivated as the aerosol flows through the pAB solution. The effluent air traveling toward
the suction device is spore-free downstream of the trap. See Figure 4.
The aerosol trap should be assembled inside a biological safety cabinet (BSC) or chemical fume
hood.
Aerosol trap
Vortexer
Figure 4. Aerosol trap
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Miscellaneous Operating Procedure (MOP) 3161-LD:
Aerosol Deposition of Spores onto Material Coupon Surfaces using
the Aerosol Deposition Apparatus (ADA) – Low Dosing
Prepared by: _________________________________________
Date: 4/29/2013
Matt Clayton, ARCADIS Work Assignment Leader
Reviewed by: _________________________________________
Date: 4/29/2013
Dahman Touati, ARCADIS Project Manager
Approved by: __________________________________________
Worth Calfee, EPA Work Assignment Manager
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
Date: 4/29/2013
MOP-3161-LD
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MOP 3161-LD
TITLE:
AEROSOL DEPOSITION OF SPORES ONTO MATERIAL COUPON
SURFACES USING THE AEROSOL DEPOSITION APPARATUS (ADA) –
LOW DOSING
SCOPE:
This MOP outlines the procedure for assembly and usage of the Aerosol
Deposition Apparatus (ADA) for low dosing.
PURPOSE:
To provide a procedure for repeatable aerosol deposition of bacterial spores onto
material surfaces for detection, sampling, and/or decontamination studies.
MATERIALS

Aerosol Deposition Apparatus (ADA)

Metered Dose Inhaler (MDI) preloaded with a bacterial spore suspension of known
concentration (i.e., 5 x 106 spores per puff or less, also designated as 5E6)

MDI Actuator with ADA adaptor (adaptor required for proper attachment of actuator to
ADA lid). The MDI actuator label must match the concentration and organism of the
MDI.

Material coupon (with dimensions at least that of the ADA)

ADA-coupon gasket (1 per ADA)

Clamping devices (i.e., medium-size steel binder clips, C-clamps (8 per ADA))

Vortex mixer

Aerosol trap (described in Appendix A)

PPE (gloves, lab coat, P95 dust mask, cap or hair bonnet, safety goggles)

pH-adjusted bleach (pAB; prepared per MOP 3128)

0.22µm pore-size syringe filters

PVC tubing (3/8” OD, 1/4” ID)

mass balance (with 1 mg accuracy)

canned air duster
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1.0
DISINFECTION OF MATERIALS
Prior to the start of any experiments, all components must be disinfected, and stored in as close
to a sterile environment as possible until usage. (Disinfection is not necessary for binder clips,
MDI, vortex, and the aerosol trap).
ADAs can be disinfected by autoclave, VHP, or pAB wiping with subsequent deionized water
(diH2O) and ethanol rinse/wipes. The ADA lid should be attached and in the closed position
during the disinfection.
MDI actuator with attached MDI adaptor can be wiped with pAB then rinsed with diH2O.
NOTE: DO NOT expose to ethanol, as ethanol will shatter the ADA adaptor.
Disinfection requirements for coupons vary by material. Regardless of disinfection method, QC
checks should be administered to ensure the effectiveness of the disinfection method.
Gasket disinfection may also vary by material.
2.0
PROCEDURE
1. Begin by donning PPE (gloves, lab coat, and protective eyewear).
2. Cover the tables with fresh bench liner.
3. Discard gloves and replace with fresh pair.
4. Using aseptic techniques (when possible) assemble the coupon/ADA by first placing the
disinfected material coupon onto the clean lab bench or workspace. Next place the
disinfected gasket on top of the coupon, and then seat the ADA on the coupon + gasket.
Orient each component so that it fits squarely with the previously placed item. Take care not
to touch the inside of the ADA or the coupon surface. Secure these components by attaching
medium-size binder clips, one at each corner, and one at the midpoint of each of the four
sides of the ADA. The binders should firmly secure the coupon to the ADA and apply
sufficient pressure to the gasket to seal the union. If material coupons are too large to use
binder clips, other methods may be used to secure the coupon and gasket to the ADA (i.e.,
larger clamps, weight added to the ADA, etc.).
Lastly, attach 0.2 um syringe filters to each vent tube on all ADAs (4 per ADA). Syringe
filters can be attached using PVC tubing (3/8” OD, 1/4” ID).
5. Determine the weight of the MDI canister using a balance. Record the weight (to the nearest
mg) in lab notebook. In addition, keep a record of the total number of ‘puffs’ dispensed for
each MDI canister.
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NOTE: A full MDI canister is usually approximately 20 grams (although they are
sometimes 15 grams); an empty canister is approximately 9.5 grams. To ensure
the canister contains adequate spore suspension for dosing, canisters should be
retired from use when their weight falls below 10.5 grams.
6. Next, assemble the MDI and actuator (with ADA adaptor) by inserting the MDI into the
actuator, taking care not to activate the MDI. Weigh the MDI and actuator assembly and
record the weight (to the nearest mg) in lab notebook.
7. Vortex the MDI/actuator assembly for 30 seconds (the MDI canister should be in direct
contact with the vortex mixer).
8. Holding the MDI/actuator assembly upright (MDI canister on top, MDI nozzle pointing
down, and actuator + adaptor outlet pointing horizontal, in contact with the aerosol trap inlet
adapter − see Figure 1), dispense three test ‘puffs’ into the aerosol trap to prime the MDI. It
is important to vortex the assembly 10 seconds before every puff (except 30 seconds prior to
the initial puff of the experiment, as prescribed in Step 7). Weigh the MDI/actuator assembly
after each puff and record in the Deposition Log. A weight reduction of 0.04-0.07g will
indicate a successful puff.
Figure 1. MDI orientation while dispensing test puffs into the aerosol trap.
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9. Vortex the assembly 10 seconds, and attach it to the ADA lid by mating the ADA adaptor to
the hole in the ADA lid. Loosen the lid screws enough to allow the lid to be slid into the
‘open’ position. (The ‘open’ position is achieved when the hole in the lid aligns with the hole
in the top of the ADA.) Secure the lid in the open position by tightening the lid screws.
10. With a swift firm motion, dispense the spores by activating the MDI. Hold the MDI in the
activated position for 3 seconds before releasing. Activation is best achieved by grasping the
MDI/actuator with two hands, and using a thumb to press the bottom of the MDI canister.
Weigh the MDI and record in the Deposition Log assembly.
NOTE: If the dosing puff is faulty, return to Step 8 and attempt a second puff on the
current coupon. Do not proceed to the next coupon until a ‘successful’ puff has
been delivered. A ‘successful’ puff is achieved when the plume can be seen
through the clear ADA adaptor immediately following the puff. The puff results
in audible characteristics typical of a successful puff, and the proper weight
reduction is achieved. Familiarity and professional judgment will be needed to
determine the success of a puff.
11. Follow the reverse order of the lid opening procedure to close the ADA lid. Detach the
MDI/actuator assembly from the lid.
12. Remove the MDI from the actuator and determine the weight of the MDI using a balance,
record the weight in lab notebook.
13. Place the empty actuator into the aerosol trap and remove dosing residue from the inner
surface of the actuator using the canned air duster. Take care to blow the compressed air into
the bottom of the actuator so that the residue is blown directly into the aerosol trap.
14. Vortex the assembly for 10 seconds, holding the assembly upright (MDI canister on top,
MDI nozzle pointing down, and actuator + adaptor outlet pointing horizontal, towards the
aerosol trap inlet − see Figure 1), dispense a test puff into the aerosol trap (expect a faulty
puff, as the MDI requires recharging after a puff in the non-upright orientation). Repeat once
more (vortex 10 seconds, puff). Expect the subsequent puff to be characteristic (visually,
audibly, and with expected weight loss) of a successful puff. If the second test puff is
successful, proceed to dosing the next coupon (Step 9). If the second puff is faulty, repeat
Step 13.
15. Repeat Steps 9 through 13 until all coupons have been dosed.
16. Allow spores to settle onto the coupon surface for at least 18 hours. Settling time should not
exceed 26 hours.
17. Wearing the proper PPE, carefully remove binder clips (or other attachment device), and
remove ADA and gasket from coupon surface, taking care not to disturb the surface of the
coupon.
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18. Test coupon is now ready for use.
19. Decontaminate the ADA and associated components with the same procedures utilized
during the initial disinfection.
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APPENDIX A – AEROSOL TRAP
Purpose: This device allows test puffs of the MDI to be deployed without contamination of the
surrounding area. Spores are pulled into the trap, contained, and inactivated.
This device consists of a suction source, a trap (containing pAB), and an inlet funnel.
Aerosolized spores are pulled into the funnel and forced into the trap. The spores are collected
and inactivated as the aerosol flows through the pAB solution. The effluent air traveling toward
the suction device is spore-free downstream of the trap. See Figure 2.
The aerosol trap should be assembled inside a biological safety cabinet (BSC) or chemical fume
hood.
Suction
Device
pH-adjusted bleach
Figure 2. Schematic drawing of Aerosol Trap.
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Miscellaneous Operating Procedure (MOP) 3164:
Procedure for 37MM Cassette and Trace Evidence Filter Vacuum
Sampling of Large and Small Coupons
Prepared by: _________________________________________
Date: 4/29/2013
Matt Clayton, ARCADIS Work Assignment Leader
Reviewed by: _________________________________________
Date: 4/29/2013
Dahman Touati, ARCADIS Project Manager
Approved by: __________________________________________
Worth Calfee, EPA Work Assignment Manager
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
Date: 4/29/2013
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TITLE:
PROCEDURE FOR 37MM CASSETTE AND TRACE EVIDENCE FILTER
VACUUM SAMPLING OF LARGE AND SMALL COUPONS
SCOPE:
This MOP describes the procedure for vacuum sampling of porous areas.
PURPOSE:
The purpose of this MOP is to ensure consistent and representative sampling of
such areas.
A: 37 mm filter
EQUIPMENT (starred* quantities are per sampling kit)
•
1.0
*1 – 37 mm Filter cassette loaded with desired filter
-
0.3 µm pore size PTFE membrane (SKC 225-1723)
-
0.8 µm pore size MCE membrane (SKC 225-3-01)
•
Vac-U-Go pump (SKC 228-9605)
•
Rotameter (SKC 320-100)
•
Tygon tubing, 1/4 in ID, 7/16 in OD, 50ft (SKC 225-1345)
•
*Two PVC adapters, 250/pkg (SKC 225-132A)
•
*One BD Falcon Sterile 15 mL Polyprolene Conical Tubes (fishersci.com, P/N 225352196)
•
*Two 5.5” x 9” 3.5 mil sterile sampling bags with flat-wire closures (fishersci.com,
Item# 14-955-187)
•
*10” x 14” overpack sample bag with round wire enclosure (fishersci.com, Item# 01-00253)
•
12” x 12” Template
•
Permanent marker
•
Nitrile gloves
•
Timer
PREPARATION
All materials needed for each sample to be collected will be prepared in advance. A sample kit
for a single cassette vacuum sample will be prepared using the following procedure.
1) Don nitrile gloves.
2) In laboratory, assemble nozzles:
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a) Cut the sampling nozzle from tygon tubing as follows: Using scissors, cut a section of
tygon tubing 1-inch long, then cut one end of the tubing at approximately a 45-degree
angle.
b) Place the nozzle onto a PVC adapter.
c) Prepare 25% more nozzles than required by the testing protocol.
d) Sterilize the nozzles with a 15 minute gravity cycle of the autoclave.
3) In laboratory, assemble cassette kit:
a) Label the 37 mm cassette with a unique sample ID.
b) Label the 15 mL conical tube with the same unique sample ID.
c) Aseptically remove the cassette plugs and place a PVC adaptor onto each end of the
cassette. Save the removed plugs for step f) below.
d) Cut a 20 cm long piece of tubing with the scissors.
e) Place the 20 cm tubing onto the downstream end of the cassette..
f) Place the sampling nozzle (1-inch section of tubing) onto the upstream end of the
cassette with the angled side furthest from the cassette.
g) Place the cassette with PVC adaptors, 20 cm tubing, and nozzle into a 5.5” x 9” 3.5
mil bag, making sure the end of the cassette that attaches to the vacuum tubing is
closest to the bag opening. (see Figure 1)
h) Place the red plugs removed from the cassette in step a) above into the same 5.5” x 9”
bag.
i) Place the 15 mL conical tube and the 5.5” x 9” bag with 37mm cassette assembly and
plugs into the 10” x 14” overpack bag.
j) Place an additional sterile small bag into the overpack bag.
4) Label the 10” x 14” bag with the sample ID.
5) Store sample kits in a clean dry location.
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Figure 1. 37mm Cassette Vacuum Inner Bag
2.0
VACUUM SAMPLING OF SMALL (12” by 12”) COUPONS
The following procedure will be used in this study for 37mm vacuum sampling of each coupon
surface:
1. A two-person team will be used, employing aseptic technique. The team will consist of a
sampler and a support person. In times when a third person (sample handler) is present, he or
she will act as an assistant to the support person for data recording.
2. The sampler will plug in the Vac-U-Go pump power cord, and attach the calibrated
rotameter to the pump. Turn on the pump and adjust the pump valve until the flow rate is 20
± 2 LPM. Record the flow rate in the project notebook. The sampler will then don his/her
sterile gloves.
3. Both members of the sampling team will each don a pair of sampling gloves (a new pair per
sample); the sampler’s gloves shall be sterile sampling gloves as they are the only member of
the team in contact with the sample. Both members shall wear dust masks to further
minimize potential contamination of the samples. Further respiratory protection beyond a
dust mask may be required to protect the sampling team (e.g., SAR; this will be specified in
the project QAPP).
4. The Vac-U-Go pump will be maintained on a rolling cart for easy movement into place.
5. The support person will aseptically unwrap a template from the bag and present it to the
sampler, taking care to not touch the template.
6. The sampler will place the template onto the coupon surface.
7. The support person will open the sampling supply bin and remove the 37mm cassette sample
kit from the bin.
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8. The support person or assistant will record the sample collection bag ID number on the
sampling log sheet or in the laboratory notebook.
9. The sampler and support person will ensure that the correct sample coupon has been selected,
referencing the coupon code on the sampling bag.
10. The support person or assistant will record the coupon code on the sampling log sheet next to
the corresponding 37mm cassette collection bag number that was just recorded.
11. The support person will:
a. Open the 37mm cassette sample kit outer bag and remove the unlabeled 37 mm
cassette assembly bag.
b. Open the small unlabeled bag containing the 37 mm cassette assembly.
c. Hold the bag so that the sampler can remove the kit.
d. The support person will hold the tubing for the sampler to place the 37mm cassette
assembly onto the tubing.
12. The sample handler will remove the 37 mm cassette assembly from the bag and attach to the
tygon vacuum tube held by the support person.
13. The support person or assistant will be prepared to record the duration of sampling. Prompts
should be given to the sampler so that the sample duration is close to the values in the Table
1.
Table 1: Suggested Sample Duration and Speed for 12” x 12” surface area
Material
Total Sampling
duration
Single Pass
duration
Number of passes
per direction
Concrete
300 seconds
3 seconds
50
Upholstery
300 seconds
3 seconds
50
Carpet
300 seconds
3 seconds
50
14. The sampler will:
a. Ensure that the filter is correctly placed on the tygon vacuum tube and adjust, if
necessary.
b. Turn on the vacuum.
c. Vacuum “horizontally” using S-strokes to cover the entire area of the material surface
not covered by the template, while keeping the tygon nozzle angled so that the
tapered opening of the tygon nozzle is flush with the sample surface.
Note: a target duration of time to vacuum each coupon should be determined prior to
testing for each material type. The sampler should pace the speed of the nozzle such
that the target sampling time is achieved for each coupon.
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d. Vacuum the same area “vertically” using the same technique.
e. Turn off the vacuum when sampling is completed.
f. Hold the nozzle and remove it from the cassette.
15. The support person will remove the 17 mm tube from the sample kit and open the tube.
16. The sampler will place the nozzle into the tube with the adapter end down while holding the
cassette in the opposite hand.
17. The support person will seal the tube and place in the small unlabeled bag.
18. The sampler will use the “nozzle” hand to remove the tubing from the outlet side of the
cassette and hold the cassette out to the support person.
19. The support person will
a. Don a fresh pair of gloves.
b. Seal the cassette with the two red plugs found in the small unlabeled sample
collection bag.
20. The support person will open the small unlabeled sample collection bag, and the sampler will
place the secured 37 mm cassette inside with the 15 mL conical tube.
21. The support person will then seal the small unlabeled sample collection bag and wipe it with
a Dispatch® wipe.
22. The support person will open the labeled 10” x 14” overpack bag and place the smaller
unlabeled collection bag containing the cassette inside.
23. The support person will then seal the labeled 10” x 14”overpack bag and wipe it with a
Dispatch® wipe.
24. The sampler will remove the used 20 cm length of tubing and discard.
25. The support person will then place the double contained sample into the sample collection bin.
26. All members of the sampling team will remove and discard their gloves.
27. Steps 3 – 22 will be repeated for each sample to be collected.
28. At the completion of testing, determine the final flow rate of the vacuum using the rotameter
and record in the project notebook.
B: 3M Trace Evidence Filter
Materials:

PPE (gloves, lab coat, safety goggles)
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
Sterile, sealed 10”x15” Twirl-em bags

