Assessment of control strategies against Staphylococcus aureus ...

Assessment of control strategies against

Staphylococcus aureus biofilms

potentially present in the fishery industry (Evaluación de estrategias de control frente a biopelículas de Staphylococcus aureus

potencialmente presentes en la industria pesquera)

Daniel Vázquez Sánchez

International Ph.D. Thesis

DEPARTAMENTO DE MICROBIOLOGÍA Y TECNOLOGÍA DE PRODUCTOS MARINOS

DEPARTAMENTO DE BIOQUÍMICA, GENÉTICA E INMUNOLOGÍA

Assessment of control strategies against


potentially present in the fishery industry

(Evaluación de estrategias de control frente a biopelículas de

Staphylococcus aureus potencialmente presentes en la industria pesquera)

Dissertation presented by Daniel Vázquez Sánchez to aim for the

International Ph.D. by the University of Vigo

Memoria presentada por Daniel Vázquez Sánchez para optar al título de

Doctor Internacional por la Universidad de Vigo

Vigo, Febrero de 2014

DR. PALOMA MORÁN MARTÍNEZ, CATEDRÁTICA DEL DEPARTAMENTO DE

BIOQUÍMICA, GENÉTICA E INMUNOLOGÍA DE LA UNIVERSIDAD DE VIGO,

INFORMA:

Que la presente Tesis Doctoral titulada “Assessment of control strategies against

Staphylococcus aureus biofilms potentially present in the fishery industry (Evaluación

de estrategias de control frente a biopelículas de Staphylococcus aureus potencialmente

presentes en la industria pesquera)” presentada por el Licenciado en Biología D. Daniel

Vázquez Sánchez para optar al Grado de Doctor por la Universidad de Vigo, fue

realizada bajo la dirección del Dr. Juan José Rodríguez Herrera y de la Dra. Marta

López Cabo en el Departamento de Microbiología y Tecnología de Productos Marinos

perteneciente al Instituto de Investigaciones Marinas (IIM) del Consejo Superior de

Investigaciones Científicas (CSIC).

Que el trabajo realizado tiene una base experimental suficiente y que supone una

contribución relevante en el ámbito de la microbiología y seguridad alimentaria.

Que la presente Tesis Doctoral reúne los requisitos necesarios para la obtención de la

mención de Tesis Internacional.

Que la Tesis Doctoral cumple los requisitos necesarios para presentarla en la modalidad

de “compendio de artículos”, debido a la coherencia argumental de los cinco artículos

incluidos en ella y que tres de los artículos científicos presentados han sido publicados

en revistas indexadas en el Journal Citations Report, publicado por Thomson Reuters.

En todos ellos, el doctorando ha contribuido de manera esencial junto con los coautores.

Por todo lo anterior autorizo su presentación ante el Tribunal correspondiente.

Y para que así conste a los efectos oportunos, se expide la presente en Vigo a 4 de

Febrero de 2014,

Asdo.: Dra. Paloma Morán Martínez

DR. JUAN JOSÉ RODRÍGUEZ HERRERA Y DRA. MARTA LÓPEZ CABO,

CIENTÍFICOS TITULARES DEL DEPARTAMENTO DE MICROBIOLOGÍA Y

TECNOLOGÍA DE PRODUCTOS MARINOS DEL INSTITUTO DE

INVESTIGACIONES MARINAS (IIM-CSIC),

INFORMAN:

Que la presente Tesis Doctoral titulada “Assessment of control strategies against

Staphylococcus aureus biofilms potentially present in the fishery industry (Evaluación

de estrategias de control frente a biopelículas de Staphylococcus aureus potencialmente

presentes en la industria pesquera)” presentada por el Licenciado en Biología D. Daniel

Vázquez Sánchez para optar al Grado de Doctor por la Universidad de Vigo, fue

realizada bajo nuestra dirección.

Que el trabajo realizado tiene una base experimental suficiente y que supone una

contribución relevante en el ámbito de la microbiología y seguridad alimentaria.

Que la presente Tesis Doctoral reúne los requisitos necesarios para la obtención de la

mención de Tesis Internacional.

Que la Tesis Doctoral cumple los requisitos necesarios para presentarla en la modalidad

de “compendio de artículos”, debido a la coherencia argumental de los cinco artículos

incluidos en ella y que tres de los artículos científicos presentados han sido publicados

en revistas indexadas en el Journal Citations Report, publicado por Thomson Reuters.

En todos ellos, el doctorando ha contribuido de manera esencial junto con los coautores.

Por todo lo anterior autorizamos su presentación ante el Tribunal correspondiente.

Y para que así conste a los efectos oportunos, se expide la presente en Vigo a 31 de

Enero de 2014,

Asdo.: Dr. Juan José Rodríguez Herrera Asdo.: Dra. Marta López Cabo

Este trabajo ha sido financiado por:

El proyecto PGIDIT 07 TAL 014402PR concedido por la Xunta de Galicia.

La beca predoctoral JAE-predoc concedida por el Consejo Superior de Investigaciones

Científicas (CSIC) a Daniel Vázquez Sánchez.

La beca para estancias en centros de investigación concedida por el CSIC a Daniel

Vázquez Sánchez para su estancia en el centro Nofima en Ås (Noruega).

"La vida es una unión simbiótica y cooperativa que permite triunfar a los

que se asocian”

Lynn Margulis (1938-2011)

Bióloga estadounidense y Doctora Honoris Causa por la Universidad de Vigo

A Vero

Índice / Contents

1

Índice / Contents

Resumen ....................................................................................................... 7

Abstract ...................................................................................................... 15

General Introduction ................................................................................ 23

1. Staphylococcus aureus ........................................................................................ 25

1.1. History .......................................................................................................... 25

1.2. Phylogeny ..................................................................................................... 25

1.3. General characteristics .................................................................................. 27

1.4. Pathogenicity ................................................................................................ 28

1.5. Pathogenic determinants ............................................................................... 29

2. Biofilm formation by Staphylococcus aureus .................................................... 32

2.1. Definition of biofilm ..................................................................................... 32

2.2. Significance of biofilm formation ................................................................ 32

2.3. The process of biofilm development ............................................................ 33

2.4. Ecological advantages of biofilm formation................................................. 34

2.4.1. Cell-cell communication ......................................................................... 34

2.4.2. Increased resistance to external stimulus ................................................ 35

2.4.3. Increased dispersal capacity .................................................................... 36

2.5. Staphylococcus aureus biofilms: development, composition and

regulation ...................................................................................................... 36

3. Incidence of Staphylococcus aureus in the food industry .................................. 38

3.1. Outbreaks ...................................................................................................... 38

3.2. Foods implicated in staphylococcal poisonings ........................................... 43

3.3. Food contamination by Staphylococcus aureus ............................................ 44

4. Control of Staphylococcus aureus in the food industry ...................................... 45

4.1. Hazard Analysis and Critical Control Points (HACCP) ............................... 45

4.2. Prevention strategies against biofilm formation ........................................... 45

4.3. Cleaning procedures ..................................................................................... 46

4.4. Disinfection treatments ................................................................................. 47

Justificación y Objetivos ........................................................................... 55

Justification and Objectives ..................................................................... 61

Índice / Contents

2

Chapter 1. Incidence and characterization of Staphylococcus aureus

in fishery products marketed in Galicia (Northwest Spain) ................. 67

Abstract ....................................................................................................................... 69

1.1 Introduction ......................................................................................................... 70

1.2 Materials and Methods ....................................................................................... 71

1.2.1 Sampling ....................................................................................................... 71

1.2.2 Isolation and identification of S. aureus ....................................................... 72

1.2.3 RAPD ............................................................................................................ 73

1.2.4 Detection of sea-see and seg-sei genes ......................................................... 74

1.2.5 Antibiotic susceptibility test ......................................................................... 76

1.2.6 Detection of blaZ and mecA genes ............................................................... 77

1.3 Results ................................................................................................................. 78

1.3.1 Incidence in fishery products ........................................................................ 78

1.3.2 RAPD-PCR ................................................................................................... 80

1.3.3 Presence of enterotoxin genes ...................................................................... 83

1.3.4 Antibiotic sensitivity ..................................................................................... 84

1.4 Discussion ........................................................................................................... 87

1.5 Conclusions ......................................................................................................... 93

Chapter 2. Impact of food-related environmental factors on the

adherence and biofilm formation of natural Staphylococcus aureus

isolated from fishery products ................................................................. 95

Abstract ....................................................................................................................... 97

2.1 Introduction ......................................................................................................... 98

2.2 Materials and Methods ..................................................................................... 100

2.2.1 Bacterial strains and growth conditions ...................................................... 100

2.2.2 Evaluation of bacterial cell surface physicochemical properties ................ 100

2.2.3 Measurement of the adherence ability to polystyrene at different ionic

strength conditions ...................................................................................... 101

2.2.4 Quantification of biofilm formation on polystyrene under different

environmental conditions............................................................................ 101

2.2.5 Transcriptional analysis .............................................................................. 102

2.2.6 Statistical analysis ....................................................................................... 103

2.3 Results ............................................................................................................... 105

2.3.1 Surface hydrophobicity and electron donor/acceptor character ................. 105

2.3.2 Adherence ability of S. aureus to polystyrene surfaces .............................. 105

Índice / Contents

3

2.3.3 Biofilm formation on polystyrene surfaces under different

environmental conditions............................................................................ 108

2.3.3.1 Effect of incubation temperature .......................................................... 108

2.3.3.2 Effect of glucose and NaCl addition ..................................................... 108

2.3.3.3 Effect of MgCl2 addition ....................................................................... 111

2.3.4 Multivariate analysis of the physicochemical, adhesion and biofilm-

forming properties of the 28 S. aureus strains ............................................ 112

2.3.5 Gene expression in relation to biofilm formation ....................................... 114

2.4 Discussion ......................................................................................................... 116

2.5 Conclusions ....................................................................................................... 119

Appendix 1: PCR-detection of icaA and icaD genes ................................................ 121

Appendix 2: PCR-detection of bap gene .................................................................. 122

Chapter 3. Biofilm-forming ability and resistance to industrial

disinfectants of Staphylococcus aureus isolated from fishery

products .................................................................................................... 125

Abstract ..................................................................................................................... 127

3.1. Introduction ....................................................................................................... 128

3.2. Material and Methods ....................................................................................... 129

3.2.1. Bacterial strains and culture conditions ...................................................... 129

3.2.2. Conditions for biofilm formation................................................................ 129

3.2.3. Biofilm formation assays ............................................................................ 131

3.2.3.1. Slime production on Congo red agar (CRA) ........................................ 131

3.2.3.2. Initial adherence .................................................................................... 131

3.2.3.3. Quantification of biofilm formation ...................................................... 131

3.2.3.4. Determination of growth kinetics ......................................................... 132

3.2.4. Biocide resistance assays ............................................................................ 132

3.2.4.1. Minimal biofilm eradication concentration (MBEC) ........................... 133

3.2.4.2. Minimal bactericidal concentration (MBC) .......................................... 133

3.2.5. Statistical analysis ....................................................................................... 134

3.3. Results ............................................................................................................... 134

3.3.1. Biofilm-forming ability of S. aureus .......................................................... 134

3.3.1.1. Detection of biofilm-forming ability on Congo red agar (CRA) .......... 134

3.3.1.2. Initial adhesion studies .......................................................................... 135

3.3.1.3. Quantification of biofilm biomass ........................................................ 135

3.3.2. Resistance to industrial disinfectants of S. aureus...................................... 137

3.3.2.1. Effectiveness of benzalkonium chloride ............................................... 137

Índice / Contents

4

3.3.2.2. Effectiveness of peracetic acid and sodium hypochlorite ..................... 141

3.4. Discussion ......................................................................................................... 143

3.5. Conclusions ....................................................................................................... 147

Appendix: Growth kinetics ....................................................................................... 149

Chapter 4. Single and sequential application of electrolyzed water

with benzalkonium chloride or peracetic acid for removal of

Staphylococcus aureus biofilms .............................................................. 153

Abstract ..................................................................................................................... 155

4.1. Introduction ....................................................................................................... 156


4.2.1. Bacterial strains .......................................................................................... 157

4.2.2. Antibacterial agents .................................................................................... 157


4.2.4. Single application of electrolyzed water (EW) ........................................... 159

4.2.5. Sequential application treatments ............................................................... 160


4.3. Results ............................................................................................................... 162

4.3.1. Single application of electrolyzed water (EW) ........................................... 162

4.3.1.1. Effects of pH ......................................................................................... 162

4.3.1.2. Effects of active chlorine concentration ............................................... 163

4.3.1.3. Effects of exposure time ....................................................................... 163

4.3.2. Double sequential application of NEW ...................................................... 164

4.3.3. Sequential application of NEW and BAC .................................................. 166

4.3.4. Sequential application of NEW and PAA .................................................. 168

4.4. Discussion ......................................................................................................... 170

4.5. Conclusions ....................................................................................................... 173

Appendix ................................................................................................................... 175

Chapter 5. Antimicrobial activity of essential oils against

Staphylococcus aureus biofilms .............................................................. 181

Abstract ..................................................................................................................... 183

5.1. Introduction ....................................................................................................... 184


5.2.1. Bacterial strain ............................................................................................ 185

5.2.2. Antibacterial agents .................................................................................... 185


Índice / Contents

5

5.2.4. Biocide resistance assays ............................................................................ 187

5.2.4.1. Resistance of planktonic cells ............................................................... 187

5.2.4.2. Resistance of biofilms ........................................................................... 187

5.2.4.3. Determination of growth kinetics ......................................................... 189


5.3. Results ............................................................................................................... 190

5.3.1. Effectiveness of essential oils (EOs) against planktonic cells .................... 190

5.3.2. Effectiveness of EOs against 48-h-old biofilms ......................................... 191

5.3.3. Effects of sub-lethal doses of thyme oil on bacterial growth ..................... 192

5.3.4. Effectiveness of thyme oil against biofilms formed under sub-lethal

doses of thyme oil ....................................................................................... 194

5.3.5. Effectiveness of benzalkonium chloride (BAC) against biofilms formed

under sub-lethal doses of thyme oil ............................................................ 195

5.4. Discussion ......................................................................................................... 196

5.5. Conclusions ....................................................................................................... 199

General Discussion .................................................................................. 203

Conclusiones Generales .......................................................................... 211

General Conclusions ............................................................................... 217

Bibliografía / References ......................................................................... 223

List of original publications .................................................................... 251

Agradecimientos / Acknowledgements .................................................. 253

7

Resumen

Resumen

9

Resumen

Staphylococcus aureus es uno de los principales agentes etiológicos de

intoxicaciones alimentarias en el mundo, debido a la ingestión de alimentos con

enterotoxinas. España es uno de los principales productores y consumidores de

productos pesqueros en la Unión Europea. Sin embargo, S. aureus se detecta

reiteradamente en estos alimentos como consecuencia de la contaminación cruzada con

manipuladores y superficies de contacto con el alimento. La capacidad de formar

biopelículas de S. aureus le proporciona una alta tolerancia a los antimicrobianos,

permitiéndole una persistencia en ambientes alimentarios a largo plazo. Las nuevas

tendencias en la producción de los alimentos (procesado mínimo, producción masiva,

globalización) han introducido además nuevos escenarios que pueden favorecer la

presencia y proliferación de S. aureus. Un gran incremento de cepas resistentes a

antibióticos ha sido detectado también en ámbitos extra-hospitalarios, incluido el

alimentario, lo que puede favorecer la transmisión de resistencias a la microbiota

humana a través de los alimentos ingeridos y causar infecciones difíciles de tratar. Por

tanto, la finalidad del presente trabajo consistió en mejorar el control de S. aureus en la

industria alimentaria (en particular, la pesquera) a través de la identificación de los

escenarios de contaminación de mayor riesgo y de la evaluación de prometedoras

estrategias de desinfección frente a este patógeno.

El primer objetivo fue pues la estimación de la incidencia de S. aureus en diferentes

productos pesqueros comercializados. Un total de 298 productos pesqueros de distinto

origen y tipo de procesado fueron muestreados. Los aislados fueron identificados como

S. aureus mediante específicas pruebas bioquímicas (producción de coagulasa y

ADNasa, fermentación del manitol) y genéticas (secuenciación del 23s ADNr), y

caracterizadas por RAPD-PCR con tres cebadores (AP-7, ERIC-2 y S). Asimismo, se

evaluó la capacidad de todos los aislados de producir enterotoxinas (es decir, presencia

de genes se) y su resistencia a varios antibióticos.

S. aureus fue detectado en una proporción significativa de productos (~ 25%), siendo

destacable en productos frescos (43%) y congelados (30%). Además, una proporción

significativa de ahumados, surimis, huevas y otros productos listos para consumir no

cumplieron con los límites legales vigentes. Los aislados mostraron 33 patrones

característicos, y cada uno fue atribuido a un clon bacteriano particular. No se encontró

Resumen

10

relación entre los patrones de RAPD y las categorías de productos. La mayoría de los

aislados (88%) son portadores del gen sea. Los recuentos de cepas con capacidad

enterotoxigénica mostraron que en 17 productos se alcanzaron niveles de riesgo. No se

encontró relación entre la presencia de genes se y los patrones RAPD. Todos los

aislados fueron resistentes a la penicilina, cloranfenicol y ciprofloxacina, y muchos a la

tetraciclina (82.4%), pero no se detectaron MRSA. De hecho, ninguna cepa portaba el

gen mecA, relacionado con la resistencia a antibióticos beta-lactámicos como la

meticilina.

Posteriormente, se examinó la prevalencia en instalaciones de procesado de pescado

de las cepas de S. aureus con capacidad enterotoxigénica mediante el estudio de su

capacidad para formar biopelículas y su resistencia a desinfectantes.

Todas las cepas fueron descritas como S. aureus capaces de producir

exopolisacáridos (fenotipo positivo en agar rojo Congo y portadores de los genes icaA e

icaD), pero ninguna portaba los genes bap (expresan la proteína accesoria de la

biopelícula). La mayoría mostraron una capacidad para formar biopelículas mayor que

S. aureus ATCC 6538 -cepa de referencia en los métodos bactericidas estandarizados-

sobre materiales habituales de las superficies de contacto (acero inoxidable,

poliestireno) y bajo diferentes condiciones ambientales (temperatura, contenido de

nutrientes, osmolaridad) potencialmente presentes en las plantas de procesado. En

general, la adherencia inicial de S. aureus incrementa bajo condiciones de alta fuerza

iónica, mientras que la formación de biopelículas es significativamente favorecida por la

presencia de glucosa (por ejemplo, como aditivo en surimis y ahumados), pero

moderadamente con cloruro sódico o magnesio (por ejemplo, residuos de agua de mar y

productos pesqueros). No obstante, el análisis transcriptacional de genes relacionados

con la formación de biopelículas (icaA, sarA, rbf y σB) mostró una alta variabilidad

entre las cepas en respuesta a estas condiciones ambientales. Por otra parte, parece que

el procesado del alimento pudo haber generado una presión selectiva ya que cepas con

una alta capacidad para formar biopelículas fueron aisladas de productos altamente

procesados y manipulados.

Las biopelículas formadas por todas las cepas mostraron una marcada resistencia a

desinfectantes aplicados habitualmente en la industria alimentaria (cloruro de

benzalconio o BAC, ácido peracético o PAA, hipoclorito sódico o NaClO), siendo

Resumen

11

mayor que la de ATCC 6538 en muchos casos. Como era de esperar, la resistencia de

las biopelículas de S. aureus fue significativamente mayor que la de las células

planctónicas en todos los casos. Pero no se encontró correlación entre la resistencia de

las biopelículas y las células planctónicas al BAC, PAA y NaClO, por lo que ninguna

extrapolación parece posible. Sin embargo, la mayoría de los métodos bactericidas

estándar usados en la Unión Europea están basados en cultivos en suspensión, y sólo

EN 13697 se basa en biopelículas, pero no parece simular realmente las condiciones

ambientales que se encuentran en la industria alimentaria. La resistencia antimicrobiana

aumentó con el desarrollo de la biopelícula. Además, la formación de biopelículas

parece atenuar el efecto de las bajas temperaturas sobre la resistencia al BAC. El PAA

fue el más efectivo frente a biopelículas y células planctónicas, seguido del NaClO y

BAC. Pero la resistencia de las cepas no siguió el mismo orden para cada biocida, lo

cual muestra la actual limitación de usar un reducido número de cepas de colección (y

sólo un S. aureus) en los métodos estándar para asegurar una apropiada aplicación de

los desinfectantes. En consecuencia, las dosis recomendadas por los fabricantes para

desinfectar superficies de contacto con el alimento mediante la aplicación de BAC,

PAA y NaClO fueron menores que las obtenidas en este estudio, por lo que no

garantizan la eliminación de las biopelículas. Por tanto, los microorganismos podrían

estar expuestos a dosis sub-letales de desinfectante, lo cual puede generar la emergencia

de resistencias antimicrobianas.

Por esta razón, este trabajo se centró a continuación en el estudio de la eficacia y

aplicabilidad de innovadoras estrategias de desinfección más respetuosas con el medio

ambiente para el control de biopelículas de S. aureus en plantas de procesado de

pescado. En particular, se investigó la actividad bactericida del agua electrolizada (EW)

y de diversos aceites esenciales (EOs), así como tratamientos combinados con BAC o

PAA. Las cuatro cepas de S. aureus con mayor potencial de prevalencia en plantas de

procesado del alimento y con una alta incidencia en productos pesqueros (St.1.01,

St.1.04, St.1.07 y St.1.08) fueron evaluados.

En el caso del EW, su eficacia frente a biopelículas apenas se ve afectada por

variaciones en el pH de producción. El EW neutra (NEW) fue por tanto usada

posteriormente porque tiene un mayor potencial de aplicación a largo plazo que el EW

ácida (debido a una menor corrosividad y toxicidad) y al mayor rendimiento de la

Resumen

12

unidad de producción a pH neutro. La aplicación de NEW causó una alta reducción en

el número de células viables en la biopelícula inicialmente. Sin embargo, se necesitó

una alta concentración de cloro activo (800 mg/L ACC) para alcanzar la reducción

logarítmica (LR) exigida en el método estándar EN 13697 (≥ 4 log CFU/cm2 tras 5

min). Una aplicación secuencial doble de NEW a más bajas concentraciones durante 5

min cada una permitió alcanzar una LR ≥ 4 log CFU/cm2 en la mayoría del rango

experimental. Aplicaciones secuenciales de NEW con BAC o PAA mostraron un efecto

similar, con PAA-NEW siendo la secuencia más efectiva. La combinación de NEW con

otros tratamientos antimicrobianos puede ser por tanto una alternativa eficaz a los

protocolos de desinfección tradicionalmente usados en la industria alimentaria.

Por otra parte, se determinó la efectividad de 19 EOs (anis, zanahoria, citronela,

cilantro, comino, Eucalyptus globulus, Eucalyptus radiata, hinojo, geranio, jengibre,

hisopo, limoncillo, mejorana, palmarosa, pachuli, salvia, árbol del té, tomillo y vetiver)

frente a células planctónicas de S. aureus St.1.01. Las células planctónicas mostraron

una gran variabilidad en la resistencia a EOs, siendo el aceite de tomillo el más eficaz,

seguido de los aceites de limoncillo y luego vetiver. Los 8 aceites más efectivos frente a

células planctónicas fueron a continuación testados frente a biopelículas formadas en

acero inoxidable durante 48 h. Todos los EOs redujeron significativamente el número

de células viables en la biopelícula, aunque ninguno consiguió eliminar totalmente la

biopelícula. Aceites de tomillo y pachuli fueron los más efectivos, pero se necesitaron

altas concentraciones para alcanzar LR por encima de 4 log CFU/cm2 tras 30 min de

exposición. El uso de dosis sub-letales de aceite de tomillo previno la formación de la

biopelícula y mejoró la eficacia del tomillo y BAC frente a biopelículas. Sin embargo,

cierta adaptación celular al tomillo fue detectada. Tratamientos basados en EOs deben

por tanto basarse en la rotación y combinación de distintos EOs o con otros biocidas

para prevenir la emergencia de cepas resistentes a los antimicrobianos. Tratamientos

combinados con tomillo pueden ser una alternativa eficaz, respetuosa con el

medioambiente y relativamente barata para controlar la formación de biopelículas de S.

aureus en instalaciones de procesado de alimentos.

15

Abstract

Abstract

17

Abstract

Staphylococcus aureus is one of the major bacterial agents causing foodborne

diseases in humans worldwide, due to the ingestion of food containing staphylococcal

enterotoxins. Spain is one of the largest producers and consumers of fishery products in

the European Union. However, S. aureus is repeatedly detected in these products as a

consequence of cross-contamination from food handlers and food contact surfaces.

Biofilm formation also provides S. aureus a high tolerance to biocides allowing a long-

term persistence of this pathogen in food-related environments. Novel trends in food

production (e.g. minimal processing, mass production, globalization) have additionally

introduced new scenarios that can enhance the presence and subsequent growth of S.

aureus. Moreover, an increased number of antibiotic-resistant S. aureus has been

detected in non-clinical ambits, including the food industry, which can lead to the

transmission of resistances to the human microbiome through the ingested food and

causing infections hard to be treated. Therefore, this work was aimed to improve the

control of S. aureus in the food industry (particularly, in fisheries) through the

identification of the most risky scenarios of contamination and the evaluation of

promising disinfection strategies against this pathogen.

The first objective was thus the assessment of the incidence of S. aureus in different

commercialized fishery products. A total of 298 fishery products of different origin and

type of processing were sampled. Isolates were identified as S. aureus by specific

biochemical (coagulase, DNAse and mannitol fermentation) and genetic tests (23s

rDNA sequencing), and characterized by RAPD-PCR with three primers (AP-7, ERIC-2

and S). In addition, the enterotoxin-producing ability (i.e. presence of se genes) and the

resistance to several antibiotics were also evaluated for all isolates.

S. aureus was detected in a significant proportion of products (~ 25%), being the

highest incidence in fresh (43%) and frozen products (30%). In addition, a significant

proportion of smoked fish, surimis, fish roes and other ready-to-eat products did not

comply with legal limits in force. Isolates displayed 33 fingerprint patterns, and each

one was attributed to a single bacterial clone. Cluster analysis based on similarity values

between RAPD fingerprints did not find relationship between any RAPD pattern and

any product category. Most isolates (88%) were found to be sea positive. Putative

Abstract

18

enterotoxigenic strains counts reached high risk levels in 17 products. No relationship

was found between the presence of se genes and RAPD patterns. All isolates were

resistant to penicillin, chloramphenicol and ciprofloxacin, and most to tetracycline

(82.4%), but MRSA were not detected. In fact, none strain carried mecA gen, which is

related to the resistance to beta-lactam antibiotics such as methicillin.

Subsequently, the prevalence in fishery-processing facilities of putative

enterotoxigenic S. aureus strains was examined by studying their biofilm-forming

ability and disinfectant resistance.

All strains were described as S. aureus able to produce exopolysaccharides (positive

phenotype in red Congo agar and icaA- and icaD-carriers), but none carried bap gene

(expression of biofilm accessory protein). Most strains showed a biofilm-forming ability

higher than S. aureus ATCC 6538 -reference strain in bactericidal standard tests- on

common food-contact surface materials (stainless steel, polystyrene) and under different

environmental conditions (temperature, nutrient content, osmolarity) potentially present

in processing plants. In general, initial adhesion of S. aureus was increased by the

presence of high ionic strength conditions, whereas biofilm formation was significantly

promoted by the presence of glucose (e.g., additive in surimis and smoked fish), but

moderately by sodium chloride or magnesium (e.g., wastes of seawater and seafood).

Nevertheless, transcriptional analysis of genes related with biofilm formation (icaA,

sarA, rbf and σB) showed a high variability between strains in the response to these

environmental conditions. Moreover, it seems that food-processing could have produced

a selective pressure and strains with a high biofilm-forming ability were more likely to

be found in highly handled and processed products.

Biofilms formed by all strains showed a marked resistance to disinfectants applied

commonly in the food industry (benzalkonium chloride or BAC, peracetic acid or PAA,

sodium hypochlorite or NaClO), being higher than ATCC 6538 in most cases. As

expected, the resistance of S. aureus biofilms was significantly higher than that of

planktonic cells in all cases. But no correlation was found between the resistance of

biofilms to BAC, PAA and NaClO and that of planktonic cells, so no extrapolation

seems thus feasible. However, most standard bactericidal tests used in the European

Union are based in suspension cultures, and only EN 13697 is biofilm-based, but it does

not seem to truly simulate environmental conditions found in the food industry. The

Abstract

19

antimicrobial resistance increased as biofilm aged. Biofilm formation also seemed to

attenuate the effect of low temperatures on BAC resistance. PAA was found to be most

effective against both biofilms and planktonic cells, followed by NaClO and BAC. But

the resistance of strains did not follow the same order for each biocide, which shows the

present limitation of using a few type strains (and only one S. aureus) in standard tests

in order to ensure a proper application of disinfectants. Consequently, doses

recommended by manufacturers for BAC, PAA and NaClO to disinfect food-contact

surfaces were lower than data obtained in this study, so they are not able to guarantee

biofilm removal. Microorganisms could therefore be exposed to sub-lethal doses of

disinfectants and this could generate the emergence of antimicrobial resistance.

For this reason, this work was then focused in the study of the efficacy and

applicability of innovative and more environmentally-friendly disinfection strategies to

control S. aureus biofilms on fishery-processing plants. Particularly, it was investigated

the bactericidal activity of electrolyzed water (EW) and a range of essential oils (EOs),

as well as combined treatments with BAC or PAA. The four S. aureus strains with the

highest potential prevalence in food-processing plants and with a high incidence in

fishery products (St.1.01, St.1.04, St.1.07 and St.1.08) were evaluated.

