CONTROL OF FOODBORNE PATHOGENS AND THEIR BIOFILMS BY
LEVULINIC ACID AND SODIUM DODECYL SULFATE
by
DONG CHEN
(Under the Direction of Michael P. Doyle)
ABSTRACT
The efficacy of commonly used sanitizers in food processing facilities is reduced
when organic matter is present. If the sanitizing procedures are inadequate, surviving
microorganisms could attach and form biofilms given time and nutrients. The objective
of this study was to validate the antimicrobial efficacy of levulinic acid (LVA) and
sodium dodecyl sulfate (SDS) on deli slicers, and to determine the effectiveness of this
treatment for inactivating foodborne pathogens in single- and mixed-species biofilms.
LVA + SDS was applied at three concentrations (0.5% LVA + 0.05% SDS, 1% LVA +
0.1% SDS, and 2% LVA + 0.5% SDS) on slicers pre-contaminated by Listeria
monocytogenes, Salmonella, and Escherichia coli O157:H7 (ca. 8.5 log CFU/blade) at
21oC. Sampling and enumeration were conducted after a treatment time of 0, 1, 2, 3, 5,
10, and 20 min. Single- and mixed-species biofilms were incubated at 21°C for 72 h
before being treated with different concentrations of LVA plus SDS (0.5% LVA + 0.05%
SDS, 1% LVA + 0.1% SDS, and 3% LVA + 2% SDS). The bactericidal activity of the
LVA with SDS treatment was concentration dependent on both the slicers and in
biofilms. Contaminated slicer surfaces sprayed with 1% LVA plus 0.1% SDS as a foam
(45-55 psi) reduced within 1 min 6.0 to 8.0 log CFU of the pathogens/blade. Greater than
6.9 log CFU of pathogens/coupon in single-species biofilms were reduced within 10 min
by 3% LVA plus 2% SDS. E. coli O157:H7 and Salmonella were antagonistic to each
other, being significantly (P < 0.05) more sensitive to LVA plus SDS in mixed-species
biofilms than in single-species biofilms. Microscopic images of LVA plus SDS-treated
biofilms captured by scanning electron microscopy (SEM) and transmission electron
microscopy (TEM) revealed that the cells were detached from the biofilm matrix and the
integrity of cell envelopes was disrupted. Photomicrographs captured by confocal laser
scanning microscopy (CLSM) confirmed that more pathogens in the biofilms were
detached when the treatment concentrations were increased. Results of this study provide
a better understanding of the antimicrobial behavior of LVA plus SDS to foodborne
pathogens on slicers and in their biofilms.
INDEX WORDS:
levulinic acid, sodium dodecyl sulfate, deli slicer, crosscontamination, biofilm, Listeria monocytogenes, Salmonella,
Escherichia coli
CONTROL OF FOODBORNE PATHOGENS AND THEIR BIOFILMS BY
LEVULINIC ACID AND SODIUM DODECYL SULFATE
by
DONG CHEN
B.E., China Agricultural University, China, 2009
M.S., Auburn University, 2012
A Dissertation Submitted to the Graduate Faculty of The University of Georgia in Partial
Fulfillment of the Requirements for the Degree
DOCTOR OF PHILOSOPHY
ATHENS, GEORGIA
2015
© 2015
Dong Chen
All Rights Reserved
CONTROL OF FOODBORNE PATHOGENS AND THEIR BIOFILMS BY
LEVULINIC ACID AND SODIUM DODECYL SULFATE
by
DONG CHEN
Major Professor:
Committee:
Electronic Version Approved:
Julie Coffield
Interim Dean of the Graduate School
The University of Georgia
May 2015
Michael P. Doyle
Joseph F. Frank
Mark A. Harrison
Yen-Con Hung
Jia-Sheng Wang
DEDICATION
This work is dedicated to my parents, Ronghui Chen and Qiong Lai.
iv
ACKNOWLEDGEMENTS
I would like to first thank my major advisor Dr. Michael Doyle for his advice. His
infinite knowledge in food microbiology as well as critical thinking has changed me and
made me grow as a professional. I am also very grateful to Dr. Tong Zhao for the
guidance of research, sharing of experience, and advices of career path. I would also like
to express my appreciation to Dr. Joseph Frank for giving me the invaluable information
and constructive suggestions on the research biofilms related. I would like to
acknowledge the other members of my committee, Dr. Mark Harrison, Dr. Yen-Con
Hung, and Dr. Jia-Sheng Wang, for their help, patience, and advices through the journey.
I am very grateful to Ms. Ping Zhao. Without her assistance and suggestions, I cannot
execute my research at a fast speed. I would also like to thank Ms. Gwen Hirsch and Dr.
John Shields for their technical assistance and patience when I was doing research in
Athens. I am also very grateful to all the personnel in the Center for Food Safety and
Department of Food Science and Technology. They have been so friendly and nice to me
and giving me a hand whenever I needed. Finally, I would like to thank Griffin. She is
such a boring place without any distractions, making me concentrate on my research and
thereby I can finish the study in less than three years.
v
TABLE OF CONTENTS
Page
ACKNOWLEDGEMENTS .................................................................................................v
LIST OF TABLES ............................................................................................................ vii
LIST OF FIGURES ......................................................................................................... viii
CHAPTER
1
INTRODUCTION .............................................................................................1
2
LITERATURE REVIEW ..................................................................................8
3
TRANSFER OF FOODBORNE PATHOGENS DURING MECHANICAL
SLICING AND THEIR INACTIVATION BY LEVULINIC ACID-BASED
SANITIZER ON SLICERS .............................................................................61
4
CONTROL OF PATHOGENS IN BIOFILMS ON THE SURFACE OF
STAINLESS STEEL BY LEVULINIC ACID PLUS SODIUM DODECYL
SULFATE ........................................................................................................89
5
SINGLE- AND MIXED-SPECIES BIOFILM FORMATION BY
ESCHERICHIA COLI O157:H7 AND SALMONELLA, AND THEIR
SENSITIVITY TO LEVULINIC ACID PLUS SODIUM DODECYL
SULFATE ......................................................................................................117
6
SUMMARY ...................................................................................................153
vi
LIST OF TABLES
Page
Table 3.1: Effect of different concentrations of levulinic acid (LVA) plus SDS at different
exposure times at 21°C on L. monocytogenes inoculated on slicer blades ............80
Table 3.2: Effect of different concentrations of levulinic acid (LVA) plus SDS at different
exposure times at 21°C on S. Typhimurium inoculated on slicer blades ..............81
Table 3.3: Effect of different concentrations of levulinic acid (LVA) plus SDS at different
exposure times at 21°C on E. coli O157:H7 inoculated on slicer blades ..............82
Table 4.1: Inactivation of 72-h biofilms of L. monocytogenes, S. Typhimurium, and
STEC formed on stainless steel after exposure to a sanitizer for 10 min ............107
Table 4.2: Inactivation of 72-h biofilms of L. monocytogenes, S. Typhimurium, and
STEC formed on stainless steel after exposure to a heat treatment at 60, 80 or
100°C for 10 min ................................................................................................109
Table 4.3: Bacterial counts on selective agar plates and TSA of 72-h biofilms of L.
monocytogenes, S. Typhimurium, and STEC formed on stainless steel after a 10min treatment with 3% lactic acid (pH 2.2) or 3% levulinic acid (pH 2.7) .........110
Table 4.4: Inactivation of 72-h biofilms of L. monocytogenes, S. Typhimurium, and
STEC formed on stainless steel after exposing to 80°C for 10 min and a
subsequent 10-min sanitizer treatment.................................................................111
Table 5.1: Bacterial strains used in this study and their origins, curli and cellulose
productions, and counts in single-species biofilms..............................................137
vii
LIST OF FIGURES
Page
Figure 3.1: Food contact locations that were swabbed on slicers to determine pathogen
contamination after inoculated deli foods were sliced ...........................................77
Figure 3.2: L. monocytogenes, S. Typhimurium, and E. coli O157:H7 populations
recovered from different contact locations on slicers after slicing inoculated deli
foods (ca. 3.0 log CFU/cm2) ..................................................................................78
Figure 3.3: Transfer of L. monocytogenes, S. Typhimurium, and E. coli O157:H7 from
inoculated slicer blades (ca. 8.5 log CFU/blade) to uninoculated Swiss cheese,
ham, and roast beef, respectively ...........................................................................79
Figure 4.1: Representative photomicrographs by SEM of biofilms formed by L.
monocytogenes after a 10-min treatment with water (control, A), 0.5% levulinic
acid + 0.05% SDS (B), 1% levulinic acid + 0.1% SDS (C), or 3% levulinic acid +
2% SDS (D) .........................................................................................................101
Figure 4.2: Representative photomicrographs by SEM of biofilms formed by S.
Typhimurium after a 10-min treatment with water (control, A), 0.5% levulinic
acid + 0.05% SDS (B), 1% levulinic acid + 0.1% SDS (C), or 3% levulinic acid +
2% SDS (D) .........................................................................................................102
Figure 4.3: Representative photomicrographs by SEM of biofilms formed by STEC after
a 10-min treatment with water (control, A), 0.5% levulinic acid + 0.05% SDS (B),
1% levulinic acid + 0.1% SDS (C), or 3% levulinic acid + 2% SDS (D) ...........103
viii
Figure 4.4: Representative photomicrographs by TEM of biofilms formed by L.
monocytogenes after a 10-min treatment with water (control, A), or 0.5% levulinic
acid + 0.05% SDS (B) ..........................................................................................104
Figure 4.5: Representative photomicrographs by TEM of biofilms formed by S.
Typhimurium after a 10-min treatment with water (control, A), 0.5% levulinic
acid + 0.05% SDS (B), 1% levulinic acid + 0.1% SDS (C), or 3% levulinic acid +
2% SDS (D) .........................................................................................................105
Figure 4.6: Representative photomicrographs by TEM of biofilms formed by STEC after
a 10-min treatment with water (control, A), 0.5% levulinic acid + 0.05% SDS (B),
1% levulinic acid + 0.1% SDS (C), or 3% levulinic acid + 2% SDS (D) ...........106
Figure 4.7: The temperature of the stainless steel coupons during the 10-min heat
treatment at 60, 80 or 100°C in an oven ..............................................................108
Figure 5.1: Single- and mixed-species biofilm formation by E. coli O157:H7 strain
USDA 5 and Salmonella strain 457-88 on 96-well polystyrene plates incubated at
21°C for 72 h........................................................................................................132
Figure 5.2: Bacterial counts of E. coli O157:H7 strain USDA 5 and Salmonella strain
457-88 in single- and mixed-species biofilms after a treatment for 5 min with
water (control), QAC (150 ppm), 1% levulinic acid + 0.1% SDS, or 3% levulinic
acid + 2% SDS .....................................................................................................133
Figure 5.3: Representative photomicrographs by CLSM of single- and mixed-species
biofilms formed by RFP-labeled E. coli O157:H7 strain USDA 5 and GFP-labeled
Salmonella strain 457-88 after a 5-min treatment with water (control, A, D, and
ix
G), 1% levulinic acid + 0.1% SDS (B, E, and H), or 3% levulinic acid + 2% SDS
(C, F and I) ...........................................................................................................134
Figure 5.4: Photomicrograph showing the three-dimensional modeling of 72-h biofilms
formed by mixed-species of RFP-labeled E. coli O157:H7 strain USDA 5 and
GFP-labeled Salmonella strain 457-88 ................................................................136
Figure 5.5: Representative photomicrographs by CLSM of single- and mixed-species
biofilms formed by RFP-labeled E. coli O157:H7 strain USDA 5 and GFP-labeled
Salmonella strain 457-88 after 4, 8, 24, and 48 h of incubation at 21°C .............141
Figure 5.6: Comparison of planktonic growth rates of FP-labeled and parental E. coli
O157:H7 strain USDA 5 and Salmonella strain 457-88 ......................................143
Figure 5.7: Single- and mixed-species biofilm formation by FP-labeled and parental E.
coli O157:H7 strain USDA 5 and Salmonella strain 457-88 on 96-well
polystyrene plates incubated at 21°C for 72 h .....................................................144
x
CHAPTER 1
INTRODUCTION
Foodborne pathogens such as Listeria monocytogenes, Salmonella, and Shiga
toxin-producing Escherichia coli (STEC) are major food safety concerns. L.
monocytogenes causes listeriosis, a disease that mainly affects immunocompromised
individuals, the elderly, and pregnant women (Kathariou, 2002). Salmonella and STEC
collectively cause in the United States an estimated 1.6 million foodborne illnesses
annually (Scallan et al., 2011). L. monocytogenes, Salmonella, and STEC have been
frequently associated with foodborne outbreaks. Three major listeriosis outbreaks
documented in the United States during the past two decades have been traced to
consumption of contaminated sliced turkey deli meat (CDC, 1999; Gottlieb et al., 2006;
Olsen et al., 2005). Outbreaks of Salmonella infections have been associated with a wide
range of foods, including ground beef, ground turkey, cereal, and peanut butter
(Cavallaro et al., 2011; CDC, 2007, 2013a, b; Russo et al., 2013). Cattle are a major
reservoir for E. coli O157:H7 and non-O157 STEC (Fratamico et al., 2014).
Contaminated raw ground beef has been the most frequently implicated vehicle
contributing to E. coli O157:H7 infections (Rhoades et al., 2009).
Deli slicers are among the most difficult pieces of food processing equipment to
clean and hence, are among the most microbiologically hazardous of all equipment used
in the retail food service. Spoilage microorganisms have been recovered from deli slicers,
including the Pseudomonads and Enterobacteriaceae families (Koo et al., 2013). Some
1
strains of Pseudomonads, such as with the Pseudomonas fragi species, not only have
strong attachment and biofilm-forming ability themselves (Zottola and Sasahara, 1994),
but also can support the attachment and biofilm formation of L. monocytogenes via their
extracellular polymeric substance (EPS) production (Sashara and Zottola, 1993). Once
biofilms form on slicers or in food processing facilities, it is challenging to eliminate
them due to enhanced resistance to disinfectants (Kumar and Anand, 1998; Simões et al.,
2006; Simões et al., 2010). Additionally, mature biofilms can release individual cells into
the surrounding environment, not only allowing the daughter cells to contaminate food
products, but also facilitating their colonization of new niches (Austin and Bergeron,
1995; Srey et al., 2013).
The efficacy of many sanitizers used in food processing facilities is reduced when
organic matter is present, whereby their usefulness as an antimicrobial is mitigated
(Simpson Beauchamp et al., 2012). Effective sanitizers that are practical, efficacious, and
safe to use are needed to control both planktonic cells and biofilms in food processing
environments. Levulinic acid with sodium dodecyl sulfate (SDS) is as an effective
disinfectant for inactivating pure cultures of foodborne pathogens in the presence of
organic matter (Magnone et al., 2013; Zhao et al., 2011; Zhao et al., 2009, 2010), and
bacteria in biofilms (Wang et al., 2012; Zhao et al., 2011). This study was designed to
validate the antimicrobial efficacy of the combination of levulinic acid and SDS on food
processing equipment (chapter 3), to determine both the effectiveness of this treatment
for inactivating foodborne pathogens in biofilms and the antimicrobial mechanism of
levulinic acid plus SDS (chapter 4), and to evaluate its efficacy of this treatment against
2
single- and mixed-species biofilms formed by Salmonella and E. coli O157:H7 (chapter
5).
3
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7
CHAPTER 2
LITERATURE REVIEW
Deli slicers
Deli slicers are widely used in retail food establishments at a high food contact
frequency to slice deli meats, cheese, and produce (Maitland et al., 2013). The slicing of
ready-to-eat (RTE) foods is often the last processing step and usually no thermal
intervention is applied to sliced RTE foods before consumption (Sheen, 2008; Sheen and
Hwang, 2010). Moreover, foodborne pathogens have the ability to be transferred between
contaminated foods and food contact surfaces (Gorman et al., 2002; Kusumaningrum et
al., 2004; Pérez-Rodríguez et al., 2008). Thus, deli slicers can be vehicles of pathogens to
contaminate foods by surface transfer between the equipment and sliced food products
(Lin et al., 2006; Sheen and Hwang, 2010; Vorst et al., 2006).
Because it is assumed that RTE foods are free of pathogens and standard food
handling procedures are followed, a wide variety of different deli meats, cheeses or
produce may be sliced on one deli slicer without cleaning between slicing different items.
This practice can result in cross-contamination between the slicer and the sliced foods if a
contaminated food product is involved. Both microbes that are indicators of good
manufacturing practices in food processing plants and retail environments, such as
Enterobacteriaceae family, and foodborne pathogens, such as Listeria monocytogenes,
have been recovered from slicers in retail establishments (Hudson and Mott, 1993;
Humphrey, 1990; Sauders et al., 2009; van Schothorst and Oosterom, 1984). Many
8
foodborne disease outbreaks have been associated with cross-contamination from food
slicers (Anonymous, 1966, 2007; Ash et al., 1964; Bocket et al., 2011; Burnett and
Davies, 1967; Campbell et al., 2008; Gauthier, 2011; Gilbert, 1969; Gilbert and Maurer,
1968; Howie, 1968; Jordan et al., 1973; Kaufmann et al., 1968; Ryser, 2011; Schmidt,
2013; Spitalny et al., 1984; Tilden et al., 1996; Williams et al., 2000). Previous studies
have investigated in laboratory settings the transfer of pathogens, including Listeria
monocytogenes (Aarnisalo et al., 2007; Chen et al., 2014; Keskinen et al., 2008a; Lin et
al., 2006; Sheen, 2008; Sheen et al., 2010; Sheen and Hwang, 2008; Vorst et al., 2006),
Salmonella (Chen et al., 2014; Sheen and Hwang, 2011; Yeater et al., 2015), Escherichia
coli O157:H7 (Chen et al., 2014; Perez-Rodriguez et al., 2007; Sheen and Hwang, 2010)
and Staphylococcus aureus (Perez-Rodriguez et al., 2007), between deli slicers and food
products. Understanding the transfer characteristics of foodborne pathogens between
slicers and deli foods could assist food safety professionals in better controlling pathogen
contamination of slicers. Cross-contamination pathways have been determined and
assessed within mock retail deli environments by using fluorescent compounds (Gibson
et al., 2013; Maitland et al., 2013). However, knowledge of the transfer of foodborne
pathogens between deli slicers and food products at retail settings is still limited (FDA,
2013a).
Because of the physical complexity of deli slicers (Lin et al., 2006) and the
working ambiance of deli workers (Endrikat et al., 2010; Gombas et al., 2003; Sauders et
al., 2009), deli slicers need extra attention to cleaning, sanitation and maintenance to keep
them safe. Since deli slicers have many inaccessible components and hard-to-reach
locations, they are among the most difficult pieces of food processing equipment to clean
9
and sanitize, and hence, are among the most microbiologically hazardous of all
equipment used in retail food service (Chen et al., 2014). For example, sealants and
gaskets are widely used to seal seams and spaces between assembled parts. However,
with long term use and repeated cleaning, cracks, chips, shrinkage or loss of the sealant
and gasket may occur resulting in compromise of seam integrity. Food materials can
build up in these open seams which are nearly impossible to reach and clean, leading to a
preferable location for microbial growth (FDA, 2011a). Cross-contamination of
pathogens between slicers and food products can occur resulting in foodborne illness.
Thus, all the seams on the slicers should be routinely inspected and once any signs of
damage or defects, such as cracks, scratches, corrosion, or loss are detected, the slicer
should be removed immediately from service until repairs, maintenance and inspection
are completed (FDA, 2011c).
Hygienic design is a critical factor to food processing equipment such as slicers
used in the food industry. Appropriate hygienic design can not only improve their
cleanability and durability, but also dramatically decrease the possibility of triggering
foodborne illness outbreaks (FDA, 2013b). Most slicers currently used in retail food
service are constructed of one-piece anodized aluminum, making the machine easy to
clean due to fewer seams and large radii. Blades are mostly comprised of chromiumcoated hard alloy, which results in long-lasting use, especially at the sharp edge, due to its
high density, low coefficient of friction, low wear rate, and high corrosion resistance
(Newby, 1999). However, once in service, the integrity of the slicer blade surface will be
impacted by a variety of factors such as conditions of use, repeated cleaning procedures,
use of chlorine- and acid-based sanitizers and types of foods being sliced (Lindsay et al.,
10
2013). Pits, scratches, crevices and corrosion will eventually emerge after use and
exacerbate the roughness of blades, increasing the chance of cross-contamination (Arnold
and Bailey, 2000; Bohner and Bradley, 1991). Pits on worn slicer blades can provide a
favorable environment for microbial attachment, growth and even biofilm formation
(Chmielewski and Frank, 2003; Neal, 2013; Zottola and Sasahara, 1994), without
removal by cleaners and sanitizers (Krolasik et al., 2010; Lindsay et al., 2013). Microbial
transfer is extended during slicing due to rougher blades allowing for greater microbial
attachment (Keskinen et al., 2008b; Vorst et al., 2006).
