Microbiocidal activity of selected weak organic acids
Zoltan Nack MSc. MBA. MASM.
PhD in Biology
The University of Newcastle
November 2012
Table of contents
Table of contents ...... ii In memorium ...... vi Statement of originality...... vi Acknowledgements ...... vii List of abbreviations, tables and figures ...... viii List of abbreviations ...... viii List of tables ...... xiii List of figures ...... xv Abstract ...... xix 1 Literature review ...... 1 1.1 Weak organic acids ...... 1 1.1.1 Historical perspective...... 2 1.1.2 Chemical and physical characteristics ...... 3 1.1.3 Toxicity ...... 4 1.1.4 Biodegradability ...... 6 1.2 Antiviral effect of weak organic acids ...... 8 1.3 Antibacterial effect of weak organic acids ...... 10 1.3.1 Overview of bacterial species used in this study ...... 10 1.3.2 Factors affecting antibacterial activity of weak organic acids ...... 11 1.3.3 Mechanism of antibacterial activity of weak organic acids ...... 12 1.3.4 Antibacterial activity of products containing weak organic acids ...... 14 1.3.5 Activity of weak organic acids against specific bacterial species ...... 15 1.3.6 Practical use of weak organic acids ...... 16 1.4 Development of acid tolerance ...... 17 1.4.1 Formation of small colony variant Staphylococcus aureus ...... 19 1.5 Antifungal effects of weak organic acids ...... 20 1.6 Biofilm formation ...... 21 1.6.1 Penetration of antimicrobials into biofilms...... 26 1.7 In situ testing ...... 27 1.7.1 Rational for in situ testing of disinfectant ...... 27 1.7.2 Rational for reduction of environmental contamination in hospitals ...... 28 1.8 Aims and conclusion ...... 29 2 Materials and Methods ...... 30 2.1 Microbes ...... 30 2.1.1 Viruses ...... 30 2.1.2 Intracellular bacteria ...... 30 2.1.3 Bacteria ...... 30 ii | P a g e
2.1.4 Yeast and fungi ...... 31 2.2 Disinfectant ...... 31 2.3 Reagents and microbiological media ...... 33 2.4 Antiviral efficacy study methodology ...... 36 2.4.1 Growth of virus in adherent cell lines ...... 36 2.4.2 Virus titration (infectivity titer) ...... 37 2.4.3 Disinfectant viral efficacy test ...... 37 2.5 Disinfectant intracellular bacteria efficacy study methodology ...... 38 2.6 Disinfectant antifungal efficacy study methodology ...... 39 2.7 Disinfectant antibacterial efficacy study methodology- planktonic bacteria ..... 41 2.7.1 Minimum Inhibitory Concentration ...... 41 2.7.2 Time-Kill Assay ...... 43 2.7.3 In house efficacy tests ...... 44 2.7.4 Therapeutic Goods Administration suspension test ...... 48 2.7.4 Therapeutic Goods Administration hard surface carrier test ...... 50 2.8 Disinfectant antibacterial study methodology- biofilm bacteria ...... 51 2.8.1 Microtitre plate method...... 51 2.8.2 Scanning Electron Microscopy method ...... 55 2.9 Acid tolerance study methodology (Staphylococcus aureus) ...... 56 2.9.1 Microtitre plate method...... 56 2.9.2 Microarray assay ...... 59 2.10 Disinfectant artificial and genuine in situ study ...... 66 2.10.1 Artificial in situ test in ‘dirty condition’ on tiles ...... 66 2.10.2 Disinfectant genuine in situ test validation against VRE- The Barwon Health trial ...... 67 3 Antiviral efficacy of weak organic acid disinfectants ...... 72 3.1 Results ...... 72 3.2 Discussion ...... 74 4 Efficacy of weak organic acid disinfectants against intracellular bacteria ...... 75 4.1 Results ...... 75 4.2 Discussion ...... 77 5 Antifungal efficacy of weak organic acids ...... 78 5.1 Results ...... 78 5.2 Discussion ...... 80 6 Antibacterial efficacy of weak organic acids against planktonic bacteria ...... 82 6.1 Minimum inhibitory concentrations of weak organic acids against antibiotic- resistant bacteria...... 82 6.1.1 Results and discussion ...... 83 iii | P a g e
6.2 Time-Kill Assay results of weak organic acids against antibiotic-resistant bacteria 85 6.2.1 Results and discussion ...... 85 6.3 In house suspension test results of weak organic acids against antibiotic-resistant bacteria ...... 89 6.3.1 Results and discussion ...... 89 6.4 In house test of weak organic acids against antibiotic-resistant bacteria using hard surface carriers ...... 92 6.4.1 Results and discussion ...... 92 7 Antibacterial efficacy of weak organic acid disinfectants against biofilm bacteria ...... 94 7.1 Introduction ...... 94 7.2 Microtitre plate results and discussion ...... 95 7.3 Scanning electron microscopy results and discussion...... 104 8 Development of acid tolerance by Staphylococcus aureus...... 136 8.1 Introduction ...... 136 8.2 Results ...... 138 8.2.1 Total RNA extraction ...... 139 8.2.2 c-DNA synthesis ...... 140 8.2.3 RNA degradation ...... 141 8.2.4 c-DNA purification ...... 142 8.2.5 c-DNA fragmentation and labeling ...... 142 8.2.6 c-DNA quantitation by real time PCR (Ct value) ...... 143 8.2.7 Hybridization ...... 145 8.2.8 Data extraction and analysis ...... 146 8.2.9 List of up-regulated genes categorized by function ...... 152 8.2.10 List of down-regulated genes categorized by function ...... 156 8.3 Discussion ...... 158 9 Artificial and genuine in situ studies ...... 161 9.1 Introduction ...... 161 9.2 Artificial in situ study on ceramic tiles ...... 162 9.2.1 Results and discussion ...... 162 9.3 Genuine in situ study of VRE disinfection at Barwon Health ...... 166 9.3.1 Results ...... 166 9.3.2 Discussion ...... 172 10 Overall discussion ...... 175 11 Presentations, publications and awards ...... 182 12 References ...... 183 13 Appendices ...... 225 13.1 Appendix A...... 225 13.2 Appendix B ...... 229 iv | P a g e
13.3 Appendix C ...... 239 13.4 Appendix D...... 242
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In memorium
In memory of my Mum, who was always behind me in good and bad times and passed away just a few weeks before the completion of this thesis.
Statement of originality
This thesis contains no material which has been accepted for the award of any other degree or diploma in any university or other tertiary institution and, to the best of my knowledge and belief, contains no material previously published or written by another person, except where due reference has been made in the text. I give consent to this copy of my thesis, when deposited in the University Library, being made available for loan and photocopying subject to the provisions of the Copyright Act 1968.
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Acknowledgements
The completion of this thesis would not have been possible without the help and assistance of my supervisors, Professor Tim Roberts from The University of Newcastle and Associate
Professor Stephen Graves from The Australian Rickettsial Reference Laboratory.
A special thanks to Associate Professor John Stenos from The Australian Rickettsial
Reference Laboratory for all his advice and support right throughout of this project.
I would like to thank Professor Hugh Dunstan from The University of Newcastle for giving me the opportunity to carry out this project.
I would like to thank colleagues and collaborators who helped to complete certain chapters of this thesis. To Dr. Fiona Collier from Barwon Biomedical Research, (Barwon Health) and to
Dr. Tracy Webster from AgriBioscience (La Trobe University) for the acid tolerance study including the microarray analysis.
To Dr. Andrew Sullivan from Deakin University for help in the SEM study of this project.
Special thanks to Dr. Sam Vonarx from Box Hill Institute on his advice on RNA extraction and c-DNA synthesis.
A very special thanks to the following people from Barwon Health, The Geelong Hospital for helping me on the in situ study
Stuart Marshall Environmental Services
Dorothy Marshall Environmental Services
Carling Southall Barwon Biomedical Research
Danielle Kennedy Barwon Biomedical Research
A very special thanks to my family, Margit my wife, Thomas and William my sons for all their understanding and for all the tolerance they expressed during my study.
I also greatly appreciate the help of my colleagues from the laboratory, Chelsea Nguyen and
Rhys Carlson. vii | P a g e
List of abbreviations, tables and figures
List of abbreviations
⁰C degree Celsius
A ascorbic acid
ACLM ascorbic-citric-lactic-malic acids
AOAC Association of Official Analytical Chemists
ATCC American Type Culture Collection
ATP adenosine triphosphate
Bl blank bp base pair
C citric acid c-DNA copy- DNA
CAMHB cation adjusted Muller-Hinton broth
CFU colony forming units cm centimetre
CPE cytopathic effect
CO2 carbon dioxide conc. concentration
DNA deoxyribonucleic acid d.f. degrees of freedom
DTT dithiothreitol e.g. exempli gratia
EBI European Bioinformatics Institute
EDTA ethylenediaminetetraacetic acid
EHEC enterohaemorrhagic Escherichia coli viii | P a g e
ETEC enterotoxigenic Escherichia coli
EPS extracellular polymeric substance
ESBL extended spectrum beta lactamase
ESBL+ Kp. extended spectrum beta lactamase positive
Klebsiella pneumoniae
EPA Environmental Protection Agency
ESCMID European Society of Clinical Microbiology and
Infectious Diseases et al. et alia (and others)
FAD flavin adenine dinucleotide
FAO Food and Agricultural Organisation (USA)
FDA Food and Drug Administration (USA) g gram g gravitational force
GDP guanosine diphosphate
GTP guanosine triphosphate
H+ proton h hours
HAV-4 Human adenovirus type 4
HBA horse blood agar
HCl hydrochloric acid
HIV Human Immunodeficiency Virus
HSV-1 Herpes simplex virus type 1
JECFA Joint Expert Committee of Food Additives kg kilogram ix | P a g e
K+ potassium ion
L lactic acid
LD50 50% of the lethal dose log logarithmic
M malic acid
M molarity
Mbp mega base pair
MIC minimum inhibitory concentration min. minutes
MFS major facilitator superfamily ml millilitre mM millimolar
MRSA multi (methicillin) resistant Staphylococcus
aureus
MRAB multi resistant Acinetobacter baumanii
MSDS material safety data sheet
N normal (chemistry)
NA nutrient agar
NAD nicotinamide adenine dinucleotide
NaOH sodium hydroxide
NATA National Association of Testing Authorities
NCTC National Collection of Type Cultures
NDA no data available nm nanometre x | P a g e nM nanomolar ng nanogram dNTPs deoxyribonucleotide triphosphates pH power of the concentration of the hydrogen ion orgs. organism
PCR polymerase chain reaction p probability
PWDS post-weaning diarrhoea syndrome
RNA ribonucleic acid m-RNA messenger-RNA t-RNA transfer-RNA
RPMI media ‘Roswell Park Memorial Institute’ media rpm revolutions per minute
SCV small colony variant
SD standard deviation
SEM scanning electron microscopy
TGA Therapeutic Goods Administration
TKA Time-kill assay
U unit
VRE vancomycin resistant Enterococcus spp.
VIDRL Victorian Infectious Disease Reference
Laboratory
WOA weak organic acid
WOAs weak organic acids
WHO World Health Organisation xi | P a g e
µl microliter
µ average
µg microgram v/v volume/volume w/v weight/volume
+ positive
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List of tables
Table number and title page
Table 1.1 The pKa values of selected weak organic acids at different equilibrium 3
Table 1.2 LD50 of selected weak organic acids (mg/kg) 5
Table 2.1 pH of individual and combination of WOAs 32
Table 3.1 Minimum effective concentration after 10 minutes exposure to WOAs 73 (mean of six determinations, data shown in Figure 3.1) to 2 viruses; Human adenovirus-4 (non-enveloped) and Herpes simplex virus-1 (enveloped).
Table 4.1 Minimum effective concentration after 10 minutes exposure to single 76 and mixed WOAs of an intracellular bacterium, Rickettsia honei
Table 5.1 Minimum effective concentration against yeast and mould after 30 79 minutes exposure to WOAs
Table 6.1 MIC of individual and selected WOAs against 4 antibiotic resistant 84 bacteria, MRSA, VRE, MRAB and ESBL + Kp.
Table 6.2 TKA results (log 10 reduction in numbers within 16 hours in 1 x MIC 87 and within 8 hours in 2 x MIC) of four weak organic acids individually or in combination (ACLM) against four bacteria; MRSA, VRE, MRAB and ESBL + Kp.
Table 6.3 In house suspension test results on planktonic bacteria (MRSA, VRE, 91 MRAB and ESBL + Kp.), yielding 7-8 log10 reduction in bacterial count
Table 6.4 Results of the in house carrier test showing no growth in all assays 93 with 4 antibiotic-resistant bacteria (MRSA, VRE, MRAB and ESBL + Kp.) after 15 +/- 2 min exposure time.
Table 7.1 1 Absorption at 570 nm after treatment of biofilms (containing 4 99 antibiotic-resistant bacteria, MRSA, VRE, MRAB and ESBL + Kp.) with individual WOAs and all four (ACLM) weak organic acids for 3 different exposure periods.
Table 7.2 Viability of antibiotic-resistant bacteria (MRSA, VRE, MRAB and 103 ESBL + Kp.) in biofilms after different exposure times to WOAs. (ACLM= all 4 WOAs), determined by subculturing onto HBA.
Table 8.1 Summary of nucleic acid concentrations obtained from non-treated 138 and acid treated Staphylococcus aureus by Trizol extraction (NanoDrop measurements).
Table 8.2 260: 280 ratios of protein and nucleic acids 139 xiii | P a g e
Table 8.3 Summary of known genes differently expressed (up or down- 151 regulated) in acid shock treated Staphylococcus aureus NCTC 8325 compared to control
Table 9.1 Reduction in bacterial count on ceramic tiles using the 10% (v/v or 164 w/v) combination of WOAs
Table 9.2 The effect of WOAs on an environment contaminated with VRE. 167 Summary of results from Birdsey Wing 6, The Geelong Hospital
Table 9.3 The effect of WOAs on an environment contaminated with VRE. 168 Summary of results from McKellar Centre, Barwon Health.
Table 9.4 The effect of WOAs on an environment contaminated with VRE. 169 Combined summary of results (The Geelong Hospital- McKellar Centre)
Table 9.5 Comparison of VRE positive swabs pre-and post-WOA disinfection 170 (T-test) from different locations within Birdsey Wing 6 (Geelong Hospital) and McKellar Centre.
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List of figures
Figure number and title page
Figure 1.1 Mode of antibacterial action of weak organic acids 13
Figure 2.1 Biofilm formation (24, 48, and 72 hour incubation) using Greiner 51 microtitre plates, following defined incubation times.
Figure 2.2 Differences between positive and negative biofilm formation 52
Figure 2.3 Acid stress test microtitre plate layout- direct (A) and gradual (B) 58 exposure to low pH
Figure 3.1 Minimum effective concentrations of WOAs against two viruses 73
Figure 4.1 Efficacy of WOAs against Rickettsia honei 76
Figure 6.1 TKA- Biocidal effect over time of mixed (ACLM) weak organic 88 acids against multiresistant bacteria (MRSA, VRE, MRAB and ESBL + Kp.) at 1 x MIC of disinfectant.
Figure 6.2 TKA- Biocidal effect over time of the mixed (ACLM) weak organic 88 acids against multiresistant bacteria (MRSA, VRE, MRAB and ESBL + Kp.) at 2 x MIC of disinfectant.
Figure 7.1 Positive (bacteria embedded) and negative (no bacteria embedded) 95 controls for biofilm formation.
Figure 7.2 Effect of individual and mixed WOAs on MRSA biofilm after three 100 different exposure time
Figure 7.3 Effect of individual and mixed WOAs on VRE biofilm after three 100 different exposure time
Figure 7.4 Effect of individual and mixed WOAs on MRAB biofilm after three 101 different exposure time
Figure 7.5 Effect of individual and mixed WOAs on ESBL + Kp. biofilm after 101 three different exposure time
Figure 7.6 VRE Biofilm (untreated) 107
Figure 7.7 ESBL positive Klebsiella pneumoniae biofilm (untreated) 107
Figure 7.8 MRSA biofilm (not treated with WOAs) 108
Figure 7.9 MRSA biofilm treated with 70% ethanol 108
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Figure 7.10 MRSA biofilm treated with 10% mixed WOAs- 10 minutes 109 exposure
Figure 7.11 MRSA biofilm treated with 10% mixed WOAs-30 minutes 109 exposure
Figure 7.12 MRSA biofilm treated with 10% mixed WOAs-60 minutes 110 exposure
Figure 7.13 MRSA biofilm treated with 10% malic acid-30 minutes exposure 110
Figure 7.14 MRSA biofilm treated with 10% malic acid-60 minutes exposure 111
Figure 7.15 MRSA biofilm treated with 10% lactic-acid-30 minutes exposure 111
Figure 7.16 MRSA biofilm treated with 10% lactic acid-60 minutes exposure 112
Figure 7.17 MRSA biofilm treated with 10% citric acid-30 minutes exposure 112
Figure 7.18 MRSA biofilm treated with 10% citric acid- 60 minutes exposure 113
Figure 7.19 VRE biofilm (not treated) 114
Figure 7.20 VRE biofilm treated with 70% ethanol 114
Figure 7.21 VRE biofilm treated with 10% mixed WOAs-10 minutes 115 exposure
Figure 7.22 VRE biofilm treated with 10% mixed WOAs-30 minutes 115 exposure
Figure 7.23 VRE biofilm treated with 10% mixed WOAs-60 minutes 116 exposure
Figure 7.24 VRE biofilm treated with 10% malic acid acid-30 minutes 116 exposure
Figure 7.25 VRE biofilm treated with 10% malic acid-60 minutes exposure 117
Figure 7.26 VRE biofilm treated with 10% lactic acid-30 minutes exposure 117
Figure 7.27 VRE biofilm treated with 10% lactic acid- 60 minutes exposure 118
Figure 7.28 VRE biofilm treated with 10% citric acid-30 minutes exposure 118
Figure 7.29 VRE biofilm treated with 10% citric acid-60 minutes exposure 119
Figure 7.30 ESBL + Kp. biofilm (not treated) 120
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Figure 7.31 ESBL + Kp. biofilm treated with 70% ethanol 120
Figure 7.32 ESBL + Kp. biofilm treated with 10% mixed WOAs-10 minutes 121 exposure
Figure 7.33 ESBL + Kp. biofilm treated with 10% mixed WOAs-30 minutes 121 exposure
Figure 7.34 ESBL + Kp. biofilm treated with 10% mixed WOAs-60 minutes 122 exposure
Figure 7.35 ESBL + Kp. biofilm treated with 10% malic acid-30 minutes 122 exposure
Figure 7.36 ESBL + Kp. biofilm treated with 10% malic acid – 60 minutes 123 exposure
Figure 7.37 ESBL + Kp. 10% biofilm treated with lactic acid-30 minutes 123 exposure
Figure 7.38 ESBL + Kp. biofilm treated with 10% lactic acid-60 minutes 124 exposure
Figure 7.39 ESBL + Kp. biofilm treated with 10% citric acid- 30 minutes 124 exposure
Figure 7.40 ESBL + Kp. biofilm treated with 10% citric acid- 60 minutes 125 exposure
Figure 7.41 MRAB biofilm (not treated) 126
Figure 7.42 MRAB biofilm treated with 70% ethanol 126
Figure 7.43 MRAB biofilm treated with 10% mixed WOAs-10 minutes 127 exposure
Figure 7.44 MRAB biofilm treated with 10% mixed WOAs-30 minutes 127 exposure
Figure 7.45 MRAB biofilm treated with 10 mixed WOAs-60 minutes 128 exposure
Figure 7.46 MRAB biofilm treated with 10% malic acid-30 minutes exposure 128
Figure 7.47 MRAB biofilm treated with 10% malic acid 60 minutes exposure 129
Figure 7.48 MRAB biofilm treated with 10% lactic acid- 30 minutes exposure 129
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Figure 7.49 MRAB biofilm treated with 10% lactic acid- 60 minutes exposure 130
Figure 7.50 MRAB biofilm treated with 10% citric acid -30 minutes exposure 130
Figure 7.51 MRAB biofilm treated with 10% citric acid – 60 minutes 131 exposure
Figure 7.52 Reduction in approximate surface area of the antibiotic resistant 132 bacteria based on size measurements of individual cells by SEM
Figure 8.1 Quantitation of data and Ct value determination for estimation the 144 amount of c-DNA available for fragmentation and labelling.
Figure 8.2 Changes in the concentration of RNA and c-DNA during c-DNA 144 production for microarray assay.
Figure 9.1 Horse blood agar plates inoculated with 100 µl of MRSA 165 suspension, collected from 1 cm2 ceramic tile before and after exposure to WOAs
Figure 9.2 Horse blood agar plates inoculated with 100 µl of MRAB 165 suspension, collected from 1 cm2 ceramic tile before and after exposure to WOAs
Figure 9.3 Heavy growth of VRE from toilet seat on bile aesculin agar 171 containing 6 µg/ml vancomycin (pre-treatment with WOAs)
Figure 9.4 Moderate growth of VRE from floor on bile aesculin agar containing 171 6 µg/ml vancomycin (pre-treated with WOAs)
Figure 10.1 Summary of effective concentration and exposure time required to 181 eliminate microbes
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Abstract
Organic acids occur naturally in plants and in animals and can be produced by microbial fermentation. In plants they play a major role in their defence mechanism, e.g. against viruses. Organic acids are generally weak acids, as they do not dissociate completely in water.
Ascorbic, acetic, citric, formic, lactic, malic, uric acids are members of the weak organic acid
(WOA) family and they have diverse applications in several industries such as food, oil, agriculture, and health.
Several empirical and historical data supports their antimicrobial effect; however, no disinfectant registered to be effective as a hospital grade disinfectant in Australia (at the completion of this project) consists solely of weak organic acid (WOAs). WOAs have been used in combination with other antimicrobial agents such as ethanol or benzalkonium chloride and probably have synergistic effects with these agents.
Antibacterial disinfectants may eliminate planktonic cells, but biofilm penetration is limited by factors such as surface adsorption and absorption by the biofilm matrix thus limiting the availability of the active component of the disinfectant.
The other limiting factor in the application of disinfectants is toxicity. The toxicity of WOAs is relatively low in comparison with the other disinfectants and because of that rinsing off after application is not required.
Studying the efficacy of WOAs against multi antibiotic resistant bacteria can provide useful information about their antimicrobial efficacy. In future, such a disinfectant may be used against pathogenic multi antibiotic resistant bacteria such as methicillin resistant
Staphylococcus aureus (MRSA), vancomycin resistant Enterococcus spp. (VRE), multi antibiotic resistant Acinetobacter baumanii (MRAB), or extended spectrum beta lactamase
xix | P a g e enzyme producing Gram negatives, such as Klebsiella pneumoniae. These bacteria seldom develop total resistance to organic acids but increased acid tolerance has been observed.
This project investigated the efficacy of selected WOAs and their combination against enveloped (Herpes simplex type 1) and non-enveloped (Human adenovirus type 4) viruses, against intracellular bacteria (Rickettsia honei) and concluded that concentrations as low as 6 mg/ml (1.5 mg/ml against intracellular bacteria) can be effective to eliminate viruses or intracellular bacteria.
Antibacterial efficacy against planktonic multi antibiotic resistant bacteria such as multi
(methicillin) resistant Staphylococcus aureus, vancomycin resistant Enterococcus spp., multi resistant Acinetobacter baumanii and extended spectrum beta lactamase positive Klebsiella pneumoniae was studied using a time-kill assay and an assay developed in-house. Results showed that four WOAs in combination (ascorbic, citric, lactic, and malic) at 10% (w/v or v/v) concentration was an effective disinfectant. Studying the efficacy of the above combination against biofilm embedded multi antibiotic resistant bacteria showed that these acids not only penetrate into biofilms, eliminating the bacteria but they are also able to destroy the principal structure of biofilms, the extracellular polymeric substance.
Activity against yeast (Candida albicans) and a fungus (Aspergillus niger) was moderate, and required at least 200 mg/ml.
The development of acid tolerance was also investigated using microarray assay and data obtained were indicative, but not conclusive, of gene expression changes responsible for the production of acid shock proteins.
An in situ study of a disinfectant containing all four WOAs at 10% (w/v or v/v) against VRE at Barwon Health, Geelong, Australia, showed that this formulation could eliminate bacteria from inanimate surfaces in a real life situation.
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Independent testing by an external laboratory confirmed that the proposed disinfectant (10 % mixed WOAs) could be registered as a hospital grade disinfectant and based on the low toxicity of the components it could be a low risk alternative to other commercially available disinfectants, in the future.
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1 Literature review
1.1 Weak organic acids
Weak organic acids are a heterogeneous group of acids and in their chemical structure vary in the number of carboxyl and hydroxyl groups, carbon-carbon double bounds and the presence or absence of an aromatic ring (Hsiao & Siebert, 1999).
The inhibitory effects of these weak acids on bacteria and fungi have been known for centuries and have been used to preserve foods from spoilage. They have also been used as pain-killers and anti-inflammatory agents (acetyl-salicylic acid) (Rainsford, 2007).
Certain flowering plants, especially citrus fruits belonging to the family Rutacaea, are very high in WOAs. These plants include but are not limited to bitter orange, sweet orange, grapefruit, lemon, and lime. The fruits of this group of plants are commonly known as hesperidium and contain different organic acids, including citric, ascorbic and also bioflavonoids (Raven et al., 1986).
Other fruits like grapes or apples contain mainly malic acid and bioflavonoids. The list of plants containing weak organic acids and bioflavonoids is long and these plants occur in diverse geographical habitats. The willow tree (Salix alba) grows in Europe, North and
South-America and South Africa and contains salicylic acid. Lemon and other citrus trees, including the bitter orange (Citrus aurantium) grow mainly in Mediterranean climates while grapes and apples prefer a colder or continental climate (Andrews, 1961).
1.1.1 Historical perspective
Ascorbic acid, also called Vitamin C (L-ascorbic acid, C6H8O6) was first isolated in 1928 by a Hungarian biochemist Albert Szent-Gyorgyi (Szent-Gyorgyi, 1932, 1938). It is a weak organic acid, synthesized by plants and by some mammals, such as apes and guinea pigs, but not by humans (Zetterstrom, 2009).
Citric acid (C6H8O7) is one of the best-known weak organic acids, initially discovered
(empirically) in the 8th century by a Persian alchemist, Jabir Ibn Hayya (Kristiansen et al.,
1999). The acidic nature of lemon and lime juice was well known by medieval scholars and eventually citric acid was isolated by a Swedish chemist, Carl Wilhelm Scheele in the late
18th century (Shetty et al., 2006).
th The initial extraction of malic acid (C4H6O5) by Paul Walden goes back to the end of the 19 century (McMurry, 1988). He studied the conversion of enantiomers and stereoisomers, the basis of optical isomerism. The role of malic acid together with citric acid in the conversion of carbohydrates and protein into water and carbon dioxide was highlighted again at the time of the discovery of aerobic oxidation, the Krebs, (citric acid or tricarboxylic acid) cycle (Berg et al., 2002).
In the early 19th century chemists, working on the isolation of the active ingredient of willow bark discovered salicylic acid (C7H6O3). An extract of willow bark has been used for centuries to alleviate pain. As far back as 1500 BC, Egyptians used it to relieve back pain and around 200 BC, the “father of medicine” Hippocrates used it to relieve both fever and pain.
In 1899, the German biochemist Hoffman “rediscovered” salicylic acid, creating aspirin
(acetylsalicylic acid), which has been used since as a painkiller by inhibiting the enzyme cyclo-oxygenase and reduce inflammation due to its modification of prostaglandin synthesis
(Rainsford, 2007).
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1.1.2 Chemical and physical characteristics
Organic acids are weak acids, based on their low level of ionization in water.
This is illustrated by following chemical reaction:
R-COOH ↔ R-COO- + H+ where H+ is a proton and R-COO- is the dissociated, ionized form of the acid (anion) and R-COOH is the un-dissociated, non-ionized form of the acid.
The ratio of ionized to non-ionized forms of acid molecules in aqueous solution is reflected by their pKa value (‘Ka’ is the acid dissociation constant), which is derived from the
+ - following equation: log10 Ka= log10 (H ) +log10 [(R-COO )/ (R-COOH)] where - log10 Ka = pKa (Houscroft & Sharpe, 2004).
The relationship between pKa and pH can be defined as:
- pKa = pH- log10[(R-COO )/ (R-COOH)] or pH = pKa + log10 [(salt)/ (acid)]
This equation explains why weak organic acids, which are mainly, present in a non- dissociated (acid) form (R-COOH), have a high (positive value) pKa.
The pKa values of weak organic acids used in this study are in order of 3.1 to 6.4 (Table 1.1).
Depending on the ionisable hydrogen atom, acids can be monoprotic (one pKa value), diprotic (two pKa values), or triprotic (three pKa values).
Table 1.1 pKa values of selected weak organic acids at different equilibrium
Weak organic acids pKa
Ascorbic acid 4. 2
Citric acid 3.1, 4.6, 6.4
Lactic acid 3.9
Malic acid 3.4 5.2
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For strong acids, pKa values are close to zero or negative, e.g. the pKa value of hydrochloric acid is -7, in comparison to the pKa value of water, which is 15.4.
Organic acids are classified according to their molecular structure (aromatic, non-aromatic) and on the number of carboxyl groups present. In carbolic acid or phenol, an aromatic
(phenol) ring is attached to a hydroxyl group. Monocarboxylic acids include acetic acid, acrylic acid, ascorbic acid, benzoic acid, formic acid, lactic acid, propionic acid, stearic acid, and salicylic acid. Dicarboxylic acids include glutaric acid, malic acid, malonic acid, oxalic acid, and succinic acid. Citric acid is the only tricarboxylic acid used in this study.
Most of the weak organic acids are a white or yellow crystalline powder at room temperature.
They are water-soluble and have melting points around 150 ºC (Stumm and Morgan 1970).
1.1.3 Toxicity
WOAs are Generally Recognized As Safe (GRAS) by Food and Drug Administration (FDA,
USA) for human use as they are non-toxic or have very low toxicity. Toxicity of citric, malic and lactic acids has been seen only at very high doses. In 1973, the Joint Expert Committee of Food Additives (JECFA) evaluated the safety of citric, malic and lactic acid and their calcium, potassium, sodium and ammonium salts and concluded that there was no significant toxicological hazard posed by these compounds and no limits were set for their acceptable daily intake by the World Health Organisation (WHO, 1973).
The lethal dose 50 (LD50) of some weak organic acids are summarized in Table 1.2.
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Table 1.2 LD50 of weak organic acids (mg/kg body weight)
(source: MSDS from Sigma-Aldrich and from the Australian Pesticides and Veterinary
Medicines Authority websites) (NDA- no data available)
Weak Organic acids Test animals and route of administration
Rat-oral Rat- Mouse-oral Mouse- Mouse- intraperitoneal intravenous subcutaneous
Ascorbic acid 11900 NDA 3367 518 NDA
Citric acid 5400 375 5400 NDA NDA
Lactic acid 3730 NDA NDA NDA 4500
Malic acid 1600 50 1600 NDA NDA
Ascorbic acid demonstrated very low toxicity in mammals, following oral or parenteral administration. Oral feeding showed a toxic effect at 25 mg/kg body weights in rats (Elmore,
2005). Early toxicological studies showed that signs of acute toxicity in rats due to citric acid appeared following ingestion of 11700 mg /kg body weight (Yokotani et al., 1971).
In subacute toxicological examination a 4800 mg/kg citric acid solution was administered to rats for 6 weeks. Sacrificed animals were examined but no specific deleterious effects were detected on any organs (Yokotani et al., 1971).
Mice and rats dosed with a 2500 mg/kg malic acid solution showed some signs of acute toxicity. According to WHO, the lethal oral dose of L-malic acid for rabbits was 5000 mg/kg
(WHO, 1967). Malic acid also causes a moderate skin irritation in rabbits. It is also non- mutagenic, but in higher dose can cause eye irritation (Fiume, 2001).
