efficiency of industrial disinfectants on food-contact ... · com mc-06-dbes 2provou ser sinérgica...

Efficiency of industrial disinfectants

on food-contact surfaces sanitation

Masters Dissertation of

Joana Rodrigues Russo

Integrated Masters in Bioengineering

Branch of Biological Engineering

Supervisors: Manuel Simões (PhD)

Ana Meireles (Msc)

Porto, June 2016

“Live as if you were to die tomorrow. Learn as if you were to live forever”

Mahatma Gandhi

Efficiency of industrial disinfectants on food-contact surfaces sanitation | Joana Russo

Faculty of Engineering of the University of Porto | i

Acknowledgments

First of all, I would like to thank my supervisors, Doctor Manuel Simões and Ana

Catarina Meireles for all the support, dedication, guidance and availability always

given throughout this dissertation.

I would also like to thank the Chemical Engineering Department and LEPABE for

all the resources provided. In addition, I would like to thank Enkrott (more specifically

Doctor Ana Pereira) for providing DBNPA and Cleanity for providing the MC-06-DBES

disinfectant.

I would like to express a special thanks to everyone at lab E-101, especially to

Diana, Rita, João, Manel, Rita and Ana Luísa for all the support when “everything”

went wrong. It was a pleasure entertaining and supporting them as well during this

past semester.

My thanks also go to the researchers in Lab E007 and, of course, to Carla, Silvia

and Paula at E-103 for putting up with us and for always being patience and helpful.

I would like to thank as well all the friends that Bio and Chemical Engineering

brought me in the course of the last 6 years, especially Joana, Ana Rita, Joana, Ana,

Paula, Mariana, Raquel and Joana.

Last but not least, I would like to thank my family, specially my parents and

brother, for loving and supporting me unconditionally.


Faculty of Engineering of the University of Porto | iii

Abstract

Foodborne diseases are a growing public health problem spread worldwide.

Approximately 524 million foodborne diseases (90% of the total) result from

contaminations caused by microorganisms found mainly during processing and

distribution. Obviously, the importance of hygiene in food processing cannot be over-

emphasized and so sanitation programs must be implied in all the phases of food

processing. There is no recognized sanitation and disinfection programs able to

effectively control microbial presence in the food industry.

In this dissertation selected disinfectants, 2,2-dibromo-3-nitrilopropionamide

(DBNPA), sodium bicarbonate (SB) and an enzymatic-based disinfectant (MC-06-DBES),

were evaluated in the control of Escherichia coli in both planktonic and biofilm states.

The biofilm was formed during 1 day (coupons in 48-well microtiter plates) and 4 days

(coupons in an agitated reactor). Different materials were used as adhesion surfaces:

polystyrene (PS), polyvinyl chloride (PVC) and stainless steel (SS). The disinfectants

were tested individually and combined (DBNPA + SB and DBNPA + MC-06-DBES). For the

planktonic tests it was possible to define the minimal inhibitory concentration (MIC)

and minimal bactericidal concentration (MBC) for DBNPA as 20 ppm. The combinations

were defined as antagonic, since the MIC and MBC values were higher than the

individual values determined for DBNPA.

Regarding the 1 day old biofilm results, PVC surfaces were the most favorable for

biofilm formation (7 log CFU/cm2), and SS was the surface that allowed less biofilm

formation (6 log CFU/cm2). Accordingly, it is advisable the use of SS surfaces in the

food industry in detriment of PVC and PS surfaces. Concerning the individual action of

DBNPA and MC-06-DBES, in biofilm control, both eliminated 3 log CFU/cm2, but were

only able to remove 1 log cells/cm2. Additionally, SB was considered very inefficient

against E. coli biofilms as it removed and eliminated less than 3 log cells/cm2. On the

other hand, while the combination of DBNPA with SB showed no removal of biofilm but

promoted the elimination of 6 log CFU/cm2, the combination of DBNPA with MC-06-

DBES proved quite synergic since it eliminated 6 log CFU/cm2 and removed 7 log

cells/cm2.

When the removal and elimination of 4 day old biofilms on PVC coupons was tested,

the combination of DBNPA with MC-06-DBES caused 4 log CFU/cm2 reduction, proving

to be the most efficient combination tested. This combination of DBNPA with MC-06-

DBES was considered as the only one appropriate for future applications in the food


iv | Faculty of Engineering of the University of Porto

industry, as it was very effective in eliminating and removing 1 and 4 days old biofilm

(attested with topography and thickness analysis for the 4 days old biofilm).

Keywords: Biofilms; DBNPA; Disinfectants; Disinfectant combinations; Escherichia coli; Food

industry; Sodium bicarbonate.


Faculty of Engineering of the University of Porto | v

Resumo

As doenças transmitidas por alimentos são um problema de saúde pública que se

tem desenvolvido e espalhado mundialmente. Cerca de 524 milhões das doenças

transmitidas por alimentos (90% do total) são provocadas por contaminações

microbiológicas, maioritariamente durante as fases de processamento e distribuição.

Obviamente, a higiene tem um papel fundamental no processamento de alimentos e é

de elevada importância que programas de sanitização sejam implementados em todas

as fases de processamento alimentar. Infelizmente, ainda não está disponível nenhum

programa de sanitização e desinfeção que consiga controlar de maneira eficiente as

contaminações microbiológicas na indústria alimentar.

Nesta dissertação, foi avaliada a ação de diversos desinfetantes, 2-2-dibromo-3-

nitrilopropionamida (DBNPA), bicarbonato de sódio e um desinfetante de base

enzimática (MC-06-DBES) no controlo de Escherichia coli em estado planctónico e em

biofilme. O biofilme foi formado durante 1 dia (cupões em microplaca de 48 poços) e

4 dias (cupões em reator agitado). Diferentes materiais foram usados como superfícies

de adesão: poliestireno, policloreto de vinilo e aço inoxidável. Os desinfetantes foram

testados tanto individualmente como em combinação (DBNPA + Bicarbonato de sódio

e DBNPA + MC-06-DBES). Para os testes com células planctónicas foi possível definir a

concentração mínima de inibição (CMI) e a concentração mínima bactericida (CMB) do

DBNPA como 20 ppm. As combinações foram definidas como antagónicas, tendo em

conta que os valores de CMI e CMB eram mais elevados que os valores individuais

determinados para o DBNPA.

Relativamente ao biofilme formado nas placas de 48 poços, as superficies de

policloreto de vinilo foram as mais favoráveis para a formação de biofilme (7 log UFC

(unidades formadores de colónias) /cm2). Por outro lado, o aço foi a superfície que

possibilitou a formação de menor quantidade de biofilme (6 log UFC/cm2). Em

conseguinte, é aconselhável o uso de superfícies de aço na indústria alimentar em

detrimento das superfícies de poliestireno e de policloreto de vinilo. No que diz

respeito à ação individual de DBNPA e MC-06-DBES no controlo de biofilme, ambos

eliminaram 3 log UFC/cm2. No entanto, só foram capazes de remover 1 log

células/cm2. Adicionalmente, o bicarbonato de sódio foi considerado ineficiente no

controlo de E. coli, removendo e eliminando menos de 3 log células/cm2. Enquanto a

combinação de DBNPA com bicarbonato de sódio não demonstrou nenhuma remoção

de biofilme mas promoveu a eliminação de 6 log UFC/cm2, a combinação de DBNPA


vi | Faculty of Engineering of the University of Porto

com MC-06-DBES provou ser sinérgica com a eliminação de 6 log UFC/cm2 e a remoção

de 7 log células/cm2.

Quanto à remoção e eliminação de biofilme de 4 dias formado em cupões de

policloreto de vinilo, a combinação de DBNPA com MC-06-DBES promoveu a redução de

4 log UFC/cm2, provando ser a combinação mais eficiente. A combinação de DBNPA

com MC-06-DBES foi considerada como a única apropriada para futuras aplicações na

indústria alimentar, visto ter sido muito eficiente tanto na remoção como eliminação

de biofilmes de 1 e 4 dias (comprovado com análise da topografia e da espessura do

biofilme de 4 dias)

Palavras-chave: Bicarbonato de sódio; Biofilmes; Combinação de desinfetantes; DBNPA;

Desinfetantes; Escherichia coli; Indústria Alimentar.


Faculty of Engineering of the University of Porto | vii

Table of Contents

Acknowledgments ................................................................................. i

Abstract ........................................................................................... iii

Resumo ............................................................................................ v

List of Figures .................................................................................... xi

List of Tables ................................................................................... xiii

List of Abbreviations ............................................................................ xv

1. Work Outline ................................................................................... 1

1.1. Background and project presentation ................................................ 1

1.2. Thesis organization ...................................................................... 2

2. Literature Review ............................................................................. 3

2.1. Food Industry Problems ................................................................. 3

2.2. Sanitation ................................................................................. 4

2.2.1. Cleaning and Disinfection .......................................................... 4

2.2.1.1. Types of Disinfectants ........................................................ 5

2.2.1.1.1. Chlorine Releasing Compounds ...................................... 7

2.2.1.1.2. Iodine .................................................................... 8

2.2.1.1.3. Hydrogen Peroxide ..................................................... 9

2.2.1.1.4. Peracetic Acid .......................................................... 9

2.2.1.1.5. Ozone .................................................................... 9

2.2.1.1.6. Quaternary Ammonium Compounds ................................ 10

2.2.1.1.7. Acid Anionic Surfactants ............................................. 11

2.2.1.2. Disinfectants Addressed in this Thesis ..................................... 12

2.2.1.2.1. DBNPA .................................................................. 12

2.2.1.2.2. Sodium Bicarbonate .................................................. 13

2.2.1.2.3. MC-06-DBES ............................................................ 14

2.2.1.3. Factors Affecting Disinfectant Effectiveness ............................. 15


viii | Faculty of Engineering of the University of Porto

2.2.1.4. Effectiveness of Disinfectants Against Different Organisms on Food

Contact Surfaces ........................................................................ 16

2.2.1.4.1. Bacillus cereus ........................................................ 16

2.2.1.4.2. Cronobacter sakazakii ................................................ 17

2.2.1.4.3. Enterobacter sakazakii ............................................... 17

2.2.1.4.4. Escherichia coli ....................................................... 18

2.2.1.4.5. Listeria monocytogenes .............................................. 19

2.2.1.4.6. Salmonella enteritidis ............................................... 19

2.2.1.4.7. Staphylococcus aureus ............................................... 19

2.2.1.5. Biofilms and Reasons for the Ineffectiveness of Disinfectants Against

Biofilms ................................................................................... 22

3. Behavior of E. coli planktonic cells in the presence of DBNPA, sodium bicarbonate

and the enzymatic disinfectant MC ........................................................... 23

3.1. Introduction ............................................................................. 23

3.2. Materials and Methods ................................................................. 23

3.2.1. Microorganism and culture conditions .......................................... 23

3.2.2. Disinfectants ........................................................................ 24

3.2.3. Determination of minimum inhibitory concentration for DBNPA ........... 24

3.2.4. Determination of MIC for DBNPA combined with SB and MC ................. 25

3.2.5. Determination of minimum bactericidal concentration for DBNPA ......... 25

3.2.5.1. MBC in the presence of MHB ................................................ 25

3.2.5.2. MBC in the absence of MHB ................................................. 25

3.2.6. Statistical Analysis ................................................................. 26

3.3. Results and Discussion .................................................................. 26

3.3.1. Determination of the MIC for DBNPA ............................................ 26

3.3.2. Determination of the MBC for DBNPA ........................................... 26

3.3.3. Determination of the synergetic activity of DBNPA with SB and DBNPA with

MC ............................................................................................ 27

3.4. Conclusions .............................................................................. 28


Faculty of Engineering of the University of Porto | ix

4. Control of E. coli biofilms in different surfaces ......................................... 29

4.1. Introduction ............................................................................. 29

4.2. Materials and Methods ................................................................. 29

4.2.1. Microorganism and culture conditions .......................................... 29

4.2.2. Disinfectants ........................................................................ 29

4.2.3. Biofilm formation in microtiter plates .......................................... 30

4.2.3.1. Biofilm control with DBNPA, SB, MC and the combination.............. 30

4.2.3.2. Biofilm mass quantification by crystal violet ............................. 30

4.2.3.3. Biofilm cells culturability .................................................... 30

4.2.4. Biofilm formation on coupons in 48-well microtiter plates .................. 31

4.2.4.1. Biofilm control with DBNPA, SB, MC and the combination.............. 31

4.2.4.2. Effect of DBNPA, SB, MC and the combination in biofilm bacteria

removal ................................................................................... 31

4.2.4.3. Effect of DBNPA, SB, MC and the combination in biofilm cells

culturability .............................................................................. 31

4.2.4.4. Regrowth of biofilm cells after exposure to DBNPA, SB, MC and the

combination .............................................................................. 32

4.2.5. Biofilm formation on coupons in an agitated reactor ........................ 32

4.2.5.1. Biofilm control with DBNPA, SB and MC and the combination ......... 32


culturability .............................................................................. 32

4.2.5.3. Effect of DBNPA, SB, MC and the combination in biofilm thickness ... 33

4.2.5.4. Observation of biofilm images in Optical Coherence Tomography (OCT)

............................................................................................. 33

4.2.6. Statistical Analysis ................................................................. 33

4.3. Results and Discussion .................................................................. 33

4.3.1. Efficiency of the elimination and removal of biofilm in microtiter plates 33

4.3.2. Efficiency of the elimination and removal of biofilm formed on coupons in

48-well microtiter plates ................................................................. 35


x | Faculty of Engineering of the University of Porto

4.3.3. Efficiency of the elimination and removal of biofilm formed on coupons in

an agitated reactor ........................................................................ 38

4.3.4. Topography of the biofilm formed on coupons in an agitate reactor ...... 40

4.3.5. Visualization of the biofilm formed on coupons in an agitated reactor ... 42

4.4. Conclusions .............................................................................. 43

5. Concluding remarks and future work ..................................................... 45

5.1. General Conclusions .................................................................... 45

5.2. Future Work ............................................................................. 46

References ....................................................................................... 47

Appendix........................................................................................... 1


Faculty of Engineering of the University of Porto | xi

List of Figures

In the Main Text:

Figure 1. Decomposition pathways of DBNPA (based on Exner et al., 1973). .......... 13

Figure 2. Graphic representation of the evolution of log CFUs with the increasing

concentrations of DBNPA. The line indicates the method detection limit (2 log

CFU/mL). ......................................................................................... 27

Figure 3. Evaluation of the combination of DBNPA with SB and MC through O.D.

variation analysis. ............................................................................... 28

Figure 4. Quantification of E. coli biofilm mass by CV assay. ............................ 34

Figure 5. Log CFU/cm2 reduction of biofilm cells formed in 96-well microtiter plates

for the individual disinfectants and for their combinations with DBNPA. .............. 35

Figure 6. Log cells/cm2 removal for biofilms formed on PS, SS and PVC coupons in 48-

well plates for the individual disinfectants and for the combinations with DBNPA (tests

with DAPI). The line indicates the method detection limit (4.22 log cells/cm2). The

symbol * represents that no cells were detected. .......................................... 35

Figure 7. Microscopic visualization of biofilm cells on PVC without treatment (a),

treated with DBNPA at 10 ppm and SB at 37 800 ppm (b), treated with DBNPA at 20

ppm (c) and SB at 75 600 ppm (d). Magnification x1000 and scale bar 10 µm. ........ 36

Figure 8. Log CFU/cm2 reduction for biofilms formed on PS, SS and PVC coupons in 48-

well plates for the individual disinfectants and for the combinations with DBNPA (CFU

enumeration). The line indicates the method detection limit (2.66 log CFU/cm2). The

symbol * represents that no CFU was detected. ............................................ 38

Figure 9. Log CFU/cm2 for biofilms formed on PVC coupons in an agitated reactor for

the individual disinfectants and for the combinations with DBNPA (CFU enumeration).