In the case of EW, its efficacy against biofilms was hardly any affected by variations

in the pH of production. Neutral EW (NEW) was therefore used in subsequent studies

as it has a higher potential for long-term application than acidic EW (due to a lower

corrosiveness and toxicity) and due to the higher yield rate of the production unit at

neutral pH. The application of NEW caused a high reduction in the number of viable

biofilm cells initially. However, a high available chlorine concentration (800 mg/L

ACC) was needed to achieve logarithmic reductions (LR) demanded by the European

quantitative surface test of bactericidal activity (≥ 4 log CFU/cm2 after 5 min). A double

sequential application of NEW at much lower concentrations for 5 min each allowed LR

≥ 4 log CFU/cm2 to be reached in most of the experimental range. Sequential

applications of NEW and either BAC or PAA showed a similar effect, with PAA-NEW

being most effective. The combination of NEW with other antimicrobial treatments can

thus be an effective alternative to disinfection protocols traditionally used in the food

industry.

Abstract

20

Otherwise, the effectiveness of nineteen EOs (anise, carrot, citronella, coriander,

cumin, Eucalyptus globulus, Eucalyptus radiata, fennel, geranium, ginger, hyssop,

lemongrass, marjoram, palmarosa, patchouli, sage, tea-tree, thyme and vetiver) was

assessed against planktonic cells of S. aureus St.1.01. Planktonic cells showed a wide

variability in resistance to EOs, with thyme oil as the most effective, followed by

lemongrass oil and then vetiver oil. The eight EOs most effective against planktonic

cells were subsequently tested against 48-h-old biofilms formed on stainless steel. All

EOs reduced significantly the number of viable biofilm cells, but none of them could

remove biofilms completely. Thyme and patchouli oils were the most effective, but high

concentrations were needed to achieve LR over 4 log CFU/cm2 after 30 min exposure.

The use of sub-lethal doses of thyme oil prevented biofilm formation and enhanced the

efficiency of thyme oil and BAC against biofilms. However, some cellular adaptation to

thyme oil was detected. Therefore, EO-based treatments should be based on the rotation

and combination of different EOs or with other biocides to prevent the emergence of

antimicrobial-resistant strains. Combined thyme oil-based treatments can be an

effective, environmentally-friendly, safe-to-use and relatively inexpensive alternative to

control the formation of S. aureus biofilms on food-processing facilities.

23

General Introduction


25


1. Staphylococcus aureus

1.1. History

Staphylococcus aureus was discovered in Aberdeen (Scotland) in 1880 by A.

Ogston, who described grape-like clusters of bacteria in stained slide preparations of

pus from patients with post-operative wound infections and abscesses (Ogston, 1882).

Four years later, A.J. Rosenbach achieved pure cultures of this pathogen on solid media

and named it as Staphylococcus aureus for the golden appearance of the colonies

(Rosenbach, 1884).

In 1884, V.C. Vaughan and J.M. Sternberg were the first in associate a food

poisoning outbreak in Michigan (USA) with the ingestion of cheese contaminated by

staphylococci. But this link was not confirmed until 1914 by M.A. Barber, who showed

that consuming milk from a cow with staphylococcal mastitis caused illness (Barber,

1914). Later, Dack et al. (1930) demonstrated that staphylococcal food poisonings were

caused by a toxin and not by the microorganism itself. Despite the great advances in

food safety, Staphylococcus aureus is still a major human pathogen capable of causing

food poisoning outbreaks worldwide.

1.2. Phylogeny

Staphylococcus aureus is a member of the monophyletic genus Staphylococcus,

which comprises Gram-positive bacteria of low DNA G + C content (32-36%)

belonging to the family Staphylococcaceae, order Bacillales, class Bacilli, phylum

Firmicutes (Schleifer and Bell, 2009). As shown in Figure 1, this genus is closely

related to bacilli and other Gram-positive bacteria with low DNA G + C content such as

enterococci, streptococci, lactobacilli and listeria (Götz et al., 2006). Although S. aureus

is the responsible to cause most of foodborne intoxications reported, other species and

subspecies of the genus Staphylococcus have also shown a real and potential risk for the

human health as they are able to produce coagulase, nuclease and/or enterotoxins

(Figure 2).


26

Figure 1. Maximum likelihood

phylogram of the genus

Staphylococcus into the order

Bacillales based on 16S rRNA

analysis performed by Götz et al.

(2006). The bar length indicates 5%

estimated sequence divergence.

Figure 2. Inference of the staphylococcal phylogeny using Bayesian estimation of species trees

(BEST) methodology on 16S rRNA and dnaJ gene fragments (modified from Lamers et al.,

2012). The bar length indicates a sequence divergence of 0.1 substitutions per site.

Staphylococcal species and subspecies of real and potential risk in foodborne intoxications are

marked with an asterisk (data from Jay et al., 2005).


27

1.3. General characteristics

Staphylococcus aureus is a Gram-positive facultative anaerobe of 0.5-1.0 μm in

diameter, which grows individually, in pairs, short chains or grape-like clusters (Figure

3A) (Schleifer and Bell, 2009). Macroscopically, S. aureus form colonies in agar

medium of 6-8 mm in diameter, rounded and smooth, glistening, translucent, with entire

margins, with pigmentations varying from grey to golden yellow to orange (Figure 3B).

Figure 3A-B. Morphology of S. aureus under scan electron microscopy (A) (from the Centers

for Disease Control and Prevention's Public Health Image Library, ID#:6486) and S. aureus

colonies on tryptic soy agar (B).

S. aureus is a ubiquitous bacteria detected in numerous environmental sources (e.g.

soil, air, water, dust, sand, organic wastes, paper, clothing, furniture, blankets, carpets,

linens, utensils, vegetal surfaces), though warn-blooded animals are the main reservoirs.

As shown in Figure 4, S. aureus form part of the normal human microbiome associated

with an asymptomatic commensal colonization of skin, throat, nose, hair, nails, axillae

and perineum (Wertheim et al., 2005).

S. aureus can grow at a wide range of temperatures (6-48ºC, optimal growth at 35-

41ºC), pHs (4-10, optimum at 6-7), water activities (aw = 0.83 ≥ 0.99, optimum at 0.99)

and salt content (0-20%, optimum at 0%), and it is also tolerant to desiccation

(Hennekinne et al., 2012; Schelin et al., 2011). Nutrients required for bacterial growth

are achieved by the secretion of diverse enzymes (e.g. thermonucleases, proteases,

lipases, hyaluronidases, catalases, collagenases) and cytolytic exotoxins such as

haemolysins (DeLeo et al., 2009; Dinges et al., 2000). Thus, S. aureus can metabolize

aerobically fructose, galactose, glucose, glycerol, lactose, maltose, mannitol, mannose,

ribose, trehalose, turanose and sucrose generating acid, whereas anaerobically produces

D- and L-lactate from glucose (Schleifer and Bell, 2009).


28

Figure 4. Colonization sites of S. aureus in the human body (Wertheim et al., 2005).

1.4. Pathogenicity

S. aureus is the most human-pathogenic species in the genus Staphylococcus. It acts

as an opportunistic bacterial pathogen in human carriers, who might also serve as spread

vectors (Chambers and DeLeo, 2009). S. aureus can infect individuals through skin

injuries (e.g. acne, styes, burns, wounds) and foreign bodies (e.g. sutures, intravenous

lines, prosthetic devices), or associated to the infection with other pathogenic agents

(e.g. viruses), chronic underlying diseases (e.g. cancer, alcoholism) or heart diseases

(Lowy, 1998; Moreillon and Que, 2004; Van-Belkum and Melles, 2005). As shown in

Figure 5, S. aureus can cause from relatively minor skin infections to life-threatening

systemic illnesses (Wertheim et al., 2005). Although staphylococcal infections do not

necessary impedes the infected person from working, it implies a serious risk of cross-

contamination in the case of food-handlers.


29

Figure 5. Major infections caused by S. aureus in humans (Wertheim et al., 2005).

Besides in humans, S. aureus is also capable of producing diverse infections in

animals (e.g. mastitis, synovitis, arthritis, endometritis, furuncles, suppurative

dermatitis, pyaemia, and septicaemia), which may have considerable economic losses in

the food industry (Schleifer and Bell, 2009).

1.5. Pathogenic determinants

S. aureus carries a wealth of extracellular and cell-wall-associated virulence factors

(specified in Figure 6), which are encoded in phages, plasmids, pathogenicity islands

and in the staphylococcus cassette chromosome. They are coordinately expressed during

the different stages of infection, comprising the colonization and invasion of host


30

tissues, protection against host defences, bacterial proliferation and spread (Bien et al.,

2011). The production of pathogenic determinants during infection is mediated by

global regulators such as the accessory gene regulator (agr), the staphylococcal

accessory regulator (sarA) and the sigma factor B (σB), which activity is influenced by

different environmental signals (e.g. changes in nutrient availability, temperature, pH,

osmolarity, oxygen tension) (Bien et al., 2011; Cheung et al., 2004).

Figure 6. Major pathogenic determinants of S. aureus.

The colonization and invasion of host tissues by S. aureus is mainly mediated by the

surface-associated adhesins MSCRAMMs (microbial surface component recognizing

adhesive matrix molecules), including fibronectin-binding proteins (Fnbp), collagen-

binding proteins (e.g. CnA) and fibrinogen-binding proteins (e.g. Clf) (Bien et al., 2011;

Foster and Höök, 1998; Garzoni and Kelley, 2009). S. aureus also produces different

exotoxins and enzymes that enhance the invasion of host tissues, such as exfoliative

toxins (e.g. ETA, ETB), proteases, lipases, hyaluronidases and thermonucleases

(TNase) (Sandel and McKillip, 2004; Bukowski et al., 2010). Thus, S. aureus may

internalize into host cells and persist indefinitely, or even multiply and further

disseminate or produce a rapid cellular apoptosis or necrosis (Garzoni and Kelley,

2009).


31

In addition, S. aureus shows different evasive responses that protects bacteria from

the immune system, such as the synthesis of surface-associated staphylococcal protein

A (SpA), extracellular capsular polysaccharides, extracellular adherence proteins (Eap),

chemotaxis inhibitory proteins (CHIPS), formyl peptide receptor-like-1 inhibitory

proteins (FLIPr), extracellular complement-binding proteins (Ecb), staphylococcal

complement inhibitors (SCIN), extracellular fibrinogen-binding proteins (Efb),

staphylokinases (SAK), haemolysins, leukocidins, phenol-soluble modulins (PSMs),

catalases and coagulases (Bokarewa et al., 2006; DeLeo et al., 2009; Dinges et al., 2000;

Haas et al., 2004; Haggar et al., 2004; Jongerius et al., 2007, 2010; O´Riordan and Lee,

2004; Prat et al., 2006; Sandel and McKillip, 2004).

Most S. aureus strains also generate pyrogenic toxin superantigens (PTSAgs), which

cause the deregulation of the immune response. Apart from the TSST-1 responsible of

most toxic shock syndrome cases (Dinges et al., 2000; Lappin and Ferguson, 2009),

most S. aureus are able to produce staphylococcal enterotoxins (SEs) that cause most

food poisonings in humans (Hennekinne et al., 2012; Le-Loir et al., 2003). SEs can be

produced at a wide range of temperatures (10-46ºC, optimal production at 34-45ºC), pH

(4.0-9.6, optimum at 7-8), water activity (aw = 0.85 ≥ 0.99, optimum at aw ≥ 0.98) and

salt content (< 12%) without affecting the sensory characteristics of the contaminated

food (Hennekinne et al., 2012; Schelin et al., 2011). Moreover, the heat-stability and the

resistance to proteolytic degradation allow SEs to retain their emetic activity after food

consumption (Bergdoll and Wong, 2006; Jablonski and Bohach, 2001; Omoe et al.,

2005). Food poisoning symptoms appear approximately 1-6 h after ingestion of SEs,

depending on the amount of toxin consumed and the sensitivity of the individuals

involved (Pinchuk et al., 2010).


32

2. Biofilm formation by Staphylococcus aureus

2.1. Definition of biofilm

Although a number of definitions have been proposed over the years with the

increased understanding of biofilms (Characklis and Marshall, 1990; Costerton et al.,

1978, 1987, 1995; Costerton and Lappin-Scott, 1995; Davies and Geesey, 1995;

Marshall, 1976; Prigent-Combaret and Lejeune, 1999), the next definition is the most

widely accepted:

“A biofilm is a microbially derived sessile community characterized by cells that are

irreversibly attached to a substratum or interface or to each other, embedded in a

matrix of extracellular polymeric substances that they have produced, and with an

altered phenotype with respect to growth rate and gene transcription”

Donlan and Costerton (2002)

2.2. Significance of biofilm formation

Biofilms are the prevailing microbial lifestyle in natural habitats due to their

protector ability during the growth, allowing the survival under extreme environmental

conditions of temperature (e.g. thermal waters, glaciers), acidity (e.g. sulphuric pools

and geysers) and humidity (e.g. deserts, rainforests) (Dufour et al., 2012).

Microorganisms also grow predominantly in form of biofilms on practically any kind of

industrial surface, causing food and water contamination, metal surface corrosion and

the obstruction of equipments (Beech et al., 2005; Srey et al., 2013). Particularly in the

food industry, biofilm formation may contribute to the persistence of spoilage and

pathogenic bacteria in food-processing environments, consequently increasing cross-

contamination possibilities, which may involve a serious risk for the consumer health as

well as subsequent economic losses due to recalls of contaminated food products.

Concerning the medical ambit, the US National Institute of Health estimates that

biofilms are involved in up to 75% of microbial infections in humans (Table 1 compiled

some examples). However, most laboratory studies and many standard methods are still

performed using only planktonic cells, though this state is considered a transitory phase

in which aims to translocate bacteria to other surfaces.


33

Table 1. Diversity of human infections involving biofilms (based on Fux et al., 2005)

Infection or disease Common bacterial species involved

Dental caries Acidogenic Gram-positive cocci

Periodontitis Gram-negative anaerobic oral bacteria

Otitis media Non-typeable Haemophilus influenzae

Chronic tonsillitis Various species

Cystic fibrosis pneumonia P. aeruginosa, Burkholderia cepacia

Endocarditis Viridans group streptococci, staphylococci

Necrotizing fasciitis Group A streptococci

Musculoskeletal infections Gram-positive cocci

Osteomyelitis Various species

Biliary tract infection Enteric bacteria

Infectious kidney stones Gram-negative rods

Bacterial prostatitis E. coli and other Gram-negative bacteria

Infections related to medical devices

Contact lens P. aeruginosa, Gram-positive cocci

Sutures Staphylococci

Ventilation-associated pneumonia Gram-negative rods

Mechanical heart valves Staphylococci

Vascular grafts Gram-positive cocci

Arteriovenous shunts Staphylococci

Endovascular catheter infections Staphylococci

Cerebral spinal fluid-shunts Staphylococci

Peritoneal dialysis (CAPD) peritonitis Various species

Urinary catheter infections E. coli, Gram-negative rods

IUDs Actinomyces israelii and others

Penile prostheses Staphylococci

Orthopaedic prosthesis Staphylococci

2.3. The process of biofilm development

The development of bacterial biofilms is a dynamic process affected by the

substratum (i.e. texture or roughness, hydrophobicity, surface chemistry, charge, pre-

conditioning film), the medium (i.e. nutrient levels, ionic strength, temperature, pH,

flow rate, presence of antimicrobial agents) and intrinsic properties of cells (i.e. cell

surface hydrophobicity, extracellular appendages, extracellular polymeric substances

(EPS), signalling molecules) (Donlan, 2002; Renner and Weibel, 2011).


34

As shown in Figure 7, the process of biofilm formation comprises 1) a reversible

attachment of bacterial cells by weak interactions (i.e., Van der Waals forces) to a pre-

conditioning film formed on the abiotic or biotic surface (Bos et al., 1999; Donlan,

2002); 2) an irreversible adsorption to the surface by hydrophilic/hydrophobic

interactions, electrostatic forces and Lewis acid-base interactions mediated by several

attachment structures (e.g., flagella, fimbriae, lipopolysaccharides, adhesive proteins)

(Bos et al., 1999; Donlan, 2002); 3) the proliferation of adsorbed cells and production of

a self-produced EPS matrix mainly composed by polysaccharides, proteins and

extracellular DNA (Branda et al., 2005; Flemming et al., 2007); 4) the formation of a

mature biofilm, whose structure can be flat or mushroom-shaped and that contains water

channels that effectively distribute nutrients and signalling molecules within the biofilm

(Dufour et al., 2012; Hall-Stoodley et al., 2004); 5) the detachment of biofilm cells

individually or in clumps as a response to external or internal factors and, finally, 6) the

spread and colonization of other niches (Srey et al., 2013).

Figure 7. Processes involved in the development of a bacterial biofilm.

2.4. Ecological advantages of biofilm formation

2.4.1. Cell-cell communication

Biofilms provide microorganisms a great adaptability to the different environmental

conditions (Davey and O´Toole, 2000). The establishment of bacteria on surfaces

generates a higher degree of stability in the cell growth and enables beneficial cell-cell

interactions, such as quorum sensing and genetic exchange (Daniels et al., 2004; Davey

and O´Toole, 2000; Elias and Banin, 2012; Hall-Stoodley et al., 2004; Watnick and


35

Kolter, 2000). In the case of quorum sensing, low-molecular mass signalling molecules

called auto-inducers (AI) regulate gene expression, metabolic cooperativity and

competition, physical contact and bacteriocin production, providing thus a mechanism

for self-organization and regulation of biofilm cells (Daniels et al., 2004; Donlan, 2002;

Elias and Banin, 2012; Parsek and Greenberg, 2005; Van-Houdt and Michiels, 2010).

The quorum sensing effects depend on the concentration of AI, which increases in a

cell-density-dependent manner (Parsek and Greenberg, 2005). Meanwhile, the

transmission of mobile genetic elements between nearby biofilm cells allows the

acquisition of new antimicrobial resistance, virulence factors and environmental

survival capabilities (Madsen et al., 2012).

2.4.2. Increased resistance to external stimulus

Bacterial cells in biofilms are protected against a wide range of environmental

stresses, leading to a higher survival and persistence when compared with planktonic

state, which is particularly problematic in clinical and food environments. Several

physiological characteristics and mechanisms are responsible of this increase of

resistance:

Presence of extracellular matrix. It acts as a barrier that slows down the infiltration,

neutralizes, binds and effectively diffuses to sub-lethal concentrations antimicrobial

and antibiotic agents (e.g. chlorine species, oxacillin, vancomycin) before they can

reach cell targets (Bridier et al., 2011a; Singh et al., 2010), and also reduce other

adverse external effects such as UV light, toxic metals, acidity, desiccation, salinity

and host defences (Hall-Stoodley et al., 2004).

Specific physiology of biofilm cells. Changes in the expression of specific genes

associated with sessile growth and physiological differentiation between biofilm cells

actively growing located in the outer surface and dormant or even in the oxygen- and

nutrient-deprived inner interfaces (Bridier et al., 2011a) represents the basis of the

biofilm-specific adaptive response and it can explain their reduced susceptibility to

antimicrobials (Bridier et al., 2011a; Gilbert et al., 2002; Høiby et al., 2010). As an

example, persistent cells are able to produce particular cellular toxins that block

cellular processes like translation, thus rendering protection against biocides that act

only against active cells (Lewis, 2010).


36

Expression of specific mechanisms of defence. In between others, biofilm cells can

respond to antimicrobials by the activation of chromosomal β-lactamases and efflux

pumps (e.g. QAC efflux system of S. aureus), the induction of mutations in

antimicrobial target molecules or, indirectly, by the release of extracellular DNA that

promotes the synthesis of biofilm matrix (Anderson and O’Toole, 2008; Bridier et

al., 2011a; Høiby et al., 2010; Kaplan et al., 2012).

2.4.3. Increased dispersal capacity

The dispersal of biofilm cells in clumps may provide a sufficient number of cells for

an infective dose that is not typically found in bulk fluid, enabling an enhanced

transmission and infection of bacterial pathogens (Hall-Stoodley and Stoodley, 2005).

Detachment can be promoted by environmental changes such as nutrient starvation or

by internal biofilm processes such as endogenous enzymatic degradation, or the release

of EPS or surface-binding proteins (Srey et al., 2013).

2.5. Staphylococcus aureus biofilms: development, composition and

regulation

S. aureus biofilms have been detected on diverse biotic and abiotic surfaces,

including human tissues, indwelling medical devices (e.g., implanted catheters, artificial

heart valves, bone and joint prostheses), food products and food-processing facilities

(Devita et al., 2007; Herrera et al., 2006; Sattar et al., 2001; Simon and Sanjeev, 2007;

Trampuz and Widmer, 2006).

The initial attachment of S. aureus to abiotic surfaces is mostly mediated by

hydrophobic or electrostatic interactions, but bacterial surface molecules such as

autolysins or teichoic acids can be also participate in this process (Gross et al., 2001;

Houston et al., 2011). In contrast, S. aureus adhere to biotic surfaces through much

more specific interactions governed by the surface-anchored proteins MSCRAMMs

(Foster and Höök, 1998).


37

Once adhered, S. aureus develop a multi-layered biofilm embedded within a matrix

composed by secreted polysaccharides (e.g. PIA-PNAG), lipids and proteins,

extracellular DNA originating from lysed cells, and some molecules trapped from the

environment (Branda et al., 2005; Cramton et al., 1999; Fitzpatrick et al., 2005;

Flemming et al., 2007; Høiby et al., 2010; Maira-Litrán et al., 2002), though the

composition can vary between strains and at different environmental conditions.

The extracellular polysaccharide intercellular adhesins (PIA) participates in the

quorum-sensing contacts that coordinate the biofilm formation process, as well as in the

cell detachment induced by changes in environmental conditions (Fitzpatrick et al.,

2005). PIA is formed by poly-β(1,6)-N-acetyl-d-glucosamine glycans (PNAG), which

are synthetized by the products of the chromosomal intercellular adhesion (ica) operon

carried by most S. aureus strains (Cramton et al., 1999; Fitzpatrick et al., 2005; Maira-

Litrán et al., 2002). IcaA and IcaD form a transmembrane protein N-acetyl-glucosamine

transferase that produce N-acetyl-glucosamine oligomers, which are elongated and

translocated to the cell surface by the membrane protein IcaC (Gerke et al., 1998),

where the surface-attached protein IcaB finally de-acetylates the polymers, introducing

positive charges that enhance the adhesion of PIA to the bacterial surface (Vuong et al.,

2004). The ica operon is fundamentally repressed by the icaR gene products (Jefferson

et al., 2003), which are regulated by the stress-induced sigma factor B (σB), the

staphylococcus accessory regulator A (SarA), the LuxS/AI-2 quorum-sensing system

and, indirectly, by the rbf gene (Cerca et al., 2008; Cue et al., 2009; Yu et al., 2012).

The role of the ica operon in the biofilm formation of S. aureus is complex and likely to

be both strain- and environment-dependent (O’Gara, 2007). In fact, the expression of

the ica operon is influenced by different environmental factors such as anaerobic

conditions, glucose, ethanol, osmolarity, temperature and antibiotics such as tetracycline

(Cramton et al., 2001; Fitzpatrick et al., 2005).

The extracellular DNA, meanwhile, has been found to play a structure-stabilizing

role in the extracellular matrix of S. aureus biofilms. Thus, the negative charge of

extracellular DNA seems to aid in interacting with other surface structures and lead to

the formation of an adherent staphylococcal biofilm (Izano et al., 2008). The

extracellular DNA is released from lysed adhered cells since the early stages of biofilm


38

formation under the control of cid/lgr operons (Mann et al., 2009; Rice et al., 2007).

Furthermore, balance of the extracellular DNA content by the degrading activity of

secreted thermonucleases (e.g. nuc1, nuc2) seems to be important in the maintenance of

mature biofilms (Mann et al., 2009).

The presence of extracellular carbohydrate-binding proteins and surface adhesins

also contribute to the formation and stabilization of the polysaccharide matrix as well as

to enhance the intercellular adhesion. Several biofilm adhesive proteins are implicated

in biofilm accumulation by S. aureus, including the multifactorial virulence factor SpA

(Merino et al., 2009), FnbpA and FnbpB (O’Neill et al., 2008), the biofilm-associated

protein (Bap) expressed by bovine strains of S. aureus (Lasa and Penadés, 2006) and the

surface proteins SasG (Corrigan et al., 2007; Geoghegan et al., 2010) and SasC

(Schroeder et al., 2009). The expansion and dispersal of S. aureus biofilms is also

controlled by secreted proteins, particularly the phenol-soluble modulins (PSMs), which

originate the characteristic channels of mature biofilms (Periasamy et al., 2012). The

proteases aureolysins seem to be also implicated in these processes, though there is no

clear evidence currently (Otto, 2013). The production of both effectors is strictly

regulated by the quorum-sensing system Agr (accessory gene regulator), which is

stimulated by auto-inducing peptides (Wang et al., 2007).

3. Incidence of Staphylococcus aureus in the food

industry

3.1. Outbreaks

A selection of the most notable foodborne outbreaks worldwide caused by the

ingestion of S. aureus enterotoxins is shown in Table 2.


39

Tab

le 2

. E

xce

rpt

of

rep

ort

ed f

oo

dborn

e in

toxic

atio

ns

cause

d b

y S

. aure

us

ente

roto

xin

s

Ref

eren

ce

Joh

nso

n e

t al

. (1

990

)

CD

C (

19

68

)

Mo

rris

et

al. (1

972

)

Eis

enber

g e

t al

. (1

975

)

CD

C (

19

76

)

To

dd

et

al. (1

981

)

CD

C (

19

83

)

Wat

erm

an e

t al

. (1

98

7)

De-

Bu

yse

r et

al.

(1

98

5)

Bo

ne

et a

l. (

19

89

)

Wo

ola

way

et

al.

(19

86)

Ev

enso

n e

t al

. (1

98

8)

Kér

ou

anto

n e

t al

. (2

00

7)

Lev

ine

et a

l. (

19

96

)

Th

aikru

ea e

t al

. (1

995

)

Ric

har

ds

et a

l. (

19

93)

Kér

ou

anto

n e

t al

. (2

00

7)

FD

A (

19

92

)

Kér

ou

anto

n e

t al

. (2

00

7)

Kér

ou

anto

n e

t al

. (2

00

7)

SE

in

vo

lved

Un

spec

ifie

d

Un

spec

ifie

d

SE

E

Un

spec

ifie

d

SE

D

SE

A,

SE

C

SE

A

Un

spec

ifie

d

SE

A,

SE

D

SE

A

SE

A

SE

A

SE

A

SE

A

SE

A,

SE

C

SE

A

SE

A

Un

spec

ifie

d

Un

spec

ifie

d

Un

spec

ifie

d

Ca

ses

20

0

13

00

10

0

19

7

80

62

12

1

21

5

20

27

50

86

0

70

10

2

48

5

10

0

32

13

64

87

47

Incr

imin

ate

d f

ood

Raw

mil

k c

hee

se

Chic

ken

sal

ad

Sau

sages

roll

s, h

am s

andw

iches

Ham

Choco

late

écl

airs

Chee

se c

urd

Ham

/chee

se s

andw

ich,

stuff

ed c

hic

ken

Des

sert

cre

am p

astr

y

Far

m e

ve

chee

se

Shee

p’s

mil

k c

hee

se

Dri

ed l

asag

ne

Choco

late

mil

k

Spag

het

tis

Can

ned

mush

room

s

Écl

airs

Ham

, bea

ns

and c

orn

Pota

to a

nd r

ice

sala

ds

Chic

ken

sal

ad

Raw

mil

k c

hee

se

Raw

mil

k c

hee

se

Lo

cali

zati

on

US

A

US

A

UK

Fli

ght

Jap

an-D

enm

ark

Fli

ght

Bra

zil-

US

A

Can

ada

US

A

Car

ibbea

n c

ruis

e sh

ip

Fra

nce

Sco

tlan

d

Fra

nce

, U

K, It

aly

, L

ux

embourg

US

A

Fra

nce

US

A

Th

aila

nd

US

A

Fra

nce

US

A

Fra

nce

Fra

nce

Yea

r

1958

1968

1971

1975

1976

1980

1982

1983

1983

1984

1985

1985

1988

1989

1990

1990

1990

1992

1997

1997


40

Tab

le 2

(co

nti

nu

ati

on

). E

xce

rpt

of

report

ed f

oodborn

e in

toxic

atio

ns

cause

d b

y S

. aure

us

ente

roto

xin

s

Ref

eren

ce

Car

mo e

t al

. (2

002

)

Co

lom

bar

i et

al.

(2

00

7)

Car

mo e

t al

. (2

002

)

Kér

ou

anto

n e

t al

. (2

00

7)

Asa

o e

t al

. (2

00

3)

Iked

a et

al.

(2

005

)

Kér

ou

anto

n e

t al

. (2

00

7)

Kér

ou

anto

n e

t al

. (2

00

7)

Kér

ou

anto

n e

t al

. (2

00

7)

Kér

ou

anto

n e

t al

. (2

00

7)

Nem

a et

al.

(2

00

7)

Hen

nek

inn

e et

al.

(2

00

9)

Fit

z-Ja

mes

et

al.

(20

08

)

Sch

mid

et

al.

(20

09

)

Kit

amoto

et

al.

(20

09

)

Ost

yn

et

al.

