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Accepted Manuscript
Activity of thyme and tea tree essential oils against selected foodborne pathogens in
biofilms on abiotic surfaces
Mohammad Sadekuzzaman, Md Furkanur Rahaman Mizan, Hyung-Suk Kim,
Sungdae Yang, Sang-Do Ha
PII:
S0023-6438(17)30795-8
DOI:
10.1016/j.lwt.2017.10.042
Reference:
YFSTL 6605
To appear in:
LWT - Food Science and Technology
Received Date: 14 April 2017
Revised Date:
16 October 2017
Accepted Date: 20 October 2017
Please cite this article as: Sadekuzzaman, M., Mizan, M.F.R., Kim, H.-S., Yang, S., Ha, S.-D., Activity of
thyme and tea tree essential oils against selected foodborne pathogens in biofilms on abiotic surfaces,
LWT - Food Science and Technology (2017), doi: 10.1016/j.lwt.2017.10.042.
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Title: Activity of Thyme and Tea Tree Essential Oils Against Selected Foodborne
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Pathogens in Biofilms on Abiotic Surfaces
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Authors: Mohammad Sadekuzzamanab, Md. Furkanur Rahaman Mizana, Hyung-Suk Kima,
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Sungdae Yanga, Sang-Do Haa*
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a
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Anseong, Gyunggido 456-756, South Korea
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b
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School of Food Science and Technology, Chung-Ang University, 72-1 Nae-Ri, Daedeok-Myun,
Department of Livestock Services, People’s Republic of Bangladesh
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*Corresponding author:
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Sang-Do Ha,
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School of Food Science and Technology,
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Chung-Ang University, 72-1 Nae-Ri, Daedeok-Myun,
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Anseong-Si, Gyunggido 456-756, South Korea
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Tel.: +82 031-670-4831
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Fax: +82 031-675-4853
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Running head: Essential oil antibiofilm activity
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Abstract
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Owing to their preservative and antimicrobial effects, essential oils (EOs) are promising natural
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ingredients for the food industry. The main objective of this study was to investigate the activity
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of thyme and tea tree oils against selected foodborne pathogens in biofilm mode. The major
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compounds of these EOs were analyzed by gas chromatography-mass spectrometry (GC-MS)
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and their antimicrobial activity was determined by a standard broth dilution assay. Biofilms were
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formed by Escherichia coli O157:H7, Listeria monocytogenes, and Salmonella spp. on abiotic
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surfaces and were treated with EOs at the minimum inhibitory concentration (MIC) and 0.1 %
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(v/v) for 2 h. Our results demonstrate that EO treatment reduced biofilm cells up to 3.5 log
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CFU/cm2, 2.1 log CFU/cm2, and 2.5 log CFU/peg on stainless steel (SS), rubber, and minimum
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biofilm eradication concentration (MBECTM) surfaces, respectively. Structural changes of the
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biofilm after exposure to EOs was confirmed by field-emission scanning electron microscopy
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and viability of biofilm cells was observed using a confocal laser scanning microscope. Overall,
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these results suggest that EOs could be used to reduce foodborne pathogens in biofilms.
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Keywords: Essential oil, Biofilm, Escherichia coli O157:H7, Listeria monocytogenes,
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Salmonellaspp.
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1.Introduction
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One of the predominant causes of morbidity and mortality worldwide is foodborne diseases,
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which pose a considerable impediment to socio-economic development
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Approximately one in ten people falls ill with foodborne diseases and, as a result, 420,000 people
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(WHO, 2015).