The line indicates the method detection limit (2.66 log CFU/cm2....................... 39

Figure 10. Thickness of biofilm formed on PVC coupons in an agitated reactor evaluated

by microscopy. .................................................................................. 41

Figure 11. Microscopic visualization of the biofilm topography in the E. coli control

coupon, without treatment. ................................................................... 41

Figure 12. Microscopic visualization of the E. coli biofilm topography when treated with

10 ppm DBNPA and 25 000 ppm MC. .......................................................... 41


xii | Faculty of Engineering of the University of Porto

Figure 13. Macroscopic observation of the biofilm formed on the PVC coupons (the one

on the left was visualized in OCT). ........................................................... 42

Figure 14. Visualization in the OCT of biofilm formed on the PVC coupons. The three

representations are different biofilms samples on the same coupon visualized under

different angles. The different color is simply to enhance contrast. ................... 42

In the Appendix

Figure A1. Microscopic visualization of MC residues after biofilm cells on PVC were

treated with MC at 75 600 ppm (a) and DBNPA at 10 ppm + MC at 37 800 ppm (b).

Magnification x1000 and scale bar 10 µm. .................................................. A1

Figure A2. Microscopic visualization of biofilm cells on PS without treatment (a),


ppm (c) and SB at 75 600 ppm (d). Magnification x1000 and scale bar 10 µm. ....... A1

Figure A3. Microscopic visualization of biofilm cells on SS without treatment (a),


ppm (c) and SB at 75 600 ppm (d). Magnification x1000 and scale bar 10 µm. ....... A2

Figure A4. Pump calibration curve. ......................................................... A3

Figure A5. Agitated reactor installation.1) feed; 2) waste residues; 3) reactor with

agitation; 4) feed pump; and 5) aeration pump. .......................................... A4


Faculty of Engineering of the University of Porto | xiii

List of Tables

In the Main Text:

Table 1. Advantages and limitations of the disinfectants most used in the food industry

...................................................................................................... 6

Table 2. Disinfection methods applied in the food industry on food contact surfaces20

Table 3. Neutralizer solution composition ................................................... 24


Faculty of Engineering of the University of Porto | xv

List of Abbreviations

2D Two dimensional

3D Three dimensional

AISI American Iron and Steel Institute

BDA Bis-(3-Aminopropyl)-Dodecylamine

CECT Spanish Type Culture Collection

CIP Clean in Place

CFU Colony Forming Units

CV Crystal Violet

DAPI 4’,6-Diamidino-2-Phenylindole

DBNPA 2,2-Dibromo-3-Nitrilopropionamide

DDAC Didecyldimethylammonium chloride

DSM German Collection of Microorganisms

EN European Standards

EOW Electrolyzed Oxidizing Water

EPS Extracellular Polymeric Substances

FAO Food and Agriculture Organization of the United Nations

FDA Food and Drug Administration

GL2A 2-O-palmitoyl-3-O-(6'-sulfoquinovopyranosyl)-glycerol

GRAS Generally Recognized As Safe

MBC Minimum Bactericidal Concentration

MHB Mueller Hinton Broth

MIC Minimum Inhibitory Concentration

OCT Optical Coherence Tomography

http://www.uv.es/uvweb/spanish-type-culture-collection/en/spanish-type-culture-collection-1285872233521.html


xvi | Faculty of Engineering of the University of Porto

OD Optical Density

PAA Peracetic Acid

PC Polycarbonate

PCA Plate Count Agar

PP Polypropylene

PS Polystyrene

PVC Polyvinyl Chloride

QAC Quaternary Ammonium Compounds

SB Sodium Bicarbonate

SH Sodium Hypochlorite

SS Stainless Steel

UK United Kingdom

US Ultrasound

USA United States of America

UV Ultra - Violet

WHO World Health Organization


Faculty of Engineering of the University of Porto | 1

Chapter 1

1. Work Outline

1.1. Background and project presentation

There is a global concern in the food industry that all world population should have

access to microbiological safe food. Unfortunately, the prevention of foodborne

diseases is a process without a universal solution and for the majority of pathogens no

vaccines have been developed (Tauxe, 1997). As a result of poor hygiene practices in

food processing plants, foodborne illness outbreaks and food spoilage can be a

consequence. Microbiological food spoilage may occur due to the production of

enzymes by the microorganisms that lead to unacceptable by-products in the food

(Benner, 2014). Moreover, the contamination of food products with pathogens may

arise, carrying serious risks for the health of consumers (Gutierrez et al., 2012).

Foodborne diseases are a significant cause of morbidity and mortality, especially

among children and the elderly and represent a significant impairment to social

development worldwide (WHO, 2015b). These problems are also an impediment to

economic development, since the food industry loses many millions of dollars annually

due to the presence of unacceptably high numbers of microorganisms, both spoilage

or pathogenic (Brooks and Flint, 2008). However, the complete eradication of these

pathogens from food processing plants is a difficult task. The major cause is the

bacterial attachment to the industrial surfaces leading to the formation of biofilms,

that increase microbial resistance to sanitation (Yang et al., 2012). In addition to the

creation of problems associated with public health and product spoilage, biofilms are

also responsible for: i) mechanical problems; ii) hampering the heat transfer processes;

iii) increasing the corrosion rate of surfaces; iv) increasing the fluid frictional

resistance (Chmielewski and Frank, 2003, Mogha et al., 2014).

Scientists and food technologists have been focusing on the development of new

procedures/techniques that are more sensitive, accurate, rapid, and cost-effective to

eliminate microbial risks in order to reduce waste and promote safer, healthier foods

(Böhme et al., 2014, Todd, 2014). Consequently, the main purposes of this thesis were

to study E. coli elimination and removal in the planktonic and sessile states (1 and 4

days old biofilm), using three different disinfectants (DBNPA, SB and MC-06-DBES)


2 | Faculty of Engineering of the University of Porto

individually and combined (DBNPA + SB and DBNPA + MC-06-DBES). Disinfection from

different surfaces materials (PVC, PS and SS) was also evaluated.

1.2. Thesis organization

This work is organized in five chapters. The present one, Chapter 1, presents the

motivations and the objectives of the overall project.

Chapter 2 provides a brief literature review about the current state of the

problematic associated to the presence of biofilms and pathogenic microorganism in

the food industry. This chapter also comprises an overview of the several disinfectants

used in the food industry as well as the reasons and factors that can affect its

efficiency.

Chapter 3 is the first chapter with experimental work and comprises the study of

E. coli in planktonic state. Furthermore, the antimicrobials tested are DBNPA, sodium

bicarbonate and MC-06-DBES, a formulation based in anionic and non-ionic surfactants,

with enzymes and essential oils. Their combined activity was also tested in this

chapter.

Chapter 4 shows the behavior of 24 h old biofilms in 96-well plates and on coupons

in 48-well plates, as well as 4 d old biofilms formed on coupons in an agitated reactor,

when exposed to the disinfectants mentioned above.

Finally, Chapter 5 describes the general conclusions from this study and provides

some suggestions for future work.



Chapter 2

2. Literature Review

2.1. Food Industry Problems

Foodborne diseases are usually infectious and frequently caused by either bacteria

(e.g. Salmonella or Listeria), viruses (e.g. norovirus), parasites (e.g. Toxoplasma

gondii or Cryptosporidium), or chemical substances through contaminated food or

water (WHO, 2015a). According to the World Health Organization (WHO), in 2010, 582

million cases of foodborne diseases had been reported, of which 351 000 represent

deaths (WHO, 2015c). It is estimated that about 90% of the foodborne diseases are

caused by microorganisms (CUF, 2013). The contamination of food may occur at any

stage in the process from food production to consumption (“farm to fork”) and can be

a result of environmental contamination, including the pollution of water, soil or air

(WHO, 2015a). The primary concern of food manufacturers should be to produce a

product that is healthy and safe, i.e. free from pathogenic microorganisms and

chemical and foreign body contamination (Holah, 2014b). Moreover, food companies

should also take into account that the contamination of food can result in serious

commercial setbacks if they do not enforce strict sanitary conditions (Berk, 2013).

Keeping in mind that contamination can arise “from farm to fork” it is vital that

hygiene considerations are implied in all the phases of food processing: from the way

the farmer handles the raw materials, passing through the equipment selection and

training of all personnel and ending at the consumer (Berk, 2013). Fresh produce, such

as lettuce or spinach, require very little processing. Consequently, they are not

submitted to any effective microbial elimination step. Consequently, food industry can

release into the market food products that may transport pathogenic or spoilage

microorganisms (Harris et al., 2003). When it comes to the preparation of the produce,

water disinfection is one of the most critical processing steps (Gil et al., 2015).



2.2. Sanitation

Food contact surfaces, according to the FDA (1998), refer to surfaces that contact

with fresh produce, and also that contact with drainage onto the produce or onto

surfaces that will afterwards contact with the fresh produce. These may include

equipment, such as conveyor belts and containers used along the food chain in

harvesting, post-harvesting and packaging. Sanitizing food contact surfaces is an

essential task in food processing plants, as pathogens and food spoilage bacteria can

spread to food contact surfaces during preparation of raw products. Since the

pathogens can be transmitted, the possibility of contamination of food materials can

lead to food safety issues (Kusumaningrum et al., 2003).

In the food industry, surfaces contain debris (food and dirt particles) which can

harbor microorganisms and potentiate biofilm formation (Chmielewski and Frank,

2003). Microorganisms tend to attach to wet surfaces, multiply and then produce a

matrix of extracellular material (produced by the organisms themselves) commonly

known as extracellular polymeric substances (EPS), forming biofilms (Flemming and

Wingender, 2010). Microorganisms in biofilms can reduce product quality and limit the

shelf-life of products, in a number of food industry sectors such as fresh produce, dairy,

poultry and red meat processing (Simões et al., 2010). However, despite affecting

product quality and shelf-life, the most serious problem associated with biofilms is

that they represent a safety threat (Grinstead, 2009).

Sanitation is a process with two main steps: cleaning and disinfection (Holah,

2009). The main objective of sanitation is to reduce undesirable material (food

residues, microorganisms, etc.) to levels considered safe, based on established

parameters and in an economical manner (Holah, 2009). However, sanitation must be

done in a way that it does not interfere with the quality and safety of the product.

The problem, though, relies on the fact that a number of sanitation strategies are not

efficient enough to provide a complete eradication of hazardous microorganisms

without affecting product quality (Srey et al., 2013).

2.2.1. Cleaning and Disinfection

Cleaning is the first step in sanitation and enables the removal of all physical

contaminants such as debris, organic substances (e.g. proteins) and inorganic materials

(e.g. hard water deposits) from walls, floors, tools and the surface of equipment (Berk,

2013). In order to prevent the contamination of food products and the formation of



biofilms, regular cleaning is necessary. However, cleaning does not eliminate the

bacteria, as it only allows the removal of the bacteria (approximately 90%) from the

surfaces. This may promote the later re-attaching to other surfaces and formation of

biofilm, thus disinfection is indispensable in order to eradicate them (Gram et al.,

2007). Cleaning can be achieved using mechanical or kinetic energy by: i) physically

removing soil (resorting to brushes); ii) using thermal energy by raising the

temperature to aid the removal of fats by promoting emulsification; iii) or using

chemical energy by applying detergents (Holah, 2014a).

Regarding disinfection, it allows the removal and/or reduction of the viability of

the microorganisms, which remained after cleaning, to a level at which they represent

no significant risk. This process occurs by means of chemical agents – disinfectants –

which should not affect human health, by leaving hazardous residues, and should not

cause corrosion of equipment (Holah, 2009). Furthermore, disinfection will only be

successful if the surfaces have been thoroughly cleaned to remove interfering matters,

unless disinfection is done by heat (Holah, 1995a).