(20

10)

So

lan

o e

t al

. (2

01

3)

SE

in

vo

lved

SE

A,

SE

B,

SE

D

SE

A

SE

A,

SE

B,

SE

C

SE

C

SE

A,

SE

H

SE

A

SE

A

SE

D

SE

A

SE

B,

SE

D

SE

A,

SE

D

SE

A,

SE

D

SE

A,

SE

D

SE

A,

SE

B,

SE

C

SE

E

SE

A,

SE

D

Case

s

4000

180

50

160

13420

21

85

17

45

> 1

00

17

15

166

75

23

42

Incr

imin

ate

d f

ood

Ch

icken

pan

cake,

ri

ce,

bea

ns,

to

mat

o

sauce

an

d

mas

hed

chic

k-p

eas

Veg

etab

le sa

lad w

ith m

ayonnai

se,

bro

iled

ch

icken

,

pas

ta i

n t

om

ato s

auce

Raw

mil

k w

hit

e ch

eese

Mix

ed s

alad

Lo

w-f

at m

ilk p

ow

der

Pan

cakes

Cre

am

Raw

mil

k s

emi-

har

d c

hee

se

Raw

shee

p´s

mil

k c

hee

se

Fri

ed p

ota

to b

alls

Co

co n

ut

pea

rls

(Chin

ese

des

sert

)

Ham

burg

er

Mil

k,

caca

o m

ilk, van

illa

mil

k

Cre

pes

Raw

mil

k c

hee

se

Mac

aroni

Lo

cali

zati

on

Bra

zil

Bra

zil

Bra

zil

Fra

nce

Jap

an

Fra

nce

Fra

nce

Fra

nce

Fra

nce

Ind

ia

Fra

nce

Bel

giu

m

Au

stri

a

Jap

an

Fra

nce

Sp

ain

Yea

r

19

98

19

98

19

99

20

00

20

00

20

01

20

01

20

01

20

02

20

05

20

06

20

07

20

07

20

09

20

09

20

11


41

The European Food Safety Authority (EFSA) informed that staphylococcal toxins

comprise 4.6% of food poisoning outbreaks reported since 2005 (Table 3), being the

fourth main causative agent after Salmonella spp. (42.3%), foodborne viruses (12.5%)

and Campylobacter spp. (8.1%) (EFSA, 2006, 2007, 2009a, 2010, 2011, 2012).

Moreover, the hospitalization rate average due to staphylococcal intoxications (14.1%)

was higher than that of foodborne viruses (5.5%) and Campylobacter spp. (7.1%),

causing also a higher proportion of deaths (7% out of total deaths against 2.2% for

foodborne viruses and 1.1% for Campylobacter spp.). Particularly in Spain,

staphylococcal toxins had higher impact in foodborne outbreaks (5.9%) than in the EU

between 2008 and 2010 (previous data were not included in EFSA reports), but a lower

hospitalization rate average (3.4%) and none death was reported.

Similar results were reported in Japan, where staphylococcal intoxications involve

4.3% of foodborne outbreaks with identified causative agent reported in 2009,

producing 690 cases but none deaths (MHLW, 2011). S. aureus enterotoxins were

associated with approximately 7.8% of foodborne outbreaks reported in China between

1994 and 2005, causing 3055 cases but none deaths (Wang et al., 2007). In contrast, the

Food and Drug Administration (FDA) estimated that staphylococcal toxins are

responsible for causing only 0.5% of food poisoning outbreaks reported in the USA,

with a hospitalization rate of 0.4% and 6 deaths each year (FDA, 2012).

However, it is known that the number of reported cases is underestimated by the

healthcare services due to misdiagnosis of the illness (symptomatically similar than

other types of food poisoning, e.g., Bacillus cereus emetic toxin), the self-limiting

nature of the illness and the rapid recovery of people intoxicated by staphylococcal

toxins (within 24 to 48 h after onset). In fact, only 40% out of the foodborne outbreaks

caused by staphylococcal toxins in the EU were verified. Furthermore, the notification

of staphylococcal intoxications is not mandatory in a number of member states of the

EU. Therefore, the actual incidence of staphylococcal food poisonings is assumed to be

much higher than reported (Lawrynowicz-Paciorek et al., 2007; Smyth et al., 2004).


42

Tab

le 3

. F

oo

db

orn

e o

utb

reak

s re

port

ed i

n t

he

Euro

pea

n U

nio

n b

etw

een 2

005 a

nd 2

010

, in

clu

din

g t

hose

cau

sed

by

sta

ph

ylo

cocc

al t

oxin

s (E

FS

A,

200

6, 2

00

7, 2

00

9a,

20

10

, 20

11

, 2

012).

Ou

tbre

ak

s ca

use

d b

y s

tap

hy

lococc

al

tox

ins

Dea

ths

1

no

2

no

4

no

2

0

3

0

0

0

12

0

no

, d

ata

not

incl

ud

ed i

n E

FS

A r

eport

s

Ho

spit

ali

sed

36

5

no

37

8

no

27

2

no

36

2

10

30

3

13

35

7

0

20

37

23

Ca

ses

16

92

no

23

73

no

27

28

no

27

49

13

2

26

71

47

7

27

96

16

6

15

00

9

77

5

% v

erif

ied

no

no

no

no

90

.70

no

27

.84

46

.88

30

.03

36

.67

13

.87

37

.50

40

.61

40

.35

N

16

4

no

24

0

no

25

8

no

29

1

32

29

3

30

27

4

24

15

20

86

Tota

l ou

tbre

ak

s

Dea

ths

24

7

50

1

19

3

32

1

46

2

25

7

196

21

Hosp

itali

sed

5330

23

5525

no

3291

362

6230

181

4356

297

4695

378

29427

1241

Case

s

47251

7682

53568

3491

39727

6705

45622

2372

48964

5584

43473

4025

278605

29859

% v

erif

ied

92.0

0

100.0

0

83.7

0

100.0

0

31.1

2

41.0

3

16.6

9

38.8

4

17.6

0

33.8

9

13.2

6

40.6

6

42.4

0

59.0

7

N

5311

460

5710

351

5733

619

5332

551

5550

416

5262

482

32898

2879

Geo

gra

ph

ic a

rea

EU

Sp

ain

EU

Sp

ain

EU

Sp

ain

EU

Sp

ain

EU

Sp

ain

EU

Sp

ain

EU

Sp

ain

Yea

r

20

05

20

06

20

07

20

08

20

09

20

10

To

tal


43

Diseases caused by food intoxications additionally generate considerable economic

costs and a wide impact in public health worldwide. For example, the Centres for

Disease Control and Prevention (CDC) estimate that foodborne diseases generate costs

of over 1.4 trillion of dollars in the USA each year (Roberts, 2007). Moreover, the recall

of contaminated products, factory closing, extraordinary cleanings and compensation to

infected people can generate additional expenses to the food industry.

3.2. Foods implicated in staphylococcal poisonings

Different food vehicles have been incriminated in staphylococcal intoxications. As

shown in Figure 8, the largest proportion of verified outbreaks with known food vehicle

caused by staphylococcal toxins in the EU since 2006 (no data included in the EFSA

report of 2005) was attributed to meat products (32.9%), milk and dairy products

(23.5%) and mixed or buffet meals (10.9%), followed by fishery products (3.8%).

Figure 8. Identified food vehicles incriminated in verified foodborne outbreaks (n = 340)

caused by staphylococcal toxins in the European Union between 2006 and 2010 (data from

EFSA, 2007, 2009a, 2010, 2011, 2012).

However, foods incriminated in staphylococcal intoxications varying widely among

countries due to differences in the food consumption and habits (Bhatia and Zahoor,

2007; Le-Loir et al., 2003). For instance, in France are frequently reported

staphylococcal poisonings associated with the consumption of milk and dairy products

(Cretenet et al., 2011), whereas meat products are mostly involved in outbreaks reported


44

in Anglo-Saxon countries (Gormley et al., 2011), and cereals (above all rice) and fishery

products in Asiatic countries such as Japan (MHLW, 2011). In Spain, staphylococcal

intoxications possibly have a higher incidence in fishery products than the European

mean, as the second largest consumer in the European Union (FAO, 2012). However,

from our knowledge, only two studies have evaluated the presence of S. aureus in

fishery products in the last decade, particularly in vacuum-packed cold‐smoked salmon

(González-Rodríguez et al., 2002) and fresh marine fish (Herrera et al., 2006).

3.3. Food contamination by Staphylococcus aureus

Human handlers are considered the major source of transference of S. aureus to food

and food-contact surfaces during the preparation and processing of food products

(Devita et al., 2007; Sattar et al., 2001; Simon and Sanjeev, 2007), as S. aureus form

part of the normal human microbiome (see Figure 4). Nevertheless, foods of animal

origin (e.g. milk from dairy animals with mastitis) (Kérouanton et al., 2007; Morandi et

al., 2010) as well as endemic strains present in the food-processing environment (Le-

Loir et al., 2003) can be also a potential source of primary contamination.

S. aureus is a poor competitor in complex microbial populations, being often

inhibited or displaced by food-spoilage microorganisms of faster growing such as

Acinetobacter, Aeromonas, Bacillus, Pseudomonas, the Enterobacteriaceae or the

Lactobacillaceae, among others. Thus, the greatest risk of staphylococcal food

poisoning is often associated with food contaminated with S. aureus after the normal

microflora has been destroyed (e.g. cooked products) or inhibited (e.g. frozen/salted

foods) (Bore et al., 2007).

Food-related environmental conditions such as the physicochemical characteristics of

food and food-contact surfaces, as well as inadequate temperatures and hygiene

practices during processing, storage or distribution of products can favour the growth of

S. aureus and stimulate the production of toxins (Hennekinne et al., 2012). Finally,

novel trends in food production such as minimal processing, mass production and

globalization, among others, have additionally introduced new factors and conditions

that can enhance the staphylococcal proliferation on food environments (Abee and

Wouters, 1999; Cebrián et al., 2007; Rendueles et al., 2011).


45

4. Control of Staphylococcus aureus in the food industry

4.1. Hazard Analysis and Critical Control Points (HACCP)

HACCP is an effective management system that assures food safety from growing to

consumption, by anticipating and preventing health hazards before they occur.

Prerequisite programs provide the basic environmental and operating conditions needed

for the production of safe, wholesome food. It combines the Good Manufacturing

Practice (GMPs), Good Hygienic Practices (GHPs) and Good Agricultural Practices

(GAPs) to determine the strategy ensuring hazard control. Some of these basic

conditions are the good quality of raw material, the effective training of employees in

food hygiene, and the appropriate design of the factory and equipment to assure a

correct cleaning and disinfection of the surfaces. All prerequisite programs should be

documented and regularly audited, and are established and managed separately from the

HACCP plan.

Once prerequisites are controlled, critical points in the process where contamination

can occur should be identified and controlled. In the case of S. aureus, the lack of

hygiene in food handlers and food-contact surfaces have been found to be the major

factors involved in the contamination of food products (DeVita et al., 2007; Sattar et al.,

2001; Simon and Sanjeev, 2007), so their control is essential to provide an appropriate

food safety conditions.

4.2. Prevention strategies against biofilm formation

Several approaches were proposed to prevent or limit bacterial attachment in food

contact surfaces, including the incorporation of antimicrobial products in the surface

materials themselves (Weng et al., 1999; Park et al., 2004; Knetsch and Koole, 2011),

the coating of surfaces with antimicrobials (Gottenbos et al., 2002; Knetsch and Koole,

2011; Thouvenin et al., 2003), the modification of the physicochemical properties of

surfaces (Chandra et al., 2005; Mauermann et al., 2009; Rosmaninho et al., 2007) or the

pre-conditioning of surfaces with surfactants (Chen, 2003; Choi et al., 2011), among

others.

In another line of research, different biofilm detectors were developed to monitor the

bacterial colonization of surfaces, allowing thus the control of biofilms in the early


46

stages of development when cells are more susceptible to antimicrobials (Pereira et al.,

2008; Philip-Chandy et al., 2000). However, even though the efforts to solve the

problem, there is currently no known technique that is able to successfully prevent or

control the formation of unwanted biofilms without causing adverse side effects

(Simões et al., 2010).

4.3. Cleaning procedures

Cleaning procedures aim to remove any food residues and other compounds that may

promote bacterial proliferation and biofilm formation (Simões et al., 2010). Cleaning

agents are frequently combined with disinfectants to synergistically enhance

disinfection efficiency due to the low efficacy of most disinfectants in the presence of

organic materials (Forsythe and Hayes, 1998; Simões et al., 2010).

Mechanical cleaning procedures (e.g., water turbulence, brushing and scrubbing) and

high temperature treatments have been traditionally used as efficient cleaning methods

in the food industry (Chmielewski and Frank, 2006; Gibson et al., 1999; Maukonen et

al., 2003), but the former usually are an arduous task while the latter may favour the

growth of opportunistic pathogens such as S. aureus.

Food residues can also be removed through the application of surfactants and

alkaline products, which decrease surface tension, emulsify fats and denature proteins

(Forsythe and Hayes, 1998; Maukonen et al., 2003), or acid cleaners, in the case of food

residues with high mineral content (Marriott and Gravani, 2006). However, the use of

these cleaning agents usually produce corrosiveness to metal surfaces, harmful effects in

workers and a high environmental impact (Marriott and Gravani, 2006).

Enzyme-based detergents and quorum-sensing inhibitors have been proposed as a

promising alternative (Høiby et al., 2010; Thallinger et al., 2013), but the complexity to

achieve a wide-spectrum enzymatic formulation and the high production costs have

reduced their potential application in the food industry (Lequette et al., 2010; Simões et

al., 2010).

Dry-ice cleaning was recently proposed as an effective procedure to remove

impurities adhered to surfaces by local undercooling (Otto et al., 2011), although it must


47

be still fully automated, and sound levels and physical stresses generated by

compression units should be greatly reduced.

4.4. Disinfection treatments

Disinfection aims to kill the remaining surface population left after cleaning and thus

prevent the microbial regrowth before production restart (Simões et al., 2010).

Physical treatments such as the application of heated solutions (e.g. hot water) and

ultraviolet (UV) radiation have been frequently used in the food industry (Marriott and

Gravani, 2006), but the limited effectiveness against spores and biofilms as well as the

possibility to form films or scale on equipment has questioned their validity. Pulsed UV

light (Gómez-López et al., 2007), ultrasonication (Chemat et al., 2011; Oulahal-Lagsir

et al., 2000), ionizing radiations (Byun et al., 2007; Niemira and Solomon, 2005;

Niemira, 2008) and low temperature atmospheric pressure plasmas (Ehlbeck et al.,

2011) have shown promising results for the eradication of bacteria attached to surfaces,

but they require high initial investments and have low consumer acceptance.

Many chemical disinfectants have been also used in the food industry. They

generally interact with multiple cellular targets (Figure 9), leading to lethal bactericidal

effects. Thus, the rotation and combination of biocides is widely accepted to prevent the

emergence of antimicrobial-resistant strains.

The choice of a chemical disinfectant depends on efficacy, safety, toxicity, corrosive

effects and ease of removal, among other factors (Marriott and Gravani, 2006).

However, the efficacy of chemical disinfectants are often affected by the presence of

organic materials, pH, temperature, water hardness, chemical inhibitors, concentration,

exposure time and bacterial resistance (Bessems, 1998; Marriott and Gravani, 2006).

Advantages and disadvantages of some disinfectants widely used in the food industry

are summarized in Table 4.


48

Tab

le 4

. A

dvan

tages

and d

isad

van

tages

of

som

e dis

infe

ctan

ts w

idel

y u

sed i

n t

he

food i

ndust

ry (

Wir

tanen

and S

alo,

2003;

Mar

riott

and

Gra

van

i, 2

006).

Dis

ad

van

tages

Bio

stat

ics.

Inef

fect

ive

agai

nst

spore

s.

Toxic

, ir

rita

ting, unst

able

, pote

nti

ally

explo

sive

and

corr

osi

ve.

In

acti

vat

ed in

th

e pre

sence

of

org

anic

mat

eria

l. p

H s

ensi

tive.

D

isco

lora

tion o

f pro

du

cts.

Res

ista

nce

dev

elopm

ent.

Bio

stat

ic.

Not

bio

deg

radab

le.

Low

pen

etra

tion in

bio

film

s.

Fla

vour

and

odour.

S

tain

pla

stic

s an

d

poro

us

mat

eria

ls.

Hig

hly

foam

ing,

unsu

itab

le f

or

clea

nin

g-

in-p

lace

(C

IP).

Red

uce

d e

ffic

acy a

t hig

h p

H a

nd a

t

tem

per

ature

s ab

ove

50°C

. E

xpen

sive.

Loss

of

effe

ctiv

enes

s in

th

e pre

sence

of

org

anic

mat

eria

l an

d s

om

e m

etal

s co

nta

ined

in w

ater

. M

ay

corr

ode

som

e m

etal

s. L

ow

eff

icac

y a

gai

nst

yea

sts

and m

ould

s. R

elat

ivel

y e

xpen

sive.

Lim

ited

ef

fect

iven

ess,

w

hic

h is

af

fect

ed by h

ard

wat

er,

low

tem

per

ature

s an

d l

ow

pH

. In

com

pat

ible

wit

h m

ost

det

ergen

ts.

Hig

hly

foam

ing,

unsu

itab

le

for

CIP

. R

esid

ual

an

tim

icro

bia

l fi

lm

form

ing.

Res

ista

nce

dev

elopm

ent.

Rel

ativ

ely e

xpen

sive.

Ad

van

tages

Chea

p,

fast

-act

ing

bio

cides

of

bro

ad

mic

robia

l

spec

trum

, non

-toxic

, ea

sy-t

o-u

se,

colo

url

ess,

har

mle

ss o

n s

kin

, so

luble

in w

ater

and v

ola

tile

.

Chea

p,

fast

-act

ing

oxid

izer

s of

bro

ad

mic

robia

l

spec

trum

. E

asy

-to

-use

an

d

unaf

fect

ed

by

har

d

wat

er.

Eff

ecti

ve

agai

nst

pla

nkto

nic

cel

ls a

nd s

pore

s,

even

at

lo

w

tem

per

ature

s.

Support

s m

icro

bia

l

det

achm

ent.

N

on

-fil

m

form

ing

wit

hout

resi

du

al

acti

vit

y.

Chea

p b

ioci

de

of

bro

ad m

icro

bia

l sp

ectr

um

, no

n-

corr

osi

ve.

San

itiz

ers

of

bro

ad

acti

vit

y

spec

trum

, non

-

corr

osi

ve,

non

-irr

itat

ing

an

d

easy

-to

-use

. L

ow

toxic

ity a

nd s

table

at

a v

ery l

ow

pH

. L

ittl

e af

fect

ed

by o

rgan

ic m

ater

ials

.

Str

ong,

fast

-act

ing

oxid

izer

s of

bro

ad

mic

robia

l

spec

trum

, re

lati

vel

y

non

-toxic

an

d

easy

-to

-use

.

Low

fo

amin

g,

suit

able

fo

r C

IP.

Eff

ecti

ve

agai

nst

bac

teri

al

bio

film

s an

d

spore

s,

even

at

lo

w

tem

per

ature

s. N

on-c

orr

osi

ve

to s

tain

less

ste

el a

nd

alum

iniu

m.

Sta

ble

, su

rfac

e-ac

tive

agen

ts,

non

-toxic

, n

on

-

irri

tati

ng,

non

-corr

osi

ve,

odourl

ess,

fla

vourl

ess

and

colo

url

ess.

L

ittl

e af

fect

ed

by

org

anic

m

ater

ials

.

Support

s m

icro

bia

l det

achm

ent.

Dis

infe

ctan

t

Alc

ohols

(e.g

., e

than

ol)

Chlo

rine-

bas

ed c

om

poun

ds

(e.g

., s

odiu

m h

ypoch

lori

te)

Glu

tara

ldeh

yde

Iodophors

Per

oxygen

s

(e.g

., p

erac

etic

aci

d,

hydro

gen

per

oxid

e)

Quat

ernar

y a

mm

oniu

m

com

pounds

(QA

Cs)

(e.g

., b

enza

lkoniu

m c

hlo

ride)


49

Figure 9. Cellular targets and effects of disinfectants commonly used in the food industry

(based on Denyer and Stewart (1998) and Maillard (2002)).

Interestingly, numerous working-safe, environmentally-friendly and cost-effective

disinfection options have been introduced ‒or at least proposed to be introduced‒ in the

food industry to be in concordance with present and future regulatory landscapes,

highlighting:

Ozone: it is considered a suitable and safety sanitizer for decontaminating food

products, food-contact surfaces, equipments and the food-processing environment

(Graham et al., 1997; Khadre et al., 2001). Likewise EW, ozone is generated on-site

and it poses low environmental impact because it decomposes rapidly without

leaving toxic residues, but it is more expensive, unstable and corrosive than EW

(O’Donnell et al., 2012). Ozone acts as a powerful and non-selective oxidant and

disinfectant against a wide range of microorganisms including food-related bacteria

(Restaino et al., 1995). Ozone kills cells by oxidizing polyunsaturated fatty acids and

the sulfhydryl groups of certain enzymes, which leads to the disruption or

disintegration of the cell envelope and subsequent leakage of cellular contents

(Victorin, 1992); or by the destruction and damage of nucleic acids (Guzel-Seydim et


50

al., 2004). As a consequence, bacteria seems to cannot develop resistance to ozone

disinfection (Pascual et al., 2007). However, the stability of ozone is affected by

temperature and pH conditions and its efficacy is decreased by the presence of

organic matter and depends on the type of microorganism targeted (Khadre et al.,

2001; O’Donnell et al., 2012).

Electrolyzed water (EW): this disinfectant is considered safety, environmentally-

friendly and non-corrosive to stainless steel surfaces (Huang et al., 2008). Moreover,

EW is more cost effective than traditional disinfectants because, once the initial

capital investment is made to purchase an EW generator unit, the only operating

expenses are water, salts and electricity to run the unit (Walker et al., 2005). The

bactericidal activity of EW derives from the combined action of pH, oxidation-

reduction potential (ORP) and available chlorine concentration (ACC). Thus, EW

damages the bacterial protective barriers, increases membrane permeability leading

to the leakage of intracellular DNA, K+ and proteins, and causes an activity decrease

on critical enzymatic pathways (Liao et al., 2007; Marriott and Gravani, 2006; Zeng

et al., 2010). The efficacy of EW against foodborne pathogens has been widely

demonstrated in suspension cultures and foods, including S. aureus (Deza et al.,

2005; Fenner et al., 2006; Guentzel et al., 2008; Horiba et al., 1999; Issa-Zacharia et

al., 2010; Rahman et al., 2010; Vorobjeva et al., 2004). Nonetheless, a reduced

number of studies were performed against bacterial biofilms (Ayebah and Hung,

2005; Huang et al., 2008; Liu et al., 2006; Monnin et al., 2012; Park et al., 2002;

Phuvasate and Su, 2010; Venkitanarayanan et al., 1999; Walker et al., 2005), which

may question the antimicrobial potential of EW in contaminated food-processing

facilities.

Bacteriophages: the application of viruses infecting bacteria along the food chain

(phage therapy, biosanitation, biopreservation) is considered as a versatile biofilm

control tool, highly active and specific (without adverse effects on the intestinal

microbiota and innocuous to mammalian cells), relatively low cost, and genetically

amenable (García et al., 2008; Loc-Carrillo and Abedon, 2011). Bacteriophages are

often engineered to express specific proteins such as endolysins and other hydrolases

that directly degrade or interfere in the biosynthesis of bacterial peptidoglycans

(Sutherland et al., 2004; Lu and Collins, 2007; García et al., 2010). Several studies


51

reported the capability of bacteriophages to remove S. aureus biofilms (García et al.,

2007; Obeso et al., 2008; Sass and Bierbaum, 2007). However, the limited spectrum

of infectivity of each bacteriophage necessitates the identification of causative

bacterial pathogens and their susceptibility to phages (Lu and Koeris, 2011). In fact,

some bacteria can evolve resistance to phages by blocking phage adsorption,

inhibiting the injection of phage genomes, restriction-modification systems and

abortive infection systems (Labrie et al., 2010). Moreover, bacteriophages should be

genetically well-characterized to avoid the dissemination of undesirable traits

(virulence and antibiotic-resistance genes) that could involve safety concerns (Loc-

Carrillo and Abedon, 2011). In food systems, different factors such as the

concentration and diffusion rate of phages, the physiological status and proportion of

cells immersed into the biofilm, as well as environmental conditions (e.g.,

temperature, pH, presence of inhibitory compounds) can additionally affect the

infection of biofilm cells by phages (García et al., 2008).

Essential oils (EOs): they comprise a wide variety of aromatic oily liquids

(approximately 3000 EOs are known nowadays) extracted from different plant

materials such as flowers, fruits, herbs, leaves, roots and seeds (Bakkali et al., 2008;

Burt, 2004). EOs show a versatile composition, which may vary depending on

geographical source, climate, harvesting season, soil composition, plant organ, age

and vegetative cycle stage, as well as the extraction method used (Angioni et al.,

2006; Ennajar et al., 2010; Guan et al., 2007; Hussain et al., 2010; Paolini et al.,

2010). EOs acts as antioxidants (Brenes and Roura, 2010) and possess broad-range

antibacterial (Oussalah et al., 2007), antiparasitic (George et al., 2009), insecticidal

(Nerio et al., 2010), antiviral (Astani et al., 2011), antifungal (Tserennadmid et al.,

2011) properties. In fact, several studies have used EOs against S. aureus biofilms

(Table 5). EOs cause changes in cell morphology, in the physicochemical properties

of membranes, as well as in the transcriptome, proteome and toxin production;

disruptions in the membrane potential, intracellular pH and Ca+2

homeostasis, and

cellular respiration; alterations in the thiol groups; and the inhibition of essential

enzymatic pathways and cell division (Hyldgaard et al., 2012). Nonetheless, the

practical application of EOs has been limited due to their strong flavour, poor

solubility and partial volatility (Delaquis et al., 2002; Kalemba and Kunicka, 2003).


52

Tab

le 5

. E

xam

ple

s o

f es

senti

al o

ils

appli

ed a

gai

nst

S.

aure

us

bio

film

s.

Ref

eren

ce

Qu

ave

et a

l. (

20

08

)

Kav

anau

gh

an

d R

ibb

eck

(20

12

)

Mil

lezi

et

al. (2

01

2)

Kav

anau

gh

an

d R

ibb

eck

(20

12

)

Qu

ave

et a

l. (

20

08

)

Qu

ave

et a

l. (

20

08

)

Sch

illa

ci e

t al

. (2

00

8)

Qu

ave

et a

l. (

20

08

)

Qu

ave

et a

l. (

20

08

)

Bu

dzy

nsk

a et

al.

(2

01

1)

Mil

lezi

et

al. (2

01

2)

Bu

dzy

nsk

a et

al.

(2

01

1)

Aie

msa

ard

et

al.

(20

11)

Qu

ave

et a

l. (

20

08

)

No

stro

et

al. (2

007

)

Kav

anau

gh

an

d R

ibb

eck

(20

12

)

Qu

ave

et a

l. (

20

08

)

Bu

dzy

nsk

a et

al.

(2

01

1)

Qu

ave

et a

l. (

20

08

)

Qu

ave

et a

l. (

20

08

)

MB

EC

, M

inim

um

Bio

film

Era

dic

atio

n C

once

ntr

atio

n.

IC5

0, co

nce

ntr

atio

n t

hat

red

uce

s th

e 50%

of

bio

film

bio

mas

s.

Eff

ecti

ven

ess

again

st S

. au

reu

s b

iofi

lms

IC5

0 =

8 m

g/L

MB

EC

=

500 m

g/L

Appli

cati

on o

f 1000 m

g/L

red

uce

bio

film

cel

ls i

n 2

.2 l

og

CF

U/c

m2

MB

EC

= 1

700 m

g/L

IC5

0 =

8 m

g/L

IC5

0 =

32 m

g/L

Appli

cati

on o

f 1500 m

g/L

red

uce

s 75%

of

bio

film

bio

mas

s

IC5

0 =

128 m

g/L

IC5

0 =

64 m

g/L

MB

EC

= 1

560 m

g/L

Appli

cati

on o

f 1000 m

g/L

red

uce

bio

film

cel

ls i

n 2

.2 l

og

CF

U/c

m2

MB

EC

= 3

80 m

g/L

Appli

cati

on o

f 4000 m

g/L

for

1 h

era

dic

ates

bio

film

s co

mple

tely

IC5

0 =

32 m

g/L

MB

EC

= 5

00-1

000 m

g/L

MB

EC

= 1

700 m

g/L

IC5

0 =

16 m

g/L

MB

EC

= 7

80 m

g/L

IC5

0 =

16 m

g/L

IC5

0 =

8 m

g/L

Ess

enti

al

oil

Bla

ck h

ore

ho

un

d (

Ba

llota

nig

ra)

Cas

sia

(Cin

na

mo

mu

m a

rom

ati

cum

)

Cit

ron

ella

(C

ymb

opo

go

n n

ard

us)

Clo

ve

(Syz

ygiu

m a

rom

ati

cum

)

Cycl

amen

(C

ycla

men

hed

erif

oli

um

)

Dog

ro

se (

Rosa

ca

nin

a)

Fra

nkin

cen

se (

Bo

swel

lia

pap

yrif

era)

Gia

nt

cane

(Aru

nd

o d

on

ax)

Gra

pe

vin

e (V

itis

vin

ifer

a)

Lav

end

er (

La

van

du

la a

ng

ust

ifoli

a)

Lem

on

(C

itru

s li

mo

nia

)

Lem

on

bal

m (

Mel

issa

off

icin

ali

s)

Lem

on

gra

ss (

Cym

bo

po

gon

cit

rate

s)

Mal

low

(M

alv

a s

ylve

stri

s)

Ore

gan

o (

Ori

ga

nu

m v

ulg

are

)

Red

th

ym

e (T

hym

us

vulg

ari

s)

Rose

mar

y (

Ro

smari

nus

off

icin

ali

s)

Tea

tre

e (M

ela

leu

ca a

lter

nif

oli

a)

Wal

nu

t (J

ugla

ns

regia

)

Wil

d b

lack

ber

ry (

Ru

bu

s ulm

ifoli

us)


53

Nanoparticles: several engineered nanomaterials have also shown strong

antimicrobial properties against S. aureus, including those made of silver (Li et al.,

2011), gold (Bresee et al., 2011) and chitosan (Xing et al., 2009) nanoparticles. This

high efficacy of nanoparticles is accounted by their high reactivity, unique

interactions with biological systems, small size and large surface to volume ratio (Li

et al. 2008; Weir et al. 2008; Taylor and Webster 2009). Moreover, these

nanoparticles can additionally load other antimicrobials by physical encapsulation,

adsorption or chemical conjugation, controlling their release and improving their

bactericidal activity against microbial pathogens (Zhang et al., 2010).