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die every year (WHO, 2015). According to the Center for Disease Control (CDC), fifteen
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pathogens are associated with approximately 95% of foodborne illness, hospitalization, and
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death in the United States (Scallan et al., 2011), among which Escherichia coli O157:H7,
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Listeria monocytogenes, and Salmonella spp. are the main causal agents. The control of these
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foodborne pathogens . gained much attention because these pathogens can adhere, colonize, and
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produce resilient biofilms on different surfaces such as stainless steel (SS), rubber, and glass
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(Brandl, 2006; Gandhi & Chikindas, 2007; Murphy, Carroll, & Jordan, 2006). Biofilm formation
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is a microbial activity that poses threats to food industry. Microbial colonization and subsequent
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biofilm formation in food-processing environments represent potential sources of contamination,
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which causes spoilage of food products, thereby reducing shelf life and favoring disease
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transmission and, ultimately, foodborne outbreaks. Moreover, it can also reduce sanitizer
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efficacy and heat transfer efficiency, produce metal corrosion in pipelines and tanks, or even
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make obstructions in heat equipment.
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Biofilm eradication is challenging because microorganisms in biofilms are much more resistant
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to the conventional antimicrobial treatments currently employed in the food industry than
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microorganisms in isolated colonies (Simões, Simões, & Vieira, 2009). The resistance
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mechanism of bacterial biofilms might be due to the complex architecture of the exopolymeric
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substances with which the microorganisms are associated, which shift the phenotype and
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metabolic activity of the microbial population in the biofilm state (Meyer, 2003; Sharma
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&Anand, 2002). Currently, several strategies have been proposed and evaluated to inactivate or
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reduce biofilms from food-processing environments. However, owing to their limited efficacy,
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emergence of microbial resistance, high cost, and food safety issues, new safe, cost-effective,
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and environmentally friendly antibiofilm strategies such as cold oxygen plasma, ultraviolet
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irradiation, ultrasound, natural substances, quorum sensing inhibition, antimicrobial coating, and
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bacteriophages are being extensively evaluated (Srinivasan et al., 2008). Several studies
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demonstrating the necessity for new strategies for the elimination of biofilms and/their effective
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control in food industries have been reported.
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Molecules with strong antimicrobial activity mostly belong to phytoalexins, of which essential
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oils (EOs) are the most important members. EOs are aromatic volatile secondary metabolites
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derived from the roots, seeds, flowers, fruits, and leaves of plants, which have been used for
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therapeutic purposes for nearly 6,000 year as natural medicines. Despite it’s aromatic flavor,
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essential oils which exert specific antimicrobial activities against many foodborne pathogens, are
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a promising natural antimicrobial alternative to chemical sanitizers. Among EOs, thyme and tea
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tree oils have been extensively reported as strong antimicrobial agents (Brady, Loughlin, Gilpin,
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Kearney, & Tunney, 2006; Budzyńska, Wieckowska-Szakiel, Sadowska, Kalemba, &Rózalska,
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2011; Szczepanski& Lipski 2014). Thyme EO is extracted from the leaf of the thyme plant and
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have a high in thymol content, which is commonly used to add flavor to a variety of meals. This
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oil has cleansing properties that can be exploited to clean surfaces and remove dirt, grime, and
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unpleasant odors without the use of chemicals harmful to humans. Additionally, tea tree oil,
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which is extracted from the shrub-like tree called Melaleuca alternifolia, is also a popular EO
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used worldwide. This oil is comprised mainly by terpenes, monoterpenes, sesquiterpenes, and
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their corresponding alcohols, and its uses throughout history have been extensive.
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These EOs have been evaluated against various pathogens such as E. coli, L. monocytogenes,
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Pesudomonas aeruginosa, and Staphylococcus aureus( Brady et al.,2006). However, study on
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the effectiveness of thyme and tea tree oils against food borne pathogens in single species
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biofilm of E. coli O157:H7, L. monocytogenes and Salmonella spp. has not been studied
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extensively. In this context, the main objective of this study was to determine whether these EOs
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could be used to reduce selected foodborne pathogens in biofilms on abiotic surfaces at 30 °C,
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which is a favorable temperature for biofilm formation on food contact surfaces (Else, Pentle &
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Amy, 2003)
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2. Materials and methods
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2.1. Bacterial strains and culture condition
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In this study, E. coli O157:H7 (NCCP 11090), L. monocytogenes(ATCC19113), Salmonella ser.