Simple design of all parts of deli slicers was most desirable for cleaning. A flat,
simple and smooth surface designed for carriage trays was much easier to clean than
those with compartments or/and ridges. The food contact surfaces, such as gauge plates
and blade covers, with large grooves and smooth small ridges were better designs than
surfaces with small grooves and large ridges. Moreover, gradual ending grooves were
easier to clean than those that ended abruptly. Blades with fewer indentations were also
easier to clean. The distance between the lower edge of the blade guard ring and the base
of a slicer should be large enough to be reached by hands and brushes. Blade guard ring
with a sharp and deep inner side were difficult to clean. For meat grips, less tooth density
and smooth grease-phobic surfaces improved cleanability.
Reduction of microbial cross-contamination on deli slicers is best accomplished
by a concerted effort among regulators, manufacturers and users. Newly manufactured
slicers certified by the National Sanitation Foundation (NSF) are evaluated to meet the
requirements of NSF/ANSI (American National Standard Institute) Standard 8:
Commercial Powered Food Preparation Equipment and this is included in the NSF
11
compliant slicer listings (NSF International, 2010b). The latest version of NSF/ANSI 8
puts greater focus on the design of slicer blades, carriage trays, gauge plates, joints,
seams and electrical components to reduce microbial cross-contamination (NSF
International, 2010a). Under the new standard, manufacturers are required to provide
detailed instructions on how to clean and sanitize their deli slicers. The FDA recommends
cleaning and sanitizing deli slicers at least once every four hours in order to prevent the
growth of foodborne pathogens (FDA, 2011b). Frequent and thorough cleaning and
maintenance can prevent the accumulation of food soils and juices that can serve as
reservoirs of foodborne pathogens (FDA, 2011b). Deli slicer users are encouraged to use
the slicing equipment certified by the latest version of NSF standards to maintain the
highest levels of hygiene and safety. Slicing operations should strictly adhere to the
instructions provided by slicer manufacturers (Powitz, 2009). After use, the slicers should
be cleaned by the cleaning and sanitizing chemicals recommended by the manufacturers
(FDA, 2011a, b) after being completely disassembled (FDA, 2009). All slicer users
should follow the instructions provided by their manufacturers for proper cleaning and
maintenance procedures.
Lack of cleanliness in the working environment is another reason why deli slicers
are prone to being the source of foodborne pathogen cross-contamination of RTE foods
(Lin et al., 2006; Neal, 2013). Unlike other food processing equipment, deli slicers are
used at ambient temperature in a random and intermittent way throughout the working
day (Powitz, 2009). Plus, the food contact area of deli slicers is large and open to the
environment. If the cleaning and sanitizing programs are inadequate, the situation will be
even worse. Thus, cross-contamination that can occur between food contact surfaces of
12
slicers and deli foods is a major food safety concern in food processing plants and retail
establishments.
Pathogen-contaminated deli food is also a factor contributing to deli slicers being
one of the most microbiologically hazardous pieces of equipment used in retail food
service. Contaminated RTE foods have been a major source of human listeriosis cases
(Hitchins and Whiting, 2001; Yang et al., 2006). In particular, deli meats have been
reported to be a leading food vehicle of listeriosis in the United States (Hammons et al.,
2015; Zhang et al., 2012). Listeriosis, which is caused by Listeria monocytogenes, is a
rare human disease but the fatality rate is as high as approximately 16%, particularly in
vulnerable populations, such as the elderly, pregnant women, and immunocompromised
individuals (FDA, 2013a; Lianou and Sofos, 2007; Yang et al., 2006). It was estimated
that deli meats caused approximately 1,600 cases of listeriosis annually, leading to 300
deaths (Endrikat et al., 2010).
Outbreaks associated with contaminated deli slicers
Deli slicers have been associated with several foodborne illness outbreaks. For
example, a typhoid outbreak in Aberdeen, Scotland in which there were 507 cases and 3
deaths, was caused by Salmonella Typhi-contaminated corned beef that was sliced on a
contaminated slicer (Anonymous, 1966). The slicer had been in constant operation for
several days without intermittent cleaning, and subsequently cross-contaminated other
deli food products (Ash et al., 1964; Howie, 1968). The contaminated cold meats were
then stored in a display window directly exposed to sunlight without the benefit of
cooling until purchased by customers (Anonymous, 1966).
13
In October 1964 and in May 1965, two salmonellosis outbreaks occurred in
Edinburgh, Scotland, causing total 94 illnesses. Chopped pork was taken from the shelf
for customer orders, sliced on a slicer and then put back on the shelf which also held
other cold meats, including minced pork and boiled ham (Burnett and Davies, 1967).
Another salmonellosis outbreak occurred in the Washington D.C. metropolitan area
during May and June, 1965, with more than 600 people becoming ill (Kaufmann et al.,
1968). Salmonella meleagridis was isolated from patients, contaminated deli meats and
one slicer used in the implicated caterer-delicatessen-restaurant (Gilbert, 1969). The
slicer was continuing to contaminate food samples until it was completely dismantled,
thoroughly cleaned and decontaminated with an iodine solution (Kaufmann et al., 1968).
In August, 1970, more than 300 cases of salmonellosis occurred following consumption
of contaminated deli meats served in a large restaurant. A meat slicer contaminated with
Salmonella enteritidis was believed as the major contributing vehicle from both
epidemiologic and bacteriologic evidence (Jordan et al., 1973). Because of personnel
safety concerns, the blade was wiped with a damp cloth instead of being disassembled
and properly cleaned. Thus, pathogens and meat debris could reside and accumulate in
the slicer seams and the bacteria thereby persisted.
In September 1981, yet another slicer-associated outbreak of salmonellosis
occurred, which was at a hospital in Vermont. The main symptom was diarrhea and
Salmonella cultures isolated from precooked roast beef which was related to the illness as
were of serotypes Chester, Tennessee, and Livingston. A meat slicer was determined to
be a source of cross-contamination (Spitalny et al., 1984). This mode of transmission was
also suspected in two Escherichia coli O157:H7 outbreaks associated with fermented dry
14
salami. A variety of deli food products were sliced on the same slicer which was not
properly cleaned and sanitized in between products, thereby resulting in 39 outbreakrelated cases in Canada and 17 in Washington and California states (Tilden et al., 1996;
Williams et al., 2000). In 2006 and 2007, a chain fast-food restaurant was associated with
salmonellosis outbreaks in Georgia and Washington states, with both two outbreaks
traced back to cross-contamination by slicers (Anonymous, 2007). Not only bacterial
pathogens such as Salmonella and Escherichia coli O157:H7 were transferred between
the deli meats and slicers, but hepatitis C virus was also transmit from an employee to
another by a meat slicer (Bocket et al., 2011). The workers shared one slicer and hand
injuries were likely caused by the sharp blade, thereby providing the opportunity for the
bloodborne pathogen to be transferred by the slicers. In 2008, the “worst in the world”
listeriosis outbreak was associated to deli meats in Canada, causing 57 hospitalizations
and 22 deaths (Attaran et al., 2008; Campbell et al., 2008; Gauthier, 2011; Ryser, 2011;
Schmidt, 2013). Listeria monocytogenes contamination was linked to a deli meat slicer
that was not properly cleaned and sanitized and the slicer parts with cracks on the food
contact surface were not replaced promptly (Schmidt, 2013; Warriner, 2011).
Biofilms
Bacteria in nature are primarily found either flowing freely in a liquid or attaching
to a surface in a large number to form a three-dimensional structure known as a biofilm.
Biofilms are single or multi-layers of microorganisms embedded in their own
extracellular polymeric substances (EPS) which associate with a solid surface (Donlan
and Costerton, 2002). Biofilm formation can occur on a variety of both biotic and abiotic
surfaces, including teeth, fresh produce, cement, ceramic, nylon, polyester, polystyrene,
15
stainless steel, and glass (Adetunji et al., 2014; Blackman and Frank, 1996; Fett, 2000;
Soni et al., 2013; Wang et al., 2012). A biofilm is mainly comprised of water (80-90%),
EPS that contributes 85-98% of the organic matter, the microorganisms, and entrapped
organic and inorganic particles (Flemming, 1998). Planktonic and biofilm modes of
growth are switchable when there is a change in environmental conditions (O'Toole et al.,
2000). Not like planktonic cells, bacterial cells in biofilms are capable of making a series
of adjustments, including tolerance to nutrient deficiency, slow growth, differences in
gene expression and enhanced stress resistance (Simões et al., 2010).
Biofilm formation is a dynamic process and involves a series of steps. The first
step is attachment, which can be active or passive depending on bacterial motility or
transportation of the cells by gravity, diffusion or fluid dynamic forces (Kumar and
Anand, 1998). In this process, bacterial cells first adhere to a substratum reversibly by
van der Waals electrostatic forces and hydrophobic interactions. The bacterial cells have
Brownian motion, and can be readily removed by shear force (Marshall et al., 1971).
Once appendages on the cell surface are anchored and extracellular polymers are
produced, the cells irreversibly attach to the substratum (Sutherland, 1983). At this stage,
Escherichia coli synthesizes curli and colonic acid, which eventually become an integral
part of its biofilm matrix (Frank, 2009). Although repulsive forces usually prevent direct
bacterial contact with the surface, since both the cell surface and substratum are
negatively charged, contact still occurs due to bonding between bacterial appendages
(e.g., pili, fimbriae, flagella, and adhesion proteins) and the substratum by various shortrange forces, including dipole-dipole interaction, hydrogen, ionic and covalent bonding,
and hydrophobic interactions (Briandet et al., 1999; Kumar and Anand, 1998). Removal
16
of irreversibly attached cells requires the application of a strong shear force, such as
scrubbing and scrapping, and chemicals or heat which can break the attachment forces
(Chmielewski and Frank, 2003).
The degree of bacterial adhesion is influenced by physiochemical properties of
the bacterial cell and substratum surfaces, nutrient availability in the surrounding medium,
and the growth stage of the bacterial cells. Attached cells then begin to multiply while
sending out chemical signals that intercommunicate among the attached cells and
planktonic cells in the surrounding medium. Once the signal intensity exceeds a certain
threshold level, the genetic mechanisms underlying exopolysaccharide production are
activated (Chmielewski and Frank, 2003). Flagella may assist with recruitment of
planktonic cells into the biofilms by helping overcome repulsive forces between the cells
and surfaces. At this step, the cells multiply within the embedded exopolysaccharide
matrix, thus giving rise to formation of a microcolony, which is comprised of a large
number of cells with metabolic and physiological heterogeneity. Microcolonies will then
mature to form biofilms (Klausen et al., 2003).
Biofilms in nature usually have a high level of organization that allows diffusion
of nutrients and oxygen to underlying cell layers and diffusion of metabolic waste
products away from cells (Frank, 2001). Typical biofilm structures include columns of
cells separated by channels through which water flows, delivering nutrients and removing
waste products. The columns may further develop to mushroom-like shape filaments, or a
honeycomb-like network. However, the structural development of mixed-species biofilm
and food system biofilms that incorporate food soils needs further elucidation. Dispersion
of cells from biofilms is a serious concern of the food industry, because this process can
17
not only allow the daughter cells to contaminate food products, but also facilitates these
cells to colonize new niches (Sauer et al., 2002). External shear force, endogenous
enzymatic degradation, release of EPS, and starvation are all possible causes of biofilm
detachment (Kaplan et al., 2003; Kaplan et al., 2004; Stoodley et al., 2002).
Cells in biofilms are more resistant to cleaning and disinfection processes than
their planktonic counterparts (Frank and Koffi, 1990; Krysinski et al., 1992). Many
researchers have determined that no disinfectants are able to reduce the microbial
populations in a biofilm within the same time frame that is needed to inactivate
planktonic counterparts. Corcoran et al. (2014) reported that commonly used disinfectants,
including sodium hypochlorite (500 ppm), sodium hydroxide (1M), and benzalkonium
chloride (0.02%), failed to eradicate Salmonella biofilms on food contact surfaces. There
are many possible mechanisms that could account for the increased resistance of biofilms
to antimicrobial agents. The EPS, in which the cells are embedded, may retard the
penetration of antimicrobial agents by either reaction or sorption (Stewart et al., 2001;
Suci et al., 1994). The EPS plays a critical role in increasing bacterial resistance to
adverse environmental conditions and antimicrobial treatments, and this resistance is lost
when this structure is disrupted (Kumar and Anand, 1998). De Beer et al. (1994)
examined the penetration of chlorine into biofilms formed by Pseudomonas aeruginosa
and Klebsiella pneumoniae. Chlorine concentrations to which biofilms are exposed can
be only 20% of the concentration of the bulk liquid. Incubation for one to two hours did
not enable complete equilibration of the chlorine between the bulk phase and the biofilm.
After exposure to 2.5 ppm of chlorine for one hour, only upper 100 µm of a biofilm was
penetrated by the chlorine. The researchers determined that chlorine reacted with the EPS
18
mitigating its antimicrobial activity while it was moving through the biofilm matrix.
Stewart et al. (2000) reported that hydrogen peroxide was unable to penetrate the biofilm
formed by wild-type Pseudomonas aeruginosa. However, when catalase-deficient
mutants of Pseudomonas aeruginosa were used to form the biofilm, hydrogen peroxide
was able to penetrate the biofilm and inactivate all the resident cells. Even those
antimicrobial chemicals that were able to rapidly penetrate the biofilms failed to
inactivate cells in biofilms at the same rate as the planktonic counterparts (Cochran et al.,
2000; Zheng and Stewart, 2002). There may be some irreversible adsorption of the
antimicrobials to the EPS or a rapid reaction that inactivates the antimicrobial as it
diffuses through the biofilm (Stewart, 1996). Cells in biofilms may be growing more
slowly than planktonic cells (McDonnell and Russell, 1999), or some other phenotypic
change may make the microbes more resistant to disinfectants. However, Pan et al. (2006)
determined that, although Listeria cells that were grown in a biofilm were resistant to
peroxides, quaternary ammonium compounds (QACs) and chlorine when the cells were
removed from the surface, these cells had no increased resistance to these sanitizers.
Therefore, even if phenotypic changes do occur in cells in biofilms, the changes may be
not stable and removal of cells from the biofilm may cause them to become susceptible to
sanitizers. In addition, the central regulator of stress response, including rpoS and algT,
may also have a role in their increased resistance to sanitizers (Foley et al., 1999; Liu et
al., 2000).
Methods for imaging the structure of biofilms
Biofilm is a three-dimensional structure in which microorganisms and EPS are
distributed in a way that affects, and perhaps optimizes, their functions (Lewandowski
19
and Beyenal, 2009). In a dual-species biofilm, aerobic bacteria Ralstonia insidiosa
exhibit a tendency of dominating top layers, whereas facultative anaerobic Escherichia
coli O157:H7 mostly resides in the bottom layers. Such layered spatial distribution could
confer better protection for facultative Escherichia coli O157:H7 to various
environmental stresses than aerobic bacteria (Liu et al., 2014). Thus, to understand the
function and phenotypic behavior of biofilms, it is imperative not only to determine the
presence of various physiological groups of microorganisms within the biofilm, but also
to specify their spatial relations with respect to each other and with respect to the
substrata, which can be facilitated by the use of several types of microscopic techniques.
It is well believed that optical microscopy was invented by the Dutchman Antony
van Leeuwenhoek in the 17th century. The maximum magnification of his device was
400×, and the quality of his convex lens was limited. Since then, the improvement of
optical microscopy has been far reaching, with the maximum magnification of a modern
light microscope being 1000× (Bozzola and Russell, 1999). However, there are two main
factors restricting its use in the research of biofilms. First, the resolving power of the light
microscope is not only limited by the number and quality of the lenses, but also by the
wavelength of the light used for illumination. Second, out-of-focus objects degrade the
images of thicker specimens.
In the 1920s, accelerated electrons were discovered that they can travel in straight
lines in vacuum and have a wavelength which is about 100,000 times smaller than that of
light. Furthermore, it was determined that electric and magnetic fields have the same
effect on electrons as glass lenses and mirrors have on visible light (Lewandowski and
Beyenal, 2009). Based on these discoveries, transmission electron microscopy (TEM)
20
and scanning electron microscopy (SEM) were invented, and they are now the two main
types of electron microscopy that are widely used in the research of biofilms.
In TEM, an electron gun accelerates electrons, which are collimated by a
condensing lens and passed through a very thin sample. Objective and projector lenses
magnify the image and pass it to a fluorescent screen, or a photographic film (Bozzola
and Russell, 1999). The samples prepared for TEM examination must be fixed by
gluteraldehyde to crosslink proteins and by osmium tetroxide to fix lipids, dehydrated by
freeze drying or a series of increasingly concentrated ethanol solutions, infused with
acrylic or other liquid resin for solidification, thin-sectioned with a diamond knife, and
stained with heavy metal solutions, such as uranyl acetate or silver nitrate. The specimens
for TEM are embedded in a resin which physically stabilizes the EPS matrix (Walker et
al., 2001). However, TEM is not applicable for observing the extent and form of surfaceassociated growth of biofilms. In addition, the specimen must be very thin (< 0.5 µm) to
allow the electrons to penetrate it, but not all the specimens can be made thin enough for
examination by TEM.
The SEM has been used for high resolution visualization of bacterial biofilms
(Walker et al., 2001). In SEM, biofilm samples are prepared by fixation, drying, and
conductively coating prior to visualizing under high vacuum (Priester et al., 2007). A
narrow beam of electrons is moved back and forth across the specimen surface in a
rastering pattern. The beam is not transmitted as in TEM, but rather scattered from the
sample. SEM operates under a vacuum, hence water must be removed because it could
vaporize in the chamber. Biological samples are not conducive to scattering electrons,
hence they are usually sprayed with a thin layer of gold (~ 10 nm thick). A heavy metal
21
spray at an angle provides a shadow effect that is more like a 3D image. The sample stage
can be moved in different directions and even tilted to give the best image. However,
SEM examines only surface details. Additionally, the drying step in the sample
preparation can significantly alter the structure of biofilms due to EPS collapse (Kachlany
et al., 2001).
To overcome the limitations that each types of electron microcopy has, cryo-SEM,
environmental SEM (ESEM), and environmental TEM (ETEM) have been developed
recently. In traditional electron microscopy, volatile molecules such as water can
vaporize and interfere with the electron beam, so they must be removed from the sample.
In cryo-SEM, the sample is frozen so rapidly that there is little distortion of the sample.
Little water vapor can escape from the samples. The ESEM/ETEM allows operation at a
moderate vacuum without a conductive coating and high humidity (Danilatos, 1993). A
secondary electron capture detector can operate in the presence of water vapor. Samples
can be maintained in humid environments. Hence, sample preparation is much simpler
compared to that of traditional electron microscopy.
The application of confocal laser scanning microscopy (CLSM) has led to a better
understanding of the architecture of biofilms and their temporal development
(Nancharaiah et al., 2005). CLSM, in addition with fluorophores, the investigation in
real-time of living, hydrated microbial biofilms without structural destruction (Tomlin et
al., 2004). CLSM can eliminate out-of-focus haze, and it enables horizontal and vertical
optical sectioning, determination of three-dimensional relationships of cells, and 3D
computer reconstruction from optical thin sections (Lawrence et al., 1991). CLSM can be
used to measure the depth of biofilms, determine distinct viability of cells, and identify
22
specific species within the biofilm matrix by labelling cell surface components with a
fluorescent-staining solution (Frank, 2001). A popular method for labeling microbes is
using fluorescent protein (FP). FP-labeled bacteria have been widely used in monitoring a
cell population in a biofilm matrix (Nancharaiah et al., 2005; Tomlin et al., 2004). The
use of FP from the jellyfish Aequorea victoria began in the late 1960s and early 1970s,
and the green fluorescent protein was cloned and expressed in bacteria in the 1990s
(Chalfie et al., 1994; Prendergast and Mann, 1978). Since then, several other FPs have
been discovered and isolated from various organisms, such as the red fluorescent protein
from Aequorea victoria (Matz et al., 1999). Many FPs have been optimized by genetic
manipulation prior to the use in the CLSM. When stimulated with light with an
appropriate wavelength, the proteins emit fluorescent light of different colors, such as
green, orange, yellow and red, depending on the structure of the protein. The advantage
of labelling bacteria with FPs is that the FPs can be genetically expressed and tracked in
living cells. However, encoding FPs in originally FP-free wild-type cells can introduce a
metabolic burden to the cells that leads to decreased growth rate. Additionally, FPs need
oxygen to properly enfold, but the oxygen may not be available in deeper layers of a
biofilm matrix (Hansen et al., 2001).