Justificación y Objetivos


57


Teniendo en cuenta que Staphylococcus aureus es uno de los principales agentes

etiológicos de intoxicaciones alimentarias en el mundo y que España es uno de los

mayores productores y consumidores de productos pesqueros en la Unión Europea, hay

dos motivos concretos que justifican la consecución de los objetivos científicos

propuestos en esta tesis:

Que S. aureus es repetidamente detectado en productos pesqueros como

consecuencia de la contaminación cruzada de los manipuladores y superficies de

contacto con el alimento.

Que la capacidad de formación de biopelículas le proporciona a S. aureus una alta

tolerancia a biocidas, permitiéndole persistir a largo plazo en ambientes alimentarios.

En este contexto, y con el objetivo final de mejorar el control de S. aureus en la

industria alimentaria mediante la identificación de los escenarios de mayor riesgo de

contaminación y la evaluación de estrategias de desinfección prometedoras frente a este

patógeno, los siguientes objetivos específicos fueron realizados:

1. Evaluar la incidencia de S. aureus en diferentes productos pesqueros

comercializados. Este objetivo comprende el aislamiento, la caracterización química

y genética de los S. aureus encontrados (incluyendo la tipificación y diferenciación

de las cepas bacterianas mediante RAPD-PCR) y estudios de la capacidad para

producir enterotoxinas y resistir antibióticos.

2. Examinar la prevalencia en plantas de procesado de productos pesqueros de S.

aureus con capacidad para producir enterotoxinas, distinguiendo dos objetivos más

específicos:

2.1. Determinar su capacidad para formar biopelículas sobre materiales habituales

de superficies de contacto con el alimento (acero inoxidable, poliestireno) y

bajo diferentes condiciones ambientales (temperatura, presencia de nutrientes,

osmolaridad) potencialmente presentes en las plantas de procesado.


58

2.2. Evaluar la resistencia de las biopelículas de S. aureus a desinfectantes aplicados

comúnmente en la industria alimentaria (cloruro de benzalconio, ácido

peracético, hipoclorito sódico).

3. Estudiar la eficacia y aplicabilidad de dos estrategias de desinfección innovadoras y

más respetuosas con el medioambiente para controlar las biopelículas de S. aureus

en las plantas de procesado de productos pesqueros:

3.1. Efectividad de la aplicación individual y secuencial de agua electrolizada con

cloruro de benzalconio o ácido peracético.

3.2. Eficacia de la aplicación de aceites esenciales frente a biopelículas de S. aureus

y de la aplicación combinada del más efectivo con cloruro de benzalconio.

61

Justification and Objectives


63


Bearing in mind that Staphylococcus aureus is one of the major bacterial agents

causing foodborne diseases in human worldwide and that Spain is one of the largest

producers and consumers of fishery products in the European Union, two specific

reasons justify the accomplishment of the scientific objectives proposed in this PhD

thesis:

That S. aureus is repeatedly detected in fishery products as a consequence of cross-

contamination from food handlers and food contact surfaces.

That biofilm formation provides S. aureus a high tolerance to biocides allowing a

long-term persistence of this pathogen in food-related environments.

In this context, and with the final aim of improving the control of S. aureus in the

food industry through the identification of the most risky scenarios of contamination

and the evaluation of promising disinfection strategies against this pathogen, the

following specific objectives were carried out:

1. Assessment of the incidence of S. aureus in different commercialized fishery

products. This aim comprises isolation, chemical and genetic characterization of the

S. aureus found (including the typing and differentiation of bacterial strains by

RAPD-PCR) and studies of enterotoxin-producing ability and antibiotic resistance.

2. Examine the prevalence in fishery-processing facilities of putative enterotoxigenic S.

aureus strains, with two more specific objectives:

2.1. Determine their biofilm-forming ability on common food-contact surface

materials (stainless steel, polystyrene) and under different environmental

conditions (temperature, nutrient content, osmolarity) potentially present in

processing plants.

2.2. Assess the resistance of S. aureus biofilms to disinfectants applied commonly in

the food industry (benzalkonium chloride, peracetic acid, sodium hypochlorite).

3. Study the efficacy and applicability of two innovative and more environmentally-

friendly disinfection strategies to control S. aureus biofilms on fishery-processing

plants:


64

3.1. Effectiveness of single and sequential application of electrolyzed water with

benzalkonium chloride or peracetic acid.

3.2. Effectiveness of the application of essential oils against S. aureus biofilms and

combined application of the most effective with benzalkonium chloride.

67

Chapter 1. Incidence and characterization of

Staphylococcus aureus in fishery products

marketed in Galicia (Northwest Spain)

Incidence and characterization of S. aureus in fishery products

69

* Corresponding author at: Instituto de Investigaciones Marinas (CSIC), Eduardo Cabello 6, 36208, Vigo Spain. Tel.: +34 986 231 930; fax: +34 986 292 762.

E-mail address: [email protected] (J.J. Rodríguez-Herrera).

Abstract

A total of 298 fishery products purchased from retail outlets in Galicia (NW Spain)

between January 2008 and May 2009 were analysed for the presence of Staphylococcus

aureus. S. aureus was detected in a significant proportion of products (~ 25%).

Incidence was highest in fresh (43%) and frozen products (30%), but it was high in all

other categories: salted fish (27%), smoked fish (26%), ready-to-cook products (25%),

non-frozen surimis (20%), fish roes (17%) and other ready-to-eat products (10%). A

significant proportion of smoked fish, surimis, fish roes and other ready-to-eat products

did not comply with legal limits in force.

RAPD-PCR of 125 S. aureus isolated from fishery products was carried out using

three primers (AP-7, ERIC-2 and S). Isolates displayed 33 fingerprint patterns. Each

pattern was attributed to a single bacterial clone. Cluster analysis based on similarity

values between RAPD fingerprints did not find relationship between any RAPD pattern

and any product category.

Isolates were also tested for se genes and susceptibility to a range of antibiotics

(cephalothin, clindamycin, chloramphenicol, erythromycin, gentamicin, oxacillin,

penicillin G, tetracycline, vancomycin, methicillin, ciprofloxacin and trimethoprim-

sulfamethoxazole). Most isolates (88%) were found to be sea positive. Putative

enterotoxigenic strains counts reached high risk levels in 17 products. No relationship

was found between the presence of se genes and RAPD patterns. All isolates were

resistant to penicillin, chloramphenicol and ciprofloxacin, and most to tetracycline

(82.4%), but none was methicillin-resistant.

Chapter 1

70

A revision of pre-requisite programs leading to improve hygienic practices in

handling and processing operations from fishing or farming to retail is recommended to

ensure fishery products safety.

Keywords: Staphylococcus aureus; fishery products; retail level; enterotoxin genes;

antibiotic resistance.

1.1 Introduction

Although it is necessary to ensure food safety for the health of consumers and

industry, Salmonella spp., Escherichia coli, Listeria monocytogenes, Staphylococcus

aureus and pathogenic vibrio species have been repeatedly detected in a diverse variety

of fishery products (EFSA, 2010; Garrido et al., 2009; Herrera et al., 2006; Kumar et

al., 2009; Novotny et al., 2004; Papadopoulou et al., 2006; Yang et al., 2008). Novel

trends in food production such as minimal processing, mass production and

globalization, among others, have additionally introduced new factors and conditions

that can enhance the presence and subsequent growth of bacterial pathogens (Abee and

Wouters, 1999; Cebrián et al., 2007; Rendueles et al., 2011).

S. aureus is one of the major bacterial agents causing foodborne diseases in humans

worldwide (EFSA, 2010; Le-Loir et al., 2003). Staphylococcal food poisoning is usually

self-limiting and resolves within 24 to 48 h after onset. Most cases are therefore not

reported to healthcare services. As a result, the actual incidence of staphylococcal food

poisoning is known to be much higher than reported (Lawrynowicz-Paciorek et al.,

2007; Smyth et al., 2004). In addition, the notification of staphylococcal intoxications is

not mandatory in a number of member states of the European Union. Staphylococcal

food poisonings result from the ingestion of food containing staphylococcal

enterotoxins (SEs) preformed by enterotoxigenic strains (Kérouanton et al., 2007; Le-

Loir et al., 2003). SEs are resistant to proteolysis and heat-stable, so the presence of SEs

involves a significant food safety risk (Omoe et al., 2005).

The widespread use of antibiotics has evolved the emergence of multidrug resistant

strains, and it makes eradication more difficult and incidence to increase. Multi-resistant

S. aureus is rather common in hospital settings and farms (Livermore, 2000; Sakoulas

and Moellering, 2008). Community-associated multi-resistant S. aureus is becoming an

emerging problem too (Popovich et al., 2007; Ribeiro et al., 2007; Stankovic et al.,


71

2007). Antibiotic-resistant strains of S. aureus have been detected in food animals (Lee,

2003) and food like meat (Normanno et al., 2007; Pesavento et al., 2007), milk and

dairy products (Gündoğan et al., 2006; Peles et al., 2007; Pereira et al., 2009) and also

fishery products (Beleneva, 2011), and it may be very hazardous for human health.

The identification of bacterial clones with enhanced virulence or increased ability to

spread is important. Nowadays, PCR-based techniques are commonly used for typing,

as they are easy, fast and cost-effective. Among such techniques, random amplified

polymorphic DNA (RAPD-PCR) has been considered a very useful tool for rapid

differentiation of clones with no prior information of the gene sequence (Van-Belkum et

al., 1995; Fueyo et al., 2001; Nema et al., 2007; Nikbakht et al., 2008; Shehata, 2008).

Nowadays, Spain is the largest fishery producer, particularly in Galicia (NW Spain),

and the second largest consumer in the European Union (Eurostat, 2007). However, no

results have been found on the incidence of bacterial pathogens in fishery products

made or sold in Galicia, apart from one study on molluscan shellfish farmed in Galician

waters (Martínez et al., 2009). The situation is not different for fishery products

marketed in other parts of Spain, and only two studies on smoked fish (Garrido et al.,

2009; Herrera et al., 2006) and another study on freshwater fish (González-Rodríguez et

al., 2002) have been carried out in the last decade.

Therefore, the present study was aimed to determine the incidence of S. aureus in

fishery products marketed in Galicia and subsequently identify most common clones by

RAPD-PCR, as well as cases of increased risk according to the presence of enterotoxin

genes and antibiotic-resistance of isolates.

1.2 Materials and Methods

1.2.1 Sampling

A total of 298 fishery products marketed at different retail outlets in Vigo (Galicia,

Northwest Spain) were purchased and analysed between January 2008 and April 2009.

Fourteen samplings (approximately one each month) were carried out. Products were

classified into eight different categories: fresh products, frozen products, salted fish,

ready-to-cook products, smoked fish, fish roes, non-frozen surimis and other ready-to-

eat products (seafood salads, pâtés and anchovies in oil). Between 24 and 43 products of

each category were analysed.

Chapter 1

72

1.2.2 Isolation and identification of S. aureus

About 50 g of product mixed with 200 mL of peptone water was homogenized in a

stomacher masticator (IUL instruments, Spain). Subsequently, homogenates were

serially diluted in peptone water (1:50 and 1:500). Aliquots (0.5 mL) of each dilution

spread onto Baird Parker agar supplemented with egg yolk tellurite emulsion (Biolife,

Italy) (BP-EY). Plates were incubated at 37ºC for 48 h.

Typical colonies of S. aureus as well as non-typical colonies (showing no white

margin and smaller than 2 mm) were counted. Between 1 and 9 colonies from each

product were selected and sub-cultured twice on BP-EY agar for isolation of single

colonies (isolates).

Isolates cultured in Brain Heart Infusion broth (Biolife) (BHI) for 24 h at 37ºC were

subjected to three different biochemical tests: coagulase, DNAse and mannitol

fermentation.

Coagulase production was tested by adding 100 µL of bacterial culture into 300 µL

of reconstituted rabbit plasma with EDTA (Bactident® Coagulase rabbit, Merck,

Germany) followed by incubation of tubes at 37ºC. Clotting of plasma was assessed at

1-h intervals during 6 h and after 24 h of incubation.

Isolates were streaked onto DNAse agar (Cultimed, Panreac Quimica, Spain)

supplemented with D-mannitol and bromothymol blue and plates were incubated at

37ºC for 24 h. The surface of the plates was then flooded with 0.1 N HCl during 15-20

min for DNA precipitation. DNAse activity was observed by the presence of a

transparent halo around the colonies on the agar. Mannitol fermentation was observed

as a colour change of the pH indicator -from blue to yellow- due to acid production.

Colonies found to be coagulase positive, DNAse positive and able to ferment

mannitol (suspected S. aureus) were confirmed to be S. aureus by species-specific 23S

rDNA PCR. Genomic DNA was extracted from 24 h cultures in BHI using an

InstaGene™ Matrix kit (Bio-Rad Laboratories, S.A., Spain) following manufacturer’s

instructions. DNA was quantified by assuming that an absorbance value at 260 nm of

0.100 corresponds to 5 µg/mL of DNA. Primers staur4 (5´-ACGGAGTTACAAAGGA

CGAC-3´) and staur6 (5´-AGCTCAGCCTTAACGAGTAC -3´) (Straub et al., 1999)

were used for each strain. Expected size of amplified PCR products was 1250 bp. Each

PCR mixture contained 100 ng DNA, 1x Taq Buffer Advanced, 2.5 U Taq DNA


73

polymerase (5 Prime, Germany), 40 nmol of each dNTP (Bioline, UK), 0.25 nmol of

forward and reverse primers (Thermo Fisher Scientific, Germany) and sterile Milli-Q

water up to a final volume of 50 µL. PCR was performed with a MyCycler™

Thermocycler (Bio-Rad). The conditions proposed by Vautor et al. (2008) were used to

target the 23S rDNA gen. An initial step of 5 min at 94ºC was followed by 30 cycles of

30 s at 94ºC, 30 s at 58ºC and 75 s at 72ºC, and a final step at 72ºC for 5 min. PCR

products were subjected to electrophoresis on 1.5% agarose gel containing ethidium

bromide for 90 min at 75 V and 100 mAmp. Gels were photographed in a Gel Doc XR

system (Bio-Rad) using the Quantity One® software (Bio-Rad). A DNA ladder of 50-

2000 bp (Hyperladder II, Bioline) was included as a molecular size marker.

Stock cultures of S. aureus isolates were maintained in 50% glycerol (w/w) at

─80ºC. When needed, stock cultures were thawed and subcultured twice in tryptic soy

broth (Cultimed) for 24 h at 37ºC prior to being used.

1.2.3 RAPD

Genotypic characterization of isolates was performed by RAPD-PCR. DNA was

extracted and quantified as previously described from two different cultures of each

isolate to check reproducibility of banding profiles. Primers S (5´-TCACGATGCA-3´)

(Martín et al., 2004), AP-7 (5´-GTGGATGCGA-3´) and ERIC-2 (5´-AAGTAAGTGAC

TGGGGTGAGCG-3´) (Van-Belkum et al., 1995) were individually used in separate

reactions with each isolate. Each PCR mixture consisted of 200 ng DNA; 1x Taq Buffer

Advanced and 2.5 U Taq DNA polymerase (5 Prime); 40 nmol of each dNTP (Bioline);

0.25 nmol primer (Thermo Fisher Scientific) and sterile Milli-Q water up to a final

volume of 50 µL. PCR mixtures with primers AP-7 and ERIC-2 were further

supplemented with 1mM MgCl2 (5 Prime). RAPD-PCR was performed with a

MyCycler™ Thermocycler (Bio-Rad). PCRs containing primer S consisted of an initial

cycle at 95ºC for 5 min, followed by 35 cycles of 95ºC for 1 min, 37ºC for 1 min and

72ºC for 2 min, with a final extension of 5 min at 72ºC. Amplification conditions for

AP-7 and ERIC-2 included a denaturation cycle at 94ºC for 4 min, 35 cycles of 94ºC for

1 min, 25ºC for 1 min and 72ºC for 2 min, and a last extension at 72ºC for 7 min. PCR

products were subjected to electrophoresis on 1.5% agarose gel containing ethidium

bromide as aforementioned. A DNA ladder of 50-2000 bp was included in all gels.

Chapter 1

74

A second-order polynomial relationship between molecular size and mobility was

obtained for each gel (r > 0.99) and used to determine the molecular size of DNA

bands. A RAPD pattern was described as different when at least one band difference

was found. Reproducibility of patterns was checked twice using independent DNA

samples. Variations in band intensity were not considered. Bands too faint to be

reproduced were not considered. A binary value (0 or 1) denoting absence or presence

of each band was assigned to each pattern. Similarity analysis determining the Dice

coefficients (Struelens et al., 1996) was performed by IBM SPSS 19.0. Cluster analysis

by UPGMA (Sneath and Sokal, 1973) and dendrograms were performed with StatistiXL

1.8.

1.2.4 Detection of sea-see and seg-sei genes

A slight modification of the method described by Omoe et al. (2002) was followed to

analyse the presence of staphylococcal enterotoxin (se) genes. Two multiplex PCR

detecting sea-see and seg-sei genes were performed for each strain. DNA was extracted

and quantified as previously described. Primer nucleotide sequences and expected sizes

of amplicons are shown in Table 1.1. Each PCR mixture contained 100 ng DNA; 10x

KCl reaction buffer and 40 nmol of each dNTP (Bioline); 40 pmol SEC-3/SEC-4

primers, 80 pmol SEB-1/SEB-4 primers and 20 pmol for other primers (Thermo Fisher

Scientific); 2.5 U Taq DNA polymerase (5 Prime) and sterile Milli-Q water up to a final

volume of 50 µL. S. aureus ATCC 12600 was used as a negative control in all PCRs,

whereas S. aureus ATCC 13565 was used as a positive control for sea and sed, S.

aureus ATCC 19095 for sec, seg, seh and sei, S. aureus ATCC 14458 for seb, and S.

aureus ATCC 27664 for see. All these strains were obtained from the Spanish Type

Culture Collection (CECT, Valencia, Spain). All PCRs were performed with a

MyCycler™ Thermocycler (Bio-Rad) as follows: an initial cycle of 95ºC for 2 min,

55ºC for 1 min and 68ºC for 2 min, followed by 28 cycles of 95ºC for 1 min, 55ºC for 1

min and 68ºC for 2 min, and a final cycle of 95ºC at 1 min, 55ºC for 1 min and 68ºC for

5 min. PCR products were subjected to electrophoresis on 2.5% agarose gel containing

ethidium bromide. Run conditions and gel display were as aforementioned. A DNA

ladder of 50-2000 bp was also included in all gels.


75

Ta

ble

1.1

. N

ucl

eoti

de

sequen

ces

of

pri

mer

pai

rs a

nd p

redic

ted s

izes

of

resu

ltin

g P

CR

pro

du

cts.

Ref

eren

ce

Bec

ker

et

al. (1

998

)

Om

oe

et a

l. (

20

02)

Am

pli

con

s si

ze (

bp

)

12

7

47

7

27

1

31

9

17

8

28

7

21

3

45

4

Nu

cleo

tid

e se

qu

ence

s (5

´→3´)

CC

T T

TG

GA

A A

CG

GT

T A

AA

AC

G

TC

T G

AA

CC

T T

CC

CA

T C

AA

AA

A C

TC

G C

AT

CA

A A

CT

GA

C A

AA

CG

GC

A G

GT

AC

T C

TA

TA

A G

TG

CC

T G

C

CT

C A

AG

AA

C T

AG

AC

A T

AA

AA

G C

TA

GG

TC

A A

AA

TC

G G

AT

TA

A C

AT

TA

T C

C

CT

A G

TT

TG

G T

AA

TA

T C

TC

CT

T T

AA

AC

G

TT

A A

TG

CT

A T

AT

CT

T A

TA

GG

G T

AA

AC

A T

C

CA

G T

AC

CT

A T

AG

AT

A A

AG

TT

A A

AA

CA

A G

C

TA

A C

TT

AC

C G

TG

GA

C C

CT

TC

AA

G T

AG

AC

A T

TT

TT

G G

CG

TT

C C

AG

A A

CC

AT

C A

AA

CT

C G

TA

TA

G C

GT

C T

AT

AT

G G

AG

GT

A C

AA

CA

C T

GA

C C

TT

TA

C T

TA

TT

T C

GC

TG

T C

GG

T G

AT

AT

T G

GT

GT

A G

GT

AA

C

AT

C C

AT

AT

T C

TT

TG

C C

TT

TA

C C

AG

Pri

mer

SE

A-3

SE

A-4

SE

B-1

SE

B-4

SE

C-3

SE

C-4

SE

D-3

SE

D-4

SE

E-3

SE

E-2

SE

G-1

SE

G-2

SE

H-1

SE

H-2

SE

I-1

SE

I-2

Gen

e

sea

seb

sec

sed

see

seg

seh

sei

Chapter 1

76

1.2.5 Antibiotic susceptibility test

Minimal inhibitory concentration (MIC) of twelve antibiotics was determined against

S. aureus isolated. The EUCAST guidelines (2003) for broth microdilution and disk

diffusion testing were followed.

Broth microdilution test was performed with nine antibiotics: cephalothin, oxacillin,

penicillin G and vancomycin (Sigma-Aldrich Química, Spain); clindamycin and

erythromycin (Acofarma, Spain); and chloramphenicol, gentamicin and tetracycline

(Fagron Iberica, Spain). Adjusted cultures of each isolate to 5·105 CFU/mL in Muller

Hinton broth (Cultimed) supplemented with CaCl2·2H2O (25 mg/mL) and MgCl2·6H2O

(12.5 mg/mL) were exposed to each antibiotic into a microtiter plate (Falcon®, Becton

Dickinson Labware, USA) for 18-20 h (or 24h for oxacillin and vancomycin) at 35ºC.

The OD655nm was measured in an iMark Microplate Reader through Microplate Manager

6® software (Bio-Rad). MIC was defined as the minimum antibiotic concentration at

which no growth was observed.

Disk diffusion test was performed with methicillin (Oxoid, UK), ciprofloxacin and

trimethoprim-sulfamethoxazole (bioMerieux España, Spain). Adjusted cultures were

evenly spread on Muller Hinton agar (Cultimed) and commercially prepared antibiotic

disks were placed on the agar surface. The length of the inhibition halo was measured

after 16-18 h (24 h for methicillin) at 35ºC.

S. aureus ATCC 29213 and S. aureus ATCC 43300 (CECT) was used as negative

control and positive control, respectively. Antibiotic susceptibility was classified as

sensitive, intermediate or resistant on the basis of the breakpoints reported in Table 1.2.

Table 1.2. Antibiotic breakpoints used for interpretation of susceptibility tests.

Antibiotic Breakpoints (µg/mL)

Sensitive ≤ Resistant >

a Cephalothin 8 32

b Chloramphenicol 8 8

b Ciprofloxacin,

b Gentamicin 1 1

b Clindamycin 0.250 0.500

b Erythromycin,

b Tetracycline 1 2

a Methicillin 8 16

a Oxacillin,

b Trimethoprim-Sulfamethoxazole 2 4

b Penicillin G 0.125 0.125

b Vancomycin 2 2

a Breakpoints from the CLSI (2011);

b

breakpoints from the EUCAST (2011).


77

1.2.6 Detection of blaZ and mecA genes

A slight modification of the methods described by Baddour et al. (2007) and Olsen et

al. (2006) was followed for detecting genes encoding penicillin (blaZ) and methicillin

resistance (mecA), respectively.

DNA was extracted with DNeasy® kit (Qiagen, Germany) according to the

manufacturer. Extraction was tested by using λ HindIII DNA Ladder as a reference

(564-23130 bp) (New England BioLabs™, USA).

Primers blaZF487 (5´-TAAGAGATTTGCCTATGCTT-3´) and blaZR373 (5´-

TTAAAGTCTTACCGAA AGCAG-3´) for blaZ gen, and mecA1-F (5´-TGGCTAT

CGTGTCACAATCG-3´) and mecA2-R (5´-CTGGAACTTGTTGAGCAGAG-3´) for

mecA gen, were used. Expected sizes of amplified PCR products were 377 bp for blaZ

gen and 309 bp for mecA gen.

PCR mixtures were composed of 20 ng of DNA; 5 nmol of each dNTP (Invitrogen

Corporation, USA); 2.5 µL of Dynazym buffer 10x and 1.2 U of Dynazym Hot Start

(Bio-Rad); 10 pmol of forward and reverse primer and sterile Milli-Q water up to a final

volume of 25 µL.

PCRs were performed with an 80 Gene Amp PCR System 9700 (Applied

Biosystems, USA). For mecA gen, PCR consisted of an initial denaturation at 94ºC for 5

min, followed by 30 cycles at 94ºC for 1 min, 54ºC for 1 min and 72ºC for 1 min, and a

final cycle at 72ºC for 7 min. Conditions for blaZ gen detection consisted of

denaturation at 94ºC for 5 min, 35 cycles of 94ºC for 1 min, 54ºC for 1 min and 72ºC

for 1 min, and a last extension at 72ºC for 10 min. S. aureus ATCC 29213 was used as

negative control, whereas S. aureus ATCC 43300 was used as positive control.

Amplicons were subjected to electrophoresis on 1.2% agarose gel containing

ethidium bromide for 30 min at 100 V and 200 mAmp. Gels were visualized and saved

in a Typhoon Scanner 8600 (Molecular Dynamics, GE Healthcare, UK). A DNA ladder

of 154-2176 bp (DNA Molecular Weight Marker VI, Roche Applied Science, USA)

was included in all gels.

Chapter 1

78

1.3 Results

1.3.1 Incidence in fishery products

Colonies were observed on BP-EY for 167 out of 298 fishery products. A total of

728 colonies were picked up, isolated and subjected to phenotypic (coagulase, DNAse

and mannitol fermentation) confirmation tests. Out of them, 125 were positive by all

tests and thus identified as S. aureus.

Additionally, one representative isolate of each RAPD global pattern (see below)

was analysed by species-specific 23S rDNA PCR. All of them were confirmed as S.

aureus (Figure 1.1). These isolates were obtained from 75 fishery products, which

represented an incidence of 25.16%.

Figure 1.1. Agarose gels showing the identification of isolates as S. aureus by species-specific

23S rDNA PCR. Lane 1 and 19, DNA molecular size marker (HyperLadder II, 50-2000 bp;

Bioline); lanes 2-18 and 20-35, PCR products for different isolates.


79

As shown in Figure 1.2, incidence was different in each product category, and it

decreased in the following order: fresh products (43%), frozen products (30%), salted

fish (27%), smoked fish (26%), ready-to-cook products (25%), non-frozen surimis

(20%), fish roes (17%), and lastly other ready-to-eat products (10%).

Figure 1.2. Incidence (%) of S. aureus in fishery products marketed at retail level in Galicia.

The number of products surveyed in each category is shown (n).

A significant proportion of surimis (9%), fish roes (4%) and other ready-to-eat

products (5%) as well as ready-to-cook products (13%) exceeded 102 CFU/g of food,

which was the maximum number of S. aureus allowed by legislation in force when the

study was conducted (O. 2/8/1991, RD 3484/2000 and Commission Regulation (EC) No

2073/2005). Additionally, counts were higher than 103 CFU/g in 9 out of 35 surimis, 5

out 41 other ready-to-eat products and 8 out of 40 ready-to-cook products. In the same

way, S. aureus must have been absent in anchovies in oil, but it was detected in 2 out of

4 products.

The present results have also shown that a significant proportion (18.6%) of smoked

products exceeded the limit set for smoked fish products in O. 2/8/1991, that is, 2·101

CFU/g of food. Later, Commission Recommendation 2001/337/EC proposed that

counts higher than 102 CFU/g (M value) should not be permitted in smoked fish. The

value set for M value was exceeded in 16.3% of the smoked products tested in this

study. Additionally, counts were higher than exceeded 103 CFU/g of food in 3 out of 43

smoked products (7%).

Chapter 1

80

Although it is not subject to legal regulations, it is also worthy to mention that the

number of S. aureus was higher than 102 CFU/g of food in 19.4% of fresh products,

14% of frozen products and 13.3% of salted fish, and 103 CFU/g of food in 7% of

frozen products, 4.8% of fresh products and 6.7% of salted fish.

1.3.2 RAPD-PCR

Genotypic characterization of isolates was performed by RAPD-PCR with three

different primers (S, AP-7 and ERIC-2). As shown in Figure 1.3., RAPD analysis with

primer S yielded 13 visually different banding profiles, whereas 12 profiles were

obtained with each of the two other primers. A total of 31 different bands with a size

between 115 and 2292 bp were amplified by primer S, whereas primer AP-7 and ERIC-

2 amplified 19 and 18 different bands ranging from 127 to 2023 bp and 125 to 1420 bp,

respectively. A good reproducibility of patters was achieved when DNA from different

cultures of each isolate was used as a template.