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Lactic acid is a metabolic intermediate of glucose and glycogen breakdown and occurs naturally in small quantities in the body where it is non-toxic at physiological concentrations.
Skin irritation has been reported with application of a high concentration of lactic acid at 20%
(v/v) solution (Guillot, 1982). Rats given lactic acid at 1300 m g/kg body weight showed signs of acute toxicity including difficulty breathing, and abdominal bloating (Morotomi,
1981).
No data is available on human toxicity of weak organic acids and MSDS data from Sigma-
Aldrich website (http://www.sigmaaldrich.com/australia.html- viewed on the 14th of April,
2012) and from the Australian Pesticides and Veterinary Medicines Authority website
(http://www.apvma.gov.au/use_safely/material_safety.php- viewed on the 14th of April,
2012), showed only mild to moderate toxicity of the above four organic acids resulting in skin, eye irritation or allergy
1.1.4 Biodegradability
During biodegradation, organic matter is converted into carbon dioxide and water, (or other oxidised products under conditions involving anaerobic microbial metabolism) (Berg et al.,
2002). Biodegradation occurs when microbes use the substrate as a carbon source in a metabolic pathway (Diaz, 2008). The rate (speed) of biodegradation can affect the residual effect of a disinfectant and limit its use. In the past, there were no guidelines or regulations for products to be called biodegradable and they were labelled biodegradable without any justification.
The biodegradation of weak or other organic acids takes place if bacteria or fungi can use these acids as a source of carbon for oxidation or fermentation.
Certain Streptomyces strains can use humic acid as a carbon source (Kontchuo & Blondeau,
1992).
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Mycorrhizae, symbiotic soil fungi present in most soils, attach themselves directly onto the roots of most plants and use organic acids from soil as a carbon source (e.g. acetic, citric, malic and lactic acids) (Van Hees et al., 2003).
Micrococcus spp. is able to utilize benzoic and salicylic acid via different metabolic pathways and certain Bacillus spp.and Nocardia species use organic acids as carbon sources (Haribabu
& Kamath, 1984).
Some members of the alpha-subgroup of Proteobacteria, including Chelatobacter or
Chelatococcus, use amino-polycarboxilic acid (e.g. EDTA-ethylenediaminetetraacetic acid) as a carbon source (Bucheli-Witschel & Egli, 2001).
The biodegradation of organic acid mixtures, containing more than one organic acid has not been studied yet in detail. One organic acid (malic acid) can be a source of carbon for one microbe ( Botrytis cinerea), while being toxic for another microbe ( methicillin resistance
Staphylococcus aureus or vancomycin resistant Enterococcus spp.) (Wackett and
Hershberger, 2001).
Based on the biodegradability data, presented above, high level, mixed contamination
(bacteria and fungi) could affect (reduce) the efficacy of WOA based sanitizer.
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1.2 Antiviral effect of weak organic acids
This study was carried out on two viruses, one enveloped, and one non-enveloped virus to evaluate the scale of antiviral efficacy.
Herpes simplex type 1 virus is an enveloped, double stranded DNA virus. It is responsible for diseases such as cold sores or aseptic meningitis (Koelle and Corey, 2008).
Human adenovirus type 4 is a non-enveloped, double stranded DNA virus and responsible for diseases such as conjunctivitis, viral meningitis or encephalitis (Rux and Burnett, 2004).
The inactivation of viruses by weak organic acids is well known and has been used for many years. The mechanism of this inactivation is not fully understood, but is probably due to denaturation of protein (capsid), and is different for enveloped and non-enveloped viruses.
Enveloped viruses with a lipid coat (acquired from the host cell) are more sensitive to disinfectants than non-enveloped viruses (Wood and Payne, 1998). This is probably due to the lack of a target on the surface on naked viruses. In contrast, an enveloped virus with a lipid coat is an excellent target for attack by disinfectant. The antiviral action of WOAs appears to involve changes in the virion structure (protein denaturation) that may result in loss of one of the outer layers of viral protein with an associated loss of virion infectivity, because capsid proteins-such as haemagglutinin in flu virus- participate in the attachment to the host cells.
Earlier in situ studies of WOAs revealed that salicylic acid or benzoic acid had potent virucidal activity against Rhinovirus on the surface of human hands and that activity persisted for 3 hours after application (Turner & Hendley, 2005 and Turner et al., 2010).
Salicylic acid also plays an important role in plant defence responses to virus attack.
The suggested mechanism of antiviral action of salicylic acid is the inhibition of the plant enzymes catalase and ascorbate peroxidise, resulting in elevated levels of peroxide, which would subsequently activate some defence-related genes (Durner & Klessig, 1995).
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Mato et al. (2003) highlighted the importance of the antiviral activity of non-aromatic WOAs
(malic, lactic, and citric) in honey.
Gallic acid, an aromatic organic acid found in tealeaves and oak bark, also has a strong antioxidant effect, scavenging reactive oxygen species and expressing anti-herpetic activity
(Savi et al., 2005).
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1.3 Antibacterial effect of weak organic acids
1.3.1 Overview of bacterial species used in this study
Four multi antibiotic resistant bacteria, two Gram positive and two Gram negative, were used in this study.
Methicillin resistant Staphylococcus aureus (MRSA) is a Gram positive coccus and was first detected in the United Kingdom in 1961. Today in the USA 60% of staphylococcal infections in Intensive Care Units are caused by MRSA (Rice, 2006). But a recent publication (2011) on
CDC website: http://www.cdc.gov/mrsa/statistics/index.html (viewed on the 15th of
February, 2012) shows that “MRSA infections in healthcare settings are declining although the exact burden of the disease caused by MRSA is still largely unknown (Harbath, 2006).
The treatment of surfaces contaminated with MRSA is an important preventive strategy in hospitals and other health care facilities around the world. Vancomycin resistant
Enterococcus spp. (VRE) are Gram positive cocci and were first isolated in 1985. The therapeutic options for serious infections due to VRE remain unclear (Kvirikadze et al. 2006).
CDC recommendation on colonization by VRE is as follows: “people with colonized VRE
(bacteria are present, but have no symptoms of an infection) do not need treatment”, stated on their website: http://www.cdc.gov/HAI/organisms/vre/vre.html (viewed on the 14th of
April, 2012).
Extended spectrum beta-lactamase positive bacteria were first isolated in Germany in 1983.
A large variety of ESBL producing bacteria including Klebsiella spp.(Gram negative bacilli) have been detected, producing different types of plasmid encoded beta-lactamases such as
TEM, SHV, etc. (Paterson et al, 2003).
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Multi resistant Acinetobacter baumanii (MRAB) is a Gram negative bacilli and was first reported in alarming and increasing numbers by the CDC in 2004. The intrinsic resistance of
MRAB to a variety of antibiotics has been well known for decades (Quinn, 1998).
1.3.2 Factors affecting antibacterial activity of weak organic acids
Historically, WOAs have been used for decades as preservatives in food.
Empirical observations show that the use of salicylic or citric acid as a preservative in homemade jams is a common practice and has proven to be very effective at keeping jams free of microbes for many years at room temperature: http://ec.europa.eu/food/fs/sc/sccp/out170_en.pdf- (viewed on the 22nd of October, 2012).
The antibacterial effect of weak organic acids can be influenced by temperature, pH and the presence or absence of organic matter (e.g. protein, serum, etc.). Cherrington et al. (1992) studied the effect of short-chain organic acids, including formic and propionic acid on
Salmonella spp. at different temperatures, using horse blood, milk, yeast extract, and serum as organic matter. They concluded that high temperature (50 º C) and the presence of yeast extract could provide some protection for Salmonella thus increasing the required effective concentration of different organic acid disinfectants. Indirectly, they also suggested that a combination of short-chain organic acids (acetic or butyric) could show a synergistic effect and be more effective than individual butyric or acetic acid.
In vitro studies with Campylobacter spp. showed that organic acids in combination had a pH dependant synergistic effect with an optimum around pH 4.5 (Chaveerarch et al., 2002 and
2003).
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1.3.3 Mechanism of antibacterial activity of weak organic acids
The antibacterial effect of WOAs is based on several mechanisms.
WOAs (proton donors) in undissociated forms can diffuse across bacterial membranes, disrupting the intracellular pH, by increasing intracellular acidity. The intracellular acidity affects the integrity of purines in nucleic acids, leading to denaturation.
The released protons (H+) decrease the intracellular pH, inhibiting essential enzyme functions and forcing the bacterial cell to release additional protons. This leads to the intracellular accumulation of acid anions, e.g. chloride (Warnecke & Gill, 2005). The increased intracellular anion concentration forces potassium ion transport into the cell to maintain electrical balance. To balance the resulting increased osmotic pressure, bacterial cells attempt to increase glutamate transfer from the cell, but the increased osmolality of the cytoplasm eventually leads to cell death (Warnecke & Gill, 2005). This intracellular accumulation of acid anions seems to be an important mechanism of the antibacterial effect of WOAs (Figure 1.1).
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Figure 1.1 Mode of antibacterial action of weak organic acids
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It was also proposed that increased intracellular acidity and increased intracellular anion concentration effects enzymes, thus affecting protein synthesis, amino acids uptake, DNA binding, etc. (Kirkpatrick et al., 2001).
Another antimicrobial mechanism is the reduction of bacterial metabolism due to deprivation of essential substrates for bacterial growth.
1.3.4 Antibacterial activity of products containing weak organic acids
Several fruits, containing WOAs, expressed antibacterial activity. e.g., the antimicrobial effect of berry fruits against Salmonella spp. and Staphylococcus spp. (Puupponen-Pimia et al., 2005).
Papaya has been used as a traditional medicine for centuries, especially for its antihelmintic activity, but it is also bacteriostatic against several enteropathogens. According to Osato et al. (1993), this is due to its high ascorbic, citric, and malic acid content and strong antioxidative properties involving scavenging oxygen, superoxide, hydroxyl radicals.
Seeds from the bakuchiol tree - a native from China- are high in citric and malic acid and have strong antibacterial effect on oral pathogens, such as Streptococcus spp. and
Actinomyces spp. (Katsura et al., 2001).
Oil, extracted from tomato pulp contains different WOAs, such as citric and tartaric. These showed strong antibacterial efficacy against Staphylococcus spp, Streptococcus spp,
Klebsiella spp, and a strong antifungal efficacy against Candida albicans (Vorobe’ev et al.,
1998).
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1.3.5 Activity of weak organic acids against specific bacterial species
Considering the different physical and chemical properties of organic acids, Nakai and
Siebert (2003) tried to predict the bacterial growth inhibition of WOAs, based on their minimum inhibitory concentration (MIC). Predicted MIC showed very good correlation with experimentally determined MIC, validating their model, which can help to predict efficacy of activity against different bacteria (Nakai & Siebert, 2003).
Numerous studies showed the efficacy of WOAs against enterohaemorrhagic Escherichia coli (EHEC). Some of the short-chain fatty acids, including acetic or propionic acids have bactericidal effect on EHEC at pH 5.5 (Shin et al., 2002). The efficacy of lactic and propionic acid against EHEC was temperature dependant, lower temperature reducing efficacy
(McWilliam et al., 2002).
Smulders and Greer (1998) showed that Yersinia and EHEC to a lesser extent expressed high susceptibility towards certain WOAs, including lactic and acetic acid.
They also suggested that organic acids could serve as decontamination agents in abattoirs, as they demonstrated effectiveness against some meat borne pathogens.
Other studies also demonstrated the effectiveness of organic acids against EHEC and
Salmonella in combination with conventional, non-organic disinfectant, such as hydrogen peroxide (Schurman, 2001).
Recent studies on WOAs, including citric, lactic, and malic demonstrated their strong antibacterial efficacy against highly pathogenic foodborne pathogens such as EHEC,
Salmonella typhimurium and Listeria monocytogenes (Sagong et. al., 2011).
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1.3.6 Practical use of weak organic acids
WOAs can sometimes successfully replace antibiotic treatment (Tsiloyiannis et al., 2001).
Piglets with post-weaning diarrhoea syndrome (PWDS), caused by enterotoxigenic
Escherichia coli (ETEC) were studied. Low concentrations (1.0-1.6 % w/v or v/v) of WOAs significantly reduced the incidence of PWDS. Lactic acid was one of the most effective acids at controlling this disease.
Another study on ascorbic, citric, and lactic acid proved that they could be very effective food-preservative in seafood (Rey et.al. 2011). In a review article on WOAs as meat preservatives by Theron and Lues (2007), they concluded that; “because of their many advantages it is, therefore, crucial that the organic acids be revisited as food preservatives, in preventing them becoming obsolete as a result of resistance (acid) development”. This statement also confirms the importance of this current study on the mechanism of the development of acid tolerance.
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1.4 Development of acid tolerance
One of the problems associated with the extensive use of different organic acids such as preservatives in the food industry, is the induction of acid tolerance in bacteria.
Bacteria thrive in diverse environments with fluctuating proton concentrations due to fermentation, digestion etc. On the surface of the teeth, bacteria metabolize carbohydrates anaerobically and create acidic by-products, lowering the pH. Several studies concluded that there was a strong correlation between changes in tolerance to acidity and changes in pathogenicity (Conte et al., 2000 and Merrell and Camilli, 2002). It was observed that acid tolerant pathogens (e.g. Staphylococcus spp. or Listeria spp.) could develop higher acid tolerance and an increase in virulence at the same time (Ricke, 2003). Acid tolerance is a survival mechanism for bacteria, necessary when they exist in the environment dominated by anaerobic respiration (e.g. biofilms) or pass through the stomach acid on their way to the small intestine. At the same time, acid tolerance can be considered to increase pathogenicity, assisting in the survival of pathogens such as Listeria monocytogenes in the intestine (Cotter and Hill, 2003). Several mechanisms participate in acid tolerance including trans-membrane protein structures such as the F1F0-ATPase proton pump or the production of alkaline products by the urease system (Cotter and Hill, 2003). Treatment of bacteria with sub lethal concentration of chloramphenicol (which inhibits protein synthesis) prevented the development of acid tolerance indicating that development of acid tolerance requires protein synthesis (O’Driscoll et al., 1996).
Several proteins are induced during the acid adaptation process (Foster and Moreno, 2007).
When bacteria are first exposed to mild acids, they produce so called pre-shock proteins.
When they are suddenly moved from alkaline to acidic condition they produce acid shock proteins. The majority of acid shock proteins are different from the pre- shock proteins
(Foster, 1991). The inhibition of synthesis of these proteins can result in the loss of
17 | P a g e adaptation to the acidic environment and at the same time may reduce pathogenicity (Foster
1991). Bacteria, including Listeria spp., were more tolerant to acids in the stationary phase of their growth cycle rather than in their exponential phase (O’Driscoll et al., 1996). Very active protein synthesis during the logarithmic or exponential growth phase is consistent with this finding. The development of acid tolerance can be induced by reducing the pH only fractionally or by sudden exposure, which induces acid tolerance and enhances the survival of the bacteria at much lower pH (Svenseter at al., 2007).
Enterohaemorrhagic Escherichia coli and Shigella spp, survived at a pH as low as 2.5 for many hours (Benjamin & Datta, 1995). An increase in acid tolerance is not just a phenomenon of planktonic bacterial cells; it also appears in bacteria associated with biofilms.
McNeill and Hamilton (2003) induced acid tolerance of Streptococcus mutans isolated from dental plaques. Exposure of a S. mutans biofilm to moderate acidity (pH 5.5) enhanced survival during the subsequent exposure to pH 3.5. Welin-Neilands and Svensäter (2007) made the same observation on other Streptococcus spp.
In summary, the problems associated with the use of WOA disinfectants is the development of induced acid tolerance. Induction can result in the synthesis of pre-shock proteins by the acidification tolerance response (Foster, 1991) or production of acid shock proteins in case of a sudden drop in pH.
A limited number of studies analysed gene expression due to acid exposure by microarray assay. Chang et al. (2006) examined the toxicogenomic response of S. aureus to peracetic acid. Their findings showed that as a result of peracetic acid exposure, membrane transport genes regulation was altered, and genes responsible for DNA repair and replication were selectively induced. They also concluded that primary metabolism-related genes were differently repressed and peracetic acid stimulated virulence factor gene expression in S. aureus (Chang et al., 2006).
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1.4.1 Formation of small colony variant Staphylococcus aureus
A specific phenotype, so called small colony variant (SCV) of S. aureus has been described many decades ago, however their role in diseases (Atalla et al. 2011) and their role in the development of chronic infection had not been noticed for many years (Looney, 2000).
The formation of this special variant is probably due to a defective electron transport system
(located in the cell membrane), which is the final process in the oxidative synthesis of ATP.
Small colony variant S. aureus show characteristics different from the non-affected variant, such as pinpoint colonies and slow growth on commercially available culture media (HBA), increased antibiotic resistance (especially against aminoglycosides), loss of haemolytic activity and sometimes reduced pigment formation on culture media such as HBA (von Eiff et al. 2000 and 2006, Melter and Radijevic, 2011). Small colony variants can persist intracellular due to a reduced toxin (alpha cytotoxin) activity or production, thus avoid the direct exposure to the immune system or to extracellular antibiotics (von Eiff et al. 2000 and
2006). Environmental pressure such as exposure to antibiotics can help in the development of such a variant (Melter and Radijevic, 2011). Indirectly it has been proposed that environmental stress can lead to the development of SCV (Clements and Foster 1999). Our experience showed that environmental pressure, such as acid exposure, also could be a contributing factor in the development of SCV forms of S. aureus.
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1.5 Antifungal effects of weak organic acids
Unicellular yeast and a multicellular fungus were used in this study. Aspergillus niger or
“black mould” is an aerobic, non-pathogenic fungus (it can be an opportunistic pathogen in immunocompromised patients) and it has been associated with some toxin production
(Schuster et al. 2002). Candida albicans is opportunistic oral or genital pathogenic, unicellular yeast that has emerged as an important pathogen in the last few decades. It is responsible for high mortality in immunocompromised patients. It is also part of the normal flora in the female genital tract (Miceli et al. 2011).
The effect of WOAs on eukaryotic cells can be seen by their activity on fungi (including yeasts). Fungi have a thick cell wall and the capacity to produce spores, making them more resistant to disinfectants, including organic acids. There are a limited number of publications on the antifungal effect of weak organic acids. Lavermicocca et al. (2003) examined the effect of some organic acids produced by probiotics. Phenyllactic acid, produced by
Lactobacillus plantarum was strongly antifungal against a large variety of fungi, such as
Aspergillus, Fusarium and Penicillium species. In another study, the authors studied the antifungal effect of WOAs produced by Propionibacterium and Lactobacillus using probiotics as a biopreservative (Jonsson & Schnurer, 2005). Succinic, citric and tartaric acid from tomato pulp oil extract showed an antifungal effect against Candida spp. (Vorob’ev et al., 1998). The spoilage effect caused by Saccharomyces cerevisiae could be reduced using benzoic acid. Eukaryotic cells such as yeast cells can survive unfavourable external conditions using a special mechanism called macroautophagy. During macroautophagy, cytosolic material undergoes lysosomal degradation and is recycled into basic molecular building blocks, such as amino acids. Macroautophagy allows cellular physiology to continue in the absence of external resources. Benzoic acid is fungicidal by inhibiting macroautophagy and by blocking this survival mechanism of the yeast cells (Hazan et al., 2004).
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1.6 Biofilm formation
In nature, bacteria exist in two forms, planktonic (free-living) and in biofilm (sessile). The formation of biofilms on surfaces occurs naturally and is influenced by biological, chemical, and physical factors such as genetic capability, bacterial density, availability of nutrients, water activity, temperature, and pressure and oxygen tension. Biofilms can be found everywhere in nature from the ocean floor to the surface of teeth. Antonie van Leeuwenhoek noted the formation of biofilm in the 17th century whilst developing the microscope, but it took three centuries to better understand the process of biofilm development (Donlan and
Costerton, 2002). By the late 1980s it was recognised that the majority of all bacterial biomass exists in biofilms, representing a massive “genetic energy”, which can be manifested in the form of increased microbial pathogenicity or increased resistance against antimicrobial agents, including disinfectants (Davey and Toole, 2000).
Biofilm is an aggregation of bacteria growing on a solid surface, within an extracellular polymeric substance (EPS) immersed in fluids. Bacterial genes regulate this cellular aggregation and formation of biofilm (Beenken et al. 2004 and Resch et al. 2005). Bacteria within biofilms are generally much more resistant to antibiotics and disinfectants than their planktonic counterparts (O’Toole et al., 2000).
Conventional disinfection frequently fails to eliminate pathogenic and non-pathogenic bacteria in biofilms on surfaces. Certain biofilms, e.g. those formed by Bacillus subtilis, remain non-wetting against ethanol or other biocides due to the modification of cohesive and adhesive forces on the surface of the biofilms (Epstein et al., 2010).
The phenotypic appearance of a biofilm is determined by the genotype of the bacterium or bacteria, participating in its formation (Donlan and Costerton, 2002). Generally, the main structural components of biofilms are bacteria, water channels and EPS, which is a combination of extracellular polysaccharides with other organic elements including proteins,
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DNA, and inorganic elements such as minerals (Davey and O’Toole, 2000 and Zhiqiang et al., 2007). EPS may account for 50% to 90% of the total organic carbon of biofilms and is a dynamic structure. This dynamic nature is probably one of the main reasons for increased resistance against liquid wetting and gas penetration into biofilms (Epstein at al., 2010).
Chemical modification, e.g. deacetylation is common during development and maturation of
EPS and plays an important role in the change of the pathogenicity and resistance of the responsible bacteria (Davey and Toole, 2000; Voung et al., 2005).
EPS is normally highly hydrated because it can incorporate large amounts of water by hydrogen bonding into the biofilm structure. There are also some hydrophobic structures present, creating a mixture of a dominant hydrophilic and, to a lesser extent a hydrophobic complex (Sutherland, 2001).
It has been proposed that the transition of an infection from acute to chronic is often associated with the formation of a biofilm, which also corresponds to an increase in microbial pathogenicity and antibiotic resistance (Aparna and Sarita, 2008).
The sequence of the bacterial genome responsible for increased pathogenicity is the so-called pathogenicity islands although not all virulence-associated genes are on pathogenicity island
(Schmidt and Hensel, 2004). The changes in pathogenicity cannot be attributed solely to the structure of biofilms as another phenomenon, called quorum sensing also has to be considered. Quorum sensing, common in bacteria to coordination gene expression and resulting in the coordination of certain behaviour, using specific signalling molecules such as acylated homoserine lactones in Gram-negative bacteria or specific oligopeptides by Gram- positive bacteria (Kong et al., 2006 and Parsek and Greenberg, 2005). Quorum sensing regulation includes virulence, sporulation, or biofilm formation (Manquani et al., 2012). The production of the signalling molecules or autoinducers is a population density dependant gene regulation and the responsible gene sequence could be situated on pathogenicity island (de
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Kievit, 2000). Bacteria inside biofilms form a complex ecological system, which is dynamic with bacterial migration in and out of complexes and with strong integration into the surrounding biosystem (Parsek and Greenberg, 2005).
Increased antimicrobial resistance in bacterial biofilms is distinct from conventional antibiotic resistance associated with planktonic bacteria. In planktonic bacteria, the development of antibiotic resistance is often associated with gene expression, e.g. the production of extended spectrum beta lactamase or uptake of plasmid (transformation), carrying the genes responsible for antibiotic resistance. In biofilms the development of antibiotic resistance is predominantly a physical attribute, mainly due to poor antimicrobial penetration, increased absorption and adsorption via the water channels and the development of a special bacterial cell type inside biofilms, called persister cells (Keren et al., 2004,
Allison et al., 2011). These persister cells neither grow nor die in the presence of antimicrobials and play a major role in the antimicrobial defence together with biofilm formation (Stewart, 2002, Rachna et al., 2009 and Lewis 2008).
The sequence involving biofilm formation differs slightly for different bacteria.
In staphylococcal biofilms, specific surface proteins initiate the biofilm formation, followed by the formation of EPS (Otto, 2008). For vancomycin sensitive or resistant Enterococcus spp. the formation of biofilms is probably an adaptive response and decreases with the acquisition of vancomycin resistance (Ramadhan and Hegedus, 2005).
In Gram-negative bacteria the presence of the extended spectrum beta lactamase (ESBL) increase the formation of biofilms, but this is still not conclusive as some Gram-negative bacteria such as Acinetobacter baumanii can be ESBL positive but form weak biofilms (Rao et al., 2008, and Norouzi et al., 2010)
Penetration, disruption or elimination of biofilm bacteria by disinfectants is a major challenge as it requires higher concentrations of disinfectant or extended exposure time, and sometimes
23 | P a g e it simply does not work at all (Rao et al., 2008). Halogens are very strong oxidizing agents; iodine and chlorine based disinfectants are the most commonly used from this group (Chen and Stewart, 1996). The penetration of chlorine into biofilms raises two issues; the penetration rate and the elimination of embedded bacteria. Some studies indicate that the penetration rate is limited due to the reaction between the biomass and the disinfectant inside the biofilms (Chen and Stewart, 1996; Stewart et al., 2001). For this reason the development of other disinfectants, which may overcome this deficiency is very important. Specifically, the ideal disinfectant would have a high efficacy against planktonic bacteria and be able to penetrate and disrupt all aspects of a biofilm. WOAs, such as citric acid in combination with surfactant have shown in vitro efficacy in disruption of biofilms, created by bacteria, colonized on rabbit sinus mucosa (Tamashiro et al., 2009). Honey ('Medihoney' therapeutic honey and Norwegian Forest Honey), high in ascorbic acid, has proven to be bactericidal for both planktonic and biofilm bacteria, although this may be partly due to the presence of hydrogen peroxide (Merckoll et al., 2009). Several studies indicated that salicylic acid alone or in combination with certain antibiotics can be very effective at reducing biofilm formation by certain bacteria. A low concentration of salicylate together with vancomycin inhibits S. epidermidis biofilm formation on catheters (Polonio et al., 2001). Earlier studies by Farber and Wolff (1993) showed that the treatment of catheters with low concentration of salicylic acid prevented the adherence of E. coli and reduced the chance of biofilm formation.
The formation of biofilm can be studied in many ways in vitro or in vivo; however, the use of microtitre plates in combination with quantitative spectrophotometric method is accepted for in vitro studies (Pitts et al., 2003).
The structure of biofilms can be studied using different microscopical methods such as confocal scanning laser microscopy, transmission or scanning electron microscopy.
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Several publications highlight the importance of contemporary approaches, such as confocal laser microscopical examination of biofilm formation on innate surfaces or inside living structures (Psaltis et al., 2007); however “classical” electron microscopical examinations are still acceptable and in use (Thenmozhi et al., 2011).
The formation of a biofilm follows the process of bacterial attachment, colonization of the surface and growth and dispersion. Formation of biofilms occurs preferentially on rough surfaces and increased surface free energy favours biofilm formation (Subramani et al.,
2009). The inner region of the biofilm is quite anaerobic in comparison with the oxygenated outer region. The metabolic activity and growth rates of bacteria are higher in a planktonic form than inside the biofilm (Bhinu, 2005). Biofilms are composed primarily of bacteria and
EPS with a network of water channels. EPS of different bacteria vary in chemical and physical properties, but it is primarily composed of polysaccharides and minerals and the process of formation is also influenced by the growth medium, the substratum and the bacterial cell surface (Bhinu, 2005). Biofilms are often composed of multiple genera or species of bacteria, which interact and compete metabolically with members of phylogeneticaly unrelated microbial groups (Davey & O’Toole, 2000). If more than one species participates in the development of a biofilm, the so called “house flora” has an effect on the ability of another species, such as Listeria spp., to participate (Carpentier and
Chassaing, 2004).
Formation of biofilm is a natural process and the availability of any solid surface in contact with a bacterial population initiate changes from planktonic to sessile form of the participating bacteria (Aparna and Sarita, 2008).
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1.6.1 Penetration of antimicrobials into biofilms
The penetration of antimicrobials into biofilms is regulated by factors including solubility, absorption and ionization rate, water activity, temperature, pressure and oxygen tension.
Biofilm associated resistance against antimicrobial agents, including disinfectant starts at the attachment phase of biofilm formation and increases as the biofilm ages (Patel, 2005).
The penetration and efficacy of chlorine into biofilms has been studied. Trachoo and
Kunyaboon (2006) showed with Campylobacter jejunii that if the active chlorine concentration was less than 25 ppm, chlorine did not completely prevent the formation of biofilm. Chlorine also can induce virulence genes in Staphylococcus spp.(Chang et al. 2007)
Folsom et al. (2006) showed that Listeria monocytogenes in biofilm survived exposure to 60- ppm of chlorine, and some strains tolerated chlorine concentrations as high as 80 ppm in their biofilm form. Although there is limited knowledge about the direct penetration of WOAs into biofilms, it has been demonstrated that organic acids together with other non-organic acids are capable of reducing biofilm formation e.g. Salmonella spp (Cirkovic et al., 2007). Their result showed that acidic pH reduces the ability of Salmonella spp. to produce biofilm.
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1.7 In situ testing
To evaluate the efficacy of a disinfectant, in situ testing provides the most valuable information and reflects the real performance of the disinfectant.
1.7.1 Rational for in situ testing of disinfectant
Preliminary in vitro and in vivo tests are good indications of antimicrobial efficacy but these tests exclude environmental effects such as reduced efficacy due to organic contamination of surfaces with lipids and proteins, absorption of the disinfectant and biodegradation.
The importance of in situ testing in health care facilities is to find an effective disinfectant which is able to reduce health care associated infections, which are major cause of patient morbidity and mortality (Weber et al., 2010).
Dettenkoffer and Spencer (2007) highlight the importance of environmental decontamination by assessing the ability of the decontamination agent based on prevention of resistance, safety for patients and safety for personnel and the environment.
The authors also emphasized the potential danger in the use of a disinfectant due to its toxicity.
The majority of disinfectants currently in use world-wide e.g. chlorine, have significantly higher toxicity than WOAs. Poisoning due to chlorine gas has been reported several times and chlorine gas (in very high concentration) actually has been used as a chemical weapon in the 1st World War (Ypres, France 1915, Gilchrist and Matz, 1933).
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1.7.2 Rational for reduction of environmental contamination in hospitals
Dettenkoffer and Block (2005) raised the question of whether the use of disinfectants on surfaces rather than cleaning with detergents alone would be sufficient to reduce nosocomial infection. Some studies suggest that the use of disinfectant alone does not necessarily lower infection rate. Cleaning surfaces with detergent prior to disinfection increases the efficacy of the disinfectant (Dettenkoffer et al., 2004).
The importance of thorough cleaning in hospitals is “under increasing scrutiny from both healthcare providers and consumers because the prevalence of serious infections due to multidrug-resistant pathogens has reached alarming levels” (Carling et al. 2008).
The importance of disinfection is not just to eliminate planktonic bacteria, but the elimination of biofilm embedded bacteria which pose a greater risk than planktonic bacteria.
Pathogens including MRSA forming biofilms on surfaces are more difficult to eliminate
(Joshi et al., 2010). The EPS structure absorbs disinfectant and reduces the available, effective concentration of the disinfectant. Penetration of disinfectant via water channels inside biofilms is also limited (see Chapter 1.6.1).
Boyce (2007) highlights the risk of transmission of pathogens by healthcare workers and emphasizes that routine disinfection does not necessarily stop this transmission. There are calls for improved methods to incorporate the introduction of new, currently non-routinely used disinfectants. WOAs could be a viable alternative to current disinfectants. WOA disinfectants can be used as impregnated wipes or as a spray (liquid form) to treat areas not accessible by wipes. Low toxic, weak organic acid disinfectants may be competitive with currently and regularly used other disinfectants.
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1.8 Aims and conclusion
For better understanding of the antimicrobial efficacy of WOAs the aim of this study was to evaluate the antimicrobial efficacy of four selected WOAs (ascorbic, citric, lactic and malic) alone and in combination (all four) against selected enveloped and non-enveloped viruses, an intracellular bacterium, planktonic and biofilm embedded multi antibiotic resistant Gram positive and Gram negative bacteria and example of yeasts and fungi.