2.2.1.1. Types of Disinfectants

That is no such thing as a perfect disinfectant, therefore it is important to

understand the strengths and limitations of sanitizers and disinfectants in order to

understand how they can be used to help preventing foodborne diseases (Schmidt,

2009). There are a few characteristics that a disinfectant should have in order to be

considered as a good food industry disinfectant: i) it must be suited for food contact

surfaces application; ii) it should have a wide range of activity; iii) it should be able to

eliminate microorganisms in the shortest time possible (Schmidt, 2009); iv) it should

have low toxicity and corrosivity (toxic residues could affect the health and sensory

properties of the products) (Holah, 1995b); v) it should be non-tainting; vi) and it

should not form any toxic by-products (Ruehlen and Williams, 2014). Moreover, an

ideal sanitizer must be stable when purchased (concentrated) and when used (diluted),

and should also be inexpensive (Marriott and Robertson, 1997).

There are many different biocides that are used in food processing. The most

commonly used biocides classes are halogens, namely chlorine, chlorine dioxide and

iodine; and peroxides, such as hydrogen peroxide and peracetic acid (Grinstead, 2009).

In Table 1 a summary of the advantages and limitations of the biocides described

hereafter.



Table

1.

Advanta

ges

and l

imit

ati

ons

of

the d

isin

fecta

nts

most

use

d in t

he f

ood indust

ry

Lim

itati

ons

Corr

osi

ve t

o m

any m

eta

ls

Rela

tively

inst

able

Acti

vit

y d

ecre

ase

s ra

pid

ly i

n t

he p

rese

nce o

f org

anic

matt

er

Can d

iscolo

r su

rfaces

Slo

w a

cti

vit

y a

t pH

7 o

r above

Must

be u

sed a

te h

igh c

oncentr

ati

ons

Can

be

inacti

vate

d

by

the

mic

roorg

anis

ms’

ow

n

enzy

mes

(cata

lase

and s

upero

xid

e d

ism

uta

se)

Expensi

ve

Unst

able

under

alk

aline p

H

Pungent

aceti

c a

cid

odor

Unst

able

Expensi

ve

Reacts

quic

kly

wit

h o

rganic

matt

er

Must

be g

enera

ted a

t poin

t of

use

Acti

ve o

ver

a w

ide p

H r

ange

Form

s bacte

riost

ati

c f

ilm

Not

com

pati

ble

wit

h s

om

e d

ete

rgents

and h

ard

wate

r

Not

eff

ecti

ve a

gain

st s

pore

s

Expensi

ve

Genera

tes

foam

Slo

w a

cti

vit

y a

gain

st s

pore

-form

ing o

rganis

ms

Eff

ecti

ve a

t acid

pH

only

Advanta

ges

Str

ong b

road s

pectr

um

bio

cid

e

Rela

tively

inexpensi

ve

Not

aff

ecte

d b

y h

ard

wate

r sa

lts

Str

ong s

pectr

um

of

acti

vit

y

Rela

tively

sta

ble

Unaff

ecte

d b

y w

ate

r hard

ness

Non-c

orr

osi

ve

Low

toxic

ity b

iocid

e a

t hig

h c

oncentr

ati

ons

Hig

h e

ffic

iency

Low

toxic

ity b

iocid

e

Eff

ecti

ve a

t lo

w c

oncentr

ati

ons

Bro

ad s

pectr

um

anti

mic

robia

l acti

vit

y

Bro

ad r

ange o

f anti

mic

robia

l acti

vit

y

Low

by-p

roduct

genera

tion

Non-t

oxic

and n

on-c

orr

osi

ve

Rela

tively

st

able

in

th

e

pre

sence

of

org

anic

matt

er

Eff

ecti

ve again

st

a w

ide ra

nge of

vegeta

tive

org

anis

ms

Eff

ecti

ve a

gain

st a

wid

e r

ange o

f org

anis

ms

Rem

oves

and p

revents

the f

orm

ati

on o

f deposi

ts

Dis

infe

cta

nts

Chlo

rine r

ele

asi

ng

com

pounds

Iodin

e

Hydro

gen

pero

xid

e

Pera

ceti

c a

cid

Ozo

ne

QACs

Acid

-anio

nic

surf

acta

nts



2.2.1.1.1. Chlorine Releasing Compounds

Chlorine – in its most varied forms – is one of the most used sanitizers in the food

industry (Schmidt, 2009). It is most frequently used after thermal processing, to

sanitize the water that is used to cool the tanks (Berk, 2013). Chlorine is usually used

in the form of hypochlorite or hypochlorous acid (weak acid that dissociates into

hypochlorite ion) (Byrd and McKee, 2005). When used at high concentrations, chlorine

compounds are considered germicidal. They are also active at low temperatures, are

fairly inexpensive and leave no residues (or insignificant residues) on the surfaces

(Parkinson, 2012). Despite having a very strong biocidal activity, hypochlorous acid is

unstable, so it is necessary to be produced in situ. Hypochlorous acid also has the

disadvantage of being easily inactivated by various metals (e.g. copper) and UV light,

and also reacting with organic matter (Grinstead, 2009). Since chlorine has the

disadvantage of reacting with organic matter present in the process line, it is highly

recommended that all the organic impurities are washed before using this disinfectant

(Berk, 2013). Another disadvantage of chlorine compounds is the fact that they are

corrosive to metal surfaces such as SS, mainly at high temperatures, with possibility of

causing pitting if exposed for a prolonged time (Schmidt, 2009).

Regarding the mechanism of action, chlorine compounds target the cell membrane,

inhibiting cellular enzymes involved in glucose metabolism (Gray et al., 2013).

Moreover, these biocides have a lethal effect on DNA (inhibiting its synthesis) and, as

oxidizing agents, destroy the cellular activity of proteins (McDonnell and Russell,

1999). Sodium hypochlorite (SH) is another hypochlorite that can also be used and has

been reported to be an effective disinfectant for biofilm inactivation (Silva et al.,

2011). Sodium hypochlorite was also stated to be a potential biofilm antimicrobial

agent against Staphylococcus aureus, Prevotella intermedia, Peptostreptococcus

miros, Streptococcus intermedius, Fusobacterium nucleatum and Enterococcus

faecalis (Srey et al., 2013).

Another example is chlorine dioxide. This gaseous compound is known for its strong

antimicrobial activity (including sporicidal) (Vandekinderen et al., 2009). However, it

can also be used in the liquid form (Choi et al., 2015). Aqueous chlorine dioxide has

been applied in the disinfection of vegetables (Gómez-López et al., 2009) and has

proved its antibrowning efficacy during storage of fresh-cut vegetables and fruits (Du

et al., 2009). This disinfectant is also used to treat water involved in food processing

and to sanitize surfaces, although less frequently (Grinstead, 2009). The advantage on

using chlorine dioxide is that, when it is used as a gas, it has almost 3 times the



oxidizing power of chlorine and it does not produce by-products harmful for the

environment, as chlorine does (Martín-Belloso et al., 2012).

One of the disadvantages associated with gaseous chlorine dioxide is the fact that

this compound is a dissolved gas which limits its use in the sanitation of surfaces.

Additionally, owing to its gaseous nature and the fact that it decomposes rapidly in the

presence of light, it is advisable that it is produced in situ (Grinstead, 2009). This

chemical is an extremely selective oxidant, due to its one-electron transfer

mechanism, through which it attacks the electrons in organic molecules, promoting

the reduction of chlorine dioxide to chlorite ion (Gordon and Rosenblatt, 2005).

Chlorine dioxide is a highly active oxidizing agent and the mechanism of action

comprises the destruction of the cellular activity of proteins in the membrane of the

cell surface (McDonnell and Russell, 1999) through the oxidation of peptide links and

denaturation of proteins (Maris, 1995).

2.2.1.1.2. Iodine

Iodine is less reactive than chlorine, however, it has a very broad spectrum of

activity against bacteria, fungi, virus, spores and even tuberculosis bacteria (Gottardi,

2001). Iodine can be used as a disinfectant in many forms such as aqueous solutions of

iodine, solutions of iodine in alcohol and iodophors (Grinstead, 2009). Iodophors are

complexes of iodine with a solubilizing agent, which acts as a carrier of the active

“free” iodine (McDonnell and Russell, 1999). One big advantage of iodophors when

compared to chlorine is the fact that iodophors are less affected by organic matter

and water hardness (Parkinson, 2012). One downside when compared to iodine is the

fact that iodophors are considered to have a lower activity against certain fungi and

spores, even though germicidal activity is maintained (Selvaggi et al., 2003).

The main disadvantages of iodine compounds are: i) the staining (especially on

plastic); ii) the loss of activity at high pH; and iii) the limitation of the temperature at

which it can be used, due to the fact that it vaporizes at 50 °C (Schmidt, 2009). The

coloration limits significantly the use of iodine in the food industry. However, if a

iodophor such as povidone-iodine is used then the staining is less pronounced (Selvaggi

et al., 2003). Concerning the mechanism of action, iodine penetrates rapidly into the

microorganisms, interfering at the respiratory chain, hampering the transport of

electrons through the chain (Russell, 2001b). It is also considered that the lethal

effects of iodine come from the disruption of the structure of nucleic acids and the

impairment of its synthesis (Rutala and Weber, 2008).



2.2.1.1.3. Hydrogen Peroxide

Hydrogen peroxide is widely used as a disinfectant, especially in applications where

the non-toxicity of its degradation by-products is fundamental, which is the case of

the food industry (Linley et al., 2012). This biocide is known as a clear, colorless liquid,

environmentally friendly, with low toxicity (McDonnell and Russell, 1999). Hydrogen

peroxide is effective in a broad spectrum (even bacterial spores). However, for a

sporicidal activity, higher concentrations (up to 30%) and longer exposure times are

required (Russell, 2001a).

One of the weaknesses of hydrogen peroxide is that it is a natural by-product of

aerobic metabolism and consequently can be inactivated by enzymes – catalase or

other peroxidases – that most organisms have (McDonnell and Russell, 1999).

Regarding the mechanism of action, hydrogen peroxide acts through chemical

oxidation of cellular components (Finnegan et al., 2010), through the production of

free hydroxyl radicals which attack essential cell components, including membrane

lipids, proteins and DNA (Block, 2001, Linley et al., 2012) .

2.2.1.1.4. Peracetic Acid

Peracetic acid (PAA) is a biocide frequently used in the food industry that has many

of the advantages of hydrogen peroxide, but is not enzymatic inactivated by

peroxidases (Block, 2001). In addition, PAA dissolves easily in water and decomposes

to acetic acid and water, which are non-toxic. This biocide is known for its fungicidal,

sporicidal and bactericidal properties – active at concentrations as low as 0.3%

(McDonnell and Russell, 1999). The main disadvantages of PAA are: i) the stinging odor;

ii) the low stability and iii) its high toxicity when concentrated (40%) (Grinstead, 2009).

Moreover, the mechanism of action of PAA is based on the targeting of the cell

membrane, oxidizing and denaturing proteins and lipids, which leads to change in the

membrane organization (Russell, 2001b). This modification is associated with the

cleavage of sulfur and sulfhydryl bonds that may alter the cell wall permeability (Block,

2001).

2.2.1.1.5. Ozone

Ozone is a powerful sanitizer due to the its potential oxidizing power. A few

applications of ozone in the food industry, suggested by Guzel-Seydim et al. (2004),

are food surface sanitation, process water treatment and also reducing the biological

and chemical oxygen demand of waste water. Ozone is not available commercially and



so it must be produced on site. However, its half-life is quite short (a few minutes)

and as soon as ozone is applied it will start converting back to oxygen (Berk, 2013).

The main disadvantages in using ozone are: i) at pH 7 ozone will start decomposing

spontaneously, which will result in the production of highly reactive free radicals

(Wysok et al., 2006); and ii) at high concentrations ozone may be fatal to humans

(Guzel-Seydim et al., 2004). Furthermore, ozone is highly biocidal and its mechanism

of action comprises a progressive oxidation of vital cellular components. The cell’s

surface is usually the first to be attacked through the oxidation of sulfhydryl groups

and amino acids of enzymes, peptides and proteins (Wysok et al., 2006) with

subsequent leakage of cellular contents (Guzel-Seydim et al., 2004). In the case of

Gram-negative bacteria, the lipoprotein and lipopolysaccharide layers are the first to

be attacked, leading to an increase in the cell permeability and consequent cell lysis.

Cellular death can also occur due to the potent destruction and damage of nucleic

acids (Guzel-Seydim et al., 2004). In the case of virus, ozone can attack the protein

capsid of bacteriophages F2 and T4, releasing and inactivating the nucleic acids

(Russell, 2001b).

2.2.1.1.6. Quaternary Ammonium Compounds

Quaternary ammonium compound (QAC) is a surfactant-based biocide, more

specifically a cationic surfactant. QACs have good biocidal activity against vegetative

bacteria and fungi, but are not strongly active against spores (Grinstead, 2009).

Although QACs are more expensive than other sanitizers (e.g. chlorine and its

derivatives), these compounds possess numerous qualities that make them an

attractive alternative: i) they are very stable, both in diluted solutions and

concentrates; ii) they are not as rapidly inactivated by organic matter as chlorine; iii)

they are not corrosive, except at high concentrations; iv) they can be stored for a long

time without losing their antimicrobial activity (Chaidez et al., 2007); and v) they are

stable over a broad range of temperature and pH, although at alkaline pH the activity

is higher (Parkinson, 2012). Conversely, QACs have reduced activity in the presence of

hard water, they are not compatible with anionic detergents and they form a

antimicrobial film (Grinstead, 2009). This residual film although it can carry an

antimicrobial activity over an extended period of time it can also interfere in the

future growth of microorganisms involved in the production of food such as beer

(Banner, 2012). At high concentrations, QACs are effective against a wide range of

microorganisms, such as bacteria (generally more active against Gram positive than

gram negative bacteria), lipophilic viruses, yeasts, mold and algae (Mousavi et al.,



2015). However, they are not highly effective against bacteriophages, bacterial spores,

mycobacteria, or hydrophilic viruses (Schmidt, 2009, Hegstad et al., 2010). In addition

to having antimicrobial properties, QACs are also excellent for hard-surface cleaning

and deodorization (McDonnell and Russell, 1999).