The combination of RAPD fingerprints obtained in separate reactions with different

primers has been employed as a strategy to increase the discriminatory power of the

analysis (Byun et al., 1997; Nema et al., 2007). The combination of the patterns

obtained with the three primers generated a higher discriminatory power (D = 0.926)

than those of single primers (0.818 for primer S, 0.698 for ERIC-2 and 0.622 for AP-7)

as well as of pairwise-combinations of primers (0.883 for S and ERIC-2, 0.878 for S

and AP-7 and 0.850 for AP-7 and ERIC-2). As a result, 33 combined or global

fingerprints were distinguished. A three-digit code number were assigned to combined

patterns, each one corresponding to the number assigned to patterns obtained with

primers S, AP-7 and ERIC-2, respectively. The most common combined pattern was

4.4.1., which was characteristic of 23 isolates. Also, two other patterns (1.1.1 and

13.1.11) were shared by 12 and 16 isolates, respectively. Thus, 40.8% of isolates were

included in one of these three patterns. In contrast, 18 patterns were specific to one

isolate. Isolates showing identical combined RAPD-PCR fingerprints were considered

to be a single genetic type, that is, a single bacterial clone or strain (Fueyo et al., 2001).

Thus, strains St.1.10 and St.1.31 were the most prevalent in fishery products, being

found in 15 and 16 products, respectively.


81

Figure 1.3A-C. Agarose gels showing RAPD-PCR fingerprints obtained for S. aureus isolates

using primers S (A), AP-7 (B) and ERIC-2 (C). Lane 1: DNA Molecular Weight Marker

(HyperLadder II, 50-2000 bp; Bioline). Pattern number is shown on the top of each fingerprint.

Chapter 1

82

Cluster analyses based on similarity measurements from RAPD fingerprints were

carried out with the aim of finding possible relationships between RAPD patterns and

product categories. Cluster analysis of combined RAPD patterns classified isolates into

17 groups at a relative genetic similarity of 0.84 (Figure 1.4). Clusters 1 and 2 were the

largest ones and contained 4 and 11 patterns, respectively, which included most isolates

(31 and 57, respectively) from all product categories. In contrast, there were 10 single

clusters which were formed by only one isolate. The other two single clusters (13.5.11

and 9.4.7) were composed of 3 and 4 isolates.

Figure 1.4. Dendrogram from cluster analysis based on the global combination of RAPD-PCR

patterns obtained with all three primers. Combined patterns were assigned a three-digit code

number, with digits corresponding to numbers assigned to patterns obtained with primers S, AP-

7 and ERIC-2, respectively.

The validity assessment of cluster analyses rendered low values of hierarchical F-

measures for banding patterns generated with primers S (0.314), ERIC-2 (0.311) and

AP-7 (0.289). Similarly, low values were also obtained for F-measure, precision and


83

recall in all cases. Hierarchical F-measure did not increase when banding patterns

generated with each primer were combined (0.329). Values of F-measure, precision and

recall did not increase in this case either. This assessment showed that no cluster was

relatively pure and included most isolates of only one product category. No relationship

was therefore found between any RAPD pattern and any product category.

1.3.3 Presence of enterotoxin genes

Over 91% of S. aureus isolates (n = 114) carried enterotoxin genes, from which 110

were sea positive. However, only four isolates carried several enterotoxin genes. In two

isolates, the seg and sei genes were detected, whereas two others carried the sea, sec and

seh genes. Each of these isolates shared a different combined RAPD pattern, so they

were different bacterial strains. Nevertheless, no further relationship was found between

RAPD patterns of se-carrying isolates.

All isolates sharing an identical global RAPD pattern carried the same se gen pattern,

and this supports the thesis that they are bacterial clones. A total of 26 out of the 33

strains identified by RAPD-PCR analysis were se gene carriers. S. aureus St.1.10 and

St.1.31, which were most prevalent, were se positive. No se-carrying strain was

characteristic of a single product category.

Non-se-carrying strains showed distinct RAPD fingerprints, which were not shared

by any se-carrying strain. However, no further relationship was found between the

presence of se genes and any RAPD pattern. Cluster analysis did not discriminate any

cluster comprising either all non-se-carrying strains or all multi-se-carrying strains only.

A significant proportion of the fishery products tested (23.5%) were contaminated

with se-positive S. aureus. Furthermore, counts of se-carrying S. aureus exceeded 102

and 103 CFU/g of food in 34 and 18 products, respectively, and were even higher than

105 CFU/g of food in two dried-salted tuna loin products (i.e. mojama) and one surimi

product. The incidence of se-positive S. aureus was different in each product category

(Figure 1.5). Isolates carrying se genes were found in all salted fish (n = 8), surimis (n =

7) and fish roes (n = 4) which were contaminated with S. aureus. The presence of se-

positive isolates was also very high in fresh products (94%), smoked fish (91%), ready-

to-cook products (90%), frozen products (85%) and other ready-to-eat products (75%)

in which S. aureus was detected.

Chapter 1

84

Figure 1.5. Presence of se genes (%) in S. aureus isolated from fishery products marketed at

retail level in Galicia. Presence of se genes was defined as the number of se-carrying S. aureus-

containing products respect to the number of S. aureus-containing products. The number of

fishery products carrying se positive S. aureus in each category is shown (n).

The presence of se-positive and se-negative isolates was not detected in a same

fishery product, except in one frozen hake nuggets product and one fresh perch fillet, in

which both types were found. Moreover, one cod pâté with pepper carried sea positive

and sea, sec and seh positive isolates.

1.3.4 Antibiotic sensitivity

No differences were found among isolates with regard to sensitivity profiles, except

for tetracycline. All strains were thus found to be resistant to penicillin G,

chloramphenicol and ciprofloxacin. However, differences were found with regard to

MIC values. Thus, strains St.1.11 and St.1.18 were the most resistant to penicillin G,

St.1.01 and St.1.31 were the most resistant to chloramphenicol and St.1.26 and St.1.33

showed the highest MIC value (1.58 µg/mL) for ciprofloxacin. On the contrary, none of

the strains was found to be resistant to beta-lactam antibiotics such as oxacillin and

methicillin, but they all had intermediate resistance to methicillin (MIC from 9.5 to 14.5

µg/mL). Strains St.1.11 and St.1.13 showed the highest levels of resistance to

methicillin (MIC = 14.5 µg/mL). No strain was resistant to vancomycin, cephalothin,

clindamycin, erythromycin, gentamicin or trimethoprim-sulfamethoxazole.


85

Tetracycline was the only antibiotic on which both resistant- and sensitive-strains

were found. Thus, 17 out of the 33 strains (51.5%) identified by RAPD-PCR analysis

were tetracycline-resistant, being S. aureus St.1.20 and St.1.31 the most resistant.

However, over 82% of isolates were tetracycline-resistant. Tetracycline-resistant

isolates were detected in 65 out of 75 fishery products contaminated with S. aureus

(86.7%). St.1.10 and St.1.31, which were most prevalent, were also tetracycline-

resistant. No differences in tetracycline susceptibility of isolates sharing identical global

RAPD patterns with all three primers were found. However, no relationship was found

between any RAPD pattern and susceptibility to tetracycline.

The incidence of tetracycline-resistant S. aureus was different in each product

category (Figure 1.6). Tetracycline-resistant isolates were detected in all salted fish and

fish roes which were found to be contaminated with S. aureus. Tetracycline-resistant

isolates were also found in most contaminated fishery products of other categories, with

an incidence decreasing in the following order: surimis (93.7%), ready-to-cook products

(87.5%), smoked fish (82.4%), other ready-to-eat products (80.0%), fresh products

(76.0%) and frozen products (64.3%).

Figure 1.6. Tetracycline resistance (%) of S. aureus isolated from fishery products marketed at

retail level in Galicia. Resistance was defined as the number of tetracycline-resistant isolates

with respect to the number of isolates. The number of isolates from each category is shown (n).

Additionally, one representative isolate of each se-carrying strain was tested for the

presence of blaZ and mecA genes, which are major determinants of the resistance of

staphylococci to penicillin (Olsen et al., 2006; Vesterholm-Nielsen et al., 1999) and

methicillin and all other beta-lactam antibiotics (Baddour et al., 2007; Strommenger et

Chapter 1

86

al., 2003), respectively. No PCR product was detected for mecA in any of the 26 isolates

tested (Figure 1.7A), whereas blaZ was detected in all of them (Figure 1.7B). These

results are in agreement with those of antibiotic sensitivity testing.

Figure 1.7A-B. Agarose gels showing PCR products obtained for the putative enterotoxigenic

S. aureus during detection of genes mecA (A) and blaZ (B). Lane 1 and 17, DNA molecular size

marker (DNA Molecular Weight Marker VI, Roche Applied Science, USA); lanes 2-3,

reference strains S. aureus ATCC 29213 and S. aureus ATCC 43300, respectively; lanes 4-16

and 18-30, amplicons of each putative enterotoxigenic strain; lane 31, blank.


87

1.4 Discussion

A high incidence of S. aureus was found in fishery products marketed in Galicia

(25.16%) in the present study. Although data on the incidence of S. aureus in fishery

products was scarce, a high incidence had also been reported over the last ten years in

some studies (Abrahim et al., 2010; Herrera et al., 2006; Oh et al., 2007; Papadopoulou

et al., 2006; Simon and Sanjeev, 2007), and only one work on fishery products collected

at retail outlets in Italy found a low incidence, i.e. < 3% (Normanno et al., 2005). Only a

few studies on microbial safety of fish products marketed in Spain have been carried out

in the last decade (Garrido et al., 2009; González-Rodríguez et al., 2002; Herrera et al.,

2006; Martínez et al., 2009). Considering the importance of fishery products at national

level, the lack of information available on microbial safety in fishery products made or

sold in Spain, and particularly in Galicia, was found surprising. Only Martínez et al.

(2009) tested for the presence of bacterial pathogens in fishery products made in

Galicia, specifically in farmed molluscan shellfish.

A wide range of product categories has been examined in the present work,

comprising between 24 and 43 products of each one. In contrast, previous studies on the

incidence of S. aureus in fish products have focused in only one or two product

categories. Thus, Da-Silva et al. (2010), Herrera et al. (2006), Oh et al. (2007) and

Papadopoulou et al. (2007) only tested fresh fishery products obtained from retail stores

in Brazil, Spain, Korea and Greece, respectively, Simon and Sanjeev (2007) focused on

frozen products and dried fish products sold in India, and Basti et al. (2006) examined

smoked and salted Iranian fish products. Similarly, González-Rodríguez et al. (2002)

only surveyed vacuum-packed cold-smoked freshwater fish.

It was also found surprising that most previous studies dealt with low risk fishery

products, such as fresh fish, frozen products and salted or dried fish products, and

hardly a work on the incidence of S. aureus in fishery products having to comply with

legal regulations in Spain or other European Union member states was found to be

published in the last ten years. As an exception, a study on vacuum-packed cold-smoked

freshwater fish by González-Rodríguez et al. (2002) reported that 3 packages out of 54

were contaminated with S. aureus. In addition, Normanno et al. (2003) reported an

incidence of 10% in a particular raw-eaten Italian fishery product, i.e. strips of

cuttlefish, and Alarcón et al. (2006) detected the presence of S. aureus in 1 out of 10

Chapter 1

88

ready-to-eat products within a study aimed to establish a RTQ-PCR procedure suitable

for detection and quantification of S. aureus in food.

Incidence was found to be high in all categories (10-43%), but notable differences

were found among them. Interestingly, the incidence was higher in those categories not

covered by legislation, that is, fresh products, frozen products and salted fish, than in

those having to comply with legal regulations in force at the time the study was

conducted, i.e. ready-to-eat products and ready meals. This result seems to underline the

effectiveness of regulations on the efforts of the industry to ensure food hygiene.

Although incidence was lower in fishery products subject to legal regulations, the

presence of S. aureus was also detected in a high proportion of ready-to-cook products,

smoked fish, non-frozen surimis, fish roes and other ready-to-eat products. Fishery

products of these categories do not need to be cooked prior to being consumed.

Therefore, the risk for the consumer can become significant if S. aureus is above

regulatory limits, and food exceeding legal limits cannot be placed on the market or

must be recalled. However, this study has revealed that a significant proportion of

fishery products marketed in Galicia (11.3%) did not comply with regulatory limits in

force.

A previous work on cold-smoked fish obtained at retail level in a nearby location

showed that 3 out of 54 packages (5.5%) were contaminated with S. aureus at levels

lower than 4 log CFU/g (González-Rodríguez et al., 2002), but authors did not report if

levels were higher than regulatory limits (O. 2/8/1991) or laid down in Commission

Recommendation 2001/337/EC). In the present study, a higher proportion of smoked

fish (18.6%) was found not to comply with legal regulations.

Incidence was highest in fresh products, followed by frozen products. Incidence

values reported for fresh products in previous studies were also high, ranging between

10 and 30%, with the highest value for those marketed in Northwest Spain (Herrera et

al., 2006). These results were slightly lower to those presented in this work. Incidence

was also reported to be slightly lower in frozen products sold in India, i.e. 17% (Simon

and Sanjeev, 2007). In the present work, samples were homogenized in a lower volume

of diluent and the volume of bacterial suspension spread on agar plates was higher than

or equal to those used in all aforementioned studies. These slight differences in the


89

methodology resulted in a higher limit of detection, and it could account for at least part

of the differences.

Conditions such as low-temperature storage, particularly in frozen fish, a low water

activity typical of frozen and salted products or the activity of specific spoilage

microorganisms of fresh fish prevent S. aureus to grow and, as a result, enterotoxin

production. Also, fresh, frozen and salted products are commonly cooked prior to being

consumed and it should destroy all or most S. aureus. The risk for the consumer is thus

lower and no regulatory limits have been laid down for these fishery products neither in

Spanish nor European legislation. However, an improper storage (temperature abuse) or

processing (e.g. long desalting), can enable SEs to be formed. For instance, desalting at

20ºC was found to result in unsafe levels of S. aureus (≥ 106 CFU/g) in cod, and

possible toxin formation (Pedro et al., 2004). The risk can increase if the number of

microorganisms is low (such as in thawed products), shelf-life is long (such as in salted

products) or the product is consumed raw, undercooked or, in general, lightly processed.

Additionally, staphylococcal enterotoxins (SEs) are highly heat-resistant and therefore,

in many cases, thermal processes cannot be used as a measure to prevent staphylococcal

food poisoning (Balaban and Rasooly, 2000; Cremonesi et al., 2005).

Most of the S. aureus isolates found in this work carried se genes so their incidence

in fishery products marketed in Galicia was high (23.5%). A lower proportion of S.

aureus se positive had been previously found among isolates from fishery products in

other geographical regions (Normanno et al., 2005; Oh et al., 2007; Simon and Sanjeev,

2007).

At present, nine different serological types of SEs (SEA-SEE and SEG-SEJ) have

been proven to have emetic activity (Ortega et al., 2010). Except for two, all se-carrying

isolates found in this study carried the sea gene, and therefore could produce SEA, but

none was found to be seb-see positive. Classical staphylococcal enterotoxins (SEA-

SEE) have been reported to cause 95% of staphylococcal food poisoning. Among them,

SEA is the most common in staphylococcus-related food poisoning (Pinchuk et al.,

2010), probably due to a very high resistance to proteolytic enzymes (Le-Loir et al.,

2003). Several studies have reported that a high proportion of isolates from outbreaks of

staphylococcal food poisoning occurring in South Korea, France, Japan and UK could

produce SEA, either alone or with another toxin (Cha et al., 2006; Kérouanton et al.,

Chapter 1

90

2007; Shimizu et al., 2000; Wieneke et al., 1993). In contrast, sea had been found not to

be the most predominant se gene in fishery products in some previous studies

(Normanno et al., 2005; Simon and Sanjeev, 2007).

Only four of the isolates were multi-se-carriers, two harboured the seg and sei genes

and two others, the sea, sec and seh genes. In contrast, Cha et al. (2006) found that most

isolates (ca. 85%) from staphylococcal food poisoning incidents in South Korea were

multi-se-carriers and detected seg-sei genes in a significant proportion of isolates, either

alone (ca. 4%) or along with other se genes (17.5%). Nonetheless, SEG and SEI have

been considered to play minor role in food poisoning (Chen et al., 2004).

A dose lower than 1 µg of SE has been reported to make symptoms of

staphylococcal food poisoning to appear within 1-6 h after consumption of

contaminated food in an adult healthy individual (FDA, 1992; Pinchuk et al., 2010).

This toxin level can be reached when cell number exceeds 105 CFU/g of food (Bhatia

and Zahoor, 2007). As a preventive measure, legal limits of 102-10

3 CFU/g had been set

for S. aureus in different fishery products. However, in this study, counts of se-carrying

S. aureus exceeded such limits in a significant number of products and in some of them

even by one or two orders of magnitude.

S. aureus has been reported as the third major causative agent of foodborne illness by

fish and fishery products in the European Union (EFSA, 2009a). Moreover, the actual

incidence of staphylococcal food poisoning is known to be much higher than reported

(Lawrynowicz-Paciorek et al., 2007; Smyth et al., 2004). Thus, the notification of

staphylococcal intoxications is not mandatory in many member states of the European

Union and most cases are not reported to healthcare services as they resolve within 24

to 48 h after onset -hospitalization rate was 19.5% for verified outbreaks caused by S.

aureus in 2008 (EFSA, 2010)-. However, microbiological criteria laid down in national

regulations (O. 2/8/1991 and RD 3484/2000) have been recently repealed (RD

135/2010) following Commission Regulation (EC) No 2073/2005 on microbiological

criteria for foodstuff. Coagulase positive staphylococci (mainly S. aureus) are thus no

longer a microbiological criterion for ready-to-cook products and most ready-to-eat

products. At present, S. aureus is set as a process hygiene criterion only for shelled and

shucked products of cooked crustaceans and molluscan shellfish, with a value for M of

103 CFU/g. In the present study, 3 out of 12 products (25%) exceeded this value.


91

The emergence of multidrug resistant pathogens is recognized as an environmental

hazard to the food supply and human health, as it makes eradication more difficult and

incidence to increase (Livermore, 2000; Popovich et al., 2007; Ribeiro et al., 2007). S.

aureus has developed multidrug resistance worldwide, but wide variations in incidence

exist regionally (Gündoğan et al., 2006; Normanno et al., 2007; Peles et al., 2007;

Pesavento et al., 2007). All isolates found in fishery products marketed in Galicia were

resistant to penicillin, chloramphenicol and ciprofloxacin and most of them were

resistant to tetracycline too. Beleneva (2011) also found a high incidence of

ciprofloxacin-resistant S. aureus (84.7%) in fishery products from the Sea of Japan and

South China Sea, but the percentage of penicillin- and tetracycline-resistant strains was

lower (47.2% and 27.5%, respectively). Variations in antibiotic resistance are also the

result of other different factors. For instance, Pereira et al. (2009) isolated a high

number of penicillin-resistant S. aureus (73%) from meat and dairy products in a nearby

geographical area (North of Portugal), but the number of chloramphenicol-,

ciprofloxacin- and tetracycline-resistant isolates was extremely low (0-2%).

Tetracycline was the only antibiotic on which both resistant- and sensitive-strains were

found. Tetracycline-resistance seemed to enhance the presence of S. aureus in fishery

products obtained at retail level in Galicia.

Methicillin-resistant S. aureus (MRSA) are being increasingly found outside clinical

settings (Popovich et al., 2007; Ribeiro et al., 2007; Stankovic et al., 2007). MRSA have

thus been found in food animals (Lee, 2003) and different foods (Gündoğan et al., 2006;

Peles et al., 2007; Pesavento et al., 2007; Pereira et al., 2009) and also in fishery

products recently (Beleneva, 2011). In the present study, however, no MRSA was

isolated from fishery products and no isolate carried the mecA gene, though

intermediate resistance to methicillin was detected in all isolates. Although EFSA

(2009b) reported that there is no current evidence that eating food contaminated with

MRSA may lead to an increased risk of humans becoming healthy carriers or infected

with MRSA strains, the risk of multidrug-resistant MSSA, which are present in food

more frequently than MRSA, has not been examined yet. In fact, it has been recently

underlined the need of incidence studies of multidrug-resistant strains in food to clarify

their public health relevance (Waters et al., 2011). Meanwhile, it is important to take

some preventive control measures.

Chapter 1

92

The identification of bacterial clones with enhanced virulence or increased ability to

spread is important. RAPD is a fast and cost-effective PCR method for typing and

differentiation of bacterial strains with no prior information of the gene sequence.

However, a lack of standardization and a low reproducibility have been claimed as

major drawbacks of RAPD (Deplano et al., 2000; Van-Belkum et al., 1995).

Nevertheless, RAPD-PCR binding patterns were found to be reproducible in this study

when DNA from different cultures of a same isolate was used as a template. A good

reproducibility has been also observed in epidemiological studies using RAPD for

typing S. aureus isolates (Aras et al., 2012; Fueyo et al., 2001; Nikbakht et al., 2008;

Shehata, 2008). Accordingly, RAPD could be used for initial screening of isolates in

public health epidemiological studies (outbreak and endemic strains), prior to other

complementary typing methods such as pulsed-field gel electrophoresis (PFGE) and

multilocus sequence typing (MLST), which must be used for global epidemiology and

population genetic studies of S. aureus (Al-Thawadi et al., 2003; Byun et al., 1997;

Deplano et al., 2006).

The use of RAPD to discriminate strains has been also questioned. Morandi et al.

(2010) has recently reported that multilocus variable number tandem repeat analysis

(MLVA) is more powerful than RAPD-PCR for typing of S. aureus (discriminatory

power of 0.99 and 0.94, respectively). Nonetheless, the discriminatory power of RAPD

was determined by using only one primer (AP-4). In the present study, the combination

of RAPD fingerprints obtained in separate reactions with three different primers

allowed the discriminatory power of the analysis to be increased (increases of 11.7%,

24.6% and 32.8% for primers S, ERIC-2 and AP-7, respectively). Other authors have

used even a much higher number of primers for strain differentiation (Byun et al., 1997;

Nema et al., 2007). This allows strains to be differentiated when RAPD patterns are

rather similar, i.e. low-yield patterns. However, the number of polymorphic bands

generated by primers S, AP-7 and ERIC-2 was considered to be high enough to

distinguish S. aureus strains among isolates found in this study. Nonetheless, it is not

unlikely that the use of a higher number of primers had increased the number of

different strains, but this had been time-consuming.

Isolates sharing a same global RAPD fingerprint also showed identical enterotoxin

gene and antibiotic susceptibility patterns, and this supports the thesis that RAPD


93

analysis allowed bacterial clones to be distinguished. Nema et al. (2007) also found that

S. aureus isolates with a same RAPD fingerprint had identical se gen patterns and

suggested a clonal origin of isolates.

Cluster analyses based on similarity measurements between RAPD patterns found no

relationship between any RAPD pattern and any product category and, though se-

negative and se-positive strains did not shared RAPD fingerprints, no further

relationship was found between the presence of se genes and any RAPD pattern either.

The use of RAPD-PCR fingerprinting with several primers to assess the genetic

relationship between S. aureus isolated from fishery products marketed in Galicia has

therefore exclude a clonal origin.

1.5 Conclusions

A significant proportion (~ 25%) of fishery products surveyed from retail sector in

Galicia in 2008 and 2009 was found to be contaminated with S. aureus, mostly with se-

carrying strains. About 12% of products did not comply with regulatory limits, and a

higher proportion of products not subject to regulations were contaminated too. These

results suggest some effect of regulations on the efforts of the industry to ensure food

hygiene.

However, a number of microbiological criteria laid down in national regulations have

been recently repealed and coagulase positive staphylococci (mainly S. aureus) are thus

no longer a microbiological criterion for ready-to-cook products and most ready-to-eat

products. However, S. aureus has been reported as the third major causative agent of

foodborne illness by fish and fish products in the European Union (EFSA, 2009a). In

addition, the actual incidence of staphylococcal food poisoning is known to be much

higher than reported -the notification of is not mandatory in many member states of the

European Union and most cases are not reported to healthcare services as they resolve

within 24 to 48 h after onset-.

A revision of pre-requisite programs and an improvement of hygienic practices in

handling and processing operations from fishing or farming to retail outlet is therefore

recommended in order to ensure the safety of fishery products marketed in Galicia.

Nonetheless, at present, S. aureus is still under surveillance by a significant part of the

industrial sector.

95

Chapter 2. Impact of food-related

environmental factors on the adherence and

biofilm formation of natural Staphylococcus

aureus isolated from fishery products

Impact of food-related environmental factors on the biofilm formation of S. aureus

97

Abstract

Staphylococcus aureus is a pathogenic bacterium capable of developing biofilms on

food-processing surfaces, a pathway leading to cross-contamination of foods. The

purpose of this study was to investigate the influence of environmental stress factors

found during seafood production on the adhesion and biofilm-forming properties of S.

aureus. Adhesion and biofilm assays were performed on 26 S. aureus isolated from

seafoods and two S. aureus reference strains (ATCC 6538 and ATCC 43300). Cell

surface properties were evaluated by affinity measurements to solvents in a partitioning

test, while adhesion and biofilm assays were performed in polystyrene microplates

under different stress conditions of temperature, osmolarity and nutrients content. The

expression of genes implicated in the regulation of biofilm formation (icaA, sarA, rbf

and σB) was analysed by reverse transcription and quantitative real time PCR.

In general, S. aureus isolates showed moderate hydrophobic properties and a marked

Lewis-base character. Initial adhesion to polystyrene was positively correlated with the

ionic strength of the growth medium. Most of the strains had a higher biofilm

production at 37ºC than at 25ºC, promoted by the addition of glucose, whereas NaCl

and MgCl2 had a lower impact markedly affected by incubation temperatures. Principal

Component Analysis revealed a considerable variability in adhesion and biofilm-

forming properties between S. aureus isolates. Transcriptional analysis also indicated

variations in gene expression between three characteristic isolates under different

environmental conditions. These results suggested that the prevalence of S. aureus

strains on food-processing surfaces is above all conditioned by the ability to adapt to the

environmental stress conditions present during food production.

Chapter 2

98

These findings are relevant for food safety and may be of importance when choosing

the safest environmental conditions and material during processing, packaging and

storage of seafood products.

Keywords: Staphylococcus aureus; Polystyrene; Hydrophobicity; Adhesion;

Biofilm; icaA.

2.1 Introduction

Staphylococcus aureus is a common human pathogen responsible for food-borne

intoxications worldwide, caused by the ingestion of food containing staphylococcal

heat-stable enterotoxins (Lowy, 1998; Le-Loir et al., 2003). The greatest risk of

staphylococcal food poisoning is associated with food contaminated with S. aureus after

the normal microflora has been destroyed or inhibited (Bore et al., 2007). In 2009, the

European Union witnessed staphylococcal outbreaks which led to a hospitalization rate

of 16.9% (EFSA, 2011). Both food products and food-contact surfaces are often

contaminated through handling during processing and packaging (Sattar et al., 2001;

DeVita et al., 2007; Simon and Sanjeev, 2007), as S. aureus is part of the normal

microbiota associated with human skin, throat and nose. Consequently, S. aureus has

been repeatedly detected in a diverse variety of food, including seafood (Novotny et al.,

2004; Herrera et al., 2006; Papadopoulou et al., 2007). One recent study (Vázquez-

Sánchez et al., 2012) reported a high incidence of S. aureus (~ 25%) in seafood

marketed in Spain, which is the largest seafood producer and the second largest

consumer in the European Union (Eurostat, 2007).

Biofilm is considered as part of the normal life cycle of S. aureus in the environment

(Otto, 2008), in which planktonic cells present attach to solid surfaces, proliferating and

accumulating in multilayer cell clusters embedded in an organic polymer matrix. This

structure protects the bacterial community from environmental stresses, from the host

immune system and from antibiotic attacks, as opposed to the situation for vulnerable

and exposed planktonic cells (Costerton et al., 1987). This may contribute to the

persistence of S. aureus in food-processing environments, consequently increasing

cross-contamination risks as well as subsequent economic losses due to recalls of

contaminated food products.


99

Several studies have shown the attachment of S. aureus on work surfaces such as

polystyrene, polypropylene, stainless steel and glass, and also in food products

(Costerton et al., 1978; Sattar et al., 2001; Herrera et al., 2006; DeVita et al., 2007;

Simon and Sanjeev, 2007). However, changes in surface physicochemical properties

and substratum topography, as well as in environmental factors such as osmolarity,

nutrient content and temperature may lead to staphylococcal biofilm development and,

consequently, influence their persistence on food-contact environments (Kumar and

Anand, 1998; Ammendolia et al., 1999; Bos et al., 1999; Poulsen, 1999; Akpolat et al.,

2003; Møretrø et al., 2003; Planchon et al., 2006; Rode et al., 2007; Pagedar et al.,

2010; Xu et al., 2010).

The extracellular matrix of S. aureus is mainly composed by polysaccharide

intercellular adhesins (PIA), i.e., poly-β(1,6)-N-acetyl-d-glucosamines (PNAG), which

are synthetized by N-acetyl-glucosamine transferase (Cramton et al., 1999; Maira-Litrán

et al., 2002; Fitzpatrick et al., 2005; O’Gara, 2007). This enzyme is induced by the co-

expression of icaA with icaD, products of the chromosomal intercellular adhesion (ica)

operon carried by most S. aureus strains (Cramton et al., 1999; Maira-Litrán et al.,

2002; Jefferson et al., 2003; Fitzpatrick et al., 2005). The expression of the ica operon is

controlled by the repressor icaR, which is regulated by the staphylococcus accessory

regulator (sarA), the stress-induced sigma factor B (σB) (Cerca et al., 2008) and,

indirectly, by the rbf gene (Cue et al., 2009), among others. These genes are also

involved in the resistance of S. aureus to various environmental stresses (Gertz et al.,

2000; Rachid et al., 2000; Lim et al., 2004).