Several “escape mechanisms” are available for bacteria to avoid the lethal exposure to disinfectants (including disinfectant based on WOAs) such as biofilm formation, becoming intracellular the development of small colony variants (especially in Staphylococcus spp.), and the development of acid resistance with associated gene expression.
Biofilm formation was studied by scanning electron microscopy, and the development of acid tolerance (resistance) was studied using a commercial microarray assay.
Apart from in vitro studies, in situ studies were carried out on inanimate surfaces to test the efficacy of WOAs in a real world situation in a hospital.
After comparing several concentrations and combinations, it was concluded that an equal combination of ascorbic, citric, lactic and malic acid (recommended in 10% concentration) produced an effective disinfectant against enveloped and non-enveloped viruses, intracellular bacteria and multi antibiotic resistant bacteria, such as methicillin resistant Staphylococcus aureus, vancomycin resistant Enterococcus spp., multi antibiotic resistant Acinetobacter baumanii and extended spectrum beta lactamase positive Klebsiella pneumoniae.
The efficacy against yeast and mould was only moderate.
Independent testing by a Therapeutic Goods Administration (TGA, Australia) registered laboratory (Siliker) also confirmed that this concentration (10%) of the combination passed all tests required for a disinfectant to be registered as a commercial or hospital grade disinfectant (Appendix D).
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2 Materials and Methods
2.1 Microbes
2.1.1 Viruses
Herpes simplex type 1 virus was obtained from the Australian Rickettsial Reference
Laboratory, Geelong, Australia.
Human Adenovirus type 4 was obtained from the Victorian Infectious Disease Reference
Laboratory (VIDRL), Melbourne, Australia.
2.1.2 Intracellular bacteria
Rickettsia honei was obtained from the Australian Rickettsial Reference Laboratory,
Geelong, Australia.
2.1.3 Bacteria
Staphylococcus aureus NCTC 8325 was obtained from the Brien Holden Vision Institute
(Sydney, Australia) and confirmed by colony characteristics on HBA, Gram stain, DNAse and tube coagulase tests using rabbit plasma.
Methicillin resistant Staphylococcus aureus (ATCC 43300), was obtained from the
Australian Collection of Microorganisms, University of Queensland and confirmed by colony characteristics on HBA, Gram stain, DNase and tube coagulase tests using rabbit plasma .
Vancomycin resistant Enterococcus faecium (VRE), extended spectrum beta- lactamase positive Klebsiella pneumonia (ESBL + Kp.) and multi resistant Acinetobacter baumanii
(MRAB) resistant to imipenem, ciprofloxacin, and some aminoglycosides were obtained with full identification and sensitivity certification from the Hunter Area Pathology Service,
(Newcastle, Australia). VRE isolate was confirmed by colony characteristics on HBA, Gram
30 | P a g e stain and disk diffusion sensitivity test. ESBL + Kp. isolate was confirmed by colony characteristics on MacConkey agar, Gram stain and disk diffusion test (Cefotaxime) as recommended by the Clinical and Laboratory Standards Institute (CLSI).
MRAB isolate was confirmed by colony characteristics on MacConkey agar, Gram stain and disk diffusion sensitivity test.
Bacteria were subcultured and stored according to the National Association of Testing
Authorities (NATA) recommendations (NATA Technical Note 14, 1992).
2.1.4 Yeast and fungi
Candida albicans (ATCC 10231) and Aspergillus niger (ATCC 16404) were obtained from the Australian Collection of Microorganisms, University of Queensland
2.2 Disinfectant
The 10% (w/v) working dilutions of ascorbic, citric and malic acids and 10% (v/v) of lactic acid (using 1.176 multiplying factor in preparation as an 85% concentration was supplied) and their subsequent dilutions and mixtures were prepared using sterile distilled water and stored at room temperature. Freshly made WOAs were used for each experiment, because stability of the disinfectant was not part of this study. Organic acids were sourced from the same supplier (Sigma-Aldrich) to avoid any manufacturers’ differences.
The pH of the individual and mixed WOA solutions are detailed in Table 2.1
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Table 2.1 pH of individual and combination of WOAs
Concentration WOA pH
10% w/v Ascorbic (A) 2.0
10% w/v Citric (C) 1.8
10% v/v Lactic (L) 1.9
10% w/v Malic (M) 1.9
10% v/v A-C-L-M 1.9
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2.3 Reagents and microbiological media
Reagent or media Supplier Country of origin
Annealing buffer Qiagen Germany
Antibiotic-Antimycotic Sigma-Aldrich USA
Array holding buffer Qiagen Germany
Ascorbic acid Sigma-Aldrich USA
Bile-aesculin agar with vancomycin Media Preparation Unit Australia
Butterfield diluent Media Preparation Unit Australia
Citric acid Sigma-Aldrich USA
Cation adjusted Muller-Hinton broth Oxoid Australia
Chloroform Sigma-Aldrich USA
Crystal violet ACR Australia
Control oligonucleotide (3 nM) Qiagen Germany dNTPs USB USA
Dnase-I USB USA
Dnase-I buffer 10 X USB USA
DTT Sigma-Aldrich USA
Ethanol Biolab Australia
Foetal calf serum Invitrogen Australia
GeneChip DNA Labelling Reagent Affymetrix USA
Gentamicin Sigma-Aldrich USA
Glutamax Invitrogen USA
HCL Sigma-Aldrich USA
Horse blood agar Media Preparation Unit Australia
Hybridization mix Qiagen Germany
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Isopropyl alcohol Sigma-Aldrich USA
Lactic acid Sigma-Aldrich USA
Luria broth Oxoid Australia
Malic acid Sigma-Aldrich USA
Membrane Wash Solution Promega USA
Muller-Hinton broth Oxoid Australia
NaOH Sigma-Aldrich USA
NaCl Chem-Supply Australia
Nutrient agar Media Preparation Unit Australia
Nutrient broth No.2 Oxoid Australia
Nuclease free H2O Promega USA
Poly-A RNA Control stock Affymetrix USA
Poly-A Control dilution buffer Affymetrix USA
PCR Purification Column Qiagen Germany
RNeasy Mini Purification Kit Qiagen USA
Random Primers Invitrogen USA
Ringer’s solution Oxoid Australia
RPMI-1640 Gibco USA
Sabouraud dextrose agar Media Preparation Unit Australia
Stain cocktail 1 and 2 Qiagen USA
Superase.in Invitrogen USA
SuperScript III/RNase OUT Enzyme Mix Invitrogen USA
Terminal Deoxynucleotidyl Transferase Promega USA
Tween 80 Sigma-Aldrich USA
Trizol Invitrogen USA
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5 X 1st Standard Buffer Affymetrix USA
2 X First-Strand reaction Mix Invitrogen USA
10 X Dnase buffer Qiagen Germany
5 x Reaction Buffer Qiagen Germany
0.5 M EDTA ( Ethylenediaminetetraacetic acid)
Sigma Aldrich USA
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2.4 Antiviral efficacy study methodology
2.4.1 Growth of virus in adherent cell lines
Different adherent cell lines were used, including Vero cells for Herpes simplex type-1 and
A549/88 cell line for Human adenovirus type 4 in 48 well microtitre plates. RPMI-1640 maintenance medium was used for each culture supplemented as follows: 500 ml RPMI-1640 with 5 mM Hepes buffer and 5 ml Glutamax with the addition of 5% foetal calf serum and
5 ml Antibiotic-Antimycotic and 1 ml Gentamicin (50 µg/10 ml).
Each cell type was grown in 25 cm2 cell culture flasks until the cell monolayer was approximately 90% confluent. The growth medium in each flask was discarded and 0.5 ml of a 1:10 dilution of virus in the appropriate maintenance medium was added to each flask.
The flasks were incubated at 37C for 60 minutes with occasional rocking to distribute the virus across the entire monolayer and maximise intracellular uptake. The remaining inoculum medium was discarded and the monolayer rinsed three times with fresh RPMI-1640 medium
(10 ml). 10 ml of the appropriate maintenance medium (RPMI-1640) was added to each flask and incubated at 37C. Cultures were observed daily for signs of virus replication (cytopathic effect, CPE) for a minimum of 5 days. When CPE was complete, or after a minimum incubation period of 5 days, the supernatant was decanted, aliquoted into 1 ml volumes and stored at -90C in cryovial with cryoprotection. This virus-containing supernatant was used for the WOA titration.
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2.4.2 Virus titration (infectivity titer)
10-fold serial dilution of viral suspension was added to confluent growth cell lines. Plates incubated at 37C in 5% CO2 for a minimum of 5 days. All wells were examined daily for
CPE. Viral titre (50% tissue culture infectious dose TCID50) of the original suspension was calculated (highest dilution showing CPE in 50% of the cell cultures).
2.4.3 Disinfectant viral efficacy test
100 µl viral suspension (107 or 108 viral unit per ml) was added to appropriate cell line and incubated overnight in a 37⁰C incubator with 5% CO2. Cell lines were washed three times with 500 µl maintenance media before exposure to WOAs.
Infected cell lines were exposed for 10 minutes to serial dilutions (10-5-2.5-1.25-0.6-0.3-0.15 w/v or v/v) of mixed WOAs. After 10 minutes contact time, the exposure was terminated by washing the cell lines with the maintenance media three times. Exposed cell lines were incubated for up to 14 days at 37 ºC in a 5% CO2 incubator and CPEs were recorded daily
For test validation the following controls were used; positive or viability control containing infected cell lines without exposure to WOAs, negative control: containing uninfected cell lines with maintenance media only and cytotoxicity control, testing the cytotoxicity of the
WOAs in different concentration (10-5-2.5-1.25-0.6-0.3-0.15 v/v or w/v) on adherent cell lines, showing any CPE due to acidity.
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2.5 Disinfectant intracellular bacteria efficacy study methodology
The genus Rickettsia is obligate intracellular bacterium, carried by fleas, ticks, mites, or lice that cause human disease such as Murine typhus, Flinders Island spotted fever, Rocky
Mountain spotted fever, and scrub typhus (Unsworth et al. 2007).
Monolayers of adherent cell lines were established in 96 well microtitre plates, using Vero cell-lines and RPMI-1640 growth media. The RPMI-1640 was supplemented with 25 mM
Hepes buffer, 10% foetal calf serum and 1% L-glutamine.
TCID50 for Rickettsia honei was determined as detailed in Chapter 2.4.2. Established monolayer was exposed to Rickettsia honei (Spotted Fever Group) for 24 hours. Initial bacterial counts for Rickettsia, using 10-fold serial dilutions were up to 107 bacteria/ml.
Infected monolayers in 96 wells microtitre plates were exposed to different concentrations
(2.5-1.2-0.6-0.3-0.15 % v/v or w/v) of single or combinations of WOAs for 10 minutes.
Monolayers were then washed with 200 µl growth media 5 times before incubating with growth media for 14 days.
Post treatment growth was monitored microscopically for cytopathic effect (CPE) for 14 days.
For test validation the following controls were used; Viability (positive) control, Rickettsia honei infected Vero cells not exposed to WOAs. Sterility (negative) control, Vero cell line with growth media only and cytotoxicity control, Vero cell line exposed to different concentration of WOAs but not exposed to Rickettsia.
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2.6 Disinfectant antifungal efficacy study methodology
The in-house assay using sterile 24 wells microtitre plates- was based on challenging the fungi or yeast with different concentrations of the diluted weak organic acid disinfectant, withdrawing a sample after a given time, and detecting viable fungal/yeast cells by culturing the acid-exposed sample in recovery medium (nutrient broth No.2).
Candida albicans was grown on Sabouraud dextrose agar slope at 25⁰ C for 48 hours.
Yeast cells were harvested and cell count was adjusted in sterile water to 2-8 x 105 cells/ml.
Aspergillus niger was grown on Sabouraud dextrose agar slope at 25⁰ C for 5 days and incubated at room temperature for another 5 days to induce spore formation.
Spore formation was checked microscopically in sterile saline (wet preparation).
Yeast and spore suspension was made in sterile water and adjusted to 2-3 x 105 spores/ml.
The concentration of viable yeast cells or spores in the inoculum was determined using 10- fold serial dilutions in quarter strength Ringer’s solution combined with growth control on
Sabouraud agar and spectrophotometric measurement. A standard curve was created by surface spread testing on horse blood agar (HBA), incubation at 25 °C and counting colonies after 2 and 5 days incubation.
600-µl aliquots of the diluted WOAs (30, 20, 10, 5, and 2.5% w/v or v/v) were transferred into the first well and 200 µl of yeast/fungal suspension was added. The final yeast and spore cell concentration was about 5 x 104 to 2 x 105 per ml for yeast cells and 5 x 104 to 7 x 104 per ml for Aspergillus spores. After the contact time, (10, 20, and 30 minutes) 20 µl aliquots were transferred into wells containing 2 ml of recovery broth (nutrient broth No.2).
This approximately 100-fold dilution (20 µl to 2 ml) eliminated the residual effect of the
WOA disinfectant. The microtitre plates containing the recovery broths were incubated at 25°
C for 48 to 72 hours and examined for growth. All tests were run with four replicates.
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For test validation, the following controls were used: sterility control for the disinfectant and the recovery medium, viability test of the yeast and the fungus and a control to test for the efficacy of inactivating the disinfectant at the end of the exposure period.
For a test to be valid the sterility control had to have no growth and, the viability and inactivator efficacy test had to show growth.
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2.7 Disinfectant antibacterial efficacy study methodology- planktonic
bacteria
Four different methodologies, Minimum Inhibitory Concentration (MIC), Time-Kill Assay
(TKA) and in house developed suspension and hard surface carrier assays were used to assess the antibacterial efficacy of the WOAs.
Two methods were used to confirm the findings of this study by an independent, TGA accredited laboratory (Silliker, Australia). One was a semi quantitative suspension test initially described by Kelsey and Maurer in 1966 and the other one was a carrier test
(Therapeutic Goods Administration Suspension and Carrier Test).
2.7.1 Minimum Inhibitory Concentration
This was a semi quantitative method measuring the in vitro activity of antimicrobials against specific bacteria. The methodology of this test involved inoculating a dilution series of an antimicrobial agent with standardized inoculant of selected bacteria, incubating overnight at
37⁰ C and determining the MIC value by detecting the lowest concentration of the antimicrobials visibly inhibiting the growth of the selected bacteria.
Bacterial colonies were grown overnight at 37⁰ C on a non-selective agar (HBA). Several colonies were suspended in 0.9% NaCl and vortexed. Turbidity was adjusted to give a bacterial concentration of 1-2 X 108 CFU/ml as determined spectrophotometrically.
Doubling dilutions of WOA disinfectant in Muller Hinton broth were made in 96 wells microtitre plates. Bacterial suspension in water-Tween 80 diluents was added to each well resulting in the final organism concentration of 2-5 x 105 orgs/ml, and microtitre plates were incubated in at 35 ⁰C for up to 24 hours. Optical density was read in a spectrophotometer
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Controls were used as follows; growth (viability) control and sterility control including the sterility of Muller-Hinton broth and the water-Tween 80 diluents.
The MIC was the lowest concentration of the disinfectant showing complete inhibition of growth
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2.7.2 Time-Kill Assay
The time-kill assay examines the rate at which a specified concentration of an antimicrobial agent eliminates/kills a microbe. This assay can be used to study both concentration and time- dependent bactericidal activity of a biocide.
Prior to the time-kill assay, the determination of MIC value of the disinfectant was necessary
(see chapter 2.7.1). Standard bacterial inocula were added to various dilutions (1 x MIC, and
2 x MIC) of antimicrobials and after specified exposure times of 2,4,8,16 and 24 hours bacterial suspensions exposed to WOAs were sampled to detect viable bacteria. Generally, a
3-log10 CFU/ml reduction in bacterial count indicates an adequate antibacterial effect.
Modified (small volume) version of the assay detailed in the “Procedure Notes” of the
Clinical Microbiology Procedure Handbook (Garcia et al., 2000) was adopted in this work and carried out in 24 wells microtitre plate. Initial bacterial count for MRSA was 5 x 105
CFU/ml; for VRE it was 6 x 105 CFU/ml; for MRAB and ESBL+Kp it was 4 x 105 CFU/ml, all prepared in cation adjusted Muller-Hinton broth (CAMHB). WOAs were prepared in sterile distilled water and 20 µl of bacterial suspension (containing 5 and 6 x 107 CFU/ml of bacteria for MRSA and VRE and 4 x 107 CFU/ml for MRAB and ESBL+Kp) was transferred into 2 ml of WOA, containing either 1 x MIC or 2 x MIC concentration of disinfectant. The zero time samples contained an average of 105 CFU/ml bacteria.
The bacterial suspension- weak organic acid mixture was incubated at 35 ºC and with the removal of 20 µl at 2, 4, 8, 16, and 24 hours later. A 10-fold serial dilution in microtitre plate was performed in 180 µl CAMHB. Serial dilutions were incubated at 35 ⁰C for 24 hours and growth was detected spectrophotometrically to estimate bacterial count. Time-growth
(bacterial count) values (CFU/ml) at 1 x MIC and 2 x MIC of disinfectant were plotted graphically.
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2.7.3 In house efficacy tests
2.7.3.1 In house suspension test
Time-kill assay evaluates the efficacy of a disinfectant in low concentration and against low bacterial count in a time-frame of 24 hours. In comparison the in house test evaluates the disinfectant efficacy at high concentration against high bacterial count and within a time- frame of up to 10 minutes only.
In principal, this methodology follows the dilution rate specified in suspension tests, making this test compatible with tests applied by regulatory authorities.
Tests were carried out in microtitre plates rather than in large volume McCartney bottles (10 ml) which made this test more temperature controlled and reduced the chance of contamination. Sterile 24 well microtitre plates were used for these tests.
The assay was based on challenging different concentrations of the diluted disinfectant with a bacterial inoculum, withdrawing a sample after a given time and detecting viable bacterial cells by culturing the disinfectant exposed bacterial suspension into recovery medium
(nutrient broth No.2).
Test organisms were grown on nutrient agar. The recovery medium, nutrient broth No.2 was freshly prepared a day before testing and checked for sterility. The concentration of viable bacterial cells in the inoculum was determined using 10-fold serial dilutions in quarter strength Ringer’s solution and surface spread testing on HBA, incubated at 37 ºC and colonies counted after 2 days incubation. The concentration of bacteria in the test suspension was adjusted to 109 orgs/ml and 600-µl aliquots of the diluted WOAs (30, 20, 10, 5 and 2.5% w/v) were transferred into the first well and 200 µl of a bacterial suspension was added
(giving about 108 to 109 orgs/ml concentration, considering the dilution factor). Following contact time of 1, 2, 4, 6, 8, 10 minutes, 20 µl aliquots were transferred into wells containing
2 ml of recovery broth.
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Considering the dilution factor (1/100) in sampling the exposed bacterial suspension and transferring into a recovery broth, the reduction rate was calculated as follows: 108-9 orgs/ml reduced to equal or less than 102 orgs/ml.
The microtitre plates containing the recovery broths were incubated at 37 ºC for 48 hours and examined for growth. All tests were run with 4 replicates.
For test validation, the following controls were used; control to test for the sterility of the broth and disinfectant, a control to test for the viability of the testing organism and control to test for the efficacy of inactivating the disinfectant.
For a test to be valid both sterility controls had to be “no growth”, and the viability and inactivator efficacy test had to show “growth”.
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2.7.3.2 In house test using hard surface carriers
The hard surface carrier test is a qualitative test for disinfectant efficacy.
The principle of the tests is that bacterial cells are dried on a surface of glass carriers, exposed to disinfectant for a specified contact time, and assayed quantitatively for the presence of any surviving cells. Controls performed, including bacterial viability, sterility and inactivation efficacy.
The carriers were prepared using glass, cloning cylinders (10 x 10 mm) obtained from the
Danish Agricultural Research Institute (Tjele, Denmark). The carriers were soaked overnight in 1 N NaOH and rinsed several times with tap water and autoclaved and stored at room temperature aseptically until use.
For the preparation of test culture, nutrient broth was inoculated with bacteria (MRSA, VRE,
MRAB and ESBL + Kp.) and incubated at 35 ºC for 24 hours, and initial cell count was determined by 10-fold serial dilution in ¼ strength Ringer solution.
Disinfectant was prepared using 10% (w/v and v/v) mixed WOA disinfectant on the day of testings.
Using 24 wells microtitre plate, 2 ml bacterial culture (containing 108 to 109 orgs/ml) was transferred into each well and using a sterile forceps and one carrier was aseptically transferred into each well. After 15 ± 2 min contact time with bacteria the carriers were removed using sterile forceps, shaken vigorously to remove excess culture, and placed in a vertical position in sterile petri dish matted with 2 layers of Whatman No. 2 filter paper.
Once all of the carriers were transferred, they were incubated at 36 ± 1° C and dried for 40 ±
2 min. Inoculated carriers were used on day of preparation. The cell count on the dried glass carriers were determined by rinsing them into ¼ strength Ringer’s solution and 10-fold serial dilution was carried out.
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After drying, the carriers were deposited into 10% (w/v and v/v) mixed weak organic acid disinfectant, and after 10 minutes exposure time, the carriers were then transferred sequentially into the subculture wells in a 24 well microtitre plate containing 2 ml recovery media (nutrient broth No.2). The microtitre pales were incubated at 35 ± 1° C incubator for
48 ± 2 hours and results were reported as growth or no growth, determined by the presence or absence of turbidity.
20 carriers per organism were tested and tests were repeated three times, and growth in tubes was checked by Gram stain to exclude contamination.
The following controls (carried out in 24 wells microtitre plate) were used in this test:
Positive carriers (viability control) were examined for the presence of the test organism by inoculating into nutrient broth media and plates were incubated for 18–24 h at 35 ± 1° C.
After incubation the plates were examined for turbidity and subcultured onto nutrient agar to check colonial morphology of the tested organism. Growth from nutrient broth was checked by Gram stain.
For neutralization control, dried carriers (2) inoculated with bacteria were placed into wells containing 2 ml of nutrient broth and 0.1% mixed weak organic acids, and incubated for 48
± 2 h at 36 ± 1° C. Positive growth indicated that neutralization of the weak organic acid was not necessary as sublethal concentration did not stop bacterial growth.
For negative control, non-inoculated carriers (2) were placed into nutrient broth (2 ml), incubated for 18–24 h at 36 ± 1° C, and checked for growth (turbidity), and no growth indicated a valid test.
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2.7.4 Therapeutic Goods Administration suspension test
This test was carried out by an external laboratory (Silliker, Melbourne). It uses a semi quantitative suspension test, initially recommended by Kelsey and Maurer (Kelsey and
Maurer, 1974 and Christensen et al. 1982). This test is based on challenging the disinfectant with bacterial inocula, withdrawing a sample after a given time and culturing this sample in a recovery medium. The disinfectant containing the initial bacterial inoculum is then challenged again with a second bacterial inoculum and transferred into a second recovery medium. The disinfectant passes or fails depending on the extent of growth detected in the two cultured recovery medium.
For disinfectants to be classified as commercial or hospital grade have to show efficacy against the following bacteria: Escherichia coli and Staphylococcus aureus for commercial grade disinfectant and Escherichia coli, Staphylococcus aureus, Pseudomonas aeruginosa and Proteus vulgaris for hospital grade disinfectant.
1 ml of bacterial suspension (108-9 orgs/ml) added to 3 ml disinfectant, diluted in standard hard water (dissolve 0.304 g of anhydrous calcium chloride and 0.139 g of magnesium chloride hexahydrate in distilled water and make up to 1 litre, providing water with a hardness of 342 mg/L calculated as calcium carbonate: http://www.who.int/whopes/quality/en/MethodM29.pdf- (viewed on the 21st of October,
2012), and at 8 minutes, 20 µl subcultured into 5 tubes containing recovery broth (nutrient broth). At 10 minutes, the disinfectant solution was inoculated again with a further 1 ml of bacterial suspension, and at 18 minutes, 20 µl subcultured into 5 tubes containing recovery broth (nutrient broth). Tubes were mixed with recovery broth by vortexing and incubated at
37 ⁰C for 48 hours, growth was observed and recorded.
Tests were repeated on all organisms on each of 2 subsequent days (triplicate determination)
To validate the test the following controls were used:
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Sterility test was carried out for the recovery broth and disinfectant, and bacterial viability test, using recovery broth with 10-7 bacterial dilution.
Inactivator efficacy was tested using recovery broth and 20 µl of disinfectant with 10-7 bacterial dilution.
The test to be valid must satisfy the following criteria; the recovery broth control and the disinfectant control should show no growth at 37 ⁰C after 48 hours on incubation, and the viability test and inactivator efficacy controls should show growth at 37 ⁰C after 48 hours incubation.
The disinfectant passes the test if no growth occurs in at least 2 out of the 5 recovery broths on all three repeats for all the tested organisms.
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2.7.4 Therapeutic Goods Administration hard surface carrier test
This test was performed by an external laboratory (Silliker).
In principle, the carrier test is a transient attachment of organisms to surfaces that need to be disinfected. The carriers may be glass, stainless steel, or porcelain depending on the organism under evaluation. Typically 60 replicates are used and after exposure to the disinfectant under conditions of proposed use (clean or dirty), the carriers are transferred to a growth medium with neutraliser and incubated before checking for growth or no growth.
For the carrier test following bacteria were used: Salmonella choleraesuis ATCC 10708,
Pseudomonas aeruginosa ATCC 15442 and Staphylococcus aureus ATCC 6538.
The glass carriers were inoculated with 100 µl inoculum of microorganism, and the disinfectant was applied onto the carriers. Disinfectant was left on the carriers for a specified amount of time. (10 minutes.), and carriers were sampled for recovery.
The samples were neutralized and plated or filtered and transferred onto recovery broth media, and incubated at 37 ⁰C for 48 hours, and colonies were enumerated. The log 10 reductions were determined for treated carriers versus untreated carriers.
The following controls were used to validate the test; positive controls, using untreated carriers and sterility test to exclude recovery broth and disinfectant contamination.
The test must satisfy the followings to be valid; the recovery broth control and the disinfectant control should show no growth at 37 ⁰C after 48 hours on incubation and the positive control must show growth.
The goal for acceptable bactericidal activity was at least a 3 log10 CFU reduction in bacterial count.
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2.8 Disinfectant antibacterial study methodology- biofilm bacteria
2.8.1 Microtitre plate method
Studying the effect of WOAs on biofilms, artificial biofilms were created using multi antibiotic resistant bacteria in microtitre plates. Biofilms were formed using techniques described by O’Toole (1998) and Djordjevic (2002). Modifications to these methodologies were based on the recommendation by the ESCMID Study Group for Biofilms from the
European Conference of Clinical Microbiology and Infectious Diseases in Barcelona, 2008.
Several types of microtitre plates were tested for producing the best biofilm formation.
Greiner (Germany), polystyrene, U-bottom, 96 well was found to establish optimal biofilms for the four tested bacteria, multi resistant Staphylococcus aureus (MRSA), vancomycin resistant Enterococcus spp. (VRE), multi resistant Acinetobacter baumanii (MRAB) and extended spectrum beta lactamase positive Klebsiella pneumoniae (ESBL+ Kp.).
Biofilm formation was monitored by staining sample wells with 1% crystal violet after 24,
48, 72, and 96 hours incubation (Figure 2.1).
Figure 2.1 Biofilm formation (24, 48, and 72 hour incubation) using Greiner microtitre plates, following defined incubation times. Blank (BL) represents negative control.
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After 72 hours incubation biofilm development was complete with no significant changes to the intensity observed after this time. Figure 2.2 shows the intensity of the biofilm formation for the four different multi resistant bacteria, using 1% (w/v) crystal violet staining (labelled
“POS”). Negative control (labelled “NEG”) contained the culture media (RPMI-1640), 5% foetal calf sera, 1% NaCl and 1% ethanol, but no bacteria. For confirming biofilm formation, the positive control (established biofilm) had to be at least three fold more intense (optical density-OD) than the negative control after staining with crystal violet and measured spectrophotometrically (Djordjevic et al., 2002). Single (ascorbic, citric, lactic, malic) and combination of WOAs in 10% (v/v or w/v) concentration, found to be effective on planktonic bacteria (chapter 2.4), were screened for their efficacy on biofilms formed by the multi antibiotic resistant bacteria at three different exposure times (30, 45 and 60 minutes).
Figure 2.2 Differences between positive and negative biofilm formation. Non-specific staining of the wells by crystal violet occurs in the negative controls. (MRSA- methicillin resistant Staphylococcus aureus, VRE- vancomycin resistant Enterococcus spp., MRAB- multi resistant Acinetobacter baumanii, ESBL-Kl. – extended spectrum beta lactamase positive Klebsiella pneumoniae). Positive biofilms contains RPMI-1640, NaCl, ethanol, and bacteria. Negative biofilms contain RPMI-1640, NaCl and ethanol but no bacteria.
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Preliminary work on shorter exposure time (10 and 20 minutes) showed no efficacy, so were excluded from the main study.
Bacteria were cultured and incubated up to 6 hours (log-phase) in nutrient broth in a 35 °C incubator. A 200 µl volume of bacterial suspension was added to 10 ml RPMI-1640 media to provide the working suspension. The working suspension of RPMI-1640 was supplemented with 1% absolute ethanol, 1% NaCl and 5% foetal calf serum. Microtitre plates with 96 wells were sprayed with 70% ethanol prior to use, dried and each well was inoculated with 200 µl of the working suspension. The positive controls used the same working suspension, while negative controls were bacteria free, consisting of the culture media with supplements (NaCl and ethanol) only. Plates were incubated at 30 °C for 72 hours to allow for biofilm formation, and bacterial growth was detected using a spectrophotometer (TECAN, Genios Pro,
Switzerland) at 570 nm and by plating onto horse blood agar. Based on literature (Biotek
Tech Resources, 2008), the 570 nm absorbance is directly proportional to the number of bacterial cells, however bacterial cell density can be monitored at a variety of wavelengths
(e.g. 600 nm), not only at 570 nm.
Biofilm formation was detected by staining wells with 1% crystal violet for 30 minutes and rinsed with distilled water 5 times. A mixture of 80% absolute ethanol, and 20% acetone was used to solubilise the EPS, and biofilm formation was confirmed when the OD of the positive control was at least three times greater than that of the negative control at 570 nm.
Following biofilm formation, the medium from each well was removed and 200 µl of 10%
WOAs added and incubated for 30, 45, and 60 minutes at 30 °C. WOAs were then removed and wells were washed 5 times with distilled water. Biofilms were broken up mechanically using pipette tips, releasing any viable embedded bacteria and RPMI-1640 growth medium was added to detect bacterial growth. Microtitre plates were incubated for 48 hours at 30 °C and the amount of bacterial growth was determined by spectrophotometer at 570 nm.
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In parallel, growth media (100 µl, using surface spread method) was subcultured onto horse blood agar and incubated for 48 hours at 30 °C to confirm the presence or absence of bacteria.
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2.8.2 Scanning Electron Microscopy method
Scanning electron microscopy (SEM) was utilized in this study, although there are some criticisms for the use of this technique, as the preparation requires a dehydration step by ethanol, which could result in the collapse of the extracellular polymeric substance (EPS).
However using appropriate controls, this approach was still a useful tool to study biofilm formation (Davey and O’Toole, 2000).
Aluminium SEM stubs (Deakin University, Australia) were submerged into working suspension of tested bacteria and exposed for 72 hours at 30 °C to allow the formation of a biofilm on the surface of the stubs. Biofilm formation was detected by staining a sample stub with 1% crystal violet (see section 2.8.1). The stubs were transferred into different composition of WOA solutions already found to be most effective by the microtitre plate method and exposed for 10, 30, 45 and 60 minutes. Stubs were then fixed in 70% ethanol for
15 minutes and dried. Biofilms on stubs were also exposed to 70% ethanol for 30 minutes to study the effect of dehydration alone. This was also used as a control as 70% ethanol had been shown previously to penetrate biofilms.
Stubs were gold-coated using EMITECH sputter coater SC7620 (Australia) for 120 seconds.
LEICA S 440 (Germany) and GEMINI-LEO 1530 (Germany) electron microscopes were used in this study.