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Enteritidis (ATCC13076), and Salmonella ser. Typhimurium (ATCC14028) were used. Bacteria
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were stored at −70 °C in tryptic soy broth (TSB; BD Diagnostics, Franklin Lakes, NJ, USA)
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containing 15% (v/v) glycerol . Individually, each strain was consecutively sub-cultured twice
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aerobically at 37 °C for 24 h. Cultures were then centrifuged (4000 ×g at 4 °C for 20 min) and
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washed twice with sterile phosphate-buffered saline (PBS; pH 7.4).
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2.2. Thyme and tea tree EO preparation
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Essential oils (thyme and tea tree) were kindly provided by Bolak Co., Ltd, South Korea. Each
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essential oil emulsion was prepared as described previously (Turgis, Vu, Dupont, & Lacroix,
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2012) with some modifications. EO was mixed with TSB containing 2% dimethyl sulfoxide
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(DMSO, Sigma- Aldrich, St. Louis, MO, USA) and 0.5% Tween 80 (Sigma- Aldrich, St. Louis,
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MO, USA). The mixture was then stirred for 30 min to form a stable emulsion for 24 h. DMSO
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and Tween 80 were added in medium to enhance EOs solubility and stabilize the emulsion.
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2.3. Quantitative analysis of the EOs constituents by gas chromatography/mass
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spectroscopy (GC-MS)
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Determination of the detailed chemical composition of the EOs was performed by GC-MS (Gas
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chromatography Agilent Co Ltd, 6890 N, USA; Mass spectrometer Agilent Co Ltd, 5975 C,
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USA),as previously described (Kim, Lee, Kim, Baek, & Lee, 2015). Briefly, the gas
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chromatographer was equipped with a fused silica capillary column DB-WAX (60 m ×
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250µm×0.25µm, Agilent Technologies (Santa Clara, California, United States), the ionizing
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energy was set at 70 eV, and helium was used as the carrier gas at a constant flow rate of 1
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mL/min. The oven temperature was maintained at 50 °C for 5 min and then increased to 230 °C
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(3 °C/min). EOs samples of 1 µL (split ratio 100:1) were injected manually. The relative
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percentages of EO components were calculated by normalizing the peak areas. The chemical
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constituents of the EOs were identified according to their GC retention times and computer
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matching with the mass spectral libraries of Wiley (7n.1) and National Institute of Standards and
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Technology (NIST).
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2.4. Determination of the minimum inhibitory concentration (MIC) and minimum
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bactericidal concentration (MBC)
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MIC determination was performed according to the microbroth dilution method as previously
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described (Branen & Davidson, 2004). Overnight bacterial culture was diluted to about 106
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CFU/mL and 120 µL of culture diluted in TSB was added into each well of a 96-well microtiter
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plate (Corning Incorporated, Corning, NY, USA). The EO solutions were serially diluted in TSB
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from 0.001 to 5% (v/v). The diluted EO solutions were mixed with equal volumes of bacterial
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cultures in each well. Plates were incubated at 30 °C (for L. monocytogenes) or 37 °C (for E.coli
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O157:H7 and Salmonella spp.). The MIC was defined as the lowest concentration that inhibits
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visible bacterial growth (Clinical and Laboratory Standards Institute guidance, 2010), . The
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MBC was determined by spreading 100 µL onto MacConkey agar containing sorbitol (for E. coli
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O157:H7), PALCAM agar, (for L. monocytogenes), and xylose-lysine deoxycholate (XLD)
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agar(CM0469, for Salmonella spp.) and incubated at 30 °C for 48 h (L. monocytogenes) or 37 °C
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for 24 h (E.coli O157:H7 and Salmonella spp.). . MBC was defined as the EO concentration
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corresponding to complete growth inhibition of bacteria (Branen & Davidson, 2004). For both
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MIC and MBC determination in positive control diluted DMSO and Tween in TSB were mixed
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with equal volume of bacterial culture, and as a negative control essential oils were mixed with
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TSB without bacterial culture. For each case of MIC and MBC six replicates have been
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evaluated .