The present study aimed at investigating the persistence of 26 natural S. aureus

isolates on polystyrene surfaces, a material frequently used in the food industry, through

the evaluation of their physicochemical, adhesion and biofilm-forming properties under

different environmental stress conditions found during processing, packaging and

storage of food products. Moreover, the variability of the expression of genes

implicated in the regulation of biofilm formation between three strains selected during

the screening was also investigated under different stress conditions.

Chapter 2

100

2.2 Materials and Methods

2.2.1 Bacterial strains and growth conditions

Twenty six S. aureus isolates from seafood marketed in Galicia (Northwest Spain)

were investigated. They were previously identified as S. aureus by specific biochemical

(coagulase, DNAse and mannitol fermentation) and genetic tests (23s rDNA) and

characterized by RAPD-PCR (Vázquez-Sánchez et al., 2012). These isolates carried sea

(n = 22), sea-c-h (n = 2) or seg-i (n = 2) genes, whose expression produce enterotoxins.

S. aureus ATCC 6538 (a known biofilm former) and ATCC 43300 (MRSA strain),

provided from the Spanish Type Culture Collection (CECT, Valencia, Spain), were

used as reference strains. Stock cultures were maintained in 20% glycerol at ─80ºC. All

strains were thawed and subcultured in tryptic soy broth (TSB, Oxoid, UK) for 24 h at

37ºC, 200 rpm prior to use.

2.2.2 Evaluation of bacterial cell surface physicochemical properties

Microbial Adhesion to Solvents (MATS) was used as a method to determine the

hydrophobic character of the cell surface of S. aureus strains and their Lewis acid-base

properties (Bellon-Fontaine et al., 1996). This method is based on the comparison

between microbial cell surface affinity to a monopolar solvent and an apolar solvent,

which both exhibit similar Lifshitz-van der Waals surface tension components.

Chloroform (an electron-acceptor solvent), hexadecane (nonpolar solvent), ethyl

acetate (an electron-donor solvent) and decane (nonpolar solvent) were used of the

highest purity grade (Sigma-Aldrich, USA). Experimentally, overnight bacterial

cultures were washed twice in phosphate buffer (7.6 g/L NaCl, 0.2 g/L KCl, 0.245 g/L

Na2HPO4 and 0.71 g/L K2HPO4; Merck, Inc.) and resuspended to a final OD400nm of 0.8

(~ 108 CFU/mL). Individual bacterial suspensions (2.4 mL) were first mixed with 0.4

mL of the respective solvent and then manually shaken for 10 s prior to vortexing for 50

s. The mixture was allowed to stand for 15 min to ensure complete separation of phases.

1 mL from the aqueous phase was removed and the final OD400nm measured. The

percentage of cells residing in the solvent was calculated by:


101

where (ODi) was the optical density of the bacterial suspension before mixing with the

solvent and (ODf) the absorbance after mixing and phase separation. Each measurement

was performed in triplicate and the experiment was performed twice using independent

bacterial cultures.

2.2.3 Measurement of the adherence ability to polystyrene at different

ionic strength conditions

The ability of S. aureus strains to adhere to polystyrene was evaluated in terms of

biomass using the crystal violet method described by Giaouris et al. (2009), but with

some modifications.

Overnight cultures were washed twice and resuspended to a final OD600nm of 0.8 in

150 mM NaCl or 1.5 mM NaCl. 200 µL of each sample was added in a flat-bottomed

96-well microtiter plate with Nunclon Surface (Nunc, Denmark) and then incubated for

4 h at 25ºC. After measuring the OD600nm, the microplates were washed three times with

peptone water (Oxoid, UK), using an automatic microplate washer (Wellwash AC,

Thermo Electron Corporation, Inc.), and air-dried for 2 h. Wells were then stained for

15 min using 150 µL of 0.5% (w/v) Crystal Violet (CV) (Merck, Inc.) followed by three

rinsing steps with distilled water. The microplates were air-dried for 15 min and the

bound CV was extracted with 150 µL of 33% (v/v) Glacial Acetic Acid (Merck, Inc.)

for 30 min at room temperature. 100 µL of the mixture was diluted in a new microplate

with 100 µL of 33% Glacial Acetic Acid prior to read the OD562nm. Each measurement

was performed in triplicate and the experiment was repeated twice using independent

bacterial cultures.

2.2.4 Quantification of biofilm formation on polystyrene under different

environmental conditions

The biofilm-forming ability of S. aureus strains on polystyrene microtiter plates was

also investigated in terms of biomass using an optimized protocol based on previously

described methods (Song and Leff, 2006; Rode et al., 2007; Peeters et al., 2008).

Each well was added with 100 μL of growth medium and 100 μL of an overnight

bacterial culture diluted 1:100 in TSB. Negative control wells contained TSB only.

Biofilm formation was evaluated after 24 and 48 h in TSB with or without 5% glucose,

Chapter 2

102

5% NaCl, 5% glucose + 5% NaCl, 0.1 mM MgCl2 or 1 mM MgCl2 (Merck, Inc.) at 25

and 37ºC. After measuring the OD600nm, the microplates were washed three times with

peptone water using the automatic microplate washer and air-dried for 2 h. The

microplates were then stained with 150 µL of 0.5% (w/v) CV for 15 min followed by

three rinsing steps with distilled water. After air-dried for 15 min, the bound CV was

extracted with 150 µL of 33% (v/v) Glacial Acetic Acid for 30 min. The mixture added

to a new microplate was then diluted 1:1 in 33% Glacial Acetic Acid prior to read the

OD562nm. Each measurement was performed in triplicate and the experiment was

repeated twice using independent bacterial cultures.

2.2.5 Transcriptional analysis

To assess the expression levels of the genes reported in Table 2.1, RNA was

extracted from St.1.07, St.1.14 and St.1.29 grown in TSB with or without 5% glucose,

5% NaCl or 5% glucose + 5% NaCl. An overnight culture was diluted 1:100 in each

medium and cultivated at 37ºC with 200 rpm of agitation until an OD600 ~ 0.5. After

incubation, two volumes of bacterial culture were diluted in four volumes of

RNAprotect Bacteria Reagent (Qiagen, Hilden, Germany). The mixture was vortexed

for 15 s, incubated for 5 min at room temperature and centrifuged (5000 × g) for 10 min

at room temperature. The supernatant was discarded and 200 μL of a mixture containing

TE buffer, 40 mg/mL lysozyme and 1 mg/mL lysostaphin (Sigma, USA) was added for

enzymatic lysis of bacteria. RNA was isolated using the Qiagen RNeasy Mini Kit

(Qiagen, Hilden, Germany), following the manufacturer's instructions and including a

DNase treatment. The concentration and purity of total RNA were analysed using a

NanoDrop, ND-1000 spectrophotometer (NanoDrop Technologies, Inc.).

Reverse transcription of the RNA isolated was carried out using random primers, as

previously described (Rode et al., 2007) with slight modifications. A reaction mixture

(13 μL) with 300 ng RNA, 100 ng Random Primers and 10 mM of each dNTP

(Invitrogen) was denatured at 65ºC for 5 min, incubated on ice immediately for at least

1 min and centrifuged briefly. A mixture (6 μL) of 5x first strand buffer, 0.1 M DTT

and 200 U Superscript III reverse transcriptase (Invitrogen) was then added to the

reaction. The samples were incubated at 25ºC for 5 min, heated at 50ºC for 45 min and

immediately incubated at 70ºC for 15 min to inactivate the reaction. A brief

centrifugation between each step was done. Six reverse transcriptase reactions were


103

made for each biological replicate of RNA, of which three were without enzyme as

negative controls.

Quantitative real-time PCR (qRT-PCR) was performed in an Abi Prism 7900 HT

Sequence Detection System (Applied Biosystems, Inc.). The PCR mixture contained 1×

TaqMan Buffer A, 5 mM MgCl2, 0.2 mM of dATP, dCTP and dGTP, 0.4 mM dUTP,

0.2 μM primer, 0.1 μM probe, 0.1 U AmpErase uracil N-glycosylase, 1.25 U Ampli-Taq

Gold DNA Polymerase (Applied Biosystems, Roche, Inc.), 10 ng of cDNA and dH2O

ultrapure DNAse and RNAse free (Gibco, Invitrogen Corporation) up to a final volume

of 25 μL. Primers and Taqman® probes were designed previously by Rode et al. (2007).

Reaction mixtures were subjected to an initial cycle of 50ºC for 2 min and 95ºC for 10

min, followed by 40 cycles of 95ºC for 30 s and 60ºC for 1 min.

CT values were estimated on SDS 2.2 software (Applied Biosystems, Inc.). The

difference between CT of the reference gene 16S and CT of other gene analysed (ΔCT)

were calculated to see possible changes in gene expression. One unit change represents

a log of 2-fold change.

2.2.6 Statistical analysis

Results from the analytical determinations were statistically treated with the software

package IBM SPSS 19.0. They were averaged and the standard error of the mean was

calculated. Data of the adhesion and biofilm formation assays were normalized and

expressed as OD562nm/OD600nm, due to the variation in total growth at 25ºC and 37ºC and

to have a clearer view of biofilm formation for the conditions where growth was

limited, as Rode et al. (2007) proposed.

Significance of the data was determined using a one way ANOVA and the

homogeneity of variances was examined by a post-hoc least significant difference

(LSD) test. Otherwise, a Dunnett´s T3 test was performed. An independent-samples T

test was also done to compare strains in pairs. Bivariate correlations were analysed

using the Pearson correlation coefficient. Significance was expressed at the 95%

confidence level (P < 0.05) or greater.

Principal Components Analysis (PCA) was performed to group the 28 S. aureus

strains by their similar physicochemical, adhesion and biofilm formation properties

showed on polystyrene. Varimax normalization method with Kaiser was used to build

the rotated component matrix.

Chapter 2

104

Ta

ble

2.1

. N

ucl

eoti

de

seq

uen

ces

of

pri

mer

pai

rs (

F:

forw

ard;

R:

rew

ard)

and T

aqm

an p

robes

(P

r) u

sed

in

th

e tr

ansc

ripti

on

al a

nal

ysi

s.

Ref

eren

ce

Bo

re e

t al

. (2

00

7)

Val

le e

t al

. (2

003

)

Lim

et

al. (2

00

4)

Wei

nri

ck e

t al

. (2

00

4)

Wei

nri

ck e

t al

. (2

00

4)

Nu

cleo

tid

e se

qu

ence

5′ →

3′

CG

T A

GG

TG

G C

AA

GC

G T

TA

TC

C G

GA

CC

A G

CA

GC

C G

CG

GT

A A

T

CG

C G

CT

TT

A C

GC

CC

A A

TA

TG

G A

TG

TT

G G

TT

CC

A G

AA

AC

A T

TG

GG

A G

TG

A A

CC

GC

T T

GC

CA

T G

TG

CA

C G

CG

TT

G C

TT

CC

A A

AG

A

TG

G A

TG

TT

G G

TT

CC

A G

AA

AC

A T

TG

GG

A G

TT

A G

AA

GG

A A

TC

TT

T A

AA

AC

C T

TA

TT

G A

AT

AA

TT

G T

GA

AT

T T

TT

CT

T C

TT

CG

G A

CA

TC

T T

TC

AT

C A

TG

CT

C A

TT

AC

G T

TT

TT

T A

TC

GA

A G

TA

AT

C T

TT

T T

TT

TA

C G

TT

GT

T G

TG

CA

T T

AA

CA

CA

T T

TA

AA

C T

AC

AA

A C

AA

CC

A C

AA

GT

T G

AG

A A

GT

GT

T A

GA

AG

C A

AT

GG

A A

AT

GG

G A

CA

AA

G T

TA

TA

A T

AT

A G

CT

GA

T C

GA

TT

A G

AA

GT

C T

CA

GA

A G

TC

A A

TG

GA

A T

GA

TC

A A

CA

CT

T A

AC

G

Na

me

16

S-P

r

16

S-F

16

S-R

ic

aA-P

r

icaA

-F

icaA

-R

rb

f-P

r

rbf-

F

rbf-

R

sa

rA-P

r

sarA

-F

sarA

-R

si

gB

-Pr

sig

B-F

sig

B-R

Gen

e

16

S

ica

A

rbf

sarA

σB


105

2.3 Results

2.3.1 Surface hydrophobicity and electron donor/acceptor character

The physicochemical surface properties of the 28 S. aureus strains were studied to

estimate their potential for adhesion and subsequent biofilm formation on surfaces.

Affinities of the strains to different polar and apolar solvents are shown in Figure 2.1.

Considerable variations in the percentage of adhesion to decane between S. aureus

tested strains reveal the degree of diversity in their hydrophobic character. Affinity to

decane ranged from 22.32% to 74.82%. However, affinity to hexadecane were less

variable ranging from 56.40% to 84.14%, revealing a moderate hydrophobic character

for the majority of S. aureus tested strains. High percentage of adhesion to chloroform

was observed for all tested strains (ranging between 74.37% and 95.75%), which in all

cases were higher than that to hexadecane. This also reveals the diversity in electron

donor (Lewis base) properties among tested S. aureus, highlighting the strain St.1.19

with the highest electron donor character. S. aureus tested strains generally expressed

non electron acceptor (Lewis acid) properties, as seen by the higher affinity to decane

compared to ethyl acetate with values below 19.75%.

2.3.2 Adherence ability of S. aureus to polystyrene surfaces

Initial adhesion to polystyrene surfaces of the 28 S. aureus strains to polystyrene was

quantified in terms of biomass at two different ionic strengths (1.5 mM and 150 mM

NaCl) to evaluate their electrostatic interactions.

The results showed that initial adhesion to polystyrene was positively correlated (r =

0.577, P < 0.01) with ionic strengths presented in the suspension. Thus, initial adhesion

to polystyrene was reduced at lower ionic strength conditions compared to high ionic

conditions, except for strains St.1.08 and St.1.21 (Figure 2.2). Moreover, the variability

of adhesive properties to polystyrene among S. aureus strains at low ionic strength

medium may also be an indication of the diversity in cell wall electronegativity among

the tested S. aureus strains, as previously described (Giaouris et al., 2009). The strains

St.1.08 and St.1.09 showed the most remarkable adherence ability under low and high

ionic strength conditions, respectively.

Chapter 2

106

Fig

ure

2.1

. A

ffin

ity o

f S

. a

ure

us

stra

ins

(n =

28)

to t

he

solv

ents

chlo

rofo

rm,

hex

adec

ane,

dec

ane

and

eth

yl

acet

ate.

Mea

n a

nd S

D v

alu

es:

thre

e

repli

cate

s o

f ea

ch s

amp

le.

Dif

fere

nt

lett

ers

on t

he

top o

f ea

ch c

olu

mn s

how

sig

nif

ican

t dif

fere

nce

s (P

< 0

.05

) in

aff

init

y t

o e

ach

solv

ent

bet

wee

n t

he

stra

ins

test

ed.


107

Fig

ure

2.2

. In

itia

l ad

hes

ion t

o p

oly

styre

ne

surf

aces

of

S.

aure

us

stra

ins

(n =

28)

under

dif

fere

nt

ionic

str

ength

co

nd

itio

ns

(NaC

l 1

.5 m

M a

nd

15

0 m

M).

Adhes

ion

abil

ity o

f ea

ch s

trai

n w

as e

xpre

ssed

in t

erm

s of

bio

film

bio

mas

s af

ter

4 h

at

25ºC

. M

ean

an

d S

D v

alu

es:

thre

e re

pli

cate

s o

f ea

ch s

amp

le.

Sig

nif

ican

t dif

fere

nce

s (P

< 0

.05)

bet

wee

n t

he

adher

ence

abil

ity o

f st

rain

s at

eac

h c

ondit

ion w

ere

ind

icat

ed b

y d

iffe

ren

t le

tter

s o

n t

he

top

of

each

colu

mn.

Chapter 2

108

2.3.3 Biofilm formation on polystyrene surfaces under different

environmental conditions

The ability of the 28 S. aureus strains to develop biofilms on polystyrene surfaces

under different conditions of temperature (25ºC and 37ºC), osmolarity and nutrient

content (TSB with or without 5% glucose, 5% NaCl, 5% glucose + 5% NaCl, 0.1 mM

MgCl2 and 1 mM MgCl2) was investigated after 24 and 48 h to understand the direct

effects of environmental factors in staphylococcal biofilm formation. These two

temperatures were selected by their relevance to the food industry and hospitals (25ºC)

and in infectious disease (37ºC). To compensate for variations in cell mass at stationary

phase at the two different temperatures, the biofilm formation values were expressed as

OD562nm/OD600nm. Significant differences (P < 0.05) between strains for each treatment

and viceversa were observed, as indicated by the different letters showed in Figure 2.3.

2.3.3.1 Effect of incubation temperature

Biofilm formation in a medium without nutrient addition (TSB only) was positively

correlated (r = 0.386, P < 0.01) with the temperature of incubation. Thus, incubation at

37ºC increased biofilm-forming ability for the majority of tested isolates (84%),

compared to incubation at 25ºC. S. aureus St.1.22 and St.1.11 showed the highest

biofilm formation at 37ºC, while St.1.31 was able to form biofilms with high cell

densities at 25ºC (Figure 2.3A).

Biofilm formation was also positively correlated (P < 0.01) with incubation

temperature when TSB was added with 5% glucose (r = 0.522), 5% glucose + 5% NaCl

(r = 0.637), 0.1 mM MgCl2 (r = 0.487) or 1 mM MgCl2 (r = 0.405), but addition of 5%

NaCl generated a negative correlation (r = ‒0.418, P < 0.01). In fact, 78.5% of the

strains showed a higher biofilm formation in TSB with 5% NaCl when they were

incubated at 25ºC than at 37ºC.

2.3.3.2 Effect of glucose and NaCl addition

Addition of 5% glucose to TSB generally led to enhanced staphylococcal biofilm

formation (Figure 2.3B), as shown its positive correlation (P < 0.01) with biofilm

formation under all tested conditions (Table 2.2). However, these increases on biofilm

development with the addition of glucose were affected by incubation temperatures. The


109

highest increases in biofilm formation with the addition of 5% glucose were produced in

the first 24 h at 37ºC and after 48 h at 25ºC, with 3-fold and 2-fold increases

respectively. In the presence of 5% glucose, isolates St.1.01, St.1.02, St.1.04 and St.1.08

expressed a 4-fold biofilm increase after 24 h at 37ºC, while isolates St.1.05 and St.1.06

showed 5-fold increases after 48 h at 25ºC.

The effect of NaCl on biofilm formation was markedly affected by incubation

temperatures. Thus, a negative correlation (P < 0.01) at 37ºC between biofilm formation

and the addition of 5% NaCl was observed (Table 2.2). In fact, 75% of tested isolates

expressed lower biofilm formation in environments with supplemented salt than those

grown in the absence of salt (Figure 2.3C). Nevertheless, a positive correlation (P <

0.01) was reported at 25ºC for the first 24 h between NaCl addition and biofilm

formation, slightly improving the production of biofilm by most isolates (75%). Under

similar conditions, isolates St.1.02, St.1.21 and St.1.29 grown in the presence of 5%

NaCl showed a remarkable 2-fold increase in biofilm formation compared to those

grown in the absence of salt. These isolates were isolated from a Paella (containing

mussels and squids), frozen shelled prawns and a Panga fillet respectively, three

seafood products with high amounts of NaCl (> 100 mg per 100 g of product) (FEM

and FROM, 2012). No significant correlation was observed after 48 h at 25ºC between

biofilm formation and the addition of NaCl.

Comparing with individual effects, no synergy was observed between the addition of

glucose and NaCl (Figure 2.3D). Moreover, no significant correlations were observed

between biofilm formation and the addition of both nutrients, except a negative

correlation (P < 0.01) reported when the strains were incubated for 24 h at 37ºC. The

addition of 5% glucose + 5% NaCl therefore slightly increased the biofilm formation

compared to growth in the absence of glucose and NaCl in 64.3% of all tested isolates

after 48 h growth for both 25ºC and 37ºC. Two-fold biofilm increases were observed in

non-supplemented TSB for isolates St.1.07, St.1.12 and St.1.28 grown at 25ºC, and

isolates St.1.03, St.1.05, St.1.06, St.1.08 and St.1.14 grown at 37ºC.

Chapter 2

110

Fig

ure

2.3

A-F

. B

iofi

lm f

orm

atio

n o

f S.

aure

us

stra

ins

(n =

28)

on p

oly

styre

ne

in T

SB

on

ly (

A)

or

added

wit

h 5

% g

luco

se (

B),

5%

NaC

l (C

),

5%

glu

cose

+ 5

% N

aCl

(D),

0.1

mM

MgC

l 2 (

E)

or

1 m

M M

gC

l 2 (

F).

Mea

n a

nd S

D v

alu

es:

nin

e re

pli

cate

s o

f ea

ch s

amp

le.

Dif

fere

nt

lett

ers

on

each

co

lum

n i

ndic

ate

sign

ific

ant

dif

fere

nce

s (P

< 0

.05)

in b

iofi

lm f

orm

atio

n b

etw

een s

trai

ns

for

each

co

ndit

ion

.


111

2.3.3.3 Effect of MgCl2 addition

Generally, addition of 0.1 mM MgCl2 did not significantly affect biofilm formation

compared to growth in the absence of MgCl2 (Figure 2.3E). No correlation was

observed between the addition of 0.1 mM MgCl2 and biofilm formation, except a

positive correlation (P < 0.05) for growth at 37ºC after 48 h (Table 2.2). In fact, 57.1%

of the strains increased significantly their biofilm formation with the addition of MgCl2

under these conditions, highlighting St.1.05, St.1.07, St.1.14, St.1.20 and St.1.31 with a

3-fold biofilm increase.

Otherwise, the increment of the MgCl2 concentration from 0.1 to 1 mM did not

induce an increase on the biofilm formation (Figure 2.3F). Consequently, only 21.4%

isolates showed on average a 2-fold increase with the addition of 1 mM MgCl2 after 48

h at 37ºC, highlighting St.1.01 and St.1.03 isolated from smoked swordfish, a seafood

with high magnesium levels (57 mg per 100 g of product) (FEM and FROM, 2012).

Biofilm formation was positively correlated (P < 0.05) with the addition of 1 mM

MgCl2 after 48 h at 37ºC, but no significant correlations were observed under the other

conditions tested (Table 2.2).

Table 2.2. Correlations between the biofilm formation and nutrient content expressed as r

values. An r value of zero indicates no correlation, whereas a value of 1 or ‒1 indicates a perfect

positive or negative correlation.

Nutrient added Incubation condition

25ºC 24 h 25ºC 48 h 37ºC 24 h 37ºC 48 h

5% glucose 0.478b 0.588

b 0.733

b 0.470

b

5% NaCl 0.499b 0.082 ‒0.521

b ‒0.439

b

5% glucose + 5% NaCl ‒0.031 ‒0.040 ‒0.356b 0.133

0.1 mM MgCl2 0.148 0.033 0.149 0.176a

1 mM MgCl2 0.139 0.049 0.068 0.161a

a P < 0.05;

b P < 0.01

Chapter 2

112

2.3.4 Multivariate analysis of the physicochemical, adhesion and

biofilm-forming properties of the 28 S. aureus strains

The variables (n = 30) defined during adhesion and biofilm formation assays as well

as two additional variables (the type of seafoods from which isolates were sampled and

the type of processing used during their production) were used to perform a Principal

Components Analysis (PCA) for the 28 S. aureus strains. However, the two principal

components (PC) obtained only accounted for 33% of total variance. The selection of

the most significant parameters (n = 8) indicated in the rotated component matrix for

each PC allowed increase up to 79.3% the total variance accounted (Table 2.3). PC1 and

PC2 accounted individually a variance of 55.7% and 23.6%, respectively. PC1 was

positively correlated (P < 0.01) with biofilm formation in TSB with 5% glucose at 25ºC

and in TSB added with or without 5% glucose, 5% glucose + 5% NaCl or 1 mM MgCl2

at 37ºC. PC1 was also correlated with type of product (r = 0.444, P < 0.05) and

processing (r = 0.625, P < 0.01). Meanwhile, PC2 was positively correlated (P < 0.01)

with biofilm formation in TSB with 5% NaCl or 5% glucose + 5% NaCl at 25ºC.

Table 2.3. Component score coefficients matrix obtained from the PCA for the eight relevant

parameters selected, which account for 79.3% of the total variance.

Indicator Condition PC 1 PC 2

TSB 37ºC 24h 0.843 ‒0.254

TSB + 5% glucose 25ºC 48h 0.754 0.122

TSB + 5% glucose 37ºC 24h 0.921 ‒0.055

TSB + 5% glucose 37ºC 48h 0.928 ‒0.003

TSB + 5% NaCl 25ºC 48h 0.030 0.938

TSB + 5% glucose + 5% NaCl 25ºC 48h ‒0.072 0.949

TSB + 5% glucose + 5% NaCl 37ºC 48h 0.885 ‒0.127

TSB + 1 mM MgCl2 37ºC 24h 0.825 0.108


113

S. aureus isolates were located in a scatter plot based on the results from both PC

obtained (Figure 2.4). They were distributed in four groups, each one corresponding to a

defined quadrant. Considerable variations in the ability to develop biofilms on

polystyrene were showed by the isolates under environmental conditions selected.

Figure 2.4. PCA score plot of the S. aureus strains (n = 28) for first two components. PC1:

impact on biofilm formation of glucose at 25ºC and glucose, glucose + NaCl and MgCl2 at

37ºC. PC2: impact of NaCl and glucose + NaCl on biofilm formation at 25ºC. First quadrant

delimited by a solid line; second quadrant delimited by dots; third quadrant delimited by broken

lines; fourth quadrant delimited by dots inserted between broken lines.

Five isolates were distributed in the first quadrant (delimited by a solid line), which

showed a biofilm formation ability significantly influenced by the addition of 5% NaCl

alone or together with 5% glucose at 25ºC. Both the biofilm former reference strain

ATCC 6538 as well as the two isolates carrying sea, sec and seh genes (St.1.07 and

St.1.24) tested in this study were located in this quadrant. However, the five strains had

a different origin: ATCC 6538 were isolated from a human lesion, St.1.07 and St.1.28

from fresh fish and St.1.20 and St.1.24 from precooked products.

Chapter 2

114

The second quadrant (delimited by dots) included six isolates which biofilm

development on polystyrene was highly influenced by the environmental conditions

selected, highlighting St.1.04 and St.1.12. However, they were isolated from seafood

with a different processing: St.1.01, St.1.06 and St.1.12 were isolated from smoked fish,

St.1.02 and St.1.05 from precooked products and St.1.04 from a salted product.

The third quadrant (delimited by broken lines) clustered the highest number of

strains (n = 10), including the antibiotic resistant strain ATCC 43300 and the two strains

carriers of seg and sei genes (St.1.16 and St.1.19) of this study. Biofilm formation of

these isolates was not significantly affected by the environmental conditions selected.

Given that most of them were isolated from frozen (5) and fresh (2) products, other

conditions such as cold temperatures could be the environmental limiting factor during

biofilm formation of these isolates. Moreover, two strains (St.1.13 and St.1.15) of this

group were isolated from shellfish growth by aquaculture, where the application of

antibiotics is widely used.

Finally, the fourth quadrant (delimited by dots inserted between broken lines)

grouped seven strains which biofilm formation was mainly influenced by the addition to

TSB of 5% glucose at 37ºC. They were isolated from precooked (St.1.10, St.1.11 and

St.1.14), smoked (St.1.22) and salted (St.1.23) products, and two from products made

with squids (St.1.09 and St.1.30).

From these results, S. aureus St.1.07, St.1.14 and St.1.29 strains were selected for

their characteristic biofilm-forming ability under food-related environmental stresses

tested to investigate the expression of different genes involved in biofilm formation.

2.3.5 Gene expression in relation to biofilm formation

The genes icaA, rbf, sarA and σB

are reported to be involved in the regulation of

biofilm formation. Their statistical significant (P < 0.05) changes in expression were

investigated under different biofilm promoting growth conditions (TSB with 5%

glucose, 5% NaCl or 5% glucose + 5% NaCl) and compared with expression in TSB by

reverse transcriptase real-time PCR for the three selected strains. All the genes were

highly expressed in TSB (CT ≤ 30), with significant (P < 0.05) differences between the

strains. Thus, St.1.14 showed the highest expression of icaA (CT = 26.9) and rbf (CT =


115

28.4) genes, whereas St.1.07 expressed remarkably genes sarA (CT = 22.7) and σB (CT =

24.6).

Each strain showed a different expression pattern of the analysed genes under the

different growth conditions tested (Figure 2.5). The most variable expression was

observed in icaA gene. An additive effect on icaA expression was seen in St.1.07 when

both NaCl and glucose were added, whereas icaA expression in St.1.29 was down-

regulated in high NaCl conditions (without glucose additions) and up-regulated by the

presence of glucose in the medium. In contrast, icaA expression in St.1.14 was highly

affected by the presence of NaCl, while an up-regulation was observed upon glucose

addition.

Figure 2.5. ΔCt for the expression of genes icaA, rbf, sarA and σB in three S. aureus strains

(St.1.07, St.1.14 and St.1.29) under different conditions compared to expression in TSB.

Different letters on the top of each column show significant differences (P < 0.05) in the

expression of these genes at each condition tested between the selected strains.

Chapter 2

116

Otherwise, the genes rbf, sarA and σB were also highly expressed by the three strains

selected. In St.1.07, the expression of these genes was up-regulated by NaCl with a

dominant down-regulating effect of glucose. For strain St.1.14, σB

expression was

increased when glucose, NaCl or both were added, whereas the presence of glucose in

the medium had a dominant effect in the expression of the other genes, leading to up-

regulation of rbf and down-regulation of sarA. Finally, an additive effect on rbf

expression was seen in St.1.29 when both NaCl and glucose were added, whereas

expression of sarA and σB was up-regulated by glucose addition.