Methods for biofilm control and removal
Surfaces
Surface materials of equipment used in food processing facilities have a large
effect on the level of bacterial attachment and biofilm formation that can occur
(Marouani-Gadri et al., 2009). Microorganisms can adhere to these surfaces, such as
stainless steel, floors, belts and rubber seals, and form biofilms after a period of time
23
(Costerton et al., 1995). Non-food contact surfaces can be a source of contamination
because microorganisms have the ability to transfer to food contact surfaces or foods by
liquids or aerosols in the environment. The materials made for surfaces must be hygienic,
and they should limit soil and microorganism accumulation and facilitate their removal
during cleaning. In addition, the materials must be resistant to mechanical action and
corrosion, especially during cleaning and disinfection processes (FDA, 2014).
A variety of materials are used in the construction of food equipment. These
materials vary in workability, cleanability, and sanitary design features. Among the
materials widely used in food processing facilities, stainless steel remains the first choice,
as it is not only resistant to corrosion in alkaline or acidic solutions, but is also highly
hygienic (Boulange‐Petermann et al., 1997). Faille et al. (2002) determined that
stainless steel was one of the less contaminated materials when challenged for Bacillus
spores adhesion. The 300 series (iron-chromium-nickel) of stainless steel is
recommended by the American Iron and Steel Institute (AISI) for food contact surfaces.
The most common alloys AISI 304 and 316 are durable in typical food processing
environments, and in domestic kitchens. 3A Sanitary Standards require 316 grade
stainless steel for most surfaces in the food industry, and 304 grade for utility usage. The
different grades can be found in different surface finishes according to the treatment to
which the stainless steel has been subjected. When stainless steel surfaces do not comply
with required roughness values, a subsequent polishing treatment on one or both sides
may be applied. The #4 finish is recommended in the US for stainless steel that comes
directly into contact with food products (Faille and Carpentier, 2009).
24
Two major strategies have been developed to control microbial attachment on
surfaces, i.e., the limitation of bacterial contamination and the inactivation of adherent
microorganisms. The first strategy is to produce anti-adhesive materials which would be
resistant to microbial adhesion. Manipulation of surface energy or topography has been
investigated in order to meet this requirement. The use of hydrophobic coatings (low
surface energy) on glass or of coatings with intermediate surface energies of
approximately 24 mN/m can reduce adhesion of some bacteria, but not of the highly
hydrophobic Bacillus cereus spores (Rosmaninho et al., 2007; Tsibouklis et al., 1999;
Zhao, 2004). Due to thermodynamic reasons, and to the fact that surfaces are quickly
conditioned in real environments, it would be very difficult to develop one surface which
would exhibit absolute anti-adhesive properties to all microbes. Another strategy is to
incorporate antimicrobial agents into materials. The antimicrobial compounds should be
selected to provide a broad activity against target microorganisms and maintain their
antimicrobial activity for long periods of time. Among the inorganic agents, silver is most
commonly selected since it is safe for humans, but has strong antimicrobial properties, as
it interferes with electron transport causing protein and DNA damage (Srey et al., 2013).
A main advantage of the use of inorganic compounds to limit the microbial
contamination of surfaces is that the reaction between the microorganism and inorganic
compounds is highly non-specific, making it rare for microorganisms to develop
resistance.
Cleaning
The best way to control biofilms is effective cleaning so as to not allow the cells
to firmly attach to surfaces (Simões et al., 2006). Sanitizing alone cannot control biofilms
25
if there are no effective cleaning operations. Effective cleaning has a major impact on
biofilms. It removes the cells in a biofilm, helps clean away the EPS produced by the
biofilm, removes soils that can adversely affect sanitizer and disinfectant activity, and
may even make microorganisms more susceptible to sanitizers. Four factors, including
time, mechanical action, chemical, and temperature, should be controlled during cleaning
in order to obtain a satisfactory total performance.
In general, the more time that is spent cleaning a surface, the cleaner a surface
will be. This can be accomplished by circulating a cleaning solution through a closed
system such as a clean-in-place (CIP) process, or soaking soiled items in a detergent
solution in a clean-out-of-place (COP) tank (Srey et al., 2013). Foam cleaners are
commonly used for general environmental cleaning in food processing facilities since
they have higher viscosity and will therefore cling to surface longer compared to their
liquid counterparts (Chen et al., 2014).
Another variable that can be controlled to influence cleaning is mechanical action.
The more energy input into cleaning, the cleaner a surface will become. Manual
scrubbing, automated impingement, high pressure spray, turbulent flow in CIP systems,
and pumping cleaning solutions through properly positioned spray balls are all the ways
to increase mechanical action when cleaning. Solid evidence provided by many
researchers reveals that mechanical action and chemical treatments can destroy biofilms
(Gibson et al., 1999; Schwach and Zottola, 1984; Wirtanen et al., 1996). However, too
much or improper mechanical action on a surface may damage the surface and
inadvertently spread the biofilm cells by creating aerosols of microbes which may result
in cross contamination to other surfaces (Grinstead, 2009).
26
The type of chemicals used to clean a surface has a tremendous impact on
cleaning effectiveness. Cleaners are formulated to clean particular soils and surfaces
under certain conditions. Most cleaning agents used in the food industry are alkali
compounds that are detergents for fat and protein. Chelators can be formulated into
cleaners to remove mineral ions. Addition of chelators, such as EDTA, in alkali cleaners
was more effective than acid cleaning solutions in removing biofilms (Wirtanen et al.,
1996). The concentration of cleaners also has a large impact on cleaning performance.
Typically, a higher concentration of cleaner will remove more soil than a lower
concentration, but most cleaners have an optimum concentration. Cleaning compounds
must be formulated with care, because many components are incompatible or most
effective if applied separately. Surfaces like glass, plastics and ceramics are suggested to
be cleaned with alkali or nonionic detergents (Lewis, 1980).
The use of high temperatures in cleaning can reduce the energy input, such as
water turbulence and scrubbing (Maukonen et al., 2003). Application of cleaning
solutions at temperatures between 40°C and 90°C is widely recommended. Redeposition
of soil can occur if the temperature is not high enough. However, proteins, in particular,
can be more difficult to remove at very high temperatures. If the cleaning temperature is
high enough to denature proteins, it can make them more difficult to remove and the
proteins may cook onto or attach more firmly to the surfaces. In addition, the chemistry
of cleaners and the surface being cleaned also affect the optimum cleaning temperature.
Cleaners containing volatile components or enzymes can be inactivated by excessive
temperatures (Grinstead, 2009).
27
In most food processing facilities, food contact surfaces are cleaned and sanitized
every day, but many environmental surfaces such as walls, ceilings, storage tanks, pump
and valve exteriors are cleaned infrequently. These locations provides bacteria favorable
niches to form biofilms if moisture (e.g. condensation) and nutrients are present (Frank,
2009). Thus, an ideal cleaning scheme should include environmental zones and be
effective in breaking up biofilm matrixes, whereby sanitizing agents can access and kill
the cells.
Sanitizing
Cleaning procedures enable the removal of approximately 90% of bacteria from
surfaces, but cannot be relied on to kill them (Srey et al., 2013). The detached cells might
later reattach to other surfaces and form a biofilm given time, water and nutrients. The
implementation of a sanitizing procedure is indispensable in controlling biofilms in food
processing facilities (Gram et al., 2007).
The most commonly used classes of sanitizers are oxidizing sanitizers, including
halogens such as chlorine, chlorine dioxide, and iodine. Other oxidizing agents include
peroxides such as hydrogen peroxide and peracetic acid (PAA, CH3COOOH). Sodium
hypochlorite (NaClO) is an effective disinfectant for biofilm inactivation (Ozdemir et al.,
2010). Hypochlorous acid (HClO, pKa = 7.4) is the active moiety responsible for the
bactericidal activity of hypochlorite solution and it is more stable in solutions between
pH 4 and 7. Hypochlorous acid disrupts oxidative phosphorylation and other membraneassociated activity (Barrette et al., 1989; Camper and McFeters, 1979). De Beer et al.
(1994) reported that chlorine is inactivated by EPS of the mixed-species biofilms formed
by Pseudomonas sp. and Klebsiella sp. since it failed to fully penetrate the matrix of a
28
400-µm thick biofilm. Compared to QACs and iodine, chlorine is better at removing the
EPS in biofilms formed by Listeria and Salmonella on stainless steel (Ronner and Wong,
1993). Increasing the contact time of chlorine sanitizers from 5 to 30 min can
substantially enhance their antimicrobial efficacy against biofilms formed by
Pseudomonas on stainless steel (Gélinas et al., 1984). Hypochlorous acid is unstable, so it
must be produced at the point of use. Additionally, hypochlorous acid can be inactivated
by metals such as copper or nickel, and ultraviolet light, and it can react with organic
matter. Although iodine is an effective disinfectant, its tendency to discolor surfaces
limits its use in food processing facilities (Grinstead, 2009). Peroxygen compounds have
also been used as sanitizers in the food industry. Some nearly unique properties of
hydrogen peroxide are its low toxicity and that it does not cause allergic reactions
(Rideout et al., 2005). Hydrogen peroxide possesses a highly oxidizing capacity that is
based on the production of free radicals which attack essential cell components, including
lipids, proteins, and DNA (McDonnell and Russell, 1999). Hydrogen peroxide is
effective against biofilms (de Carvalho, 2007; de Carvalho and da Fonseca, 2007).
However, one of the disadvantages of hydrogen peroxide is that it is a natural byproduct
of aerobic metabolism, therefore most organisms that respire have enzymes (e.g., catalase
and superoxide dismutase) that can detoxify it. A peroxygen compound that is commonly
used in food processing plants is peracetic acid (PAA). Compared to hydrogen peroxide,
PAA is a more potent disinfectant, and enzymatic inactivation of PAA is not a concern.
Hence, PAA is biocidal at lower concentrations than hydrogen peroxide. PAA also
maintains its efficacy in the presence of organic matter (McDonnell and Russell, 1999).
Peroxide-based sanitizers are more effective against Listeria monocytogenes and
29
Salmonella in biofilms than is hypochlorite (Fatemi and Frank, 1999; Härkönen et al.,
1999).
The other broad class of sanitizers is based on surfactants. QACs are cationic
surfactants that have biocidal activity against vegetative bacteria, molds, and yeast, but it
is not strongly active against spores (McEldowney and Fletcher, 1987). QACs are
membrane-active agents with a target site predominantly at the cytoplasmic membrane in
bacteria or the plasma membrane in yeasts (Hugo and Frier, 1969). QACs have reduced
activity in the presence of hard water, but they are very stable and are not rapidly
inactivated by organic matter. Therefore, QACs are recommended to be applied for long
contact time on non-food contact surfaces, such as floors, walls, and storage tanks.
However, QACs are not very effective against biofilms since the compounds could not
fully inactivate Listeria monocytogenes in biofilms within 20 min at a concentration of
800 ppm (Frank and Koffi, 1990).
Other methods
For equipment with poor design or that contains niches where nutrients can
accumulate but cleaning and sanitizing agents cannot adequately access, a heat treatment
may have the potential to control biofilms formed by foodborne pathogens. Chmielewski
and Frank (2004) developed a model which could predict the reduction of Listeria
monocytogenes in biofilms on stainless steel by heat treatment. The susceptibility of
Listeria monocytogenes in biofilms to inactivation by heat was strain specific. Effective
cleaning is still important when heat is used to eliminate biofilms after cleaning. The
presence of food soils can significantly increase the heat resistance of some strains of
Listeria monocytogenes.
30
Radiation is also used by some researchers to control biofilms. Niemira and
Solomon (2005), and Niemira (2007) found that some strains of Salmonella and
Escherichia coli O157:H7 had increased sensitivity to ionizing radiation compared to
their planktonic counterparts, whereas others had decreased sensitivity. Thus, the
sensitivity of bacteria in biofilms to ionizing radiation may depend on the specific strain,
hence it is difficult to predict how a biofilm will respond to a radiation. UV radiation is
not able to penetrate a biofilm due to its poor penetrating power. Alginate, a common
component of the EPS of Pseudomonas biofilms, only transmits less than 1/3 of the UV
light to which it is exposed (Elasri and Miller, 1999). UV radiation is not able to
penetrate a biofilm so would not be a very effective biocide, especially against microbes
that are deep inside a biofilm. The antimicrobial efficacy of both ionizing and photonic
radiations against cells in biofilms needs more study.
Enzymes may be useful in biofilm control. Since EPS is a heterogenic matrix, a
combination of enzymes is required to degrade the complex (Simões et al., 2010).
Lactoperoxidase and glucose oxidase enzymes were used together and they reduced 3 and
2 logs of Pseudomonas and Staphylococcus, respectively, in a biofilm grown on stainless
steel. A mixture of polysaccharide hydrolyzing enzymes removed biofilms from stainless
steel and polypropylene, but did not have significant bactericidal activity (Johansen et al.,
1997). The performance of enzymes can be enhanced by the inclusion of surfactants and
chelating agents, as the combination revealed enhanced efficiency in removing the
biofilms formed by Bacillus and Pseudomonas fluorescens (Lequette et al., 2010).
Another possible biofilm control method is bacteriophage. Pretreatment of the
substratum with bacteriophage that could infect Staphylococcus epidermidis reduced
31
nearly 4.5 log CFU of the pathogen (Curtin and Donlan, 2006). One of the advantages in
using bacteriophage is that the EPS cannot prevent the phage from entering the biofilm
matrix and diffusing through it (Briandet et al., 2008). Some phage may even induce
production of enzymes that degrade EPS polymers. Hughes et al. (1998) reported that a
phage can possess a polysaccharide depolymerase that is specific for the EPS produced
by the bacteria that the phage could infect. The phage was able to disrupt a biofilm by a
combination of degradation of the EPS and lysis of the cells in the biofilms.
Zhao et al. (2006) evaluated the efficacy of two lactic acid bacterial isolates
(Lactococcus lactis subsp. lactis C-1-92 and Enterococcus durans 152) that produced
antimicrobials for reducing Listeria spp. contamination of floor drains in a poultry
processing plant. The drains had detectable Listeria in biofilms before application of the
competitive exclusion (CE) microorganisms. After the drains were treated with 107
CE/ml in a cleaning agent for 10 times in 4 weeks, two of the drains no longer had
detectable Listeria and the other three had reduced Listeria levels. The Listeria
reductions for all five drains tested ranged from 2.3 to 4.1 log CFU/100 cm2. Although
the Listeria levels in the drains were reduced, the total aerobic microbial load in the
drains was not changed. CE may have colonized the drains and not only reduced Listeria
spp. populations, but also subsequently formed their own biofilms and controlled
bacterial populations that were sensitive to the antagonistic metabolites of the CE
(Borucki et al., 2003; Costerton et al., 1999; Goeres et al., 2005; Toledo-Arana et al.,
2001). Although more studies are needed to determine how well the CE microorganisms
can persist in the environment and how they would respond to recontamination with
Listeria or other pathogens, this approach provides a way to address biofilms in those
32
areas where the problem is not the biofilm itself, but rather the constituents of that
biofilm (Grinstead, 2009).
Levulinic acid and sodium dodecyl sulfate
Levulinic acid (4-oxopentanoic acid, C5H8O3, molecular mass: 116.12 Da) is a 5carbon organic acid which has been designated as Generally Recognized as Safe (GRAS,
21 CFR, 172.515) by FDA for direct addition to food products as a flavoring agent
(Wang et al., 2012). It can be produced at a low cost and in a high yield from renewable
feedstocks (Bozell et al., 2000b). Levulinic acid is used as a cigarette additive to
desensitize the upper respiratory tract, which can mask the irritation caused by smoke and
increase the potential for cigarette smoke to be inhaled deeper into the lungs (Keithly et
al., 2005). Addition of sodium levulinate in RTE meats was effective at inhibiting growth
of spoilage bacteria and Listeria monocytogenes without noticeable different flavor
(Thompson et al., 2008; Vasavada et al., 2003). Salmonella Enteritidis populations in
pure culture held at 21°C were reduced by 3.4 log CFU/ml when exposed to 0.3%
levulinic acid for 30 min (Zhao et al., 2009). The application of levulinic acid in fresh
produce may extend shelf life because the organic acid can arrest light-induced
chloroplast development during greening and can be removed by washing the leaves to
restore the developmental process without any apparent toxic effect (Jilani et al., 1996).
There were no visual differences in leafy colors between lettuce treated with the
combination of levulinic acid and sodium dodecyl sulfate (SDS) and lettuce rinsed with
water only (Zhao et al., 2009). The antimicrobial activity of levulinic acid could be
attributed to the reduction of medium pH, decrease of the cytoplasmic pH of microbes by
ionization of undissociated acid molecules, chelation of metal ions, disruption of
33
substrate transport by altering cell membrane permeability and/or reduction of proton
motive force (Cannon et al., 2012; Eswaranandam et al., 2004; Kreske et al., 2008;
Raybaudi-Massilia et al., 2009).
SDS is also designated as GRAS by FDA for multipurpose additives (21 CFR
172.822). It is an anionic surfactant and is widely used in household products such as
toothpastes, shampoos, shaving creams, and bubble baths. SDS is approved for use in a
variety of foods, including egg whites, fruit juices, vegetable oils, and gelatin as a
whipping or wetting agent (Zhao et al., 2009). The SDS molecule has a 12-carbon tail
attached to a sulfate group, giving the molecule amphiphilic properties. SDS can denature
proteins and damage cell membranes, and its bactericidal effect can be increased when
pH is reduced to between 1.5 and 3.0 (Anderson et al., 1990; Byelashov et al., 2008;
Williams and Payne, 1964). The detergent properties of SDS is able to facilitate
detachment of cells from surfaces, thereby making the cells more likely to be inactivated
by other disinfectants that are present. However, static treatment of noroviruscontaminated stainless steel surfaces with water containing 2% SDS was no more
effective at inactivating noroviruses than treatment with water alone (Cannon et al., 2012).
The application of levulinic acid and SDS alone had limited antimicrobial
efficacy. However, combining levulinic acid with SDS greatly increased the bactericidal
activity of these two chemicals. Application of 3% levulinic acid for 2 min to pure
cultures was shown to reduce Escherichia coli O157:H7 by 1.7 log, whereas treatment
with 0.5% levulinic acid plus 0.05% SDS for less than 1 min reduced the population of
all Shiga toxin-producing Escherichia coli (STEC) strains tested, including E. coli
O157:H7, to an undetectable level (> 6-log reduction) (Zhao et al., 2014). At higher
34
concentrations of 3% levulinic acid plus 2% SDS, more than 4-log reduction of
Salmonella on treated chicken carcasses was achieved after a 5-min immersion at 21°C
(Zhao et al., 2011). Stelzleni et al. (2013) reported that lower concentrations of 1%
levulinic acid plus 0.1% SDS had a great effect on reducing Salmonella than either 2%
(w/v) liquid buffered vinegar or 2.5% (w/w) powdered buffered vinegar, but levulinic
acid and SDS-treated beef patties had increased growth of psychrotrophic organisms and
reduced color scores after 3 days of retail display. Although levulinic acid and SDS are
very bactericidal, there are some factors that can interfere with their combined efficacy.
First, the antimicrobial effect of levulinic acid and SDS is concentration dependent.