Concerning the mechanism of action, QACs target the cytoplasmic (inner)

membrane (Carmona-Ribeiro and Carrasco, 2013) and bind irreversibly to the

membrane phospholipids and proteins (Maris, 1995). This is followed by loss of

structural organization, integrity of the cytoplasmic membrane and consequent

leakage of intracellular material and degradation of proteins and nucleic acids

(McDonnell and Russell, 1999).

2.2.1.1.7. Acid Anionic Surfactants

Acid anionic surfactants are also surfactant-based biocides. However, these

sanitizers are negatively charged unlike QACs (Schmidt, 2009). The antimicrobial

activity of acid anionics is satisfactory, particularly against vegetative microorganisms

and virus but not bacterial and fungus spores (Troller, 1993, Grinstead, 2009). They

are also not very effective against molds and yeasts (Featherstone, 2015). In the food

industry, they are used in the dairy industry and are specially effective in controlling

milkstone (a carbonate formed during the processing of dairy-based products) (Singh,

2009). The advantage of using acid-anionic surfactants relies on: i) the stability at high

temperatures; ii) the low odor that they possess; and iii) the fact that they are not

corrosive or cause staining (Schmidt, 2009). Due to the acidic nature, these biocides

are suited for use in the presence of carbon dioxide, for example in breweries or soft

drinks plants (Featherstone, 2015).

The main disadvantages of acid-anionic surfactants are: i) the small pH range of

activity (pH 2 to 3.5); ii) excessive foaming in CIP systems (Schmidt, 2009); and iii) the

possible inactivation by cationic surfactant detergents (Grinstead, 2009). Depending

on the nature of the surfactant, acid-anionic surfactants may produce foam

(Featherstone, 2015).



2.2.1.2. Disinfectants Addressed in this Thesis

2.2.1.2.1. DBNPA

Dibromonitrilopropionamide (DBNPA) is a non-oxidizing biocide known for its fast

biocidal action when removing and eliminating bacteria (Harrison and Sisk, 2004).

DBNPA has a density of 1190 g/L, molecular weight of 241.9 g/mol and a solubility in

water of 1.5 g in 100 mL (NCBI, 2016b). DBNPA is widely used in industrial circulating

water systems, large air-condition systems and also in sewage treatment to eliminate

microorganisms and algae (Klaine et al., 1996, Gao et al., 2001). DBNPA is also utilized

in the papermaking industry to prevent the reduction of the paper quality through the

degradation of starch by amylase enzymes produced by the microorganisms (Pirttijarvi

et al., 2001, Huang et al., 2009, Kolari et al., 2014).

Regarding the mechanism of action, DBNPA is an electrophilic active compound

that binds strongly and rapidly to the cell walls of the bacteria, reacting with

cytoplasmic constituents such as thiol groups of proteins, thus inhibiting cellular

metabolism (Maillard, 2002, Tommel, 2007). The basis of its mode of action relies on

the fact that DBNPA reacts through its bromine chemistry with sulphur-containing

nucleophiles, common to microorganisms, such as cysteine. This process occurs with

the inactivation of thiol-based (R-SH) amino-acids and enzymes by converting their

functional –SH groups to the oxidized S-S form, and with the formation of disulfide

bridges thus oxidizing the amino-acids beyond the formation of disulfide species. This

reaction irreversibly disrupts the function of cell-surface components, interrupting

transport across cell membranes and inhibiting key biological functions (Tommel,

2007). The use of DBNPA carries a few disadvantages such as rapid degradation under

alkaline conditions and sensitivity to UV light and nucleophilic substances. Under these

conditions it must be used as a quick kill substance (Huber et al., 2010).

The mechanism for the degradation of DBNPA has been described by Exner et al.

(1973). An overall scheme is presented in Figure 1 which illustrates two possible

pathways: i) formation of malonic acid and ii) formation of oxalic acid. The first

pathway was defined by the author as occurring only in the presence of nucleophiles

or sunlight.



Figure 1. Decomposition pathways of DBNPA (based on Exner et al.,

1973).

2.2.1.2.2. Sodium Bicarbonate

SB is a weak base commonly used as a food addictive, listed as Generally Regarded

As Safe (GRAS) by the Food and Drug Administration (FDA) (Qin et al., 2015). This salt

is considered as non-toxic as well as inexpensive and readily available (Palou et al.,

2001). SB has a density of 2100 g/L, a molecular weight of 84 g/mol and a solubility in

water of 8.5 g in 100 mL (NCBI, 2016a). SB is frequently used as a leavening (Gibbs et

al., 1999) and whitening agent in the dental field (Joiner, 2010). Furthermore, it can

also be applied as a disinfectant against bacteria (Drake, 1997) and more commonly

against molds in plants and fruits (Nigro et al., 2006, Smilanick et al., 2006).



Bicarbonates have been demonstrated to effectively control a wide range of fungi,

including food spoilage organisms and plant pathogens (Palmer et al., 1997).

Concerning the mechanism of action of SB, it has not been completely elucidated,

however, there is evidence that it may act against fungi by buffering an elevated pH

environment, or by increasing osmotic levels on surfaces, as well as through the

inactivation of extracellular enzymes (Palmer et al., 1997, Cerioni et al., 2012).

2.2.1.2.3. MC-06-DBES

MC-06-DBES, henceforward known as MC, is a formulation based in anionic and non-

anionic surfactants with enzymes and essential oils, such as carvacrol and 2-O-

palmitoyl-3-O-(6'-sulfoquinovopyranosyl)-glycerol (GL2A). MC has a density of 1060 g/L

and a pH of 7.5.

Enzymes are well established in the industry as a way to control protein biofilms

on surfaces and in closed pipelines (Augustin et al., 2004). Their use in detriment of

other disinfectants, such as chlorine dioxide, came from the need to overcome the

negative environmental impact that these chemical disinfectants carry, as well as the

resistance that microorganisms are developing for such disinfectants (Molobela et al.,

2010)

Regarding MC’s mechanism of action, the enzymes present in the solution target

primarily the EPS matrix. However, this matrix has heterogeneous composition and it

varies from biofilm to biofilm (Meyer, 2003, Xavier et al., 2005). As a result of this

characteristic, a mixture of enzymes may be necessary to completely degrade the

biofilm (Simões et al., 2010). Therefore, a combination of specific enzymes with other

treatments, such as antimicrobial agents, is required in order to achieve better results

(Lequette et al., 2010).

Some enzymes such as protease, DNase I, alginate lyase, amylase and cellulase

have proved that can successfully remove biofilms (Molobela et al., 2010, Nijland et

al., 2010, Lamppa and Griswold, 2013). A major drawback from the use of enzymes as

part of the disinfection process is their high cost, when compared with the low prices

of the chemical disinfectants (Augustin et al., 2004, Thallinger et al., 2013). Regarding

carvacrol, the essential oil present in this disinfectant, it is a major component of the

essential oil fractions of oregano and thyme (Knowles and Roller, 2001). Carvacrol has

GRAS status and it is mostly used as a savoring agent in baked goods, beverages, and



chewing gum (Burdock, 2004). On the subject of the antimicrobial activity of carvacrol,

this compound has been proved to be effective against a wide range of foodborne fungi

and bacteria including E. coli, Enterococcus faecalis, Listeria monocytogenes,

Salmonella enterica sv. Typhimurium, S. aureus, and Bacillus cereus (Ultee et al.,

2000, Guarda et al., 2011, Soni et al., 2013, Benbelaïd et al., 2014, Silva-Angulo et

al., 2015).

2.2.1.3. Factors Affecting Disinfectant Effectiveness

The activity of sanitizers against microorganisms depends on a variety of factors:

some factors are related to the chemical itself, or external conditions in which the

disinfection is performed, and others are even related to the type of microorganisms

present in the surface. All these factors should be considered in order to have a more

effective disinfection (Schmidt, 2009). Furthermore, the characteristics of the surface

also need to be considered. For example if the surface has cracks or pits the sanitation

effectiveness decreases, since these modifications can shelter the microorganisms

(Schmidt, 2009). Besides, to have an effective sanitation it is necessary that all the

involved surfaces have been cleaned previously (Seier-Petersen, 2013). In order to have

an eliminating action on the microorganisms the sanitizer requires direct contact with

them (Schmidt, 2009). This way, one factor that contributes to a reduction in the

activity of sanitizers is the presence of organic matter. It can act as a physical barrier

between microorganisms and the disinfectants (Lewis and Arens, 1995) and it can also

react chemically with the disinfectants reducing its bactericidal potential (Rutala and

Weber, 2008). Another important issue is the exposure time. There is a minimum

contact time that is necessary in order to reduce the microorganisms’ viability.

Usually, the longer time the sanitizer is in contact with the surface, the more effective

is the sanitization. However, in some cases, the prolongation of the contact time does

not improve the effectiveness of the disinfection (Marriott and Robertson, 1997).

Temperature is also a contributing factor and, generally, the activity of sanitizers

increase as the temperature of the solution increases (Marriott and Robertson, 1997).

However, very high temperatures may cause the degradation of the disinfectant and

thereby decrease its activity (Rutala and Weber, 2008), as well as damage the

equipment/surface.

Concomitantly, an increase in the concentration of the disinfectant leads to a

stronger activity and subsequently to a shorter time necessary to eliminate the

microorganisms (Russell and McDonnell, 2000). Nevertheless, applying the sanitizer at



a concentration higher than the one recommended by the manufacturer usually does

not improve disinfection and, in contrast, can be corrosive to equipment and cause

food damage if not properly eliminated (Schmidt, 2009).

When it comes to pH, sanitizers are greatly affected by the pH of the solution. For

some sanitizers, an increase in pH results in the improvement of the antimicrobial

activity, since the higher pH causes a change in the molecule or on the cells’ surface

(Russell, 2004). For example, in the activity of QACs a rising pH improves these

sanitizers’ activity. However in some cases (hypochlorites and iodine) the increased

pH can decrease the antimicrobial activity (Rutala and Weber, 2008). In the case of

hypochlorites, raising the pH leads to an increasing difficulty in the releasing of

hydroxyl ions, and the subsequent attack to the cytoplasmic membrane (Estrela et al.,

2002).

Moreover, certain disinfectants are significantly affected by the water properties,

such as hardness (Schmidt, 2009). A rise in the hardness of the water causes an increase

in the concentration of ions such as calcium or magnesium. These ions interact with

the disinfectant and result in the formation of insoluble precipitates that can reduce

the efficiency of certain disinfectants (Rutala and Weber, 2008).

There are also a few biological factors that may interfere with the sanitation

efficiency. The microbiological load is an important factor, since the higher the

number of microorganisms is, the more time it will be to eliminate all (Schmidt, 2009).

Also, with different types of microorganisms the resistance to disinfectants also varies

– e.g. bacterial spores are more resistant than vegetative cells and, apart from prions,

spores possess the highest resistance to chemical sanitizers (Russell, 2001b).

2.2.1.4. Effectiveness of Disinfectants Against Different Organisms on Food

Contact Surfaces

2.2.1.4.1. Bacillus cereus

A study was undertaken by Peng et al. (2002b) to evaluate the efficacy of SH and

a QAC, separately, against planktonic and biofilm cells of B. cereus, on SS. It was

reported a poor reduction of B. cereus biofilm by 300 ppm of QAC, since the biocide

only reduced 1.5 log CFU, after 30 s. For the same exposure time, 25 ppm SH achieved

the same reduction. For planktonic cells, a 5 log CFU reduction was achieved after 15

s of exposure to 25 ppm SH. For the same reduction and exposure time, it was

necessary 100 ppm of QAC. Thus, proving that hypochlorite is more effective than QAC



in the elimination of both planktonic and biofilm cells of B. cereus and also that

planktonic cells are easier to eliminate (Peng et al., 2002b).

Sudhaus et al. (2013) studied the effect of a PAA based disinfectant on spore

suspensions of B. cereus. With this purpose the spore suspensions were treated with

various concentrations of PAA at three different temperatures (10, 15 and 20 °C) and

with different exposure times (5, 30 and 60 min). On one hand, the treatment of spores

of B. cereus (strain DSM 4384) with 2.0% PAA, for 30 min at 10 °C, inactivated all

spores. On the other hand, for the same conditions it was achieved a reduction of less

than 1 log CFU for a different strain (DSM 4313). For the complete removal of this

strain it was necessary a 60 min exposure time with PAA at 5% and at 10°C, which

indicates that different strains have different resistances to the antimicrobials applied

(Sudhaus et al., 2013).

2.2.1.4.2. Cronobacter sakazakii

Park et al. (2015) investigated the effect of ultrasounds and SH on reducing

Cronobacter sakazakii on the surface of head lettuce. Ultrasounds (37 kHz, 1200 W)

reduced the population of C. sakazakii in 0.58 log CFU/g after 100 min. On the other

hand, with SH (200 ppm) treatment, the authors were able to achieve a significant

reduction of C. sakazakii after 5 min (2.84 log CFU/g). The synergic combination of

ultrasound and SH in the previous conditions attained a 4 log CFU/g reduction. Kim et

al. (2006) studied the efficacy of chlorine dioxide and a PAA based sanitizer (Tsunami

200®) in eliminating C. sakazakii from apples and lettuce. After 5 min of exposure,

chlorine dioxide (10 μg/mL) reduced 3.77 log CFU/apple of C. sakazakii and

Tsunami 200 (40 μg/mL) treatment achieved a ≥ 4 log CFU/apple reduction of

C. sakazakii. For samples of lettuce leaves (9 × 9 cm) and an exposure time of 5 min,

chlorine dioxide and Tsunami 200 reduced C. sakazakii in 4 log CFU/sample and ≥ 5 log

CFU/sample, respectively. Overall, treatment of apples and lettuce with chlorine

dioxide and Tsunami 200 resulted in significant reductions of C. sakazakii.