2.4 Discussion

The present study showed considerable variations between the adhesion and biofilm

formation properties of 26 natural S. aureus isolates from seafoods on polystyrene

surfaces under different food-related environmental stress conditions. This surface is

frequently used in the food industry, above all in the packaging of products, and its

bacterial colonization may cause food-spoilage, consequently increasing risk for the

consumer health as well as subsequent economic losses due to recalls of contaminated

food products.

Bacterial adhesion to surfaces is directly correlated with cell surface hydrophobicity

(Rosenberg, 1981; Pagedar et al., 2010). According to our results, all S. aureus strains

expressed moderate hydrophobicity, suggesting a lower initial adhesion to hydrophobic

polystyrene compared to hydrophilic surfaces such as glass. Mafu et al. (2011) also

reported a moderate hydrophobicity and a low tendency to attach to polystyrene in S.

aureus, but a single strain was used.

The electrostatic interactions between the tested S. aureus strains and polystyrene

surface showed a significantly (P < 0.01) higher adhesion when the ionic strength

conditions were increased from 1.5 mM NaCl to 150 mM NaCl, except for strains

St.1.08 and St.1.21. As previously reported (Habimana et al., 2007; Giaouris et al.,

2009), adhesion at high ionic conditions was probably caused by the attenuation of

repulsive electrostatic interactions between the highly negatively charged bacteria and

the negatively charged polystyrene surface. The initial adhesion of S. aureus to

polystyrene could therefore be enhanced in situations involving the use of seawater

during seafood-processing, consequently increasing the risk of biofilm formation and


117

cross-contamination. Therefore, the use of fresh water as a mean to reduce the

attachment of negatively charged bacteria to polystyrene should be considered.

Moreover, obtained results showed that adhesion was dependent on both tested strain

and ionic strength conditions. A high variability in adhesion to polystyrene among S.

aureus strains was observed for both ionic conditions, hence suggesting possible

differences in cell wall electronegativity, as described by Giaouris et al. (2009) in

Lactococcus lactis. To our knowledge, this is the first time that such variability of

surface physicochemical properties is described for natural S. aureus strains from

fisheries. Therefore, these findings provide important information for the development

of novel surfaces and control strategies against the adhesion of natural S. aureus during

processing, packaging and storage of food products, especially in fisheries.

Principal Components Analysis also showed a considerable variability in biofilm

formation between the 26 S. aureus strains tested under relevant environmental

conditions of temperature, osmolarity and nutrient content found during seafood

production. Thus, isolates had generally higher biofilm production at 37ºC as expected,

although four strains (St.1.14, St.1.16, St.1.24 and St.1.31) showed a significantly

higher biofilm development during the first 48 h at 25ºC. Pagedar et al. (2010) also

reported a higher cell count of S. aureus growth in TSB at 25ºC than at 37ºC after 48 h,

but these biofilms were formed on stainless steel surfaces.

Meanwhile, the presence of glucose increased biofilm formation of all tested S.

aureus, although significant differences between isolates were observed. This nutrient is

considered a limiting factor of biofilm formation due to its requirement during the

production of the extracellular matrix components (Ammendolia et al., 1999).

Therefore, our results are in totally agreement with those obtained by Rode et al. (2007),

considering that the presence of glucose promotes biofilm formation in S. aureus. In

fisheries, glucose is an additive frequently used to reduce the water activity of products,

above all in surimis and smoked fish. Data obtained in this study showed that the

presence of glucose significantly influenced biofilm formation of most isolates (70%)

from surimis and smoked fish, as shown their distribution in the PCA score plot. Thus,

the presence of glucose in these products could potentially increase the contamination

by S. aureus, involving a serious risk for the health of consumers and probable

economic losses.

Chapter 2

118

Another important environmental factor is the amount of NaCl present on food-

processing surfaces, which could be increased by the presence of seawater and seafood

wastes generated during seafood production. Different authors showed that NaCl could

promote bacterial aggregation and enhanced the stability of biofilms in polystyrene

(Møretrø et al., 2003; Rode et al., 2007). However, the addition of NaCl generally

decreased the biofilm formation of tested S. aureus strains at 37ºC, whereas it was

improved at 25ºC. Xu et al. (2010) also reported that the number of adhered cells of S.

aureus ATCC 12600 in polystyrene was higher in a medium without NaCl for the first

48 h at 37ºC. A possibility proposed by Lim et al. (2004) could be the repression of

biofilm formation either directly or through overexpression of rbf gen with

concentrations of 5% NaCl approximately. However, a rather average expression of rbf

gen was observed in this study during transcriptional analysis by qRT-PCR of S. aureus

St.1.07, St.1.14 and St.1.29 ‒isolates selected by their characteristic biofilm-forming

properties for PCA‒ when they were growth in TSB added with 5% NaCl. Rachid et al.

(2000) described an osmotic stress resistance and biofilm formation induced by σB, but a

lower expression of σB was reported in S. aureus St.1.29, which had a remarkable

biofilm formation in the presence of 5% NaCl compared to those grown in the absence

of salt. Similarly, the presence of 5% NaCl also caused the down-regulation of the

biofilm-forming promoter sarA in St.1.29, whereas a higher expression was detected in

the other two strains tested under these osmolarity conditions. Therefore, these results

indicate a great variability of regulatory responses against osmolarity stress conditions

during the development of staphylococcal biofilms. Further investigations (e.g. using

knock-out mutants) should be done in the future to deepen this study. Results of such

studies could lead to new biofilm control strategies on food-contact surfaces.

Several authors also indicated the influence of MgCl2 in the adhesion to food-contact

surfaces of Staphylococcus spp. (Barnes et al., 1999; Dunne, 2002; Akpolat et al., 2003;

Planchon et al., 2006). In fisheries, both seawater and seafood wastes are an important

source of magnesium. However, biofilm formation of S. aureus isolates tested in this

study generally was not affected by the presence of MgCl2, although rather favoured

after 48 h at 37ºC. These results are in accordance as those previously reported,

suggesting that MgCl2 are implicated in biofilm stabilization at optimal growth

conditions.


119

The results obtained in this study hence supported that environmental conditions

found in the food industry affected the adhesion and biofilm formation in S. aureus.

Different regulatory pathways are involved in biofilm development of S. aureus

highlighting the ica operon, which is associated in the regulation of extracellular matrix

synthesis (Cramton et al., 1999). Several authors reported that the addition of glucose,

NaCl or both together promote biofilm formation by inducing the ica operon in S.

aureus (Møretrø et al., 2003; Rode et al., 2007). In this study, all the tested strains

carried icaA and icaD (see Appendix 1). Moreover, an increase in icaA expression with

the addition of glucose was also observed during transcriptional analysis by qRT-PCR

of the selected S. aureus isolates St.1.07, St.1.14 and St.1.29. However, although icaA

expression remained high, biofilm formation was lowered when both glucose and NaCl

were added, suggesting that other ica-independent pathways are implicated as proposed

previously different authors (Fitzpatrick et al., 2005; Kogan et al., 2006). Other internal

factors supposedly involved in the initial adhesion to surfaces and host molecules and in

the intercellular adhesion are the biofilm-associated proteins or Bap (Cucarella et al.,

2002). However, none of the natural S. aureus isolates from seafoods carried bap gen

(see Appendix 2). These results are in accordance with Vautor et al. (2008), which

concluded that the prevalence of this gene among S. aureus isolates should be very low.

In fact, the bap gene has only been identified in a small proportion of S. aureus strains

originating from bovine mastitis (Cucarella et al., 2001).

2.5 Conclusions

According to results obtained in the present study, natural S. aureus seems to show a

high ability to adhere and form biofilms on polystyrene surfaces. Food-contact surfaces

made of this material can thus be a hazardous reservoir for S. aureus in the food

industry and, therefore, an important source of food contamination unless appropriate

food safety procedures are applied.

Our results also support that staphylococcal biofilm formation is influenced by

environmental conditions relevant for the food industry such as temperature, osmolarity,

nutrients content and cell surface properties. In fact, considerable variations in biofilm-

forming ability were observed between the different strains tested under these

environmental conditions. Therefore, the prevalence of S. aureus isolates on food-

Chapter 2

120

contact surfaces may be linked to their ability to adapt to the environmental stresses

present during food production.

These findings are relevant for food safety and may be of importance when choosing

the safest environmental conditions and material during processing, packaging and

storage of seafood products. The maintenance of thermal conditions that avoid or reduce

the bacterial growth in food products, the use of low-adherent materials in food-

processing facilities as well as the application of proper cleaning and disinfection

procedures to food-contact surfaces are essentials to ensure food safety.


121

Appendix 1: PCR-detection of icaA and icaD genes

Two genes encoded in ica operon (icaA, icaD) were studied for all S. aureus strains.

Genomic DNA was extracted from 24 h cultures in BHI using an InstaGene™ Matrix

kit (Bio-Rad Laboratories, S.A., Spain) following manufacturer’s instructions. DNA

was quantified by assuming that an absorbance value at 260 nm of 0.100 corresponds to

5 µg/mL of DNA. Primers ICAAF (5´-CCTAACTAACGAAAGGTAG-3´) and ICAAR

(5´-AAGATATAGCGATAAGTGC-3´) for icaA gen, ICADF (5´-AAACGTAAGAGA

GGTGG-3´) and ICADR (5´-GGCAATATGATCAAGATAC-3´) for icaD gen were

used for each strain (Cramton et al., 1999). Expected size of amplified PCR products

were 1315 bp for icaA gen, 381 bp for icaD. Each PCR mixture contained 100 ng DNA,

1x Taq Buffer Advanced, 2.5 U Taq DNA polymerase (5 Prime, Germany), 40 nmol of

each dNTP (Bioline, UK), 0.25 nmol of forward and reverse primers (Thermo Fisher

Scientific, Germany) and sterile Milli-Q water up to a final volume of 50 µL. PCR was

performed with a MyCycler™ Thermocycler (Bio-Rad). PCR conditions consisted of

denaturation at 92ºC for 5 min, 30 cycles at 92ºC for 45 s, 49ºC for 45 s and 72ºC for 1

min, and a final cycle at 72ºC for 7 min (Vasudevan et al., 2003). PCR products were

subjected to electrophoresis on 1.5% agarose gel containing ethidium bromide for 90

min at 75 V and 100 mAmp. Gels were photographed in a Gel Doc XR system (Bio-

Rad) using the Quantity One® software (Bio-Rad). A DNA ladder of 50-2000 bp

(Hyperladder II, Bioline) was included as a molecular size marker.

Figure Ap.1A-B. Agarose gels showing PCR products obtained for all S. aureus strains during

detection of genes icaA (A) and icaD (B). Lane 1 and 17, DNA molecular size marker

(HyperLadder II, 50-2000 bp; Bioline); lanes 2-3, reference strains S. aureus ATCC 29213 and

S. aureus ATCC 43300, respectively; lanes 4-16 and 18-30, amplicons of each strain; lane 31,

blank.

Chapter 2

122

Appendix 2: PCR-detection of bap gene

The presence of gene involved in the production of biofilm-associated proteins (bap)

was investigated for all S. aureus strains. DNA was extracted with DNeasy® kit

(Qiagen, Germany) according to the manufacturer. Extraction was tested by using λ

HindIII DNA Ladder as a reference (564-23130 bp) (New England BioLabs™, USA).

Primers sasp-6m (5´-CCCTATATCGAAGGTGTAGAATTGCAC-3´) and sasp-7c (5´-

GCTGTTGAAGTTAATACTGTACCTGC-3´) were used (Cucarella et al., 2004). The

expected size of amplified PCR products was 971 bp. PCR mixtures were composed of

20 ng of DNA; 5 nmol of each dNTP (Invitrogen Corporation, USA); 2.5 µL of

Dynazym buffer 10x and 1.2 U of Dynazym Hot Start (Bio-Rad); 10 pmol of forward

and reverse primer and sterile Milli-Q water up to a final volume of 25 µL. PCRs were

performed with an 80 Gene Amp PCR System 9700 (Applied Biosystems, USA). PCR

conditions consisted of denaturation at 94ºC for 2 min, 40 cycles of 94ºC for 30 s, 55ºC

for 30 s and 72ºC for 75 s, and a last extension at 72ºC for 5 min (Vautor et al., 2008).

S. aureus ATCC 29213 was used as negative control, whereas S. aureus ATCC 43300

was used as positive control. Amplicons were subjected to electrophoresis on 1.2%

agarose gel containing ethidium bromide for 30 min at 100 V and 200 mAmp. Gels

were visualized and saved in a Typhoon Scanner 8600 (Molecular Dynamics, GE

Healthcare, UK). A DNA ladder of 154-2176 bp (DNA Molecular Weight Marker VI,

Roche Applied Science, USA) was included in all gels.

Figure Ap.2. Agarose gel showing PCR products obtained for all S. aureus strains during

detection of bap gene. Lane 1 and 17, DNA molecular size marker (DNA Molecular Weight

Marker VI, Roche Applied Science); lanes 2-3, reference strains S. aureus ATCC 29213 and

ATCC 43300; lanes 4-16 and 18-30, amplicons of each strain; lane 31, blank; lane 32, bap-

positive strain S. aureus V329 (Cucarella et al., 2001).

125

Chapter 3. Biofilm-forming ability and

resistance to industrial disinfectants of

Staphylococcus aureus isolated from fishery

products

Biofilm-forming ability and resistance to industrial disinfectants of S. aureus

127

* Corresponding author at: Instituto de Investigaciones Marinas (CSIC), Eduardo Cabello 6, 36208, Vigo Spain. Tel.: +34 986 231 930; fax: +34 986 292 762.

E-mail address: [email protected] (J.J. Rodríguez-Herrera).

Abstract

Adhesion and biofilm-forming ability of twenty six S. aureus strains previously

isolated from fishery products on stainless steel was assessed. All strains reached counts

higher than 104 CFU/cm

2 after 5 h at 25°C. Most strains also showed a biofilm-forming

ability higher than S. aureus ATCC 6538 ‒reference strain in bactericidal standard

tests‒ by crystal violet staining. In addition, it seems that food-processing could have

produced a selective pressure and strains with a high biofilm-forming ability were more

likely found in highly handled and processed products.

The efficacy of the industrial disinfectants benzalkonium chloride (BAC), sodium

hypochlorite (NaClO) and peracetic acid (PAA) against biofilms and planktonic

counterparts was also examined in terms of minimum biofilm eradication concentration

(MBEC) and minimum bactericidal concentration (MBC), respectively. Biofilms

showed an antimicrobial resistance higher than planktonic cells in all cases. However,

no correlation was found between MBEC and MBC, likely due to differences in biofilm

extracellular matrix (composition, content and architecture) between strains. BAC

resistance increased as biofilms aged. Generally, biofilm formation seemed to attenuate

the effect of low temperatures on BAC resistance. PAA was found to be most effective

against both biofilms and planktonic cells, followed by NaClO and BAC. Resistance did

not follow the same order for each biocide, which remarks the need of using a wide

collection of strains in standard tests of bactericidal activity to ensure a proper

application of disinfectants. Doses recommended by manufacturers for BAC, PAA and

Chapter 3

128

NaClO to disinfect food-contact surfaces were lower than doses for complete biofilm

removal (i.e. MBEC) under some environmental conditions common in the food

industry, which questions bactericidal standard tests and promotes the search for new

strategies for biofilm removal.

Keywords: Staphylococcus aureus; biofilm; benzalkonium chloride; peracetic acid;

sodium hypochlorite.

3.1. Introduction

Many disinfectants are used to kill microorganisms in the food industry, being

quaternary ammonium compounds, chlorine-based biocides and peroxides among most

used. However, bacteria form biofilms on any kind of surface, which allows them to

tolerate the presence of biocides much better than free-living counterparts (Bridier et al.,

2011a; Donlan and Costerton, 2002; Van-Houdt and Michiels, 2010). The resistance of

biofilms to biocides can increase the persistence of bacterial pathogens in the food

environment (Srey et al., 2013; Van-Houdt and Michiels, 2010). In fact, an improper

use of biocides (due to short exposure times, sub-lethal doses or unsuitable equipment

designs) can provoke an increase in incidence or even the emergence of antimicrobial-

resistance (Gibson et al., 1999; Langsrud et al., 2003; Sharma and Anand, 2002).

Controversy has therefore arisen over the validity of official standardised tests EN 1040

(CEN, 2005), EN 1276 (CEN, 2009) and EN 13697 (CEN, 2002) applied in the

European Union to determine the bactericidal efficacy of disinfectants, particularly if

they truly simulate conditions found in the food industry as well as if they cover the

spectrum of biological variation that can be found in nature (Briñez et al., 2006;

Langsrud et al., 2003; Meyer et al., 2010).

Staphylococcus aureus is a major foodborne bacterial pathogen causing food

poisoning due to the ingestion of food containing staphylococcal enterotoxins (Bhatia

and Zahoor, 2007; EFSA, 2010; Le-Loir et al., 2003). S. aureus can spread from food

handlers and food-contact surfaces to food products throughout the food chain (DeVita

et al., 2007; Sattar et al., 2001; Simon and Sanjeev, 2007; Sospedra et al., 2012).

Nonetheless, recent changes in Spanish national regulations (RD 135/2010) following

Commission Regulation (EC) No 2073/2005 have revoked the use of S. aureus as a

microbiological criterion for a number of foods, including several fishery products.


129

However, a high incidence of S. aureus (~ 25%) has been recently found in fishery

products marketed in Spain (Vázquez-Sánchez et al., 2012). Being Spain the largest

producer and the second largest consumer of seafood in the European Union (Eurostat,

2010; FAO, 2012), the safety of seafood produced or consumed in Spain is of outmost

importance.

The present study was therefore aimed to examine if the formation of biofilms could

be a potential source of contamination with S. aureus in industries processing fishery

products. With this aim, the biofilm-forming ability of the strains previously isolated

from fishery products and the resistance of such biofilms to three traditional biocides

(benzalkonium chloride, peracetic acid and sodium hypochlorite) was determined.

3.2. Material and Methods

3.2.1. Bacterial strains and culture conditions

Twenty six S. aureus strains isolated from fishery products from different countries

of origin were investigated (Table 3.1). They have been previously identified as S.

aureus by specific biochemical (coagulase, DNAse and mannitol fermentation) and

genetic tests (23s rDNA) (Vázquez-Sánchez et al., 2012). Strains were also formerly

characterized by RAPD-PCR and considered to be putative enterotoxigenic strains

because they carried se genes (Vázquez-Sánchez et al., 2012). S. aureus ATCC 6538,

which is used as a reference gram-positive strain in United States and European

bactericidal standard tests, was provided by the Spanish Type Culture Collection.

Bacterial stocks of each strain were maintained at ‒80°C in tryptic soy broth (TSB)

(Panreac Química, Spain) containing 20% glycerol (v/v). All strains were thawed and

subcultured twice in TSB at 37°C for 24 h under static conditions prior to each

experiment.

3.2.2. Conditions for biofilm formation

Stainless steel coupons (AISI 304, 2B finish) (Markim Galicia S.L., Spain) of

approximately 10 mm × 10 mm (and 0.8 mm thickness) were used as experimental

surfaces. Coupons were soaked in 2 M NaOH to remove residues, rinsed several times

with distilled water, air-dried and autoclaved before use. One sterile coupon was placed

into each well of a sterile 24-well flat-bottom microtiter plate (Falcon®, Becton

Dickinson Labware, USA).

Chapter 3

130

Table 3.1. Origin and putative enterotoxigenicity (as se gene carried) of S. aureus strains

studied.

Strain Fishery product Type of processing Producing country se gene

St.1.01 Smoked swordfish Smoking Spain (Andalucía) a

St.1.02 Shellfish Paella Precooking Spain (Valencia) a

St.1.03 Smoked swordfish Smoking Spain (Andalucía) a

St.1.04 Mojama (salted dry tuna) Salting Spain (Valencia) a

St.1.05 Herring in Madeira sauce Precooking Denmark a

St.1.06 Smoked angel fish Smoking Spain (Andalucía) a

St.1.07 Fresh cuttlefish Fresh France a, c, h

St.1.08 Surimi elver Precooking Spain (Basque Country) a

St.1.09 Fresh squid Fresh France a

St.1.10 Surimi elver Precooking Portugal a

St.1.11 Elver with shrimps Precooking Denmark a

St.1.12 Smoked tuna Smoking Spain (Andalucía) a

St.1.13 Frozen shellfish paella Freezing Spain (Galicia) a

St.1.14 Surimi elver Precooking Spain (Galicia) a

St.1.15 Frozen shellfish paella Freezing Spain (Galicia) a

St.1.16 Perch fillet Fresh France g, i

St.1.19 Frozen cod Freezing Spain (Galicia) g, i

St.1.20 Elver with shrimps Precooking Portugal a

St.1.21 Frozen Shelled prawn Freezing Spain (Valencia) a

St.1.22 Smoked trout Smoking Spain (Catalonia) a

St.1.23 Salted cod Salting Spain (Galicia) a

St.1.24 Cod pâté with peppers Precooking Spain (Valencia) a, c, h

St.1.28 Panga fillet Fresh Vietnam a

St.1.29 Panga fillet Fresh Vietnam a

St.1.30 Frozen Squid ring Freezing Spain (Galicia) a

St.1.31 Frozen hake nuggets Freezing Spain (Galicia) a

Overnight bacterial cultures were adjusted to an absorbance value at 700 nm of 0.100

± 0.01 with phosphate buffer saline (PBS, composed by 7.6 g/L NaCl, 0.2 g/L KCl and

0.245 g/L Na2HPO4 (BDH Prolabo, VWR International Eurolab, Spain); and 0.71 g/L

K2HPO4 (Panreac Química, Spain)). This value corresponds to a cell concentration of

approximately 108 CFU/mL for all strains. PBS-suspended cells were then serially

diluted in TSB, and a 700 μL aliquot (containing approximately 7×105 CFU) was added

into each well with a coupon. Inoculum size was checked in all cases by plating on

tryptic soy agar (TSA) (Cultimed, Panreac Química, Spain). A negative control with no

inoculum was included in all assays. Microplates were incubated at 25°C under static

conditions until analysis.


131

3.2.3. Biofilm formation assays

3.2.3.1. Slime production on Congo red agar (CRA)

A slight modification of the method proposed by Freeman et al. (1989) was used. CRA

was composed of 37 g/L brain heart infusion broth (Biolife, Italy), 36 g/L sucrose

(Probus, Spain), 10 g/L agar-agar (Scharlau, Spain) and 0.8 g/L Congo red stain

(Panreac Química, Spain). Congo red solution was autoclaved separately and then

added to sterile agar medium at 55°C. A 0.1 mL inoculum of each S. aureus strain was

spread on CRA (by duplicate), incubated at 37°C for 72 h. Two plates were visually

inspected for slime production each 24 h. The assays were repeated twice using

independent bacterial cultures.

3.2.3.2. Initial adherence

After 5 h of incubation at 25°C, coupons were removed from microplates and

washed with 1 mL of PBS for 10 s to eliminate non-adhered cells. Adhered cells were

collected by thoroughly rubbing with two sterile swabs (Deltalab, Spain). Cells were

resuspended by vortexing swabs vigorously for 1 min in 9 mL of peptone water (10 g/L

triptone (Cultimed, Panreac Química, Spain) and 5 g/L sodium chloride (BDH Prolabo,

VWR International Eurolab, Spain)). Ten-fold serial dilutions of resuspended cells were

made in peptone water and aliquots of 0.1 mL of appropriate dilutions were spread on

TSA plates. Plates were incubated at 37°C for 24 h. Four coupons were analysed for

each strain and each assay was repeated twice using independent bacterial cultures.

3.2.3.3. Quantification of biofilm formation

Biofilm formation was quantified in terms of biomass after 5, 24 and 48 h of

incubation at 25°C. At each sampling time, coupons were placed into a new microplate

and washed with 1 mL of PBS for 10 s to remove non-adhered cells. Coupons were then

air-dried and biofilms were fixed in Bouin´s solution (Sigma-Aldrich Química, Spain)

for 45 min. Subsequently, coupons were further washed with 1 mL of PBS for 10 s to

discard excess fixative. Once fixed, biofilms were stained with 0.5 mL of 0.5% (w/v)

crystal violet solution (Panreac Química, Spain) for 15 min. Excess stain was rinsed off

by placing microplates under running tap water. Coupons were air-dried and biofilm-

bound crystal violet was solubilized in 1 mL of 33% (v/v) glacial acetic acid (BDH

Chapter 3

132

Prolabo, VWR International Eurolab, Spain) by shaking at 100 rpm for 30 min. Crystal

violet acid solution was then transferred to a 96-well round-bottom microplate (Falcon®,

Becton Dickinson Labware, USA) and absorbance was read out at 595 nm using a

iMark™ Microplate Absorbance Reader equipped with Microplate Manager software

6.0 (Bio-Rad Laboratories, Spain). Coupons without biofilms were used in each assay

as a negative control to subtract background staining. Six coupons were analysed for

each strain at each sampling time and all assays were repeated twice using independent

bacterial cultures.

3.2.3.4. Determination of growth kinetics

A set of test tubes containing 5 ml TSB each was inoculated with a 100 μL aliquot

(containing approximately 105 CFU/mL) from bacterial cultures prepared as

aforementioned. Cultures were incubated at 12°C and 25°C for 14 and 8 days,

respectively, under static conditions. Absorbance was read out at 700 nm after each 24 h

of incubation. Three replicates were used each day for each strain. A negative control

without inoculum was included in all assays. Growth kinetics was determined using two

independent bacterial cultures. Once obtained, experimental data of growth were fitted

to the Gompertz equation proposed by Zwietering et al. (1990):

(

) { [(

) ]}

where ODt is the optical density at 700 nm at time t (days), OD0 is the optical density at

the time of inoculation, A is a dimensionless asymptotic value, µmax is the maximum

specific growth rate (1/days) and λ is the detection time (days), i.e., the time at which

OD at 700 nm is firstly noted to increase.

3.2.4. Biocide resistance assays

Three traditional chemical disinfectants were tested: benzalkonium chloride (50%

(v/v) BAC solution, Guinama Absoluta Calidad, Spain), peracetic acid (40% (v/v) PAA

solution in acetic acid:water, Fluka, Sigma-Aldrich Química, Spain) and sodium

hypochlorite (NaClO, Scharlau, Spain). BAC was studied in a greater detail than PAA

and NaClO as BAC is more widely used for disinfection of stainless steel surfaces in the

food industry. Disinfectants were diluted in ultrapure water to working concentrations


133

just before each assay. Each concentration was evaluated in triplicate in two

independent experiments.

3.2.4.1. Minimal biofilm eradication concentration (MBEC)

The resistance of biofilms was assessed in terms of the minimal biofilm eradication

concentration (MBEC), which was defined as the lowest disinfectant concentration that

kills all biofilm cells under the experimental conditions tested. Once non-adhered cells

were removed, three coupons were exposed to 1.5 mL of each disinfectant concentration

for 30 min. Thirteen different concentrations were tested for BAC (5000, 7500, 10000,

12000, 14000, 16000, 18000, 20000, 22000, 24000, 26000, 28000 and 30000 mg/L),

twelve for PAA (1000, 1500, 2000, 2500, 3000, 3500, 4000, 5000, 7000, 9000, 11000

and 13000 mg/L) and eleven for NaClO (5000, 6000, 7000, 8000, 9000, 10000, 12000,

14000, 16000, 18000, 20000 mg/L). Afterwards, coupons were placed into sterile glass

vials and 9 mL of neutralizing broth (0.34 g/L KH2PO4 and 5 g/L Na2S2O3 (Probus,

Spain); and 3 g/L soy lecithin, 1 g/L L-histidine and 3% (v/v) polysorbate 80 (Fagron

Iberica, Spain)) was added and left to stand for 10 min at room temperature, as proposed

by Luppens et al. (2002). Finally, each coupon was incubated in TSB at 37°C for 24 h

and bacterial growth was monitored visually. In addition, visually undetectable growth

was checked by plating an aliquot of 0.1 mL of culture medium on TSA and searching

for the presence of colonies after 24 h at 37°C. In any case, bacterial growth indicated

the presence of viable cells in disinfectant-treated biofilms.

3.2.4.2. Minimal bactericidal concentration (MBC)

The resistance of planktonic cells was assessed in terms of the minimum bactericidal

concentration (MBC), which was considered to be the lowest disinfectant concentration

necessary to kill all free-living bacterial cells under the experimental conditions tested.

Planktonic counterparts (i.e. non-adhered cells) were diluted in TSB to achieve a similar

cell concentration to that in biofilms. Planktonic cells (0.1 mL) were exposed to each

disinfectant concentration (0.1 mL) for 30 min (by triplicate) and then immediately

neutralized (2.0 mL). Eleven different concentrations were tested for BAC (750, 1000,

1500, 2000, 2500, 3000, 4000, 5000, 6000, 7000 and 8000 mg/L), PAA (100, 150, 200,

250, 300, 350, 400, 450, 500, 600, 750 mg/L) and NaClO (500, 550, 600, 650, 700, 750,

800, 850, 900, 950 and 1000 mg/L). An aliquot of 0.3 mL of each neutralized

Chapter 3

134

disinfectant-treated bacterial culture was added to 1.7 mL of TSB and incubated at 37°C

for 24 h. Bacterial growth was monitored as for MBEC.

3.2.5. Statistical analysis

Experimental results were statistically analysed with the software packages

Microsoft Excel 2010 and IBM SPSS 19.0. Statistical significance analysis was carried

out using a one-way ANOVA. Homogeneity of variances was examined by a post-hoc

least significant difference (LSD) test. Otherwise, a Dunnett´s T3 test was performed.