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2.9 Acid tolerance study methodology (Staphylococcus aureus)
2.9.1 Microtitre plate method
An in-house designed microtitre plate (96 wells) assay was used for the study of acid tolerance. Each well contained 90 µl recovery broths (Luria broth) and the acidic environment was created using WOA solution.
In the study, two types of acid exposure were undertaken, a direct exposure, exposed to low pH in one step and an adaptive, fractional or gradual exposure to gradually lower pH.
The layout of these microtitre plates are shown in Figure 2.3 A and B. Staphylococcus aureus
NCTC 8325 was used in this study (arrays were available for this strain only), grown in minimal essential media (Luria broth) overnight at 35⁰C.
In the direct exposure study, the bacteria- grown overnight in Luria broth- (T) were transferred into the well containing a specific pH (from 2.6 to 3.9) (Figure 2.3 A).
After 10 minutes exposure 10 µl was transferred into 190 µl recovery broths (Luria broth) (B) and incubated overnight at 35⁰C. If growth was detected- by spectrophotometer at 570 nm- bacteria were subcultured onto HBA and into 10 ml Luria broth. Staphylococcus growing in the Luria broth was utilized in the microarray testing protocol and the HBA culture was used to evaluate viability and assess colony morphology.
In the gradual exposure experience, 10 µl bacterial suspension from the test well (T) was transferred into the first well (190 µl Luria broths), representing the highest pH environment and exposed for 10 minutes. After exposure, bacteria were transferred (10 µl) into a well containing 190 µl Luria broth (B) and incubated for 24 hours overnight in a 35⁰C incubator.
After this “recovery” time the exposed bacteria were transferred into the next lowest pH and exposed for a further 10 minutes (Figure 2.3 B).
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The previous steps were repeated several times until growth was detectable in the “recovery” broth (Luria broth). The lowest pH tube containing detectable growth was subcultured in parallel onto HBA and 10 ml Luria broth and incubated overnight in a 35 ⁰C incubator.
Total-RNA extracted from Staphylococcus aureus NCTC 8325 growing in the Luria broth was used in the microarray assay. Bacterial growth on the HBA was used to evaluate purity and examine for colony formation. The negative control (N) was used to exclude contamination of the recovery broth and the positive, growth control was used to check the viability of the bacteria used in the study (Staphylococcus aureus NCTC 8325).
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(A)
1 2 3 4 5 1 2 3 4 5
N G T 3.9 B N G T 3.1 B N G T 3.8 B N G T 3.0 B N G T 3.7 B N G T 2.9 B N G T 3.6 B N G T 2.8 B N G T 3.4 B N G T 2.6 B N G T 3.3 B N G T 3.2 B
(B)
1 2 3 4 5 4 5 4 5 4 5
N G T 3.9 B 3.8 B 3.7 B 3.6 B N G T 3.4 B 3.3 B 3.2 B 3.0 B N G T 2.9 B 2.7 B
N G T 3.9 B 3.8 B 3.7 B 3.6 B N G T 3.4 B 3.3 B 3.2 B 3.0 B N G T 2.9 B 2.7
1- N- negative control
2- G- positive (growth) control
3- T- test wells
4- pH of wells containing WOAs
5- B- recovery broth (Luria broth)
Figure 2.3 Acid stress test microtitre plate layout- direct (A) and gradual (B) exposure to low pH
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2.9.2 Microarray assay
The goal of this study was to obtain and assess the quality and quantity of RNA (extracted from the WOA exposed and non-exposed Staphylococcus aureus NCTC 8325) and the subsequent analysis of synthetized c-DNA by microarray. Several steps were involved in obtaining sufficient amounts of c-DNA (detailed below) for microarray analysis.
A NanoDrop spectrophotometer (Thermo Scientific), PCR thermal cycler (Corbett Research) and Bioanalyzer (Agilent) were used to assess the quality and quantity of total- RNA and c-DNA.
The Agilent 2100 Bioanalyzer is a microfluidics-based platform for the sizing, quantification and quality control of DNA, RNA, proteins, and cells (Lightfoot, 2002).
The Bioanalyzer was used only at AgriBioscience (La Trobe University) but the NanoDrop was used on daily base for the assessment of total-RNA and c-DNA purity.
The real time PCR method was used after fragmentation to assess the quantity of c-DNA available for hybridization.
NanoDrop measures the concentration of macromolecules absorbing light at a specific wavelength.
Nucleotides, RNA, ssDNA (e.g. c-DNA), and dsDNA have absorbance peak at 260 nm wavelength and cumulatively contribute to the total absorbance.
The primary measure for nucleic acid purity versus protein contamination used in this study was the 260/280 absorbance ratio as the maxima for nucleic acids absorbance was at 260 nm wavelength and for proteins was at 280 nm wavelength.
This ratio should be around 1.8 to be considered satisfactory for the purity of DNA, including c-DNA (Glasel, 1995).
Carbohydrates and phenols (from Trizol®) have absorbance at 230 nm wavelength.
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The 260/230 ratio was used as a secondary measurement of nucleic acid purity and it is commonly in the range of 2.
The slope of the absorbance and the amplitude of the peak on the NanoDrop diagram (X- axis) at specific wavelength together with the 260/280 ratio was taken into consideration to assess the quality, purity and quantity (concentration) of nucleic acids (c-DNA) and was confirmed by Bioanalyzer and by real time PCR (Ct value) prior to hybridization.
Appendix A shows examples (graphs) of NanoDrop results from total- RNA extraction to c-DNA purification. These figures reflect each step of the total- RNA isolation and c-DNA synthesis and purification and they are examples of typical results displaying the changes in quality and quantity of the c-DNA, as used for the microarray analysis.
2.9.2.1 Extraction of total-RNA
Total-RNA was extracted using the “Trizol” (Invitrogen, USA) monophasic solution of phenol and guanidine isothiocyanate. The methodology was modified to obtain sufficient total RNA from the staphylococcal suspension. Staphylococcus aureus NCTC 8325 was grown overnight in nutrient broth at 35 ⁰C, and the final bacterial cell count was between
1010 -1011 CFU/ml, determined by 10-fold serial dilution.1 ml of bacterial suspension was centrifuged at 4000 g for 10 minutes to obtain a visible pellet of bacteria, and 1 ml of Trizol reagent was added to the bacterial pellet, vortexed for 10 seconds to obtain a homogeneous suspension. The bacterial suspension were sonicated (Vibra Cell, USA) on ice, three times for
5-10 seconds. The sonicated bacterial suspension was incubated at room temperature (25 ⁰C) for 5 minutes and was placed in a -90⁰C freezer for overnight (18-24 hours). After thawing the sample, 200 µl chloroform was added and vortexed for 5- 10 seconds, incubated at room temperature (25 ⁰C) for 2-3 minutes and centrifuged at 18 000 g for 15 minutes at 2-8⁰C.
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The “aqueous” phase from the top was removed and mixed with 500 µl isopropyl alcohol and incubated at room temperature (25 ⁰C) for 5-10 minutes. The precipitate was centrifuged at
18 000 g for 10 minutes at 2-8 ⁰C and the supernatant removed. The pellet was washed with
1 ml 75% ethanol, vortexed and centrifuged at 11 500 g for 5 minutes. The ethanol was removed and the pellet was dried for 5-10 minutes, completely removing the ethanol.
The pellet obtained, was dissolved in RNase free water (20-30 µl) and incubated at 55 ⁰C for
10 minutes. Total RNA content was measured using a NanoDrop spectrophotometer and the sample was stored at -70⁰C for further work.
2.9.2.2 c-DNA (copy-DNA) synthesis
Poly-A RNA control was prepared using Invitrogen Poly-A RNA control Kit. The 49 Array
Format was used in the assay. 2 µl of the Poly-A RNA Control Stock was added to 38 µl of
Poly-A Control Dilution Buffer (1:20), mixed and spun at 4000 g for 10 second ( stored at -
20°C for up to six weeks and frozen-thawed up to 8 times). The 2 µl of the previous dilution was added to 24 µl of Poly-A Control Dilution Buffer (1:13).
2 µl of the second dilution was used in the c-DNA synthesis. The total-RNA sample was thawed on ice and the following mixture prepared in a 200 µl thin PCR tube on ice; 12 µl of sample RNA and 2 µl of Primer (Random Hexamers) and 2 µl of the second dilution of Poly-
A RNA control and 2 µl of annealing buffer. The above sample was incubated in a thermocycler at 65 °C for 5 minutes and placed on ice immediately afterwards for at least 1 minute. Sample was spun at 18 000 g for 5-10 second and the following mixture on ice was added; 10 µl of 2 X First-Strand reaction Mix to the previous mix and 2 µl of SuperScript
III/RNase OUT Enzyme Mix. Sample was vortexed, spun at 1800 g for 5-10 second and
61 | P a g e incubated in a thermocycler at 25 °C for up to 10 minutes followed by at 50 °C for 50 minutes. The reaction was terminated at 85 °C for 5 minutes and chilled on ice and c-DNA was stored at -20°C
2.9.2.3 RNA degradation for the removal of non-coding RNA
20 µl of 1 N NaOH was added and incubated at 65 °C for 30 minutes, and neutralized with the addition of 20 µl of 1 N HCL.
2.9.2.4 c-DNA purification
For purification, the “Wizard SV Gel and PCR Clean-Up System” (Promega, USA) was used.
Equal volume of membrane binding solution was added to the RNA degraded c-DNA sample. The so called SV minicolumn was inserted into a collection tube and the previous mixture was transferred to the minicolumn and incubated at room temperature (25⁰C) for 1 minute. The minicolum was centrifuged at 16 000 g for 1 minute and 700 µl membrane wash solution was added and centrifuged at 16 000 g for 1 minute. The previous step was repeated with 500 µl membrane wash solution again, and centrifuged at 16000 g for 5 minutes. The
SV Column was re-centrifuged for 1 minute at 16 000 g to allow the evaporation of any residual ethanol. The minicolum was transferred into a 1.5 ml microcentrifuge tube and 30 µl of nuclease free water (eluent) was added, incubated at room temperature for 1 minute and centrifuged at 16 000 g for 1 minute. The purified c-DNA was stored at -20 ⁰C until further work.
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2.9.2.5 Quantitation of c-DNA, Ct value determination
The Ct value was determined using PCR to assess to quantity of available c-DNA for fragmentation and hybridization.
The following primers and probe was used for real-time quantitative PCR:
QNuc-S CCTGAAGCAAGTGCATTTACG
QNuc-AS CTTTAGCCAAGCCTTGACGAACT
QNuc-probe CATCAGCATAAATATACGCTAAGCCACGTCCA
PCR was used to amplify a sequence of the nuc gene, which encodes for the thermostable nuclease of Staphylococcus aureus (Graber et al., 2007 and Brakstad et al., 1992).
2.9.2.6 c-DNA fragmentation
The purpose of the fragmentation was to obtain c-DNA fragments of about 200-300 bp.
These fragment sizes were acceptable for hybridization with the complementary array sequences. Following fragmentation, the fragmented single-stranded DNA can then be labeled for detection of hybridization using terminal deoxynucleotidyl transferase (TdT) in the presence of biotinylated compound.
The following mixture was prepared; 2 µl of 10 X Dnase buffer (Qiagen) and 10 µl of c-
DNA and 0.6 U Dnase I (Qiagen) for each µg of c-DNA. Up to 20 µl of nuclease free water was added, giving the total volume of 20 µl.
Using a thermocycler, the reaction was incubated at 37°C for 10 minutes, followed by the inactivation of Dnase I at 98 °C for 10 minutes, and the fragmented c-DNA (expected to be between 50 to 200 bp range- Qiagen protocol information) was stored at -20°C for later use.
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2.9.2.7 Terminal labelling
Qiagen, GeneChip DNA Labelling Reagent was used to label the 3’ termini of the fragmented c-DNA. The following mixture was prepared; 10 µl of 5 x Reaction Buffer, 2 µl of GeneChip DNA Labelling Reagent, 7.5 m, 2 µl of terminal Deoxynucleotidyl Transferase, up to 20 µl of fragmented c-DNA, and 16 µl of nuclease free water given a total volume of 50
µl. The mixture was incubated at 37°C for 60 minutes and the reaction was stopped by adding 2 µl of 0.5 M EDTA (greater than 90% of the fragments should be labelled-product information by Qiagen).
This product was used with the microarray or stored at -20 °C for later use.
2.9.2.8 Hybridization
The hybridization cocktail was prepared as follows; 75 µl fragmented and labelled c-DNA,
3.3 µl control oligonucleotide (3 nM), and 2 x hybridization mix (containing 2µl LNA dT blocker and 29µl 2x enhanced hybridization buffer) with the addition of nuclease free water to a final volume was 200 µl.
Arrays were inoculated with 100 µl of the above mixture using one septa and “airing” at the other septa. Arrays were placed into Hybridization oven (Affymetrix, Appendix B) at 45 ⁰C for 16 hours and rotated at 60 rpm
2.9.2.9 Washing and staining
Stain reagents were prepared using 600 µl of Stain cocktail 1 into amber microcentrifuge vial,
600 µl of Stain Cocktail 2 into clear microcentrifuge vial and 800 µl of array holding buffer into clear microcentrifuge vial. Arrays and stain cocktails were placed into the appropriate station and stained for 90 minutes (Appendix B).
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2.9.2.10 Data extraction and analysis
Two lots of arrays were analyzed. Firstly, one array examining the extract from non-treated
Staphylococcus aureus NCTC 8325 and one array examining the acid exposed
Staphylococcus aureus NCTC 8325(two genotypes).
The second run was carried out on extracts from two non-treated and two acid treated
Staphylococcus aureus NCTC 8325 (two different genotypes).
This analysis was carried out at AbriBioscience, La Trobe University, Melbourne, Australia.
Data files for analysis were put through, using the Agilent’s Gene Spring expression analysis program. GeneSpring is an expression analysis tool that assists in analyzing genes which are differentially expressed. Data obtained from microarray assay was analyzed by GeneSpring.
The data analysis workflow using GeneSpring as follows:
1. Import and set up experimental data
2. Quality control
3. Filter for differential expression
4. Advanced data analysis
5. Reference of known biology
(Source: Agilent technologies, GeneSpring Manual and Dresen et al., 2003))
The analyses was carried out on genes commonly expressed in the first and second run, and were considered to be differently expressed between the two genotypes at p <= 0.05.
Using the Analysis Centre from the Affymetrix website, Probe Set IDs were searched from the Affymetrix Gene Chip Catalog:
(https://www.affymetrix.com/analysis/netaffx/batch_query.affx?netaffx=netaffx4_annot
(viewed on the 14th of April, 2012)
The level of fold changes in gene expression, resulting in up-or down-regulation was also used in the data analysis.
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2.10 Disinfectant artificial and genuine in situ study
2.10.1 Artificial in situ test in ‘dirty condition’ on tiles
This test was carried out in triplicates on all four multi antibiotic resistant bacteria, using 10%
(w/v and v/v) mixed WOA disinfectant.
Bacterial suspension in Mueller Hinton broth with a bacterial count of >1012 orgs/ml, supplemented with 1% foetal calf serum (dirty condition) was prepared. Inanimate surface
(porcelain tile) exposed for 10 minutes with 1 ml of bacterial solution which allows drying on of bacteria. Surface was swabbed before applying disinfectant (pre-exposure), 10 x 1 cm2 area. Swabs were placed into 10 ml diluent (Butterfield), and 100 µl samples from the
Butterfield diluent was transferred to appropriate solid, non-selective media (HBA).
Parallel with subculturing, a 10-fold serial dilution was made in a microtitre plate, from each swab, to estimate the detected number of organism per cm2 of tile.
After the pre-exposure study, 1 ml WOA disinfectant was added to surface previously inoculated with bacteria and exposed for 10 minutes to disinfectant (10% w/v or v/v mixed
WOAs). A 10 x 1 cm2 surface area was swabbed (post-exposure) and swabs were placed into
10 ml diluent (Butterfield). 100 µl of sample was subcultured from the Butterfield diluent to appropriate solid media (HBA). From each swab a 10-fold serial dilution was also made in a microtitre plate, incubated in a 35⁰C incubator for up to 24 hours to estimate the number of organism per cm2 of tile, post-exposure.
Controls were used as follows; multi antibiotic resistant bacteria exposed surface without disinfectant treatment (positive control) and 70% alcohol cleaned surface without exposure to bacterial solution (negative control).
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2.10.2 Disinfectant genuine in situ test validation against VRE- The
Barwon Health trial
In situ, a Latin word meaning “in position” and the main purpose of an in situ study is to evaluate the performance of a material under investigation in a position where it is going to perform in its everyday use.
The objective of this study was to establish the effectiveness of the 10% mixed WOA disinfectant against VRE in a real life situation, and confirm the validity and practicality of the in vitro data, obtained in the previous experiments.
Sites for the study were selected on the basis of high exposure to VRE.
At The Geelong hospital (Barwon Health, Geelong, Victoria, Australia), an oncology ward
(Birdsey Wing 6) was allocated to the study. The combined en-suite bathroom/ toilet part of a
4 patient room accommodating VRE-colonized patients was used for testing.
At the McKellar Centre (Barwon Health), three rooms were selected, each having accommodated one or twoVRE positive patient(s) for several weeks.
Areas within the bathroom/ toilet , expected to have heavy exposure to VRE were swabbed and treated with the WOA disinfectant on three consecutive mornings.
To obtain a significant number of precleaning VRE positive swabs,sites were swabbed early morning (about 5 am) as the last routine cleaning was normally carried out using Viroclean® at about 6 pm on the night before.
The bile- aesculin agar, used for the detection of VRE , a specifically develped selective culrure media to differentiate Enterococcus spp. from Streptococcus spp. (Oxoid Manual).
Bile stops the growth of Streptococcus spp. and the aesculin will be hydrolysed to glucose and esculetin by Enterococcus spp.,blackening the medium. On the bile-aesculin agar, supplemented with 6 µg/ml vancomycin, VRE appears as a brownish-black colony, blackening the microbiological medium around colonies (Figure 9.3 and 9.4). 67 | P a g e
Only typical colonies were counted and incorporated in the data. The presence of vancomycin in the media excluded the growth of any other Enterococcus spp. that may have been part of the normal faecal flora.
To demonstrate the significance of difference in VRE positivity before and after cleaning,
Paired T-test analysis (Student’s T-test) was used which analyses whether the means (µ) of two groups (control and treatment group) were statistically different.
The control group was the VRE positive swabs collected before the application of the disinfectant and the treatment group was the VRE positive swabs collected after the application of the disinfectant.
Room at Geelong Hospitalwas at Birdsey Wing 6, whichh is an oncology ward.
Three rooms at the McKellar Centre were; South Wing, Room 1, single room, South Wing,
Room 15, double room and Central Wing, Room 32, single room.
400 swabs were collected during this study from bathrooms and toilets; 200 swabs prior to disinfection and 200 swabs post-disinfection with WOAs.
Pre and post-swabbing and cleaning were carried out independently from the researcher and evaluations were cross- checked by an independent scientist.
The following describes the methodology used for the in situ study.
Using preliminary prepared templates, 5 x 5 cm squares were swabbed prior to cleaning and swabs were inserted into 1 ml buffered liquid. Sites were sprayed with 10% (mixed) WOA disinfectant and exposed for 10 minutes. The excess of the disinfectant was removed by gently wiping off with a cloth, and 5 x 5 cm squares were swabbed again and swabs were placed into 1 ml buffered liquid, agitated for a few seconds and poured onto bile-aesculin agar containing 6 µg/ml vancomycin (Media Preparation Unit, Melbourne University,
Melbourne, Australia). Plates were incubated at 37 ºC incubator for 48 hours, and were
68 | P a g e checked and counted (CFU/ml) for brownish-black colonies surrounded by black rings, assumed to be VRE.
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Sites tested at The Geelong Hospital (bathroom/toilet):
1. floor
2. toilet seat
3. toilet cover
4. portable toilet seat
5. hand rail next to toilet
6. hand rail next to shower
7. bathroom tap- hot
8. bathroom tap- cold
9. hand basin
10. door handle
11. toilet flush
12. shower handle
13. white, plastic chair
Sites tested at McKellar Centre (bathroom/toilet):
1. floor
2. toilet seat
3. toilet cover
4. bathroom tap- hot
5. bathroom tap- cold
6. hand basin
7. door handle
8. shower handle
9. toilet flush
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For test validation and control, the test described above was carried out using a commercially available disinfectant, Viraclean (Whiteley Corporation, Australia) (Friedman et. al 2012).
2.10.2.1 Statistical analysis of in situ test
'Student's T-test is one of the most commonly used statistical techniques for testing a hypothesis on the basis of a difference between sample means.
‘Student’ T-test compares two small sets of quantitative data when samples are collected independently of one another.
Calculation of T-test:
where µ0 is the population mean, is the sample mean, ‘s’ is the sample standard deviation of the sample and ‘n’ is the sample size. The degrees of freedom used in this test is n − 1.
The calculation was carried out using an online calculator on the following website: http://www.physics.csbsju.edu/stats/t-test_bulk_form.html, (viewed on the 14th of April,
2012).
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3 Antiviral efficacy of weak organic acid disinfectants
The purpose of this study was to evaluate the efficacy of WOAs against viruses, including an envelope and a non-enveloped. The elimination of non-enveloped or naked viruses is more difficult, as no target ‘surface’ is available for the disinfectant to attack. Efficacy proven against a specific naked virus can be extrapolated to other naked viruses and to enveloped viruses (Steinmann, 2004).
3.1 Results
Antiviral efficacy tests were run in parallel and repeated three times. Data (Table 3.1) represent the mean (µ) of the results obtained and rounded up towards the higher concentration, (e.g. 12 and 12 mg/ml, 12 and 6 mg/ml, and 6 and 12 mg/ml result recorded 12 mg/ml). After 10 minutes exposure the WOAs alone or in combination were effective against both an enveloped Herpes simplex virus type 1(HSV-1) and a non-enveloped Human
Adenovirus type 4 (HAV-4) viruses. No toxicity was observed on the host cell-lines by
WOAs up to 50 mg/ml concentration.
The WOAs were able to eliminate viruses even if they were present at concentrations as high as 107-108 virions/ ml. Individual WOAs were less effective against the non-enveloped
HAV- 4, requiring a concentration of at least 12 mg/ml for virus inactivation (Figure 3.1).
For the elimination of the enveloped HSV-1, a concentration of WOAs as low as 3 mg/ml was sufficient for inactivation. In combination (double, triple or all four) the WOAs were effective at lower concentration, generally around 6 mg/ml or less. Double combinations such as ascorbic-malic, citric-lactic, citric-malic or lactic-malic showed strong efficacies against both viruses in concentration as low as 3 mg/ml.
Triple combinations were also effective, especially ascorbic-lactic-malic and citric-lactic- malic.
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Table 3.1 Minimum effective concentration after 10 minutes exposure to WOAs (mean of six determinations, data shown in Figure 3.1) to 2 viruses; Human adenovirus-4 (non-enveloped) and Herpes simplex virus-1 (enveloped).
HAV-4 HSV-1 (non-enveloped) (enveloped) WOAs %v/v or w/v mg/ml %v/v or w/v mg/ml
A (ascorbic) 2.5 25 0.6 6 C (citric) 1.2 12 0.6 6 L (lactic) 1.2 12 0.3 3 M ( malic) 1.2 12 0.3 3 AC 0.3 3 1.2 12 AL 0.3 3 1.2 12 AM 0.3 3 0.3 3 CL 0.3 3 0.3 3 CM 0.3 3 0.6 6 LM 0.3 3 0.6 6 ACL 0.3 3 1.2 12 ALM 0.3 3 0.6 6 CLM 0.3 3 0.6 6 ACLM 0.6 6 0.6 6
Minumum effective concentration of WOAs against two viruses
30 25 20 15 10
5 Concentration(mg/ml) 0
WOAs
Human adenovirus type 4 HSV-1
Figure 3.1 Minimum effective concentrations of WOAs (A-ascorbic, C-citric, L-lactic- malic and combination) against two viruses (Data tabulated in Table 3.1.
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3.2 Discussion
Generally 3 mg/ml or 6 mg/ml concentration of WOA was sufficient to eliminate both viruses.
The combination of the all four WOAs (ascorbic-citric-lactic-malic, ACLM) destroyed both viruses at 6 mg/ml (triplicate determination). Due to the lack of a target surface, the elimination of non-enveloped viruses by disinfectant is usually more difficult than enveloped viruses (Wood and Payne, 1998), however no obvious differences in efficacy was demonstrated in this study (except with ascorbic acid).
This study demonstrated that WOAs could be very effective agents to eliminate viruses, even if present in very high numbers. The effective concentrations of these acids were very low, significantly below any concentration which could be regarded as toxic.
The demonstrated efficacy against non-enveloped viruses can make WOAs effective disinfection agents against viruses such as Human Rhinovirus (responsible for common cold),
Norovirus, Enterovirus, or Rotavirus, which are common in environments such as kindergartens and nursing homes and responsible for the majority of non-bacterial gastroenteritis (Widdowson, 2005). These findings encourage further studies on other viruses such as HIV, Hepatitis B or C to evaluate specific efficacy of this disinfectant. If efficacy against these viruses could be proven (they are all enveloped viruses) then the use of WOAs as rinsing agents in or on medical devices or as a hand disinfectant, may expand its practical, medical use. To register a disinfectant by TGA (Australia) with virucidal activity, the product must pass the requirements detailed in TGA Guidelines 54; http://www.tga.gov.au/pdf/archive/consult-disinfectants-050731-att2.pdf- (viewed on the
22nd of October, 2012).
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4 Efficacy of weak organic acid disinfectants against intracellular bacteria
The efficacy of WOA disinfectants against intracellular pathogenic bacteria is important especially in veterinary microbiology as several intracellular pathogens cause severe diseases in animals (Wattanaphansak and Singer 2009).
Utilizing a similar methodology (cell culture) as used in the viral studies, this project investigated the efficacy of WOAs against an intracellular pathogen, Rickettsia honei.
4.1 Results
Tests, evaluating the efficacy of WOAs against intracellular bacteria were run in parallel and repeated three times. Data represent the mean (µ) of the results obtained and rounded up towards the higher concentration (Table 4.1).
Rickettsia honei showed high sensitivity towards WOAs (Figure 4.1) and apart from ascorbic acid, individual or combinations of WOAs were effective in concentration as low as 1.5 mg/ml.
Individual WOAs required between 6 and 12 mg/ml concentration to eliminate the intracellular bacteria, but malic acid showed an exceptionally strong efficacy against
Rickettsia honei. Malic acid alone and in combination with citric acid and in the combination of all four proved to be effective in concentration as low as 1.5 mg/ml.
Ascorbic acid alone was the least effective and required concentration as high as 12 mg/ml, but in combination with others, it showed better efficacy.
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Table 4.1 Minimum effective concentration after 10 minutes exposure to single and mixed
WOAs of an intracellular bacterium, Rickettsia honei (Data also shown in Figure 4.1)
Minimum effective WOAs concentration (mg/ml) (µ) A (ascorbic) 12
C (citric) 6 L (lactic) 6 M (malic) 3 AC 3 AL 6
AM 3 CL 6 CM 1.5 LM 3 ACL 6 ALM 6 CLM 6 ACLM 1.5
Minimum effective concentration of WOAs against Rickettsia honei (mg/ml)
14
12
10 conc. 8 mg/ml 6
4
2
0 A C L M AC AL AM CL CM LM ACL ALM CLM ACLM
WOAs
Figure 4.1 Efficacy of WOAs against Rickettsia honei (Data also shown in Table 4.1)
(A-ascorbic, C-citric, L-lactic, M-malic acid and combinations)
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4.2 Discussion
Some of the individual WOA such as malic acid and all double and triple combinations worked very effectively against this intracellular bacterium and the combination of all four
(ascorbic-citric-lactic-malic, ACLM) was even more effective at eliminating Rickettsia honei in very low concentration (as low as 1.5 mg/ml) after 10 minutes exposure.
Other disinfectant with significantly higher toxicity (containing heavy metals, such as
Stalosan , required longer exposure time, up to 30 minutes (Wattanaphansak et al., 2009)
It was demonstrated that disinfectant, containing WOAs can be used efficiently to eliminate intracellular bacteria such as Rickettsia honei and may be a potential disinfectant (further work will be required on other intracellular pathogen) as a carcass wash in abattoirs where other intracellular bacterial exposure such as Anaplasma spp. may occur.
Results on viruses and intracellular bacteria showed that a WOA based sanitizer could be very effective in very low concentration to eliminate these microbes from surfaces.
Obtaining approval for a disinfectant to be used in abattoirs in Australia, the requirements by
PrimeSafe (Australia) have to be satisfied, detailed on their website; http://www.primesafe.vic.gov.au/licensing/meat/abattoir/food-safety-programs (viewed on the 22nd of October, 2012)
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5 Antifungal efficacy of weak organic acids
This study was carried out to demonstrate the efficacy of WOAs, on eukaryotic pathogens such as unicellular yeast and multicellular fungi.
5.1 Results
The antifungal efficacy of the tested WOAs was significantly lower than the antiviral or intracellular bacterial efficacy (Table 5.1).
The antifungal efficacy results show (Table 5.1) that even very high concentrations (over 300 mg/ml) and extended exposure times (30 minutes or over) were in many cases unable to eliminate yeast cells or spores. Exposure time less than 30 minutes (10 and 20 minutes) was insufficient for antifungal efficacy. A 3-log reduction of either of the fungi was difficult to achieve even with high WOA concentration and long exposure times.
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Table 5.1 Minimum effective concentration against yeast and mould after 30 minutes exposure to WOAs
Minimum effective concentration of WOAs. WOAs Candida albicans Aspergillus niger ______mg/ml mg/ml A (ascorbic) >300 >300 C (citric) >300 >300 L (lactic) 300 300 M (malic) >300 >300 AC >300 >300 AL 300 200 AM >300 >300 CL 300 200 CM >300 >300 LM 300 200 ACL 300 200
ALM >300 200 CLM >300 >300 ACLM 300 >300
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5.2 Discussion
The presence of the complex polymers of glucose and the thick chitin layer in the Candida albicans cell wall (Kapteyn et al., 2002) may be one of the reasons why these eukaryotic organisms were poorly eliminated by WOA disinfectants.
The spore formation of fungi (a haploid reproductive structure) produces a very acid- resistant structure that is difficult to eliminate by disinfectants (Trail 2007).
Aspergillus niger spores showed some limited susceptibility towards double combinations of
WOAs, such as ascorbic-lactic, citric-lactic, or lactic-malic and towards triple combinations such as ascorbic-citric-lactic or ascorbic-lactic-malic. The presence of lactic acid in these combinations may play an important role in increasing the antifungal efficacy (Batish et al.,
1997). The reduced efficacy of WOAs against Aspergillus niger may be explained that it can produce citric acid, and has been used for decades for the production of this weak organic acid (Schuster et al. 2002). Based on this finding, citric acid could be removed from any of the above combinations as it is unlikely to be effective.
The efficacy of the tested WOAs against Candida albicans was minimal, as none of the tested combinations expressed any real effect. Based on some limited publications (see
“Literature review”) addition of other WOAs such as benzoic, succinic or phenyllactic acid may have had some effect on increased efficacy, but were not tested in this study.
There is a veterinary ear wash product on the market (Otoclean®) with limited antifungal efficacy. It contains WOAs such as salicylic, lactic and oleic, but the main purpose of this product is in maintaining ear hygiene and prevention of fungal otitis externa. No claim of any antifungal activity was made. The organic acid contents are mainly responsible for dissolving wax and acidifying the external ear canal making it unfavorable for bacterial or fungal colonization or infection. Further work on antifungal efficacy would need to concentrate on other WOAs and their combinations.
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This study on WOAs antifungal effect has showed very limited activity and this may limit their use as a disinfectant to remove fungal contamination from surfaces, such as floors, shower cubicles and any other area exposed to heavy fungal contamination.