2.2.1.4.3. Enterobacter sakazakii

Kim et al. (2007) examined the effectiveness of QAC and PAA against E. sakazakii

strain 3231 in suspension and in a 6 day biofilm on coupons of SS. Both biocides were

able to significantly reduce the number of planktonic cells in 1 min of exposure time.

For QAC the reduction was nearly 7 log CFU (reduction to almost undetectable levels)

and for PAA the reduction was nearly 6 log CFU. PAA disinfection allowed undetected

levels after 5 min of exposure. The performance of the biocides against E. sakazakii in



biofilms, however, was much different. None of the biocides were able to reduce the

microbial levels in the biofilm to undetectable levels within 1 min of exposure. After

10 min of exposure QAC showed no significant reduction of the microbial load at any

exposure time. At the same time PAA was able to achieve 1 log CFU reduction

(statistically significant) after 5 min and 2.4 log CFU reduction after 10 min of

exposure. Consequently, PAA was better suited to eliminate E. sakazakii biofilms (Kim

et al., 2007).

2.2.1.4.4. Escherichia coli

Cabeça et al. (2012) compared the efficacy of various disinfectants – iodine, QACs,

PAA and SH – on planktonic and biofilm cells of E. coli on SS coupons. After 10 min of

exposure, no growth of planktonic cells was detected for all the biocides. However,

biofilm cells showed to be more difficult to eliminate. For the same time of exposure,

iodine reduced 5.1 log CFU/cm2, QACs reduced 4.2 log CFU/cm2, PAA reduced 3.8 log

CFU/cm2 and SH reduced 5.6 log CFU/cm2. SH proved to be the most effective biocide

against biofilm cells, while PAA was the least effective (Cabeça et al., 2012).

Another study was carried out by Oulahal-Lagsir et al. (2003) to evaluate the

efficiency of ultrasounds (US) and enzymes in the removal of E. coli biofilms developed

on SS surfaces. In this experiment US (40 kHz, 10s) allowed a complete removal of the

biofilm. On the other hand, the enzyme trypsin (7600 U/mL, pH 7, 20°C) required

30 min to achieve a reduction of biofilm of 96%. In this study, the combination of US

and trypsin was also tested. The SS slides were immersed in the enzymatic preparation

and then the US (40 kHz) were applied for a total time of 10s. This combination allowed

a reduction of 76%. Thus, improving the enzyme efficiency as it achieved a good

reduction well under 30 min.

Ban et al. (2012) tested the effect of lactic acid on E. coli O157:H7 formed on PVC

and SS coupons. The biofilm was developed on 5 x 2 cm coupons for 6 days at 25°C

under static conditions. The quantity of cells reached was 5.91 and

5.84 log CFU/coupon for PVC and SS coupons, respectively. The application of lactic

acid (2% v/v, 30 s) allowed a 0.9 log CFU/coupon reduction on PVC and

0.8 log CFU/reduction on SS. When lactic acid was combined with 5 s of steam

exposure an additional reduction of 2.9 and 3.1 log CFU/coupon was achieved for PVC

and SS (respectively), indicating a synergic effect.



2.2.1.4.5. Listeria monocytogenes

Robbins et al. (2005) studied the efficacy of ozone and hydrogen peroxide to

destroy L. monocytogenes planktonic and biofilms cells. The study showed that using

2.0 ppm of ozone for 3 min lead to the reduction of over 8 log CFU of unattached

L. monocytogenes cells. The same concentration and exposure time resulted in only

4-4.5 log CFU reduction of L. monocytogenes in biofilms. Similarly, hydrogen peroxide

at 3.0% reduced planktonic cells of L. monocytogenes to undetectable levels in 10 min.

However, if the treatment was used to eliminate L. monocytogenes biofilm, this

concentration (3.0%) of hydrogen peroxide would not be able to completely eliminate

the biofilm even after 15 min of exposure. The authors needed a 5.0% concentration

of hydrogen peroxide for 15 min to achieve complete elimination of biofilm (Robbins

et al., 2005). Ayebah et al. (2006) studied the ability of a solution of hypochlorous acid

generated through an electrochemical reaction of sodium chloride in water (also

known as electrolyzed oxidizing water or EOW) to inactivate L. monocytogenes in

suspension and in biofilms. The EOW is a broad-spectrum and high-performance

bactericide and is being increasingly applied in the food industry. The EOW reduced

planktonic L. monocytogenes to undetectable levels (over 7 log CFU reduction) in one

minute. However, the reduction of L. monocytogenes biofilm was less than 4.5 log

CFU.

2.2.1.4.6. Salmonella enteritidis

Martínez-Hernández et al. (2015) examined the effects of SH, PAA, ultraviolet C

light (UV-C) and the PAA+UV-C combination in the sanitizing treatments to reduce

S. Enteritidis growth on fresh-cut kalian-hybrid broccoli. For a concentration of PAA

and SH of 100 mg/L and exposure time of 2 min and an illumination of 7.5 kJ UV-C m-2

the reduction of S. Enteritidis determined was 2.1, 2.7 and 2.2 log CFU/g for SH, PAA

and UV-C, respectively. In this case PAA was the disinfectant that allowed a higher

S. Enteritidis elimination. The combination PAA+UV-C allowed a reduction of 3 log

CFU/g. According to Limpel’s formula (Richer, 1987) the combination of PAA with UV-

C promoted the S. Enteritidis elimination, but it is considered as an addictive effect

and not a synergic action.

2.2.1.4.7. Staphylococcus aureus

Ding et al. (2016) studied the disinfection efficacy of SH and hydrochloric acid on

S. aureus. SH (1 min exposure) allowed a 3.26 log CFU reduction of S. aureus and with

hydrochloric acid a lower reduction of 2.73 log CFU was obtained. At the same time,

slightly acidic electrolyzed water was tested for 1 min obtaining a much higher



reduction of 5.8 log CFU. Abdallah et al. (2015) undertook a study to investigate the

relationship between the environmental conditions of biofilm formation and the

resistance to disinfectants. The antibiofilm efficacy of disinfectants was investigated

on S. aureus biofilms grown at 20, 30 and 37 °C during 24 h, on SS and polycarbonate

(PC). The increase of temperature significantly increased the S. aureus biofilm

resistance to disinfectants. It was tested a combination of bis (3-aminopropyl)

dodecylamine (BDA) and didecyldimethylammonium chloride (DDAC). The results

underlined that the treatment with the mixture of disinfectants completely removed

the initial population of S. aureus biofilms in both surfaces tested. However, the

treatment of S. aureus biofilms, grown at 30 °C and 37 °C, only reduced the initial

population to 1.2 and 2.8 log CFU/cm2, respectively on SS and to 1.7 and 2.3 log

CFU/cm2, respectively on PC.

In Table 2 a list of some of the disinfection methods applied in the food industry

surfaces is summed up.

Table 2. Disinfection methods applied in the food industry on food contact surfaces

Disinfectant Method Results Reference

Sodium hypochlorite

25 ppm; 15s Reduction of 5 log CFU/mL of B. cereus

Peng et al. (2002b)

25 ppm; 30s Reduction of 1.5 log CFU/cm2 of B. cereus on SS surfaces

Peng et al. (2002b)

1.5%; 10 min Reduction of 5.6 log CFU/cm2 of E. coli on SS surfaces

Cabeça et al. (2012)

30 ppm; 1 min Reduction of 3.3 log CFU/mL of S. aureus

Ding et al. (2016)

250 ppm; 30 s Reduction of 2.7 log CFU/cm2 of S. aureus on SS surfaces

de Souza et al. (2014b)

250 ppm; 30 s Reduction of 1.9 log CFU/cm2 of S. aureus on Polypropylene (PP) surfaces


Hydrochloric acid 0.1%; 1 min Reduction of 2.7 log CFU/mL of S. aureus

Ding et al. (2016)

QACs 100 ppm; 15 s Reduction of 5.6 log CFU/mL of B.cereus

Peng et al. (2002b)

300 ppm; 30 s Reduction of 1.4 log CFU/cm2 of B.cereus on SS surfaces

Peng et al. (2002b)

0.11% ; 1 min Reduction of 7 log CFU/mL of E. sakazakii

Kim et al. (2007)

0.11% ; 1 min Reduction of 1 log CFU/cm2 of E. sakazakii on SS surfaces

Kim et al. (2007)

0.11% ; 5 min Reduction of 2.4 log CFU/cm2 of E. sakazakii on SS surfaces

Kim et al. (2007)

0.5%; 10 min Reduction of 4.2 log CFU/cm2 of E. coli on SS surfaces




Disinfectant Method Results Reference

Iodine 0.2% ; 10 min Reduction of 5.1 log CFU/cm2 of E. coli on SS surfaces


PAA 0.5%; 10 min Reduction of 3.8 log CFU/cm2 of E. coli on SS surfaces


0.4%; 15 min Reduction of 5.6 log CFU/cm2 of P. aeruginosa on PS surfaces

Martín-Espada et al. (2014)

1.6%; 15 min Reduction of 5.6 log CFU/cm2 of P. aeruginosa on PS surfaces

Martín-Espada et al. (2014)

30 ppm; 30 s Reduction of 3.2 log CFU/cm2 of S. aureus on SS surfaces


30 ppm; 30 s Reduction of 3.7 log CFU/cm2 of S. aureus on PP surfaces


Ozone 2 ppm; 3 min Reduction of 8 log CFU/mL of L. monocytogenes

Robbins et al. (2005)

2 ppm; 3 min Reduction of 4-4.5 log CFU/cm2 of L. monocytogenes on SS surfaces


Hydrogen peroxide

3%; 10 min Reduction of 6 log CFU/mL of L. monocytogenes


5%; 15 min Reduction of 5.6 log CFU/cm2 of L. monocytogenes on SS surfaces


DDAC + BDA 490 ppm + 180 ppm; 15 min; 37 ºC

Reduction of 2.8 log CFU/cm2 of S. aureus on SS surfaces

Abdallah et al. (2015)

490 ppm + 180 ppm; 15 min; 37 ºC

Reduction of 2.3 log CFU/cm2 of S. aureus on PC surfaces

Abdallah et al. (2015)

SB 5% ; 1 min Reduction of 99.22% of feline calicivirus on food contact surfaces

Malik and Goyal (2006)

SB + hydrogen peroxide

2% + 2%; 10 min Reduction of 99.68% of feline calicivirus on food contact surfaces


SB + glutaraldehyde

1% + 1.3%; 1 min

Reduction of 99.99% of feline calicivirus on food contact surfaces


Anionic surfactant +α-amylase

50 mM + 5-10%; 30 min; 45 ºC

Reduction of 3 log CFU/cm2 of B. mycoides on SS surfaces

Lequette et al. (2010)

50 mM + 5-10%; 30 min; 45 ºC

Reduction of 1.6 log CFU/cm2 of P. fluorescens on SS surfaces

Lequette et al. (2010)

Spezyme GA 300 7.3 mg protein/mL; 30 min

Reduction of 4 log CFU/mL of P. aeruginosa

Augustin et al. (2004)

Resinase A 2X 2.34 mg protein/mL; 15 min

Reduction of 3 log CFU/mL of P. aeruginosa

Augustin et al. (2004)



2.2.1.5. Biofilms and Reasons for the Ineffectiveness of Disinfectants Against

Biofilms

The main reason for the ineffectiveness of biocides rests in the fact that the EPS,

that microbial cells produce in biofilms, interfere with the penetration of the biocide

(Grinstead, 2009). De Beer et al. (1994) examined the penetration of chlorine into

biofilms of P. aeruginosa and Klebsiella pneumoniae. They observed that chlorine

concentrations in biofilms were less than 20% of the concentration in the bulk liquid.

After exposure to 2.5 ppm chlorine for 1-2 hours a complete chlorine equilibrium

between the bulk phase and the biofilm was not achieved. After 1 hour, only the top

layer (100 μm) of the biofilm had been penetrated by this disinfectant. Stewart et al.

(2000) also observed that hydrogen peroxide was unable to penetrate a biofilm of

P. aeruginosa. However, when hydrogen peroxide was applied on a biofilm of a

catalase deficient mutant of P. aeruginosa it achieved full penetration and was able

to eliminate the resident cells (Stewart, 1996). The data presented by Stewart et al.

(2000) and De Beer et al. (1994) suggested that biocides are inactivated during passage

through the EPS and this way the biocide does not have the “strength” to attack the

microorganisms when it finally reaches the target.

McDonnell and Russell (1999) hypothesized that the production of enzymes, such

as catalase and superoxide dismutase, may be intensified in microorganisms in

biofilms. For this reason, cells in biofilms may grow more slowly than planktonic cells.

Slow-growing bacteria have been known to be less susceptible to disinfectants (Dodd

et al., 1997). However, in a study performed by Pan et al. (2006) it was found that

despite the fact that Listeria cells grown in a biofilm were resistant to biocides, such

as peroxides, QACs and chlorine, the resistance did not persist when these cells were

removed from the biofilm. Thus, proving what McDonnell and Russell (1999) stated,

that if cells are removed from a biofilm and re-cultured they tend to be as resistant as

the planktonic cells of that species and not more resistant.



Chapter 3

3. Behavior of E. coli planktonic cells in the presence of DBNPA, sodium

bicarbonate and the enzymatic disinfectant MC

3.1. Introduction

Due to its high prevalence in the gastrointestinal tract of humans and warm-blooded

animals, E. coli is used as the preferred indicator to detect and measure faecal

contamination in the assessment of food and water safety (Odonkor and Ampofo,

2013). For this reason, this Gram-negative bacterium was chosen for this project. Most

E. coli are harmless commensal organisms when contained in their natural intestinal

habitat. Nonetheless, some strains can be pathogenic and they are distinguished from

other E. coli by their ability to cause illness through genetically controlled mechanisms

such as toxin production, adhesion and invasion of host cells, interference with cell

metabolism and tissue destruction (FAO, 2013).