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2.5. Biofilm formation and EO effect on biofilm cells on stainless steel (ss) and rubber
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surfaces
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Effectiveness of the EOs in removing biofilm cells from SS and rubber coupons was tested after
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inoculation with different organisms. SS (2 × 2 × 0.1 cm, type: 304) coupons were processed as
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described previously (Shen et al., 2012). Rubber (Latex, natural rubber, 0.06 cm thickness) was
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aseptically cut into pieces (2 × 2 cm). The pieces were sterilized by dipping into 70% ethanol for
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10 minutes followed by switching on UV light for 1 hour and air-drying in a biosafety cabinet.
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The overnight bacterial culture culture was diluted 1:50 and inoculated in TSB in 50-mL Falcon
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tubes; each tube contained one SS and one rubber coupon that were completely submerged in 7
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mL of TSB to enable biofilm formation. The tubes were incubated without shaking at 30 °C for
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72 h with a change of medium every 24 h to develop mature biofilms on both coupons. After
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biofilm formation, coupons were removed from the tube and washed three times with PBS to
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remove planktonic bacteria. The coupons were submerged in a tube containing 7 mL of TSB
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with previously prepared EOs at MIC and0.1 % (v/v) concentrations. Control coupons were
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submerged in TSB containing 2% DMSO and 0.5% Tween 80. Then, the coupons were
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incubated for 2 h at the same conditions (30 °C) at which the biofilms were developed.
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Following incubation, each SS and rubber coupon was removed from the tube and transferred to
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a small petri dish (55 × 12 mm) containing5 mL of 0.1% buffered peptone water (BPW),
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scrubbed, and transferred to a test tube, and vortexed for 1 min to disperse the biofilm. The
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solution was centrifuged for 2 minutes at 12000 × g, then cells were ten fold diluted in BPW for
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counting. The samples were spread onto MacConkey agar containing sorbitol (for E. coli
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O157:H7), PALCAM agar, (for L. monocytogenes), and xylose-lysine deoxycholate (XLD) agar
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(CM0469, for Salmonella spp.) and incubated at 30 °C for 48 h (L. monocytogenes) or 37 °C for
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24 h (E.coli O157:H7 and Salmonella spp.).
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2.6. Antibiofilm activity of EOs tested on mimimum biofilm eradication concentration
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(MBECTM) Biofilm device
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The effectiveness of EOs against biofilm cells was tested using the MBECTM (Innovotech Inc.,
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Edmonton Canada) biofilm inoculator consisting of a lid with 96 pegs and individual wells.
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Biofilms were established on the polystyrene pegs (108.9 mm2/peg) of the MBECTM with some
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modification as described early (Ceri et al., 2001). Briefly, 150 µL of each bacterial inoculum
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(105 CFU/mL) was added to each well of the base. Then, the peg lid was placed onto the
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microtiter base. The device was placed on a platform shaker set at 110 rpm in a humidified
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incubator at 30°C.The plate was incubated for 16 h. After biofilm formation, the lid was placed
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onto another 96-well plate containing 200 µL of normal saline solution in each well for 10 s to
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remove planktonic bacteria attached to the pegs. Immediately after rinsing, the MBECTM lid was
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transferred onto the challenge plate containing 150 µL of EOs preparations (MIC level and 0.1%
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v/v) in each well and 150 µL of normal saline solution in the control well. The plate was
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incubated for 2h at 30°C or 37°C. Then the plate(s) were placed in a dry stainless steel insert tray
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which sits on the water of the sonicator (Power Sonic 505, Hwashin Technology Co. Repupublic
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of Korea) and sonicate at room temperature (21± 1°C) on high for 10 minutes for dislodge the
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biofilm cells and plated on selective agar plates.
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2.7. Field-emission scanning electron microscopy(FE-SEM) for biofilms
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Samples of the EO-treatment and control groups were prepared as previously described (Jahid,
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Lee, Kim, & Ha, 2013), with slight modifications. The samples were fixed with 2.5%
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glutaraldehyde in PBS at room temperature for 4 h. The coupons were then serially treated with
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ethanol (i.e., 50% for 15 min, 60% for 15 min, 70% for 15 min, 80% for 15 min, 90% for 15 min,
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and 100% twice for 15 min) and successively dehydrated by soaking in 33, 50, 66, and 100%
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hexamethyldisilazane in ethanol for 15 min each. The dehydrated coupons were sputter-coated
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with platinum and visualized using an FE-SEM (Hitachi/Baltec, S-4700).