To register a disinfectant in Australia to be antifungal, a TGA registration has to be obtained as detailed on the following website;
http://www.tga.gov.au/pdf/disinfectants-evaluation-guidelines.pdf- (viewed on the 23rd of
October, 2012).
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6 Antibacterial efficacy of weak organic acids against planktonic bacteria
The antibacterial effect of WOAs were studied utilizing several different methodologies, including MIC, Time-Kill assay, in house suspension and carrier test and results obtained by these methodologies were confirmed by external laboratory tests. After the confirmation of the efficacy against planktonic bacteria, this project investigated the efficacy of WOAs against biofilm embedded bacteria.
6.1 Minimum inhibitory concentrations of weak organic acids against antibiotic-resistant bacteria
Minimum inhibitory concentration is the lowest concentration of an antimicrobial agent that will inhibit the visible growth of a microorganism after overnight incubation.
MIC is regarded as one of the most basic microbiology laboratory measurements of the activity of an antimicrobial agent. MIC can be determined by agar or broth dilution methods.
Broth dilution method was utilized in this study.
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6.1.1 Results and discussion
The MIC values of the tested WOAs were between 0.7 and 12.5 mg/ml (Table 6.1).
The MIC values for the two Gram negative bacilli (MRAB and ESBL + Kp.) were fractionally higher than for the Gram positive cocci (MRSA and VRE).
ESBL + Kp MIC values were the highest followed by VRE, MRAB and MRSA had the lowest MIC, as low as 0.7 mg/ml for the combination of WOAs. The combination of WOAs had the highest efficacy against the tested bacteria, based on MIC values. Citric, lactic and malic acids showed similar MIC values and ascorbic acid proved to be the least effective, with the highest MIC values. This lower efficacy of ascorbic acid was also demonstrated in results obtained by the in-house method.
The generally low MIC values for individual or mixed WOAs demonstrated their strong antimicrobial efficacy against both Gram positive and Gram negative bacteria and these MIC values were used in the time-kill assay, which is a more specific and sensitive measurement of antimicrobial efficacy than MIC.
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Table 6.1 MIC of individual and selected WOAs against 4 antibiotic resistant bacteria,
MRSA, VRE, MRAB and ESBL + Kp.
MIC (mg/ml)
MRSA VRE MRAB ESBL+Kp
Ascorbic (A) 3.2 6.5 6.5 12.5
Citric (C) 1.5 3.2 1.5 6.5
Lactic (L) 1.5 3.2 1.5 6.5
Malic (M) 1.5 3.2 1.5 6.5
ACLM (combination of all 4) 0.7 1.5 3.2 6.5
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6.2 Time-Kill Assay results of weak organic acids against antibiotic-resistant bacteria
Time-Kill Assay (TKA) is used for examining the rate at which concentrations of an antimicrobial agent kill bacteria. This assay can be used to study both concentration and time- dependent antimicrobial activity.
6.2.1 Results and discussion
In this study TKA- based on MIC results- was used to assess the microbiocidal activity of selected WOAs against multi antibiotic resistant bacteria. WOAs were tested in the TKA at two different concentrations, MIC breakpoint (1 x MIC) and double (2 x MIC) the MIC breakpoint value. TKA gives a good indication of the correlation between of a biocide concentration and its time-dependent antibacterial (bactericidal) activity
The 1 x MIC and the 2 x MIC values with the corresponding log10-reductions, based on the
TKA of the individual and mixed WOAs are summarized in Table 6.2.
TKA using individual WOAs resulted in longer kill times than the combination of all four weak organic acids (Table 6.2). Individual WOAs in 1 x MIC produced 2-5 log10 reductions in bacterial numbers within 4 hours (Figure 6.1). In 2 x MIC the minimum reduction was
4-log10 but in general, 5-log10 reductions in bacterial numbers were achieved within less than
8 hours (Figure 6.2).
TKA results for the combination of WOAs (ACLM) showed 4 to 5-log10 and a 5-log10 reduction in bacterial numbers in 1 x and 2 x MIC respectively against all bacteria. These graphs also indicate the speed at which these reductions occurred. In the 1 x MIC mixed
WOAs achieved a 5-log10 reduction in bacterial numbers within 16 hours for MRSA and
VRE and within 4 hours for MRAB and ESBL + Kp.
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In the 2 x MIC mixed WOAs achieved a 5-log10 reduction in bacterial numbers within 8 hours against VRE, 4 hours against MRSA and MRAB, and 2 hours against ESBL + Kp.
Within a reasonable time frame, it is generally accepted that a 3-log10 reduction in bacterial count is an adequate bactericidal response for a biocide (Ogunmwonyi et al., 2010). All the
TKAs achieved this reduction with the exception of malic acid against MRSA. Although most of the TKAs produced the minimum 3-log10 reduction, greater effect was noticed with the 2 x MIC TKA. The equal mixture of WOAs (10% Ascorbic-Citric-Lactic-Malic) at either
1 x MIC or 2 x MIC TKAs was more effective than individual WOAs.
Although these data reflect in vitro results against the multi antibiotic resistant bacteria, the efficacy of these WOAs in combination, under in situ conditions (on inanimate surfaces against VRE, in chapter 9) support the in vitro results. Inanimate surfaces are one of the major sources of bacteria for outbreaks of nosocomial infections and food contamination leading to food poisoning. Earlier studies indicated that most Gram-negative or Gram- positive bacteria could survive for many months on these surfaces (Neely and Maley, 2000).
The long survival of multi antibiotic resistant bacteria on commonly used surfaces in hospitals, underscores the need for meticulous contact control procedures and careful disinfection to limit the spread of these bacteria (Neely and Maley, 2000).
In conclusion, the study has demonstrated that WOA based disinfectants displayed strong antibacterial activity in the TKA and achieved better than the expected 3-log10 reduction in bacterial count.
Following these observations, in house developed (modified) suspension and carrier tests were used to further assess the activity of the tested WOAs against multi antibiotic resistant bacteria within 8 hours.
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Table 6.2 TKA results (log 10 reduction in numbers within 16 hours in 1 x MIC and within 8 hours in 2 x MIC) of four weak organic acids individually or in combination (ACLM) against four bacteria; MRSA, VRE, MRAB and ESBL + Kp.
TKA was performed at 1 x and 2 x Minimum Inhibitory Concentration
TKA log10 reduction in bacterial numbers ------MRSA VRE MRAB ESBL+Kp.
Ascorbic (A)
MIC (mg/ml) 3.2 6.5 6.5 12.5
1 x MIC 3 5 5 5
2 x MIC 5 5 5 5
------
Citric (C)
MIC (mg/ml) 1.5 3.2 1.5 6.5
1 x MIC 3 4 4 5
2 x MIC 5 4 4 5
------
Lactic (L)
MIC (mg/ml) 1.5 3.2 1.5 6.5
1 x MIC 3 4 3 4
2 x MIC 5 5 5 5
------
Malic (M)
MIC (mg/ml) 1.5 3.2 1.5 6.5
1 x MIC 2 5 4 5
2 x MIC 4 5 5 5
------
ACLM (combination of all 4)
MIC (mg/ml) 0.7 1.5 3.2 6.5
1 x MIC 5 5 4 5
2 x MIC 5 5 5 5
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6
5
1 x MIC
4 MRSA VRE 3 MRAB ESBL+ Kp. 2
Colony Forming Units Forming Colony Units 10) (log 1
0 0 5 10 15 20 25 30 Time (hours)
Figure 6.1 TKA- Biocidal effect over time of mixed (ACLM) weak organic acids against multiresistant bacteria (MRSA, VRE, MRAB and ESBL + Kp.) at 1 x MIC of disinfectant.
6
5
2 x MIC 4 MRSA VRE 3 MRAB ESBL + Kp. 2
1 ColonyForming Units 10) (log
0 0 5 10 15 20 25 30 Time (hours)
Figure 6.2 TKA- Biocidal effect over time of the mixed (ACLM) weak organic acids against multiresistant bacteria (MRSA, VRE, MRAB and ESBL + Kp.) at 2 x MIC of disinfectant.
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6.3 In house suspension test results of weak organic acids against antibiotic-resistant bacteria
The main purpose of the in house test was to determine the effective concentration of the
WOA disinfectant against high concentrations of bacteria (109 orgs/ml) with short exposure times.
This test provided an opportunity to further evaluate the efficacy of different combinations of the WOAs. Results obtained from previous experiments (6.1 and 6.2) and this experiment
(Table 6.3) was used to select the most effective combination of WOAs for further work
(biofilm study).
6.3.1 Results and discussion
A 1/100 dilution (20 µl to 2 ml) of the disinfectant (in nutrient broth No.2) stopped any residual activity of the disinfectant, because of the dilution factor. Ascorbic acid alone showed moderate efficacy against all four multi antibiotic resistant bacteria but required relatively high concentration to eliminate them. Citric acid was more effective against the
Gram negative multi antibiotic resistant bacteria and was exceptionally effective against
ESBL+ Kp. Lactic acid was the most effective among all four individual WOA, especially against the Gram negative bacteria.
Malic acid effectively eliminated Gram negative bacteria, but showed only moderate efficacy against Gram positive bacteria such as MRSA.
Amongst the double combinations (A-C, A-L, A-M, C-L, C-M, and L-M) citric with lactic and malic and lactic were exceptionally effective against MRAB and ESBL pos. Kp.
At the same time they had only moderate efficacy against the Gram positive multi antibiotic resistant bacteria, such as MRSA and VRE.
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The triple combinations (A-C-L, C-L-M, and A-L-M) were also effective in low concentration and short exposure time against the Gram negatives (MRAB and ESBL+Kp.) but less effective against the Gram positive bacteria.
The combination of all four WOAs (A-C-L-M) was the most bactericidal, showing very high efficacy against the two Gram negative bacteria in low concentration and with short exposure time (Table 6.3).
The time and concentration needed to eliminate the Gram positive, multi antibiotic resistant bacteria was as high as 10 % (w/v and v/v) with at least 10 minutes exposure time.
Referring back to Table 6.3, for WOA disinfectant (mixture of all four), to be effective against both Gram positive and Gram negative multi antibiotic resistant bacteria, 10 % (w/v and v/v) concentration was necessary because the higher concentration and longer exposure time was needed to eliminate the Gram positive bacteria in comparison with the more susceptible Gram negative bacteria.
Results obtained by the previous tests were supported by the results from the TGA accredited laboratory (Silliker, Melbourne, Australia), using the TGA approved suspension test, and confirming that the combination of ascorbic-citric-lactic-malic at 10% concentration was sufficiently effective to be classified as a “Hospital Grade Disinfectant”. Tested against
Pseudomonas aeruginosa, Escherichia coli, Proteus vulgaris, and Staphylococcus aureus with an average bacterial count of 108 orgs/ml, the combination of WOA in 10% concentration achieved 8-log10 reduction in triplicate tests.
Having a disinfectant with negligible toxicity (very high LD50, see section 1.1.3) and very strong efficacy (as low as 2.5 % with 1 minutes exposure time-Table 6.3), even against multi antibiotic resistant bacteria, could make this combination a successful replacement for disinfectants with elevated toxicity, especially in environments where toxic residuals must be avoided, such as in kindergartens, nursing homes, and food processing facilities.
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Table 6.3 In house suspension test results on planktonic antibiotic resistant bacteria (MRSA, VRE,
MRAB and ESBL + Kp.) yielding 7-8 log10 reduction in bacterial count.
Conc./time values represent the minimum concentration (%) and the minimum exposure time
(min) necessary to achieve 7-8 log10 reduction in bacterial count.
Methicillin Vancomycin Multiresistant ESBL + Klebsiella resistant resistant Acinetobacter pneumoniae Staphylococcus Enterococcus baumanii aureus (43300) spp.
Conc./time Conc./time Conc./time Conc./time (%/min) (%/min) (%/min) (%/min)
Ascorbic acid 30/10 30/10 20/2 20/6
Citric acid 20/4 20/10 5/8 2.5/8
Lactic acid 20/10 10/8 2.5/1 2.5/1
Malic acid 20/4 10/8 2.5/1 2.5/4
A-C 30/8 25/10 10/4 10/4
A-L 20/4 20/2 2.5/1 10/2
A-M 10/10 10/8 10/4 2.5/6
C-L 20/4 20/2 2.5/4 2.5/4
C-M 25/4 10/8 2.5/6 2.5/8
L-M 20/6 20/2 2.5/1 2.5/1
A-C-L 10/8 20/6 2.5/4 2.5/1
C-L-M 20/4 20/4 2.5/4 2.5/1
A-L-M 20/10 10/10 2.5/1 2.5/4
A-C-L-M 10/10 10/10 2.5/1 2.5/1
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6.4 In house test of weak organic acids against antibiotic-resistant bacteria using hard surface carriers
This test involves carriers, such as glass cylinders submerged in a bacterial suspension, dried and then brought into contact with a disinfectant for a given time (Best et al., 1994). This test has been required by regulatory bodies (e.g. TGA or FDA) to be carried out for a registration of disinfectants. These carriers, with dried on bacteria, are exposed to a disinfectant to assess its efficacy.
6.4.1 Results and discussion
Tests were carried out using the 10% mixed WOAs only, as previous results showed (section
6.3) that this concentration was effective in suspension test. The cell count on the carriers was between 106 to 107 orgs/carrier. Results obtained demonstrated that the 10% concentration of the mixed WOAs was effective at eliminating dried on bacteria (between 106 to 107 orgs/carrier) from carriers and as a result of this it could be considered for registration as a commercial or hospital grade disinfectant (Table 6.4).
The elimination (at least 3-log10 reduction- recommended by TGA) of dried on bacteria from surfaces has two implications for further work. First, this result could be a good indication of the future performance of the disinfectant in situ testing where the majority of the microbes are present in semi-dried form on surfaces. Secondly, it could be an indication to predict the performance of the disinfectant at eliminating biofilm embedded bacteria, although in biofilms there are other factors which need to be considered such as absorption and penetration, not just, on the surface exposure.
Results obtained from this project were confirmed later by a TGA accredited laboratory
(Silliker, Melbourne, Australia), passing the AOAC hard surface carrier test against the
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In summary these in vitro observations (MIC, TKA, in house suspension and carrier test) on the efficacy of WOAs on microbes growing planktonically established the basis for further experiments on the efficacy of weak organic acids on sessile microbes existing inside biofilms.
Table 6.4 Results of the in house carrier test showing no growth in all assays with 4 antibiotic-resistant bacteria (MRSA, VRE, MRAB and ESBL + Kp.) after 15 +/- 2 min exposure time.
Bacteria Test result No. 1 Test result No. 2 Test result No. 3
MRSA No growth of 20 carriers No growth of 20 carriers No growth of 20 carriers
VRE No growth of 20 carriers No growth of 20 carriers No growth of 20 carriers
MRAB No growth of 20 carriers No growth of 20 carriers No growth of 20 carriers
ESBL +Kp. No growth of 20 carriers No growth of 20 carriers No growth of 20 carriers
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7 Antibacterial efficacy of weak organic acid disinfectants against biofilm bacteria
7.1 Introduction
The purpose of this study was to investigate the efficacy of selected WOAs on biofilm formation and elimination of established biofilms produced by multi antibiotic resistant bacteria.
Previous work and results obtained on planktonic, multi antibiotic resistant bacteria (chapter
6) encouraged further investigation on biofilm-embedded bacteria.
To assess the antibacterial efficacy of a disinfectant, it is important to study how disinfectants affect bacteria in both a planktonic and in a biofilm associated form. An effective disinfectant must be able to eliminate bacteria from both environments or at least significantly reduce (4 log10 or higher) the number of viable bacteria present in either of these forms (Bridier et al.,
2011).
This part of the study concentrated on the effect of WOAs on biofilm embedded bacteria.
Biofilms were generated artificially in microtitre plates and the formation was confirmed by scanning electron microscopy, as detailed in chapter 7.3. Please note, that the effect of ascorbic acid, which had limited efficacy on biofilms, was not investigated by SEM.
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7.2 Microtitre plate results and discussion
This methodology was developed and validated in house using recommendation from relevant literature (see chapter 1, section 1.6 and chapter 2, section 2.8).
Biofilm formation in microtitre plates was done in duplicate and repeated three times.
Biofilm formation was confirmed by spectrophotometric measurement calculating the mean
(µ) absorbance at 570 nm for positive and negative controls for all four multi antibiotic resistant bacteria. The mean (µ) of the positive control biofilm was required to be greater than three times the negative control threshold (Figure 7.1).
The three-fold differences requirement between positive and negative control for successful biofilm formation is demonstrated by the following graph (Figure 7.1).
1.6
1.4
1.2
1
Positive 0.8 Negative 0.6
0.4 Absorbance Absorbance 570 at nm
0.2
0 MRSA VRE MR-AB ESBL Kl. Multi antibiotic resistant bacteria
Figure 7.1 Positive controls (biofilm bacteria-MRSA, VRE, MRAB and ESBL + Kp.- in
RPMI-1640 with ethanol, NaCl, and serum) and negative controls (RPMI-1640 with ethanol,
NaCl and serum, but no bacteria) in biofilm formation, represented by optical density at 570 nm after staining with 1% crystal violet. (Data represents means of 6 replicates.)
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The following four graphs (Figure 7.2, 7.3, 7.4, and 7.5) show the effect of individual and mixed WOAs on multi antibiotic resistant bacterial biofilms after three different exposure times (30-45 and 60 min). Spectrophotometric results, detecting bacterial growth (Table 7.1), along with the results of subculture onto HBA (Table 7.2), were used to determine the efficacy of the WOAs on bacterial biofilms.
Absorbance readings to detect bacterial growth in the biofilm formation study, including controls, are summarized in Table 7.1.
The MRSA data (Figure 7.2) shows that normal biofilm formation without exposure to
WOAs gives the absorption reading of about 1.4 for the positive control and the negative control absorption values were around 0.6. Readings of absorption after 45 minutes exposure to citric, lactic and the combination (ACLM) of WOAs were as low as 0.2. These values were equal or lower than the negative absorption values (Figure 7.2). Thirty minutes exposure did not seem to be effective for single WOAs. However the combination of WOAs (ACLM) on biofilms produced very low absorbance reading after 30 minutes exposure. Subculturing (100
µl, surface spread method) the exposed bacteria after 45 or 60 minutes exposure produced no growth on HBA (Table 7.2). For the combination (ACLM) of WOAs a period as short as 30 minutes exposure resulted in loss of viability and no growth on HBA (Table 7.2).
No growth on HBA following subculture confirmed lack of viability of the bacteria, tested.
VRE biofilms were affected by ascorbic, citric, lactic acid and by the combination (ACLM) of WOAs after 30 minutes exposure, but malic acid was only effective after 60 minutes exposure (Figure 7.3).
MRAB biofilms were also rendered sterile by ascorbic, citric, lactic acid and by the combination (ACLM) of WOAs after 30 minutes exposure, but malic acid required 60 minutes exposure (Figure 7.4 and Table 7.2).
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The ESBL+Kp biofilm showed greater resistance against the WOAs and generally 60 minutes exposure was required to produce sterility. Lactic acid and the combination (ACLM) of WOAs proved to be the most effective to eliminate biofilms formed by ESBL+Kp. even after 30 minutes exposure (Figure 7.5). Lactic acid alone appeared to be as effective as the combination was(ACLM).
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MRSA 30 min MRSA 45 min MRSA 60 min Positive cont. 1.40 1.40 1.40 Negative cont. 0.58 0.56 0.57 10% Asc. 0.70 0.23 0.30 10% Citric 0.78 0.25 0.25 10% Lactic 0.71 0.25 0.22 10% Malic 0.90 0.29 0.28 10%ACLM 0.31 0.22 0.25
VRE 30 min VRE 45 min VRE 60 min Positive cont. 1.10 1.12 1.14 Negative cont. 0.50 0.50 0.50 10% Asc. 0.30 0.22 0.25 10% Citric 0.31 0.33 0.29 10% Lactic 0.41 0.30 0.27 10% Malic 0.84 0.72 0.25 10%ACLM 0.30 0.28 0.27
MRAB 30 MRAB 45 min min MRAB 60 min Positive cont. 1.42 1.42 1.42 Negative cont. 0.49 0.50 0.50 10% Asc. 0.45 0.23 0.42 10% Citric 0.49 0.23 0.22 10% Lactic 0.48 0.31 0.29 10% Malic 1.05 0.75 0.31 10%ACLM 0.35 0.37 0.24
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ESBL + Kp ESBL + K 45 30 min min ESBL + Kp 60 min Positive cont. 1.31 1.29 1.30 Negative cont. 0.59 0.58 0.60 10% Asc. 0.75 0.58 0.29 10% Citric 0.82 0.76 0.25 10% Lactic 0.42 0.42 0.28 10% Malic 0.82 0.46 0.22 10%ACLM 0.40 0.41 0.24
Table 7.1 Absorption at 570 nm after treatment of biofilms (containing 4 antibiotic-resistant bacteria, MRSA, VRE, MRAB and ESBL + Kp.) with individual WOAs and all four
(ACLM) weak organic acids for 3 different exposure periods.
(The values represent the mean of 4 determinations)
(These data are shown in Figure 7.2, 7.3, 7.4 and 7.5)
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Effect of individual and mixed WOAs on MRSA biofilm after different exposure time 1.6
1.4
1.2 MRSA 30 min 1 MRSA 45 min 0.8 MRSA 60 min 0.6
0.4 Absorption at 570 nm 570at Absorption 0.2 0
WOAs
Figure 7.2 Effect of individual and mixed (ACLM) WOAs on MRSA biofilm after three different exposure times (ACLM; ascorbic, citric, lactic and malic acid)
Effect of individual and mixed WOAs on VRE biofilm after different exposure time 1.4
1.2
1 VRE30 min 0.8 VRE 45 min 0.6 VRE 60 min 0.4
Absorption at 570 nm 570at Absorption 0.2
0
WOAs
Figure 7.3 Effect of individual and mixed (ACLM) WOAs on VRE biofilm after three different exposure times (ACLM; ascorbic, citric, lactic and malic acid)
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Effect of individual and mixed WOAs on MRAB biofilm after different exposure time
1.6 1.4 1.2 MRAB 30 min 1 MRAB 45 min 0.8 0.6 MRAB 60 min 0.4 Absorption at 570 nm 570at Absorption 0.2 0
WOAs
Figure 7.4 Effect of individual and mixed (ACLM) WOAs on MRAB biofilm after three different exposure times (ACLM; ascorbic, citric, lactic and malic acid)
Effect of individual and mixed WOAs on ESBL + Kp. biofilm after different exposure time
1.6 1.4
1.2 ESBL + Kp.30 min 1 ESBL + Kp.45 min 0.8 ESBL + Kp.60 min 0.6
Absorption at 570 nm 570at Absorption 0.4 0.2 0
WOAs
Figure 7.5 Effect of individual and mixed (ACLM) WOAs on ESBL + Kp. biofilm after three different exposure times(ACLM; ascorbic, citric, lactic and malic acid)
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Positive controls contained bacterial suspension in RPMI-160 supplemented with 5% foetal calf serum, 1% NaCl and 1% ethanol. Negative controls contained RPMI-1640 with foetal calf serum, NaCl, ethanol but no bacteria. Spectrophotometer, capable to measure adsorption of bacterial suspensions-optical density- in flat bottom microtoitre paltes (TECAN) was used.
Absorbance at 570 nm (Y-axis) and the WOAs together with positive and negative controls
(X-axis) with the “positive” on the X-axis being the growth of a normal biofilm without exposure to any WOAs (Fig. 7.2, 7.3, 7.4 and 7.5).
The negative control (which contained RPMI-1640, 1% ethanol, 1% NaCl and 5% foetal calf serum but no bacteria) was also used to detect any contamination of the culture media.
Artificially created biofilms in microtitre plates were satisfactory to study the penetration of
WOAs into biofilms and the elimination of biofilm embedded bacteria.
The controls (Fig.7.1) confirmed the succesful formation of biofilms on the surface of microtitre wells by the four multi-antibiotic resistant bacteria.
Spectrophotometric measurement of bacterial growth after exposure and subculturing of the growth media from each well onto HBA (Fig. 7.3) showed no viable bacteria after 30 minutes exposure to the combination of the four WOAs (ACLM).
Lactic acid alone was also very effective (after 30 minures exposure) with the only exeption of MRSA.
The combination of the four WOAs was effective to eliminate biofilm embedded bacteria and the following SEM study (chapter 7.3) confirmed this efficacy, exploring the structural changes of the exposed biofilms (see more detailed discussion on pages 133-136).
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Table 7.2 Viability of antibiotic-resistant bacteria (MRSA, VRE, MRAB and ESBL + Kp.) in biofilms after different exposure times to WOAs. (ACLM= all 4 WOAs), determined by subculturing onto HBA. Growth: bacterial killing not achieved. No growth: bacterial killing achieved
MRSA VRE MRAB ESBL + Kp.
Positive (growth) control Growth Growth Growth Growth
Negative (no growth) control No growth No growth No growth No growth
10% (w/v) Ascorbic- 30 min. Growth No growth No growth Growth
10% (w/v) Ascorbic- 45 min. No growth No growth No growth Growth
10% (w/v) Ascorbic- 60 min. No growth No growth No growth No growth
10% (w/v) Citric- 30 min. Growth No growth No growth Growth
10% (w/v) Citric- 45 min. No growth No growth No growth Growth
10% (w/v) Citric- 60 min. No growth No growth No growth No growth
10% (v/v) Lactic- 30 min. Growth No growth No growth No growth
10% (v/v) Lactic- 45 min. No growth No growth No growth No growth
10% (v/v) Lactic- 60 min. No growth No growth No growth No growth
10% (w/v) Malic- 30 min. Growth Growth Growth Growth
10% (w/v) Malic- 45 min. No growth Growth Growth No growth
10% (w/v)Malic- 60 min No growth No growth No growth No growth
10% (w/v or v/v) ACLM- 30 min. No growth No growth No growth No growth
10% (w/v or v/v) ACLM- 45 min. No growth No growth No growth No growth
10% (w/v or v/v) ACLM- 60 min. No growth No growth No growth No growth
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7.3 Scanning electron microscopy results and discussion
Figure 7.6 and 7.7 show typical (three dimensional) bacterial biofilm formation on the surface of aluminium stubs after 72 hours incubation for Gram positive cocci and for Gram negative bacilli.
These photos demonstrate the complexity of biofilms formed by bacteria, using organic and non-organic substances as building blocks.
Figure 7.6 and 7.7 show clusters of embedded bacteria (1) are integrated into the EPS (2) which also include water channels (3). Water channels play an essential role in biofilm function, delivering nutrient, and oxygen from outside, removing toxic materials from inside
Water channels also allowing migration of bacteria in and out of the biofilm (Jain et al.,
2007). These structures also increase the surface area of biofilms, which plays an important role in the development of antibiotic resistance due to absorption of antibiotics. This increased absorption of antibiotics and antimicrobials on biofilm surfaces increases the difficulty of treating biofilm related infections with antibiotics or remove them from surfaces.
Non- treated MRSA biofilms showed normal cocci formation about 0.8-1µ in size, together with a well-structured EPS with several water channels (Figure 7.8). The 70% ethanol treated
MRSA biofilm appeared mildly affected, with no obvious sign of dehydration (Figure 7.9).
Ten minutes exposure of MRSA biofilms to mixed WOA showed a moderate effect on biofilm formation or structure (Figure 7.10).
Mixed WOAs (ACLM) at 10% concentration after 30 minutes and 60 minutes exposure produced severely affected MRSA biofilms, including damaged bacterial cells, disappearance of water channels and severe degradation of EPS (Figure 7.11 and Figure 7.12). MRSA biofilms, exposed to malic acid for 30 minutes (Figure 7.13) still maintained a relatively intact ESP structure together with normal coccal morphology, but 60 minutes exposure produced visible damage to the EPS and the bacteria (Figure 7.14).
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The exposure of MRSA biofilms to lactic acid produced very similar results to malic acid exposure, including EPS degradation (Figure 7.15 and Figure 7.16).
MRSA biofilms exposed to citric acid also showed severe effects on the EPS and water channel formation, although relatively normal clusters of cells were still visible (Figure 7.17 and Figure 7.18).
Untreated VRE biofilms were also well structured and ethanol exposure had no serious detrimental effect on structure (Figure 7.19 or Figure 7.20). Ten minutes exposure to mixed
WOAs showed no dramatic changes in the structure of biofilms formed by VRE (Figure
7.21). Mixed (ACLM) WOAs after 30 minutes exposure produced severely affected EPS structure and 60 minutes exposure resulted in damaged bacterial cells (Figure 7.22 and 7.23) which were no longer viable as confirmed by the culture attempts.
Malic, lactic and citric acids had similar effects on VRE biofilms, resulting in destruction of
EPS structure and elimination of water channels (Figure 7.24, 7.25, 7.26, 7.27, 7.28, and
7.29).
ESBL +Kp. biofilms were also well structured after ethanol fixation, with minimal effect on the EPS structure (Figure 7.30 and 7.31). As described for the other bacteria, 10 minutes exposure of ESBL + Kp. biofilms to mixed WOAs showed minimal effect on the biofilm structure (Figure 7.32).
However 10% mixed WOAs caused detrimental effect on ESBL+Kp biofilms (Figure 7.33 and Figure 7.34). Lactic, malic, and citric acids eliminated the normal three-dimensional structure of these biofilms and caused conformational changes on bacterial cells (bacilli changed to cocci shape) (Figure 7.35, 7.36, 7.37, 7.38, 7.39, 7.40).
Untreated or ethanol treated MRAB biofilms maintained their natural, structured appearance, including EPS, water channels and large clusters of bacilli (Figure 7.41 and 7.42).
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It was also observed that 10 minutes exposure to mixed WOAs resulted in no obvious changes in biofilm structure, showing well-structured EPS and the presence of water channels
(Figure 7.43)
Similar to the other biofilms formed by multi antibiotic resistant bacteria, treatment with 10% mixed (ACLM) WOAs for 30 or 60 minutes, resulted in a severely damaged EPS. Although some poorly structured EPS material was still recognisable, but there were no bacteria visible by SEM (Figure 7.44 and 7.45).
Malic, lactic and citric acid treatment of the MRAB biofilms showed partial destruction and partial conformational changes in the shape of bacteria (cocci rather than bacilli (Figure 7.46,
7.47, 7.48, 7.49, 7.50 and 7.51).
In summary, the effect of WOAs alone or in combination on biofilm structures was limited after 10 minutes exposure but significant after 30 minutes exposure, resulting in severe structural changes of the EPS, the disappearance of water channels and conformational changes in the size and shape of the participating bacteria. These results indicate that WOA based disinfectants have significant effect on bacterial biofilms and disinfectant, containing
WOAs could be a very effective to eliminate both planktonic and biofilm embedded bacteria from surfaces, significantly extending the scope of practical use of this antimicrobial product.