Regarding food pathogens they tend to exist more within or on structured matrices

(biofilms) and seldom as planktonic cells in free water or other liquids (Jakobsen et

al., 2011). However, when it comes to biofilm formation there are 5 stages: i)

reversible attachment, ii) irreversible attachment, iii) early development of biofilm

architecture, iv) maturation v) and finally dispersion. As the first step includes the

attachment of planktonic cells to the surface (Srey et al., 2013), the elimination of

those planktonic cells can reduce the probability of biofilm formation.

3.2. Materials and Methods

3.2.1. Microorganism and culture conditions

The microorganism studied was E. coli CECT 434. This bacterium was already used

as model microorganism for antimicrobial tests with disinfectants by Meireles et al.

(2015a) and Borges et al. (2014), since it is referenced to be used for the development

of disinfection strategies by the European Committee for Standardization (EN-1276,

1997). Prior to use, E. coli stored at –80 ºC ± 2 ºC in a cryovial containing 30 % (v/v) of

glycerol (Panreac, Spain) was subcultured in Plate Count Agar (PCA) (Oxoid, England)

plates at 30 ºC during 24 h before testing.



The inoculum was obtained from overnight cultures grown in 100 mL flasks with 25

mL of Mueller-Hinton broth (MHB) (Oxoid, Hampshire), incubated (VELP Scientifica FOC

225l cooled incubator, Italy) at 30 °C in an orbital shaker (Orbital IKA KS 130 basic,

Germany) at 120 rpm. For all experiments, the aforementioned conditions of

temperature and agitation were maintained.

3.2.2. Disinfectants

The disinfectants tested were DBNPA 10% (v/v) (Enkrott, Portugal), SB 1M (Merck,

Germany) and MC-06-DGES 100% (Cleanity, Spain). In order to prepare the

concentrations tested, for each of the disinfectants, microcentrifuge tubes (1.5 mL,

Graduated, BIOplastics, Netherlands) were used and the liquid volume was completed

with sterile distilled water.

3.2.3. Determination of minimum inhibitory concentration for DBNPA

The MIC for DBNPA was determined in 96-wells PS microtiter plates (Orange

Scientific, Belgium). The bacteria were grown overnight at 30 °C in MHB, and then the

inoculum was adjusted to an OD600nm of 0.1 in a V-1200 spectrophotometer (VWR

International, EUA). In each microtiter plate well, 180 µL of bacterial inoculum were

added to 20 µL of increasing concentrations of DBNPA (5 - 70 ppm), for a total volume

of 200 µL. Cell suspension and MHB (200 µL) were used as controls. After 24 h of contact

time, the suspensions were subjected to a process of biocide neutralization, by

removing 180 µL of the suspension and pipetting 180 µL of the neutralizer solution into

each well, according to the method of Johnston et al. (2002). The disinfectants were

chemically neutralized (10 min contact time) by the solution which is composed by the

elements presented in Table 3 (the compounds were added by the order present in the

table and solubilized in phosphate buffer 0.25 M pH 7.2).

Table 3. Neutralizer solution composition

Compounds Concentration (g/L)

Lecithin (Alfa Aesar, Germany) 3

Sodium thiosulfate pentahydrate (Labkem, Spain) 5

L-histidin (Merck, Germany) 1

Saponin (VWR chemicals Prolabo, Belgium) 30

Tween 80 (VWR chemicals Prolabo, France) 30



Before and after the incubation, the absorbance was measured at 600 nm using the

SynergyTM HT 96-well microplate reader (Biotek Instruments, USA). The MIC was

determined by the microdilution method and recorded as the lowest concentration of

DBNPA that inhibits the growth of the microorganism (Andrews, 2001). Three

independent experiments were performed for each condition tested.

3.2.4. Determination of MIC for DBNPA combined with SB and MC

The determination of the MIC for the disinfectant combinations was performed

going according to 3.2.3 with a difference in the preparation of the microtiter plate.

Hence, disinfectant combinations were assayed by applying increasing concentrations

of DBNPA (10 µL) and 37 800 ppm SB and 25 000 ppm MC, for a total volume of 20 µL.

In each microtiter plate well, 180 µL of bacterial inoculum was added to 20 µL of the

disinfectants combination. Cell suspension and MHB (200 µL) were used as controls.

3.2.5. Determination of minimum bactericidal concentration for DBNPA

3.2.5.1. MBC in the presence of MHB

After the overnight-growth of the culture, the OD600 was adjusted to 0.1, using

fresh medium. Afterwards, after pipetting the bacterial suspension and the

disinfectants, in the same amounts as previously described in Section 3.2.3, using cell

suspension and MHB as controls, the plate was incubated for 24 h. After the incubation

time, the suspensions were subjected to a process of biocide neutralization, as

mentioned in 3.2.3. Later, bacterial suspensions undertook serial dilutions in 0.85%

saline solution (Scharlau, Spain), from 100 to 10-6. Then, the motion drop method (Reed

and Reed, 1948) was applied by transferring 10 µL of each suspension onto PCA plates

which were then incubated at 30 ˚C. Colony enumeration was carried out after 24 h.

The MBC was taken as the lowest concentration of the disinfectant at which no CFU

were detected on solid medium (Ferreira et al., 2011). Three independent experiments

were performed for each condition tested.

3.2.5.2. MBC in the absence of MHB

In order to understand the effect of the medium in the determination of the MBC,

this essay was repeated under different conditions. The overnight-grown culture was

centrifuged (Eppendorf centrifuge 5810R, Germany) in a 50 mL falcon tube (VWR

International, EUA) at 3220 g for 15 min, the MHB medium was discarded and the cells

were washed with 0.85% saline solution and then centrifuged once more. Afterwards,



bacteria were resuspended in saline solution to obtain an OD600 of 0.1. Subsequently,

in each microtiter plate well, 180 µL of the bacterial suspension was added to 20 µL of

increasing concentrations of the disinfectant, for a total volume of 200 µL, using cell

suspension as control. The plate was incubated for 1 h and, afterwards, the suspensions

were then subjected to a process of biocide neutralization and the dilution and colony

enumeration process was the one indicated in Section 3.2.5.1.

3.2.6. Statistical Analysis

The mean and standard deviation within samples were calculated. The results were

compared using Student’s t-test with the aim to understand if the differences between

the experimental values obtained under different conditions could be considered

significant. Significant difference between means was assumed for P < 0.05.

3.3. Results and Discussion

3.3.1. Determination of the MIC for DBNPA

The effect of DBNPA in bacterial suspension growth was studied and several

concentrations were tested (between 5 and 70 ppm). From the results obtained, it was

possible to observe that the minimal concentration that inhibited the cellular growth

was 20 ppm. Therefore, this concentration was considered the MIC.

3.3.2. Determination of the MBC for DBNPA

After the determination of the MIC, the MBC value was obtained, plating several

concentrations sub and above-MIC. To represent these results, a graphic with the

evolution of the log CFU/mL with the increasing concentrations of DBNPA is presented

in Figure 2. From the figure it is possible to observe a natural tendency of the number

of log CFU/mL to decrease as the concentration of DBNPA increases. Moreover, it is

also possible to conclude that the MBC for DBNPA is 20 ppm, since at this concentration

no CFU was detected. These results differ from the results obtained by Kim and Park

(2016) that determined the MIC and MBC of DBNPA for E. coli as 125 ppm, however, in

that study E. coli was grown in tryptic soy broth at 37 ˚C and the pH of DBNPA was

adjusted to 7.0 which could affect the result. Additionally, Huang et al. (2009) results

indicated that for P. aeruginosa, in MHB, the MIC and MBC for DBNPA was 64 ppm.

However, and even though it is also a Gram-negative bacterium, different bacteria can

be affected in different ways by the same disinfectant.



Figure 2. Graphic representation of the evolution of log CFUs with the increasing concentrations of DBNPA. The line indicates the method detection limit (2 log CFU/mL).

When comparing with a sanitizer typically used in the food industry, such as SH,

Meireles et al. (2015a) determined that the MBC was 447 ppm, thus proving that DBNPA

can be effective under very low concentrations, when compared to SH. This can be

very useful since low concentrations of biocide can be applied.

With the purpose of understanding the effect of MH medium in the determination

of the MBC, an assay with saline solution was performed. The MBC was determined as

3 ppm, thus confirming that the medium influences the results, probably by reacting

with the disinfectant instead of the cells. Huang et al. (2009) tested DBNPA in P.

aeruginosa in simulated white water (papermaking process water) and determined that

the MBC was 16 ppm. Although this value does not confirm the results obtained, the

fact that the authors achieved a different MBC in MHB and in a simulated water

complies with the results presented in this thesis.

3.3.3. Determination of the synergetic activity of DBNPA with SB and DBNPA

with MC

In order to understand, the synergic activity of SB and MC with DBNPA in planktonic

E. coli cells, a graphic with the variation of the O.D. at 600 nm is presented in Figure

3. It is possible to conclude that in order to obtain inhibition of the cellular growth it

is necessary either the same concentration as the MIC for DBNPA (in the MC case) or

even the double of the concentration (in the SB case).

0.0

1.0

2.0

3.0

4.0

5.0

6.0

7.0

8.0

9.0

0 5 10 15 20 25

log

CFU

/mL

[DBNPA] (ppm)



Figure 3. Evaluation of the combination of DBNPA with SB and MC through O.D. variation analysis.

Therefore, to confirm this result, the presence or absence of synergism was

determined by the calculation of the ratio between the MIC of DBNPA in combination

and MIC of DBNPA alone. For a MIC ratio from 0 to 1 (excluding) potentiation to modest

enhancement is occurring. In the case of a MIC ratio higher than 1 the combination is

antagonic (Meireles et al., 2015a). As expected, the MIC ratio was 1 and 2 for MC and

SB (respectively) thus proving that, for the treatment of planktonic E. coli, DBNPA

alone is more effective than the combinations.

3.4. Conclusions

In this chapter, DBNPA was found to be effective against planktonic E. coli and it

is active at concentrations as low as 20 ppm (MIC and MBC). In order to potentiate the

effect of DBNPA on planktonic cells, the combinations with SB and MC were tested. It

is possible to conclude that in the treatment of planktonic E. coli the combination of

DBNPA with SB 37 800 ppm and with MC 25 000 ppm was antagonic.

0.00

0.10

0.20

0.30

0.40

0.50

0.60

5 ppm 10 ppm 20 ppm 30 ppm 40 ppm

O.D

. 600

nm

vari

atio

n

[DBNPA] (ppm)

Combination with 37 800 ppm SB

Combination with 25 000 ppm MC



Chapter 4

4. Control of E. coli biofilms in different surfaces

4.1. Introduction

One of the biggest concerns in the food industry is the difficulty in the elimination of

pathogens, mainly due to the fact that bacteria can attach to food contact surfaces and

form biofilms, surviving even after cleaning and disinfection (Garrett et al., 2008, de Souza

et al., 2014a). A common cause of food contamination relies in the presence of biofilms.

After initial attachment of the cells to the surface, using flagella, pili, and membrane

proteins, they start to multiply and secrete a consistent matrix of EPS in which cells are

enclosed (Van Houdt and Michiels, 2005). Biofilms are a serious problem in many food

industry sectors (Gutierrez et al., 2012), since they are difficult to eradicate from surfaces.

Moreover, bacterial colonization tend to damage the equipment, as it can cause pitting

and corrosion of metal surfaces and the breakdown of plastics (Myszka and Czaczyk, 2011).

Hence there is a necessity to improve the disinfection methods for the sanitation of

surfaces. When it comes to the materials used in food-contact surfaces, the most used ones

are metals, plastics, and other materials (Lewan and Partington, 2014). Regarding metals,

SS is the material most commonly used is food processing equipment due to its durability

and resistance to corrosion. This material can be used in equipment such as pasteurizers,

heat-exchangers or storage tanks (Dewangan et al., 2015). Concerning the plastics, these

materials are widely used in the food industry for example in cutting surfaces, packages,

conveyor belts or pipes (Zeraik and Nitschke, 2012). These plastics include PS and PVC.

These three materials abovementioned were the three tested in this thesis project.

4.2. Materials and Methods

4.2.1. Microorganism and culture conditions

The microorganisms and culture conditions used for biofilm development were

described in Section 3.2.1.

4.2.2. Disinfectants

The disinfectants tested were described in Section 3.2.2. In order to prepare the

concentrations tested, for each of the disinfectants, sterile 0.85% saline solution was

used to complete de volume.



4.2.3. Biofilm formation in microtiter plates

In order to understand if DBNPA, SB and MC can control the formation of biofilms,

the crystal violet (CV) assay was undertaken. In this regard, the biofilm developing

process in microtiter plates was adapted from the one proposed by Stepanović et al.

(2000). The bacteria were grown overnight in 25 mL of MHB at 30 °C in an orbital

shaker (120 rpm). Then, the OD600nm of this inoculum was adjusted to 0.04, using a 50

mL falcon tube and fresh MHB. In each microtiter plate well, 200 µL of the bacterial

inoculum was added. The microtiter plates were then incubated for 24 h at 30 °C and

120 rpm.

4.2.3.1. Biofilm control with DBNPA, SB, MC and the combination

After the incubation time all the plate content was discarded and each well was

washed with 250 µL of saline solution, which was then discarded. Afterwards, 180 µL

of MHB and 20 µL of DBNPA, SB e MC as well as the disinfectants combinations were

added. 20 minutes of contact time was applied.