Alcohol wipes

Dry wipes moistened with 3% sodium thiosulfate

Sharpie and/or pre-printed labels

Secondary containment such as a large Tupperware bin

Lab notebook

QAPP for project that is utilizing the Trace Evidence Filter samples

3M Forensic Vacuum Filter (Trace Evidence Filter) (Precision Data Products, Grand Rapids,
MI catalog #FF-1), referred to as TEF
1.0
PROCEDURE
1.1
Assembly of Trace Evidence Filter Kits
TEF kits are assembled in the following manner:
1. TEF kits can be assembled outside of the biological safety cabinet, in a dry, clean area.
Make certain to use proper PPE, including gloves, while handling all TEF kit materials.
Gather all materials to assemble the kits before assembly. These materials include:
- 3M Forensic Vacuum Filters
- 10”x15” Twirl-em bags
- Sharpie or pre-printed labels
2. Obtain a copy of the labeling scheme for the samples. This may be detailed in the QAPP.
For each TEF kit, use a Sharpie and label a large 10” x 15” Twirl-em bag with the correct
sample ID. Using 2 pre-printed labels, label the bag containing the TEF with a permanent
label and a label that can be easily removed. The removable label will be placed on the TEF
after sampling.
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Figure 1: Example of a permanent and removable label on a TEF
3. Open the labeled, 10” x 15” Twirl-em bags one at a time. Place the labeled, TEF in the 10” x
15” Twirl-em bags that have the corresponding label. Add a non-labeled, 10” x 15” Twirlem bag in the labeled 10” x 15” Twirl-em bag containing the TEF. This completes the TEF
kit assembly. In the lab notebook, record the date and project for which the TEF kits were
assembled.
4. Place the assembled TEF kits into a secondary containment, such as a large Tupperware bin.
When moving the kits to a sampling location, always have them in secondary containment.
Figure 2: Example of an assembled TEF kit
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2.0
VACUUM SAMPLING OF SMALL (12” by 12”) COUPONS
The following procedure will be used in this study for TEF vacuum sampling of each coupon
surface:
A two-person team will be used, employing aseptic technique. The team will consist of a
sampler and a support person. In times when a third person (sample handler) is present, he or she
will act as an assistant to the support person for data recording.
1) The Omega vacuum will be maintained on a rolling cart for easy movement into place.
2) The sampler will plug in the Omega Vacuum into a variable transformer (Staco Energy
Products Co., Model 3PN1010B) set on 50%. The sampler will wipe down the nozzle
(inside and out) and end of the tubing first with a Dispatch® wipe, next with a wipe premoistened with 3% Sodium thiosulphate, next with a wipe pre-moistened with 70 %
ethanol, then don his/her sterile gloves.
3) Both members of the sampling team will each don a pair of sampling gloves (a new pair
per sample); the sampler’s gloves shall be sterile sampling gloves as they are the only
member of the team in contact with the sample. Both members shall wear dust masks to
further minimize potential contamination of the samples. Further respiratory protection
beyond a dust mask may be required to protect the sampling team (e.g., SAR; this will be
specified in the project QAPP).
4) The support person will aseptically unwrap a template from the bag and present it to the
sampler, taking care to not touch the template.
5) The sampler will place the template onto the coupon surface.
6) The support person will open the sampling supply bin and remove the TEF sample kit from
the bin.
7) The support person or assistant will record the sample collection bag ID number on the
sampling log sheet or in the laboratory notebook.
8) The sampler and support person will ensure that the correct sample coupon has been
selected, referencing the coupon code on the sampling bag.
9) The support person or assistant will record the coupon code on the sampling log sheet next
to the corresponding TEF collection bag number that was just recorded.
10) The support person will:
a. Open the TEF sample kit overpack bag and remove the labeled TEF assembly
bag.
b. Open the labeled assembly bag containing the TEF cassette assembly.
c. Hold the bag so that the sampler can remove the TEF cassette without touching
the outside of the bag.
d. The sampler will remove the red round plug from the TEF cassette, and return it
to the inside of the bag.
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e. The sampler will attach the TEF to the vacuum hose.
f. The sampler will remove the black end cap from the TEF cassette, and return it to
the inside of the bag.
11) The support person or assistant will be prepared to record the duration of sampling.
Prompts should be given to the sampler so that the sample duration is close to the values in
the Table 2.
Table 2: Suggested TEF Sample Duration and Speed for 12” x 12” surface area
Material
Total Sampling
duration per
coupon
Single Pass
duration
Number of passes
per direction
Concrete
90 seconds
3 seconds
15
Upholstery
90 seconds
3 seconds
15
Carpet
90 seconds
3 seconds
15
12) The sampler will:
a. Ensure that the filter is correctly placed on the vacuum hose and adjust, if necessary.
b. Turn on the vacuum.
c. Vacuum “horizontally” using S-strokes to cover the entire area of the material surface
not covered by the template, while keeping the nozzle flush with the sample surface.
d. Vacuum the same area “vertically” using the same technique.
e. Turn off the vacuum when sampling is completed.
f. Reseal the inlet with the black end cap. This will be provided to him by the support
person working the cap to the lip of the bag, touching only the outside of the bag,
until the sampler can grab it.
g. Remove the filter from the vacuum hose.
h. Reseal the outlet with the red plug. This will be provided to him by the support
person working the cap to the lip of the bag, touching only the outside of the bag,
until the sampler can grab it.
13) The support person will remove the second removable label from the assembly bag and
place on the filter housing.
14) The support person will hold open the evidence bag while the sampler places the filter
inside.
15) The support person will seal the evidence bag with the self-adhering strip and wipe the
outside with a Dispatch wipe.
16) The support person will open the labeled 10” x 14” overpack bag and place the evidence
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collection bag containing the cassette inside.
17) The support person will then seal the labeled 10” x 14”overpack bag and wipe it with a
Dispatch® wipe.
18) The support person will then place the double contained sample into the sample collection bin.
19) All members of the sampling team will remove and discard their gloves.
20) The sampler will wipe down the nozzle (inside and out) and end of the tubing first with a
Dispatch® wipe, next with a wipe pre-moistened with 3% STS, next with a wipe premoistened with 70 % ethanol.
21) Steps 4 – 24 will be repeated for each sample to be collected.
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Miscellaneous Operating Procedure (MOP) 3165:
Sponge Sample Collection Protocol
Prepared by:
__________________________________________
Date: 11/15/2012
Stella McDonald, ARCADIS Work Assignment Leader
Reviewed by: __________________________________________
Dahman Touati, ARCADIS Project Manager
Date: 11/15/2012
Approved by: __________________________________________
Worth Calfee, EPA Work Assignment Manager
Date: 11/15/2012
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
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MOP 3165
Title:
SPONGE SAMPLE COLLECTION PROTOCOL
Scope:
This MOP outlines the procedure for collecting spores using a 3M Sponge-Stick™.
Purpose:
To provide a procedure for the collection of spore samples using a Sponge-Stick™
in a consistent and repeatable manner.
MATERIALS
•
3M Sponge-Sticks™ (P/N SSL10NB), hereafter referred to as ‘sponge’
•
One Seward stomacher bag (P/N BL6041/CLR) per kit
•
Disposable gloves
•
Sterilized sampling templates
•
One Fisher Sterile sampling bag with flat wire enclosure (7” x 12”, P/N 14-955-194) per kit
•
One Fisher Sterile sampling bag with flat wire enclosure (10” x 14”, P/N 01-002-53) per kit
for overpack
•
Dispatch wipes
1.0
PREPARATION
All materials needed for collection of each sample will be prepared in advance using aseptic
technique. A sample kit for a single sponge sample will be prepared as follows:
1.1
One stomacher bag will be uniquely labeled as specified in the project QAPP.
1.2
A 10” x 14” bag will be labeled with the same ID as the stomacher bag.
1.3
One stomacher bag, and one 9.5” x 12” unlabeled bag will be placed in the overpack bag.
1.4
A sterile Sponge-Stick will be added to the overpack bag.
1.5
Each prepared bag is one sampling kit.
2.0
PROCEDURE
A two person team will be used, employing aseptic technique throughout. The team will consist of
a sampler and a sample handler. In some cases, a third person may be needed to move samples.
Throughout the procedure, the support person will log anything they deem to be significant into the
laboratory notebook.
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In general, the team works from the least contaminated sample set (i.e., control blanks) towards the
most contaminated sample set (i.e.,positive controls).
All members shall wear dust masks to minimize potential contamination of the samples.
Depending on the situation, respiratory protection beyond a dust mask may be required to protect
the sampling team (e.g., SAR; this will be specified in the project QAPP). New disposable lab
coats are required for the sample handler when changing between different types of materials or
when direct contact between the coupon and lab coat occurs.
2.1
Wearing a clean pair of gloves over existing gloves, the sampler will place the disposable
template over the area to be sampled.
2.2
The support person will remove a sample kit from the sampling bin and record the sample
tube number on the sampling log sheet next to the corresponding coupon code just
recorded.
2.3
The sampler and support person will verify the sample code and ensure that the correct
coupon and location are being sampled.
2.4
The support person will:
a)
Open the outer sampling bag touching the outside of the bag.
b)
Touching only the outside of the (10” x 14”) bag, remove the sponge, and hand it to
the sampler.
c)
Remove the stomacher bag, being careful to not touch the inside of the outer sampling
bag, and open it touching only the outside.
2.5
The sampler will remove the sterile sponge from its package. Grasp the sponge near the top
of the handle. Do not handle below the thumb stop.
2.6
The sampler will wipe the surface to be sampled using the moistened sterile sponge by
laying the widest part of the sponge on the surface, leaving the leading edge slightly lifted.
Apply gentle but firm pressure and use an overlapping ‘S’ pattern to cover the entire
surface with horizontal strokes (Figure 1). Use the other hand to hold the template during
sampling, being careful not to touch the surface.
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Figure 1. First pass with sponge – horizontal strokes using one side of the sponge
2.7
The sampler will turn the sponge over and wipe the same area again using vertical ‘S’strokes (Figure 2).
Figure 2. Second pass with sponge – vertical strokes using the other side of the sponge
2.8
The sampler will the use the edges of the sponge (narrow sides) to wipe the same area using
diagonal ‘S’-strokes (Figure 3). The sponge will be flipped to use the opposite side
immediately after the longest stroke at opposite corners.
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Figure 3. Third pass with sponge – diagonal strokes using the edges of the sponge
2.9
The sampler will use the tip of the sponge to wipe the perimeter of the sampling area
(Figure 4).
Figure 4. Final (fourth) pass with sponge – perimeter wipe using the tip of the sponge
2.10 The sample handler will open the stomacher bag, careful not to touch the inside of the bag.
2.11 The sampler will place the end of the sponge in the bag, holding the handle outside the
opening of the bag.
2.12 The sample handler will grasp the sponge from outside of the bag, and help the sample break
off the handle of the sponge. The handle below the thumbstop should not touch the inside of
the stomacher bag.
2.13 The sample handler will securely seal the stomacher bag and wipe the outside with a
disinfecting wipe.
2.14 The sample handler will then place the stomacher bag inside the unlabeled sterile bag.
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2.15 The sample handler will place this in the overpack bag and wipe the overpack bag with
disinfecting wipes.
2.16 The sample handler will place the overpack bag in the sample bin.
NOTE: Remove excessive air from the re-sealable plastic bags to increase the number of
samples that can be shipped in one container.
2.17 The sampler will dispose of the template.
2.18 Both members will remove outer gloves and discard. Clean gloves should be worn for each
new sample.
3.0
REFERENCES
Sponge sample collection protocol adapted from:
National Validation Study of a Cellulose Sponge Wipe-Processing Method for Use after Sampling
Bacillus anthracis Spores from Surfaces. Rose, Laura J.; Hodges, Lisa; O’Connell, Heather;
Noble-Wang, Judith. Appl. Environ. Microbiol. 2011, 77(23):8355.
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MOP 3168
TITLE:
AGGRESSIVE AIR SAMPLING (AAS) FOR WA 3-28: PHASE I SAMPLING
APPROACH
SCOPE:
This MOP outlines the setup, operation, and timeline schedule for conducting
AAS testing in COMMANDER.
PURPOSE:
To provide a standardized and repeatable procedure for all AAS tests to be
conducted under WA 3-28’s Phase I sampling matrix using high volume (HiVol)
samplers.
1.0
INTRODUCTION
Preparations for each test will be conducted according to the schedule listed in this procedure.
Any deviations will be noted in the laboratory notebook, along with the reason for the deviation.
Section 2.0 lists the preparation steps that need to be taken the Thursday before testing is to
occur (Day 1). Sections 3.0 and 4.0 detail the pre- and post-decon procedures to be followed for
AAS testing.
2.0
PREPARATION
NOTE: Materials needed before Day 1 include material coupons and gaskets, placed in
VHP bags, and exposed to a VHP sterilization cycle. These should be left to
degas for a minimum of 3 days before use.
Day 1 – Thursday
1. Ensure the following materials for inoculation are in airlock:





(2) 3 x 3 Grids
(6) ADAs and clamps for single coupons
(18) ADAs and clamps for 3 x 3 coupons
Oscillating fan
Tables for coupon staging come Monday
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2. Ensure all equipment needed for testing is in COMMANDER and COMMANDER is setup
(see Figure 1):