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2.8. Confocal laser scanning microscopy (CLSM)
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The Film Tracer Live/Dead biofilm viability kit (Molecular probes®, USA) was used to
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distinguish live and dead bacteria in biofilms according to the manufacturer’s protocol. The
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LIVE/DEAD biofilm viability kit contains a mixture of SYTO® 9 green-fluorescent nucleic acid
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stain specific for intact live bacteria, and propidium iodide red fluorescent nucleic acid stain
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specific for membrane-damaged or non-viable bacteria. In this experiment, two types of cells
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were observed: green cells representing intact or viable cells, and differently red-shaded cells
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representing damaged or non-viable bacterial cells after treatment. The formation of static
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biofilms was visualized using a confocal laser microscope (Carl Zeiss LSM 710) equipped with a
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40× objective. Argon laser excitation was at 488 nm and emission was at 500–550 nm. For each
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experiment, at least 10 random fields of four independent cultures were chosen for microscopic
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analysis.
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2.9. Statistical Analysis
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One-way analysis of variance (ANOVA) followed by Bonferroni’s post-test was performed
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using GraphPad Prism 5.03 for Windows (GraphPad Software, San Diego California USA;
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www.graphpad.com). All experiments were conducted at least twice and plotted with mean ± SE.
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***P<0.001, **P<0.01, and *P<0.05 compared to the control group.
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3. Results and Discussion
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The major constituents of thyme and tea tree EO, which were analyzed by GC-MS, are listed in
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Supplementary tables 1 and 2. The results showed that cymene , thymol , alpha pinene and
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carvacrol , were the main components of thyme oil, whereas terpinen , gamma terpinen, alpha
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terpine , and beta fenchol were predominant in tea tree EO. Thesedata agree with previously
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published results with some variations (Szczepanski & Lipski, 2014; Millezi, Cardoso, Alves, &
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Piccoli, 2013; Carson, Hammer, & Riley, 2006). Several studies have confirmed that EOs from
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plants of the same species are composed of different chemicals depending on the geographical
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region where the plant grew. This compositional difference is attributed to several factors such as
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cultivation conditions, genetics, climate, soil type, and rainfall amount (Gobbo-Neto & Lopes,
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2007).
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The MIC and MBC values of thyme and tea tree EO against the pathogens tested are shown in
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Table 1. The of MIC and MBC values were lowerfor thyme oil than for tea tree oil, . . ;
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. Before treatment with EOs, average bacterial counts in biofilm on SS coupon for E. coli
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O157:H7, L. monocytogenes, S. Enteritidis, and S. Typhimurium were presentedin Fig. 1 & 2.
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The levels of E. coli O157:H7 biofilm cells were reduced by 1.4 and 3.0 log CFU/cm2 when
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treated with thyme EO and by 1.8 and 2.8 log CFU/cm2 when treated with tea tree EO at MIC
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and 0.1% v/v, respectively on SS (Fig 1 & 2). . The levels of Salmonella spp. biofilm cells were
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reduced by 1.8 and 3.3 log CFU/cm2 when treated with thyme EO and by 2.3 and 3.1 log
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CFU/cm2 when treated with tea tree EO at MIC and 0.1% v/v, respectively. The reduction of L.
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monocytogenes biofilm cells was 1.5 and 3.3 log CFU/cm2 when treated with thyme EO and 2.0
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and 3.2 log CFU/cm2 when treated with tea tree EO at MIC and 0.1% v/v, respectively.
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On rubber surface, the initial biofilm populations of E. coli O157:H7, L. monocytogenes, S.
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Enteritidis, and S. Typhimurium were shown in Fig. 3 & 4. Figures 3 and 4 show the inactivation
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effect of thyme and tea tree EOs on E. coli O157:H7, L. monocytogenes, Salmonella spp. biofilm
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cells on rubber surface. The overall reduction patterns were lesser than those found on the SS
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surface.