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2
1
3
(SEM photo by Zoltan Nack, 7th of July 2009) Figure 7.6 VRE Biofilm (untreated) (1-bacteria, 2-EPS, and 3-water channels)
3
1
2
(SEM photo by Zoltan Nack, 7th of July 2009) Figure 7.7 ESBL positive Klebsiella pneumoniae biofilm (untreated) (1-bacteria, 2-EPS, and
3-water channels)
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Figure 7.8 MRSA biofilm (not treated with WOAs)
Figure 7.9 MRSA biofilm treated with 70% ethanol
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Figure 7.10 MRSA biofilm treated with 10% mixed WOAs- 10 minutes exposure
Figure 7.11 MRSA biofilm treated with 10% mixed WOAs-30 minutes exposure
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Figure 7.12 MRSA biofilm treated with 10% mixed WOAs-60 minutes exposure
Figure 7.13 MRSA biofilm treated with 10% malic acid-30 minutes exposure
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Figure 7.14 MRSA biofilm treated with 10% malic acid-60 minutes exposure
Figure 7.15 MRSA biofilm treated with 10% lactic-acid-30 minutes exposure
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Figure 7.16 MRSA biofilm treated with 10% lactic acid-60 minutes exposure
Figure 7.17 MRSA biofilm treated with 10% citric acid-30 minutes exposure
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Figure 7.18 MRSA biofilm treated with 10% citric acid- 60 minutes exposure
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Figure 7.19 VRE biofilm (not treated)
Figure 7.20 VRE biofilm treated with 70% ethanol
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Figure 7.21 VRE biofilm treated with 10% mixed WOAs-10 minutes exposure
Figure 7.22 VRE biofilm treated with 10% mixed WOAs-30 minutes exposure
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Figure 7.23 VRE biofilm treated with 10% mixed WOAs-60 minutes exposure
Figure 7.24 VRE biofilm treated with 10% malic acid acid-30 minutes exposure
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Figure 7.25 VRE biofilm treated with 10% malic acid-60 minutes exposure
Figure 7.26 VRE biofilm treated with 10% lactic acid-30 minutes exposure
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Figure 7.27 VRE biofilm treated with 10% lactic acid- 60 minutes exposure
Figure 7.28 VRE biofilm treated with 10% citric acid-30 minutes exposure
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Figure 7.29 VRE biofilm treated with 10% citric acid-60 minutes exposure
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Figure 7.30 ESBL + Kp. biofilm (not treated)
Figure 7.31 ESBL + Kp. biofilm treated with 70% ethanol
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Figure 7.32 ESBL + Kp. biofilm treated with 10% mixed WOAs-10 minutes exposure
Figure 7.33 ESBL + Kp. biofilm treated with 10% mixed WOAs-30 minutes exposure
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Figure 7.34 ESBL + Kp. biofilm treated with 10% mixed WOAs-60 minutes exposure
Figure 7.35 ESBL + Kp. biofilm treated with 10% malic acid-30 minutes exposure
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Figure 7.36 ESBL + Kp. biofilm treated with 10% malic acid – 60 minutes exposure
Figure 7.37 ESBL + Kp. 10% biofilm treated with lactic acid-30 minutes exposure
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Figure 7.38 ESBL + Kp. biofilm treated with 10% lactic acid-60 minutes exposure
Figure 7.39 ESBL + Kp. biofilm treated with 10% citric acid- 30 minutes exposure
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Figure 7.40 ESBL + Kp. biofilm treated with 10% citric acid- 60 minutes exposure
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Figure 7.41 MRAB biofilm (not treated)
Figure 7.42 MRAB biofilm treated with 70% ethanol
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Figure 7.43 MRAB biofilm treated with 10% mixed WOAs-10 minutes exposure
Figure 7.44 MRAB biofilm treated with 10% mixed WOAs-30 minutes exposure
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Figure 7.45 MRAB biofilm treated with 10 mixed WOAs-60 minutes exposure
Figure 7.46 MRAB biofilm treated with 10% malic acid-30 minutes exposure
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Figure 7.47 MRAB biofilm treated with 10% malic acid 60 minutes exposure
Figure 7.48 MRAB biofilm treated with 10% lactic acid- 30 minutes exposure
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Figure 7.49 MRAB biofilm treated with 10% lactic acid- 60 minutes exposure
Figure 7.50 MRAB biofilm treated with 10% citric acid -30 minutes exposure
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Figure 7.51 MRAB biofilm treated with 10% citric acid – 60 minutes exposure
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There was also an overall reduction in estimated surface area of the bacteria, over and above that due to dehydration from ethanol treatment (Fig. 7.52).
12
10
8 MRSA VRE 6 MR-AB ESBL Kl.
surface area (mm2) area surface 4
2
0 UT 70% EtOH C15 min L15 min M15 min Mixed WOAs and exposure time
Figure 7.52 Reduction in approximate surface area of individual antibiotic resistant bacteria based on size measurements of random cells seen by SEM. (UT: untreated, C:citric acid,
L:lactic acid, M:malic acid, Mixed: 10% mixed weak organic acids)
Selection of WOAs at specified concentrations for this biofilm study was based on previous work on planktonic bacteria, presented in chapter 6.
Several currently used disinfectants such as chlorine show strong efficacy on planktonic bacteria, including multi antibiotic resistant bacteria, but their effects on bacteria in biofilms are generally limited (Rao et al., 2008 and Trachoo and Kunyaboon, 2006) ).
Planktonic bacteria are directly exposed to antimicrobial agents on their surfaces, but for a disinfectant to reach biofilm-embedded bacteria there must be strong penetration into the biofilms via water channels which include reduced adsorption to the surface of the EPS and good penetration deep into the EPS (Ntsama-Essomba et al., 1997).
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Effective disinfection of surfaces requires the elimination of, or significant reduction in, the numbers of participating planktonic and biofilm associated bacteria (Bridier et al., 2011).
The effect of WOAs on biofilm destruction could be a physical or chemical effect and could be due t acidity, although other factors cannot be excluded, such as effect on gene expression or inhibition of quorum sensing of the bacteria participating in the formation of biofilms
(Roeder et al., 2010 and Bridier et al., 2007).
As showed in chapter 6, WOA based disinfectants alone or in a combination, were effective against multi antibiotic resistant planktonic bacteria. Elimination of, or reduction in, numbers of planktonic bacteria inhibits the initiation and formation of biofilms.
Experiments on artificially formed biofilms by multi antibiotic resistant bacteria demonstrated that these selected WOAs were able to modify the structure or eliminate biofilms on surfaces, including the elimination of viable, embedded bacteria.
In this study, 10% (w/v) ascorbic acid was one of the less effective of the four WOA tested and generally required at least 45 minutes exposure to eliminate any of the four bacteria in biofilms.
Malic acid in 10% (w/v) concentration showed the poorest elimination rate on the multi antibiotic resistant biofilm bacteria and generally required 60 minutes exposure.
Citric acid in 10% (w/v) was moderately effective, but showed reduced efficacy against
ESBL+Kp. and fractionally lower efficacy against MRSA.
Lactic acid in 10% (v/v) was a very effective WOA, eliminating most of the tested biofilm bacteria after 30 minutes exposure, except MRSA.
The 10% (w/v or v/v) equal combination of all four tested WOA (ACLM) was the most effective, acting on biofilms after 30 minutes exposure, including sever changes of biofilm structure and elimination of biofilm bacteria.
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The strong antimicrobial effect of the combination of WOAs (ACLM) against planktonic and biofilm bacteria confirmed that the combination could be used as a very effective surface disinfectant, and demonstrated that WOA disinfectants were very effective against embedded, multi-antibiotic resistant, biofilm bacteria. These acids can penetrate into biofilms, eliminate biofilm-associated bacteria, and destroy the biofilm structure (EPS, water channels).
Results demonstrated that weak organic acids could effectively penetrate biofilms without being compromised by the EPS biomass. The absorption of disinfectant onto the surface and adsorption by biomass present in the biofilm is one of the main factors reducing their efficacy
(Behnke et al., 2011). The SEM photos of severely affected biofilms of the multi-antibiotic resistant bacteria showed the structural destruction of the EPS structure and water channels.
After the destruction of the EPS, embedded bacteria would be exposed in planktonic form to disinfectants, becoming an easy target for elimination. This ‘doubling’ effect, good penetration and severe structural destruction, could make these WOAs a very effective disinfectant. Due to their low toxicity (section 1.1.3 and WHO. 1973), detailed in chapter 1, these disinfectants could be used as a very effective agent against conventional or multi- antibiotic resistant bacteria, present in both planktonic and biofilm embedded, sessile form.
The low toxicity of the components (ascorbic, citric, lactic and malic acid)makes this disinfectant a safe alternative for use in high risk areas such as hospitals where colonization of multi antibiotic resistant bacteria is a significant problem and in nursing homes or kindergartens where low toxicity of the disinfectant in use could be a crucial issue. The proven efficacy against biofilm-embedded bacteria could make weak organic acid based disinfectants superior to other disinfectants, such as those that are chlorine based.
The other aspect of efficacy which must be considered in efficacy evaluation is biodegradation and the corresponding retained efficacy. Weak organic acids can be used as
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Hershberger, 2001).
Because only limited microbial species can utilize weak organic acids as a carbon source, the rate of biodegradation is very slow and this would increase retained efficacy of WOAs.
Efficacy against planktonic and biofilm bacteria, slow biodegradation and long lasting retained efficacy increases the usefulness of this disinfectant for in situ use.
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8 Development of acid tolerance by Staphylococcus aureus
8.1 Introduction
The development of acid tolerance could be a deterring factor associated with the standard use of an organic acid based disinfectant, however applying the recommended effective concentration and implementing the recommended exposure time, the risk of the development of acid tolerance could be reduced significantly.
The purpose of this experiment was to study the development of acid tolerance on S. aureus when exposed to WOAs. In this study both direct (sudden) exposure and fractional (gradual) pH changes towards a more acidic environment were used in an attempt to induce acid shock response.
S. aureus NCTC 8325 isolates exposed to WOAs and after exposure, growing on HBA, were slow growing (took up to 48 hours), forming small colonies (pinpoint) with slight pigmentation, altogether showing the characteristics of a small colony variant Staphylococcus
(Proctor et al., 1998 and Melter and Radijevic , 2010). (These isolates were sent to The
University of Newcastle for further studies, not part of this project).
Other bacteria (Enterococcus spp, Acinetobacter baumanii, and Klebsiella pneumoniae) were not studied due to the limited availability of commercial microarrays.
As a result of acid exposure, bacteria react by the production of acid shock proteins (Foster and Moreno, 2007). Gene expression, indicative of enhanced acid shock protein production was investigated by microarray assay.
The aim of this study was to obtain high quality and substantial quantity of RNA from acid treated and non-treated S. aureus, c-DNA synthesis, DNA fragmentation, hybridization, and the analysis of the synthetized and fragmented c-DNA by microarray assay.
Table 8.1 summarizes the steps and results involved in obtaining sufficient amounts of
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8.2 Results
Table 8.1 Summary of nucleic acid concentrations obtained from non-treated and acid treated Staphylococcus aureus by Trizol extraction (NanoDrop measurements).
(Examples and more details, see Appendix A)
Non-treated 260/280 260/230 Total concentration Steps in isolation of Staphylococcus aureus ratio ratio of nucleic acids (ng/µl) c-DNA NCTC 8325 Total RNA 1.6-1.8 0.4-1.0 63-70
c-DNA 1.6-1.8 1.7-1.8 550-600
c-DNA after RNA 1.3-1.5 2.0-2.1 200-220
removal
Purified c-DNA 1.8-1.9 1.4-1.6 40-50
Steps in isolation of Acid treated 260/280 260/230 Total concentration c-DNA Staphylococcus aureus ratio ratio of nucleic acids (ng/µl) NCTC 8325 Total RNA 1.7-1.9 0.7-1.2 99-110
c-DNA 1.6-1.8 1.8-1.9 650-700
c-DNA after RNA 1.3-1.5 1.9-2.0 190-200
removal
Purified c-DNA 1.9-2.0 1.7-1.8 39-41
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8.2.1 Total RNA extraction
Table 8.1 shows the ratios and concentration of total RNA extracted using Trizol® reagent from non-treated and acid treated S. aureus sampled whilst in the log-phase of growth in culture.
The purity of the extract can be assessed using Table 8.2.
The 260/280 ratio was fractionally lower (around 1.6 and 1.8) than the expected value in both treated and non-treated staphylococcal extracts. This lower than expected 260/280 ratio could be the result of some protein contamination.
The low 260/230 value was probably due to the residual phenol from Trizol® or the presence of residual carbohydrates from the cell wall. This ratio was satisfactory in all the other steps
(c-DNA synthesis and purification), ranging between 1.4 and 2.1 (Table 8.1).
To avoid or minimize any protein contamination the top ‘water phase’ from the Trizol® treated Staphylococcus was carefully removed for further work after chloroform extraction and centrifugal separation. This procedure was repeated several times to obtain the highest yield and best quality of total RNA.
The highest yields of total RNA obtained were 63 to 70 ng/µl and 99 to 110 ng/µl in a total volume of 20 µl, yielding about 1.4 µg total-RNA for non-treated S. aureus and 2.20 µg total-
RNA for acid treated S. aureus.
This RNA was converted into single stranded c-DNA for further work.
Table 8.2 260: 280 ratios of protein and nucleic acids (Glasel, 1995)
% protein % nucleic acid 260:280 ratio 100 0 0.57 95 5 1.06 90 10 1.32 70 30 1.73
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8.2.2 c-DNA synthesis
Table 8.1 shows the total c-DNA synthetized from the untreated and acid treated S. aureus
RNA.
The extracted total RNA was used to synthetise c-DNA for fragmentation which was subsequently hybridized to microarrays. The high yield of synthetised c-DNA (550 to 600 ng/
µl for untreated and 650 to 700 ng/µl for acid exposed S. aureus) was satisfactory for further work but genomic DNA contamination could not be excluded.
The 260/230 ratios (1.7 to 1.9) for both synthetized c-DNAs were acceptable and the 260/280 ratios (1.6 to 1.8) were also acceptable; but both preparations may have had some minor contamination.
The peak of the measurements for both c-DNA (from untreated and from acid treated
Staphylococcus) was between 260 nm and 270 nm (see Appendix A) indicating the presence of sufficient nucleic acid, including c-DNA.
The amount of c-DNA (probably slightly contaminated with RNA and protein) available for purification was about 23 to 25 µg in a total volume of 42 µl for untreated S. aureus and 24 to 25 µg in a total volume of 36 µl for acid treated S. aureus.
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8.2.3 RNA degradation
The purpose of the RNA degradation with 1 N sodium hydroxide was the removal of non- coding RNA, enzymes such as RNA polymerases, ligases, and excess oligonucleotides that were used in c-DNA synthesis.
The reduction in concentration for both untreated (around 13 µg in 60 µl total volume) and acid treated (around 10 µg in 45 µl total volumes) staphylococcal c-DNA confirmed the initial presence but successful removal of RNA contamination (Table 8.1).
The shift of the peaks on graphs (Appendix A) towards 260 nm also indicated the successful purification of the c-DNA solution.
The 260/230 values for both isolates were acceptable, ranging between 1.9 to 2.1 but the
260/280 ratios were fractionally low, ranging between 1.3 to 1.5 (Table 8.1, c-DNA after
RNA removal)
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8.2.4 c-DNA purification
c-DNA purification process removed salts, proteins, nucleotides and small oligonucleotides from the synthetised c-DNA mixture. After purification the total amount of available c-DNA from untreated S. aureus was around 1.5 µg in 30 µl of total volume and around 1.6 µg from acid treated S. aureus in 40 µl of total volume.
The 260/280 ratios (from both untreated and acid treated S.aureus.) were 1.8 and 2.0 indicating the presence of sufficiently pure c-DNA for fragmentation. The 260/230 ratios were 1.4 to 1.8, lower than in the previous step (RNA degradation) but still acceptable. (see
Table 8.1, purified c-DNA).
8.2.5 c-DNA fragmentation and labeling c-DNA fragments (about 200-300 bp based on the product information from Qiagen DNase protocol) were obtained, labelled and these fragment sizes were acceptable for hybridization with the complementary array sequences.
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8.2.6 c-DNA quantitation by real time PCR (Ct value)
The Ct (threshold cycle) value was measured to assess the quantity of c-DNA after fragmentation, available for hybridization. Ct is the intersection of an amplification curve and a threshold line. It is a relative estimation of the concentration of template (c-DNA in this case) in a PCR reaction. Many factors can impact on a Ct value, including the availability of the initial total RNA, successful synthesis of c-DNA, successful elimination of genomic
DNA, etc. In this experiment the Ct value was used to estimate the concentration of c-DNA
(successful conversion from total RNA), available for hybridization.
Relatively high Ct values for c-DNA from both untreated (29.4- representing about 104 copies/µl) and treated (18.7-representing about 107 copies/µl) Staphylococcus indicated the availability of relatively low amounts of c-DNA for hybridization.
This was probably the consequence of the initially low efficacy of total RNA extraction.
Figure 8.2 shows the concentration changes from extracted total RNA to purified c-DNA, ready for fragmentation. This purified c-DNA was used for the determination of the Ct value.
(Figure 8.1)
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Figure 8.1 Quantitation of data and Ct value determination for estimation the amount of c-
DNA available for fragmentation and labelling.
Concentration of RNA and c-DNA (ng/µl)
700
600
500
400
300 Untreated Staph.aureus
200 Acid treated Staph.aureus
100
0 total RNA c-DNA c-DNA after Purified RNA removal c-DNA
Figure 8.2 Changes in the concentration of RNA and c-DNA during c-DNA production for microarray assay.
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8.2.7 Hybridization
The purpose of the hybridization was to attach the fragmented and labelled c-DNA (target
DNA) to the probe (or reporter) sequence on the gene chip. This target-probe hybridization was evaluated by detecting chemiluminescence using laser reader and data were evaluated using the algorithm, GeneSpring (Agilent Technologies, USA).
Data obtained by the GeneSpring analysis were used to evaluate changes in gene expression resulting from acid shock treatment of S. aureus.
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8.2.8 Data extraction and analysis
The genome of Staphylococcus aureus (NCTC 8325) contains close to 3 million base-pairs
(2.85 Mbp) (Bershcheid, 2012) consisting of approximately 2500 open reading frames (ORF) and several thousand other coding gene sequences: http://gtps.ddbj.nig.ac.jp/single/index.php?spid=Saur_NCTC8325 (viewed on the 14th of
April, 2012).
The regulation of most of these genes are well understood (Said-Salim et al, 2003).
During acid shock treatment bacteria use several mechanisms for adaptation, including modification of protein synthesis, increased signalling mechanisms, membrane transportation changes and alterations in other metabolic activities (Chang et al, 2006).
The number of up-regulated genes due to exposure to WOAs has been studied and after exposure to peracetic acid (pKa of 8.2, a weak organic acid) expression of some genes increased several hundred times compared to the initial expression level, depending on acid exposure time (Weinrick et al, 2004).
In this present study the total number of genes expressed (up or down-regulated or not effected) was close to 8000. The total genome of this strain of S. aureus (NCTC 8325) contains several thousand genes, so the expression rate achieved in this study represents an acceptable level of expression in the microarray analysis.
The table in Appendix B is the list of all genes (close to 450) with known function expressed on the 6 arrays. GeneSpring data analysis software matches sequences expressed (hybridized on the array) with sequences in the database (e.g. Affymertrix) obtained by mapping the chromosome of the bacteria. The function of a large proportion of the sequences in the staphylococcal chromosome is unknown. Out of approximately 8000 expressed (gene) sequences, only 450 enzymes with known function were identified. The table in Appendix B summarizes all the enzymes expressed in both untreated and acid shock treated S. aureus.
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In general, enzymes which play an important role in the essential function of a bacteria, such as membrane transporters, energy producers, metabolic regulators, replication, transcription and translation regulators, toxin producers and virulence factors were expressed by acid shock treatment. This list of genes (Appendix B) with known function were identified by the
Affymetrix expression database, found on the following website: http://www.affymetrix.com/analysis/netaffx/fullrecord.affx?pk=S_AUREUS:SA_I10346DR_
X_AT (viewed on the 14th of April, 2012).
This data analysis algorithm (GeneSpring) matched these genes (obtained from the arrays) with genes from the data base (Affymetrix). The ‘similarity’ of the gene expressed to the gene listed in the database was indicated by the logarithmic value next to the gene (enzyme) name (e.g. 4.5E-29 for the Major facilitator superfamily gene). A lower value (higher number) represents a closer match to the known gene in the database.
Several thousand other gene sequences were also expressed, but due to their unknown functions, no matches were available in the database.
Due to the availability of only 6 arrays the analysis of gene expression was limited to two separate runs. Altogether 3 arrays were used to study gene expression on untreated S. aureus and 3 arrays on acid exposed (? SCV) S. aureus.
To obtain useful information about genes participating in acid shock adaptation, a comparative analysis (unpaired T-test with p<=0.05) of genes differently expressed in acid shock treated S. aureus NCTC 8325 was carried out using the Agilent’s GeneSpring expression analysis program
The Agilent’s GeneSpring expression analysis analysed the significance of differences in expression values between the two staphylococcal types (acid treated and non-treated).
On the genomic level, bacterial adaptation to acid shock treatment was expressed by either up or down-regulation of certain genes.
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Altogether 525 genes were affected, 259 were up-regulated, and 266 were down-regulated
(Appendix B).
The 52 genes with known function (from the Affymetrix database) are summarized in Table
8.3, including the value of the ‘strength of matching’ to the similar gene in the database.
Out of the 52 genes, 29 of them were up-regulated and 23 of them were down-regulated.
Affymetrix Gene Chip Catalogue (NetAffx Query) was used to obtain the function of the genes expressed; using the probe set ID from the array analysis data sheet.
The results of up and down-regulation changes of genes are detailed in Table 8.3.
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Gene expressed and matching strength Gene regulation
General substrate transporter // 1.7E-5 Up
Peptidase S1 and S6, chymotrypsin/Hap // 2.7E-7 Down
SMC protein, N-terminal // 5.1E-8 Up
Amino acid permease-associated region // 0.0099 Down
Amino acid permease-associated region // 0.014 Down
RimK-like ATP-grasp // 1.6E-4 Up
UDP-glucuronosyl/UDP-glucosyltransferase // 1.4E-4 Down
UDP-glucuronosyl/UDP-glucosyltransferase // 3.7E-4 Down
Bacterio-opsin activator, HTH // 0.0069 Up
Sulphate transporter // 0.0053 Down
Oxidoreductase FAD/NAD(P)-binding // 1.2E-4 Down
Oxidoreductase FAD/NAD(P)-binding // 1.5E-4 Down
Fumarate reductase/succinate dehydrogenase flavoprotein, N- Up terminal // 6.5E-4 FAD dependent oxidoreductase // 1.0E-6 Up
FAD-dependent pyridine nucleotide-disulphide oxidoreductase // Up 4.5E-5 Aminotransferase, class V // 2.3E-4 Down
General substrate transporter // 9.4E-5 Up
Phospholipase/Carboxylesterase // 4.2E-6 Down
Phospholipase/Carboxylesterase // 2.0E-6 Down
Saccharopine dehydrogenase // 4.4E-6 Up
Saccharopine dehydrogenase // 1.2E-5 Up
Shikimate/quinate 5-dehydrogenase // 4.0E-4 Up
UDP-glucose/GDP-mannose dehydrogenase // 5.2E-7 Up
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Fumarate reductase/succinate dehydrogenase flavoprotein, N- Up terminal // 2.0E-5 HI0933-like protein // 2.1E-5 Up
FAD-dependent pyridine nucleotide-disulphide oxidoreductase // Up 1.3E-7 FAD-dependent pyridine nucleotide-disulphide oxidoreductase // Up 4.3E-6 Glyoxalase/bleomycin resistance protein/dioxygenase // 4.3E-4 Down
Glyoxalase/bleomycin resistance protein/dioxygenase // 1.7E-4 Down
3-demethylubiquinone-9 3-methyltransferase // 0.0018 Down
3-demethylubiquinone-9 3-methyltransferase // 9.0E-4 Down
TRAP C4-dicarboxylate transport system permease DctM subunit // Down 5.5E-4 Citrate transporter // 0.0074 Down
Citrate transporter // 0.0059 Down
Carboxylesterase, type B // 8.0E-4 Up
Sigma-70, region 4 type 2 // 6.9E-5 Up
Pyridoxal-dependent decarboxylase // 7.6E-4 Down
Pyridoxal-dependent decarboxylase // 4.0E-4 Down
Fungal chitin synthase // 4.3E-4 Up
Cyclopropane-fatty-acyl-phospholipid synthase // 0.0040 Down
UbiE/COQ5 methyltransferase // 8.6E-4 Down
Protein synthesis factor, GTP-binding // 3.1E-10 Up
Major facilitator superfamily MFS_1 // 4.5E-29 Up
Glycine cleavage system P-protein // 0.0084 Up
Glycine cleavage system P-protein // 0.011 Up
Carboxyl transferase // 1.7E-11 Up
Fungal chitin synthase // 4.3E-4 Down
GTP-binding protein, HSR1-related // 2.7E-8 Up
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FAD dependent oxidoreductase // 3.1E-4 Up
Phosphotransferase system, EIIC // 1.4E-4 Up
Phosphotransferase system, EIIC // 7.8E-7 Up
Table 8.3 Summary of known genes differently expressed (up or down-regulated) in acid shock treated Staphylococcus aureus NCTC 8325 compared to control.
The ‘similarity’ of the gene expressed to the gene listed in the database was indicated by the logarithmic value (e.g. 4.5E-29 for the Major facilitator superfamily gene).
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8.2.9 List of up-regulated genes categorized by function
8.2.9.1 Genes participating in membrane transport
Major facilitator superfamily (MFS) transporters are membrane transporters, very common in prokaryotes and responsible for transporting oligosaccharides, amino acids, nucleosides,
Krebs cycle metabolites, and organic and inorganic anions and cations. 17 different families of this transmembrane protein have been identified so far and each is responsible for a specific compound transport (Pao et al, 1998).
Other integral membrane proteins strongly up-regulated due to acid shock were the General
Substrate Transporters. They are part of the Major Facilitator Superfamily which “are single- polypeptide secondary carriers capable only of transporting small solutes in response to chemiosmotic ion gradients” (EBI website) and able to transport several metabolites including acid anions, present in increased concentration intracellular after acid exposure
(Pao et al, 1998).
8.2.9.2 Genes participating in translation and signalling
GTP (guanosine triphosphate) binding protein synthesis factor is responsible for the production of GTP-binding proteins. These proteins can act as translation factors and the hydrolysis of the GTP to GDP or GMP (cGMP or cyclic GMP) is a signalling (messenger) factor, transmitting signals between cells (Mittenhuber 2001).
Shikimate/quinate 5-dehydrogenase is an essential for the biosynthesis of aromatic amino acids in bacteria (Herman and Weaver 1999) and such Saccharopine dehydrogenase participates in the synthesis of proteins necessary for acid shock response.
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RimK-like ATP-grasp protein catalyses ATP-dependent binding of a carboxyl group to an amino group and contribute to the synthesis of macromolecules (Galperin and Koonin, 1997).
8.2.9.3 Genes participating in energy production
Carboxyl transferases are biotin (Vitamin B7) dependant enzymes and one of the most important is Acetyl-CoA carboxylase which play a major role in fatty acid biosynthesis and the regulation of gluconeogenesis (Meades et al, 2011). Fatty acid synthesis and gluconeogenesis stimulation providing energy for membrane transport and strongly stimulated in the acid shock response.
FAD (Flavin-adenine- dinucleotide) dependent oxidoreductase protein is a redox cofactor that can accept or donate electrons (Negri et al. 1992 and Lehninger, Principles of Biochemistry
Part II/13). The formation of FADH2 is an energy conservation step, accepting 2 electrons
- (FAD + 2 e = FADH2) and the stored energy can be converted into ATP (Heyde and
Portalier 1990).
Phosphotransferase system is common in many bacteria and its role is to phosphorylate sugars (most importantly glucose) which will not be able to leave the cell due to the negative charge of the phosphate group (Saier, 1977). A similar phosphotransferase system is responsible for the same phenomena in glycolysis in eukaryotes (Lehninger, Principles of
Biochemistry Part II/14). This mechanism supports energy production to carry out action such as membrane transport, signaling, or protein synthesis due to acid shock.
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Fumarate reductase/succinate dehydrogenase flavoprotein, N-terminal is a distinct, membrane-bound, enzyme complex that is responsible for electron carrier activity (Blaut et al, 1989).
8.2.9.4 Genes participating in different metabolic activities
Saccharopine dehydrogenase is involved in the metabolism of lysine and in acid shock treatment probably plays an important role in the recirculation of amino acids (Leon-Ramirez et al, 2010).
UDP-glucose/GDP-mannose dehydrogenase has oxidoreductase activity, acting on the CH-
OH group of donors with NAD or NADP as acceptors (Snook et al, 2003).
HI0933-like protein is a family of conserved hypothetical proteins that may include proteins such as oxidoreductases and dehydrogenases (Miller et al, 2007).
Carboxylesterase, type B is a peptide hydrolyzing enzyme (Cygler et al, 1993).
Glycine cleavage system P-protein is part of the glycine decarboxylase multienzyme complex
(Nakai et al, 2005).
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8.2.9.5 Genes participating in the regulation of transcription
Bacterio-opsin activator is a bacterial transcription regulatory protein (Reno et al, 2009).
Sigma-70, region 4 type 2 participates in the initiation and regulation of transcription (RNA polymerase) (Campbell et al, 2002).
Structural Maintenance Protein (SMC protein), Its N-terminal group reacts with other proteins in chromosome condensation, sister-chromatid cohesion, recombination, DNA repair and epigenetic silencing of gene expression (Hearing et al, 2002).
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8.2.10 List of down-regulated genes categorized by function
8.2.10.1 Genes participating in metabolic activity
Peptidase S1 and S6, chymotrypsin is a serine protease, belongs to the chymotrypsin family, which cuts peptide bonds where the carboxyl side is a tyrosine, tryptophan or phenylalanine
(EBI website).
Aminotransferase, class V is an integral trans-membrane protein participate in metabolic process (Ouzounis and Sander, 1993).
Phospholipase / Carboxylesterase is a hydrolyzing enzyme which hydrolyses phospholipids into fatty acids (Portilla et al, 1998).
Oxidoreductase FAD/NAD (P)-binding protein plays a key role in electron flow and consequently in energy production (Bortolotti et al, 2009).
UDP-glucosyltransferase has transferase activity and also specifically inactivates macrolide antibiotics (Hernandez et al, 1993).
8.2.10.2 Genes participating in biosynthesis
Glyoxalase/bleomycin resistance protein/dioxygenase catalyzes the first step of the glyoxal pathway which is the major product of DNA oxidation (Fillgrove et al, 2007).
3-demethylubiquinone-9 3-methyltransferase is a transferase and participates in ubiquinone biosynthesis which is part of the electron transport chain. (DelVecchio et al, 2002).
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Pyridoxal-dependent decarboxylase is primarily involved in the biosynthesis of amino acids and amino acid-derived metabolites (Eliot and Kirsch, 2004).
Cyclopropane-fatty-acyl-phospholipid synthase is a transferase and participates in the lipid biosynthetic process (Huang et al, 2002).
UbiE/COQ5 methyltransferase participate in the biosynthesis of ubiquinone and is involved in biotin and sterol biosynthesis (Lee et al, 1997).
Fungal chitin synthase is a plasma membrane-bound protein which plays a major role in cell wall biogenesis (Specht et al, 1996).
8.2.10.3 Genes participating in transportation
Amino acid permease is an integral membrane protein involved in the transport of amino acids into the cell (Weber et al, 1988).
Sulphate transporters are integral trans-membrane proteins involved in the transport of sulphate across a membrane (Smith et al, 1995).
Citrate transporter participates in the uptake of citrate (Krom er al, 2000).
TRAP C4-dicarboxylate transport system permease DctM subunit is part of the bacterial and archaeal TRAP C4-dicarboxylate transport system permease and allows C4-dicarboxylates like succinate, fumarate, and malate to be taken up (Janausch et al, 2002).
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8.3 Discussion
Individual cells react to an external stimulus first by activating a signaling system to deliver the specific intracellular signal which triggers a response in a form of transcription and translation, resulting in the production of proteins which play a major role in activities such as membrane transport by producing trans-membrane proteins (Weinrick et al, 2004).
Genes strongly expressed (up-regulated) in acid exposed S. aureus were those responsible for membrane transport, transcription regulation, translation and signaling, electron transport resulting in energy production and energy storage and metabolic activities such as oxido- reductase and hydrolase activity. All of these functions would be essential for the adaptation to an acid shock response (Bore et al. 2007).
Up-regulated genes that participate in transportation (trans-membrane) probably play a vital role in acid shock adaptation (section 1.3, Figure 1.1), because increased intracellular anion concentration forces potassium ion transport into the cell to maintain electrical balance and increase osmotic pressure. This cation transport is probably facilitated by the Major
Facilitator Superfamily transporters and General Substrate transporters. Therefore it would be consistent to observe up-regulation of those genes responsible for the production of transmembrane proteins.
Cell to cell signaling and regulation of transcription and translation in protein synthesis, leads to the production of pre shocked and acid shock proteins.