Generally, higher concentrations of both levulinic acid and SDS achieve greater
inactivation of bacterial populations (Zhao et al., 2014). Second, reduction of bacteria is
directly related to the temperature, with more reduction obtained at higher temperatures
and minimal bacterial inactivation at refrigeration temperature (Stelzleni et al., 2013;
Zhao et al., 2014; Zhao et al., 2009). Third, the treatment followed by mechanic actions,
such as rubbing, is generally more effective in inactivating bacteria than the chemical
treatment only (Zhao et al., 2014).
The combination of levulinic acid and SDS exhibits desirable properties besides
its high antimicrobial efficacy. First, both of the two chemicals are Generally Recognized
as Safe (GRAS). Levulinic acid has a desensitizing effect so it does not cause irritation to
the users (Wang et al., 2012). Additionally, after a treating alfalfa seeds with 0.5%
levulinic acid plus 0.05% SDS for 30 min, the seeds germinated at an equivalent rate
compared to the control seeds treated with deionized water (Zhao et al., 2009, 2010).
Second, levulinic acid can be cost effective. Levulinic acid can be produced from
35
cellulose-containing waste materials and thereby costs can be low (Wang et al., 2012).
The estimated cost of a 3% levulinic acid preparation is 20 cents per liter based on
biosynthesis (Bozell et al., 2000a), and because the bactericidal activity of the levulinic
acid plus SDS treatment is not readily neutralized in the presence of organic matter,
unlike many chemical treatments such as sodium hypochlorite, the levulinic acid plus
SDS treatment can be reused to minimize cost and water usage (Zhao et al., 2011). Third,
the formulation is readily soluble in water at ambient temperature, a property that allows
it to be used in different formats such as solutions, pastes, gels, and foams. The format of
cleaning and sanitizing chemicals can influence the amount of contact time, and
eventually affect the final antimicrobial efficacy. For example, foaming solutions
generally have higher viscosity and therefore cling to surfaces longer. Application of this
solution as a foam on contaminated deli slicers substantially reduced larger populations
of Salmonella, Listeria monocytogenes, and Escherichia coli O157:H7 than the liquid
treatment (Chen et al., 2014). Fourth, the antimicrobial activity of the combination of
levulinic acid and SDS is not mitigated by the presence of organic materials. More than 5
log CFU of Salmonella and Escherichia coli O157:H7 were reduced on fresh produce,
chicken wings and skin, and in an organic-rich environment containing fecal matter or
feathers (Zhao et al., 2011). The combination of levulinic acid and SDS was incorporated
into FIT® L Food and Vegetable Wash products by a University of Georgia Research
Foundation’s licensee in 2010. It was granted a utility patent by the U.S. Patent and
Trademark Office (patent NO. 8,722,123) in 2014 (Doyle and Zhao, 2014).
Although the combination of levulinic acid and SDS has substantial antimicrobial
activity, currently levulinic acid plus SDS do not have regulatory approval for application
36
as a sanitizer. Additional studies are needed to validate the efficacy of levulinic acid and
SDS in actual production operations (Zhao et al., 2009). Guan et al. (2010) claimed that
although the treatments inhibited cut edge browning of lettuce pieces that develops
during storage, levulinic acid (0.5% to 3%) and 0.05% SDS caused detrimental effects on
the visual quality and texture of lettuce. Levulinic acid and SDS-treated samples were
sensorially unacceptable due to the development of sogginess and softening after 7 and
14 days of storage. The researchers found the combination caused an increase in the
respiration rate of fresh-cut lettuce as indicated by higher CO2 and lower O2 in modified
atmosphere packages.
Despite the extensive research already completed on the spectrum of activity and
the levels required for successful inactivation of foodborne bacteria, more research is
needed to better elucidate the mechanisms of antimicrobial activity of levulinic acid and
SDS, the efficacy of the combination applied on food products at concentrations that do
not have adverse sensory effects, as well as an approach to effectively reduce the cost of
the application of levulinic acid and SDS in actual food processing facilities.
37
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60
CHAPTER 3
TRANSFER OF FOODBORNE PATHOGENS DURING MECHANICAL SLICING
AND THEIR INACTIVATION BY LEVULINIC ACID-BASED SANITIZER ON
SLICERS1
1
Chen, D., T. Zhao and M.P. Doyle. 2014. Food Microbiology. 38:263-269.
Reprinted here with permission of the publisher.
61
ABSTRACT
This study investigated the degree of cross-contamination between deli foods and
slicers by Listeria monocytogenes, Salmonella, and Escherichia coli O157:H7, and their
inactivation by levulinic acid (LVA) plus sodium dodecyl sulfate (SDS) on slicers. The
transfer rate of pathogens at 5 locations on the contaminated slicers (scenario I) and on
food slices (scenario II) was determined. The antimicrobial efficacy of the LVA + SDS
sanitizers applied either as a liquid or as a foam at three concentrations (0.5% LVA +
0.05% SDS, 1% LVA + 0.1% SDS, and 2% LVA + 0.5% SDS) was determined for
decontamination of the pathogens on the slicers at 21oC. After slicing 10 slices, the
pathogens recovered from slicer blades were significantly (P < 0.05) less than the
recovery from some other contact locations (scenario I). With an initial inoculum at
approximately 8.5 log CFU/blade, the populations of the pathogens transferred from
blades to slices decreased logarithmically (R2 > 0.9, scenario II). Contaminated slicer
surfaces sprayed with 1% LVA plus 0.1% SDS as a foam (45-55 psi) reduced within 1
min 6.0 to 8.0 log CFU/blade of the pathogens. Results revealed that cross-contamination
can occur between deli foods and slicers. Also, LVA-based sanitizer applied as foam can
be a useful treatment to remove microbial contamination on the slicers.
Key words: slicer, deli, transfer, levulinic acid, Listeria, Salmonella, Escherichia coli
62
1. Introduction
Listeria monocytogenes causes listeriosis, a human disease with a fatality rate as
high as approximately 16% (FDA, 2013). This psychrotrophic microorganism is widely
distributed in the retail food establishments, and can reside in some food processing
facilities for more than 1 year (Sauders et al., 2009; Tompkin, 2002). Deli meats have
been reported as the leading vehicle of foodborne listeriosis in the United States (Zhang
et al., 2012). A risk assessment by the U.S. Food and Drug Administration (FDA) and
U.S. Department of Agriculture-Food Safety and Inspection Service (USDA-FSIS) of
ready-to-eat (RTE) foods for the risk of acquiring listeriosis revealed that deli meats were
the food source of greatest risk (FDA and USDA, 2003). Three major listeriosis
outbreaks documented in the United States during the past two decades have been traced
to consumption of contaminated sliced deli meat (CDC, 1999; Gottlieb et al., 2006; Olsen
et al., 2005). Many listeriosis outbreaks have also been associated with consumption of
contaminated cheese (Cartwright et al., 2013). Salmonella and E. coli O157:H7 are also
major foodborne pathogens in the United States (Scallan et al., 2011). RTE salami was
the source of recent Salmonella and E. coli O157:H7 outbreaks, causing a total of 311
cases (CDC, 2010; Williams et al., 2000).
Slicers are commonly used in the deli department of retail food establishments
and have been known for many years to serve as a vehicle for cross-contaminating deli
foods with foodborne pathogens (Gilbert and Maurer, 1968). Outbreaks of L.
monocytogenes, Salmonella and E. coli O157:H7 infections have been associated with
deli food products, and cross-contamination by these pathogens during slicing was
suspected as the mode of transmission (Anonymous, 2007; Campbell et al., 2008;
63
Williams et al., 2000). The risk of cross-contamination of slicers is increased by contact
with contaminated food surfaces because foodborne pathogens can be transferred
between contaminated foods and food contact surfaces (Carrasco et al., 2012; PérezRodríguez et al., 2008). Moreover, the slicer itself could serve as the source of foodborne
pathogens. Working ambiance and physical complexity are the two main contributing
factors which make slicers prone to contamination. Unlike other food processing
equipment, deli slicers are mostly used in a random and intermittent way throughout
working days and are held at ambient temperature (Sheen and Hwang, 2010). The area of
food contact surfaces on deli slicers is large and exposed to an environment where L.
monocytogenes can be widely distributed (Endrikat et al., 2010; Pradhan et al., 2010;
Sauders et al., 2009). Sealants and gaskets are widely used in slicers to seal seams and
gaps between connecting parts. With long-term use and repeated cleaning, cracks, chips,
shrinkage or loss of the sealant and gasket may occur, resulting in compromise of the
seam integrity. Food soils can accumulate in these open seams, which are nearly
impossible to reach and clean, leading to a desirable location for microbial growth (FDA,
2011a, b).
Routine and proper cleaning and sanitizing procedures are thought to be effective
in preventing cross-contamination of deli slicers (Lin et al., 2006). Proper cleaning is
needed to remove organic matter before applying sanitizers, but failure of disinfection
can occur due to poor employee training in equipment cleaning (Neal, 2013) and some
locations of slicing equipment used long term are difficult to clean (Vorst et al., 2006).
Quaternary ammonium-based sanitizers are commonly used by the industry to sanitize
slicers, but their efficacy in killing foodborne pathogens is generally reduced in
64
commercial settings when organic material is present (Simpson Beauchamp et al., 2012).
Hence, it would be beneficial to develop a sanitizer that possesses stable disinfectant
activity in the presence of organic materials. A bactericide containing levulinic acid plus
sodium dodecyl sulfate (SDS), was previously determined to be an effective sanitizer in
the presence of organic matter, including chicken feces, feathers, and feed (Zhao et al.,
2011). In addition, both levulinic acid (FDA 2008, 21 CFR, 172.515) and SDS (FDA,
2007, 21 CFR, 172.822) were individually designated by FDA as Generally Recognized
as Safe (GRAS) for specific uses in foods.
Although studies of the transfer of L. monocytogenes between slicing equipment
and deli foods have been reported previously (Aarnisalo et al., 2007; Keskinen et al.,
2008; Lin et al., 2006; Sheen, 2008; Sheen et al., 2010; Sheen and Hwang, 2008; Vorst et
al., 2006), transfer of Salmonella and E. coli O157:H7 in slicing scenarios has not been
well documented. In addition, deli slicers are still being identified as the source of L.
monocytogenes cross-contamination to RTE foods according to a recent FDA report
(FDA, 2013). Since recent outbreaks have been associated with these three pathogens in
sliced deli food products, a study on the transfer of these pathogens on deli slicers and
how to better sanitize slicing equipment can provide insightful information for the
prevention of cross-contamination of pathogens to slicers. This study was designed to
determine: (1) the degree of cross-contamination by L. monocytogenes, S. Typhimurium,
and E. coli O157:H7 between deli food products and slicers, and (2) the efficacy of
levulinic acid plus SDS for inactivating these three pathogens on slicers.
65
2. Materials and methods
2.1. Bacterial strains
Five strains of each pathogen were used in this study. The five strains of L.
monocytogenes were LM101 (serotype 4b, salami isolate), LM112 (serotype 4b, salami
isolate), LM113 (serotype 4b, pepperoni isolate), H9666 (serotype 1/2c, human isolate),
and ATCC 5779 (serotype 1/2c, cheese isolate); the five isolates of S. Typhimurium
DT104 were H2662 (cattle isolate), 11942A (cattle isolate), 13068A (cattle isolate),
152N17-1 (dairy isolate), and H3279 (human isolate); and the five strains of E. coli
O157:H7 were 932 (human isolate), E009 (beef isolate), E0018 (cattle isolate), E0122
(cattle isolate), and E0139 (deer jerky isolate). All the five Salmonella strains were
ampicillin (32 µg/ml), streptomycin (64 µg/ml) and tetracycline (16 µg/ml) resistant,
whereas the E. coli O157:H7 strains were nalidixic acid (50 µg/ml) resistant. All strains
of the three pathogens were from University of Georgia Center for Food Safety culture
collection and stored at -20°C in vials containing 25% glycerol (Sigma, St. Louis, MO)
plus either brain heart infusion broth (BHI, Becton Dickinson, Sparks, MD) for L.
monocytogenes or tryptic soy broth (TSB, Becton Dickinson) for S. Typhimurium and E.
coli O157:H7. All cultures were maintained on plates of modified Oxford agar (MOX,
Becton Dickinson), xylose lysine deoxycholate agar (XLD, Becton Dickinson) with 32
µg/ml ampicillin, 64 µg/ml streptomycin and 16 µg/ml tetracycline, and Sorbitol
MacConkey agar (SMAC, Becton Dickinson) with 50 µg/ml nalidixic acid for L.
monocytogenes, Salmonella and E. coli O157:H7, respectively, and subcultured monthly.
Each strain of the three pathogens was cultured in 10 ml of BHI or TSB individually with
its specific antibiotic(s) to which it is resistant, and incubated at 37°C for 20 h. For each
66
pathogen, the optical density (OD) of each strain was adjusted in a spectrophotometer
(model 4001/4, Spectronic Instruments, Rochester, NY) with 0.1 M phosphate buffered
saline (PBS) to an OD reading of 0.9 (ca. 9.0 log CFU/ml) at 630 nm. Approximately the
same cell number of each strain of the three pathogens was combined to obtain three
bacterial mixtures. Cell numbers in the mixtures were determined by spread plating serial
dilutions (1:10 in 0.1% peptone water) onto plates as described above. The plates were
incubated at 37°C for 24 (Salmonella and E. coli O157:H7) to 48 h (L. monocytogenes)
and typical colonies were counted.
2.2. Slicers
Three retail-scale, gravity-fed slicers (model SE 12, Bizerba, Piscataway, NJ)
were used in this study. This slicer model was selected after surveying the deli
departments of local grocery stores. The main body of the slicers was forged with one
piece of anodized aluminum. The blade equipped on the mechanical slicers was 13 in. (33
cm) in diameter, running at 266 revolutions per minute (rpm) and made of chromiumcoated hard alloy. The three slicers were new and used for the first time in this study.
Before and after each use, slicers were cleaned with detergent, rinsed with sterile distilled
water, and treated with 70% ethanol. After ethanol evaporation (ca. 20 min), sterile
distilled water was sprayed on the slicers to remove the ethanol residual. The slicers were
air dried at 21°C for 24 h before use.
2.3. Transfer of the pathogens from inoculated deli foods to food contact locations on
slicers
Deli Swiss cheese (Prima Della, Bentonville, AR), roasted black forest ham
(Hormel Foods, Austin, MN), and roast beef (Charlie’s Pride Meats, Vernon, CA) were
67
purchased from a local grocery store, held at 4°C, and used within 1 week. To study the
transfer of pathogens from contaminated deli foods to clean slicers, the Swiss cheese,
ham, and roast beef were surface-inoculated with L. monocytogenes, S. Typhimurium,
and E. coli O157:H7 mixtures, respectively. The cell numbers of three bacterial mixtures
were diluted to 5.0 log CFU/ml by serial dilution (1:10 in 0.1% peptone water) and
confirmed by the direct plating method as described above. The diluted mixtures (5 ml)
of L. monocytogenes, S. Typhimurium, and E. coli O157:H7 were then evenly spread on
the surface of Swiss cheese, ham, and roast beef, respectively, by using a hockey stick.
The inoculated deli foods (ca. 3.0 log CFU/cm2) were placed in a laminar air flow hood
for 20 min allowing attachment of the pathogens, and were subsequently placed into 2.5gallon (9.5 liters) Ziploc bags. The surface-inoculated samples were held at 4°C for 24 h
before use. The Swiss cheese, ham and roast beef were sliced at a thickness of 1 to 2.5
mm and a weight of 10 to 20 g. After slicing 10 slices, the food contact areas of the
slicers were swabbed from approximately 10 cm2 with sterile 6-inch (15.2 cm) polyestertipped swabs (Fisher Scientific, Pittsburgh, PA) which were pre-saturated with 100 µl of
sterile 0.1 M PBS. The swabbing locations were identified based on our previous
research (Zhang et al., 2006) and included: 1, meat grip; 2, carriage tray; 3, gauge plate; 4,
slicer blade; and 5, blade cover. Each swab was then dipped in 500 µl of 0.1 M PBS, pH
7.25, agitated by a Vortex (G-560, Scientific Industries, Bohemia, NY) for 10 s, and 0.1
ml of undiluted and 1:10 dilutions in 0.1 M PBS were spread plated in duplicate on MOX
(for L. monocytogenes), XLD (for S. Typhimurium), and SMAC (for E. coli O157:H7)
plates with specific antibiotic(s). The plates were incubated at 37°C for 24 to 48 h, and
colonies typical of the selected pathogens were counted.
68
2.4. Transfer of the pathogens from inoculated blades to food slices
The three bacterial mixtures were separately inoculated on the front side of the
blades of the three slicers. For each pathogen, a 5-strain mixture (300 µl, ca. 9.0 log
CFU/ml) was applied on the blade by drop (10 µl/drop) inoculation with a total of 10
drops being applied as an inoculum. Three sets of the 10-drop inoculum were applied on
the slicer blade at an approximately 120° distance from each other. Each 10-drop
inoculum was 2 cm in width and 5 to 10 cm in length on the rim of the round blade. The
inoculated slicer blades were allowed to dry at 21oC for 24 h before being used for slicing.
The deli foods were sliced according to the protocol previously described. Slices were
selected for sampling at intervals of each 3 slices. Thus, the 1st, 4th, 7th… 58th and 61st
slices were selected for enumeration of the pathogens and each slice was individually
placed into a 24-oz (709.8 ml) Stomacher bag (Nasco Whirl-Pak, Fort Atkinson, WI)
with an equal weight of 0.1% peptone water. The content of each Stomacher bag was
subsequently pummeled for 1 min at 230 rpm (Stomacher model 400, Seward Ltd.,
Westberry, NY). Thereafter, 2 ml or 200 µl of undiluted and 1:10 dilutions in 0.1%
peptone water of each sample were plated in duplicate onto MOX, XLD, and SMAC
plates with specific antibiotic(s). All the plates were incubated at 37oC for 24 or 48 h
before colony enumeration.
2.5. Sanitizer efficacy determinations
The three pathogens were inoculated as individual mixtures on slicer blades
according to the protocol described above. The inoculated slicers were allowed to air dry
at 21oC for 24 h before sanitizer was applied. The blade cover of each slicer was
disassembled and removed for the study. Different concentrations (0.5% LVA + 0.05%
69
SDS; 1% LVA + 0.1% SDS; and 2% LVA + 0.5% SDS) of levulinic acid (LVA) plus
sodium dodecyl sulfate (SDS) were applied as a liquid or foam on the slicer blades to
determine decontamination efficacy in comparison with a commercial quaternary
ammonium-based sanitizer (150 ppm, 2.250% octyl decyl dimethyl ammonium chloride,
1.125% dioctyl dimethyl ammonium chloride, 1.125% didecyl dimethyl ammonium
chloride, 3.000% alkyl (50% C14, 40% C12, 10% C18) dimethyl benzyl ammonium
chloride, and 92.500% inert ingredients) which served as the positive control and sterile
distilled water was the negative control. The sanitizers in liquid form were applied with a
16-oz (473.2 ml) spray bottle (high-density polyethylene) with a sprayer nozzle (Decon
Labs, King of Prussia, PA). Sanitizer (ca. 11 ml) was applied to the slicer blade by 16 full
squeezes of the trigger of the spray bottle until the entire inoculated slicer blade was
covered. Sanitizer as a foam was applied with a portable foam unit (Pump Up Foamer FI10, Innovative Cleaning Equipment, Grand Rapids, MI) with pressure at 45 to 55 psi. The
sanitizer in foam was sprayed on inoculated blades for 5 s until the entire blade surface
was covered. Each blade was swabbed with sterile 6-inch (15.2 cm) polyester-tipped
swabs by rotating the blade clockwise for three complete rotations at 0, 1, 2, 3, 5, 10, 20,
and 30 min post-sanitizer application. The exposure time reported as “0 min” was
completed in less than 10 s after the sanitizer was in contact with the inoculated blade.
The swab was then dipped in 500 µl of 0.1 M PBS, pH 7.25, (for LVA plus SDS) or
neutralizing buffer (Becton Dickinson) (for the quat-positive control) to halt the
bactericidal activity. Each swab was agitated and surface plated as described above. In
addition, all of the swabs were transferred to enrichment media. The enrichment broths
for L. monocytogenes, S. Typhimurium, and E. coli O157:H7 were Fraser broth, Selenite
70
broth, and TSB plus nalidixic acid (50 µg/ml), respectively. Isolates from positive
enrichment cultures were confirmed by Listeria immunoassay (for L. monocytogenes) or
latex agglutination assay (for S. Typhimurium or E. coli O157:H7, all from Oxoid,
Basingstoke, UK).