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Biofilm growth on the pegs of the MBECTM device and biofilm cells susceptibility to EOs are
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shown in Figure 5 & 6. . EO treatment caused biofilm reduction up to 2.0 and 2.5 log CFU/peg
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(thyme oil), and 2.1and 2.2 log CFU/peg (tea tree EO) at MIC and 0.1 %, respectively (Fig. 5 &
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6).
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The results demonstrated that all the tested pathogens could develop biofilms on tested surfaces.
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Although bacterial populations in the single species biofilm differed (P< 0.05) depending on the
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strain, with L. monocytogenes forming the densest population and S. typhimurium forming the
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least dense population, similar biofilm inactivation patterns were observed for all the tested
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pathogens. These results indicate that thyme and tea tree EOs could reduce bacterial population
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in biofilms formed by the tested strains.
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Other studies also confirmed the effect of thyme and tea tree EOs on bacterial biofilms. Millezi,
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Cardoso, Alves, & Piccoli (2012) found that a detergent-sanitizing solution of thyme EO reduced
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the biofilm cells of Aeromonas hydrophila by 3.84 log CFU/cm2. Desai, Soni, Nannapaneni,
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Schilling, & Silva (2012) demonstrated that thyme oil at 0.5% concentration was adequate to
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completely eliminate a 4-day old L. monocytogenes biofilm (7 log CFU/coupon). In a related
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study,Soni et al. (2013) observed that 0.05%–0.1% v/v thyme oil could reduce a 7-log CFU/cm2
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S. Typhimurium biofilm mass to a non-detectable level on both polystyrene and SS within 1 h of
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incubation.
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The concentration value is crucial when EOs are used as antibiofilm agents. It is worth noting
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that high concentrations are needed when EOs are used against foodborne pathogens in
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preformed biofilms (Kavanaugh & Ribbeck, 2012). In the present study, both EO concentrations
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used ( MIC and 0.1% v/v) can reduce biofilm cells significantly (P< 0.05), even though a
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stronger effect was observed when higher concentration of essential oils used.
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The biofilms on SS and rubber surfaces were observed by FE-SEM with and without 0.1% v/v
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EOs treatment. FE-SEM images show that untreated SS and rubber surfaces were colonized with
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adherent cells forming a compact and dense biofilm (Fig 7A–C). Exposure to EOs for 2 h
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considerably altered the morphology of biofilm cells compared to that in the control group (Fig
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7D–I).
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The effect of EOs on biofilm cells was evaluated on SS coupons to facilitate confocal imaging. .
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As shown in Figure 8 A-C, a non-treated biofilm consists of thick and densely populated live
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colonies (green color), whereas many dead cells colored in red were observed in the samples
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treated with EOs (Figure 8 D-I).
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The actual antimicrobial EO action mechanism on biofilms is complex and has not been
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completely elucidated. However, EOs components can diffuse through the polysaccharide matrix
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of the biofilm and destabilize its architecture by means of their strong intrinsic antimicrobial
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properties (Oral et al., 2010). EOs initiates permeability of bacterial cytoplasmic membrane,
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which results in cell content leakage loss of essential molecules and ions causes ultimate cell
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death (Oliveira, Brugnera, Cardoso, Alves, & Piccoli, 2010).
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4. Conclusions
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In conclusion, we demonstrated that thyme and tea tree EOs are effective in reducing E. coli
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O157:H7, L. monocytogenes, and Salmonella spp. Levels in biofilms on simulated food
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processing surfaces. Future studies will include the identification of the EO active ingredients
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specifically possessing antibiofilm activity, and the study of the molecular mechanism through
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which these ingredients exhibit their action. In the present study, we tested only two EOs, but
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many other oils exist that could be used as alternatives to current antibiofilm approaches.
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Acknowledgements
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This research was supported by the Basic Science Research Program through the National
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Research Foundation of Korea (NRF), which is funded by the Ministry of Science, ICT & Future
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Planning (2016R1A2B4007960) and by the Chung –Ang University Research scholarship grants
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in 2016-2017.