4.2.3.2. Biofilm mass quantification by crystal violet

After 20 min of contact time, the plate content was discarded and the wells were

washed with sterile water, which was then discarded. The bacterial biofilms were fixed

using 250 µL/well of 96% v/v ethanol (Aga, Portugal). The fixed bacteria were stained

for 5 min with 200 µL/well of CV (Merck, Germany) which was afterwards removed and

200 µL/well of sterile distilled water was added and then discarded to remove the

excess. Finally, 200 µL/well of glacial acetic acid (Fisher Chemical, United Kingdom)

was added. The optical density of the final solution was measured at 570 nm, using a

microtiter plate reader (Simões et al., 2007b).

4.2.3.3. Biofilm cells culturability

After 20 minutes of contact time with the disinfectant solutions, the plate content

was discarded and the wells were washed with 200 µL of saline solution, which was

then discarded. Afterwards, 200 µL of saline solution was added and the biofilm (wall

and bottom of the well) were scraped for 1 minute and the content was then pipetted

into a microcentrifuge tube (Abreu et al., 2014). This procedure was repeated 4 times

for a total volume of 800 µL. Then, serial dilutions, plating in PCA plates and CFU

enumeration was performed as described in section 3.2.5.1.



4.2.4. Biofilm formation on coupons in 48-well microtiter plates

Biofilms were formed on coupons (dimensions of 1.0 x 0.9 x 0.1 cm) of PS, SS (AISI

316) and PVC. The SS coupons were washed with ethanol 96% v/v and the PS and PVC

coupons were washed first with detergent, then with water and after this they were

subjected to 1h30 of UV exposure (40 W, SterilGARD, Baker, USA). Afterwards, the

coupons were inserted in 48-wells flat-bottomed PS tissue culture plates

(Thermo Fisher Scientific, USA) and 1 mL of cell suspension – O.D. 600 nm was adjusted

to 0.1 – was added to each well. The plates were incubated for 24 hours for biofilm

formation at 30 °C in an orbital shaker (120 rpm).

4.2.4.1. Biofilm control with DBNPA, SB, MC and the combination

After the incubation period, the coupons were transferred (using a sterilized

clamp) to a new 48-well plate containing the disinfection solutions. This plate was then

incubated for 20 minutes at 30 °C in an orbital shaker (120 rpm) (Meireles et al.,

2015b). Controls were obtained by placing the coupons in saline solution without the

disinfectant solution.


removal

After the disinfectant exposure, the number of cells attached to the coupons was

accessed by 4’,6-diamidino-2-phenylindole (DAPI, Merck, Germany) staining based on

Simões et al. (2007a). Each slide was stained with 20 µL of DAPI at a concentration of

0.5 µg/mL and then incubated for 10 min in the dark. The slides were mounted with

non-fluorescent immersion oil on glass microscope slides and visualized under an

epifluorescence microscope LEICA DMLB2 (Leica Microsystems, Germany) equipped

with a filter with the following characteristics: excitation filter 340–380 nm,

dichromatic mirror of 400 nm and suppression filter LP 425 (Meireles et al., 2015a).

For each coupon, a minimum of 10 micrographs were obtained using a microscope

camera (AxioCam HRC, Carl Zeiss) resorting to the software LAS version 4.2 (Leica

Microsystems, Germany) to determine the number of attached cells per cm2 (Gomes

et al., 2016). Three independent experiments were performed for each surface.

4.2.4.3. Effect of DBNPA, SB, MC and the combination in biofilm cells

culturability

After the exposure time, the coupons were placed in 9 mL of saline solution in

15 mL falcon tubes (VWR International, EUA) and the cells were removed by vigorous



vortex for 1 minute (Gomes et al., 2016). The serial dilutions, plating in PCA plates

and CFU enumeration were performed as described in 3.2.5.1.

4.2.4.4. Regrowth of biofilm cells after exposure to DBNPA, SB, MC and the

combination

After the exposure time, the coupons were transferred to a 48-well plate and 1 mL

of fresh medium as placed in each well. The plate was then incubated for 24 h at 30

°C in an orbital shaker (120 rpm). The serial dilutions, plating in PCA plates and CFU

enumeration were performed as described in 3.2.5.1.

4.2.5. Biofilm formation on coupons in an agitated reactor

The biofilm formation was performed using a reactor similar to the one described

by Ferreira et al. (2012). A 2 L glass continuous reactor at 25 ºC, aerated and

magnetically agitated was used (Figure A4 of the Appendix). Bacteria (500 mL) from

an overnight culture (5.5 g/L glucose (Chem-Lab, Belgium), 2.5 g/L peptone (Oxoid,

England), 1.25 g/L yeast extract (Merck, Germany), 1.88 g/L monopotassium

phosphate (Chem-Lab, Belgium) and 2.6 g/L sodium phosphate dibasic (VWR

Chemicals prolabo, Belgium)) were added (with an initial number of cells of 7 × 107

CFU/mL) to 1.5 L of 0.85% saline solution. The feeding process began 2 hours after the

inoculation. The reactor was continuously fed with 0.1417 L/h of a sterile solution with

55 mg/L glucose, 25 mg/L peptone, 12.5 mg/L yeast extract, 1.88 g/L monopotassium

phosphate and 2.6 g/L sodium phosphate dibasic. Twenty-four coupons of PVC

(2.0 × 2.0 x 0.1 cm) were placed vertically in contact with the bacterial suspension for

4 days. An illustration of the installation is presented in the Figure A5 of the Appendix.

4.2.5.1. Biofilm control with DBNPA, SB and MC and the combination

After the biofilm formation, the coupons were transferred to closed sterile 100 mL

beakers containing the disinfectant solutions and 0.85% saline solution for control. The

flasks were left at 25 ºC, for an exposure time of 20 min.


culturability

Afterwards, the coupons were placed in 10 mL of saline solution in 50 mL falcon

tubes which were then vigorously vortexed for 2 minutes. The serial dilutions, plating

in PCA plates and CFU enumeration were performed as described in 3.2.5.1.



4.2.5.3. Effect of DBNPA, SB, MC and the combination in biofilm thickness

After the disinfectant exposure, the biofilm topography was accessed by 4’,6-

diamidino-2-phenylindole (DAPI, Sigma, Portugal) staining based on Simões et al.

(2007a). Each slide was stained with 100 µL of DAPI at a concentration of 0.5 µg/mL

and then incubated for 10 min in the dark. Biofilms were analyzed in an

epifluorescence microscope Nikon Eclipse LV 100 (software, NiS-Elements AR 4.13.05)

equipped with a filter block sensitive to DAPI fluorescence (359 nm excitation filter in

combination with a 461-nm emission filter). With the software, 5 fields per image were

visualized through the camera (Nikon Digital Sight DS-Ri1, Japan) and with a joystick

(Prior Scientific Ltd, UK) the bottom and top of the biofilm were defined and between

the two 5 plans were analyzed and a 3D structure was built up. For each field, 5 vertical

plans were selected and 621 thickness points were measured.

4.2.5.4. Observation of biofilm images in Optical Coherence Tomography (OCT)

After the disinfectant exposure the biofilm structure was visualized resorting to a

spectral domain Optical Coherence Tomograph (Thorlabs Ganymede OCT System, USA)

with the following characteristics: central wavelength 930 mm, axial scan rare 29 kHz,

maximum imaging depth 2.7 mm and axial resolution air/water 5.8 µm/4.4 µm. The

2D and 3D images were created using the maximum intensity profile algorithm included

in the instrument software (ThorImage OCT version 4.2.4). OCT images were recorded

keeping the samples immersed in 0.85% saline solution.

4.2.6. Statistical Analysis

The statistical analysis was performed as referred in Section 3.2.6.

4.3. Results and Discussion

4.3.1. Efficiency of the elimination and removal of biofilm in microtiter plates

To access the biofilm formation and the removal associated with every disinfectant

and the combination of disinfectants the CV assay was performed. The results obtained

are present in Figure 4.



Figure 4. Quantification of E. coli biofilm mass by CV assay.

Observing Figure 4 is possible to conclude that amongst the three disinfectants

tested individually neither was able to attain a percentage of reduction higher than

30%. However, when DBNPA is combined with MC their combination is able to achieve

a reduction of 37%, proving to be a synergic combination, statistically different from

the individual use of DBNPA at 10 ppm (P<0.05), but not statistically different from the

individual use of DBNPA at 20 ppm (P>0.05). However, the combination of DBNPA with

SB proved the opposite as it reached a reduction of only 13%, almost half of the

reduction achieved by DBNPA 20 ppm individually (P<0.05), evidencing that the

combination is antagonic. Furthermore, the reduction percentage achieved by SB, MC

and the combination of DBNPA with SB was around 15%, which is not statistically

significant (P>0.05) with all tests performed.

The log CFU/cm2 reduction achieved by the disinfectants solutions, which was

determined through CFU enumeration, is presented in Figure 5. It is noteworthy a log

CFU/cm2 reduction of 2 for the combination of DBNPA with MC, representing a

reduction of 32% of the initial population, which is similar to the previous results. When

compared with the disinfectants used individually this result allowed a higher log

CFU/cm2 reduction than DBNPA at 10 ppm and MC at 25 000 ppm (P<0.05), but not

higher than DBNPA at 20 ppm (P>0.05). Regarding the log CFU/cm2 reduction of DBNPA

combined with SB it is clear that the action of this solution did not reach the log

reduction that the previous combination did. However, the combination allowed a

higher reduction than 37 800 ppm of SB alone (P<0.05), but at the same time it was

less efficient than DBNPA 10 ppm used individually. For this reason, this combination

(DBNPA + SB) must be considered an antagonic combination, as it was previously

demonstrated.

0

5

10

15

20

25

30

35

40

45

20 ppmDBNPA

10 ppmDBNPA

75 600 ppmSB

37 800 ppmSB

50 000 ppmMC

25 000 ppmMC

10 ppmDBNPA

+37800 ppmSB

10 ppmDBNPA +

25000 ppmMC

% R

ed

uct

ion



Figure 5. Log CFU/cm2 reduction of biofilm cells formed in 96-well microtiter plates for the individual disinfectants and for their combinations with DBNPA.

4.3.2. Efficiency of the elimination and removal of biofilm formed on coupons

in 48-well microtiter plates

The elimination and removal of biofilm formed in food surfaces is of extreme

importance due to the many negative implications in the food industry, such as

foodborne illness outbreak and spoilage of the products, which have an economic

impact. Hence, the action of the chosen disinfectants was tested in 24 h old biofilm in

three different surfaces – PS, SS and PVC. With DAPI staining it was possible to observe

and enumerate biofilm cells in the different coupons. The log CFU/cm2 is presented in

Figure 6 and an example of the results for PVC surfaces are presented in Figure 7.

Additional results are in Figure A1 to A3 of the Appendix section.

Figure 6. Log cells/cm2 removal for biofilms formed on PS, SS and PVC coupons in 48-well plates for the individual disinfectants and for the combinations with DBNPA (tests with DAPI). The line indicates the method detection limit (4.22 log cells/cm2). The symbol * represents that no cells were detected.

0.00

0.50

1.00

1.50

2.00

2.50

20 ppmDBNPA

10 ppmDBNPA

75 800 ppmSB

37 800 ppmSB

50 000 ppmMC

25 000 ppmMC

10 ppmDBNPA +

37800 ppmSB

10 ppmDBNPA +

25000 ppmMC

log

CFU

/cm

2re

du

ctio

n

0.00

1.00

2.00

3.00

4.00

5.00

6.00

7.00

8.00

Control 20 ppm DBNPA 75 600 ppm SB 50 000 ppm MC 10 ppm DBNPA+ 37 800 ppm

SB

10 ppm DBNPA+25 000 ppm

MC

log

cells

/cm

2

PS

SS

PVC

* * * * * *



Figure 7. Microscopic visualization of biofilm cells on PVC without treatment (a), treated with DBNPA at 10 ppm and SB at 37 800 ppm (b), treated with DBNPA at 20 ppm (c) and SB at 75 600 ppm (d). Magnification x1000 and scale bar 10 µm.

First of all, it is possible to visualize that all surfaces presented the same number

of cells of biofilm formed (P>0.05). Regarding the action of the disinfectants it is

immediately visible that 50 000 ppm MC and the combination of DBNPA and MC

achieved a reduction of 100% (7.1, 7.0 and 7.2 log cells reduction for PS, SS and PVC,

respectively), proving to be very good disinfectant solutions in the removal of biofilm

from the three surfaces. Furthermore, when DBNPA is used individually it is possible

to detect no significant reduction from the control for PS (P>0.05) and a small

difference for SS and PVC (P<0.05), even though DBNPA achieved in a reduction of 0.4

log cells/cm2 on PS and a 0.3 log CFU/cm2 (approximately) reduction in SS and PVC.

Considering SB alone, it achieved, in all surfaces, a small yet significant reduction

when compared with the controls (P<0.05), for a reduction of 0.5 log cells/cm2 on PS

and SS and 0.6 log cells/cm2 for PVC.

Also, regarding the combination of DBNPA with SB, no difference was observed

between this combination and the disinfectants applied individually (P>0.05).

Moreover, the log cells/cm2 reduction achieved was 0.2 for all the surfaces tested. As

expected, this combination seems to be antagonic when it comes to the removal of

biofilm from surfaces. The combination of DBNPA with MC cannot be considered

a b

c d



synergic, since although it improved significantly the reduction of biofilm when

compared with DBNPA individually (P<0.05), it achieved the same reduction with MC

individually.

Resorting to CFU enumeration, whose results are presented in Figure 8, it was

possible to confirm that PVC and PS were the surfaces that allowed a higher biofilm

formation (P>0.05), followed by SS, for a total of 7.0, 6.9 and 6.2 log CFU/cm2,

respectively. Meireles et al. (2015a) also determined that PS had a higher biofilm

development than SS.