Increased translation for the production of acid shock proteins (Heyde and Portalier 1990) is a natural response for a bacterium to survive a harsh condition such as exposure to low pH.
Membrane transport, signaling, transcription or translation are all energy demanding activities and the up-regulation of genes involved in energy production such as carboxyl transferase or FAD dependent oxydoreductase or phosphotransferase or succinate dehydrogenase or fumarate reductase are essential to provide energy in the form of ATP.
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Signalling also can help to coordinate these responses amongst individual bacterial cells.
Genes up-regulated and participating in different metabolic activities such as saccharopine dehydrogenase or carboxylestarse type B or the glycine cleavage system P-protein are essential in the recirculation of amino acids, providing building blocks for the synthesis of pre-shock or acid shock proteins.
Several genes were down-regulated after exposure to WOAs. These genes can be classified into three groups, genes with specific metabolic activities, genes participating in biosynthesis, and genes participating in some specific membrane transportation.
Some peptidase and aminotransferase functions are not essential when cells are exposed to shock treatment and some specific metabolic activity such as phospholipid hydrolization or antibiotic inactivation may also not be necessary vital function for a bacterial cell to survive.
According to results obtained by this experiment, several biosynthetic activities were also down-regulated. A possible explanation for the down-regulation of certain biosynthetic activity could be to minimise any non-vital function that requires significant energy and divert this energy use towards more important (vital) activities such as membrane transport or essential protein synthesis.
The down-regulation of some non-essential transport activity, such as amino-acid permease or sulphate or citrate transporter are probably the result of the redistribution and separation of vital and non-vital cellular activity.
Bacterial cells developed several adaptation functions to adjust to environmental changes.
Adaptation to heat, radiation, and changes in chemical balance including pH changes are essential for their survival. Adaptation is not just a survival mechanism for bacteria. Stress response systems also can play an important role in the expression of virulence in pathogenic organisms (Requena, 2012). For example, exposure to benzalkonium chloride can promote the development of antibiotic resistance (Mc Cay et al, 2010).
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In summary, this work – not conclusively, but satisfactorily- confirmed previous findings by others (Bore et al., 2007, Foster 1991, 2007, Heyde, 1990) , that exposure to pH changes, specifically to WOAs can trigger an adaptive response in bacterial cells, including specific up and down-regulation of certain genes. The changes in gene expression result in altered protein synthesis and changes in transmembrane functions, such as ion transportation. The findings of this study in many aspects correlates very well with the findings by Chang et al.
(2006) including the regulation of membrane transport genes and the expression of metabolism-related genes.
All these functional changes would be necessary for the organism to survive or in some cases even could trigger an increase in virulence.
This experiment also raised several questions and indicated that further work would be necessary to investigate the differences in acid shock protein production with gradual or instant acid exposure. The time dependency of acid shock protein production, and changes in regulation of the pathogenicity islands genes of exposed bacteria, may result in altered pathogenicity.
These findings should not deter the use of disinfectants based on WOAs; rather it should highlight the importance of the responsible use of disinfectants or any other antimicrobial agent, such as antibiotics. Responsible use includes the use of correct concentration and correct exposure time, as both are necessary for the optimal effect of an antimicrobial agent.
Responsible use reduces the chance of the development of any resistance or other bacterial adaptation, such as spore formation, small colony variant formation or biofilm formation http://www.cdc.gov/mrsa/environment/- (viewed on the 20th of October, 2012).
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9 Artificial and genuine in situ studies
9.1 Introduction
Artificial in situ study was performed on ceramic tiles, and provided a good indication of the performance of the WOA disinfectant under dirty condition where the effect of organic soiling (fetal calf serum) was evaluated. Data obtained from this experiment supported the next step, a genuine in situ study on VRE.
A genuine in situ study assesses the efficacy of a disinfectant in a situation where it will be used on an regular basis. These studies evaluate the efficacy (at a given concentration and exposure time) of the disinfectant as affected by environmental factors, such as temperature, pH, light exposure, and non-organic or organic contamination of surfaces.
An in situ study carried out in an environment which is heavily contaminated (colonized) with pathogenic bacteria (including multi-antibiotic resistant such as VRE) either can prove or disprove the efficacy of a disinfectant in practice.
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9.2 Artificial in situ study on ceramic tiles
This study was carried out to assess the activity of the 10% mixed WOA disinfectant in dirty condition, when environmental contamination by organic matter, such as proteins, lipids, etc. could affect the activity of a disinfectant.
The other purpose of this study was to obtain some data from an artificially designed in situ test extrapolate this finding to the design of a genuine in situ test, detailed in the following chapter.
9.2.1 Results and discussion
The study was conducted on ceramic tile to assess the efficacy of the 10% (v/v or w/v) WOA combination supplemented with 1% foetal calf serum (to simulate dirty conditions by absorbing disinfectant and reducing efficacy) on an inanimate surface, using commercially available ceramic tiles.
Table 9.1 represents the average reduction in bacterial count per cm2. For the two Gram positive cocci, MRSA (Figure 9.1) and VRE the trial achieved an average 5-log10 reduction.
For ESBL positive Klebsiella pneumoniae the reduction was fractionally lower, around 4.5- log10 and for the other Gram negative bacillus, MRAB (Figure 9.2) the reduction was less, but still resulting in an average of 3.5-log10 reduction.
The more resistant nature of Gram negative bacilli was probably due to the presence of a mucoid capsule which provided a natural barrier against weak organic acids penetration into the bacterial cells.
The addition of 1% foetal calf serum simulates “dirty” conditions-absorbance of disinfectant by organic materials, resulting in decreased efficacy- due to contamination of the environment with organic matter (Reybrouck1992).
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The 3.5-log10 reduction or greater was sufficient to claim the disinfectant as effective in the presence of an organic load (Bailey et al., 2011 and BS EN 13697:2001- British Standard
Institution: http://www.bis.gov.uk/policies/innovation/standardisation/bsi-) (viewed on the
20th of October, 2012 ).
Organic contaminations are very common in real life situations due to the presence of fatty acids and proteins and they may be responsible for the reduced activity of commercially available disinfectants, in situ.
The outcome of the above test was two-fold; WOA based disinfectant showed to be effective in the presence of organic contamination and also this finding suggested that this disinfectant could be used without the pre-treatment of surfaces with detergent.
Pre-treatment of surfaces prior to use of any disinfectant can be time consuming, labor intensive and increase cost.
This study also provided background data for the genuine in-situ study at Barwon Health, where areas tested were heavily exposed to organic matters, especially to fecal contaminations.
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Table 9.1 Reduction in bacterial count (inoculating concentration: >1012 CFU/ml) on ceramic tiles using the 10% (v/v or w/v) combination of WOAs (with the addition of 1% fetal calf serum), for four antibiotic-resistant bacteria (MRSA, VRE, MRAB and ESBL + Kp.) in
3 replicates.
Bacteria Bacterial count and log-10 reduction
MRSA
Pre-exposure bacterial count 5 x 107 1 x 107 5 x 107 (CFU/cm2) Post-exposure bacterial count <5 x 102 <5 x 102 <5 x 102 (orgs/cm2) Log-10 reduction 5 4.5 5 VRE
Pre-exposure bacterial count 5 x 107 5 x 107 5 x 107 (CFU/cm2) Post-exposure bacterial count <5 x 102 <5 x 102 <5 x 102 (orgs/cm2) Log-10 reduction 5 5 5 ESBL positive Klebsiella pneumoniae Pre-exposure bacterial count 1 x 107 1 x 107 5 x 107 (CFU/cm2) Post-exposure bacterial count <5 x 102 <5 x 102 <5 x 102 (orgs/cm2) Log-10 reduction 4.5 4.5 5 MRAB
Pre-exposure bacterial count 1 x 106 1 x 106 5 x 106 (CFU/cm2) Post-exposure bacterial count <5 x 102 <5 x 102 <5 x 102 (orgs/cm2) Log-10 reduction 3.5 3.5 3.5
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Figure 9.1. Horse blood agar plates inoculated with 100 µl of MRSA suspension, collected from 1 cm2 ceramic tile before and after exposure to WOAs
Figure 9.2 Horse blood agar plates inoculated with 100 µl of MRAB suspension, collected from 1 cm2 ceramic tile before and after exposure to WOAs
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9.3 Genuine in situ study of VRE disinfection at Barwon Health
9.3.1 Results
Table 9.2, 9.3, 9.4, and 9.5 summarises the results obtained by the genuine in situ testing at
The Geelong Hospital and at the McKellar Centre. Due to more extensive use of the toilet and bathroom facilities at the McKellar Centre (elderly but mobile patients) the pre-swabbing contamination by VRE was much higher than in the hospital ward, where many patients were bed-bound.
At Birdsey Wing 6 the highest number of swabs positives for VRE (prior to disinfection) were detected on the toilet seat, followed by the floor surrounding the toilet. The toilet flush had visible contamination and the swabbing of the toilet cover, hand basin, the taps and the portable toilet seat also resulted in VRE growth (prior to disinfection).
Sites such as handrails, doorhandles, and shower handles tested negative for VRE.
In total 21 VRE positive swabs were obtained prior to disinfection. This represented 9.1%
VRE-positivity of the 150 swabs collected (Table 9.2).
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Site Total number of swabs Number of positive Number of positive
before-after cleaning swabs growing VRE swabs growing VRE
before cleaning after cleaning
Floor 18-18 3 0
Toilet seat 18-18 10 1
Toilet cover 18-18 1 0
Portable toilet seat 18-18 1 0
Toilet hand rail 9-9 0 0
Shower hand rail 9-9 0 0
Bathroom tap-hot 6-6 1 0
Bathroom tap-cold 6-6 1 0
Hand basin 18-18 1 0
Door handle 18-18 0 0
Toilet flush 3-3 2 0
Shower handle 3-3 0 0
White, plastic chair 6-6 1 0
Total number of swabs: 150/150
Total number of positives before cleaning: 21 21/150= 9.1%
Total number of positives after cleaning: 1 1/150= 0.007%
Total reduction 20/21= 95.2%
Table 9.2 The effect of WOAs on an environment contaminated with VRE. Summary of results from Birdsey Wing 6, The Geelong Hospital, Barwon Health.
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Site Total number of swabs Number of positive Number of positive
before-after cleaning swabs growing VRE swabs growing VRE
before cleaning after cleaning
Shower handle 2-2 0 0
Toilet seat 11-11 5 0
Floor 9-9 2 0
Hand rail 6-6 0 0
Bathroom tap-cold 3-3 0 0
Bathroom tap- hot 3-3 0 0
Hand basin 7-7 1 0
Toilet cover 6-6 1 0
Toilet flush 3-3 1 0
Total number of swabs: 50/50
Total number of positives before cleaning: 10 10/50=20%
Total number of positives after cleaning: 0 0/50= 0%
Total reduction 10/10= 100%
Table 9.3 The effect of WOAs on an environment contaminated with VRE.
Summary of results from McKellar Centre, Barwon Health.
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Site Total number of swabs Number of positive Number of positive
before-after cleaning swabs growing VRE swabs growing VRE
before cleaning after cleaning
Floor 27-27 5 0
Toilet seat 29-29 15 1
Toilet cover 24-24 2 0
Portable toilet seat 18-18 1 0
Toilet hand rail 15-15 0 0
Shower hand rail 9-9 0 0
Bathroom tap-hot 9-9 1 0
Bathroom tap-cold 9-9 1 0
Hand basin 25-25 2 0
Door handle 18-18 0 0
Toilet flush 6-6 3 0
Shower handle 3-3 0 0
White, plastic chair 6-6 1 0
Total number of swabs: 200/200
Total number of positives before cleaning: 31 31/200= 15.5%
Total number of positives after cleaning: 1 1/200= 0.005%
Total reduction 30/31= 96.8%
Table 9.4 The effect of WOAs on an environment contaminated with VRE.
Combined summary of results (The Geelong Hospital- McKellar Centre)
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Site Number of positive swabs Number of positive swabs
growing VRE before cleaning growing VRE after cleaning
(Group 1) (Group 2)
Floor 5 0
Toilet seat 15 1
Toilet cover 2 0
Portable toilet seat 1 0
Bathroom tap-hot 1 0
Bathroom tap-cold 1 0
Hand basin 2 0
Toilet flush 3 0
White, plastic chair 1 0
(Group 1) (Group 2)
Mean (µ) 3.44 0.11
Standard deviation 4.53 0.33
Standard error of means 1.51 0.11
Number of sample sites 9 9
Two-tailored P value: 0.043
T-test value: 2.201
T value shown in Student’s T test table: 1.746 df: 16 (number of samples -1)
Table 9.5 Comparison of VRE positive swabs pre-and post-WOA disinfection (T-test) from different locations within Birdsey Wing 6 (Geelong Hospital) and McKellar Centre.
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Figure 9.3 Heavy growth of VRE (left plate) from toilet seat on bile aesculin agar containing
6 µg/ml vancomycin (pre-treatment with WOAs) and no growth (right plate) post-WOA treatment.
Figure 9.4 Moderate growth of VRE (left plate) from floor on bile aesculin agar containing
6 µg/ml vancomycin (pre-treated with WOAs) and no growth (right plate) post-WOA treatment
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9.3.2 Discussion
This in situ study was carried out to asses the activity of the WOA disinfectant in a real life situation against a multi antibiotic resistant bacterium, VRE. This is one of most prevalent multi-resistant organisms with major implications for patients’ health, especially those with lowered immunity
Swabbing after disinfection resulted in only 1 positive culture in total, on the toilet seat. This positive swab had only one VRE colony growing on the selective microbiological media (see
Figure 9.3 and 9.4).
Less frequent cleaning and more frequent faecal exposure at the McKellar Centre contributed significantly for the detection of higher number of VREs prior to cleaning, providing a better opportunity to assess the efficacy of the disinfectant.
At the McKellar Centre 10 VRE positive swabs was detected prior to cleaning in all three rooms from a total of 50 swabs. No positive culture was obtained after disinfection (Table
9.3).
In total, 31 VRE positive swabs were detected out of 200 taken prior to cleaning and only one was positive out of 200 swabs taken after cleaning (Table 9.4). These 31 swabs represent
15.5% VRE positivity. After cleaning the 1 positive culture represented a 0.005% positivity rate, a reduction of 96.8% due to WOA disinfection.
Table 9.5 shows all the calculated data. The paired T-test analysed two dependent groups
(before disinfection- Group 1- and after disinfection- Group 2, (Table 9.5). The sample size from both groups was equal and the standard deviation was expected to be same in both groups.
The calculated p value was equal to 0.043 which indicated that the difference due to WOA disinfection of VRE between the two groups was statistically significant. For the statistical analysis 95% confidence interval was used.
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The calculated T value was 2.201
The ‘expected’ T value shown in Student’s T test table was 1.746 (Senn and Richardson,
1994).
The ‘null hypothesis’ for this experiment was that there was no significant difference in the number of VRE positive swabs on the tested surfaces before and after disinfection.
Because the calculated T value was greater than the expected value in a Student’s T value table, the null hypothesis was rejected. There was a significant difference in VRE count before and after treatment.
The statistical analysis together with preliminary data on the efficacy of the WOA disinfectant strongly suggests that it was an effective disinfectant able to eliminate pathogenic bacteria such as VRE from surfaces.
A reduction in VRE is likely to lead to fewer fatal infections, and reduced need for contact isolation of patients, as stated on the National Institutes of Health (NIH-US) website; http://www.niaid.nih.gov/topics/antimicrobialresistance/examples/vre/Pages/overview.aspx-
(viewed on the 21st of October 2012)
The reduction or elimination of VRE or other multi-antibiotic resistant bacteria from surfaces surrounding patients has significant social and financial impact. Colonisation affects a large percentage of patients in hospitals, especially among haematology and oncology patients.
Colonisation places them at risk of invasive infection with an increased incidence noted over the last several years. Any product that can reduce the environmental burden of antibiotic- resistant bacteria will subsequently reduce both colonisation and infection. Reduced colonisation and subsequent infections will reduce the length of stay in hospital and reduce cost of health care. The availability of a safe (low toxic) and effective product (10% WOA) to achieve proper decontamination is likely to contribute to all of the above.
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The artificial in situ study on tiles with other multi-antibiotic resistant bacteria, including
MRSA, MRAB or ESBL positive Gram negative (see chapter 9.2) demonstrated that this disinfectant had the capacity to reduce or completely eliminate bacterial contaminations from surfaces, containing organic materials and pathogenic bacteria.
The genuine in situ study demonstrated that disinfectants containing weak organic acids can be effective against multi-antibiotic resistant bacteria (VRE) in real life situations. Due to the low toxicity (section 1.1.3) and negligible safety risks, WOAs expose no significant harm to the environment, to patients or to health workers.
This project was accepted by the “Office for Research” at The Geelong Hospital, as shown by the approval in Appendix C.
Also, this part of the project was submitted to the Smart Geelong Network Research Expo and was a ‘Researcher of the Year’ finalist in 2011 (Appendix C).
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10 Overall discussion
Effective disinfection is important in preventing healthcare associated infections, and achieving appropriate disinfection through the use of disinfectants is essential for ensuring that the surrounding environments do not transmit infectious pathogens to patients in healthcare facilities, such as hospitals and nursing homes. Numerous published articles emphasise the requirement for all health care services to use appropriate disinfection as detailed in a book edited by Rutala (2010).
This thesis studied the antimicrobial efficacy of selected WOAs, and investigated the possibility of using WOAs as the basis for a new class of safer, efficacious, and more environmentally responsible disinfectants.
Disinfectants currently available for domestic or hospital use have a number of drawbacks.
These include the significant human toxicity of chlorine-based disinfectants (White and
Martin, 2010); the limited efficacy of chlorhexidine-based products (Kampf et al., 1998); relatively poor abilities to penetrate biofilms (Holah et al., 2008); and the tendency to induce antibiotic resistance in the target organisms (McDonnell and Russell, 1999).
A trial disinfectant developed through this study and containing naturally occurring WOAs was able to overcome most of those problems. Furthermore, this study demonstrated that this type of disinfectant could be very effective against a wide range of microbes, including enveloped and non-enveloped viruses, intracellular, planktonic and biofilm embedded bacteria. However, only limited efficacy against yeast and mould was found.
In further investigating the antimicrobial activity of WOA-based disinfectants, numerous aspects of the activity are addressed in this thesis. The low mammalian toxicity of WOAs is discussed in chapter 1 (section 1.1.3); the mode of antiviral action (via the denaturation of capsid proteins) is presented in section 1.2; the antibacterial action (by forced active membrane transport is examined in section 1.3; while the ability of WOAs to penetrate and
175 | P a g e eliminate biofilms is detailed in chapter 7. All these activities support the broad-spectrum efficacy of this disinfectant, succinctly summarized in figure 10.1(chapter 10).
The efficacy of the WOA based disinfectants against intracellular pathogens in a concentration as low as 1.5 mg/ml (Figure 10.1) increases the scope of the practical use of this type of disinfectant. Elimination of intracellular pathogens such as Rickettsia prowazekii is difficult and requires a combination of disinfectants such as phenol or formaldehyde
(Eremeeva et al., 1989). These disinfectants are highly toxic to eukaryotes in comparison with disinfectants containing WOAs only (Tisler and Zagorc-Koncan, 1997).
The effect of WOAs on planktonic bacteria was studied using multi-antibiotic resistant bacteria, such as multi resistant Staphylococcus aureus; vancomycin resistant Enterococcus spp.; multi resistant Acinetobacter baumani;, and extended spectrum beta lactamase positive
Klebsiella pneumoniae. These bacteria pose enormous risks to societies all around the world and the cost of their elimination is significantly higher than the elimination of any non multi resistant bacteria (Cirzet al, 2005). WOAs showed antimicrobial efficacy against these bacteria in concentrations of about 100 mg/ ml (Figure 10.1) in in-house testing during this project.
External testing by a Therapeutic Goods Administration accredited laboratory (Siliker,
Australia) confirmed the finding that the 10% mixed WOA disinfectant was effective and, based on their testing, could therefore be classified as a hospital grade disinfectant (Appendix
D). This opens the way for further commercial development as efficacy testing of a disinfectant by a TGA approved laboratory is a requirement for TGA registration (together with appropriate labelling and the production of the disinfectant under Good Manufacturing
Practice (GMP)).
In exploring the practical applications and commercial potential of this class of compounds as a new type of disinfectant, WOAs was found to be effective against both enveloped and non-
176 | P a g e enveloped virus in concentrations as low as 6 mg/ml (Figure 10.1). In fact, due to the presence of an envelope, which is a membrane fraction from the host cells-providing an excellent target surface for disinfectant- enveloped viruses are more susceptible to disinfectants and easier to eliminate than non-enveloped viruses (Block, 2000) such as influenza virus. The ability to eliminate non-enveloped viruses is very important (Kampf and
Kramer, 2004), and include important human pathogen such as norovirus, human adenovirus, and rotavirus which are responsible for the majority of non-bacterial gastroenteritis cases around the world. These viruses are also known to be relatively resistant to alcohol-based disinfectants (Widdowson, 2005 and Bernstein, 2009). In in-situ testing, WOA-based disinfectant was able to eliminate viruses from hard surfaces thus reducing the chance of their transmission via inanimate surfaces, or if used as a hand-disinfectant via direct contact
(Turner & Hendley, 2005 and Turner et al., 2010). Hospitals, kindergartens, nursing homes are highly exposed to viral transmission and the reduction or complete elimination of this transmission and the consequent reduction in nosocomial infection could have enormous health, social and financial benefits. http://www.health.nsw.gov.au/factsheets/infectious/gastroenteritis.html (viewed on the 22nd of October, 2012).
The in vitro efficacy of the WOA mixture against microbes, including multi antibiotic resistant bacteria, was supported by an in situ study at The Geelong Hospital, Victoria,
Australia. Two wards were selected for this study, both of which were accommodating patients colonized or infected with vancomycin resistant Enterococcus spp. (VRE). The study showed that the 10% mixture of the four WOAs was able to eliminate VRE from surfaces and the efficacy was comparable or greater than the commercially available disinfectant
Viraclean®, which is a benzalkonium chloride-based product produced by Whiteley
Industries Pty.Ltd., NSW, Australia (Friedman, 2012). The in situ efficacy of WOAs against
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VRE was comparable to the reported efficacy of a disinfectant containing phenol and quaternary ammonia, and exceeded that of a hydrogen peroxide based preparation (Saurina et al., 1997). In summary, data generated in this study via artificial and genuine in situ studies demonstrated the ability of this class of disinfectant to be used in hospitals and in other health care environments as an effective agent to fight multi antibiotic resistant bacteria.
In previous studies, biofilm-embedded bacteria have shown reduced susceptibility to current disinfectants, which could be the result of limited penetration into biofilms, or increased absorbance by the extracellular polymeric substance; http://cfpub.epa.gov/si/si_public_record_Report.cfm?dirEntryId=230297&CFID=104610777
&CFTOKEN=82508780&jsessionid=4e30a3e0cbbf0e59a75b465139f2153233f3 viewed on the 22nd of October, 2012 and De Beer et al., 1994).
Chlorine based disinfectants also have limited efficacy against biofilm bacteria (Rao et al.,
2008 and Holah et al., 2008). In contrast, a 10% WOA-based disinfectant mixture, developed in this project, exhibited an acceptable elimination rate of biofilm embedded, multi antibiotic resistant bacteria, although in comparison with planktonic bacteria, the exposure time had to be increased from 10 to 30 minutes.
The issue of the development of acid tolerance in bacteria (Ricke, 2003 and Alvarez-Ordóñez et al., 2009) is a potential problem which may influence the long-term efficacy of a disinfectant based on WOAs, and was raised in the literature review (chapter 1), before further investigation in this study.
To explore the development of acid tolerance in bacteria induced by exposure to WOAs, a microarray analysis was carried out on Staphylococcus aureus studying gene expression (up and down regulation) due to acid shock treatment. Although this study had only a limited success, it showed that acid exposure stimulates cellular mechanisms (up-regulation of genes) such as membrane transport, transcription and translation regulation, cell to cell signalling,
178 | P a g e energy production and other metabolic activities such as oxydo-reductase and hydrolase activity. Conversely, certain genes were down-regulated post acid exposure, largely genes which were not participating in vital biosynthetic functions. An earlier study by Bore (2007) on Staphylococcus aureus similarly confirmed up and down regulation of genes due to acid exposure.
In summary, the objective of this study was to evaluate the efficacy of WOAs against microbes, including viruses, intracellular, planktonic and biofilm embedded bacteria and fungi. Results obtained on antibacterial efficacy in this study were confirmed and compared with external test results (Appendix D). It was shown that WOAs could be used effectively as disinfectants, having low toxicity compared to other commercially available disinfectants while expressing acceptable antimicrobial efficacy and in some cases, such as in biofilm penetration, express better than average efficacy (Holah et al., 2008). Consequently, there may be potential for a commercial development of WOAs as an alternative disinfection technology for use in health and other industry sectors.
This study has also revealed opportunities for subsequent projects, for example in investigating the underlying reasons for the limited antifungal efficacy observed here.
For example, these studies showed only a modest antifungal efficacy of WOAs, and required a concentration of at least 300 mg/ml to eliminate yeast such as Candida albicans or fungus such as Aspergillus niger (Figure 10.1). The limited fungicidal activity of existing disinfectants generally is well known and published (Gupta et al., 2001 and 2002), although a recent study investigated the growth inhibition of WOAs on Saccharomyces cerevisiae (Ullah et al., 2012), and concluded that the yeast cell’s ability to restore pH was a prime determinant of inhibition, rather than simple acidification.
Further studies may evaluate the use of WOAs in areas such as cosmetics (for example as an anti-acne face wash, James, 2005) or food preservation (Ross and Morgan, 2002), and further
179 | P a g e explore the efficacy of different combinations and concentrations of WOAs with the aim of the development of a more effective antimicrobial agent, based on WOAs, including a more effective anti-fungal agent.
Shortly after the completion of this thesis, the TGA (Australia) registered this disinfectant based on WOAs as a hospital grade disinfectant.
180 | P a g e
Viruses 6 mg/ml Intracellular bacteria 10 minutes 1.5 mg/ml (Effective) 10 minutes (Effective)
Fungus and Yeast >300 mg/ml 30 minutes Weak Organic Acid disinfectant (Moderately effective)
Planktonic bacteria 100 mg/ml Biofilm bacteria 10 minutes 100 mg/ml (Effective) 30 minutes (Effective)
Figure 10.1 Summary of effective concentration and exposure time required to eliminate
microbes
181 | P a g e
11 Presentations, publications and awards
1. European Conference of Clinical Microbiology and Infectious Diseases
Helsinki 2009
- Efficacy of selected weak organic acids against enveloped and non-
enveloped viruses and an intracellular bacterium (poster presentation).
Z .Nack, S .Graves, H. Dunstan, J. Stenos, T. Roberts
- Efficacy of selected weak organic acids against multi antibiotic resistant
planktonic and biofilm bacteria (poster presentation).
Z .Nack, S .Graves, H. Dunstan, J. Stenos, T. Roberts
2. European Conference of Clinical Microbiology and Infectious Diseases
Vienna 2010
- Effect of weak organic acids on biofilms as observed by scanning electron
Microscopy (poster presentation)
Z .Nack, S .Graves, H. Dunstan, J. Stenos, T. Roberts
3. Smart Geelong Network, Researcher of the Year finalist 2011(poster presentation by
Z. Nack, Appendix C)
182 | P a g e
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13 Appendices
13.1 Appendix A
The following NanoDrop (spectrophotometer) images show the amount of total RNA, extracted from untreated and acid treated bacteria respectively and converted into c-DNA.