SAR inlet filter box, hoses and masks
(2) Blowers and needed power cords
(3) HiVol Samplers in open orientation (no air-tight seals)
HiVol samplers should have been calibrated the day prior
Hydrogen peroxide (H2O2) air monitors are calibrated and functional
Fan
HighVol – R1
HighVol – R2
HighVol – R3
To airlock
Figure 1. COMMANDER layout showing HiVol sample numbers
3. Close and seal COMMANDER door.
4. Close airlock door.
5. VHP airlock per MOP-3120 (VHP Operation).
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Day 2 – Friday
1. Aerate the airlock upon arrival in the morning.
2. VHP COMMANDER per MOP-3120 in the morning, so that external aeration can begin in
mid afternoon.
3. Clear the airlock for entry as early as possible using a Draeger tube.
4. Set up coupons in H130, outside of the enclosure, using equipment from the airlock and
previously sterilized coupons and gaskets allowed to degas in VHP bags.
5. Inoculate coupons according to MOP 6561 (Aerosol Deposition of Spores onto Material
Coupon Surfaces using the Aerosol Deposition Apparatus (ADAs)). Fill out Attachment B,
WA 3-28 Coupon Deposition Log. The list of coupon IDs follows, where M is the material
ID; L(Laminate), D(Drywall), or C(Carpet):
42” x 42” not inoculated
28-[test ID]-XM,
42” x 42”
28-[test ID]-M
14” x 14” material
28-[test ID]-M1-R[1,2,3]
14” x 14” stainless steel
28-[test ID]-S-R[1,2,3]
14” x 14” material not inoculated
28-[test ID]-XM1-F1
6. Calibrate the COMMANDER ATI H2O2 sensor outside of COMMANDER using MOP 3136
(General Procedure for Calibration of ATI Hydrogen Peroxide Gas Transmitters using
Solution Wells).
7. Place the ATI sensor back into COMMANDER.
8. Mop the enclosure with Clorox Clean-up.
9. Close enclosure doors.
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3.0
PRE-DECON AAS PROCEDURES
Day 3 - Monday
 Conduct a pre-job safety briefing prior to initiating work.
 A “buddy” equipped with appropriate respiratory protection is required to be present
outside of COMMANDER when personnel are working inside.
1. Set up the ELPI system from outside of COMMANDER and zero according to MOP 3133
(Basic Operation and Maintenance of the Dekati Electrical Low Pressure Impactor (ELPI)).
2. Wipe down the outside of the 42” x 42” coupons, the 14” x 14” test coupons, and ADA’s
with dispatch wipes.
3. Transfer the coupons from H130 to the airlock. This operation will be done with a clean team
inside the airlock, and a team in H130. The H130 team (in clean garb) will complete the wipe
down, and then change garb. The airlock team will enter the airlock in clean garb and
booties. During the moving operation, the H130 team will pass the coupon to the airlock
team with no direct contact between alternate teams. Once both coupons have been passed
into the airlock, all personnel will leave the airlock, and it will be purged for 30 minutes. An
oscillating fan will be used to prevent settling.
4. Positive control 14” x 14” coupons should be moved at this same time, but put back in their
original location. This is to simulate the movement undergone by the test coupons.
5. Verify the camera is recording, and that the DAS is operational and recording data.
6. Assemble the supplies:
a. Sterilized PM10 filters (8)
b. Sterilized collection shims (8)
c. Sterile gloves (2 packs) and laboratory gloves
d. Sterile garb x 8(coat, p95, hair net, 12 boot covers)
e. EtO’d notebook, AAS event log and pen
f. Digital timer (2)
g. Duct tape
h. Blank sample kits (Filters Sample IDs : 28-05a-XAF-1-R(1-3), and 28-05a-XAF1-F1),( Shims Sample IDs : 28-05a-XAS-1-R(1-3) and 28-05a-XAS-1-F1)
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(Sponge sticks:
and 28-05a-S-1-F1)
28-05a-M-1-R(1-3), 28-05a-XM-1-F1 28-05a-S-1-R(1-3)
i. Cooling vests
j. Dispatch wipes
k. Molykote® Grease Spray
l. Anemometer
7. Three personnel (referred to as “Clean Man”, “Dirty Man 1”, and Dirty Man 2”, or CM,
DM1 and DM2) will enter the enclosure and wipe in bins of supplies, then close the
enclosure doors.
- Plug in and record position of power cords for blowers and fans.
- Install the relative humidity (RH) and vaprous hydrogen peroxide (VHP) sensors to
the COMMANDER chamber.
- Verify supply quantities against the WA 28 packing list (found on DTRL/WA 3-28).
- ELPI should be monitoring enclosure PM, and should be changed to COMMANDER
immediately before entry.
- Personnel should spend 10 minutes minimum in enclosure before opening airlock
door.
8. Three personnel (CM, DM1, and DM2) will enter the airlock wearing cooling vests, C-suits
and carrying the supply bins:
9. Once the airlock door is closed, open the door to COMMANDER. Put on clean garb over the
C-suit. Install the sterile filters on top of the vents of the three HiVol samplers (vent seen in
Figure 2a). To do this, the support person (DM1) will grab the filter frame with sterile gloves
and hold it out of the way. The sample handler (DM2) will open the bag containing the filter
and the sampler (CM) will use sterile thumb forceps to put the filter in place. The DM1 will
then return the frame and secure it in place. Unlatch the hinge of the head unit and pull down.
Secure latches around the base of the unit (Figure 2b). CM can help with this last step before
changing into new sterile gloves.
10. The DM1 will unwrap the aluminum foil covering the collection shim, leaving the shim
inside and exposed. The DM2 will spray the shim with Molykote® Grease Spray, ensuring
full coverage of the shim. Wearing sterile gloves, the CM will aseptically pull the shim out of
the foil and place in each of the three HiVol inlet heads, minimizing contact with the greased
surface. Align the shim and secure in place with the Teflon tabs (Figure 2c).
11. CM will unlatch the top covering of the head unit and pull down. Secure in place with the
latches (Figure 2d).
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12. DM1 and DM2 will return to the airlock.
13. Wearing sterile gloves, the DM1 will remove the apparatus brace, ADAs, and gaskets from
the sterile coupon and put aside in the airlock. DM1 and DM2 will pick up the coupon, being
careful not to touch the surface. Move the coupon into COMMANDER and place in the
designated sampling position. To do this, the coupon will need to be rotated vertically so that
it can fit through the door.
14. DM1 and DM2 remove their gowns and don SAR. The CM returns to the airlock and closes
the airlock door.
 Inspect SAR masks and hoses and the filter box prior to donning.
a.
b.
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c.
d.
Figure 2. a) HiVol sampler with head unit and top open, and filter vent secured;
b) head unit closed and latched; c) placement of shim; and d) top closed and
latched.
15. Turn on the fan to the 70% speed setting. Measure the wind speed at the HiVol inlets. These
should be less than 30 mph.
16. Test the force of the leaf blower, pointed away from the coupon, using the anemometer.
17. The DM1 will agitate the surface of the coupon using a leaf blower according to MOP 3166
(Aerosolization of Contaminated Coupons Using the Toro Power Sweep Electric Blower for
Aggressive Air Sampling (AAS)). At the very moment the blower is turned on, DM2 will turn
on all three HiVol samplers. DM2 will monitor the blowing time as per MOP 3166 and
document the intervals by filling out the Aggressive Air Sampling (AAS) Event Log
(Attachment A).
18. At the completion of the agitation, the DM1 and DM2 remain in COMMANDER. After 20
minutes total (10 minutes after finishing aggressive agitation), DM2 turns off the HiVol
samplers.
19. Turn off (unplug) the fan.
20. CM will re-enter COMMANDER.
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21. Collect blank samples:
Filters Sample IDs : XAF-1-R(1-3) + XAF-1-F1 (Field blank)
Shims Sample IDs : XAS-1-R(1-3) + XAS-1-F1 (Field blank)
a. DM1 and DM2 don outer layer of clean garb.
b. The DM1 will open the first HiVol sampler and latch in it place.
c. The DM1 will don a pair of sterile gloves and remove the collection shim, being careful
not to touch the sides of the flow ports.
d. The DM2 and CM will then use a sponge kit to sample the top surface of the shim
according to MOP 3169 (Sponge Sample Collection Protocol for AAS Shims).
e. The DM1 will then discard the shim to the airlock for sterilization. The DM1 will unlatch
the base of the HiVol head unit and tilt the unit up to reveal the vent (see Figure 2a).
f. The DM1 will don a new pair of sterile gloves and remove the vent bracket and top grate,
being careful not to touch the filter.
g. Donning new gloves, the CM will remove the filter using sterile thumb forceps. Fold the
filter in half and then in half again, keeping the top side of the filter inside of the fold.
Using the forceps, transfer the filter directly to a stomacher bag. The DM2 will hold the
stomacher bag open for this transfer.
h. Sample the shim plates and remove the filters for the two remaining HiVol samplers
using the same protocol.
22. Remove the blank coupon and place against the COMMANDER wall.
23. All personnel change into new sterile garb.
24. CM will install sterile filters on top of the vents of the three HiVol samplers and secure in
place. Unlatch the hinge of the head unit and pull down. Secure latches around the base of
the unit.
25. The DM1 will unwrap the aluminum foil covering the collection shim, leaving the shim
inside and exposed. The DM2 will spray the shim with Molykote® Grease Spray, ensuring
full coverage of the shim. Wearing sterile gloves, the DM2 will aseptically pull the shim out
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of the foil and place in each of the three HiVol inlet heads, minimizing contact with the
greased surface. Align the shim and secure in place with the Teflon tabs (see Figure 2c).
26. DM2 will unlatch the top covering of the head unit and pull down. Secure in place with the
latches.
27. Personnel will return to the airlock and DM1 and DM2 will don 3 pairs of boot covers.
28. Wearing gloves, the DM1 will remove the apparatus brace, ADAs, and gaskets from the
inoculated coupon and put aside in the airlock. DM1 and DM2 will pick up the coupon, being
careful not to touch the inoculated surface, and move the coupon into COMMANDER and
place in the designated sampling position. DM1 and DM2 will remove one pair of boot
covers while crossing the threshold into COMMANDER with the help of CM.
29. In the same aseptic manner, remove ADAs from material coupons and place inside on
COMMANDER floor just above the test coupon, removing a second pair of boot covers
while entering COMMANDER. This process will make one large 42” x 56” coupon.
30. The DM1 and DM2 remove their gowns and don SAR. CM closes the airlock door.
31. CM communicates to the buddy outside to unplug blowers 2 and 3. Confirm this is done to
DM2. Alternatively, the CM can unplug as he leaves the airlock.
32. DM2 then plugs in blowers 2 and 3 (now, or after the 12 minute AAS operation).
33. DM2 turns on the fan to the 70% speed setting.
34. Agitate the surface of the inoculated coupon according to MOP 3166. At the very moment
the blower is turned on, a DM2 will turn on the first HiVol sampler as specified in the QAPP.
The DM2 will monitor the blowing time as per MOP 3166 and document the intervals by
filling out the AAS Event Log.
35. When agitation is complete, remove SAR and leave in COMMANDER.
36. CM to exit airlock, leaving clean garb in the airlock upon exit.
37. DM 1 and DM2 will open the airlock door, move to the airlock and close the airlock door.
38. DM1 and DM2 will shower out and exit the airlock.
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39. After 20 minutes of air sampling, turn off the first HiVol sampler. Immediately turn on the
second sampler.
40. After 20 minutes of sampling with the second HiVol sampler, turn off the second sampler
and immediately turn on the third. Again, sample for 20 minutes and then turn off.
41. Wearing clean garb, collect samples from the stainless steel and laminate control coupons
according to MOP 3165 (Sponge Sample Collection Protocol). A blank laminate coupon (L1F1) and a field blank for sponge sticks (S1-F1) should also be sampled at this time.
Reference Positives IDs : S-1-R(1-3), L1-R(1,2,3), L1-F1, and S1-F1
42. VHP airlock and start aeration as soon as possible (MOP-3120). Allow 18 hours for the
agitated spores to settle in COMMANDER.
Day 4 – Tuesday
1. Verify the camera is recording, and that the DAS is operational and recording data.
2. After waiting the specified spore-settling time, a three-person sampling team (CM, DM1,
and DM2) will enter COMMANDER in C-suits carrying :
a. Sterile gloves and laboratory gloves
b. Sterile garb (coat, p95, hair net, 8 boot covers)
c. EtO’d notebook, AAS event log and pen
d. Digital timer
e. Duct tape
f. Sample
kits:
Kits:
28-05a-MAF-1-R{1-3)
28-05a-MAS-1-R{1-3) and 28-05a-MAS-1-F1
g. Dispatch wipes
and
28-05a-MAF-1-F1
3. Each will don sterile garb and boot covers.
4. Enter COMMANDER, removing one pair of boot covers while crossing the threshold.
5. Remove the air filters from the HiVol samplers. The team will consist of a CM, DM2 and a
DM1.
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Filter Sample IDs: MAF-1-R(1-3), where M is material ID – C-carpet, L-laminate, and
W-drywall
Shim Sample IDs: MAS-1-R(1-3), where M is material ID – C-carpet, L-laminate, and
W-drywall
6. The DM1 will open the first HiVol sampler and latch in it place.
7. The DM1 will don a pair of sterile gloves and remove the collection shim, being careful not
to touch the sides of the flow ports.
8. The DM2 and CM will then use a sponge kit to sample the top surface of the shim according
to MOP 3169 (Sponge Sample Collection Protocol for AAS Shims). CM will manipulate the
sponge stick.
9. The DM1 will then discard the shim into the airlock for disinfection. The DM1 will unlatch
the base of the head unit and tilt the unit up to reveal the vent.
10. The DM1 will don a new pair of gloves and remove the vent bracket and top grate, being
careful not to touch the filter.
11. Donning new gloves, the CM will remove the filter using sterile thumb forceps. Fold the
filter in half and then in half again, keeping the top side of the filter inside of the fold. Using
the forceps, transfer the filter directly to a stomacher bag. The DM2 will hold the stomacher
bag open for this transfer.
12. Sample the shim plates and remove the filters for the two remaining HiVol samplers using
the same protocol.
13. All HiVol samplers should be in the fully open position.
14. All personnel shower out of the airlock.
15. VHP COMMANDER as per MOP 3120. Conditions will be determined by the WAM.
16. Start aeration of COMMANDER.
17. VHP the airlock as per MOP 3120.
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4.0
POST-DECON AAS PROCEDURES
Day 5 - Wednesday
1. Start airlock aeration as soon as possible and ensure conditions are safe for entry.
2. Verify the camera is recording, and that the DAS is operational and recording data.
3. Three personnel will enter the airlock wearing C-suits and carrying:
a. Sterilized PM10 filters (3)
b. Sterilized collection shims (3)
c. Sterile gloves and laboratory gloves
d. Sterile garb (coat, p95, hair net, boot covers)
e. EtO’d laboratory notebook, AAS event log and pen
f. Digital timer
g. Duct tape
h. Dispatch wipe
i. Sample kits Sponge stick kits 28-05a-L-2-R(1-3) and 28-05a-L-2-F1
4. Once the airlock door is closed, open the door to COMMANDER. Don boot covers (2 pairs),
being careful not to contaminate the first pair with the airlock floor.
5. Move into COMMANDER, removing one pair of boot covers when crossing the threshold.
6. Collect post-decon sponge stick surface samples (or vacuum sock samples of carpet coupons)
of the three positive coupons per MOP 3165. CM will serve as the sampler, DM1 will serve
as the support person.
Sample IDs: M-2-R(1-3), where M is material type as listed above
7. DM2 will place coupons in soapy water in airlock after sampling.
8. All personnel to change into new clean garb.
9. CM will install the sterile filters on top of the vents of the HiVol samplers and secure in
place. DM2 will unlatch the hinge of the head unit and pull down. Secure latches around the
base of the unit.
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10. The DM1 will unwrap the aluminum foil covering the collection shim, leaving the shim
inside and exposed. The DM2 will spray the shim with Molykote® Grease Spray, ensuring
full coverage of the shim. Wearing sterile gloves, the DM2 will aseptically pull the shim out
of the foil and place in each of the three HiVol inlet heads, minimizing contact with the
greased surface. Align the shim and secure in place with the Teflon tabs (see Figure 2c).
11. The DM1 and DM2 will remove their gowns and don SAR, while CM retires to airlock.
12. CM to communicate to buddy outside to verify Blower 1 power cord is plugged in, and
Blowers 2 and 3 are off.
 Inspect the SAR masks and hoses and the filter box prior to donning.
13. Turn on the fan to the 70% speed setting. Measure the wind speed at the HiVol inlets. These
should be less than 30 mph.
14. Test the force of the leaf blower, pointed away from the coupon, using the anemometer.
15. Agitate the surface of the coupon according to MOP 3166. At the very moment the blower is
turned on, DM2 will plug in the first HiVol sampler. The DM2 will monitor the blowing time
as per MOP 3166 and document the intervals by filling out the AAS event log.
16. When agitation is complete, DM1 and DM2 move to airlock, remove SAR and drop their
hose and mask in COMMANDER, close the airlock door and shower out of the
COMMANDER airlock.
17. After 20 minutes of air sampling, turn off the first HiVol sampler. Immediately turn on the
second sampler.
18. After 20 minutes of sampling with the second HiVol sampler, turn off the second sampler
and immediately turn on the third. Again, sample for 20 minutes and then turn off.
19. Allow chamber to settle for 2 hours.
20. After the settling time, a three-person sampling team will enter COMMANDER carrying
post-decon sample kits: filter kits 28-05a-LAF-2-R{1-3) and 28-05a-LAF-2-F1 and shim
kits 28-05a-LAS-2-R{1-3) and 28-05a-LAS-2-F1
21. Sample the collection shims and collect the filters.
Filter Sample IDs: MAF-2-R(1-3), where M is material code , listed above
Shim Sample IDs: MAS-2-R(1-3), where M is material code , listed above
MOP 3168
Revision 3
February 2013
Page 15 of 17
a. The DM1 will open the first HiVol sampler and latch in it place.
b. The DM1 will don a pair of sterile gloves and remove the collection shim, being
careful not to touch the sides of the flow ports.
c. The DM2 and CM will then use a sponge kit to sample the top surface of the shim
according to MOP 3169.
d. The DM1 will then discard the shim to the airlock. The DM1 will unlatch the base of
the head unit and tilt the unit up to reveal the vent.
e. The DM1 will don a new pair of gloves and remove the vent bracket and top grate,
being careful not to touch the filter.
f. Donning new gloves, the CM will remove the filter using sterile thumb forceps. Fold
the filter in half and then in half again, keeping the top side of the filter inside of the
fold. Using the forceps, transfer the filter directly to a stomacher bag. The DM2 will
hold the stomacher bag open for this transfer.
g. Sample the shim plates and remove the filters for the two remaining HiVol samplers
using the same protocol.
22. Conduct post-test flow calibration as outlined in MOP 3170 (HiVol Calibration Check).
23. Verify that two blowers are placed in COMMANDER and that the samplers are opened.
Ensure all SAR related equipment is located in COMMANDER.
24. Exit COMMANDER and shower out of the airlock.
25. VHP COMMANDER per MOP-3120.
MOP 3168
Revision 3
February 2013
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Attachment A
Aggressive Air (AA) Sampling Event Log
Test ID ______________
Agitator
DM2
Buddy 2
Reference:
Dimensions
(in2)
Agitation
Time (s)
Horizontal
Sweeps
Vertical
Sweeps
14 x 14
28 x 28
42 x 42
60
240
540
10
20
30
10
20
30
Sweep
rate
(s/sweep)
3
6
9
Test Day/
date
Coupon ID
High Vol 1
start
High Vol
2 start
High Vol 3
start
Vertical
start
Vertical
end
# sweeps
2/
2/
4/
Test Day/
date
2/
2/
3/
3/
3/
4/
Sterile
Blank
Test
Test
Coupon ID
Sterile
Blank
Test
Positive 1
Positive 2
Positive 3
Test
30
30
10
10
10
30
Horizontal Horizontal
start
end
#
sweeps
30
30
10
10
10
30
MOP 3168
Revision 3
February 2013
Page 17 of 17
Attachment B
WA 3-28 Coupon Deposition Log
Test ID
Date
Personnel / Title
Time
Coupon
ID
Page ____ of ___
MDI ID:
Initial Weight (g):
Final Weight (g):
Vortex Interval (s)
Puff Number
MDI
Weight (g)
Comments
MOP 6535a
Revision 4
January 2013
Page 1 of 8
Miscellaneous Operating Procedure (MOP) 6535a:
Serial Dilution: Spread Plate Procedure to Quantify Viable
Bacterial Spores
Prepared by: __________________________________________
Date: 2/11/2013
Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader
Reviewed by: __________________________________________
Dahman Touati, ARCADIS Project Manager
Date: 2/11/2013
Approved by: __________________________________________
Worth Calfee, EPA Work Assignment Manager
Date: 2/11/2013
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
MOP 6535a
Revision 4
January 2013
Page 2 of 8
MOP 6535a
TITLE:
SERIAL DILUTION: SPREAD PLATE PROCEDURE TO QUANTIFY
VIABLE BACTERIAL SPORES
SCOPE:
Determine the abundance of bacterial spores in a liquid extract
PURPOSE:
Determine quantitatively the number of viable bacterial spores in a liquid
suspension using the spread plate procedure to count colony-forming units (CFU)
Materials:
•
Liquid suspension of bacterial spores
•
Sterile centrifuge tubes
•
Diluent as specified in QAPP or Test Plan (e.g., sterile water, Phosphate Buffered Saline with
Tween 20 (PBST))
•
Media plates as specified in QAPP or Test Plan (e.g., Trypticase Soy Agar (TSA) plates)
•
Microliter pipettes with sterile tips
•
Sterile beads placed inside a test tube (used for spreading samples on the media surface
according to MOP 6555 (Petri Dish Media Inoculation Using Beads) or cell spreaders
•
Vortex mixer
1.0 PROCEDURE (This protocol is designed for 10-fold dilutions.)
1. For each bacterial spore suspension to be tested label microcentrifuge tubes as follows: 10-1,
10-2, 10-3, 10-4, 10-5, 10-6... (The number of dilution tubes will vary depending on the
concentration of spores in the suspension). Aseptically, add 900 uL of sterile diluent to each
of the tubes.
2. Label three media plates for each dilution that will be plated. These dilutions will be plated
in triplicate.
3. Mix original spore suspension by vortexing thoroughly for 30 seconds. Immediately after the
cessation of vortexing, transfer 100 uL of the stock suspension to the 10-1 tube. Mix the 10-1
tube by vortexing for 10 seconds, and immediately pipette 100 uL to the 10-2 tube. Repeat
this process until the final dilution is made. It is imperative that used pipette tips be
exchanged for a sterile tip each time a new dilution is started.
4. To plate the dilutions, vortex the dilution to be plated 10 seconds, immediately pipette 100
uL of the dilution onto the surface of a media plate, taking care to dispense all of the liquid
MOP 6535a
Revision 4
January 2013
Page 3 of 8
from the pipette tip. If less than 10 seconds elapses between inoculation of all replicate
plates, then the initial vortex mixing before the first replicate is sufficient for all replicates of
the sample. Use a new pipette tip for each set of replicate dilutions.
5. Carefully and aseptically spread the aliquotted dilution on the surface of the media either by
use of glass beads (MOP 6555) or cell spreader (the method used may be directed in the
QAPP or Test Plan) until the entire sample is distributed on the surface of the agar plate.
Repeat for all plates.
6. Incubate the plates for the optimum time period at the optimum growth temperature for the
target organism (incubation conditions will vary depending on the organism’s optimum
growth temperature and generation time. This information can be found in Bergey’s Manual
of Determinative Bacteriology or it will be provided with the ATCC certification.
7. Manually enumerate the colony forming units (CFU) on the media plates by manually
counting with the aid of a plate counting lamp and a marker (place a mark on the surface of
the Petri dish over each CFU when counting, so that no CFU is counted twice). A hand held
tally counter or an electronic counting pen may be used to assist the person counting, but
may not be used as the primary source for the count.
Quality control (QC) requirements for bacterial enumeration will be addressed per QAPP or
test plan. However, in general, the following QC practices should always be adhered to:
a. The arrangement of plates and tubes, and the procedure for preparing dilutions and
enumerating CFU should be done the exact same way each time. This helps prevent
systematic errors and often helps determine the cause of problems when a discrepancy is
found.
b. A visual check of the graduated pipette tip should be made during each use to ensure the
pipette is pulling properly.
c. Samples should acclimate to room temperature for 1 hour prior to plating.
d. Samples should be processed (extracted and plated) from the least contaminated to the
most contaminated.
e. When a target range of CFU is known, three dilution factors are plated to bracket the
expected results (0, -1, and -2, if the -1 dilution factor was the target).
f. Enumerated colonies and results should be verified that the results are the target
organism, and that second counts have been performed. Second counts must be
completed on 25% of significant data, and must be within 10% of the first count. If
CFUs are found to have more than a 10% difference between first and second counts,
then a third count is to be completed.
MOP 6535a
Revision 4
January 2013
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g. Pictures should be taken of any plates that are contaminated or have results out of the
normal
8. Record all quantitative data in the “Serial Dilution/Plating Results Sheet”. Target range for
statistically significant counts is 30-300 CFU. Data that fall out of the 30-300 CFU range are
addressed in MOP 6584 (Procedure for Replating Bacteria Spore Extract Samples) and MOP
6565 (Filtration and Plating of Bacteria from Liquid Extracts).
2.0
CALCULATIONS
Total abundance of spores (CFU) within extract:
(Avg CFU / volume (mL) plated) X (1 / tube dilution factor) X extract volume
For example:
Tube Dilution
10-3
10-3
10-3
Volume plated
100 µL (0.1 mL)
100 µL (0.1 mL)
100 µL (0.1 mL)
Replicate
1
2
3
CFU
150
250
200
Extract total volume = 20 mL
(200 CFU / 0.1 mL) X (1/10-3) X 20 mL =
(2000)
X (1000) X 20
=
4.0 X 107 CFU
Note: The volume plated (mL) and tube dilution can be multiplied to yield a ‘decimal factor’
(DF). DF can be used in the following manner to simplify the abundance calculation.
Spore Abundance per mL
=
(Avg CFU) X
(1 / DF) X
extract volume
MOP 6535a
Revision 4
January 2013
Page 5 of 8
Serial Dilution/Plating Results Sheet
Page 1 of ______
TEST INFORMATION
EPA Project No.
Technician Name
Technician Signature
PI
Test Date
Test No.
RESULTS
Date:
Volume Plated:
Tube Dilution
Sample ID
Plate
Repl.
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
NOTES:
100
10-1
10-2
10-3
10-4
10-5
10-6
MOP 6535a
Revision 4
January 2013
Page 6 of 8
Page 2 of ______
Sample ID
Plate
Repl.
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
NOTES:
100
10-1
10-2
10-3
10-4
10-5
10-6
MOP 6535a
Revision 4
January 2013
Page 7 of 8
Page _____ of ______
Sample ID
Plate
Repl.
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
NOTES:
100
10-1
10-2
10-3
10-4
10-5
10-6
MOP 6535a
Revision 4
January 2013
Page 8 of 8
Page _____ of ______
Sample ID
Plate
Repl.
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
A
B
C
NOTES:
100
10-1
10-2
10-3
10-4
10-5
10-6
MOP 6555
Revision 2
November 2012
Page 1 of 4
Miscellaneous Operating Procedure (MOP) 6555:
Petri Dish Media Inoculation Using Beads
Prepared by: __________________________________________
Date: 11/15/2012
Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader
Reviewed by: __________________________________________
Dahman Touati, ARCADIS Project Manager
Date: 11/15/2012
Approved by: __________________________________________
Worth Calfee, EPA Work Assignment Manager
Date: 11/15/2012
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
MOP 6555
Revision 2
November 2012
Page 2 of 4
MOP 6555
TITLE:
PETRI DISH MEDIA INOCULATION USING BEADS
SCOPE:
This MOP outlines the procedure for cleaning, assembling, and using beads to
inoculate agar plates.
PURPOSE:
To provide an easily repeatable method for spreading liquid inoculation onto
agar plates.
Equipment:

#13 test tubes

6 x 12 test tube racks (which hold 72 tubes)

Beads of various sizes (glass)

Glass autoclavable trays (stainless steel is eventually corroded by the bleach and
autoclaving processes)

Bleach

DI water

Hot gloves

Amber bottle for collecting hazardous waste (with hazardous waste label)

Funnel

Aluminum foil

Label tape

Chemical hood

Autoclave

Oven

Labeled bead container or cup (All mold and bacteria beads must be kept separately)
1.0
CLEANING BEADS
1. When a sufficient number of beads have been collected, or at least once a day when
beads are being used to spread colonies, place the used beads into a tray with a solution
containing a 1:5 ratio of bleach to deionized water.
Add the bleach to the beads first, under the protection of a chemical safety cabinet. Then
MOP 6555
Revision 2
November 2012
Page 3 of 4
add the deionized water. Cover the pan with aluminum foil and label it with the
contents (for example: “bacteria beads in 1:5 bleach to DI water solution”). Soak the
beads 12-24 hours (usually overnight) in a chemical fume hood.
2. After soaking, take a bottle brush and thoroughly scrub the beads.
3. Decant the bleach solution (collect the bleach for proper disposal) and rinse with
deionized water 6 to 8 times, collecting the rinsate after the first rinse for disposal
(subsequent rinses can be discarded in the lab sink). Rinse until the decanted liquid is
clear. Use a funnel to add the bleach waste to a labeled amber waste bottle. These
liquids must be labeled “hazardous waste” and can then be stored, collected or disposed
of properly.
4. Cover the beads with deionized water and autoclave for 1 hour on the liquid cycle.
5. Decant the deionized water and place the tray of beads in the Thelco lab oven at 121 C
until dry (a minimum of 3 hours).
6. Remove the beads from the oven using proper safety equipment (heat gloves) and cover
with clean aluminum foil to prevent contamination. Label each tray with the following
information:
“Clean bacteria (or mold) beads,” the date beads were cleaned, initials of the person
who cleaned them.
7. These beads are then ready for use as described in “PLACING BEADS IN TUBES”.
3.0
PLACING BEADS IN TUBES
1. Fill a 6 x 12 rack with tubes
2. Place clean beads into a shallow pan, and then manually fill each tube with 7-15
beads/tube.
Note: Beads vary in size and will therefore fill the tubes to different heights.
3. Tightly attach a cap to each tube
4. Autoclave for 1 hour using a gravity sterilization cycle (see MOP 6570). Autoclave tape
must be placed on the top of each rack to provide evidence that the beads have been
sterilized
MOP 6555
Revision 2
November 2012
Page 4 of 4
4.0
SPREADING BEADS
1.
To spread inoculum on the agar surface, one tube of beads should be used for each
individual plate.
2.
After the beads have been added, the plates can be stacked up to six plates high. The
plates are then shaken 10 times from side-to-side. Turn the stack of plates ¼ turn, and
again shake 10 times from side-to-side. Repeat this procedure (¼ turn and 10 shakes) two
more times, so that the beads are shaken a total of forty times.
3.
Turn the plates over (upside down), and tap the beads into the lid.
4.
Aseptically dump the beads into a labeled bead container (mold and bacteria beads must
be labeled and collected separately), which should be considered contaminated one plate
at a time, replacing the lid as quickly as possible to prevent contamination.
References
1.
http://serc.carleton.edu/microbelife/k12/LIMW/dilution.html
2. http://www.qbiogene.com/businessdivisions/platformnews/news0005-b.shtml
3. http://www.genlantis.com/objects/catalog/Product/extras/C400050.pdf
4. EPA - SHEM Chemical Hygiene Plan
MOP 6562
Revision 1
February 2013
Page 1 of 6
Miscellaneous Operating Procedure (MOP) 6562:
Preparing Pre-Measured Tubes with Aliquoted Amounts of
Phosphate Buffered Saline with Tween 20 (PBST)
Prepared by: __________________________________________
Date: 2/12/2013
Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader
Reviewed by: __________________________________________
Dahman Touati, ARCADIS Project Manager
Date: 2/12/2013
Approved by: __________________________________________
Worth Calfee, EPA Work Assignment Manager
Date: 2/12/2013
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
MOP 6562
Revision 1
February 2013
Page 2 of 6
MOP 6562
TITLE:
PREPARING PRE-MEASURED TUBES WITH ALIQUOTED AMOUNTS
OF PHOSPHATE BUFFERED SALINE WITH TWEEN 20 (PBST)
SCOPE:
This MOP provides the procedure for preparing PBST.
PURPOSE:
This procedure will ensure that that the PBST is prepared correctly and that all
measured tubes are filled aseptically.
1.0
PREPARING STERILE PHOSPHATE BUFFERED SALINE WITH TWEEN 20
(PBST)
Phosphate Buffered Saline with Tween 20 (PBST) is prepared 1 L at a time in a 1 L flask.
1. Add 1 packet of SIGMA Phosphate Buffered Saline with Tween 20 (P-3563) to 1 L of
deionized (DI) water.
2. Shake vigorously to mix until dissolved.
3. Label bottle as “non-sterile PBST” and include date and initials of person who made
PBST.
4. Filter sterilize into two 500 mL reagent bottles using 150 ml bottle top filter (w/ 33mm
neck and .22 µm cellulose acetate filter) for sterilization. Complete this by pouring the
liquid into the non-sterile PBST into the top portion of the filtration unit 150 ml at a time,
while using the vacuum to suck the liquid through the filter. Continue to do this until 500
ml have been sterilized into a 500 ml bottle. Change bottle top filter units between each
and every 500 ml bottle.
5. Change label to reflect that the PBST is now sterile. Include initials and date of
sterilization. The label should now include information on when the PBST was initially
made and when it was sterilized and by whom.
6. Each batch of PBST should be used within 90 days.
2.0
PREPARING 20 ML/5 ML PBST TUBES FOR USE DURING
EXPERIMENTATION
Twenty (20) ml or five (5) ml of the prepared PBST will be added to each sterile 50-ml
conical tube as detailed below. Each flat of conical tubes contains 25 tubes, so one 500 ml
sterile bottle of PBST should fill approximately one flat when 20 ml tubes are needed and
four flats when 5 ml tubes are needed.
MOP 6562
Revision 1
February 2013
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1. Prepare the hood by wiping down with ethanol, followed by bleach, followed by DI water
and a clean Kimwipe or Techwipe. Then stock the hood with the following items if they
are not already there:
-
The flats of sterile conical tubes you need to fill with PBST.
Sufficient bottles of sterile PBST to fill these tubes.
Ample 25 ml serological pipettes (at least 3 per flat) for 20 ml transfers and 10
ml serological pipettes for the 5 ml transfers.
Serological pipetter (automatic, hand-held pipette).
Burner and striker.
2. Light the burner and adjust the flame for a width adequate to flame the lips of the PBST
bottles.
3. Take one flat of sterile conical tubes and loosen each cap on the outside edges (about ½
turn).
4. Open a serological pipette and insert into the serological pipetter, taking care to not touch
the tip to any surface.
5. Hold the pipetter with the first three fingers of your right (or dominant) hand. With your
left hand (or non-dominant hand), pick up a bottle of the PBST and use the bottom of
your right hand to unscrew the lid. Place the lid upside down on the benchtop and
quickly flame the lip of the bottle. Turn the bottle and repeat, taking care to thoroughly
flame the lip without getting the glass so hot that it shatters.
6. Inset the tip of the pipette into the bottle and fill to the 20 ml line. Flame the bottle lip
and place the bottle on the benchtop.
NOTE: If the tip of the pipette touches the outside of the bottle or any other
surface in the hood, consider it contaminated. Discard the pipette
and reload a new one.
7. Quickly pick up one of the tubes that you have loosened the cap on, and use the bottom
of your right hand to remove the cap. Completely discharge the entire pipette into the
tube, taking care to not touch anything with the tip of the pipette. Recap the tube and
place back into the flat (the lid does not have to be tight – you will tighten the lids after
you have completed filling the 10 outside tubes).
NOTE: If the tip touches the outside or rim of the tube (or any other surface
in the hood), consider the tube and pipette contaminated. Discard
both the tube and the pipette.
MOP 6562
Revision 1
February 2013
Page 4 of 6
8. Pick up the PBST bottle and flame the lip. Repeat Steps 6 and 7 until all 10 of the tubes
on the outside of the flat have been filled. Flame the lip of the PBST bottle and replace
the cap. Slide the used pipette back into the plastic sleeve and put to the side of the hood
for disposal. Then tighten the lid of each tube you just filled. But rather than placing it
back into its original spot in the flat, switch it for the empty tube from the next row.
When this has been completed, go around the outside of the flat again and loosen the lids
of these 10 tubes. Repeat steps 4 through 7 to fill and cap these tubes.
9.
This same procedure is used to fill the middle row of tubes from the flat, and if more
than one flat of tubes is being filled, can be done at the same time as the outside rows of a
second flat.
10. When all tubes have been filled, label each flat as follows, and place on the shelf in room
E390B:
“PBST Tubes (20 ml or 5 ml)”
Date prepared
Your initials
11. These tubes should be made at least 14 days before they need to be used so that they can
be verified as sterile. Any tubes that are cloudy or that have any floating matter/turbidity
should be discarded. The tubes are stable for and should be used within 90 days.
3.0
CLEANUP FOR 20 ML/5 ML PBST TUBES
1. Dispose of the used pipettes in the nonregulated waste.
2. Plug in the serological pipetter so that it can recharge.
3. Replace any unused PBST in the liquid containment on the shelf. Make sure that the
bottle is labeled as having been opened (date opened and initials of whomever used it).
4. Turn off the burner.
5. Wipe down the hood benchtop with ethanol, followed by bleach, followed by DI water
and a clean Kimwipe or TechWipe.
4.0
PREPARING 900µL PBST TUBES FOR USE DURING EXPERIMENTATION
1. Prepare the hood by wiping down with ethanol, followed by bleach, followed by DI water
and a clean Kimwipe or Techwipe. Then stock the hood with the following items if they
are not already there:
MOP 6562
Revision 1
February 2013
Page 5 of 6
-
-
A sterile beaker of microcentrifuge tubes.
Sufficient tubes of sterile PBST to fill these tubes (PBST may be aseptically
transferred to 50 ml conical tubes for an easier aseptic transfer to the
microcentrifuge tubes- it is easier than working from a 500 ml reagent bottle.
Make certain that these 50 ml conical tubes are labeled to when the PBST was
made, sterilized, etc.).
1000 µL micropipette.
1000 µL sterile pipette tips
Microcentrifuge tube racks.
Labeled beaker or waste container used to hold non-regulated waste, such as
tips, under the hood.
2. Carefully remove the microcentrifuge tubes one at a time from the beaker and close the
top on each one before placing it in the tube rack. Place the tubes in the rack skipping
every other row. Fill up two racks doing this.
3. Add 900 µL of PBST to the microcentrifuge tubes by aseptically transferring the PBST
from the sterile 50 ml conical tube containing the PBST. Do this by using the 1000 µL
micropitte and tips. Change tips whenever after two rows of tubes are completed or
whenever a contamination event (such as touching the outside of the 50 ml tube or the
microcentrifuge tube) occurs. Put the dirty tips in the beaker or container used to contain
waste (tips, tubes) in the hood. If any 900 µL tubes are contaminated during the transfer,
dispose of them in the waste container used to hold tips under the hood. If a new box of
tips has to be opened, make certain the date it was opened and initials of the person who
opened it are clearly labeled on the box.
4. After both racks are full, carefully move all the tubes from one rack to fill in the empty
rows on the other rack. In this manner, one rack should be completely filled with tubes at
this point.
5. Label the rack of tubes as “Sterile 900 µL PBST Tubes”, along with the name of the
person who completed the transfer, along with the date. Also, include the date that the
original stock of PBST was made and the date it was sterilized, along with the initials of
the person who completed those steps.
5.0 CLEANUP FOR 900µL PBST TUBES
1. Dispose of the waste that was put in the labeled beaker or waste container (micropipette
tips and tubes) in the nonregulated waste. Then, place this beaker in the “To be
decontaminated via sterilization- contaminated glassware” bin or if it is a disposable
container, then it can be put in the non-regulated waste container.
2. Put the unused sterile tips and the micropipetter back in its original location.
MOP 6562
Revision 1
February 2013
Page 6 of 6
3. Replace any unused 50 ml conicals of PBST in the liquid containment on the shelf.
Make sure that the tube is labeled as having been opened (date opened and initials of
whomever used it). If the tube could possibly be contaminated in any way, dispose of it
in non-regulated waste.
4. Wipe down the hood benchtop with ethanol, followed by bleach, followed by DI water
and a clean Kimwipe or TechWipe.
MOP 6563
Revision 2
November 2012
Page 1 of 6
Miscellaneous Operating Procedure (MOP) 6563:
Swab Streak Sampling and Analysis
Prepared by: __________________________________________
Date: 11/15/2012
Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader
Reviewed by: __________________________________________
Dahman Touati, ARCADIS Project Manager
Date: 11/15/2012
Approved by: __________________________________________
Worth Calfee, EPA Work Assignment Manager
Date: 11/15/2012
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
MOP 6563
Revision 2
November 2012
Page 2 of 6
MOP 6563
TITLE:
SWAB STREAK SAMPLING AND ANALYSIS
SCOPE:
This MOP provides the procedure for the process of completing a swab streak
plate and subsequent qualitative analysis.
PURPOSE:
This procedure will ensure that the swab streak plate sampling and analysis
methods are standardized and that the collection and plating of samples are
free from contamination.
1.0
PREPARING THE MATERIALS
There are two types of prepared swabs that can be used in this procedure:
Environmental Transport Swabs – purchased swabs that are individually packaged
and pre-sterilized.
In-house Sterilized Swabs – swabs placed into autoclave pouches and sterilized
using a 1-hour gravity cycle.
This procedure requires the following materials and equipment:

Tryptic soy agar (TSA) media plates

32 C incubator

Nitrile (non-sterile) gloves

Sharpie for writing on plates
2.0
COLLECTING AND PLATING SAMPLES
The procedure for collecting and plating samples is dependent on the type of swab being
used. Appropriate PPE should be worn in both cases and includes a lab coat, nitrile gloves,
and safety glasses.
MOP 6563
Revision 2
November 2012
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2.1
Environmental Transport Swabs
2.1.1 Collection of Environmental Transport Swab Sample
1. Break the seal on the individually packaged and sterile swab. Collect the specimen with
the swab applicator as detailed in the specific test protocol, then replace the swab in the
tube.
2. Label the tube with what is being swabbed (sample ID), the date, time, and initials of the
person performing the procedure.
3. Place the swab into a secondary container, such as a sterile bag, and label the bag with
the same information placed on the tube label.
4. Transport the sample(s) to the Microbiology Laboratory for processing.
2.1.2 Plating of Environmental Transport Swab Sample
1. When the sample is received in the Microbiology Laboratory, label one TSA plate using
a Sharpie with the information from the swab packaging. Verify that the sample ID and
date match.
2. Place labeled plates and swab samples under the biological safety cabinet. Remove the
sample swab from the secondary container and the tube. Press onto the plate in an Sstroke motion, turning the swab as it is plated to ensure that all of the surface area of the
swab touches the plate. Press firmly, but not so hard that the surface of the media is
broken.
3. Replace the swab into its tube and discard in the non-regulated waste container.
4. Repeat steps #1 through #4 for each sample.
5. Label three TSA plates as Swab Blank A, Swab Blank B, and Swab Blank C. These
plates will serve as negative controls for the swabs.
6. Open a new/unused Environmental Transport Swab and use it to plate the three blank
plates as detailed in Step #2.
7. Stack the triplicate plates media side up and place in a 35 C ± 2 C incubator for at least
18 hours. Note the time the plates were placed in the incubator.
MOP 6563
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November 2012
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2.2
In-house Sterilized Swabs
When In-house Sterilized Swabs are being used to collect samples, they need to be plated
immediately (unlike the Environmental Transport Swabs which are transported back to the
Microbiology Laboratory for plating). Therefore, prior to travelling to the sample site, collect
the following materials and supplies which will be needed:

One TSA media plate (in a media bag) per sample to be collected plus three additional
plates to be used as negative controls for swab blanks.

One In-house Sterilized Swab (in their autoclave pouches) per sample to be collected, one
swab for the control plates, plus a few extras.

Sharpie for labeling plates.
Use the following procedure to collect and plate samples.
1. Once at the sample collection site, take the TSA plates out of the media bag and label one
plate for each sample with what is being swabbed (sample ID), date, time, and initials of
the person performing the procedure.
2. As carefully and as aseptically as possible, remove the swab from the autoclave pouch by
the stick end. Be sure and not touch the swab end to anything but the sample. If the
swab’s sterility is compromised, dispose of the swab and use one of the extras.
3. Collect the specimen with the swab applicator as detailed in the specific test protocol.
4. Press onto the plate in an S-stroke motion, turning the swab as it is plated to ensure that
all of the surface area of the swab touches the plate. Press firmly, but not so hard that the
surface of the media is broken. Because these samples are being plated in the open air
and not in a biological safety cabinet, be certain to limit the time that the lid is removed
from the TSA plate.
5. Replace the swab into the autoclave pouch it came in and discard in the non-regulated
waste container.
6. Repeat steps #1 through #6 for each sample.
7. Label three TSA plates as Swab Blank A, Swab Blank B, and Swab Blank C. These
plates will serve as controls for both the swabs and the TSA.
8. Open another in-house sterilized swab from the autoclave pouch and use it to plate the
three blank plates as detailed in Step #2.
MOP 6563
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November 2012
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9. Put the TSA plates back into the media bag and transport to the Microbiology
Laboratory.
10. When received by the laboratory, the plates should be stacked media side up and placed
in a 35 C ± 2 C incubator for at least 18 hours. Note the time the plates were placed in
the incubator.
3.0
ANALYZING THE SAMPLES
The Swab Results Template, which follows this section, is used to record the results of the
sampling. Some quantities of samples may require more than one form. Make certain that
the data is filled in completely on each page. The analyst will use the information on the
TSA plates to fill in the following blanks at the top of the form:

Swab samples taken on: (date)

Swabbed by: (person)

Plating completed on: (date)

Plated by: (person)
The following procedure is used to analyze the samples and complete the remainder of the
Swab Results Template form.
1. Fill in the final two sections at the top of the form: Plate results read on and Results
read by.
2. Observe the agar surface on the plates and note the sample IDs on the first three lines in
the Sample column.
3. For each plate, check whether there was growth (G) or no growth (NG). Growth is
indicative of an organism(s) being present, and should be described on the form. Be as
detailed as possible, noting colony morphology (size, shape, color and any other
distinctive things that can be seen concerning the growth).
4. The Swab Results Template form serves as the sample report and should be provided to
the Project Manager.
MOP 6563
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November 2012
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Swab Results Template
Swab samples taken
on:
Plating completed on:
Plate results read on:
Sample Name
Controls
Swab blank A
Swab blank B
Swab blank C
Result
G  NG 
G  NG 
G  NG 
G  NG 
G  NG 
G  NG 
G  NG 
G  NG 
G  NG 
G  NG 
G  NG 
G  NG 
G  NG 
G  NG 
G  NG 
G  NG 
G  NG 
G  NG 
G  NG 
Result
G  NG 
G  NG 
G  NG 
Swabbed by:
Plated by:
Results read
by:
If growth, describe
If growth, describe
Key
G = Growth.
NG = No Growth.
All plates are plated in triplicate resulting in sample identification of “A”, “B”, and “C”.
MOP 6565
Revision 4
November 2012
Page 1 of 4
Miscellaneous Operating Procedure (MOP) 6565:
Filtration and Plating of Bacteria from Liquid Extracts
Prepared by: __________________________________________
Date: 11/15/2012
Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader
Reviewed by: __________________________________________
Dahman Touati, ARCADIS Project Manager
Date: 11/15/2012
Approved by: __________________________________________
Worth Calfee, EPA Work Assignment Manager
Date: 11/15/2012
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
MOP 6565
Revision 4
November 2012
Page 2 of 4
MOP 6565
TITLE:
FILTRATION AND PLATING OF BACTERIA FROM LIQUID
EXTRACTS
SCOPE:
This MOP outlines the procedure for filtration and subsequent cultivation
of bacterial spores from a liquid extract.
PRUPOSE:
This method is deployed when results from spread-plate methods yield
less than 30 colony-forming units (CFU) per plate. This method allows a
lower limit of detection for bacterial recovery/survivorship assays. This
method can also be used to analyze liquid samples such as decon rinsates.
Materials:

Petri dishes with appropriate agar

0.2 µm pore-size disposable analytical filter units (2-3 per sample)

P1000 pipette and sterile tips

Sterile forceps

Pipettman and sterile serological pipettes
1.0
PROCEDURE
1. For each liquid sample to be analyzed, gather the required number of disposable
analytical filter units and Petri dishes containing the desired sterilized/QC’d media.
NOTE #1: For analysis of 5 to 30 ml extracts, 1 ml and remainder should be
filtered; for 31 to 200 ml samples, 1 ml, 10 ml, and remainder should
be filtered; for samples over 200 ml, more filter samples may be
needed.
NOTE #2: For previously plated samples where 10 – 19 CFU were observed,
replating using a 400 µl inoculum, and plates where 20 – 29 CFU
were observed, replating using a 200 µl inoculum can be executed
rather than filter plating. For inoculua greater than 200 µl, a sterile
spreader should be used rather than the bead method).
MOP 6565
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November 2012
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2. Label plates.
3. Vortex liquid extract vigorously for 2 minutes, using 10 second bursts. (for larger
volume samples, a vigorous mixing by shaking of the sample container can be
substituted for vortex mixing)
4. Using a P1000, sterile tip, and aseptic techniques, immediately following vortexing,
pipette 1 ml of the extract into one of the filter units.
5. Apply vacuum to the filter unit, to pull the liquid through the filter and collect the
spores on the surface of the filter.
6. Using a sterile serological pipette, rinse the filter unit by pipetting 10 ml of sterile
deionized water along the inner sides of the unit while it is under vacuum.
7. Aseptically remove the filter from the filter apparatus using sterile forceps, and lay
the filter onto the agar surface within the Petri dish (spore side up).
8. Vortex the liquid extract vigorously for 10 seconds.
9. Use the appropriate volume serological pipette to transfer the remaining aliquots into
their respective filtration units (one at a time).
10. Repeat steps 5 through 7 taking time to vortex or mix the sample 10 seconds
immediately before removing an aliquot.
Important: Be sure to note and record the volume of the “remainder” sample.
11. Incubate all plates at the optimal growth temperature for the organism used for 16 –
28 hours.
12. Enumerate and record the number of CFU on each plate.
2.0
DATA CALCULATIONS
Utilize the following equation to determine the total abundance of recovered spores:
N  CFU 
V Extract
V filtered
MOP 6565
Revision 4
November 2012
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Where, N is the total number of spores recovered in the extract, CFU is the abundance of
colonies on the agar plate, VExtract is the total volume of the extract (before any aliquots
were removed), VFiltered is the volume of the extract filtered.
MOP-6567
Revision 1
November 2012
Page 1 of 4
Miscellaneous Operating Procedure (MOP) 6567:
Recovery of Bacillus Spores from Wipe Samples
Prepared by:
__________________________________________
Date: 11/15/2012
Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader
Reviewed by:
__________________________________________
Dahman Touati, ARCADIS Project Manager
Date: 11/15/2012
Approved by:
__________________________________________
Worth Calfee, EPA Work Assignment Manager
Date: 11/15/2012
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
MOP-6567
Revision 1
November 2012
Page 2 of 4
MOP 6567
TITLE:
RECOVERY OF BACILLUS SPORES FROM WIPE SAMPLES
SCOPE:
This MOP outlines the procedure for recovering Bacillus spores from wipe
samples.
PURPOSE:
To aseptically extract and quantify Bacillus spores from wipe samples in order to
determine viability and obtain quantifiable data.
Materials:

PPE (gloves, lab coat, safety goggles)

Biological Safety Cabinet (Class II)

pH-amended bleach

Deionized water

70% solution of denatured ethanol

Kimwipes

Disbatch® bleach wipes

Non-regulated waste container

50 mL sterile conical tubes containing 20 mL of sterile phosphate buffered saline with Tween
20 solution (PBST) (MOP 6562)

Vortex mixer

Cart

Wire or foam rack for 50mL conical tubes

Tryptic soy agar plates

900 µL tubes of sterile PBST

Pipettor and pipette tips for dilutions

Incubator set to appropriate growth temperature for target organism (35°C or 55°C)

Light box for counting colonies

Lab notebook

QAPP for project that is utilizing the wipe samples
MOP-6567
Revision 1
November 2012
Page 3 of 4
1.0
PROCEDURE
1. Begin by donning PPE (gloves, lab coat, and protective eyewear).
2. Obtain wipe samples that may contain Bacillus spores. Wipe samples should be received as
one wipe/sponge in a sterile 50 mL conical tube delivered in secondary containment. Make
certain that all of the samples are labeled. Review any chain of custody forms that may
accompany the samples to ensure that all of the labels are consistent and that there is no notable
variation in the samples. If variation has occurred, make a note of it in the notebook.
3. Clean the workspace (biological safety cabinet) by wiping surfaces with pH-amended bleach,
next with deionized water, and lastly with a 70-90 % solution of denatured ethanol. Wipe with
a Kimwipe to remove any excess liquid. Make sure the workspace is clean and free of debris.
Gather all necessary items to perform the task, place these items on a clean cart beside the
biological safety cabinet, within arm’s reach so that, once the procedure has begun, the task
may be performed without interruptions.
4. Discard gloves and replace with fresh pair.
5. One at a time, under the biological safety cabinet, remove the sample tube containing the wipe
sample from the secondary containment bag in which it arrived. Using the Dispatch® bleach
wipes, wipe each sample tube with one wipe, and then wipe it with a clean Kimwipe. Discard
the used bleach wipe and the used Kimwipe in the secondary containment bag and place them
in the non-regulated waste container. Remove gloves and don a fresh pair of gloves. Repeat
this procedure for every sample. After each sample has been cleaned, place the tubes
containing the wipe samples in an appropriate sized wire or foam rack to hold the tubes in an
upright, vertical position.
6. Leaving the tubes in the rack underneath the biological safety cabinet, aseptically add 20 mL of
PBST solution (this should be in a pre-measured, sterile conical tube, per MOP 6562) to each
sample tube containing a wipe, one a time. Remove the rack containing wipe samples from
hood when all samples have had the PBST added. Place the rack with the samples on the cart.
7. Using the procedure to clean the biological safety cabinet, as found in Step 3, clean the
biological safety cabinet again. Afterwards don a fresh pair of gloves.
8. Using a vortex mixer, agitate the wipe samples, four at a time, in a biological safety cabinet, for
ten second bursts for two minutes total. Make certain to clean the biological safety cabinet
after each set of four samples and change gloves between each set of samples.
NOTE: The reason that four samples are done at one time is to limit the time between
agitation and plating. The samples need to be processed immediately after
MOP-6567
Revision 1
November 2012
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agitation, and agitation of more than four samples at a time leaves too much
time between agitation and spread plating.
9. Using tryptic soy agar media plates that are appropriately labeled with the sample number,
dilution set and date, complete dilution plating for the wipe samples immediately after the two
minute agitation step is completed. The samples should also be agitated again for ten seconds
directly prior to removing an aliquot from the sample tube. Each dilution tube should also be
agitated for ten seconds prior to removal of aliquots. Dilutions should be completed using the
techniques and methodology as described in MOP 6535a, and the 900 µL tubes should be
made with sterile PBST to stay consistent with materials/solutions. Plating in this manner
should be repeated for all samples, with any changes in protocol noted in the lab notebook.
10. Once the dilution plating has been completed, the plates are to be placed in an incubator. For
non-thermophilic Bacillus species, the plates should be placed at 35°C +/- 2° C for 12-24
hours. For thermophilc Bacillus species, such as Geobacillus stearothermophilis, the plates
should be incubated at 55°C ±2 °C for 12-24 hours. The target Bacillus organism that will be
used for the wipe samples will be specific to the project and noted in the QAPP.
11. After the plates have incubated for a sufficient amount of time (12-24 hours) and the growth
from any Bacillus colonies are quantifiable, the colonies should be manually counted using the
light box and the data should be properly recorded as dictated per project by the QAPP. All
results will be checked for quality assurance and all data will be reported to the proper
personnel as listed in the QAPP.
MOP 6570
Revision 3
March 2013
Page 1 of 4
Miscellaneous Operating Procedure (MOP) 6570:
Use of Steris Amsco Century SV 120 Scientific Prevacuum
Sterilizer
Prepared by: __________________________________________
Date: 3/21/2013
Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader
Reviewed by: __________________________________________
Dahman Touati, ARCADIS Project Manager
Date: 3/21/2013
Approved by: __________________________________________
Worth Calfee, EPA Work Assignment Manager
Date: 3/21/2013
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
MOP 6570
Revision 3
March 2013
Page 2 of 4
MOP 6570
TITLE:
USE OF STERIS AMSCO CENTURY SV 120 SCIENTIFIC
PREVACUUM STERILIZER
SCOPE:
Basic instructions for use of the large Steris autoclave.
PURPOSE:
To outline proper procedural use of the autoclave, using preprogrammed
cycles, to effectively sterilize items, while complying with quality control
standards.
Materials:

Amsco Century SV 120 Scientific Prevacuum Sterilizer

Items to be sterilized (liquids, solids, waste, etc)

Pouches to contain materials during sterilization and maintain sterility until use

Aluminum foil

Autoclave indicator tape

Sterilization verification ampoules (such as Raven ProSpore Ampoules)

Thermally resistant gloves

De-Ionized (DI) water
1.0
PROCEDURE
1.1
Start Up
1. Turn on the autoclave. The power switch is located behind the door in the top right
corner. The digital touch screen on the front of the unit will power up and indicate
that a memory test is in progress.
2. After the memory test is complete, the device will request that it be flushed. This
should be conducted daily to minimize scaling inside the boiler. The flush valve is
located behind the door on the bottom, left of the device (yellow handle). Move the
valve to the open position and then press the “Start Timer” button on the touch
screen. The flush will run for 5 minutes and will alert at completion with a single
chime.
3. Once the flush is complete, close the flush valve and press the “Continue” button on
the touch screen. The screen should then return to its default menu which has 2
choices “Cycle Menu” and “Options”
MOP 6570
Revision 3
March 2013
Page 3 of 4
1.2
Basic Operation
1. Prepare any items that need to be sterilized. The items must be carefully wrapped or
sealed in sterilization pouches in order to maintain sterility when removed from the
autoclave. Examples of this include: wrapping any orifices with aluminum foil,
placing whole items in autoclave pouches, loosely applying a cap on a bottle (to allow
for the pressure changes inside).
2. Once prepared, each item should be outfitted with a sterility indicator such as a small
piece of autoclave indicator tape; or by utilizing an autoclave pouch with a built-in
sterility indicator strip. These indicators provide a visual verification that the
sterilizing temperature (121°C) was reached.
3. To add items to the autoclave, open the autoclave door by pressing down on the foot
pedal on the bottom right corner on the front of the device.
4. Place items that need to be sterilized into the autoclave, adding or moving racks to
accommodate the load. If liquids are being autoclaved, then they must have secondary
containment (usually a large plastic autoclave-safe tray) to contain any fluids in the
event of a leak, spill or boil-over. Add an indicator ampoule to the first autoclave
cycle of the day, regardless of the type of cycle.
5. Once the autoclave is loaded, press the foot pedal to close the autoclave door.
6. Once the door is sealed, a menu of the cycles can be seen by pressing the button on
the touch screen labeled “Cycle Menu”. Then choose the appropriate cycle by
touching the corresponding button. If the cycle chosen is the one desired for the
sterilization process, press the “Start Cycle” button. Otherwise, press “Back” to
return to the prior menu screen.
7. After the cycle has started, the type of cycle, the number of the cycle, the items
placed in the autoclave during the cycle, the time, whether or not an indicator
ampuole was included in the load, and the initials of the person starting the cycle
must be recorded in the autoclave log book, located in the drawer across form the unit
labeled “Autoclave Supplies.”
8. Quality control (QC) indicator ampoules, usually Raven ProSpore Ampoules with
Geobacillus stearothermophilus (at a concentration 10E6), are added to one cycle
each day to ensure that the autoclave is functioning properly. These ampoules are
used according to manufacturer’s instructions. These ampoules must be properly
labeled with the date in which they were autoclaved and the initials of the individual
that completed the cycle. At the beginning of each week, a positive control ampoule
must be processed, where the ampoule is placed directly into the 55°C water bath,
without being autoclaved. The positive control indicator ampoule should change
from purple to yellow in color, indicating growth. All test ampoules should be placed
in a water bath following the end of the cycle in which they are run. These ampoules
should not change color (from purple to yellow, but instead should remain a purple to
MOP 6570
Revision 3
March 2013
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purple-brown color). Ampoules should be checked at both 24 and 48 hour intervals
for growth and then finally recorded and disposed of after 48 hours. All QC
information concerning ampoules should be recorded in the autoclave notebook.
9. Upon completion of any cycle, the autoclave will alarm with a repeating beep for
approximately one minute. Any time after this alarm starts, it is safe to open the main
door (take caution because the steam escaping the chamber will be very hot when the
door is opened). The contents from the autoclave will be very hot; use protection to
remove items from the autoclave (thermally resistant gloves).
10. Place the contents of the autoclave in an appropriate place to cool, and close the
autoclave door using the foot pedal.
1.3
1.3.1
Cycles
Gravity Cycles
Gravity cycles are used to sterilize glassware and other utensils, which are not submerged
in nor contain any volume of liquid. These cycles are typically used for “dry” materials.
Currently there are two different gravity cycles programmed for daily operations: a 1hour cycle and a 30-minute cycle. The time that the chamber is held at the sterilization
temperature (121 °C) is the only difference between these two cycles. The different
sterilization times allow for the compensation of the various sizes of materials and more
resilient organisms. The 30-minute cycle is primarily used for a small quantity of
material. The 1 hour cycle is used for large loads or items containing a large amount of
contamination. The 1 hour cycle is recommended for inactivation of gram positive sporeforming bacteria.
1.3.2
Liquid Cycles
Liquid cycles are used to sterilize a variety of liquids and solutions. The solutions are
typically mixed prior to sterilization. It is important to have secondary containment to
contain any fluids in the event of a leak, spill or boil-over. The 30-minute liquid cycle is
used to sterilize small volumes of liquid (usually less than 2L total). When attempting to
sterilize any volume larger than 2L, the 1-hour liquid cycle should be used to ensure
complete sterilization. The 1-hour liquid cycle is the preferential cycle used as the
destruction cycle for waste. In the event of materials (liquid or otherwise) being
contaminated/exposed microorganisms, the 1-hour liquid cycle will be used as the initial
means of decontamination. When completing a decontamination cycle, if there is no
liquid inside of a container, then deionized water must be added to the container or the
item must be submerge prior to the start of the cycle. Only items that are being
decontaminated can go in destruction cycles. Decontamination cycles cannot be mixed
with sterilization cycles.
MOP 6572
Revision 2
November 2012
Page 1 of 4
Miscellaneous Operating Procedure (MOP) 6572:
Recovery of Spores from Vacuum Sock Samples
Prepared by:
__________________________________________
Date: 11/15/2012
Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader
Reviewed by:
__________________________________________
Dahman Touati, ARCADIS Project Manager
Date: 11/15/2012
Approved by:
__________________________________________
Worth Calfee, EPA Work Assignment Manager
Date: 11/15/2012
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
MOP 6572
Revision 2
November 2012
Page 2 of 4
MOP 6572
TITLE:
RECOVERY OF SPORES FROM VACUUM SOCK SAMPLES
SCOPE:
This MOP outlines the procedure for recovering spores from vacuum sock samples
PURPOSE:
To aseptically extract and quantify spores from vacuum sock samples in order to
determine viability and obtain quantifiable data
MATERIALS

PPE (gloves, lab coat, safety goggles)

Biological Safety Cabinet (Class II)

pH-amended bleach

Deionized water

70% solution of denatured ethanol

Kimwipes

Disbatch® bleach wipes

Non-regulated waste container

3 oz. sterile specimen cup containing 20 mL of sterile phosphate buffered saline with Tween 20
solution (PBST) (MOP 6562)

Sterile scissors

Vortex mixer

Cart

Tryptic soy agar plates

900 µL tubes of sterile PBST

Pipettor and pipette tips for dilutions

Incubator set to appropriate growth temperature for target organism (35 °C or 55 °C)

Light box for counting colonies

Lab notebook

QAPP for project that is utilizing the vacuum sock samples
MOP 6572
Revision 2
November 2012
Page 3 of 4
1.0
PROCEDURE
1. Begin by donning PPE (gloves, lab coat, and protective eyewear).
2. Obtain vacuum sock samples that may contain Bacillus spores. Vacuum sock samples should
be received as one vacuum sock in a sterile 5.5’ x 9 bag secondarily contained in a 10’ x 15’
bag. Make certain that all of the samples are labeled. Review any chain of custody forms that
may accompany the samples to ensure that all of the labels are consistent and that there is no
notable variation in the samples. If variation has occurred, make a note of it in the notebook.
3. Clean the workspace (biological safety cabinet) by wiping surfaces with pH-amended bleach,
next with deionized water, and lastly with a 70-90 % solution of denatured ethanol. Wipe with
a Kimwipe to remove any excess liquid. Make sure the workspace is clean and free of debris.
Gather all necessary items to perform the task, place these items on a clean cart beside the
biological safety cabinet, within arm’s reach so that once the procedure has begun the task may
be performed without interruptions.
4. Discard gloves and replace with fresh pair.
5. Label a 3 oz. specimen cup to match the vacuum sock sample ID. The specimen cup contains
20 mL of sterile PBST.
6. When extracting samples, handle one sample at a time from start to finish. Begin by removing
the inner bag from the outer bag. Discard the outer bag in the non-regulated waste container.
Place the inner bag containing the vacuum sock under the hood. Loosen the cap on the 3 oz.
specimen cup and open a pack of sterile scissors. Open the bag and remove the sock, careful
not to touch the white part. Roll the non-sterile blue portion of the vacuum sock onto the
smaller cardboard ring. Dispose of the larger cardboard ring. Wet the vacuum sock by holding
the upper blue portion of the vacuum sock (around the smaller cardboard ring) and dipping the
lower 1-inch of the vacuum sock into the PBST. The vacuum sock will be allowed to absorb
the PBST for a few seconds. After wetting, the vacuum sock will be lifted up just above the
opening of the specimen bottle, and a 1-inch vertical slit will be cut up the center from the
bottom of the sock using sterile scissors (a new pair of scissors should be used for each
sample). The vacuum sock is then cut horizontally from side to side, about 1 inch from the
bottom allowing the two pieces to fall into the specimen bottle. The vacuum sock should be cut
only where the sock has been wetted. Repeat the dip/cutting procedure until the entire
collection portion of the sock has been excised. The upper top blue portion of the vacuum sock
will then be discarded. Place used scissors in a discard pan. After samples are all extracted,
scissors will be immediately autoclaved using a one hour gravity destruction cycle in
preparation for use with the next sample batch. Remove gloves and don a fresh pair of gloves.
Repeat the extraction procedure for every sample, while maintaining aseptic technique.
7. After cutting all vacuum sock samples, all specimen cups (up to sixty samples at a time) should
be loaded into the sample cup holder of the orbital shaker-incubator. The samples are then
MOP 6572
Revision 2
November 2012
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agitated in the shaker incubator at 300 rpm for 30 minutes at room temperature. The samples
are then removed from the shaker incubator and brought to the Biological Safety Cabinet for
dilution plating.
8. Using the procedure to clean the biological safety cabinet, as found in Step 3, clean the
biological safety cabinet again. Afterwards don a fresh pair of gloves.
9. Using tryptic soy agar media plates that are appropriately labeled with the sample number,
dilution set and date, complete dilution plating for the vacuum sock samples immediately after
the thirty minute agitation step is completed. The samples should also be agitated again for ten
seconds directly prior to removing an aliquot from the specimen cup. Each specimen cup
should also be agitated for ten seconds prior to removal of aliquots. Dilution-plating should be
carried out according to MOP 6535a. Dilution tubes used in MOP 6535a should contain
PBST to stay consistent with materials/solutions. Repeat procedure for all samples.
10. Once the dilution plating has been completed, the plates should be incubated. For nonthermophilic Bacillus species, the plates should be placed at 35°C ±2 °C for 18-24 hours. For
thermophilc Bacillus species such as Geobacillus stearothermophilis, the plates should be
incubated at 55°C ±2 °C for 18-24 hours. The target Bacillus organism that will be used for
the vacuum sock samples will be specific to the project and noted in the QAPP.
11. After the plates have incubated for a sufficient amount of time (18-24 hours) and the growth is
quantifiable, the colonies should be manually counted with the assistance of a light box. The
data should be properly recorded as dictated per project by the QAPP. All results will be
checked for quality assurance and all data will be reported to the proper personnel as listed in
the QAPP.
MOP 6579
Revision 4
April 2013
Page 1 of 6
Miscellaneous Operating Procedure (MOP) 6579:
Recovery of Bacillus Spores from 37MM Filter Cassettes
Prepared by: __________________________________________
Date: 4/29/2013
Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader
Reviewed by: __________________________________________
Dahman Touati, ARCADIS Project Manager
Date: 4/29/2013
Approved by: __________________________________________
Worth Calfee, EPA Work Assignment Manager
Date: 4/29/2013
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
MOP 6579
Revision 4
April 2013
Page 2 of 6
MOP:
6579
Title:
RECOVERY OF BACILLUS SPORES FROM 37MM FILTER CASSETTES
Objective:
To aseptically extract and quantify recovery of Bacillus spores from 37mm filters used to
sample porous surfaces.
MATERIALS AND EQUIPMENT
Equipment:

Biological Safety Cabinet (BSC, Class II)

Sonicating cleaner (such as Branson 8510, 40KHz)

Incubator set to appropriate growth temperature for target organism (35 °C)

Light box for counting colonies

Vortex mixer

Cassette opening tool (SKC cat# 225-8372, www.SKCshopping.com)

Specimen cup rack (cat # 5534M1, http://www.medicus-health.com)
Materials:

Personal Protective Equipment (PPE) (gloves, lab coat, safety goggles)

1:10 Bleach, pH-amended to 7.0 with acetic acid

Dispatch bleach wipes

Disposable 500 ml beaker (such as Fisher Scientific cat #14-955-111D) (for discarding used tips
and beads/spreaders)

70% Solution of denatured ethanol

Kimwipes

Waste container

Sterile Phosphate Buffered Saline with Tween 20 solution (PBST) (MOP 6562)

Tryptic Soy Agar (TSA) plates

Sterile scissors or scalpels, preferably one for each sample

Sterile 1500 µL tubes filled with 900µL of sterile PBST

15 ml centrifuge tube containing 6 ml of sterile PBST (one for each sample)

Autoclaved 2 oz polypropylene jars with lids (cat # J037, cat # L208
www.containerandpackaging.com)
MOP 6579
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April 2013
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1.0

Pipettor and pipette tips for dilutions (100 μl, 1000 μl, and 5000 μl.)

Pall Filter units 0.45 um MCE (VWR; catalog #28143-544)

Vacuum Pressure Gauge (McMaster; catalogue # 38545K16)