Regarding, the effect of the disinfectants in the elimination of the biofilm cells in

the different surfaces it is noticeable that DBNPA attained a better log CFU/cm2

reduction, achieving a significant (P<0.05) log CFU/cm2 reduction of around 3.5 for the

PS and SS surfaces and 4 for PVC, when compared with the control. The disinfectant

MC was able to eliminate 2 log CFU/cm2 (approximately) for PS and SS and 3 log

CFU/cm2 for PVC. This difference between the two disinfectants was considered

statistically different (P<0.05). Furthermore, MC was more efficient in eliminating cells

on PVC surfaces (P<0.05). Comparing the results in Figure 6 and in Figure 8, it is

possible to conclude than MC is an appropriate disinfectant in biofilm removal, but it

is not suitable for the elimination of the bacterial cells, as it did not eradicate biofilm

cells. Additionally, Borges et al. (2014) and Chen and Stewart (2000) also demonstrated

that biofilm removal and elimination are distinct phenomena, as sometimes the

amount of biofilm removal and the reduction in viable cell numbers in the biofilm are

not correlated.

SB proved to be not suitable for biofilm elimination, only achieving a log CFU/cm2

reduction of 0.5 for all the surfaces. Meireles et al. (2015a) also reached a log CFU/cm2

reduction inferior to 1 for E. coli biofilms in PS surfaces and for the same SB

concentration. Conversely, the combination of SB with DBNPA proved to be efficient

in the elimination of the biofilm cells. However, it is possible that DBNPA is the only

responsible for the elimination of bacterial cells in this mixture, since SB was a poor

biofilm remover. On the other hand, the combination of DBNPA with MC proved very

successful as it achieved 100% elimination of the biofilm cells as well as 100% removal.

After 20 minutes of contact time of the disinfectants with the biofilm cells in the

coupons, these were placed in fresh MHB to try to understand if the disinfectants

eliminated the cells or simply weakened them and the cells became uncultivable.



Figure 8. Log CFU/cm2 reduction for biofilms formed on PS, SS and PVC coupons in 48-well plates for the individual disinfectants and for the combinations with DBNPA (CFU enumeration). The line indicates the method detection limit (2.66 log CFU/cm2). The symbol * represents that no CFU was detected.

For every surface and for DBNPA (20 ppm), SB (75 600 ppm) and MC (50 000 ppm)

it was obtained a regrowth of 7 log CFU/cm2 (approximately), which was expected

since there were still biofilm cells in the surfaces. For the two combinations it was

expected no regrowth since the cells were apparently completely removed, however,

only DBNPA with MC proved to be truly efficient in the completely elimination of the

biofilm cells. The combination of DBNPA with SB attained a regrowth of 6.5 log

CFU/cm2 which means that the cells were only weakened and remained viable and re-

cultivable.

4.3.3. Efficiency of the elimination and removal of biofilm formed on coupons

in an agitated reactor

From the results in 24 h old biofilm, it can be concluded that the combination of

DBNPA with MC was synergic. However, when the biofilm is older in few days it

becomes more resistant to the disinfection process.

In this regard, and due to the fact that PVC showed a higher biofilm development,

4 day old biofilm was formed on PVC coupons in an agitated reactor and the same

disinfectants were applied. The preliminary results of the CFU enumeration are

expressed in Figure 9.

0.00

1.00

2.00

3.00

4.00

5.00

6.00

7.00

8.00

Control 20 ppm DBNPA 75600 ppm SB 50000 ppm MC 10 ppm DBNPA+ 37800 ppm SB

10 ppm DBNPA+ 25 000 ppm

MC

log

CFU

/cm

2PS

SS

PVC

* * * * * *



Figure 9. Log CFU/cm2 for biofilms formed on PVC coupons in an agitated reactor for the individual disinfectants and for the combinations with DBNPA (CFU enumeration). The line indicates the method detection limit (2.66 log CFU/cm2.

After 4 days, the PVC coupons had biofilm with 7.2 log CFU/cm2, which is a very

small difference when compared with the biofilm formed for 24 h on PVC coupons.

This is probably due to the variability of conditions in the laboratory environment, such

as temperature and humidity that did not allow a higher biofilm formation. Moreover,

the biofilm is not only formed by cells, but also by EPS that was not quantified.

Furthermore, analyzing the Figure in detail it should be noted that the combination

of DBNPA and MC achieved the highest reduction in the log CFU (4 log CFU/cm2

reduction), remaining the most efficient disinfectant solution from all the ones tested,

proving to be a synergic combination when compared with MC alone (P<0.05) and

DBNPA alone (P<0.05). Considering that DBNPA 20 ppm achieved a 1.3 log CFU/cm2

reduction, which represents a difference of 3 log CFU/cm2 reduction less than in 1 day

old biofilm, it was expected that the combination of DBNPA and MC efficiency also

decreased. However, MC applied individually proved to be more efficient against 4 day

old biofilms with 3.5 log CFU/cm2 reduction, which is almost 1 log CFU/cm2 reduction

higher that in 1 day old biofilm. These results can be explained by the fact that MC is

an appropriate removal for old biofilms. Despite the fact that the biofilm had less cells,

the 4 day biofilm must had a higher EPS content and, as MC is an enzymatic agent, it

was favorable in the removal of biofilm, allowing and facilitating the DBNPA action

afterwards. Hence, this combination was considered the most appropriate.

0.00

1.00

2.00

3.00

4.00

5.00

6.00

7.00

8.00

Control 20 ppm DBNPA 75 600 ppm SB 50 000 ppm MC 10 ppm DBNPA +37 800 ppm SB

10 ppm DBNPA +25 000 ppm MC

log

CFU

/cm

2



Additionally, the combination of DBNPA and SB proved not as efficient for 4 day

old biofilm as it was for 1 day old biofilm, achieving merely 1.1 log CFU/cm2 reduction.

The fact that DBNPA was less effective and SB continued to achieve low log reductions

(only 0.5 log CFU/cm2 reduction) corroborates this inefficiency of the combination.

The combination of DBNPA and SB proved to be antagonic, once again, as the log

CFU/cm2 reduction was minor for the combination than for DBNPA alone (P<0.05).

Furthermore, the combination of disinfectants showed better results (P<0.05) than SB

applied individually, however, the difference is minimal.

4.3.4. Topography of the biofilm formed on coupons in an agitate reactor

In order to characterize the 4 day old biofilm its topography was visualized, as well

as the biofilm thickness was measured. The values of the biofilm thickness are

presented in Figure 10. Regarding the coupons for the control (only with E. coli without

treatment), a thickness of 13 µm was determined. This result is in accordance with the

studies elaborated by Janjaroen et al. (2013), where the authors observed that for a 4

week E. coli biofilm on PVC a 30 µm thickness was measured. Furthermore, it should

be added that PVS surfaces are not smooth, on the contrary they are highly roughened.

Therefore, this may be the reason for the low values of biofilm thickness measured in

the microscope, since biofilm was formed in the cavities of the surface. In Figure 11 it

is possible to see an example of the 3D structure of the biofilm in the control coupon.

Furthermore, the preliminary results indicate that SB and DBNPA + MC are equally

efficient in removing biofilm from the coupons (P>0.05), with both reaching a thickness

of around 2.7 µm (ranging 80% of removal of biofilm).

In Figure 12 it is possible to see an example of the 3D structure of the biofilm

treated with the combination of DBNPA and MC, showing a significant decrease in the

thickness when compared with the control (Figure 11).

SB proved to be better at removing 4 day old biofilm than 1 day old biofilm.

Additionally, the biofilms treated with DBNPA presented a thickness of 5.2 µm (61%

removal). When DBNPA was combined with SB the biofilm removal was not increased

(thickness of 6.4 µm - 52% removal), which is in accordance with the previous results.

In addition, the results showed that MC used individually was able to remove 73% of

the biofilm with a final thickness of 3.5 µm.



Figure 10. Thickness of biofilm formed on PVC coupons in an agitated reactor evaluated by microscopy.

Figure 11. Microscopic visualization of the biofilm topography in the E. coli control coupon, without treatment.

Figure 12. Microscopic visualization of the E. coli biofilm topography when treated with 10 ppm DBNPA and 25 000 ppm MC.

0.00

2.00

4.00

6.00

8.00

10.00

12.00

14.00

16.00

18.00

Control 20 ppm DBNPA 75 600 ppm SB 50 000 ppm MC 10 ppm DBNPA+ 37 800 ppm

SB

10 ppm DBNPA+ 25 000 ppm

MC

Bio

film

Th

ickn

ess

(µ

m)



4.3.5. Visualization of the biofilm formed on coupons in an agitated reactor

In order to understand better the topography of 4 day old biofilm in PVC coupons,

visualization in the OCT equipment was carried out. First of all, it is noticeable that

4 day old biofilm is visible macroscopically (Figure 13). Secondly, it is observable that

it has a natural tendency to form in cracks and fissures (Figure 14) as Murray (2014)

previously stated. Furthermore, analyzing Figure 13 it is possible to say that the

samples exhibited a homogeneous biofilm structure almost mushroom-like.

Figure 13. Macroscopic observation of the biofilm formed on the PVC coupons (the one on the left was visualized in OCT).

Figure 14. Visualization in the OCT of biofilm formed on the PVC coupons. The three representations are different biofilms samples on the same coupon visualized under different angles. The different color is simply to enhance contrast.



4.4. Conclusions

It was possible to conclude that regarding removal of the biofilm from the surfaces,

in all tests, SB achieved very poor results as well as in combination with DBNPA.

Concerning the elimination potential of the disinfectants, SB proved once more to be

very inefficient. In general, the combination of DBNPA with SB proved to be inefficient,

as well, except on biofilms from the coupons in 48-well plates where it achieved a

100% reduction. However, when a regrowth was performed the cells that were inactive

were only weak and with the new media added they were activated and cultivable

once again.

Regarding the effect of the disinfectant solution in 1 day old biofilm on PS, SS and

PVC coupons, the combination of DBNPA with MC proved to be synergic because even

though it attained the same log reduction (100% removal) as MC alone it attained a log

CFU/cm2 reduction of 6.5 CFU/cm2 (approximately) for all surfaces (100% kill).

The effect of the disinfectant solutions on the 1 day old and 4 days old biofilms

was not much different. The combination of DBNPA with MC proved to be once again

the best one, and SB used individually and the combination of DBNPA with SB were not

suitable for biofilm removal.

In this chapter it was also possible to conclude that PVC has a tendency to have a

higher formation of biofilm when compared to SS and PS.



Chapter 5

5. Concluding remarks and future work

5.1. General Conclusions

The sanitation of food-contact surfaces is of extreme importance in the food

industry, since pathogenic and spoilage microorganisms must be eliminated in order to

avoid foodborne illness outbreaks and economic loss. With this study it was possible to

define that DBNPA is a suitable disinfectant, since it was able to inhibit and eliminate

planktonic E. coli using a relatively low concentration (20 ppm). Furthermore, the

combination of DBNPA with MC and DBNPA with SB was proved to be antagonic against

planktonic E. coli.

Concerning the treatment of biofilms, the enzymatic disinfectant MC proved to be

a good option. This result, using an enzymatic disinfectant, adds support for the use

of the so called “green chemicals’’ in disinfectant formulations, helping the

development of green-based antimicrobial strategies. The combination of DBNPA with

MC allows a decrease in the concentration of DBNPA, a chemical disinfectant, and

achieves excellent results as it is able to remove and eliminate completely the biofilm

cells from the surfaces. However, regarding planktonic cells, the combination was not

considered synergic but additive, due to the fact that the input of MC in the

combination is mainly in the removal part of the action, as DBNPA used individually is

able to eliminate a higher log CFU/cm2 reduction than MC used individually.

Moreover, the combination of DBNPA with SB proved to be antagonic against both

planktonic and biofilm cells which is unfortunate since SB is already accepted in the

food industry as it is safe and also cost effective.



5.2. Future Work

In a future work it would be important to quantify the EPS production, since MC

was more efficient in 4 day old biofilms and a possible justification is the presence and

levels of EPS. Therefore, this aspect should be explored.

Furthermore, since the results from the agitated reactor are preliminary, more

tests have to be performed with older biofilms and formed under different process

conditions (temperature, agitation speed, nutrient concentration, etc).

In order to validate the application of the combination of DBNPA with MC in the

food-contact surfaces the next step would be to understand the mechanism of action

of this disinfectant solution and also determinate if there is production of any by-

products that may promote food spoilage or be hazardous for the process operators.

One possibility would be to test their combination in several surfaces, afterwards

sanitize fresh produce in those surfaces and then test for any differences in the

chemical composition and organoleptic characteristics of the final product.



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Faculty of Engineering of the University of Porto | A1

Appendix A1. Efficiency of the elimination and removal of biofilm formed on coupons in 48-

well plates

a)

b)

Figure A1. Microscopic visualization of MC residues after biofilm cells on PVC were treated with MC at 75 600 ppm (a) and DBNPA at 10 ppm + MC at 37 800 ppm (b). Magnification x1000 and scale bar 10 µm.

a)

b)

c)

d)

Figure A2. Microscopic visualization of biofilm cells on PS without treatment (a), treated with DBNPA at 10 ppm and SB at 37 800 ppm (b), treated with DBNPA at 20 ppm (c) and SB at 75 600 ppm (d). Magnification x1000 and scale bar 10 µm.


A2 | Faculty of Engineering of the University of Porto

a)

b)

c)

d)

Figure A3. Microscopic visualization of biofilm cells on SS without treatment (a), treated with DBNPA at 10 ppm and SB at 37 800 ppm (b), treated with DBNPA at 20 ppm (c) and SB at 75 600 ppm (d). Magnification x1000 and scale bar 10 µm.

A.2. Efficiency of the elimination and removal of biofilm formed on coupons in an

agitated reactor


Faculty of Engineering of the University of Porto | A3

Figure A4. Pump calibration curve.

Q = 58.308 Volt - 6.6733R² = 0.9983

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Q (

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A4 | Faculty of Engineering of the University of Porto

Figure A5. Agitated reactor installation.1) feed; 2) waste residues; 3) reactor with agitation; 4) feed pump; and 5) aeration pump.

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