Extracted total RNA from untreated Staphylococcus aureus NCTC 8325
Extracted total-RNA from acid treated Staphylococcus aureus NCTC 8325 225 | P a g e
c-DNA from untreated Staphylococcus aureus NCTC 8325
c-DNA from acid treated Staphylococcus aureus NCTC 8325
226 | P a g e
RNA degraded c-DNA from untreated Staphylococcus aureus NCTC 8325
RNA degraded c-DNA from acid treated Staphylococcus aureus NCTC 8325
227 | P a g e
RNA degraded and purified c-DNA from untreated Staphylococcus aureus NCTC 8325
RNA degraded and purified c-DNA from acid treated Staphylococcus aureus NCTC 8325
228 | P a g e
13.2 Appendix B
Summary of genes with known function expressed on all 6 arrays # Notes : All Entities # Technology : Affymetrix.GeneChip.S_aureus # Owner : gxuser Signals for all genes on array
IPR001899 // Surface protein from Gram-positive cocci, anchor region // 1.1E-4 /// IPR001899 // Surface protein from Gram-positive cocci, anchor region // 5.5E-5 IPR001245 // Tyrosine protein kinase // 1.4E-14 IPR002938 // Monooxygenase, FAD-binding // 2.9E-4 /// IPR003953 // Fumarate reductase/succinate dehydrogenase flavoprotein, N-terminal // 1.6E-5 IPR013027 // FAD-dependent pyridine nucleotide-disulphide oxidoreductase // 7.3E-4 IPR005828 // General substrate transporter // 1.7E-5 IPR010656 // TRAP C4-dicarboxylate transport system permease DctM subunit // 0.0058 /// IPR010656 // TRAP C4-dicarboxylate transport system permease DctM subunit // 0.0053 IPR000192 // Aminotransferase, class V // 7.0E-4 /// IPR000192 // Aminotransferase, class V // 7.3E-4 IPR011701 // Major facilitator superfamily MFS_1 // 4.5E-29 IPR011545 // DEAD/DEAH box helicase, N-terminal // 1.3E-7 IPR001269 // Dihydrouridine synthase, DuS // 3.1E-5 IPR000835 // Bacterial regulatory protein, MarR // 3.5E-6 /// IPR011611 // PfkB // 3.1E-8 /// IPR013749 // Phosphomethylpyrimidine kinase type-1 // 9.7E-5 IPR004479 // ExsB // 1.9E-4 /// IPR011063 // PP-loop // 0.13 IPR000795 // Protein synthesis factor, GTP-binding // 3.1E-10 IPR002938 // Monooxygenase, FAD-binding // 2.9E-4 /// IPR004792 // HI0933-like protein // 5.4E-46 IPR001254 // Peptidase S1 and S6, chymotrypsin/Hap // 2.7E-7 IPR000192 // Aminotransferase, class V // 7.0E-4 IPR003395 // SMC protein, N-terminal // 5.1E-8 IPR004680 // Citrate transporter // 4.8E-4 /// IPR004680 // Citrate transporter // 3.1E-4 /// IPR010656 // TRAP C4-dicarboxylate transport system permease DctM subunit // 0.0065 /// IPR010656 // TRAP C4-dicarboxylate transport system permease DctM subunit // 0.0059 IPR004099 // Pyridine nucleotide-disulphide oxidoreductase dimerisation region // 3.2E-5 IPR006380 // Sucrose-6F-phosphate phosphohydrolase, plant and cyanobacteria // 6.2E-4 IPR008914 // PEBP // 4.5E-4 IPR005828 // General substrate transporter // 4.3E-5 /// IPR005828 // General substrate transporter // 7.4E-6 IPR004841 // Amino acid permease-associated region // 0.0099 /// IPR004841 // Amino acid permease-associated region // 0.014 IPR001296 // Glycosyl transferase, group 1 // 0.0024 /// IPR001296 // Glycosyl transferase, group 1 // 0.0034 IPR000795 // Protein synthesis factor, GTP-binding // 5.0E-24 IPR012001 // Thiamine pyrophosphate enzyme, N-terminal TPP binding region // 0.0021 /// IPR012001 // Thiamine pyrophosphate enzyme, N-terminal TPP binding region // 0.0011 IPR002917 // GTP-binding protein, HSR1-related // 2.7E-8 IPR013651 // RimK-like ATP-grasp // 1.6E-4
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IPR007848 // Methyltransferase small // 8.9E-4 IPR003806 // Protein of unknown function DUF201 // 7.4E-4 /// IPR003806 // Protein of unknown function DUF201 // 7.1E-4 IPR006143 // Secretion protein HlyD // 4.3E-4 IPR002218 // Glucose-inhibited division protein A // 2.5E-6 /// IPR002938 // Monooxygenase, FAD-binding // 4.2E-4 /// IPR003953 // Fumarate reductase/succinate dehydrogenase flavoprotein, N-terminal // 1.0E- 10 /// IPR004792 // HI0933-like protein // 2.7E-8 IPR013108 // Amidohydrolase 3 // 0.0013 IPR000115 // Phosphoribosylglycinamide synthetase // 1.8E-5 /// IPR000115 // Phosphoribosylglycinamide synthetase // 1.7E-5 /// IPR003806 // Protein of unknown function DUF201 // 7.9E-4 /// IPR003806 // Protein of unknown function DUF201 // 7.2E-4 /// IPR013651 // RimK-like ATP-grasp // 0.0053 /// IPR013651 // RimK-like ATP-grasp // 0.0040 IPR004792 // HI0933-like protein // 7.5E-6 /// IPR013027 // FAD-dependent pyridine nucleotide-disulphide oxidoreductase // 0.0042 IPR004843 // Metallophosphoesterase // 2.3E-5 IPR006123 // Staphylococcal/Streptococcal toxin, beta-grasp // 0.13 /// IPR006123 // Staphylococcal/Streptococcal toxin, beta-grasp // 0.11 IPR006076 // FAD dependent oxidoreductase // 3.1E-4 IPR011704 // ATPase associated with various cellular activities, AAA-5 // 5.0E-7 IPR004088 // KH, type 1 // 0.075 IPR004365 // nucleic acid binding, OB-fold, tRNA/helicase-type // 0.78 /// IPR006935 // Type III restriction enzyme, res subunit // 7.2E-5 IPR003869 // Polysaccharide biosynthesis protein CapD // 2.0E-4 IPR013525 // ABC-2 type transporter // 0.0018 IPR001509 // NAD-dependent epimerase/dehydratase // 3.2E-6 /// IPR002225 // 3-beta hydroxysteroid dehydrogenase/isomerase // 0.0021 IPR000277 // Cys/Met metabolism pyridoxal-phosphate-dependent enzymes // 0.0061 IPR000173 // Glyceraldehyde 3-phosphate dehydrogenase // 0.0098 IPR005242 // Conserved hypothetical protein 374 // 2.2E-5 IPR002314 // tRNA synthetase, class II (G, H, P and S) // 7.2E-4 IPR003358 // Putative methyltransferase // 0.0052 IPR001732 // UDP-glucose/GDP-mannose dehydrogenase // 0.0011 /// IPR001732 // UDP-glucose/GDP-mannose dehydrogenase // 0.0013 /// IPR006115 // 6-phosphogluconate dehydrogenase, NAD-binding // 3.6E-4 /// IPR006176 // 3-hydroxyacyl-CoA dehydrogenase, NAD-binding // 5.0E-5 /// IPR013332 // Ketopantoate reductase ApbA/PanE, N-terminal // 4.4E-5 IPR004360 // Glyoxalase/bleomycin resistance protein/dioxygenase // 0.018 IPR002213 // UDP-glucuronosyl/UDP-glucosyltransferase // 1.4E-4 /// IPR002213 // UDP-glucuronosyl/UDP-glucosyltransferase // 3.7E-4 IPR012302 // Malic enzyme, NAD-binding // 0.011 IPR013120 // Male sterility C-terminal // 1.5E-4 IPR003953 // Fumarate reductase/succinate dehydrogenase flavoprotein, N-terminal // 9.2E-9 IPR004792 // HI0933-like protein // 1.7E-5 IPR003395 // SMC protein, N-terminal // 2.6E-10 IPR013651 // RimK-like ATP-grasp // 9.9E-4 IPR006143 // Secretion protein HlyD // 0.0011
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IPR003437 // Glycine cleavage system P-protein // 0.0084 /// IPR003437 // Glycine cleavage system P-protein // 0.011 IPR002938 // Monooxygenase, FAD-binding // 1.7E-8 /// IPR003953 // Fumarate reductase/succinate dehydrogenase flavoprotein, N-terminal // 3.8E-6 IPR006935 // Type III restriction enzyme, res subunit // 2.8E-5 /// IPR006935 // Type III restriction enzyme, res subunit // 3.1E-5 IPR002086 // Aldehyde dehydrogenase // 1.0E-6 IPR002198 // Short-chain dehydrogenase/reductase SDR // 1.1E-4 /// IPR002198 // Short-chain dehydrogenase/reductase SDR // 1.3E-4 ///
IPR002225 // 3-beta hydroxysteroid dehydrogenase/isomerase // 5.5E-7 /// IPR002225 // 3-beta hydroxysteroid dehydrogenase/isomerase // 7.0E-7 /// IPR005913 // dTDP-4-dehydrorhamnose reductase // 1.6E-5 /// IPR005913 // dTDP-4-dehydrorhamnose reductase // 1.8E-5 /// IPR013120 // Male sterility C-terminal // 6.6E-4 /// IPR013120 // Male sterility C-terminal // 0.0013 IPR001279 // Beta-lactamase-like // 4.0E-9 IPR001761 // Periplasmic binding protein/LacI transcriptional regulator // 7.2E-7 /// IPR001761 // Periplasmic binding protein/LacI transcriptional regulator // 1.5E-6 IPR002225 // 3-beta hydroxysteroid dehydrogenase/isomerase // 2.6E-4 /// IPR005913 // dTDP-4-dehydrorhamnose reductase // 0.011 IPR000115 // Phosphoribosylglycinamide synthetase // 0.0053 IPR007152 // Protein of unknown function DUF354 // 0.069 IPR000522 // Bacterial transport system permease protein // 0.0059 /// IPR000522 // Bacterial transport system permease protein // 0.0046 IPR007050 // Bacterio-opsin activator, HTH // 0.0069 IPR003959 // AAA ATPase, central region // 6.2E-10 IPR000086 // NUDIX hydrolase // 0.049 IPR002562 // 3-5 exonuclease // 0.024 IPR004872 // NLPA lipoprotein // 0.0079 IPR002314 // tRNA synthetase, class II (G, H, P and S) // 0.012 IPR013221 // Mur ligase, middle region // 1.4E-5 IPR002308 // Cysteinyl-tRNA synthetase, class Ia // 2.8E-4 IPR005913 // dTDP-4-dehydrorhamnose reductase // 0.0024 /// IPR005913 // dTDP-4-dehydrorhamnose reductase // 8.0E-4 IPR000022 // Carboxyl transferase // 1.7E-11 IPR006151 // Shikimate/quinate 5-dehydrogenase // 0.0015 IPR004479 // ExsB // 0.0024 IPR010662 // Protein of unknown function DUF1234 // 2.2E-5 IPR013027 // FAD-dependent pyridine nucleotide-disulphide oxidoreductase // 1.4E-4 IPR008278 // 4-phosphopantetheinyl transferase // 0.0023 IPR004680 // Citrate transporter // 9.2E-7 IPR003402 // Protein of unknown function Met10 // 0.0025 /// IPR003402 // Protein of unknown function Met10 // 0.0018 IPR004680 // Citrate transporter // 1.3E-6 IPR010343 // Protein of unknown function DUF939, bacterial // 9.4E-23 IPR000415 // Nitroreductase // 7.9E-4 IPR001761 // Periplasmic binding protein/LacI transcriptional regulator // 0.0014 /// IPR001761 // Periplasmic binding protein/LacI transcriptional regulator // 6.5E-4 IPR003199 // Choloylglycine hydrolase // 1.3E-4
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IPR011545 // DEAD/DEAH box helicase, N-terminal // 8.2E-8 /// IPR011545 // DEAD/DEAH box helicase, N-terminal // 6.3E-8 IPR000914 // Bacterial extracellular solute-binding protein, family 5 // 4.6E-5 /// IPR000914 // Bacterial extracellular solute-binding protein, family 5 // 1.6E-13 /// IPR000914 // Bacterial extracellular solute-binding protein, family 5 // 2.3E-12 IPR013216 // Methyltransferase type 11 // 3.5E-6 IPR002933 // Peptidase M20 // 6.7E-4 /// IPR002933 // Peptidase M20 // 6.9E-4 IPR003135 // ATP-dependent carboxylate-amine ligase-like, ATP-grasp // 7.7E-4 IPR004104 // Oxidoreductase, C-terminal // 0.031 /// IPR004104 // Oxidoreductase, C-terminal // 0.029 /// IPR004104 // Oxidoreductase, C-terminal // 0.041 IPR011611 // PfkB // 9.6E-6 /// IPR011611 // PfkB // 2.8E-5 IPR013196 // Helix-turn-helix, type 11 // 0.0027 IPR003594 // ATP-binding region, ATPase-like // 9.8E-5 IPR002740 // Protein of unknown function DUF55 // 0.0036 IPR001509 // NAD-dependent epimerase/dehydratase // 0.0015 IPR003501 // Phosphotransferase system, lactose/cellobiose-specific IIB subunit // 7.1E-5 IPR011490 // Uncharacterised sugar-binding // 4.9E-6 /// IPR011490 // Uncharacterised sugar-binding // 5.5E-6 IPR013498 // DNA topoisomerase, type IA, zn finger // 0.014 IPR000835 // Bacterial regulatory protein, MarR // 0.0040 IPR002198 // Short-chain dehydrogenase/reductase SDR // 1.3E-4 IPR006092 // Acyl-CoA dehydrogenase, N-terminal // 3.7E-6 /// IPR006092 // Acyl-CoA dehydrogenase, N-terminal // 3.9E-5 /// IPR013107 // Acyl-CoA dehydrogenase, type 2, C-terminal // 3.1E-4 IPR004088 // KH, type 1 // 0.0011 IPR011547 // Sulphate transporter // 0.0053 IPR001433 // Oxidoreductase FAD/NAD(P)-binding // 1.2E-4 /// IPR001433 // Oxidoreductase FAD/NAD(P)-binding // 1.5E-4 IPR003675 // Abortive infection protein // 5.8E-4 IPR013108 // Amidohydrolase 3 // 2.2E-4 IPR003953 // Fumarate reductase/succinate dehydrogenase flavoprotein, N-terminal // 6.5E-4 IPR006076 // FAD dependent oxidoreductase // 1.0E-6 /// IPR013027 // FAD-dependent pyridine nucleotide-disulphide oxidoreductase // 4.5E-5 IPR003395 // SMC protein, N-terminal // 2.1E-5 /// IPR003395 // SMC protein, N-terminal // 3.7E-5 IPR001327 // Pyridine nucleotide-disulphide oxidoreductase, NAD-binding region // 3.1E-5 IPR001327 // Pyridine nucleotide-disulphide oxidoreductase, NAD-binding region // 3.5E- IPR006076 // FAD dependent oxidoreductase // 5.9E-4 /// IPR006076 // FAD dependent oxidoreductase // 3.5E-4 IPR002178 // Phosphotransferase system, phosphoenolpyruvate-dependent sugar EIIA 2 // 7.2E-5 /// IPR013196 // Helix-turn-helix, type 11 // 0.0043 /// IPR013196 // Helix-turn-helix, type 11 // 0.0058 IPR013525 // ABC-2 type transporter // 0.024 IPR000089 // Biotin/lipoyl attachment // 3.2E-4 /// IPR000089 // Biotin/lipoyl attachment // 1.7E-4 IPR001140 // ABC transporter, transmembrane region // 1.6E-6 ///
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IPR001140 // ABC transporter, transmembrane region // 1.4E-6 IPR000192 // Aminotransferase, class V // 2.3E-4 IPR005828 // General substrate transporter // 9.4E-5 IPR003869 // Polysaccharide biosynthesis protein CapD // 0.0072 IPR002012 // Gonadotropin-releasing hormone // 0.16 IPR005846 // Phosphoglucomutase/phosphomannomutase alpha/beta/alpha domain III // 0.28 IPR005846 // Phosphoglucomutase/phosphomannomutase alpha/beta/alpha domain III // 0.3 IPR004680 // Citrate transporter // 3.0E-4 IPR002731 // ATPase, BadF/BadG/BcrA/BcrD type // 1.1E-4 IPR013149 // Alcohol dehydrogenase, zinc-binding // 2.9E-8 IPR000524 // Bacterial regulatory protein GntR, HTH // 2.6E-9 IPR003140 // Phospholipase/Carboxylesterase // 4.2E-6 /// IPR003140 // Phospholipase/Carboxylesterase // 2.0E-6 IPR002831 // Transcriptional regulator TrmB // 0.0027 IPR000182 // GCN5-related N-acetyltransferase // 1.9E-6 /// IPR000182 // GCN5-related N-acetyltransferase // 2.1E-6 IPR006380 // Sucrose-6F-phosphate phosphohydrolase, plant and cyanobacteria // 1.4E-4 IPR013149 // Alcohol dehydrogenase, zinc-binding // 1.7E-7 IPR004669 // C4-dicarboxylate anaerobic carrier // 0.0032 /// IPR004669 // C4-dicarboxylate anaerobic carrier // 0.0033 IPR005097 // Saccharopine dehydrogenase // 4.4E-6 /// IPR005097 // Saccharopine dehydrogenase // 1.2E-5 /// IPR006151 // Shikimate/quinate 5-dehydrogenase // 4.0E-4 /// NULL // NULL // 2.6E-4 /// NULL // NULL // 2.7E-4 IPR001732 // UDP-glucose/GDP-mannose dehydrogenase // 5.2E-7 /// IPR003953 // Fumarate reductase/succinate dehydrogenase flavoprotein, N-terminal // 2.7E-5 IPR003953 // Fumarate reductase/succinate dehydrogenase flavoprotein, N-terminal // 2.0E-5 IPR004792 // HI0933-like protein // 2.1E-5 /// IPR013027 // FAD-dependent pyridine nucleotide-disulphide oxidoreductase // 1.3E-7 /// IPR013027 // FAD-dependent pyridine nucleotide-disulphide oxidoreductase // 4.3E-6 IPR001327 // Pyridine nucleotide-disulphide oxidoreductase, NAD-binding region // 5.8E-7 IPR002218 // Glucose-inhibited division protein A // 4.1E-6 /// IPR003953 // Fumarate reductase/succinate dehydrogenase flavoprotein, N-terminal // 1.2E-6 IPR004792 // HI0933-like protein // 1.1E-4 /// IPR013027 // FAD-dependent pyridine nucleotide-disulphide oxidoreductase // 3.4E-7 /// IPR013027 // FAD-dependent pyridine nucleotide-disulphide oxidoreductase // 4.2E-7 IPR008258 // Lytic transglycosylase, catalytic // 1.0E-10 IPR001509 // NAD-dependent epimerase/dehydratase // 9.3E-5 IPR006076 // FAD dependent oxidoreductase // 8.8E-5 IPR004360 // Glyoxalase/bleomycin resistance protein/dioxygenase // 4.3E-4 /// IPR004360 // Glyoxalase/bleomycin resistance protein/dioxygenase // 1.7E-4 /// IPR009725 // 3-demethylubiquinone-9 3-methyltransferase // 0.0018 /// IPR009725 // 3-demethylubiquinone-9 3-methyltransferase // 9.0E-4 IPR003352 // Phosphotransferase system, EIIC // 1.4E-4 /// IPR003352 // Phosphotransferase system, EIIC // 7.8E-7 IPR003018 // GAF // 6.0E-5 IPR000683 // Oxidoreductase, N-terminal // 0.0018 IPR001647 // Bacterial regulatory protein, TetR // 1.7E-4 IPR000801 // Putative esterase // 2.9E-5 IPR004841 // Amino acid permease-associated region // 1.1E-4 ///
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IPR004841 // Amino acid permease-associated region // 1.3E-4 IPR013525 // ABC-2 type transporter // 0.0094 IPR011116 // SecA Wing and Scaffold // 7.2E-6 /// IPR011116 // SecA Wing and Scaffold // 9.2E-6 IPR011128 // NAD-dependent glycerol-3-phosphate dehydrogenase, N-terminal // 3.6E-4 /// IPR011128 // NAD-dependent glycerol-3-phosphate dehydrogenase, N-terminal // 0.0013 IPR000741 // Fructose-bisphosphate aldolase, class-I // 7.8E-6 /// IPR000741 // Fructose-bisphosphate aldolase, class-I // 3.3E-6 IPR007197 // Radical SAM // 3.1E-7 IPR010656 // TRAP C4-dicarboxylate transport system permease DctM subunit // 5.5E-4 IPR000241 // Putative RNA methylase // 7.5E-9 IPR001258 // NHL repeat // 0.0059 /// IPR001258 // NHL repeat // 0.0059 IPR002208 // SecY protein // 1.5E-10 IPR003501 // Phosphotransferase system, lactose/cellobiose-specific IIB subunit // 4.3E-5 /// IPR007737 // M trans-acting positive regulator // 0.0079 IPR002018 // Carboxylesterase, type B // 0.01 IPR005097 // Saccharopine dehydrogenase // 1.5E-5 /// IPR005913 // dTDP-4-dehydrorhamnose reductase // 0.0061 /// IPR008030 // NmrA-like // 0.0037 /// IPR013120 // Male sterility C-terminal // 6.1E-6 IPR002938 // Monooxygenase, FAD-binding // 5.1E-7 /// IPR002938 // Monooxygenase, FAD-binding // 7.8E-7 /// IPR004792 // HI0933-like protein // 1.9E-5 /// IPR004792 // HI0933-like protein // 2.1E-5 /// IPR006076 // FAD dependent oxidoreductase // 8.1E-6 /// IPR006076 // FAD dependent oxidoreductase // 6.2E-6 IPR008286 // Orn/Lys/Arg decarboxylase, C-terminal // 6.2E-7 IPR011547 // Sulphate transporter // 0.0090 IPR002225 // 3-beta hydroxysteroid dehydrogenase/isomerase // 0.0025 /// IPR002225 // 3-beta hydroxysteroid dehydrogenase/isomerase // 0.0048 IPR006123 // Staphylococcal/Streptococcal toxin, beta-grasp // 0.0082 IPR006123 // Staphylococcal/Streptococcal toxin, beta-grasp // 0.13 /// IPR006123 // Staphylococcal/Streptococcal toxin, beta-grasp // 0.082 IPR007848 // Methyltransferase small // 7.2E-6 /// IPR007848 // Methyltransferase small // 5.4E-6 IPR003869 // Polysaccharide biosynthesis protein CapD // 1.1E-4 /// IPR013120 // Male sterility C-terminal // 7.9E-4 IPR006380 // Sucrose-6F-phosphate phosphohydrolase, plant and cyanobacteria // 0.0062 /// IPR006380 // Sucrose-6F-phosphate phosphohydrolase, plant and cyanobacteria // 0.0082 IPR002300 // Aminoacyl-tRNA synthetase, class Ia // 1.5E-10 /// IPR002300 // Aminoacyl-tRNA synthetase, class Ia // 1.1E-7 IPR006935 // Type III restriction enzyme, res subunit // 8.4E-6 IPR004268 // Virulence factor MVIN-like // 8.7E-4 /// IPR004268 // Virulence factor MVIN-like // 6.4E-4 IPR003265 // HhH-GPD // 0.03 /// IPR003265 // HhH-GPD // 0.04 IPR001466 // Beta-lactamase // 3.1E-4 /// IPR001466 // Beta-lactamase // 2.9E-4 IPR011611 // PfkB // 2.3E-4
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IPR004841 // Amino acid permease-associated region // 1.5E-4 /// IPR004841 // Amino acid permease-associated region // 1.2E-4 IPR004680 // Citrate transporter // 0.0074 /// IPR004680 // Citrate transporter // 0.0059 IPR013108 // Amidohydrolase 3 // 6.8E-5 IPR004685 // Branched-chain amino acid transport system II carrier protein // 0.0062 IPR002018 // Carboxylesterase, type B // 8.0E-4 IPR003661 // Histidine kinase A, N-terminal // 9.3E-4 IPR002218 // Glucose-inhibited division protein A // 1.4E-6 /// IPR002922 // Thiamine biosynthesis Thi4 protein // 2.0E-8 /// IPR002938 // Monooxygenase, FAD-binding // 7.6E-5 /// IPR003953 // Fumarate reductase/succinate dehydrogenase flavoprotein, N-terminal // 8.6E-8 IPR004792 // HI0933-like protein // 5.3E-7 /// IPR006076 // FAD dependent oxidoreductase // 6.6E-8 IPR010708 // 5 nucleotidase, deoxy, cytosolic type C // 3.2E-5 /// IPR010708 // 5 nucleotidase, deoxy, cytosolic type C // 3.6E-5 IPR006935 // Type III restriction enzyme, res subunit // 2.7E-6 IPR001650 // Helicase, C-terminal // 0.0023 IPR006674 // Metal-dependent phosphohydrolase, HD subdomain // 7.7E-4 IPR004680 // Citrate transporter // 6.8E-4 /// IPR004680 // Citrate transporter // 9.0E-4 /// IPR010656 // TRAP C4-dicarboxylate transport system permease DctM subunit // 0.0039 /// IPR010656 // TRAP C4-dicarboxylate transport system permease DctM subunit // 0.0063 IPR001597 // Aromatic amino acid beta-eliminating lyase/threonine aldolase // 3.1E-4 IPR001295 // Dihydroorotate dehydrogenase // 0.1 IPR002758 // Predicted cation antiporter // 1.7E-4 /// IPR002758 // Predicted cation antiporter // 9.9E-5 IPR011637 // DoxX // 0.0013 IPR011063 // PP-loop // 1.6E-4 IPR004245 // Protein of unknown function DUF229 // 4.3E-4 IPR000846 // Dihydrodipicolinate reductase // 1.8E-5 IPR002109 // Glutaredoxin // 2.9E-4 IPR013166 // Citrate lyase ligase, C-terminal // 1.4E-4 IPR001468 // Indole-3-glycerol phosphate synthase // 6.0E-6 IPR005025 // NADPH-dependent FMN reductase // 7.7E-4 IPR001962 // Asparagine synthase // 7.7E-4 /// IPR004506 // tRNA (5-methylaminomethyl-2-thiouridylate)-methyltransferase // 3.8E-11 /// IPR011063 // PP-loop // 3.9E-5 IPR000515 // Binding-protein-dependent transport systems inner membrane component // 0.0038 /// IPR000515 // Binding-protein-dependent transport systems inner membrane component // 0.0045 IPR002429 // Cytochrome c oxidase, subunit II // 2.2E-4 IPR003675 // Abortive infection protein // 0.013 /// IPR003675 // Abortive infection protein // 0.014 IPR013249 // Sigma-70, region 4 type 2 // 6.9E-5 IPR006992 // Amidohydrolase 2 // 9.7E-4 IPR000172 // Glucose-methanol-choline oxidoreductase // 8.7E-4 NULL // NULL // 2.4E-5 IPR002129 // Pyridoxal-dependent decarboxylase // 7.6E-4 ///
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IPR002129 // Pyridoxal-dependent decarboxylase // 4.0E-4 IPR005913 // dTDP-4-dehydrorhamnose reductase // 0.011 IPR000524 // Bacterial regulatory protein GntR, HTH // 0.0036 /// IPR000524 // Bacterial regulatory protein GntR, HTH // 0.0053 IPR005834 // Haloacid dehalogenase-like hydrolase // 4.2E-6 /// IPR013200 // Haloacid dehalogenase-like hydrolase, type 3 // 8.6E-7 IPR001327 // Pyridine nucleotide-disulphide oxidoreductase, NAD-binding region // 2.1E-4 IPR006076 // FAD dependent oxidoreductase // 4.0E-4 IPR013027 // FAD-dependent pyridine nucleotide-disulphide oxidoreductase // 6.3E-4 IPR000523 // Magnesium chelatase, ChlI subunit // 3.0E-4 /// IPR011703 // ATPase associated with various cellular activities, AAA-3 // 1.6E-5 IPR004835 // Fungal chitin synthase // 4.3E-4 IPR003953 // Fumarate reductase/succinate dehydrogenase flavoprotein, N-terminal // 2.0E-4 IPR013027 // FAD-dependent pyridine nucleotide-disulphide oxidoreductase // 3.8E-5 IPR000310 // Orn/Lys/Arg decarboxylase, major region // 0.23 /// IPR001597 // Aromatic amino acid beta-eliminating lyase/threonine aldolase // 9.9E-4 IPR003783 // Regulatory protein RecX // 0.0027 /// IPR003783 // Regulatory protein RecX // 0.0032 IPR004455 // NADP oxidoreductase, coenzyme F420-dependent // 8.0E-5 IPR002810 // Nodulation efficiency, NfeD // 2.0E-5 /// IPR002810 // Nodulation efficiency, NfeD // 3.9E-5 IPR013684 // Miro-like // 9.2E-5 IPR000005 // Helix-turn-helix, AraC type // 2.7E-4 /// IPR000005 // Helix-turn-helix, AraC type // 2.7E-4 IPR001296 // Glycosyl transferase, group 1 // 0.0097 IPR004479 // ExsB // 1.9E-4 /// IPR011063 // PP-loop // 0.13 IPR001509 // NAD-dependent epimerase/dehydratase // 0.0045 IPR000212 // UvrD/REP helicase // 7.3E-5 IPR006090 // Acyl-CoA dehydrogenase, type 1 // 5.2E-5 /// IPR013107 // Acyl-CoA dehydrogenase, type 2, C-terminal // 2.3E-5 IPR005828 // General substrate transporter // 1.0E-9 IPR011701 // Major facilitator superfamily MFS_1 // 4.8E-5 IPR001163 // Like-Sm ribonucleoprotein, core // 0.0020 IPR013746 // Hydroxymethylglutaryl-coenzyme A synthase C terminal // 5.6E-5 IPR000835 // Bacterial regulatory protein, MarR // 9.1E-5 IPR003333 // Cyclopropane-fatty-acyl-phospholipid synthase // 0.0040 /// IPR003333 // Cyclopropane-fatty-acyl-phospholipid synthase // 0.0040 /// IPR004033 // UbiE/COQ5 methyltransferase // 8.6E-4 IPR003358 // Putative methyltransferase // 0.014 IPR002904 // Lysyl-tRNA synthetase (archaeal), class 1c // 0.0099 /// IPR002904 // Lysyl-tRNA synthetase (archaeal), class 1c // 0.0099 IPR013105 // Tetratricopeptide TPR_2 // 0.0021 IPR001450 // 4Fe-4S ferredoxin, iron-sulfur binding // 0.0044 IPR000835 // Bacterial regulatory protein, MarR // 3.5E-6 /// IPR011611 // PfkB // 3.3E-7 /// IPR013749 // Phosphomethylpyrimidine kinase type-1 // 9.7E-5 IPR010289 // Protein of unknown function DUF893, YccS/YhfK // 0.0040 /// IPR010289 // Protein of unknown function DUF893, YccS/YhfK // 0.0043 IPR001899 // Surface protein from Gram-positive cocci, anchor region // 1.1E-4 /// IPR001899 // Surface protein from Gram-positive cocci, anchor region // 5.5E-5
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IPR001899 // Surface protein from Gram-positive cocci, anchor region // 1.1E-4 IPR001899 // Surface protein from Gram-positive cocci, anchor region // 1.1E-4 /// IPR001899 // Surface protein from Gram-positive cocci, anchor region // 5.5E-5 IPR001584 // Integrase, catalytic region // 4.6E-20 IPR001245 // Tyrosine protein kinase // 0.0048 IPR008258 // Lytic transglycosylase, catalytic // 8.9E-4 IPR002938 // Monooxygenase, FAD-binding // 2.9E-4 IPR006173 // Staphylococcal/Streptococcal toxin, OB-fold // 1.2E-4 IPR005828 // General substrate transporter // 1.7E-5 IPR011545 // DEAD/DEAH box helicase, N-terminal // 1.3E-7 IPR001584 // Integrase, catalytic region // 4.2E-14 IPR000999 // Ribonuclease III // 0.043 /// IPR002904 // Lysyl-tRNA synthetase (archaeal), class 1c // 0.0099 IPR003680 // Flavodoxin-like fold // 1.5E-5 IPR006444 // Phage major capsid protein, HK97 // 0.0024 IPR001899 // Surface protein from Gram-positive cocci, anchor region // 0.0013 IPR001279 // Beta-lactamase-like // 1.3E-6 IPR011545 // DEAD/DEAH box helicase, N-terminal // 1.3E-7 IPR003594 // ATP-binding region, ATPase-like // 2.1E-6 IPR006123 // Staphylococcal/Streptococcal toxin, beta-grasp // 0.1 IPR006123 // Staphylococcal/Streptococcal toxin, beta-grasp // 0.1 IPR005913 // dTDP-4-dehydrorhamnose reductase // 0.0024 /// IPR005913 // dTDP-4-dehydrorhamnose reductase // 8.0E-4 IPR001387 // Helix-turn-helix type 3 // 3.7E-5 IPR008160 // Collagen triple helix repeat // 5.6E-6 IPR005828 // General substrate transporter // 2.1E-4 IPR001584 // Integrase, catalytic region // 4.2E-14 IPR006123 // Staphylococcal/Streptococcal toxin, beta-grasp // 0.1 IPR003594 // ATP-binding region, ATPase-like // 2.1E-6 IPR004841 // Amino acid permease-associated region // 1.7E-5 IPR001254 // Peptidase S1 and S6, chymotrypsin/Hap // 2.7E-7 IPR013105 // Tetratricopeptide TPR_2 // 0.0021 IPR006090 // Acyl-CoA dehydrogenase, type 1 // 5.2E-5 /// IPR013107 // Acyl-CoA dehydrogenase, type 2, C-terminal // 2.3E-5 IPR005828 // General substrate transporter // 1.0E-9 IPR010289 // Protein of unknown function DUF893, YccS/YhfK // 0.0040 /// IPR010289 // Protein of unknown function DUF893, YccS/YhfK // 0.0043 IPR001245 // Tyrosine protein kinase // 1.4E-14 IPR001899 // Surface protein from Gram-positive cocci, anchor region // 5.5E-5 IPR001899 // Surface protein from Gram-positive cocci, anchor region // 1.1E-4 /// IPR001899 // Surface protein from Gram-positive cocci, anchor region // 5.5E-5 IPR000795 // Protein synthesis factor, GTP-binding // 3.1E-10 IPR001450 // 4Fe-4S ferredoxin, iron-sulfur binding // 0.0044 IPR003034 // DNA-binding SAP // 3.3E-5 IPR005828 // General substrate transporter // 1.7E-5 /// IPR011701 // Major facilitator superfamily MFS_1 // 1.1E-14 IPR011701 // Major facilitator superfamily MFS_1 // 4.5E-29 IPR010656 // TRAP C4-dicarboxylate transport system permease DctM subunit // 0.0058 /// IPR010656 // TRAP C4-dicarboxylate transport system permease DctM subunit // 0.0053 IPR003395 // SMC protein, N-terminal // 5.0E-5
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IPR005913 // dTDP-4-dehydrorhamnose reductase // 0.0024 /// IPR005913 // dTDP-4-dehydrorhamnose reductase // 8.0E-4 IPR001295 // Dihydroorotate dehydrogenase // 0.1 IPR002758 // Predicted cation antiporter // 1.7E-4 /// IPR002758 // Predicted cation antiporter // 9.9E-5 IPR004841 // Amino acid permease-associated region // 1.5E-4 /// IPR004841 // Amino acid permease-associated region // 1.2E-4
Summary of signals expressed on all 6 arrays
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13.3 Appendix C
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Research Committee approvals for the in situ project
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Smart Geelong Network, Researcher of the Year finalist, 2011
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13.4 Appendix D
Antibacterial efficacy results obtained by this project were confirmed by external, independent laboratory (Silliker, Australia).
Making this disinfectant acceptable to be registered by TGA in Australia as a Hospital Grade
Disinfectant, it was required to be tested for efficacy by TGA accredited laboratory..
Appendix D shows all certificates in details on external test results.
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Commercial Grade disinfectant test for Therapeutic Goods Administration approval
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Hospital Grade disinfectant test for Therapeutics Goods Administration approval
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AOAC Hard Surface Carrie test result
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