2.6. Statistical analysis
Each experiment was repeated twice with duplicate plates. The mean population
of pathogens per ml was converted to log CFU/ml. Data were analyzed for analysis of
variance (ANOVA) by SAS software (SAS 9.3, SAS Institute, Cary, NC) to determine
least significant differences (P < 0.05) among the treatments.
3. Results and discussion
3.1. Transfer of the pathogens from inoculated deli foods to food contact locations on
slicers
Listeria monocytogenes, S. Typhimurium, and E. coli O157:H7 were recovered
from all of the contact locations of slicers that were tested after slicing the surfaceinoculated deli foods (Figure 3.1). The three pathogens were transferred at a rate of 1.0 to
4.0 log CFU/10 cm2 from the surface of contaminated deli foods (ca. 3.0 log CFU/cm2) to
slicer surfaces, with no significant (P > 0.05) difference among the mean or total transfer
rates. The populations of L. monocytogenes and E. coli O157:H7 recovered from the
blades were significantly (P < 0.05) less than the cell numbers recovered from the meat
grips and carriage trays, whereas the least (P < 0.05) cell numbers of S. Typhimurium
were recovered from the blade among the 5 contact locations (Figure 3.2). The meat grips
had teeth that contacted the food surface, thereby increasing the contact area. Meat juice
was released when the meat grip was pressed on the deli meats to secure them,
71
subsequently contributing to additional transfer of the pathogens to the grips. The
carriage tray had a large area that contacted the deli foods. The groove and ridge design
on the surface of the carriage tray increased the contact area with the deli foods.
Moreover, food juice and debris accumulated in these areas. Like the carriage tray, the
gauge plate and blade cover also had grooves and ridges, but neither of them directly
contacted the exterior surface of the deli foods. The blades largely came in contact with
the surface-inoculated deli foods at the blade edge, so the population of pathogens
brought along to the inner blade surface while slicing was limited. Sheen et al. (2010)
reported that many pathogen cells are killed or rendered nonviable on a slicer blade due
to the impact of the surface shear force and the instantly lethal high temperature produced
during the slicing operation. However, this mechanism of lethality has not been definitely
elucidated.
3.2. Transfer of the pathogens from inoculated slicer blades to food slices
This study was designed to simulate a worst-case scenario to determine
contamination transfer of the three pathogens on the slicers. When the slicer blades were
contaminated with L. monocytogenes, S. Typhimurium, or E. coli O157:H7 initially at ca.
8.5 log CFU/blade, the slicing operation contaminated up to 61 consecutive slices (Figure
3.3). The first slice fell off from the blade before making contact with the inoculated
upper-side of the blade, so it was not contaminated by the pathogens. The transfer rate
was 2.0 to 4.0 log CFU of pathogens/slice for the first 25 slices, and transfer
logarithmically (R2 > 0.9) decreased to a low level (< 2.0 log CFU/slice) and, did in some
trials, continued for up to 100 slices (data not shown). Enrichment cultures of pathogens
of slices were sporadically pathogen-positive for those slices that were close to the 100
72
slices of the deli foods that had been cut. There was no significant difference (P > 0.05)
among the three deli foods in the total counts of the three pathogens recovered from the
slices selected.
Our data also revealed that the three pathogens had the ability to survive on dried
surfaces of the slicers for at least 6 days (data not shown). Lin et al. (2006) suggested that
pathogens could adhere to some locations on slicers and subsequently contaminate meat
during slicing. Examining slicer blade surfaces by scanning electron microscopy (SEM)
revealed that even new slicer blades had numerous micro-ridges and grooves, and
repeated use and cleaning exacerbated the roughness (Vorst et al., 2006). These microridges and grooves could serve as niches for pathogen attachment and enhance crosscontamination. Improper cleaning and sanitization of meat and cheese slicers in retail
establishments could result in pathogens residing in these niches and forming a biofilm
which can be 1000 times more resistant to certain sanitizers than planktonic cells of the
pathogens (Simões et al., 2010). A biofilm also has the ability to release planktonic
pathogen cells for an extended period of time, thereby consistently contaminating delisliced foods (Srey et al., 2013). The Swiss cheese had a higher fat content (29%) than the
ham (4%) and roast beef (5%), and a layer of cheese was observed to form on the slicer
blade after the cheese had been sliced. Previously reported studies revealed that a fat
layer was formed on blades after slicing high fat-content salami and the fat layer was
thought to contribute to prolonged Listeria transfer and a higher prevalence of listeriae on
slicer surfaces (Lin et al., 2006; Vorst et al., 2006). However, there was no significant
difference (P > 0.05) in the mean transfer rate of the three pathogens from slicer blades to
cheese or meat slices in this study. Since cheese was inoculated with Listeria in our study,
73
instead of salami, the food composition and texture may have influenced the rate of
transfer of the pathogens.
3.3. Determination of the efficacy of levulinic acid plus SDS-based sanitizer on slicers
Applying 0.5% LVA plus 0.05% SDS or 1% LVA plus 0.1% SDS as a liquid on
slicer blades at 21°C achieved 3.6- and 5.8-log CFU/blade reduction of L. monocytogenes
populations, respectively (Table 3.1). Inactivation was greater when the concentrations of
levulinic acid and SDS were increased. The L. monocytogenes population was reduced to
an undetectable level (< 1.5 log CFU/blade) within 1 min when treated with quaternary
ammonium-based sanitizer (150 ppm; > 6-log CFU/blade reduction), 2% LVA plus 0.5%
SDS as a liquid, or all the three foam treatments (> 8-log CFU/blade reduction) with
different concentrations (0.5% LVA + 0.05% SDS; 1% LVA + 0.1% SDS; and 2% LVA
+ 0.5% SDS). Listeriae were undetected within 10 s after applying 2% LVA plus 0.5%
SDS as foam. The quat sanitizer (150 ppm) did not provide within 1 min a significant
killing effect (P > 0.05) on S. Typhimurium (Table 3.2). Salmonella cell numbers were
reduced by 3.5 and 5.2 log CFU/blade within 1 min when treated with 0.5% LVA plus
0.05% SDS in liquid and 1% LVA plus 0.1% SDS as a liquid, respectively. Greater than
a 6.0-log reduction was achieved when the Salmonella-inoculated slicer blades were
exposed for 1 min to 2% LVA plus 0.5% SDS as a liquid or all three foam treatments
with the different concentrations of LVA plus SDS. The quat sanitizer (150 ppm), 0.5%
LVA plus 0.05% SDS as a liquid, and 1% LVA plus 0.1% SDS as a liquid did not
significantly (P > 0.05) decrease within 1 min the E. coli O157:H7 population inoculated
on slicer blades (Table 3.3). Increasing the concentration of LVA to 2% and SDS to 0.5%
substantially increased the antimicrobial efficacy of the LVA plus SDS treatment when
74
applied as a liquid, with E. coli O157:H7 only detected by enrichment method (ca. 6.0log CFU/blade reduction) after 1 min of exposure. When applied as a foam, within 2 min
E. coli O157:H7 cell numbers were reduced to an undetectable level (E. coli O157:H7negative by enrichment culture) by the 0.5% LVA plus 0.05% SDS treatment, within 1
min with 1% LVA plus 0.1% SDS, and within 10 s with 2% LVA plus 0.5% SDS.
The deli slicer is one of the most difficult pieces of equipment to clean in the food
industry. Improper cleaning and sanitization can lead to cross-contamination of
pathogens to foods and biofilm formation by pathogens on slicers. Combinations of
levulinic acid and SDS at appropriate concentrations, and especially when applied as
foam, are highly effective in killing L. monocytogenes, S. Typhimurium, and E. coli
O157:H7 on slicers. Treating slicer blades with a liquid solution of 2% levulinic acid plus
0.5% SDS killed all three of the pathogens by 6.0 to 8.0 log CFU/blade within 1 min.
Foaming solutions (45-55 psi), applied either at equivalent or lower concentrations of
levulinic acid plus SDS, significantly (P < 0.05) reduced the three pathogens at all of the
contact times evaluated. The difference in the antimicrobial efficacy of the liquid versus
the foam treatments may have been related to their penetration rate, with the foam
treatment providing a greater rate of penetration. In addition, it was observed that the
foaming solutions were able to penetrate hidden areas, such as backside of the blade and
the space between the blade and blade guard. Since the deli slicers have numerous hidden
areas, which are hard to clean and on which contamination is not always visually
apparent, application of sanitizers as foaming solutions may be more practical to reduce
the risk of cross-contamination. E. coli O157:H7 was the least sensitive pathogen to the
levulinic acid plus SDS treatments, whereas L. monocytogenes was the most sensitive
75
(Tables 3.1-3.3). Sanitizer type, form of application (liquid or foam) and concentration,
and contact time are all important factors in killing the three pathogens on slicer blades.
In conclusion, this study determined the dynamics of cross-contamination of L.
monocytogenes, S. Typhimurium, and E. coli O157:H7 from contaminated deli foods to
slicers and from contaminated slicers to deli foods. After slicing surface-inoculated deli
food products, the pathogens were recovered from all of the 5 food contact locations on
slicers, with a significantly (P < 0.05) less transfer rate on blades than meat grips and
carriage trays. With an initial inoculation of ca. 8.5 log CFU/blade, the transfer of
pathogens decreased logarithmically from 4.0 to <1.5 log CFU/slice after 60 slices. Even
low concentrations of levulinic acid (1%) plus SDS (0.1%) as foam can reduce 6.0 to 8.0
log CFU of the three pathogens/blade within 1 min on slicer surfaces. Hence, this
combination of chemicals may have the potential to be an effective sanitizer for largescale applications in food processing facilities.
Acknowledgements
We thank Ping Zhao for technical assistance. This study was supported by grants
from the Center for Food Safety, University of Georgia, and the National Institute of
Food and Agriculture, U. S. Department of Agriculture.
76
1
3
2
4
5
Figure 3.1. Food contact locations that were swabbed on slicers to determine pathogen
contamination after inoculated deli foods were sliced. 1, meat grip; 2, carriage tray; 3,
gauge plate; 4, slicer blade; and 5, blade cover.
77
L. monocytogenes
S. Typhimurium
E. coli O157: H7
4.0
Log CFU/10 cm2
3.5
3.0
2.5
2.0
1.5
1.0
0.5
0.0
Grip
Carriage tray Gauge plate
Blade
Blade cover
Contact locations on slicer
Figure 3.2. L. monocytogenes, S. Typhimurium, and E. coli O157:H7 populations
recovered from different contact locations on slicers after slicing inoculated deli foods
(ca. 3.0 log CFU/cm2). Values are means ± standard deviations. Minimum detection limit
by the direct plating method was 1.5 log CFU/10 cm2.
78
Log CFU/slice
5
4.5
4
3.5
3
2.5
2
1.5
1
0.5
0
L. monocytogenes
S. Typhimurium
E. coli O157:H7
1
11
21
31
41
51
61
Slice number
Figure 3.3. Transfer of L. monocytogenes, S. Typhimurium, and E. coli O157:H7 from
inoculated slicer blades (ca. 8.5 log CFU/blade) to uninoculated Swiss cheese, ham, and
roast beef, respectively. Minimum detection limit by the direct plating method was 0.7
log CFU/slice.
79
Table 3.1. Effect of different concentrations of levulinic acid (LVA) plus SDS at different exposure times at 21°C on L.
monocytogenes inoculated on slicer blades.
Listeria monocytogenes count (log CFU/blade) ata:
0 minb
1 min
2 min
3 min
5 min
Liquid
H2O
8.4 ± 0.3 8.1 ± 0.3 7.9 ± 1.0 7.3 ± 1.1 7.3 ± 0.5
Commercial quat (150 ppm) 4.4 ± 0.5c
+c
-c
-c
-c
c
c
c
c
0.5% LVA + 0.05% SDS
5.4 ± 0.7 4.5 ± 0.9 3.6 ± 0.8 2.4 ± 1.4
-c
c
c
c
c
1% LVA + 0.1% SDS
3.2 ± 0.4 2.3 ± 0.6
+
-c
2% LVA + 0.5% SDS
+c
-c
-c
-c
-c
Foam
0.5% LVA + 0.05% SDS
1.5 ± 0.0c
-c
+c
-c
-c
c
c
c
c
1% LVA + 0.1% SDS
+
-c
c
c
c
c
2% LVA + 0.5% SDS
-c
a
Values are means ± standard deviations. Each sample was tested in duplicate. The minimum detection limit by the direct plating
Chemical Treatment
method was 1.5 log CFU/blade. “+” indicates that L. monocytogenes was not detected by the direct plating method but was enrichment
culture-positive. “-” indicates that both the direct plating method and enrichment cultures were negative.
b
The exposure time of “0” was completed in less than 10 s after the chemicals were sprayed onto the slicer blades.
c
Significantly different from the negative control (P < 0.05).
80
Table 3.2. Effect of different concentrations of levulinic acid (LVA) plus SDS at different exposure times at 21oC on S. Typhimurium
inoculated on slicer blades.
Salmonella Typhimurium count (log CFU/blade) ata:
0 minb
1 min
2 min
3 min
5 min
Liquid
H2O
6.6 ± 0.7 6.7 ± 0.7 5.6 ± 0.3 5.3 ± 0.9 5.2 ± 0.3
Commercial quat (150 ppm) 5.1 ± 0.2d 4.8 ± 0.1d 2.0 ± 0.7c 1.8 ± 0.5c
-c
d
c
c
c
0.5% LVA + 0.05% SDS
4.6 ± 0.4 3.2 ± 0.4 2.8 ± 0.8 1.7 ± 0.5
-c
1% LVA + 0.1% SDS
3.9 ± 0.4d 1.5 ± 2.1c
+c
-c
-c
c
c
c
c
2% LVA + 0.5% SDS
-c
Foam
0.5% LVA + 0.05% SDS
+c
-c
-c
-c
-c
c
c
c
c
1% LVA + 0.1% SDS
+
-c
2% LVA + 0.5% SDS
+c
-c
-c
-c
-c
a
Values are means ± standard deviations. Each sample was tested in duplicate. The minimum detection limit by the direct plating
Chemical Treatment
method was 1.5 log CFU/blade. “+” indicates that S. Typhimurium was not detected by the direct plating method but was enrichment
culture-positive. “-” indicates that both the direct plating method and enrichment cultures were negative.
b
The exposure time of “0” was completed in less than 10 s after the chemicals were sprayed onto the slicer blades.
c
Significantly different from the negative control (P < 0.05).
d
Not significantly different from the negative control (P > 0.05).
81
Table 3.3. Effect of different concentrations of levulinic acid (LVA) plus SDS at different exposure times at 21oC on E. coli O157:H7
inoculated on slicer blades.
Escherichia coli O157:H7 count (log CFU/blade) ata:
0 minb
1 min
2 min
3 min
5 min
10 min
20 min
Liquid
H2O
6.7 ± 0.5 6.4 ± 0.3 5.5 ± 1.0 6.2 ± 0.5 5.9 ± 0.2 4.3 ± 0.7 3.9 ± 0.3
Commercial quat (150 ppm) 6.1 ± 0.1d 4.7 ± 0.9d 3.8 ± 0.2d 2.1 ± 0.5c
-c
-c
-c
d
d
d
c
c
c
0.5% LVA + 0.05% SDS
6.2 ± 0.2 6.1 ± 0.4 4.9 ± 0.2 4.5 ± 0.2 3.1 ± 0.2
+
-c
c
d
c
c
c
c
1% LVA + 0.1% SDS
4.3 ± 0.7 3.8 ± 0.3 2.4 ± 1.4 1.9 ± 0.8
+
-c
2% LVA + 0.5% SDS
-c
+c
-c
-c
+c
-c
-c
Foam
0.5% LVA + 0.05% SDS
2.4 ± 0.9c
+c
-c
-c
-c
-c
-c
c
c
c
c
c
c
1% LVA + 0.1% SDS
3.1 ± 0.1
-c
c
c
c
c
c
c
2% LVA + 0.5% SDS
-c
a
Values are means ± standard deviations. Each sample was tested in duplicate. The minimum detection limit by the direct plating
Chemical Treatment
method was 1.5 log CFU/blade. “+” indicates that E. coli O157:H7 was not detected by the direct plating method but was enrichment
culture-positive. “-” indicates that both the direct plating method and enrichment cultures were negative.
b
The exposure time of “0” was completed in less than 10 s after the chemicals were sprayed onto the slicer blades.
c
Significantly different from the negative control (P < 0.05).
d
Not significantly different from the negative control (P > 0.05).
82
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88
CHAPTER 4
CONTROL OF PATHOGENS IN BIOFILMS ON THE SURFACE OF STAINLESS
STEEL BY LEVULINIC ACID PLUS SODIUM DODECYL SULFATE2
2
Chen, D., T. Zhao and M.P. Doyle. Accepted by International Journal of Food Microbiology.
Reprinted here with permission of the publisher.
89
ABSTRACT
The efficacy of levulinic acid (LVA) plus sodium dodecyl sulfate (SDS) to
remove or inactivate Listeria monocytogenes, Salmonella Typhimurium, and Shiga toxinproducing Escherichia coli (STEC) in biofilms on the surface of stainless steel coupons
was evaluated. Five- or six-strain mixtures (ca. 9.0 log CFU/ml) of the three pathogens
were separately inoculated on stainless steel coupons. After incubation at 21°C for 72 h,
the coupons were treated for 10 min by different concentrations of LVA plus SDS (0.5%
LVA + 0.05% SDS, 1% LVA + 0.1% SDS, and 3% LVA + 2% SDS) and other
commonly used sanitizers, including a commercial quaternary ammonium-based sanitizer
(150 ppm), lactic acid (3%), sodium hypochlorite (100 ppm), and hydrogen peroxide
(2%). The pathogens grew in the biofilms to ca. 8.6 to 9.3 log CFU/coupon after 72 h of
incubation. The combined activity of LVA with SDS was bactericidal in biofilms for
cells of the three pathogens evaluated, with the highest concentrations (3% LVA + 2%
SDS) providing the greatest log reduction. Microscopic images indicated that cells were
detached from the biofilm matrix and the integrity of cell envelopes were decreased after
the treatment of LVA plus SDS. This study is conducive to better understanding the
antimicrobial behavior of LVA plus SDS to the foodborne pathogens within biofilms.
Key words: biofilm, levulinic acid, sodium dodecyl sulfate, Listeria, Salmonella,
Escherichia coli
90
1. Introduction
Foodborne pathogens such as Listeria monocytogenes, Salmonella, and Shiga
toxin-producing Escherichia coli (STEC) are major food safety concerns. L.
monocytogenes causes listeriosis, a disease that mainly affects immunocompromised
individuals, the elderly and pregnant women (Kathariou, 2002). The symptoms of
listeriosis include encephalitis, meningitis, and abortion (Schlech, 2000). Salmonella and
STEC collectively cause in the United States an estimated 1.6 million foodborne illnesses
annually (Scallan et al., 2011). Salmonella causes fever, diarrhea and abdominal cramps 8
to 72 h after infection (Li et al., 2013), whereas STEC has been implicated in numerous
outbreaks, with symptoms including bloody diarrhea and hemolytic uremic syndrome
(HUS) (Durso et al., 2005).
In food processing facilities, some surfaces such as dead-end microscopic cracks
in gaskets, drip pan within refrigerators, and damp walls and ceilings due to condensation
are favorable sites for bacteria to grow in static biofilms (Chmielewski and Frank, 2004).