Disposable cell spreaders or glass beads for cell spreading

Disposable sterile thumb forceps
REMOVING FILTER AND DUST FROM CASSETTE
1. Begin by donning PPE (gloves, lab coat, and protective eyewear).
2. Each cassette should be received separately in a small sealable bag (primary containment), then
placed within secondary containment. Make certain that all of the samples are labeled. Review any
chain of custody forms that may accompany the samples to ensure that all of the labels are consistent
and that there is no notable variation in the samples. If discrepancies are found, make a note of each
one in the notebook.
3. Clean the workspace (BSC) by wiping surfaces with pH-amended bleach, next with deionized water,
and lastly with a 70-90 % solution of denatured ethanol. Wipe with a kimwipe to remove any excess
liquid. Make sure the workspace is clean and free of debris. Gather all necessary items to perform
the task.
4. Discard gloves and replace with fresh pair.
5. For each cassette sample, label one 2 oz polypropylene jar with the sample identifier.
6. One at a time, under the BSC, remove the conical tube and cassette from the primary containment
bag in which it arrived.
7. First wipe down the outside of the conical tube containing the vacuum tubing and adapter with a
disinfecting wipe, and place it into a rack. Then, aseptically add 5ml of premeasured PBST. Set the
rack to the side.
8. Remove the band from around the cassette using a sterile scalpel or sterile pair of scissors. Using
the Dispatch bleach wipes, wipe each cassette with a wipe, and then a clean kimwipe. Discard the
used bleach wipe and the used kimwipe in the secondary containment bag and place them in the nonregulated waste container. Remove gloves and don a fresh pair of gloves before proceeding.
9. With the plug in place on the back side of the filter, remove the red plug from the front filter side of
the cassette, use a pipette and aseptically dispense about 1mL μl of PBST (taken from the tube
containing 6ml PBST) into the cassette. Roll the cassette around to allow the liquid to touch all
surfaces of the inside of the cassette. If there is a large quantity of dust inside, more PBST may be
needed to dampen the dust before opening the cassette.
10. Once a visual check has been completed to assure that the dust is dampened enough to prevent its
aerosolization, use the cassette tool to pry open the top section of the cassette by carefully rotating
the cassette while using the tool to pry the edges up. Be careful to hold the cassette right side up
while prying open, so as not to spill the liquid inside.
MOP 6579
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April 2013
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11. When the cassette is open, use a pipette to aseptically rinse the inside walls of the cassette with 1 to
2 ml of PBST (additional aliquots from the same 5ml tube used previously), then remove the rinse
eluent (using the same transfer pipette) and place it in the 2 oz sterile cup labeled with the sample
identifier.
12. Use the cassette opening tool to then remove the middle section of the cassette in the same manner
as the top section, thereby allowing the filter to be removed. Using sterile forceps, aseptically
remove the filter, being careful not to pick up the support filter underneath, and place it in the 2 oz
polypropylene jar with the rinse eluent.
13. Use the remainder of the 6 ml PBST to rinse the walls of the middle and top sections of the cassette
while holding them over the open 2 oz cup. The entire 6 ml PBST, the filter and as much dust as can
be rinsed from the cassette should now be inside the 2 oz polypropylene jar.
14. Discard the cassette sections, support filter, cassette plugs, disposable pipettes and disposable thumb
forceps into the proper waste. Close the 2 oz jar tightly. Wipe outside of the jar with disinfecting
wipe and place in specimen cup rack or sample retention bin.
15. Wipe down the cassette opening tool thouroughly with a disinfecting wipe, and wipe down the
surface of the BSC as described in Step 3 above.
16. Change gloves, and repeat from Step 5 for each remaining sample set.
2.0
PROCESSING 37 MM FILTERS
1. Place the rack of conical tubes containing the 5mL of PBST with the vacuum tubing and adapters
into the sonicating bath with a weighted, waterproof, rectangular, flat surface on top of the tubes to
keep them from floating when submerged. Sonicate the conical tubes for 1 minute.
2. Remove the tubes from the sonicator and dry them with a kimwipe and wipe them with a
disinfecting wipe.
3. Vortex the conical tubes for 2 continuous minutes.
4. Aseptically transfer the 5mL of PBST from the conical tube samples to the 2oz specimen cup
containing the filter sample that corresponds with the conical tube sample. The labels on the conical
tube should match the label on the specimen cup.
5. After the aseptic transfer of extraction liquid from the conical tube to the specimen cup, using a 10
ml serological pipette, measure the amount of PBST extraction liquid that has been collected in the
2oz specimen cup. Record this number.
6. Place all 2 oz polypropylene cups containing the 37mm filters and 5 ml PBST in the specimen cup
rack, place a weighted waterproof, rectangular, flat surface on top of the jars to keep them from
floating when submerged in the sonicating bath.
7. Place the rack with the cups and the weighted surface into the bath so that they are submerged about
1 – 2 inches, leaving about 1 inch between the level of the bath water and the jar lid.
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April 2013
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NOTE: This may require one or two wire racks (test tube racks or additional cup holders) in the
bottom of the sonicating bath to achieve the correct level for the cups. Make sure the
sonicating bath is filled with water to the correct level as designated by the line on the
inside of the bath. A series of open racks is preferable to a basket for this purpose, as
the baskets provided by the bath manufacturers tend to have more solid walls with
small holes, which can restrict the sonic waves more than a wire rack.
8. When the correct level of submersion is achieved, turn on the sonicating bath (no heat) and sonicate
for 3 minutes, rotating the cup rack within the bath at the 1 min and 2 min mark (pick up, turn 90
degrees, re-submerge).
9. Remove the rack of cups from the sonicating bath and place on an absorbent pad. Remove each 2 oz
cup from the rack, wipe with a kimwipe to dry and place back inside the BSC.
3.0
FILTRATION AND PLATING OF BACTERIA FROM LIQUID EXTRACTS
1. Follow MOP 6535a for spread-plating samples in triplicate from the 2 oz cups.
2. Filtration – detailed in the steps below – is required only if extremely low recovery is expected.
a. Label three TSA plates with information specific to the sample and amount to be filtered.
b. Set up filter manifold with three filter unit cups.
c. Set the vacuum, with proper gauge, to no more than 10 inches Hg (4.9 psi).
d. Add 10 ml PBST to filter cup.
e. Add 1 ml sample eluent from sample cup to each filter cup on the manifold.
f. Open the vacuum valve and vacuum liquid through filter.
g. Rinse sides of filter cup with 10 ml of sterile PBST, vacuum through again.
h. Pop off sides of the filter cup by squeezing and lifting.
i. Use sterile forceps to lift filter and place it on the TSA, making sure good contact with the
agar surface is made and air pockets are minimized.
3. Repeat for subsequent sample jars, completing samples in batches of no more than 4. This will help
to avoid too much settling time between vortexing and plating.
4. Place all TSA plates in incubator set at 35 ºC and incubate 18 – 24 hrs before counting colonies.
MOP 6579
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April 2013
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4.0
CALCULATIONS
For spread plates: total CFU recovered = (mean CFU on plate / 0.1ml volume plated) x (1/dilution
factor) x (5 ml total in jar)
For vacuum filter plates: total CFU recovered = (mean CFU on vacuum filter / volume filtered) x 5 ml
total in jar
MOP 6580
Revision 2
February 2013
Page 1 of 7
Miscellaneous Operating Procedure (MOP) 6580:
Recovery of Bacillus Spores from 3M Sponge-Stick™ Samples
Prepared by: __________________________________________
Date: 2/12/2013
Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader
Reviewed by: __________________________________________
Dahman Touati, ARCADIS Project Manager
Date: 2/12/2013
Approved by: __________________________________________
Worth Calfee, EPA Work Assignment Manager
Date: 2/12/2013
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
MOP 6580
Revision 2
February 2013
Page 2 of 7
MOP 6580
TITLE:
RECOVERY OF BACILLUS SPORES FROM 3M SPONGE-STICK™
SAMPLES
SCOPE:
This MOP provides the procedure for recovering spores from 3M SpongeStick™ samples.
PURPOSE:
To extract and quantify bacterial spores from 3M Sponge-Stick™ samples
using a highly repeatable procedure.
MATERIALS

pH-amended bleach

70-90 % Solution of denatured ethanol

Deionized (DI) water

Kimwipes

3M Sponge-Stick™ samples (P/N SSL10NB), hereafter referred to as ‘sponge’

Seward Stomacher® bags (P/N BA6041/CLR)

Phosphate buffered saline with 0.05% TWEEN®20 (PBST) (SIGMA-ALDRICH, Co,
P/N P3563-10PAK)

MicroFunnel Disposable Filter Funnels, Pall Life Sciences (VWR P/N 55095-060) or
Nalgene Sterile Analytical Filter Unit (Fisher P/N 130-4020)

Disposable polystyrene serological pipettes (5mL and 10mL)

Tryptic Soy Agar (TSA) plates

Vortex mixer

Disposable sterile 10µl loops

Disposable sterile forceps

Disposable gloves

Cell spreaders or glass beads for spreading

Racks for 15 mL and 50 mL centrifuge tubes

Sterile, plastic, screw-cap 50 mL centrifuge tubes (e.g. Fisher Cat# 14-959-49A)
MOP 6580
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February 2013
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
Sterile, plastic, screw-cap 15 mL centrifuge tubes (e.g. Fisher Cat# 14-959-49D)

Pipette tips with aerosol filter for 1 mL and 200 μL
1.0
PREPARATION
Personnel must be familiar with all procedures prior to start.
1.1
Equipment Preparation
a) Begin by donning personal protective equipment (PPE) such as gloves, lab coat,
and protective eyewear.
b) Clean the workspace (Biological Safety Cabinet; BSC) by wiping surfaces with pHamended bleach, next with DI water, and lastly with a 70-90% solution of denatured
ethanol. Allow any excess liquid to dry prior to beginning procedure. Make sure
the workspace is clean and free of debris.
c) Assemble equipment in the BSC as needed: vortex mixer, filtration manifold,
automatic pipettors, tips, racks, etc.
d) Assemble extra supplies, such as stomacher and reagents, near BSC.
1.2
Supply Preparation
a) Unpack shipping containers directly into a BSC.
b) If sponges are not in Stomacher® bags, label one Stomacher® bag for each sponge
and place in a bag rack.
c) Label two sterile 50 mL centrifuge tubes for each sponge sample and place in tube
rack.
d) For each sample, label TSA plates on the agar side of the plate with the sample
number and the appropriate dilution factors, as per MOP 6535a (Serial Dilution:
Spread Plate Procedure to Quantify Viable Bacterial Spores).
e) Label two additional plates for filter-plate analysis.
MOP 6580
Revision 2
February 2013
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2.0
PERFORM SPORE EXTRACTION, ELUTION, AND CULTURE PROCEDURE
2.1
Dislodge Spores from the Sample Sponges
a) Begin by donning a new pair of gloves. All subsequent procedures involving
manipulation of sponges or spore suspensions must be carried out in a BSC.
(Stomaching may occur outside the BSC when samples are double-contained inside
the indicated bags.)
b) If the sponges are not in Stomacher® bags, aseptically transfer each sponge to a
Stomacher® bag (labeled during step 1.2b) using sterile disposable forceps. Change
forceps between samples.
c) Aseptically add 90 mL of PBST to each bag that contains a sponge.
d) Stomach sponges in the PBST by completing the following:

Make certain the Stomacher® is set to MANUAL. Program the Stomacher®
speed to 260 RPM and the timer to 1 minute.

Open the Stomacher® door by raising the lid fully upward and back. The
DOOR OPEN icon will be displayed.

Place the stomacher bag containing the sponge sample into a second stomacher
bag to contain any leakage in the event the primary containment is
compromised. Place the combined bags such that 50 to 60 mm of the top
portions protrude above the bag clamp, while making certain that the sponge
sample rests evenly between the homogenizer paddles.

Close the door to the Stomacher®. The DOOR OPEN icon will no longer be
illuminated.

Stomach each sponge for 1 min by pressing the START button.

When the cycle ends, the Stomacher® will stop. If there is an emergent reason
to stop the stomacher during the 1 minute stomaching period, press the red
button or the power button to do so prior to opening the Stomacher®. Stopping
the Stomacher® by opening the door can damage the equipment.

Open the door of the Stomacher® and remove the bags containing the sponge.
Grab the sponge from the outside of the bag with your hands. Move the sponge
MOP 6580
Revision 2
February 2013
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to the top of the bag while using your hands to squeeze excess liquid from the
sponge.

Remove and discard the sponge using sterile forceps.
e) Repeat steps (b) through (d) for all samples.
f) Allow bags to sit for 10 min to allow elution suspension foam to settle before
beginning the concentration step.
2.2
Remove Sponge Elution Suspension
a) Gently mix elution suspension up and down with a 50 mL pipette three times.
b) Split elution suspension volume equally.

Remove half of the suspension volume (~45 mL) with a sterile 50 mL pipette
and place it in a 50 mL screw capped centrifuge tube.

Place remaining suspension (~45 mL) into a second 50 mL tube.
c) Record suspension volumes on tubes and data sheet.
d) Repeat steps (a) through (c) for all samples.
2.3
Concentrate Sponge Elution Suspension (Optional)
a)
Centrifuge 50 mL centrifuge tubes

Prior to daily use and before placing tubes into centrifuge, follow MOP 6558
(Centrifuge Cleaning Procedure) for cleaning this equipment.

Add centrifuge tubes to rotor, evenly distributing weight.

Centrifuge tubes at 3500 x g for 15 min. Do not use the brake option on the
centrifuge to slow the rotor, as re-suspension of pellet may occur.
b) Carefully remove about 42mL of supernatant with a 50 mL pipette and discard to
leave approximately 3 mL in each tube. The pellet may be easily disturbed and not
visible, so place pipette tip away from the tube bottom or side.
MOP 6580
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c) Vortex and sonicate tubes as follows:

Set vortex mixer to level 10 and touch activation.

Turn on sonicator water bath.

Vortex tubes for 30 sec.

Transfer tubes to sonicator bath and sonicate for 30 sec.

Repeat vortex and sonication cycles two additional times.
d) Remove suspension from one tube with a sterile 5 mL pipette and place it in the
other tube of the same sample. The combined result is the final sponge elution
suspension.
e)
Measure final volume of the final sponge elution suspension with 5 mL pipette
and record on tube and data sheet.
f) Repeat steps (e) through (i) for all samples.
2.4
Serially Dilute and Plate the Final Spore Elution Suspension
a) Use MOP 6535a to serially dilute and plate samples.
NOTE: If the samples are turbid, wide-orifice pipette tips may be used to prevent
clogging of pipette tips.
b) Place all plates in an incubator set at 35 ± 2 ºC for a maximum of 3 days. Plates
should be examined within 18-24 hours after start of incubation. Manually
enumerate CFU of target organism and record data.
2.5

If the CFU is <300/plate, record actual number.

If the CFU is >300/plate, record as “too numerous to count” (TNTC)
Capture Spores on Filter Membranes and Culture on TSA
Choose one of the following to methods to filter the final spore elution suspension:
a)
Complete filter plating using MOP 6565 (Filtration and Plating of Bacteria from
Liquid Extracts).
MOP 6580
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b)
Complete filter plating using the following method:
1) Place two 0.45 μm (pore-size) Microfunnels on a Pall vacuum manifold (Pall Cat#
15403).
2) Moisten Microfunnel membranes with 5 ml PBST, open vacuum, and vacuum
through the filter. All filtering should be done with a vacuum pressure <20 cm Hg.
3) Make certain that the manifold vacuum valve is closed. Turn on the vacuum.
4) With the vacuum valve closed, place 10 mL of PBST into each filter cup.
5) Add 1.0 mL of the final sponge elution suspension to each filter cup.
6) Open valves and allow the suspension to flow through the filter, close the valve.
7) Rinse the walls of each Microfunnel cup with 10 mL of PBST. Reopen the valve to
allow the suspension to flow through the filter.
8) Close the valve, turn off the vacuum pump. Slowly reopen the valve to equalize the
pressures.
9) Squeeze the walls of the Microfunnel cup gently and separate the walls from the
base holding the filter. Remove each filter membrane with sterile disposable
forceps and place grid-side up on a TSA plate. Make sure that the filter is in good
contact with the surface of the agar. If an air pocket occurs under the filter, use the
sterile forceps to lift the edge of the filter to release the air pocket for better contact
with the agar.
10) Record exact volume of the sponge elution suspension filtered on each plate. It
should be 1 mL. (Greater sample volumes may be used to lower detection limits)
11) Repeat steps (1) through (8) for all each sample.
12) Incubate TSA plates with filter membranes at 35 ± 2 ºC for a maximum of 3 days.
Plates should be examined within 18-24 hours after start of incubation. Manually
enumerate CFU of target organism and record data.

If the CFU is <300/plate, record actual number.

If the CFU is >300/plate, record as “too numerous to count” (TNTC)
MOP 6582
Revision 2
April 2013
Page 1 of 10
Miscellaneous Operating Procedure (MOP) 6582:
Recovery of Bacillus Spores from Trace Evidence Filters
Prepared by: __________________________________________
Date: 4/29/2013
Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader
Reviewed by: __________________________________________
Dahman Touati, ARCADIS Project Manager
Date: 4/29/2013
Approved by: __________________________________________
Worth Calfee, EPA Work Assignment Manager
Date: 4/29/2013
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
MOP 6582
Revision 2
April 2013
Page 2 of 10
MOP:
6582
Title:
RECOVERY OF BACILLUS SPORES FROM TRACE EVIDENCE FILTERS
Objective:
To aseptically extract and quantify recovery of Bacillus spores from 3M Trace Evidence
filters (TEF) used to sample porous surfaces.
MATERIALS AND EQUIPMENT
Equipment:

Biological Safety Cabinet (BSC, Class II)

Incubator set to appropriate growth temperature for target organism (35 °C)

Light box for counting colonies

Vortex mixer

Stomacher (Seward Stomacher® 400 Circulator (Seward; catalog #0400/001/AJ)

Closure bags (catalog # BA6141/CLR)

Stomacher bag rack (Seward catalog # BA6090)

Specimen cup rack

50 mL Tube rack
Materials:

Personal Protective Equipment (PPE) (gloves, lab coat, safety goggles)

1:10 Bleach, pH-adjusted (pAB) to 7.0 with acetic acid

Dispatch bleach wipes

Disposable 500 mL beaker (such as Fisher Scientific cat #14-955-111D) (for discarding used tips
and beads/spreaders)

70% Solution of denatured ethanol

Kimwipes

Waste container

Sterile Phosphate Buffered Saline with Tween 20 solution (PBST) (MOP 6562)

Tryptic Soy Agar (TSA) plates

3M Forensic Vacuum Filter (Trace Evidence Filter) (Precision Data Products, Grand Rapids, MI
catalog #FF-1)

Stomacher bags, closure type (Seward catalog # BA6141/CLR)
MOP 6582
Revision 2
April 2013
Page 3 of 10
1.0

Sterile scalpels, preferably one for each sample

Sterile 50 mL centrifuge tubes filled with 45 mL of sterile PBST, two for each sample

Pipettor and sterile 50 mL pipettes

Pipettor and pipette tips for dilutions (100 μl, 1000 μl, and 5000 μl.)