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Conflict of Interest
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No conflict of interest declared.
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Figure Legends
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Fig. 1 Biofilm cells grown at 30 °C for 72 h on stainless steel surface before and after EOs
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treatment (MIC level) for 2 h. Values represent means ± standard error (SE, n=3). *
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Fig. 2 Biofilm cells grown at 30 °C for 72 h on stainless steel surface before and after EOs
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treatment (0.1 % v/v) for 2 h. Values represent means ± standard error (SE, n = 3).
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Fig. 3 Biofilm cells grown at 30 °C for 72 h on rubber surface before and after EOs treatment
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(MIC level) for 2h. Values represent means ± standard error (SE, n = 3).
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Fig. 4 Biofilm cells grown at 30 °C for 72 h on rubber surface before and after EOs treatment
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(0.1% v/v) for 2h. Values represent means ± standard error (SE, n = 3).
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Fig. 5 Biofilm eradication from the MBECTM Biofilm device upon EOs treatment (MIC level).
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Values represent means ± standard error (SE, n = 6). *
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Fig. 6 Biofilm eradication from the MBECTM Biofilm device upon EOs treatment (0.1% v/v).
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Values represent means ± standard error (SE, n = 6). *
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Fig. 7 Field-emission scanning electron micrographs of biofilms formed over 72 h on stainless
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steel before and after EOs treatment (0.1% v/v for 2 h).(A) Escherichia coli O157:H7 biofilm
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(control), (B) Listeria monocytogenes biofilm (control), (C) Salmonella spp. biofilm (control),(D)
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E. coli O157:H7 biofilm (thyme oil treatment), (E) L. monocytogenes biofilm (thyme oil
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treatment), (F) Salmonella spp. biofilm (thyme oil treatment), (G) E. coli O157:H7 biofilm (tea
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tree oil treatment), (H) L. monocytogenes biofilm (tea tree oil treatment), (I) Salmonella spp.
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biofilm (tea tree oil treatment).
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Fig 8. Live/dead confocal laser scanning microscopic images of biofilms treated with thyme and
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tea tree essential oils. Green cells represent live cell and red cells represent dead cells.(A)
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Escherichia coli O157:H7 biofilm (control), (B) Listeria monocytogenes biofilm (control), (C)
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Salmonella spp. biofilm (control), (D) E. coli O157:H7 biofilm (thyme oil treatment), (E) L.
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monocytogenes biofilm (thyme oil treatment), (F) Salmonella spp. biofilm (thyme oil treatment),
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(G) E. coli O157:H7 biofilm (tea tree oil treatment), (H) L. monocytogenes biofilm (tea tree oil
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treatment), (I) Salmonella spp. biofilm (tea tree oil treatment).
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Table 1. MICs and MBCs of thyme and tea tree oils against E. coli O157:H7, L. monocytogenes,
S. Enteritidis, and S. Typhimurium.
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MBC (%)
0.03 (0.02-0.04)
0.12 (0.09-0.2)
0.07 (0.05-0.09)
0.07 (0.05-0.09)
0.25 (0.1-0.3)
0.30 (0.20-0.50)
0.15 (0.9-0.2)
0.15 (0.9-0.2)
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0.01(0.008-0.02)
0.06 (0.04-0.07)
0.03 (0.02-0.05)
0.03 (0.02-0.04)
0.08 (0.05-0.09)
0.09 (0.07-0.1)
0.07 (0.05-0.08)
0.07 (0.05-0.08)
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Bacteria
E. coli O157:H7
L. monocytogenes
S. Enteritidis
S. Typhimurium
E. coli O157:H7
L. monocytogenes
S. Enteritidis
S. Typhimurium
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Thyme
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Highlights:
Chemical composition of thyme and tea tree essential oil.
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Antimicrobial activity of thyme and tea tree EO E.coli O157:H7, L. monocytogenes
and Salmonella spp.
Thyme and tea tree oils reduce biofilms significantly.
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