Biofilms are single or multi layers of microorganisms embedded in their own
extracellular polymeric substances (EPS) which associate with a solid surface (Donlan
and Costerton, 2002). It has been suggested that biofilms are the predominant matrix
resulting from bacterial growth, and approximately 80% of all bacterial infections are
biofilm-associated (de la Fuente-Nunez et al., 2012; Janssens et al., 2008). Biofilms
formed by foodborne pathogens can pose a substantial hygienic risk for the food industry
because biofilms with pathogens can serve as a contamination source and have an
enhanced resistance to mechanical actions and commonly used sanitizers (Carpentier and
Cerf, 1993). Corcoran et al. (2014) reported that commonly used disinfectants, including
91
sodium hypochlorite (500 ppm), sodium hydroxide (1 M), and benzalkonium chloride
(0.02%), failed to eradicate Salmonella biofilms on food contact surfaces. The sanitizer
applied on biofilms should not only possess antimicrobial activity, but also should be able
to penetrate the EPS barrier such that with sufficient concentration and exposure time it
will contact all of the cells in the biofilm. The efficacy of many sanitizers used in food
processing facilities is reduced when organic matter is present, whereby their usefulness
as an antimicrobial is mitigated (Simpson Beauchamp et al., 2012). Effective sanitizers
that are practical, efficacious, and safe to use are needed to control biofilms in food
processing. Levulinic acid (LVA) with sodium dodecyl sulfate (SDS) has been reported
previously to be an effective sanitizer for inactivating foodborne pathogens in the
presence of organic matter (Magnone et al., 2013; Zhao et al., 2011; Zhao et al., 2009), as
this treatment can reduce cell populations in biofilms by > 6 log within 1 min (Wang et
al., 2012; Zhao et al., 2011). To our knowledge, no studies have evaluated the
antimicrobial efficacy of a LVA with SDS combination on inactivating and removing the
foodborne pathogens L. monocytogenes and STEC growing as biofilms on stainless steel.
Hence, the goal of this study was to determine the effectiveness of LVA plus SDS for
inactivating L. monocytogenes, Salmonella, and STEC cells in biofilms formed on
stainless steel coupons.
2. Materials and methods
2.1. Bacterial strains
Five strains of L. monocytogenes, including LM101 (serotype 4b, salami isolate),
LM112 (serotype 4b, salami isolate), LM113 (serotype 4b, pepperoni isolate), H9666
(serotype 1/2c, human isolate), and ATCC 5779 (serotype 1/2c, cheese isolate); five
92
isolates of S. Typhimurium DT104, including H2662 (cattle isolate), 11942A (cattle
isolate), 13068A (cattle isolate), 152N17-1 (dairy isolate), and H3279 (human isolate);
and six strains of STEC, including O26:H11 (DEC10B, cattle isolate), O45:H2 (human
isolate), O103:H2 (human isolate), O111:NM (0944-95, cattle isolate), O121-Hunt
(human isolate), and O157:H7 (932, human isolate), were used. The cultures were
collected and incubated as described previously (Chen et al., 2014). Briefly, each strain
of the three pathogens was cultured in 10 ml of BHI (L. monocytogenes) or TSB
(Salmonella and STEC) individually, and incubated at 37°C for 20 h. The cultures were
then washed three times with 0.1 M phosphate buffered saline (PBS, pH 7.2, Sigma, St.
Louis, MO) by centrifugation at 3,000 × g for 10 min at 4°C and re-suspended in BHI or
TSB medium. For each pathogen, the optical density (OD) of each strain was adjusted in
a spectrophotometer (model 4001/4, Spectronic Instruments, Rochester, NY) with BHI or
TSB to an OD reading of 0.9 (ca. 9.0 log CFU/ml) at 630 nm. Approximately the same
cell number of each strain of the pathogen was combined to obtain three bacterial
mixtures. Cell numbers in the mixtures were determined by spread plating serial dilutions
(1:10 in 0.1 M PBS) onto TSA plates. The plates were incubated at 37°C for 24
(Salmonella and STEC) to 48 h (L. monocytogenes), and typical colonies were counted.
2.2. Preparation of stainless steel coupons
Stainless steel (type 304; Tull Metals Company, Atlanta, GA) coupons (4 cm ×
2.5 cm) were prepared according to the protocol described by Zhao et al. (2004), with
minor modifications. Prior to use, the coupons were washed by a 12-h immersion in
1,000 ml of an aqueous 2% RBS 35 detergent concentrate solution (20 ml of RBS 35
concentrate per liter of sterile distilled water at 21°C; Pierce, Rockford, IL), and rinsed
93
three times by a 10-min immersion in 1,000 ml of sterile distilled water at 21°C. The
washed stainless steel coupons were air dried, and an area 1.27 cm in diameter was
encircled by a permanent marker. The coupons were then wrapped individually with
aluminum foil and autoclaved at 121°C for 15 min.
2.3. Biofilm formation
Each sterile stainless steel coupon was individually transferred into a tissue
culture dish base (60 mm × 15 mm, Falcon, Franklin Lakes, NJ) which was then placed
in an extra-deep Petri dish (100 mm × 25 mm, Thermo Scientific, Rochester, NY)
containing 10 ml of sterile water. An inoculum of 0.1 ml of the mixtures (ca. 9.0 log
CFU/ml) of L. monocytogenes, S. Typhimurium, or STEC was deposited within the
marked area of the stainless steel coupon and incubated at 21°C. Every 24 h, the marked
area was aspirated to remove spent media, washed five times with 0.1 ml of 0.1 M PBS to
remove unattached cells, and replaced with fresh BHI (L. monocytogenes) or TSB
(Salmonella and STEC) medium. All the biofilms in this study were grown for 72 h
before sampling.
2.4. Efficacy of sanitizer treatments
Before the coupons were treated with sanitizers, the marked area on coupons was
aspirated and washed five times with 0.1 M PBS as described above, to remove
unattached cells. The sanitizers evaluated included a commercial quaternary ammoniumbased sanitizer (QAC, 150 ppm) containing a mixture of dimethylammonium chlorides
with various even-numbered alkyl chain lengths as active ingredients, lactic acid (LA, 3%;
Sigma), sodium hypochlorite (SHC, 100 ppm; Becton Dickinson, Sparks, MD), hydrogen
peroxide (HP, 2%; Becton Dickinson), levulinic acid (LVA, 3%; Sigma), sodium dodecyl
94
sulfate (SDS, 2%; Sigma), and three different concentrations of LVA plus SDS (0.5%
LVA + 0.05% SDS, 1% LVA + 0.1% SDS, and 3% LVA + 2% SDS). Sterile distilled
water was used as the control. All of the sanitizers were prepared according to the
manufacturers’ instructions, immediately before use. After rinsing with 0.1 M PBS as
described previously and air dried for 5 min, 0.1 ml of sanitizer was placed on the marked
area on each coupon. After exposure to the sanitizer for 10 min, the marked area was
aspirated to remove the sanitizers and unattached cells. The residual sanitizers were
neutralized with 0.1 ml of neutralizing buffer (Becton Dickinson) for 10 min. After
aspiration, the coupons were subject to bacterial enumeration or added with 0.1 ml of
BHI or TSB medium in the encircled area as a 24-h enrichment culture.
2.5.Bacterial enumeration
Each coupon bearing pathogenic bacteria in biofilms was washed with 0.1 ml of
0.1 M PBS as described previously and then individually placed in a 50-ml centrifuge
tube containing 9.9 ml of 0.1 M PBS and 30 glass beads (5-mm diameter; Fisher
Scientific, Norcross, GA). The tubes were agitated by a Vortex mixer (Fisher Scientific)
for 2 min to detach the cells from the stainless steel surface. One milliliter of the
suspension and 0.1 ml of serial dilutions (1:10 in PBS) were plated in duplicate on TSA
plates. The plates were incubated at 37°C for 24 (Salmonella and STEC) to 48 h (L.
monocytogenes) before bacterial counts.
2.6. Scanning electron microscopy (SEM)
Biofilm formation by the three pathogens on the surface of stainless steel coupons
after treatment with LVA plus SDS was visualized using scanning electron microscopy
(SEM). Biofilms were grown at 21°C for 72 h and then treated with sterile distilled water
95
(control) and different concentrations of LVA plus SDS for 10 min as described
previously. After adding with neutralizing buffer, the coupons were fixed with 2%
glutaraldehyde (Sigma) for 1 h at room temperature, rinsed three times for 15 min each
with PBS, air dried for 30 min, and sputter coated with gold (model 11428-AB, Spi
Supplies, West Chester, PA). The samples were subsequently examined with a Zeiss
1450EP scanning electron microscope (Zeiss, Scotts Valley, CA).
2.7. Transmission electron microscopy (TEM)
The effect of LVA plus SDS on the structures of the pathogens in biofilms was
investigated to determine this treatment’s influence on cell viability and cellular injury.
After treatment with LVA plus SDS, the coupons bearing pathogenic bacteria in biofilms
were aspirated to remove chemicals and neutralizing buffer was added on the marked
area as described previously. Each coupon was swabbed with a sterile 6-inch (15.2 cm)
polyester-tipped swab (Fisher Scientific), and the swab was then dipped in 900 µl of 0.1
M PBS, pH 7.2. After agitation by a Vortex mixer for 2 min, the suspensions were fixed
with 2% glutaraldehyde for 1 h at room temperature, rinsed three times with PBS for 15
min each time, then secondarily fixed with 1% OsO4 for 1 h at room temperature, and
dehydrated in an ethanol series of 25%, 50%, 75%, 100%, 100% and 100% for 15 min
each. The samples were then soaked in 50% (PO:ethanol) and 100% propylene oxide
(PO) for 5 min each, and infiltrated with EmBed 812 resin (EMS, Hatfield, PA) in a
series of 25%, 50% and 75% (resin:PO) for 1 h each. The samples were then allowed to
polymerize in 100% resin overnight at 60°C. Hardened blocks were trimmed and
sectioned on an ultramicrotome (RMC/Boekeler, Tuscon, AZ) to a thickness of
approximately 50 nm collected on a copper grid. Sections were post-stained with 2%
96
uranyl acetate for 30 min and lead citrate for 5 min. The samples were washed with
distilled water and air dried, then examined using a Technai 20 transmission electron
microscope (FEI, Eindhoven, Netherlands).
2.8. Statistical analysis
Each experiment was repeated three times, with duplicate samples each time. The
mean population of pathogens per ml or coupon was converted to log CFU/ml or log
CFU/coupon. Data were analyzed for one-way analysis of variance (ANOVA) by SAS
software (SAS 9.3, SAS Institute, Cary, NC) to determine least significant differences at
P < 0.05 among the treatments.
3. Results and discussion
After static incubation at 21°C for 72 h, L. monocytogenes, S. Typhimurium, and
STEC grew to ca. 8.6, 9.0, and 9.3 log CFU/coupon, respectively, in biofilms on the
surface of the stainless steel coupons (Table 4.1). For all of the sanitizers tested, complete
elimination/inactivation of the pathogens in biofilms did not occur, except for the
combination of the highest concentrations of LVA plus SDS (3% LVA + 2% SDS) on
STEC in biofilms. There is a synergistic antimicrobial effect between LVA and SDS
since the application of LVA or SDS individually at equivalent concentrations is
considerably less effective, with only a 0.2 to 1.2-log CFU/coupon reduction.
Dramatically enhanced antimicrobial efficacy when combining LVA with SDS was also
observed previously (Zhao et al., 2014; Zhao et al., 2009, 2010). Surfactants, like SDS,
can act as antiadhesive agents and LVA may assist in removal of the attachment polymer
by chelating divalent cations required to link the polymer at the surface (Frank, 2001).
Hence, the combination of an organic acid and surfactant may promote each other to
97
detach bacterial cells from a biofilm matrix. Once the cells are directly exposed to the
chemicals without the protection of EPS, SDS can chelate divalent cations, such as Ca2+
and Mg2+, leading to instability of outer membrane of Gram-negative bacteria (Hancock
and Rozek, 2002). LVA, in addition to its antimicrobial activity due to lowering pH, may
also damage cell membranes by releasing lipopolysaccharide from the outer membrane of
Gram-negative bacteria (Alakomi et al., 2000; Ricke, 2003). LVA and SDS are both
permeabilizers that complement each other enabling penetration of cells, leading to
increased susceptibility (Alakomi et al., 2000; Helander et al., 1997).
The other evaluated chemical sanitizers, including 150 ppm QAC, 2% lactic acid,
100 ppm sodium hypochlorite, and 2% hydrogen peroxide, were not effective against all
the three pathogens in the biofilms on stainless steel. QAC predominantly target cell’s
membrane, and Gram-positive bacteria are more sensitive to QACs at low concentrations
than Gram-negative (McDonnell and Russell, 1999). The undissociated form of lactic
acid can freely diffuse across the bacterial cell membrane and lower the cytoplasmic pH
upon dissociation (Virto et al., 2005). Lactic acid is generally effective to inactivating
enteric bacterial pathogens (Zhao et al., 2014). In hypochlorite solution, hypochlorous
acid readily reacts with organic matters in cells and is more bactericidal to Gram-negative
(Corcoran et al., 2014; Virto et al., 2005). Hydrogen peroxide acts as an oxidant by
producing hydroxyl free radicals attacking essential cell components (McDonnell and
Russell, 1999). All three of the pathogens used in this study can produce catalase, which
provides protection to the embedded cells by preventing full penetration of hydrogen
peroxide into the biofilm (Stewart et al., 2000). These commonly used sanitizers failed to
fully inactivate bacterial cells within biofilms, perhaps because the cells in the biofilm
98
matrix do not contact these sanitizers due to insufficient penetration and neutralization
with constituents of the biofilms (Corcoran et al., 2014; Stewart et al., 2001; Stewart et al.,
2000).
The SEM images (Figures 4.1-4.3) revealed that most of the cells in the biofilms
were detached from the biofilm matrix after a 10-min treatment with LVA plus SDS.
Once the bacterial cells detach from the biofilm matrix, they are more vulnerable to
antimicriobial agents (Kumar and Anand, 1998). The SEM images revealed the
occurrence of abundant EPS in the biofilms formed by the three pathogens, and that a
greater number of the pathogens in biofilms were inactivated when the treatment
concentrations of LVA and SDS were increased. The TEM analysis revealed structural
changes to bacterial cells treated with LVA plus SDS (Figures 4.4-4.6). The untreated
cells of L. monocytogenes (Figure 4.4A), S. Typhimurium (Figure 4.5A), and STEC
(Figure 4.6A) had smooth and well-defined cell walls or outer membranes. Treatment of
L. monocytogenes with 0.5% LVA + 0.05% SDS produced a loss of definition in the cell
wall and aggregation of cytoplasmic cellular component. We did not observe any cells in
the L. monocytogenes samples (n = 12) that were treated with higher concentrations of
LVA plus SDS, likely because of cellular lysis and loss of cells during the sample
preparation for TEM. For S. Typhimurium and STEC, aggregation of the cytoplasmic
components of cells followed by less defined cell morphology was observed as the
treatment concentrations of LVA plus SDS were increased. Cellular leakage of STEC
was observed (Figure 4.6B), indicating the cell membrane was damaged in the presence
of low concentrations of LVA plus SDS. Bacterial cell counts by the plating method
performed in this study revealed that L. monocytogenes was more susceptible to LVA
99
plus SDS compared to the other two Gram-negative pathogens. This finding is consistent
with our previous study (Chen et al., 2014). Unlike Gram-negative bacteria, Grampositives such as L. monocytogenes do not possess an outer membrane. Hence, they are
more susceptible to the action of chemicals interfering with the transport of ions across
the cell membrane (Feliciano et al., 2012; Skrivanova et al., 2006).
In conclusion, the combination of LVA and SDS was most effective against the
three pathogens in biofilms compared to the other evaluated sanitizers, with the highest
concentrations (3% LVA + 2% SDS) providing the greatest log reduction. SEM images
confirmed that the combination of LVA and SDS was effective in inactivating bacterial
cells in biofilms, and the TEM images revealed that LVA with SDS cause changes to the
permeability of the cell membrane and aggregation of the cytoplasmic components.
Results of this study may be useful to mitigate the presence of biofilms in food
processing facilities.
Acknowledgments
We thank Ping Zhao and Dr. John Shields for technical assistance. This study was
supported by grants from the Center for Food Safety, University of Georgia, and the U. S.
Department of Agriculture, National Institute of Food and Agriculture, Food Research
Initiative Grant No. 2011-68003-30012.
100
A
B
C
D
Figure 4.1. Representative photomicrographs by SEM of biofilms formed by L.
monocytogenes after a 10-min treatment with water (control, A), 0.5% levulinic acid +
0.05% SDS (B), 1% levulinic acid + 0.1% SDS (C), or 3% levulinic acid + 2% SDS (D).
Scale bar = 2 μm.
101
A
B
C
D
Figure 4.2. Representative photomicrographs by SEM of biofilms formed by S.
Typhimurium after a 10-min treatment with water (control, A), 0.5% levulinic acid +
0.05% SDS (B), 1% levulinic acid + 0.1% SDS (C), or 3% levulinic acid + 2% SDS (D).
Scale bar = 2 μm.
102
A
B
C
D
Figure 4.3. Representative photomicrographs by SEM of biofilms formed by STEC after
a 10-min treatment with water (control, A), 0.5% levulinic acid + 0.05% SDS (B), 1%
levulinic acid + 0.1% SDS (C), or 3% levulinic acid + 2% SDS (D). Scale bar = 2 μm.
103
A
B
Figure 4.4. Representative photomicrographs by TEM of biofilms formed by L.
monocytogenes after a 10-min treatment with water (control, A), or 0.5% levulinic acid +
0.05% SDS (B). The black arrow highlights the morphological differences after the 10min treatment of levulinic acid plus SDS. Scale bar = 100 nm.
104
A
B
C
D
Figure 4.5. Representative photomicrographs by TEM of biofilms formed by S.
Typhimurium after a 10-min treatment with water (control, A), 0.5% levulinic acid +
0.05% SDS (B), 1% levulinic acid + 0.1% SDS (C), or 3% levulinic acid + 2% SDS (D).
Scale bar = 100 nm.
105
A
B
C
D
Figure 4.6. Representative photomicrographs by TEM of biofilms formed by STEC after
a 10-min treatment with water (control, A), 0.5% levulinic acid + 0.05% SDS (B), 1%
levulinic acid + 0.1% SDS (C), or 3% levulinic acid + 2% SDS (D). The black arrow
highlights the morphological differences after the 10-min treatment of levulinic acid plus
SDS. Scale bar = 100 nm.
106
Table 4.1. Inactivation of 72-h biofilms of L. monocytogenes, S. Typhimurium, and
STEC formed on stainless steel after exposure to a sanitizer for 10 min.
Counts (log CFU/coupon) b
L. monocytogenes
S. Typhimurium
STEC
DW (control)
8.6 ± 0.2ac
9.0 ± 0.0a
9.3 ± 0.1a
QAC (150 ppm)
4.2 ± 0.9d
8.2 ± 0.3a
7.6 ± 0.5c
LA (3%)
4.4 ± 0.2d
3.1 ± 0.0de
5.7 ± 0.1e
SHC (100 ppm)
7.9 ± 0.3bc
4.4 ± 0.9c
7.8 ± 0.0c
HP (2%)
8.3 ± 0.0ab
8.3 ± 0.2a
7.8 ± 0.6c
LVA (3%)
8.3 ± 0.2ab
8.6 ± 0.2a
8.8 ± 0.1b
SDS (2%)
7.4 ± 0.5c
8.8 ± 0.0a
9.0 ± 0.0ab
LVA (0.5%) + SDS (0.05%)
1.9 ± 0.3e
5.8 ± 0.5b
7.0 ± 0.3d
LVA (1%) +SDS (0.1%)
+e
3.1 ± 0.8d
6.1 ± 0.1e
LVA (3%) + SDS (2%)
+e
2.1 ± 0.5e
-f
a
DW, distilled water; QAC, quaternary ammonium compound; LA, lactic acid; SHC,
Treatmenta
sodium hypochlorite; HP, hydrogen peroxide; LVA, levulinic acid; SDS, sodium dodecyl
sulfate.
b
Values are means ± standard deviations. The minimum detection limit by the direct
plating method was 1.7 log CFU/coupon. “+” indicates that pathogen was not detected by
the direct plating method but was enrichment culture-positive. “-” indicates that both the
direct plating method and enrichment cultures were negative.
c
Values in the same column that are not followed by the same lower case letters are
significantly different (P < 0.05).
107
Appendix
Figure 4.7. The temperature of the stainless steel coupons during the 10-min heat
treatment at 60, 80 or 100ºC in an oven. The temperature was determined using a twochannel thermocouple (model HH23, Omega Engineering, Stamford, CT).