Disposable cell spreaders or glass beads for cell spreading

Disposable sterile thumb forceps

Disposable sterile 2 mL bulb pipettors (such as Fisher Sci cat #13-711-7M)
REMOVING FILTER AND DUST FROM TRACE EVIDENCE FILTER
1. Begin by donning PPE (gloves, lab coat, and protective eyewear).
2. Each TEF should be received separately in a small sealable bag (primary containment), and within
secondary containment. Make certain that all of the samples are labeled. Review any chain of
custody forms that may accompany the samples to ensure that all of the labels are consistent and that
there is no notable variation in the samples. If discrepancies are found, make a note of each one in
the notebook.
3. Clean the workspace (BSC) by wiping surfaces with pAB, next with deionized water, and lastly with
a 70-90% solution of denatured ethanol. Wipe with a kimwipe to remove any excess liquid. Make
sure the workspace is clean and free of debris. Gather all necessary items to perform the task.
4. Discard gloves and replace with fresh pair.
5. For each TEF sample, label one closure-type stomacher bag and two 50 ml centrifuge tubes
(containing 45 ml PBST each) with the sample identifier.
6. One at a time, under the BSC, remove the TEF from the primary containment bag in which it
arrived. Using the Dispatch bleach wipes, wipe each cassette with a wipe, and then a clean
kimwipe. Discard the used bleach wipe and the used kimwipe in the secondary containment bag and
place them in the non-regulated waste container. Remove gloves and don a fresh pair of gloves
before proceeding.
7. With the plug in place on the back side of the filter, remove the black nozzle cover from the front
filter side of the cassette, use a pipette and aseptically dispense 5 mL of PBST (taken from the
centrifuge tubes containing 45 ml PBST) into the cassette (Figure 1).
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Figure 1. Dispensing 5 mL PBST into TEF nozzle
8. Replace the black nozzle cover and roll the TEF around to allow the liquid to touch all surfaces of
the inside of the cassette. If there is a large quantity of dust inside, more PBST may be needed to
dampen the dust before opening the TEF. Be careful not to hold or squeeze the black nozzle cover,
as this may cause the seal to be disrupted, resulting in loss of PBST.
9. Once a visual check has been completed to assure that the dust is dampened enough to prevent its
aerosolization, use the sterile scalpel to cut through the red tape seal around the circumference of the
TEF.
10. Holding the TEF nozzle side up, open the plastic case of the filter. Turn the top of the filter case
upside down (nozzle side down with the black nozzle cover in place) and set aside for a moment,
inside the BSC on a rack (specimen cup rack will work) so that it will not be touching anything.
(Figure 2)
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Figure 2. Place the top half of the filter case onto a rack so that the filter can be removed from
the bottom half of the filter case.
11. Position the labeled stomacher bag in the bag rack so that the closure wires can be bent in a manner
so that the bag remains open while standing up in the rack (Figure 3).
Figure 3. Stomacher bag in bag holder with wires in open position, ready to accept filter and
liquid.
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12. Remove the moistened filter with sterile forceps and place in the stomacher bag (Figure 4).
Figure 4. Placing moistened filter into open stomacher bag.
13. Use a pipette to aseptically remove liquid that has collected under the filter during the moistening
step (Step 7 above). Place the liquid into the stomacher bag with the filter (Figure 5).
Figure 5. Transferring moistening liquid to stomacher bag.
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14. Use a disposable bulb pipette to aseptically rinse the inside walls of the bottom portion of the filter
case with 1 to 2 mL of PBST (additional aliquots from the same 45 mL tubes used previously), then
remove the rinse eluent (using the same transfer pipette) and place it into the stomacher bag along
with the filter (Figure 6).
Figure 6. Rinsing inside walls of filter case.
15. Repeat the rinsing step for the top portion of the filter case that was set aside (Step 10).
16. Pipette or pour the remainder of the 90 mL PBST (two tubes of 45 mL each) into the stomacher bag
that contains the filter and the rinsate of the filter case (Figure 7).
Figure 7. Pouring remainder of 90 mL PBST into stomacher bag.
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17. Discard the filter case, red plug nozzle cover, disposable pipettes and disposable thumb forceps into
the proper waste containers. DO NOT discard the 50 mL centrifuge tubes, as you will use these
again for centrifuging the eluent after processing in the stomacher.
18. Close stomacher bags by folding multiple times and folding the wire closures into the center of the
bag. Wipe the outside of the bag with a disinfecting wipe and place in a secondary stomacher bag
rack or sample retention bin.
19. Wipe down the surface of the BSC as described in Step 3 above.
20. Change gloves, and repeat from Step 5 for each remaining TEF sample.
2.0
PROCESSING TRACE EVIDENCE FILTERS
1. Set the stomacher to 1 min, 260 rpm (setting P3). Place one stomacher bag from Section 1.0, Step 18
(above) into the stomacher and run the cycle. Remove the stomacher bag and place back in the bag
rack.
2. Repeat for each sample.
3. Hold the samples at room temp for 10 min to allow the foam to settle in the stomacher bag (Figures
8 and 9).
Figure 8. Stomacher bag with sample; foam in stomacher bag immediately after 1 min
stomacher cycle.
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Figure 9. Stomacher bag with sample; settled foam after 10 min holding time.
4. Using a 50 mL pipette, transfer the eluent evenly (so the tubes will be balanced in the centrifuge)
into the same two labeled 50 mL centrifuge tubes saved from Section 1.0, Step 17 above (~43 mL in
each tube, since ~ 4 mLs is absorbed by the filter).
5. Weigh the tubes to guarantee that they will be balanced (+/- 0.25g).
6. Centrifuge the eluent at 3500g for 15 min.
7.
Carefully remove all but approximately 3 mL (use the 5mL gradient on the side of the tube to guide
the amount) from each tube using a 50 mL pipette, being careful not to disturb the pellet. Vortex
vigorously to disperse the pellet.
8. Vortex the remaining liquid in the tubes and then aseptically combine the remaining eluent from the
two tubes so that approximately 6 mL of eluent is now in one tube. Vortex and sonicate to break up
aggregates before plating.
9. Measure and record the exact volume of the remaining liquid within each sample.
3.0
PLATING AND FILTRATION OF BACTERIA FROM LIQUID EXTRACTS
1. Follow MOP 6535a for serial dilutions and spread-plating samples in triplicate from the ~6mL
solution containing the concentrated sample.
2. Filtration – detailed in the steps below – is required only if extremely low recovery is expected.
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a. Label three TSA plates with information specific to the sample and amount to be filtered.
b. Set up filter manifold with three filter unit cups.
c. Set the vacuum, with proper gauge, to no more than 10 inches Hg (4.9 psi).
d. Add 10 mL PBST to filter cup.
e. Add 1 mL sample eluent from sample cup to each filter cup on the manifold.
f. Open the vacuum valve and vacuum liquid through filter.
g. Rinse sides of filter cup with 10 mL of sterile PBST, vacuum through again.
h. Pop off sides of the filter cup by squeezing and lifting.
i. Use sterile forceps to lift filter and place it on the TSA, making sure good contact with the
agar surface is made and air pockets are minimized.
3. Repeat for subsequent sample tubes, completing samples in batches of no more than 4. This will help
to avoid too much settling time between vortexing and plating.
4. Place all TSA plates in incubator set at 35 ºC and incubate 18 – 24 hrs before counting colonies.
4.0
CALCULATIONS
For spread plates:
total CFU recovered = (mean CFU on plate / 0.1 mL volume plated) x (1/dilution factor) x (total
volume in centrifuge tube after decanting)
For vacuum filter plates:
total CFU recovered = (mean CFU on vacuum filter /volume filtered) x total volume in
centrifuge tube after decanting
MOP 6586
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Miscellaneous Operating Procedure (MOP) 6586:
Recovery of Bacillus Spores from Quartz Filters
Prepared by: __________________________________________
Date: 2/12/2013
Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader
Reviewed by: __________________________________________
Dahman Touati, ARCADIS Project Manager
Date: 2/12/2013
Approved by: __________________________________________
Worth Calfee, EPA Work Assignment Manager
Date: 2/12/2013
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
MOP 6586
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MOP:
6586
Title:
RECOVERY OF BACILLUS SPORES FROM QUARTZ FILTERS
Objective:
To aseptically extract and quantify recovery of Bacillus spores from quartz filters used to
sample air.
MATERIALS AND EQUIPMENT
Equipment:

Biological Safety Cabinet (BSC, Class II)

Seward 400 Circulator Stomacher®

Incubator set to appropriate growth temperature for target organism (35 °C ± 2 ºC)

Light box for counting colonies

Vortex mixer

Stomacher bag rack

Serological pipette
Materials:

Personal Protective Equipment (PPE) (gloves, lab coat, safety goggles)

1:10 Bleach, pH-amended to 7.0 with acetic acid (pAB)

Dispatch bleach wipes

Disposable 500 ml beaker (such as Fisher Scientific, cat #14-955-111D) (for discarding used tips
and beads/spreaders)

70% Solution of denatured ethanol

Kimwipes

Waste container

Sterile Phosphate Buffered Saline with Tween 20 solution (PBST) (MOP 6562)

Tryptic Soy Agar (TSA) plates

Sterile 1500 µL tubes filled with 900µL of sterile PBST

120 mL specimen cup containing 100 mL of sterile PBST
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1.0

Pipettor and pipette tips for dilutions (100 μl, 1000 μl, and 5000 μl.)

Disposable cell spreaders or glass beads for cell spreading

100 mL serological pipette tips

Disposable sterile thumb forceps

10” x 15” Twirl’em bag (Fisher, cat # 01-002-53)

Quartz filter (Whatman, cat # 18209932)

Seward stomacher 400 bags (BA6141/CLR closure bags x 10)
PREPARATION
Personnel must be familiar with all procedures prior to start.
1.1
Equipment Preparation
a) Begin by donning personal protective equipment (PPE) such as gloves, lab coat, and protective
eyewear.
b) Clean the workspace (Biological Safety Cabinet; BSC) by wiping surfaces with pH-amended
bleach (pAB), next with deionized (DI) water, and lastly with a 70-90% solution of denatured
ethanol. Allow any excess liquid to dry prior to beginning procedure. Make sure the
workspace is clean and free of debris.
c) Assemble equipment in the BSC as needed: vortex mixer, filtration manifold, automatic
pipettors, tips, racks, etc.
d) Assemble extra supplies, such as stomacher and reagents, near BSC.
e) Wipe down workspace once again with 70-90% solution of denatured ethanol.
1.2
Supply Preparation
a) Unpack shipping containers onto a cart near a BSC.
b) For each sample, label TSA plates on the agar side of the plate with the sample number and the
appropriate dilution factors, as per MOP 6535a (Serial Dilution: Spread Plate Procedure to
Quantify Viable Bacterial Spores).
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c) Pre-label sterile, empty 120 mL specimen cups to collect the extraction liquid from each
sample.
2.0
PERFORM SPORE EXTRACTION AND CULTURE PROCEDURE
2.1
Dislodge Spores from the Sample Filters
a) Begin by donning a new pair of gloves. All subsequent procedures involving manipulation of
the filters or spore suspensions must be carried out in a BSC. (Stomaching may occur outside
the BSC when samples are double-contained inside the indicated bags.)
b) Outside of the BSC, open the secondary (10” x 15”) containment bag containing the labeled
stomacher bag containing the sample filter. Check to make sure the secondary bag and the
stomacher bag have matching labels. Wipe down the stomacher bag with a dispatch wipe and
place in the stomacher rack located in the BSC. Discard the outer 10” x15” bag in the nonregulated waste container.
c) Aseptically add 100 mL of PBST to each labeled stomacher bag containing a filter. The 100
mL of PBST should be pre-measured in a 120 mL specimen cup. Discard this specimen cup
when empty.
d) Place the stomacher bag containing the quartz filter sample and PBST into a second stomacher
bag to contain any leakage in the event the primary containment is compromised. Place the
combined bags such that 50 to 60 mm of the top portions protrude above the bag clamp, while
making certain that the quartz filter rests evenly between the homogenizer paddles.
e) Stomach filters in the PBST by completing the following:
1) Make certain the Stomacher® is set to MANUAL. Program the Stomacher® speed to 230
RPM and the timer to 2 minutes.
2) Open the Stomacher® door by raising the lid fully upward and back. The DOOR OPEN
icon will be displayed.
3) Close the door to the Stomacher®. The DOOR OPEN icon will no longer be illuminated.
4) Stomach each filter for 2 min by pressing the START button.
5) When the cycle ends, the Stomacher® will stop.
Note: If there is an emergency, stop the stomacher by pressing the red button or
the power button prior to opening the Stomacher®. Stopping the Stomacher® by
opening the door can damage the equipment.
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6) Open the door of the Stomacher® and remove the bags containing the filter. Remove the
labeled stomacher bag containing the filter sample and place it into the stomacher bag rack.
The same secondary stomacher bag may be used for each sample provided there was no
leak. If a leak was detected, leave the labeled stomacher bag in the secondary bag and
place both into the stomacher bag rack.
f) Repeat steps (b) through (d) for all samples.
g) Allow bags to sit for 10 min to allow the suspension foam to settle before beginning the
extraction step.
2.2
Recovery of Suspension
a) Obtain a new, empty, pre-labeled 120 mL specimen cup with the same sample ID as that of the
stomacher bag.
b) The suspension will be removed using a 100 mL pipette. Open the stomacher bag and remove
the liquid by first tilting the bag to one side, moving the liquid away from the filter. Remove
as much liquid as possible using the pipette.
c) When liquid can no longer be collected this way, simultaneously squeeze the filter and collect
liberated liquid via pipette. Make attempts to locate pipette tip in a location within the bag that
reduces collection of filter debris.
NOTE: The filter particles can become lodged in the pipette tip. If this happens it may be
necessary to eject some of the liquid to dislodge the particle.
d) After all the liquid has been removed from the bag, place it into the pre-labeled specimen cup.
Make sure to note the volume collected on the specimen cup as well.
e) Discard the labeled stomacher bag with the filter into a non-regulated waste container.
f) Repeat steps (a) through (e) for each sample.
2.3
Serially Dilute and Plate the Final Spore Elution Suspension
a) Use MOP 6535a to serially dilute and plate samples.
NOTE: If the samples are turbid, wide-orifice pipette tips may be used to prevent clogging of
the pipette tips.
MOP 6593
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Miscellaneous Operating Procedure (MOP) 6593:
Recovery of Spores from HVAC Filters
Prepared by:
__________________________________________
Date: 4/30/2013
Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader
Reviewed by:
__________________________________________
Date: 4/30/2013
Dahman Touati, ARCADIS Project Manager
Approved by:
__________________________________________
Worth Calfee, EPA Work Assignment Manager
Prepared for
National Homeland Security Research Center
Office of Research and Development
U.S. Environmental Protection Agency
Research Triangle Park, NC 27711
Prepared by
ARCADIS U.S., Inc.
4915 Prospectus Drive, Suite F
Durham, NC 27713
Date: 4/30/2013
MOP 6593
Revision 1
April 2013
Page 2 of 3
MOP: 6593
Title:
RECOVERY OF SPORES FROM HVAC FILTERS
Scope:
This MOP outlines the procedure for recovering spores from HVAC filter sections
Purpose:
To aseptically extract and quantify spores from sections of HVAC filters in order to
determine viability and obtain quantifiable data
1.0
MATERIALS

PPE (gloves, lab coat, safety goggles)

Class II Biological Safety Cabinet (BSC)

pH-amended bleach

Deionized (DI) water

70% solution of denatured ethanol

Kimwipes

Dispatch bleach wipes

Non-regulated waste container

Bottles of sterile Phosphate Buffered Saline with Tween 20 solution (PBST) (MOP 6562)

Vortex mixer

Shaker table with clamps or brackets to hold Nalgene bottles

Cart

Trypticase soy agar (TSA) plates

900uL dilution tubes of sterile PBST

Pipettor and pipette tips for dilutions

Incubator set to appropriate growth temperature for target organism (35°C+/-2°C)

Light box for counting colonies

Lab notebook

Sterile graduated cylinder

Parafilm
2.0
PROCEDURE
1. Begin by donning PPE (gloves, lab coat, and protective eyewear).
2. Samples will be received as follows: Half portions of HVAC filters will be in 32oz white capped Nalgene
jars (Fisher cat no. 2118-0032) and quarter portions of HVAC filters will be in 1L clear bottles (Fisher cat
no. 02-893D). The samples should be secondarily contained by large sterile bags (10” x 15” sterile
bags Fisher cat no. 01-002-53). Chain of custody forms that accompany the samples will need to be
MOP 6593
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reviewed to ensure that all of the samples IDs are consistent and that there is no notable variation in the
samples. If variation has occurred, make a note of it in the notebook.
3. Clean BSC workspace by wiping surfaces with pH-amended bleach, followed by DI water, and lastly
with a 70% solution of denatured ethanol. Wipe with a kimwipe to remove any excess liquid. Make
sure the workspace is clean and free of debris. Gather all necessary items to perform the task, place
these items on a clean cart beside the BSC, within arm’s reach so that once the procedure has begun
the task may be performed without interruptions.
4. Discard gloves and replace with a fresh pair.
5. Place the samples one at a time, under the BSC. Samples should be handled in order of least
contaminated to most contaminated, with any negative controls or blank samples being handled first,
test samples being handled next, and lastly the positive controls. Carefully unwrap the secondary
containment bag from around the jar or bottle, discard in non-regulated waste container and then wipe
down the jars or bottles with a dispatch wipe, followed by a Kimwipe. Change gloves after handling
each sample.
6. Once all of the samples are under the BSC and disinfected, change gloves and place the sterile
graduated cylinder under the BSC, along with several bottles of sterile PBST. Again, in order from
least contaminated to most contaminated, handle the samples individually, and carefully open the jars
or bottles to aseptically add 700 mL of PBST to the sample. The PBST is first aseptically poured into
the graduated cylinder and measured at 700 mL, and then it is aseptically poured from the graduated
cylinder into the sample. Carefully transfer all liquid as to not spill any or disturb/touch the filter or its
container in any way. Once the 700 mL of PBST has been added, carefully place the jar lid or bottle top
back on the sample, and make certain it is tightly closed.
7. Again, wipe the outside of the samples with Dispatch wipes, followed by a Kimwipe to remove any
moisture. After all samples have 700 mL of sterile PBST added, wrap the sample lids/bottle tops with
Parafilm to ensure a tight seal and to prevent sample leakage.
8. Place all sample jars or bottles into an orbital shaker table, using the specially designed platform
holders. Make certain all samples are secure, and shake at room temperature for 30 min at 300 rpm.
9. Immediately after the thirty minute agitation step is completed, remove the samples from the shaker and
place them, again from least contaminated to most contaminated, into the BSC for dilution plating. Don
a fresh pair of gloves, remove and discard the Parafilm, and follow MOP 6535a to complete serial
dilutions using TSA media plates that are appropriately labeled with the sample ID, dilution set, and
date. Each sample should also be manually agitated for ten seconds prior to removal of aliquots.
Dilution tubes used in MOP 6535a should contain PBST to stay consistent with materials/solutions.
10. Once the dilution plating has been completed, incubate the plates at 35°C +/- 2°C for 18-24 hours, and
archive the samples in at refrigerator at 4°C+/-2°C.
11. Count the colonies with the assistance of a light box. Record data.
12. Colony forming units for each 6” x 12” half section or for each 6” x 6” quarter section will be determined
and the sum of the CFUs for the two or four sections making up one 12” x 12” sample will be
determined. Data will be reported as total CFU for the 12” x 12” sample.
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
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