108
Table 4.2. Inactivation of 72-h biofilms of L. monocytogenes, S. Typhimurium, and
STEC formed on stainless steel after exposure to a heat treatment at 60, 80 or 100ºC for
10 min. Before the coupons were treated by different temperatures, the marked area on
coupons was aspirated and washed five times with 0.1 M PBS, to remove unattached
cells. The coupons were then placed into an oven (model J BS07 M1BB, General Electric,
Louisville, KY) which was preset at 60, 80, or 100°C and temperature was confirmed by
an interior thermometer. After exposure to the preset temperatures for 10 min, the
coupons were cooled at 21°C for 30 min before bacterial enumeration.
Counts (log CFU/coupon) a
L. monocytogenes
S. Typhimurium
STEC
b
21 ± 1°C (control)
8.6 ± 0.2a
9.0 ± 0.0a
9.3 ± 0.1a
60 ± 5°C
7.7 ± 0.2b
8.3 ± 0.1ab
8.5 ± 0.0b
80 ± 5°C
7.6 ± 0.5b
7.5 ± 0.3bc
7.5 ± 0.3c
100 ± 5°C
6.9 ± 0.3b
7.4 ± 0.1b
6.6 ± 0.4d
a
Values are means ± standard deviations. The minimum detection limit by the direct
Treatment
plating method was 1.7 log CFU/coupon.
b
Values in the same column that are not followed by the same lower case letters are
significantly different (P < 0.05).
109
Table 4.3. Bacterial counts on selective agar plates and TSA of 72-h biofilms of L. monocytogenes, S. Typhimurium, and STEC
formed on stainless steel after a 10-min treatment with 3% lactic acid (pH 2.2) or 3% levulinic acid (pH 2.7). The percentage of
injured cells after the treatment of lactic acid (3%, pH 2.2) and levulinic acid (3%, pH 2.7) was determined by:
% injured cells = 100 × (TSA counts – selective agar counts) / TSA counts
Count (log CFU/coupon)a
% injured cells
Selective agar platesb
TSA
Water (control)
8.6 ± 0.3
8.6 ± 0.2
-c
L. monocytogenes
3% Lactic acid
2.1 ± 0.2
4.4 ± 0.2
52.3%
3% Levulinic acid
8.3 ± 0.1
8.3 ± 0.2
Water
8.9 ± 0.0
9.0 ± 0.0
S. Typhimurium
3% Lactic acid
< 1.7
3.1 ± 0.0
> 45.2%
3% Levulinic acid
4.4 ± 0.3
8.6 ± 0.2
48.8%
Water
9.2 ± 0.1
9.3 ± 0.1
STEC
3% Lactic acid
< 1.7
5.7 ± 0.1
> 70.2%
3% Levulinic acid
6.4 ± 0.4
8.8 ± 0.1
27.3%
a
Values are means ± standard deviations. The minimum detection limit by the direct plating method was 1.7 log CFU/coupon.
Pathogen
b
c
Treatment
Selective plates were MOX, XLD, and MAC for L. monocytogenes, S. Typhimurium, and STEC, respectively.
A difference of at least 25% between counts on the TSA and selective agar plates was considered evidence of acid injury.
110
Table 4.4. Inactivation of 72-h biofilms of L. monocytogenes, S. Typhimurium, and
STEC formed on stainless steel after exposing to 80°C for 10 min and a subsequent 10min sanitizer treatment. Coupons were washed, placed in an oven (80°C) for 10 min, and
then cooled at 21°C for 30 min. The encircled area was then washed with 0.1% peptone
water for five times before a 10-min sanitizer application. The marked area was washed
again before bacterial enumeration.
Counts (log CFU/coupon) b
L. monocytogenes
S. Typhimurium
STEC
c
DW (control)
7.6 ± 0.5a
7.5 ± 0.3a
7.5 ± 0.3a
QAC (150 ppm)
-c
6.6 ± 0.3a
6.2 ± 0.6a
LA (3%)
-c
-c
-b
SHC (100 ppm)
3.7 ± 1.2bc
2.0 ± 0.7c
7.2 ± 0.4a
HP (2%)
5.1 ± 1.3b
4.9 ± 0.6b
7.1 ± 0.1a
a
DW, distilled water; QAC, quaternary ammonium compound; LA, lactic acid; SHC,
Treatmenta
sodium hypochlorite; HP, hydrogen peroxide.
b
Values are means ± standard deviations. Each sample was tested in triplicate (n = 3).
The minimum detection limit by the direct plating method was 1.7 log CFU/coupon. “-”
indicates that both the direct plating method and enrichment cultures were negative.
c
Values in the same column that are not followed by the same lower case letters are
significantly different (P < 0.05).
111
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CHAPTER 5
SINGLE- AND MIXED-SPECIES BIOFILM FORMATION BY ESCHERICHIA COLI
O157:H7 AND SALMONELLA, AND THEIR SENSITIVITY TO LEVULINIC ACID
PLUS SODIUM DODECYL SULFATE3
3
Chen, D., T. Zhao and M.P. Doyle. Accepted by Food Control.
Reprinted here with permission of the publisher.
117
ABSTRACT
The development of single- and mixed-species biofilms formed by Escherichia
coli O157:H7 and Salmonella was observed, and the antimicrobial effectiveness of
levulinic acid (LVA) plus sodium dodecyl sulfate (SDS) on the cells in single- and dualspecies biofilms was determined. Biofilm-forming ability of single- and mixed-species
cultures was observed by crystal violet staining and their resistance to levulinic acid plus
SDS was determined by enumeration. Fluorescent protein-labeled E. coli O157:H7 and
Salmonella were constructed and the bacterial composition of the biofilms after treatment
with levulinic acid plus SDS was visualized by confocal laser scanning microscopy
(CLSM). E. coli O157:H7 and Salmonella were antagonistic to each other, being more
sensitive to levulinic acid plus SDS in mixed-species biofilms. Images captured by
CLSM revealed that E. coli O157:H7 and Salmonella were distributed evenly in the
single- and dual-species biofilms, and confirmed that the combination of levulinic acid
and SDS was effective in inactivating bacterial cells in biofilms. Results revealed that
levulinic acid with SDS may be used as a potential biofilm control intervention.
Key words: mixed-species biofilm, Escherichia coli O157:H7, Salmonella, levulinic acid,
sodium dodecyl sulfate
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1. Introduction
Among Shiga toxin-producing Escherichia coli serotypes, O157:H7 is the one
most commonly associated with human disease in North America (Foley, et al., 2004). E.
coli O157:H7 has been implicated as the causative agent of many foodborne outbreaks
since it was first identified as a human pathogen in the 1980s (Ryu & Beuchat, 2005).
The pathogen can cause bloody diarrhea and target organ systems, such as the kidney,
resulting in hemolytic uremic syndrome (HUS) (Mead & Griffin, 1998). Salmonella
causes an estimated one million illnesses annually, and is one of the leading causes of
bacterial foodborne illness in the United States (Scallan, et al., 2011).
Both E. coli O157:H7 and Salmonella have animal reservoirs and have been
associated with contamination of meat and meat products (CDC, 2006, 2013). Studies
have revealed E. coli O157:H7 and Salmonella contamination of beef hides at slaughter
(Arthur, et al., 2008; Barkocy-Gallagher, et al., 2003; Rivera-Betancourt, et al., 2004),
and both pathogens can be isolated from feces (Bosilevac, et al., 2015). In recent years,
fresh produce has been increasingly implicated in foodborne outbreaks caused by E. coli
O157:H7 and Salmonella (Deering, Mauer, & Pruitt, 2012; Mohle-Boetani, et al., 2001).
E. coli O157:H7 and Salmonella may coexist and occur simultaneously at different meat
and produce processing sites, which serve as potential sources of cross contamination
(Wang, Kalchayanand, Schmidt, & Harhay, 2013). Biofilms formed by or harboring these
foodborne pathogens may be conducive to cross contamination since biofilms commonly
occur in fresh produce and meat processing environments (Giaouris, et al., 2014; Jahid &
Ha, 2012). In addition, biofilms confer to pathogens enhanced resistance to cleaning and
119
sanitization operations (van der Veen & Abee, 2011), thereby posing an increased food
safety concern.
Under appropriate conditions, E. coli O157:H7 and Salmonella can form singleand mixed-species biofilms on food and food contact surfaces (Wang, et al., 2013).
Inactivation of pathogens in biofilms with commonly used sanitizers has been extensively
studied, but complete elimination is seldom achieved (Corcoran, et al., 2014). Levulinic
acid (LVA) with sodium dodecyl sulfate (SDS) has been reported previously to be an
effective sanitizer for inactivating foodborne pathogens (Chen, Zhao, & Doyle, 2014;
Magnone, Marek, Sulakvelidze, & Senecal, 2013; Zhao, Zhao, Cannon, & Doyle, 2011;
Zhao, Zhao, & Doyle, 2009, 2010), as this combination can reduce Salmonella cell
counts in biofilms by > 6 log within 1 min (Wang, Hong, Ciancio, Zhao, & Doyle, 2012;
Zhao, et al., 2011). In addition, both levulinic acid (FDA 2008, 21 CFR, 172.515) and
SDS (FDA, 2007, 21 CFR, 172.822) have been individually designated by FDA as
Generally Recognized as Safe (GRAS) for specific uses in foods (Chen, et al., 2014). Our
recent study revealed that 3% levulinic acid plus 2% SDS was effective in reducing > 6.9
log CFU/coupon of Listeria monocytogenes, Shiga toxin-producing E. coli (STEC) and S.
Typhimurium in single-species biofilms formed on stainless steel coupons.
Although biofilms formed by mono-species have been studied extensively, the
sensitivity of E. coli O157:H7 and Salmonella formed in a mixed-species biofilm to the
combination of LVA and SDS is unknown. The purpose of this study was to determine
whether synergistic or antagonistic interactions occur between E. coli O157:H7 and
Salmonella in mixed-species biofilms, and their sensitivity to levulinic acid plus SDS.
120
2. Materials and methods
2.1. Bacterial strains
Escherichia coli O157:H7 strain USDA 5 and Salmonella strain 457-88 were used
in this study because the two strains were determined as strong biofilm producers in
previous experiment (Table 5.1). The cultures were stored at -20°C in tryptic soy broth
(TSB, Becton Dickinson, Sparks, MD) containing 25% glycerol (Sigma, St. Louis, MO)
and streaked on tryptic soy agar (TSA, Becton Dickinson) plates from which a single
colony was inoculated into 10 ml of TSB and incubated at 37°C. Three consecutive 20-h
transfers were made before use.
2.2. Preparation of single- and mixed-species cultures
Individual bacterial strains were grown in TSB at 37°C for 20 h. The singlespecies cultures were prepared by 1:100 dilution of the overnight culture (ca. 8.0 log
CFU/ml) in sterile TSB. An equal volume of the two1:100 diluted single-species cultures
at the same bacterial population was combined to make the mixed-species culture. The
cell numbers were confirmed by the direct plating method on TSA agar plates.
2.3. Single- and mixed-species biofilm formation on 96-well polystyrene plates
Single- and mixed-species cultures 100 μl per well were added to 96-well flatbottom polystyrene plates (Costar, Corning, NY). Plates were incubated statically for 72
h at 21°C. After incubation, the supernatant fluid in each well was aspirated, washed
three times with 200 μl of 0.1 M phosphate buffer saline (PBS, Sigma) per well, and
stained with 100 μl of 0.1% crystal violet (CV) per well for 20 min at 21°C. The CV
solution was removed by aspiration and the plates were then washed again with PBS. The
remaining CV was dissolved in 100 μl of 85% ethanol per well. After 10 min, the amount
121
of CV in each well was determined by measuring the optical density at 655 nm (OD655nm)
using a microplate reader (model 680XR, BioRad, Berkeley, CA). Wells containing only
100 μl of sterile TSB served as the control and the OD655nm was used as the background
value, which was subtracted from OD655nm values obtained from other wells with bacteria
(Burmølle, et al., 2006).
2.4. Sensitivity of E. coli O157:H7 and Salmonella in single- and mixed-species biofilms
to levulinic acid plus SDS
Single- and mixed-species cultures were added in 24-well flat-bottom polystyrene
plates (Costar) at 1 ml per well. Control wells contained only 1 ml of sterile TSB. The
plates were incubated statically at 21°C. After 72 h of incubation, the supernatant fluid
was removed, and the wells were rinsed three times with 2 ml of PBS. The plates were
then air dried for 5 min at 21°C. Two concentrations of levulinic acid and SDS (1% LVA
+ 0.1% SDS and 3% LVA + 2% SDS) were used in this study. Sterile water was the
negative control and a commercial quaternary ammonium compound-based sanitizer
(QAC, 150 ppm, Chemstar, Lithia Springs, GA) was the positive control. The QAC
contained a mixture of dimethylammonium chlorides with various even-numbered alkyl
chain lengths as active ingredients. The water or chemicals were removed by aspiration
after the 5-min treatment. The wells were then filled with 2 ml of N/E neutralizing broth
(Becton Dickinson) per well, held for 10 min before aspiration, and 1 ml of PBS was then
added to each well. The content of each well was harvested by scraping the surface
thoroughly with a sterile 6-inch (15.2 cm) polyester-tipped swab (Fisher Scientific,
Pittsburgh, PA). The PBS containing released biofilm cells and the swab were added to 9
ml of 0.1 M PBS, pH 7.25, and vigorously agitated by a Vortex (G-560, Scientific
122
Industries, Bohemia, NY) at maximum speed for 1 min. One milliliter of the suspension
or appropriate dilutions (0.1 ml) in 0.1 M PBS were spread plated in duplicate on
MacConkey agar (MAC, Becton Dickinson) plates, with E. coli O157:H7 and Salmonella
forming red and white colonies, respectively. The plates were incubated at 37°C for 48 h,
and colonies typical of the selected pathogens were counted. The limit of detection by the
direct plating method was 0.7 log CFU/ml.
2.5. Confocal laser scanning microscopy (CLSM)
Fluorescent protein (FP)-labeled pathogens were constructed to better observe the
phenotypic behavior of E. coli O157:H7 and Salmonella in the biofilms under CLSM and
minimize any disturbance caused by staining procedures. Plasmid pDsRed-Express2
(Clontech Laboratories, Mountain View, CA), containing genes to produce red
fluorescent protein (RFP), and plasmid pGFPuv (Clontech Laboratories), containing
genes to encode green fluorescent protein (GFP), were transformed into USDA 5 and
457-88, respectively. Both of the two plasmids had an ampicillin marker. The resulting E.
coli O157:H7 strain USDA 5 produced red fluorescence and the resulting Salmonella
strain 457-88 produced green fluorescence under UV light (Webb, Davey, Erickson, &
Doyle, 2013). The two plasmids remained stable in the respective strains after five
consecutive days of sequential propagation in TSB medium without ampicillin.
Sterile glass slides were used as the biofilm substratum. An area of 1.27 cm in
diameter was encircled at the center of glass slides by a hydrophobic liquid blocker pen
(Sigma). An inoculum of 0.1 ml of the diluted (1:100) single- or mixed-species culture
(containing 100 µg/ml of ampicillin) of FP-labeled USDA 5 and 457-88 was deposited in
the encircled area and the slide was then covered by a glass coverslip. The coverslip-
123
covered slides were then placed in a 90% relative humidity growth incubator (Fisher
Scientific) so that the medium did not dry out during incubation. After incubation at 21°C
for 72 h, the marked area on the glass slides was aspirated, washed with 100 µl of PBS
for three times, treated with 100 µl of water (control) or the different concentrations of
levulinic acid and SDS (1% LVA + 0.1% SDS and 3% LVA + 2% SDS) for 5 min,
neutralized with N/E broth for 10 min, and washed again with PBS. The encircled area on
the glass slides was aspirated and then examined with CLSM. The samples were
visualized with a Zeiss LSM 700 CLSM (Carl Zeiss Microscopy, Thornwood, NY) using
a Plan-Apochromat 63× oil-immersion, numerical aperture 1.4 objective (Carl Zeiss), a
488-nm argon laser and a 555-nm diode-pumped solid-state laser, with band-pass 490 to
550 nm for detection of Salmonella (green) and 560 to 700 nm for detection of E. coli
O157:H7 (red). Z-stack scanning was performed to visualize the spatial distribution of
the two strains in the biofilm structure. Serial images were captured and processed by
Zeiss Zen 2012 software (Carl Zeiss) using maximal projection.
2.6. Statistical analysis
The experiment of single- and mixed-species biofilm formation on 96-well
polystyrene plates was repeated twice with 12 wells each time (n = 24), and the
experiment of sensitivity of E. coli O157:H7 and Salmonella in biofilms to levulinic acid
with SDS was repeated twice with triplicate wells each time (n = 6). Data were analyzed
for analysis of variance (ANOVA) by SAS software (SAS 9.4, SAS Institute, Cary, NC)
to determine least significant differences (P < 0.05) among the treatments. One-way
ANOVA was used to analyze the biomass production in single- and mixed-species
biofilms and their sensitivity to levulinic acid plus SDS.
124
3. Results
3.1. Single- and mixed-species biofilm formation as determined by crystal violet staining
Escherichia coli O157:H7 strain USDA 5 and Salmonella strain 457-88 were
examined for synergistic or antagonistic effects when coincubated with each other on 96well polystyrene plates at 21°C for 72 h. Significantly (P < 0.05) more biomass was
produced by Salmonella strain 457-88 (OD655nm = 0.28 ± 0.06) than E. coli O157:H7
strain USDA 5 (OD655nm = 0.03 ± 0.01). The OD655nm value of this mixed-species biofilm
was 0.20 ± 0.04, which was significantly (P < 0.05) less than that of the biofilms formed
by strain 457-88 only (Figure 5.1). Thus, E. coli O157:H7 strain USDA 5 and Salmonella
strain 457-88 exhibited an antagonistic effect with each other in dual-species biofilms.
3.2. Bacterial counts of single- and mixed-species biofilm formation and their sensitivity
to levulinic acid plus SDS
Bacterial enumeration results of single- and mixed-species biofilms formed by E.
coli O157:H7 strain USDA 5 and Salmonella strain 457-88, and their resistance to
levulinic acid plus SDS are presented in Figure 5.2. In single-species biofilms, bacterial
counts of USDA 5 and 457-88 in control groups were not significantly (P > 0.05)
different (9.2 ± 0.2 and 9.4 ± 0.1 log CFU/ml, respectively). QAC (150 ppm), 1%
levulinic acid + 0.1% SDS, and 3% levulinic acid + 2% SDS treatments significantly (P <
0.05) reduced USDA 5 and 457-88 cells in single-species biofilms by 0.6, 1.1 and 4.1 log
CFU/ml, and 0.8, 1.6 and 7.6 log CFU/ml, respectively. In mixed-species biofilms, the
initial population of USDA 5 (8.3 ± 0.5 log CFU/ml) was approximately 10-fold less than
the count of 457-88 cells (9.1 ± 0.4 log CFU/ml), and also approximately 10-fold less
than the count of USDA 5 in single-species biofilms. Thus, culture conditions for both
125
single- and mixed-species significantly affected biofilm formation by USDA 5, but not
by 457-88. QAC (150 ppm), 1% levulinic acid + 0.1% SDS, and 3% levulinic acid + 2%
SDS treatments significantly (P < 0.05) reduced USDA 5 and 457-88 cells in mixedspecies biofilms by 0.4, 2.6 and 7.1 log CFU/ml, and 0.6, 3.4 and 8.4 log CFU/ml,
respectively. Two concentrations of levulinic acid and SDS significantly reduced both
pathogenic cells in single- and mixed-species biofilms, with a greater number of the
pathogens in biofilms inactivated when the treatment concentrations of levulinic acid and
SDS were increased. Cells of the two pathogens in mixed-species biofilms were more
sensitive to the combination of levulinic acid and SDS compared to the respective cells in
single-species biofilms. Although the initial population of USDA 5 in mixed-species
biofilms was approximately 10 times less than the initial count of 457-88 cells, after a 5min treatment of either of the two concentrations of levulinic acid and SDS, there was no
significant (P > 0.05) difference between the bacterial counts of these two pathogens in
the mixed-species biofilm.
3.3. Microscopic examination of single- and mixed-species biofilms
Single- and mixed-species biofilms formed by FP-labeled USDA 5 and 457-88 on
glass slides after a 5-min treatment of sterile water (control), 1% levulinic acid + 0.1%
SDS, or 3% levulinic acid + 2% SDS were examined using CLSM (Figure 5.3). In both
single- and dual-species biofilms, the cells were distributed evenly in the 72-h biofilm
matrix which consisted of multiple layers of cells, and the thickness of the biofilm matrix
reached 6 to 8 µm (Figure 5.4). The CLSM images revealed that more of the pathogens in
both single- and mixed-species biofilms were inactivated when the treatment
concentrations of levulinic acid and SDS were increased, which was in concordance with
126
the bacterial counts of viable cells in the biofilms after receiving the levulinic acid plus
SDS treatment.
4. Discussion
Biofilm formation is one of the strategies bacteria use to withstand environmental
stresses. In nature, mixed-species biofilms predominate in most environments (O'Toole,
Kaplan, & Kolter, 2000). In food processing environments, biofilms consisting of
multispecies of bacteria, including foodborne pathogens, can occur (Kostaki,
Chorianopoulos, Braxou, Nychas, & Giaouris, 2012). A better understanding of the
species interaction and how to inactivate the cells of multispecies within biofilms formed
on abiotic surfaces in food processing facilities could be helpful to prevent their
formation, and thereby reduce the potential for product contamination. In the singlespecies biofilms, significantly (P < 0.05) more biomass production by Salmonella strain
457-88 than E. coli O157:H7 stain USDA 5 was observed by CV staining, whereas
bacterial counts of strain USDA 5 and 457-88 were not significantly (P > 0.05) different.
Methods based on CV staining and CFU enumeration have been used to assess biofilmforming ability, and no or low statistically significant correlation between the two
methods was determined (Lourenco, Rego, Brito, & Frank, 2012). In this study, static
biofilms were used because there are many favorable sites, such as dead-end microscopic
cracks in gaskets, drip pans within refrigerators, and condensation damp walls and
ceilings, for bacteria to grow statically in biofilms in the food processing environment
(Chmielewski & Frank, 2004).
Bacterial counts revealed that significantly more Salmonella strain 457-88 than E.
coli O157:H7 strain USDA 5 grew in the mixed-species biofilms. Esteves, Jones, and
127
Clegg (2005) determined that Salmonella can outgrow E. coli on the epithelial surface in
mixed biofilms. Interspecies competition is common in mixed-species biofilms (Norwood
& Gilmour, 2001). In this study, E. coli O157:H7 strain USDA 5 and Salmonella strain
457-88 were capable of forming single-species biofilms with no significant (P > 0.05)
difference of bacterial counts. Hence, the competition between these two strains was not
simply because of an inability to form biofilms or a slower growth rate for USDA 5 than
457-88. Infrequent curli/cellulose expression by E. coli O157:H7 may result in being
outcompeted by Salmonella in mixed-species biofilms (Liu, Nou, Lefcourt, Shelton, &
Lo, 2014; Uhlich, Keen, & Elder, 2001). CLSM examination of both 8-h postinoculation
single- and mixed-species biofilms revealed that considerably less E. coli O157:H7 cells
were attached to the glass slides than Salmonella (Figure 5.5). This observation is
consistent with that of other researchers (Dewanti & Wong, 1995; Liu, et al., 2015). At
the initial stage of biofilm formation, Salmonella can attach to locations where nutrients
and certain environmental conditions are preferred. In contrast, Wang et al. (2013)
observed enhanced resistance of both E. coli O157:H7 and Salmonella in mixed-species
biofilms to sanitizers. These differences may be due to several factors. First, biofilm
formation by E. coli O157:H7 and Salmonella is strain specific (Bang, et al., 2014; Patel,
Singh, Macarisin, Sharma, & Shelton, 2013; Solomon, Niemira, Sapers, & Annous,
2005). Mixed-species biofilms consisting of different strains can exhibit different
phenotypic behaviors. Second, levulinic acid plus SDS used in this study has a different
mode of bacterial inactivation than either chlorine or QAC solutions. Hence, the
sensitivity of cells in biofilms can differ between studies. Third, biofilm formation and its
resistance to sanitizers could be influenced by other factors, such as surface
128
characteristics of substratum, environmental conditions, harvest methods of biofilm cells,
and other influences.
To better understand the species interaction within the biofilms and their spatial
and temporal development, E. coli O157:H7 strain USDA 5 and Salmonella strain 457-88
were transformed with plasmids expressing red and green fluorescent protein,
respectively. FP-labeled bacteria have been used previously in tracking a cell population
in a biofilm matrix (Nancharaiah, Venugopalan, Wuertz, Wilderer, & Hausner, 2005;
Tomlin, Clark, & Ceri, 2004). In this study, E. coli O157:H7 and Salmonella can be
effectively labeled with the FP plasmids and be stable in the respective two chosen strains
for at least 50 generations (Ma, Zhang, & Doyle, 2011). There was no significant (P >
0.05) difference in planktonic cell growth rates between the parental and respective FPlabeled strains (Figure 5.6). However, the biofilm formation test measured by CV
staining and absorbance at 655 nm on 96-well polystyrene plates revealed that the
biomass production by GFP-labeled Salmonella strain 457-88 (OD655nm = 0.16 ± 0.07)
was significantly less than that of its parental strain (OD655nm = 0.28 ± 0.06), and the
OD655nm value of mixed-species biofilm formed by FP-labeled USDA 5 and 457-88
(OD655nm = 0.14 ± 0.03) was significantly less than that of mixed-species biofilm formed
by the plasmid-free pathogens (OD655nm = 0.20 ± 0.04) (Figure 5.7). It is likely the
populations of RFP-labeled E. coli O157:H7 strain USDA 5 were less than the parental
strain in both single- and mixed-species biofilms on 24-well plates (data not shown).
Additional metabolic burden may be posed by the expression of the label, causing a
lower growth rate of FP-labeled bacteria in biofilms. However, planktonic cell growth
rate of transformed bacteria was not affected, indicating there was no relationship
129
between planktonic cell growth rate and biofilm growth rate (Folsom, Siragusa, & Frank,
2006). Hence, underestimation could occur in the quantification of FP-labeled cells
instead of wild-type bacteria in biofilms.
Biofilms generally have enhanced resistance to antimicrobial treatments (Meyer,
2003). Bacteria may achieve high numbers on a surface due to high attachment with no or
slow growth or low attachment with growth. Since it is not possible to entirely prevent
bacterial attachment due to the ubiquitous nature of some bacteria and their ability to
form biofilms, our studies have focused on reducing the growth rate of bacteria by
developing a novel disinfectant consisting of levulinic acid and SDS (Chen, et al., 2014;
Zhao, et al., 2014). There is a synergistic antimicrobial effect between levulinic acid and
SDS because the application of levulinic acid or SDS individually at equivalent
concentrations is considerably less effective (Zhao, et al., 2011; Zhao, et al., 2014; Zhao,
et al., 2009, 2010). Combination of this organic acid and surfactant may promote the
detachment of bacterial cells from a biofilm matrix and kill the detached cells by the
undissociated form of levulinic acid. The antagonistic effect between E. coli O157:H7
strain USDA 5 and Salmonella strain 457-88 in which significantly (P < 0.05) less
biomass production and lower bacterial counts of strain USDA 5 in the mixed-species
biofilm may have contributed to the enhanced sensitivity of the two strains to the
levulinic acid plus SDS compared to the single-species biofilms. Images captured
previously by transmission scanning microscopy revealed that cellular leakage was
caused by levulinic acid plus SDS. The mode of antimicrobial action of levulinic acid
with SDS needs further research.
130
In conclusion, this study revealed that interspecies interactions within mixed
biofilms formed by E. coli O157:H7 and Salmonella had a pronounced influence on the
microbial composition and their resistance to levulinic acid plus SDS. E. coli O157:H7
strain USDA 5 and Salmonella strain 457-88 were antagonistic to each other in the
mixed-species biofilm, and both of the two strains had enhanced sensitivity to levulinic
acid plus SDS compared to their resistance in single-species biofilms. Transformation of
cells with FP-encoding plasmids, in addition to the application of CLSM, enabled us to
study in real-time, live, hydrated microbial biofilms without structural destruction.
Images captured by CLSM revealed temporal and spatial development of FP-expressing
cells in the biofilm matrix and the antimicrobial effectiveness of levulinic acid plus SDS
on the pathogens in both single- and mixed-species biofilms. However, FP-labeled cells
in biofilms grew more slowly than unlabeled cells and this should be considered when
explaining the results.
Acknowledgments
The authors thank Dr. Cathy Webb for providing RFP and GFP plasmids and Ping
Zhao for technical assistance. The study was supported by grants from the USDA
National Institute of Food and Agriculture, Agriculture and Food Research Initiative
Grant No. 2011-68003-30012 and Center for Food Safety, University of Georgia.
131
Figure 5.1. Single- and mixed-species biofilm formation by E. coli O157:H7 strain
USDA 5 and Salmonella strain 457-88 on 96-well polystyrene plates incubated at 21°C
for 72 h. Biofilm-forming ability was quantified by crystal violet staining and absorbance
measurement at 655 nm. The bars represent the mean values + standard deviations (n =
24). Different letters above the bars indicate a significant difference (P < 0.05).
132
Figure 5.2. Bacterial counts of E. coli O157:H7 strain USDA 5 and Salmonella strain
457-88 in single- and mixed-species biofilms after a treatment for 5 min with water
(control), QAC (150 ppm), 1% levulinic acid + 0.1% SDS, or 3% levulinic acid + 2%
SDS. Biofilms were incubated on 24-well polystyrene plates held at 21°C for 72 h before
treatment. The bars represent the mean values + standard deviations (n = 6). The
minimum detection limit by the direct plating method was 0.7 log CFU/ml.
133
Figure 5.3. Representative photomicrographs by CLSM of single- and mixed-species
biofilms formed by RFP-labeled E. coli O157:H7 strain USDA 5 and GFP-labeled
Salmonella strain 457-88 after a 5-min treatment with water (control, A, D and G), 1%
levulinic acid + 0.1% SDS (B, E and H), or 3% levulinic acid + 2% SDS (C, F and I).
134
USDA 5 (RFP)
457-88 (GFP)
Water
1% LVA + 0.1% SDS
3% LVA + 2% SDS
135
457-88 (GFP) + USDA 5 (RFP)
Figure 5.4. Representative photomicrograph by CLSM showing the three-dimensional
modeling of 72-h biofilms formed by mixed-species of RFP-labeled E. coli O157:H7
strain USDA 5 and GFP-labeled Salmonella strain 457-88. Scale bar = 10 µm.
136
Appendix
Table 5.1. Bacterial strains used in this study and their origins, curli and cellulose productions, and counts in single-species biofilms.
Counts (log
Bacterial strains
Origin
Curlia Celluloseb
CFU/ml) in single-
Reference or sources
species biofilmsc
E. coli O157:H7
E0122
Beef
+
-
7.8 ± 0.3
(Fischer, et al., 2001)
932
Human
+weak
-
8.6 ± 0.2
(Fischer, et al., 2001)
E0139
Venison
-
-
8.4 ± 0.1
(Fischer, et al., 2001)
E009
Cattle
-
-
8.4 ± 0.2
(Fischer, et al., 2001)
E0018
Cattle
-
-
8.3 ± 0.2
(Fischer, et al., 2001)
Whitewater G
Human
+weak
-
6.8 ± 0.3
(Zhao, Doyle, Zhao, Blake, & Wu, 2001)
Whitewater H
Human
-
-
7.3 ± 0.1
(Zhao, et al., 2001)
Whitewater I
Human
-
-
7.4 ± 0.3
(Zhao, et al., 2001)
Whitewater J
Human
-
-
7.6 ± 0.3
(Zhao, et al., 2001)
Whitewater K
Human
-
-
7.8 ± 0.3
(Zhao, et al., 2001)
Whitewater L
Human
-
-
7.6 ± 0.4
(Zhao, et al., 2001)
C7927
Human
-
-
7.8 ± 0.2
(Zhao, Doyle, & Besser, 1993)
E1352
Cattle
-
-
7.7 ± 0.1
CFSd
E0143
Cattle
-
-
8.0 ± 0.3
CFS
137
E0654
Feces
-
-
8.0 ± 0.3
CFS
E0144
Cattle
+weak
-
7.8 ± 0.2
CFS
WA Lot 16
Beef
-
-
8.0 ± 0.2
CFS
C0083
Feces
-
-
8.0 ± 0.3
CFS
C0286
Feces
-
-
7.9 ± 0.2
CFS
CA Lot 9 #18
Beef
-
-
7.9 ± 0.2
CFS
USDA 1
Beef
-
-
7.5 ± 0.2
(Jadeja, Hung, & Bosilevac, 2013)
USDA 2
Beef
-
-
8.2 ± 0.3
(Jadeja, et al., 2013)
USDA 3
Beef
-
-
8.0 ± 0.2
(Jadeja, et al., 2013)
USDA 4
Human
+weak
-
7.3 ± 0.4
(Jadeja, et al., 2013)
USDA 5
Human
++
-
9.2 ± 0.2
(Jadeja, et al., 2013)
H2662
Cattle
+
+
8.6 ± 0.3
(Chen, et al., 2014)
11942A
Cattle
+
+
8.8 ± 0.4
(Chen, et al., 2014)
13068A
Cattle
-
-
7.8 ± 0.2
(Chen, et al., 2014)
152N17-1
Dairy
+
+
8.2 ± 0.1
(Chen, et al., 2014)
H3279
Human
+
+
8.2 ± 0.6
(Chen, et al., 2014)
8748A-1
Cattle
+
+
8.3 ± 0.2
CFS
H3380
Cattle
-
-
8.6 ± 0.2
CFS
43
Beef
+
+weak
8.1 ± 0.2
CFS
S. Typhimurium
138
45
Beef
+
+weak
8.2 ± 0.2
(Zhao, Doyle, Fedorka-Cray, Zhao, & Ladely, 2002)
56
Beef
+
-
8.1 ± 0.2
CFS
62
Beef
+
+weak
8.2 ± 0.2
(Zhao, et al., 2002)
65
Beef
+
+weak
8.0 ± 0.1
CFS
Human
++
+weak
8.2 ± 0.2
(Zhao, et al., 2009)
E39
Egg
++
+
8.0 ± 0.4
(Zhao, et al., 2009)
564-88
Food
++
+weak
8.1 ± 0.5
(Zhao, et al., 2009)
457-88
Poultry
++
+
9.4 ± 0.1
(Zhao, et al., 2009)
S. Enteritidis
193-88
a
Curli production was determined by streaking isolates on tryptone agar plates containing 40 µg/ml of Congo red and 20 µg/ml of
Commassie blue. After incubation at 21°C for 48 h, strains were divided into four groups based on morphotypes: -, negative, white
colonies; +weak, weak positive, light brown colonies; +, positive, red colonies; ++, strong positive, dark red, dry and rough colonies.
b
Cellulose production was determined by streaking isolates on calcofluor-containing agar plates. After incubation at 21°C for 48 h,
strains were divided into three groups based on intensity of fluorescence under UV light (365 nm). -, negative, no fluorescence; +weak,
weak positive, low intensity of fluorescence; +, positive, high intensity of fluorescence.
c
Individual bacterial strains were grown in TSB at 37°C for 20 h. Overnight cultures (ca. 8.0 log CFU/ml) were diluted 1:100 in
sterile TSB and added in 24-well flat-bottom polystyrene plates (Costar) at 1 ml per well. The plates were incubated statically at 21°C.
After 72 h of incubation, the supernatant fluid was removed, and the wells were rinsed three times with 2 ml of PBS. The content of
139
each well was harvested by scraping the surface thoroughly with a sterile 6-inch (15.2 cm) polyester-tipped swab (Fisher Scientific,
Pittsburgh, PA). The PBS containing released biofilm cells and the swab were added to 9 ml of 0.1 M PBS and vigorously agitated by
a Vortex (G-560, Scientific Industries, Bohemia, NY) at maximum speed for 1 min. Appropriate dilutions (0.1 ml) in 0.1 M PBS were
spread plated in duplicate on TSA plates, which were incubated at 37°C for 48 h before bacterial count. Values are means ± standard
deviations.
d
CFS, Center for Food Safety, The University of Georgia.
140
Figure 5.5. Representative photomicrographs by CLSM of single- and mixed-species
biofilms formed by RFP-labeled E. coli O157:H7 strain USDA 5 and GFP-labeled
Salmonella strain 457-88 after 4, 8, 24, and 48 h of incubation at 21°C. Scale bar = 10
µm.
141
4h
8h
457-88 (GFP)
USDA 5 (RFP)
457-88 (GFP) + USDA 5
(RFP)
142
24 h
48 h
Figure 5.6. Comparison of planktonic growth rates of FP-labeled and parental E. coli
O157:H7 strain USDA 5 and Salmonella strain 457-88. Growth rates of FP-labeled and
parental bacterial strains were determined by measuring absorbance at 600 nm at 0, 1, 2,
4, 6, 8, 12, and 24 h after 1:1000 inoculation in TSB without ampicillin. Point values are
the means of at least two trials.
143
Figure 5.7. Single- and mixed-species biofilm formation by FP-labeled and parental E.
coli O157:H7 strain USDA 5 and Salmonella strain 457-88 on 96-well polystyrene plates
incubated at 21°C for 72 h. Biofilm-forming ability was quantified by crystal violet
staining and absorbance measurement at 655 nm. The bars represent the mean values +
standard deviations (n = 24). Different letters above the bars indicate a significant
difference (P < 0.05).
144
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CHAPTER 6
SUMMARY
Transfer of foodborne pathogens during mechanical slicing and their inactivation
by levulinic acid-based sanitizer on slicers
The cross-contamination of L. monocytogenes, S. Typhimurium, and E. coli
O157:H7 from contaminated deli foods to slicers and from contaminated slicers to deli
foods was determined. After slicing surface-inoculated deli food products, the pathogens
were recovered from all of the 5 food contact locations on slicers, with the transfer rate
being significantly (P < 0.05) less on blades than meat grips and carriage trays. With an
initial inoculation of ca. 8.5 log CFU of the pathogens/blade, the transfer of pathogens
decreased logarithmically from 4.0 to <1.5 log CFU/slice after 60 slices. Results revealed
that the transfer rate of pathogens between the slicer and foods depends on food contact
locations on slicers, the composition of the food, and the type of pathogens. Applying 1%
levulinic acid plus 0.1% SDS as a foam can reduce 6.0 to 8.0 log CFU of the three
pathogens/blade within 1 min on slicer surfaces. Hence, this combination of chemicals
could potentially be an effective sanitizer for large-scale applications in food processing
facilities.
Control of pathogens in biofilms on the surface of stainless steel by levulinic acid
plus sodium dodecyl sulfate
The combination of levulinic acid and SDS was more effective in reducing the
populations of the three pathogens in biofilms compared to other commonly used
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sanitizers, with the highest concentrations evaluated (3% LVA + 2% SDS) providing the
greatest log reduction. SEM images confirmed that the combination of LVA and SDS
was effective in inactivating bacterial cells in biofilms, and TEM images revealed that
LVA with SDS cause changes to the permeability of the pathogens’ cell membranes and
aggregation of their cytoplasmic components. Results of this study may have practical
application for mitigating the presence of biofilms in food processing facilities.
Microscopic images enable a better understanding of the antimicrobial behavior of LVA
plus SDS to foodborne pathogens within biofilms; however, additional research on the
antimicrobial mechanism of action of the LVA with SDS is needed.
Single- and mixed-species biofilm formation by Escherichia coli O157:H7 and
Salmonella, and their sensitivity to levulinic acid plus sodium dodecyl sulfate
Interspecies interactions within mixed biofilms formed by E. coli O157:H7 and
Salmonella had a pronounced influence on the microbial composition of the biofilms and
the resistance of these pathogens to levulinic acid plus SDS. E. coli O157:H7 strain
USDA 5 and Salmonella strain 457-88 were antagonistic to each other in a mixed-species
biofilm, and both of the two strains had enhanced sensitivity to levulinic acid plus SDS
compared to their resistance in single-species biofilms. Transforming these pathogens
with fluorescent protein (FP)-encoding plasmids, then observing them in biofilms with
CLSM enabled us to study in real-time, live, hydrated microbial biofilms without
structural destruction. Images captured by CLSM revealed temporal and spatial
development of FP-expressing cells in the biofilm matrix and the antimicrobial
effectiveness of levulinic acid plus SDS on the pathogens in both single- and mixed-
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species biofilms. However, FP-labeled cells in biofilms grew more slowly than unlabeled
cells, and this should be considered when explaining the results.
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