An independent-samples Student´s t-test was also done to determine if there were

statistical differences between pairs of strains. A bivariate correlation analysis was

conducted using Pearson’s correlation coefficient to measure the strength of the linear

relationship between two variables. Statistical significance was accepted at a confidence

level greater than 95% (P < 0.05). Occasionally, a level greater than 99% (P < 0.01) is

considered to remark differences between variables. The variability among strains was

calculated by means of the coefficient of variation (CV), which is defined as the ratio of

the standard deviation to the mean.

3.3. Results

3.3.1. Biofilm-forming ability of S. aureus

The biofilm-forming ability of a number of S. aureus strains isolated from food

products was compared with that of S. aureus ATCC 6538. This strain is used as a

reference in several antimicrobial tests, including the quantitative surface test (EN

13697) of bactericidal activity of disinfectants used in food, industrial, domestic and

institutional areas, on the basis that it is a biofilm former.

3.3.1.1. Detection of biofilm-forming ability on Congo red agar (CRA)

Biofilm-forming ability of each strain was also visually investigated by the CRA

plate test following the colorimetric scale proposed by Arciola et al. (2002). Colonies

coloured from very black to almost black have been described as typical of a biofilm-

positive phenotype, whereas negative slime producers form colonies coloured from very

red to burgundy. All strains formed black colonies after 24 h, with colour changing


135

from red to black in the surrounding agar too. Thus, all strains showed slime production

ability after 24 h. All of them maintained this biofilm-positive phenotype during the

following 48 h. No strain formed red colonies onto CRA.

3.3.1.2. Initial adhesion studies

The adhesion of strains to stainless steel was determined after 5 h at 25°C. The

number of cells adhered was higher than 104 CFU/cm

2 for all strains. A small variability

was found among strains (CV of 4.0%), but significant differences (P < 0.05) were

observed between strains (Figure 3.1). S. aureus St.1.01, St.1.09 and St.1.30 showed the

highest adhesion ability (6-7×104 CFU/cm

2), which was significantly higher than that of

S. aureus ATCC 6538. On the contrary, S. aureus St.1.03, St.1.06 and St.1.13 showed

the lowest number of adhered cells (< 2×104 CFU/cm

2).

Figure 3.1. Adherence of S. aureus strains to stainless steel. Adherence was assessed after 5 h

at 25°C. Statistically significant differences (P < 0.05) are indicated by different letters.

3.3.1.3. Quantification of biofilm biomass

Biofilm formation was quantified in terms of biomass by using the crystal violet

staining method. Biofilm biomass of all strains increased continuously (and

significantly, P < 0.01) during incubation for 48 h at 25°C, except for the case of St.1.28

between 24 and 48 h (Figure 3.2A-C). The increase was higher for most of the strains

under study than for S. aureus ATCC 6538. Thus, only three strains (St.1.09, St.1.30

and St.1.31) showed a biofilm-forming ability significantly higher (P < 0.05) than S.

aureus ATCC 6538 after 5 h, but the majority showed a biofilm biomass significantly

higher (P < 0.05) after 24 h (76.9%) and 48 h (69.2%). A high inter-strain variability in

Chapter 3

136

biofilm production ‒involving statistically significant (P < 0.05) differences‒ was found

at each time of study (CV of 36.4%, 30% and 29.7% after 5, 24 and 48 h, respectively).

A positive correlation (r = 0.890, P < 0.01) between the number of adhered cells and

biofilm density was also found after 5 h at 25°C.

Interestingly, 48-h-old biofilm biomass was found to be positively correlated (r =

0.401, P < 0.01) with the type of processing applied to products that strains were

isolated from (Table 3.1). In fact, eight strains out of the ten with highest biofilm

biomass were found in highly-processed products (including smoked products, salted

fish and precooked products), whereas seven strains out of the ten with lowest biofilm-

forming ability were isolated from low-processed products (i.e., fresh and some frozen

products).

Figure 3.2A-C. Biofilm-forming ability of S. aureus on stainless steel. Biofilm-forming ability

was measured after 5 h (A), 24 h (B) and 48 h (C) at 25°C in terms of biofilm biomass by

crystal violet staining. Different letters denote statistically significant differences (P < 0.05).


137

3.3.2. Resistance to industrial disinfectants of S. aureus

3.3.2.1. Effectiveness of benzalkonium chloride

3.3.2.1.1. Variability in BAC resistance

The resistance to benzalkonium chloride (BAC) was firstly assessed in 48-h-old

biofilms and planktonic counterpart cells. As shown in Figure 3.3, the resistance of both

biofilms and planktonic cells showed a high variability between strains (CVs of 27.1%

and 34.7%, respectively). Thus, whereas nearly half of strains (44.4%) formed biofilms

that only resisted up to 14000 mg/L (included ATCC 6538), seven others showed a

much higher resistance (MBEC = 20000-26000 mg/L). In the case of planktonic cells,

most strains (65.4%) resisted up to 2000-3000 mg/L, but St.1.09, St.1.11, St.1.28 and

St.1.30 achieved MBC values of 4000 mg/L, 4-fold higher than that of ATCC 6538

(MBC = 1000 mg/L).

Figure 3.3. Effectiveness of BAC against S. aureus biofilms and planktonic cells after 48 h at

25°C. MBEC: Minimal Biofilm Eradication Concentration; MBC: Minimal Bactericidal

Concentration.

As expected, biofilms formed by S. aureus on stainless steel showed a higher

resistance to BAC than their planktonic counterpart cells, but this increase was highly

strain-dependent. Thus, S. aureus St.1.12 biofilms showed a resistance 16-fold higher

than planktonic cells, whereas St.1.11 biofilms, which had shown a high biofilm-

forming ability, were only about 3-fold more resistant than planktonic counterparts. As

an overall result, no correlation was found between MBEC and MBC, neither between

MBEC and 48 h biofilm biomass.

Chapter 3

138

From these results, those strains highly-BAC resistant (St.1.01, St.1.03, St.1.07,

St.1.14, St.1.23, St.1.24, St.1.28, St.1.30 and St.1.31) were chosen for the subsequent

studies. S. aureus St.1.04, St.1.08 and St.1.10 were also selected for these studies on the

basis of a higher incidence in fishery products (Vázquez-Sánchez et al., 2012) and a

higher risk to food safety associated to the type of product that they were isolated from

(Table 3.1).

3.3.2.1.2. Effects of growth kinetics on BAC resistance

Foreseeing a different response during the time course of biofilm formation, the

resistance to BAC of biofilms formed by the strains selected was determined after 5, 24,

48 and 168 h at 25°C. The resistance of planktonic counterparts after 24, 48 and 168 h

was also determined for the interest in studying post-exponential and stationary-phase

cells able to express antimicrobial resistance responses.

The development of biofilms induced a progressive increase in BAC resistance

(Figure 3.4A), particularly during the first 24 h, when most of strains (83.3%) showed

MBEC values at least 2-fold higher than after 5 h. MBEC was thus only found to be

positively correlated with biofilm biomass after 24 h (r = 0.558, P < 0.01), but no after 5

or even 48 h. In fact, the increase slowed down subsequently, with most strains (75%)

displaying only slight increases (2000 mg/L) or remaining similar otherwise.

Nevertheless, 168-h-old biofilms of the majority of the strains (83.3%) were able to

resist up to 20000-28000 mg/L, with St.1.01 forming the most resistant biofilms.

Figure 3.4A-B. Effectiveness of BAC against biofilms (A) and planktonic counterparts (B)

after 5, 24, 48 and 168 h at 25°C.


139

The resistance of planktonic cells also increased progressively as bacterial cultures

aged, particularly during the first 48 h (Figure 3.4B). However, likewise in biofilms,

increases slowed down or simply did not occur in most strains (75%) between 48-168 h.

Nevertheless, 168-h-old planktonic cells of five strains were able to resist up to 4000-

5000 mg/L.

These results have shown that the formation of biofilms decreased markedly the

efficacy of BAC from the first stages of biofilm development. In fact, the resistance of

5-h-old-biofilms was higher than that of 168-h-old planktonic cells in all cases.

However, MBEC and MBC were only found to be positively correlated after 168 h of

incubation (r = 0.603, P < 0.01). Interestingly, a positive correlation was found between

initial adhesion and MBEC after 24 h (r = 0.637, P < 0.01), 48 h (r = 0.683, P < 0.01)

and 168 h (r = 0.652, P < 0.01).

3.3.2.1.3. Effects of growth temperature on BAC resistance

The effects of the growth temperature on the BAC resistance of biofilms were

analysed at 12°C and 25°C after 168 h of incubation. The use of 168-h-old biofilms is

accounted for the interest in studying highly mature biofilms as examples of worst-case

scenarios as well as for the slow development of biofilms at 12°C.

As shown in Figure 3.5A, the temperature affected the resistance of biofilms, with

lower MBEC values at 12°C (10000-24000 mg/L) than at 25°C (12000-28000 mg/L).

These differences can be at least partially accounted for the growth kinetics of these

strains (see Appendix), particularly in terms of detection times (λ) much longer and

maximum specific growth rates (µmax) much lower at 12°C than at 25°C (Table 3.2). For

instance, λ at 12°C was positively correlated (r = 0.711, P < 0.01) with the decrease in

resistance of biofilms. That is, the longer the λ, the higher the decrease in resistance

between temperatures. Besides, differences in these parameters between strains were

much higher at 12°C than at 25°C. Also, MBEC and cell density were found to be

positively correlated at 12°C (r = 0.604, P < 0.01), but not at 25°C. Nevertheless, a

positive correlation (r = 0.755, P < 0.01) was found between the resistance of biofilms

at both temperatures.

Chapter 3

140

Figure 3.5A-B. Effectiveness of BAC against 168-h-old biofilms (A) and planktonic

counterparts (B) grown at 12°C and 25°C.

Table 3.2. Detection time (λ) and maximum specific growth rate (µmax) of S. aureus strains

grown at 12ºC and 25ºC. Letters denote statistically significant differences (P < 0.05).

Strain λ (days) µmax (1/days)

12ºC 25ºC 12ºC 25ºC

St.1.01 2.09 ± 0.21c

0.00 1.25 ± 0.05cd

3.52 ± 0.05a

St.1.03 8.15 ± 0.08a

0.00

1.44 ± 0.10b

2.81 ± 0.06bc

St.1.04 0.47 ± 0.12f

0.00

0.77 ± 0.01g

2.58 ± 0.10de

St.1.07 0.24 ± 0.06g

0.00

0.67 ± 0.12g

2.50 ± 0.16de

St.1.08 1.77 ± 0.06c

0.00

0.99 ± 0.06ef

3.07 ± 0.19b

St.1.10 1.83 ± 0.20c

0.00 1.59 ± 0.02a

3.89 ± 0.38a

St.1.14 1.42 ± 0.18d

0.00

1.11 ± 0.08de

2.78 ± 0.05c

St.1.23 1.36 ± 0.03d

0.00

0.91 ± 0.07f

2.39 ± 0.16e

St.1.24 0.00h

0.00 0.97 ± 0.09ef

2.39 ± 0.17e

St.1.28 0.88 ± 0.05e

0.00

1.19 ± 0.06d

3.51 ± 0.15a

St.1.30 6.47 ± 0.51b

0.00

1.58 ± 0.30abc

2.73 ± 0.13bcd

St.1.31 0.71 ± 0.17ef

0.00

0.91 ± 0.12f

2.19 ± 0.29e


141

The effect of the temperature was particularly noticeable in S. aureus St.1.03, St.1.07

and St.1.30, with decreases in MBEC of 8000-10000 mg/L from 25°C to 12°C. This

seems to be a consequence of the slow adaptation to low temperatures of these strains

(St.1.03 and St. 1.30 showed the longest λ and St.1.07 the lowest µmax in planktonic

state of all strains at 12°C). In contrast, St.1.01 and St.1.14 biofilms seemed to be

weakly affected by low temperatures as they also showed a high resistance at 12°C.

This could be partially accounted for the high µmax shown by these strains in the

planktonic state. Surprisingly, St.1.24 showed a MBEC slightly lower at 25°C than at

12°C, which was presumably due in part to the absence of λ at 12°C.

The temperature also influenced the resistance of planktonic cells to BAC (Figure

3.5B), which showed a lower MBC at 12°C (1000-3000 mg/L) than at 25°C (2000-5000

mg/L). MBC was found to be positively correlated with cell density at 12°C (r = 0.670,

P < 0.01), but not at 25°C. MBEC and MBC were also found to be positively correlated

at 12°C (r = 0.736, P < 0.01). Unlike biofilms, however, the resistance of planktonic

cells at 12°C and 25°C were not found to be correlated. The variability of BAC

resistance was higher in planktonic cells than in biofilms, particularly at 12°C (CV of

38.5% and 26.5%, respectively).

3.3.2.2. Effectiveness of peracetic acid and sodium hypochlorite

The antimicrobial efficacy of peracetic acid (PAA) and sodium hypochlorite

(NaClO) against 48-h-old biofilms and planktonic counterparts of the selected strains

grown at 25°C was assessed and compared with that of BAC. The type strain ATCC

6538 was also included in this study.

Likewise for BAC, biofilms showed a much higher resistance than planktonic cells to

PAA (up to 11-fold for St.1.01 and St.1.14) and NaClO (up to 23-fold for St.1.24) in all

cases. However, strains were ranked in a different order according to the resistance of

biofilms ‒and planktonic cells‒ to each disinfectant (Table 3.3). As a result, no

correlation was found between the resistance of biofilms or planktonic cells to different

disinfectants. No correlation was found either between MBEC and MBC or between

MBEC for PAA, NaClO or BAC and biofilm biomass after 48 h of incubation at 25°C.

Chapter 3

142

PAA was found to be the most effective disinfectant against 48-h-old biofilms in all

cases. In fact, PAA was up to 11-fold (St.1.31) and 9-fold (St.1.24) more effective

against biofilms than BAC and NaClO, respectively. The efficacy of NaClO against

biofilms was over 2-fold higher than BAC in many cases (50%) and similar in others

(St.1.04 and St.1.10), whereas BAC was more effective than NaClO in one case

(St.1.08).

PAA also showed the highest effectiveness against planktonic counterparts, up to 11-

fold higher than BAC (St.1.28 and St.1.30) and 2-fold higher than NaClO in all cases.

Meanwhile, NaClO was more effective than BAC in all cases, being up to 4-fold higher

(St.1.03, St.1.28 and St.1.30).

Lastly, the resistance of biofilms to PAA showed higher variability (coefficient of

variation of 33.4%) than to BAC (25.8%) and NaClO (14.4%). However, the resistance

of planktonic cells showed the highest variability in the case of BAC (21.6%) followed

by PAA (10.0%) and NaClO (6.1%).

Table 3.3. Effectiveness of BAC, PAA and NaClO against S. aureus biofilms and planktonic

counterpart cells after 48 h at 25°C. MBEC: Minimal Biofilm Eradication Concentration; MBC:

Minimal Bactericidal Concentration.

Strain MBEC (mg/L) MBC (mg/L)

BAC PAA NaClO BAC PAA NaClO

ATCC 6538 10000 1500 5000 1000 300 600

St.1.01 26000 4000 14000 2500 350 850

St.1.03 18000 3000 12000 3000 350 800

St.1.04 12000 4000 12000 2500 400 800

St.1.07 22000 4000 16000 2500 350 950

St.1.08 10000 2000 12000 2500 350 900

St.1.10 14000 1500 14000 2500 300 800

St.1.14 26000 4000 12000 2500 350 850

St.1.23 20000 4000 16000 2000 400 800

St.1.24 20000 2000 18000 2500 450 800

St.1.28 18000 2000 12000 4000 350 900

St.1.30 24000 2500 14000 4000 350 900

St.1.31 22000 2000 12000 2500 350 900


143

3.4. Discussion

The results obtained in the present study have shown that a number of S. aureus

strains (n = 26) isolated from fishery products have, in general, a high biofilm-forming

ability on stainless steel surfaces and a high resistance to some disinfectants commonly

used in the food industry. Accordingly, the entrance of these strains in food-processing

facilities does not only involve an immediate risk to food safety but more importantly

the risk of long-term presence (even persistence) unless appropriate measurements are

applied to kill them.

Biofilm-forming ability was assessed in terms of biomass using the crystal violet

staining method. It could thus be observed that all strains had a high biofilm-forming

ability on stainless steel under experimental conditions simulating situations normally

found in the food industry. Stainless steel, a common material in food-processing

facilities, can thus serve as an important reservoir for S. aureus in the food industry.

Several studies have also detected the presence of S. aureus biofilms on stainless steel

food-processing surfaces (Bagge-Ravn et al., 2003; Sattar et al., 2001; Sospedra et al.,

2012).

Biofilm biomass increased proportionally as biofilms aged. A high variability in

biofilm biomass was found among strains throughout the time course of biofilm

formation, which is in accordance with previous studies (Marino et al., 2011; Melchior

et al., 2007; Rode et al., 2007). The risk of the presence of S. aureus would thus be

highly dependent on both the strain and the age of the biofilm. As previously observed

by the authors (Herrera et al., 2007), the importance of defining the biofilm formation

kinetics (or at least some stages, as done here) seems thus clear to draw reliable

conclusions.

Interestingly, a statistical trend was found between 48-h-old biofilm biomass and the

type of processing applied to the fishery products that strains were isolated from. This

trend seems to show a selective pressure of food-processing conditions on S. aureus.

Thus, most strains isolated from highly-processed products showed a biofilm-forming

ability higher than those from low-processed products. Generally, a high degree of

food-processing involves a high degree of contact with surfaces and a high degree of

handling of raw materials and intermediate products. As S. aureus is a major component

Chapter 3

144

of human microbiome, a high degree of handling can enhance the spread of S. aureus to

food and food-contact surfaces (DeVita et al., 2007; Sattar et al., 2001; Simon and

Sanjeev, 2007; Sospedra et al., 2012), where it forms biofilms that increase the

resistance to external stresses such as antimicrobial compounds, high salt contents,

relatively high temperatures, etc. (Bridier et al., 2011a; Donlan and Costerton, 2002;

Van-Houdt and Michiels, 2010).

Biofilm formation was also inspected by the CRA plate test, which is easier to

perform and less time-consuming than staining methods. This method has been used

successfully for the detection of biofilm-forming strains of S. epidermidis (Handke et

al., 2004; Oliveira et al., 2006), but the interpretation of results has been controversial in

the case of S. aureus, with phenotypic colony changes taking place during incubation

and false negative being reported (Milanov et al., 2010; Vasudevan et al., 2003). In this

study, only black-coloured colonies after 72 h of incubation were considered to be

typical of a biofilm-positive phenotype, as Arciola et al. (2002) recommended. All

strains formed black-coloured colonies after 24 h, and this biofilm-positive phenotype

was maintained subsequently. These results are in accordance with those of crystal

violet staining, and it contrasts with claims that CRA test results do not always

correspond to biofilm production determined by staining methods (Jain and Agarwal,

2009; Oliveira et al., 2006). Its qualitative nature, however, limits significantly the

scope of this method. According to Arciola et al. (2002), CRA test results were

correlated to the presence of icaA and icaD genes (Vázquez-Sánchez et al., 2013),

which are usually involved in staphylococcal biofilm formation. However, recent

studies have observed a lack of correspondence between slime production and the

presence of ica genes (Ciftci et al., 2009; Zmantar et al., 2010).

Biofilms formed by all strains showed a marked resistance to benzalkonium chloride,

peracetic acid and sodium hypochlorite, being higher than ATCC 6538 in most cases.

However, a high variation in biofilm resistance was observed between strains.

Moreover, strains followed a different order according to the resistance of biofilms to

each biocide, which reveals a different response between strains. Similarly, Melchior et

al. (2007) had previously indicated that each strain requires a specific exposure time to

each antimicrobial for complete biofilm removal. Therefore, the use of a wide collection

of strains for the assessment of the bactericidal activity of disinfectants seems to be


145

necessary to ensure that they are correctly applied. However, the current European

Union standard bactericidal tests (EN 1040, EN 1276 and EN 13697) use a small

number of type strains (and only one S. aureus) to assess the efficacy of disinfectants.

Doses considerably lower than the MBECs determined in this study under some

conditions common in the food industry are thus often recommended by manufacturers

for BAC (200-1000 mg/L), PAA (50-350 mg/L) and NaClO (50-800 mg/L) (Gaulin et

al., 2011). Microorganisms could therefore be exposed to sub-lethal doses of

disinfectants and this can generate the emergence of antimicrobial resistance (Langsrud

et al., 2003; Sheridan et al., 2012). Whether or not standard bactericidal tests truly

simulate conditions found in the food industry and clinical settings has been also subject

of a long debate (Briñez et al., 2006; Langsrud et al., 2003; Meyer et al., 2010).

Likewise for biomass, the resistance of biofilms was found to increase as biofilms

aged. Therefore, ensuring biofilm removal would need to adequate antimicrobial

treatments to highly-mature biofilms, or a biofilm representing a worst-case scenario.

Accordingly, it would be expected that the higher the biofilm biomass, the higher the

MBEC. However, this was not always the case and, in fact, biomass and BAC

resistance were found not to be correlated. This lack of correlation is likely accounted

by a high variation in the expression of genes involved in biofilm formation (e.g. icaA,

rbf and σB), such as it has been recently observed between different strains used in this

study (Vázquez-Sánchez et al., 2013). This could in turn produce differences in the

composition and architecture of the extracellular matrix and, therefore, in the stability

and resistance of biofilms. A thorough knowledge of the composition and architecture

of the extracellular matrix could thus be helpful to improve the efficacy of disinfection

strategies. However, biofilms are highly complex and the extraction and purification of

extracellular matrix components is difficult, so providing a complete biochemical

profile still remains an important challenge (Flemming et al., 2007).

Although the resistance of biofilms was much higher than planktonic counterparts,

no correlation was found between MBEC and MBC. Therefore, this lack of correlation

is presumably not as much due to differences in cell resistance as to the protective role

of the extracellular matrix (Bridier et al., 2011a; Donlan and Costerton, 2002; Russell,

2003). In fact, differences between MBEC and MBC increased as biofilms aged and the

extracellular matrix developed. However, an unexpected positive correlation between

Chapter 3

146

MBEC and MBC of BAC was obtained after 168 h of incubation, probably due to cell

detachment from mature biofilms caused by nutrient depletion (Hunt et al., 2004). S.

aureus biofilms often detach in form of cell clumps which are partially protected from

antimicrobials by the extracellular matrix (Fux et al., 2004; Hall-Stoodley and Stoodley,

2005). Although MBCs were much lower than MBECs, 168-h old planktonic cultures

would have shown an antimicrobial behaviour more similar (and thereof correlated) to

that of biofilms than in earlier stages of culture, when only free-living cells (and no cell

clumps) would have been present.

The antimicrobial resistance was also affected by the temperature, as observed in

168-h-old biofilms and planktonic counterpart cells for BAC, with resistance being

lower at 12°C than at 25°C. A similar effect had been previously reported (Meira et al.,

2012; Taylor et al., 1999). This effect was highly strain dependent and this dependence

can be partially accounted for the different adaptive response of each strain to low

temperatures (as defined by the growth kinetics). The importance of the growth kinetics

explains in turn that both MBEC and MBC were found to be positively correlated with

cell density at 12°C, but no at 25°C, temperature at which all cultures were in stationary

phase after 168 h of incubation. It also explains that the correlation between MBEC and

MBC was higher at 12°C than at 25°C. Nonetheless, the development of biofilms

attenuated the adverse effect of low temperatures on BAC resistance in most cases.

Thus, the resistance of biofilms developed at 12°C and 25°C were found to be positively

correlated, but no correlation was found in the case of planktonic cells.

Significant differences in the efficacy of the industrial disinfectants tested were

observed. Peracetic acid was found to be the most effective against both biofilms and

planktonic cells, followed by sodium hypochlorite and then benzalkonium chloride.

Bridier et al. (2011b) had also reported that PAA was more effective than BAC against

planktonic cells of several S. aureus strains. Meanwhile, PAA was found to have a

higher effectiveness than NaClO against biofilms of S. aureus on different food-contact

surfaces (Meira et al., 2012). The lack of correlation between the efficacies of

disinfectants against biofilms is indicative of significant differences in the diffusion and

reactivity of each disinfectant in biofilms, which also depends on the composition of the

extracellular matrix. The high antimicrobial activity of PAA is considered to be

accounted for a high oxidative potential, a non-specificity and a high reactivity against


147

intracellular enzymes, proteins and DNA as well as against bacterial cell membrane and

extracellular matrix components, and also a small size enabling diffusion through the

biofilm matrix (Bessems, 1998; Denyer and Stewart, 1998; Kitis, 2004; Saá-Ibusquiza

et al., 2011). NaClO is a powerful oxidizing agent which disturbs the synthesis of DNA

and reacts with intracellular proteins, cell wall and extracellular matrix components

(Russell, 2003), but it has a limited diffusion in the biofilm (De-Beer et al., 1994).

Meanwhile, BAC forms mixed micellar aggregates that solubilize membrane

phospholipids and proteins and thus causes a decrease of fluidity and the appearance of

hydrophilic voids in the membrane leading to a generalized and progressive leakage of

cytoplasmic materials to the environment and eventually cell lysis (Gilbert and Moore,

2005). However, the interaction of BAC with extracellular matrix components slows

down penetration into the biofilm (Bridier et al., 2011c).

The rotation and combination of biocides is widely accepted to prevent the

emergence of antimicrobial-resistant strains. However, in practice, different

disinfectants with similar cell targets or similar mechanisms of action are often applied,

which increases the risk of cross-resistance, particularly in biofilms (Braoudaki and

Hilton, 2004; Chapman, 2003; Langsrud et al., 2004; Saá-Ibusquiza et al., 2011;

Schweizer, 2001). The introduction of novel antimicrobials (e.g. electrolyzed water,

essential oils, ozone, etc.) as well as the development of new control strategies (e.g.

incorporation of antimicrobials on surface materials, modification of the

physicochemical properties of surfaces, etc.) could be an effective alternative to avoid,

or at least reduce, the risk of biofilm formation and antimicrobial resistance. Moreover,

the use of disinfectant compounds less corrosive to metal surfaces, less hazardous for

the health of workers and more environmentally-friendly have to be imperative

nowadays.

3.5. Conclusions

S. aureus can spread from humans to food and food-contact surfaces. All strains

tested in this study showed a significant ability to adhere and form biofilms on stainless

steel surfaces. In fact, most showed a biofilm-forming ability higher than S. aureus

ATCC 6538, a common reference strain in bactericidal standard tests. Stainless steel

food-contact surfaces can thus be an important reservoir for S. aureus in the food

Chapter 3

148

industry and play an important role as a source of food contamination unless appropriate

protocols of cleaning and disinfection are applied.

It is well known that biofilms show a much higher resistance to disinfectants than

planktonic counterparts. This has driven to some controversy on the validity of standard

bactericidal tests used in the European Union, since most are suspension-based. In this

study, no correlation was found between the resistance of biofilms to BAC, PAA and

NaClO and that of planktonic cells. No extrapolation seems thus feasible. Nonetheless,

a biofilm-based standard bactericidal test (EN 13697) has been developed, but it does

not seem to truly simulate environmental conditions found in the food industry.

Moreover, the resistance of the strains did not follow the same order for BAC, NaClO

and PAA, which shows the present limitation of using a few type strains (and only one

S. aureus) in standard tests in order to ensure a proper application of disinfectants.

These different reasons could at least partially account for the fact that doses

recommended by manufacturers for BAC, PAA and NaClO to disinfect food-contact

surfaces are lower than the MBEC values determined in this study under some

conditions common in the food industry and therefore are not able to guarantee biofilm

removal.


149

Appendix: Growth kinetics

Figures showed the growth kinetics of the twelve selected S. aureus strains (St.1.01,

St.1.03, St.1.04, St.1.07, St.1.08, St.1.10, St.1.14, St.1.23, St.1.24, St.1.28, St.1.30 and

St.1.31) at 12°C and 25°C. ODt represents the optical density at 700 nm at time t (days),

whereas OD0 is the optical density at the time of inoculation. Experimental values are

represented by symbols, whereas corresponding expected values are shown as solid

lines.

Chapter 3

150

153

Chapter 4. Single and sequential application

of electrolyzed water with benzalkonium

chloride or peracetic acid for removal of


Single and sequential application of EW with BAC or PAA for removal of S. aureus biofilms

155

Single and sequential application of electrolyzed water

with benzalkonium chloride or peracetic acid for removal

of Staphylococcus aureus biofilms

Daniel Vázquez-Sánchez, Marta López Cabo, Juan José Rodríguez-Herrera

Seafood Microbiology and Technology Section, Marine Research Institute (IIM), Spanish

National Research Council (CSIC), Eduardo Cabello 6, 36208, Vigo (Spain).

Abstract

The effectiveness of electrolyzed water (EW) against biofilms formed on stainless

steel by S. aureus strains isolated from fishery products was assessed. The bactericidal

activity of EW against biofilms was hardly any affected by variations in the pH of

production. Neutral EW (NEW) was therefore used in subsequent studies as it has a

higher potential for long-term application than acidic EW (due to a lower corrosiveness

and toxicity) and due to the higher yield rate of the production unit at neutral pH.

The application of NEW caused a high reduction in the number of viable biofilm

cells initially. However, a high available chlorine concentration (800 mg/L ACC) was

needed to achieve logarithmic reductions (LR) demanded by the European quantitative

surface test of bactericidal activity (≥ 4 log CFU/cm2 after 5 min of exposure). A double

sequential application of NEW at much lower concentrations for 5 min each allowed LR

≥ 4 log CFU/cm2 to be reached in most of the experimental range. Sequential

applications of NEW and either benzalkonium chloride (BAC) or peracetic acid (PAA)

showed a similar effect, with PAA-NEW being most effective. The combination of

NEW with other antimicrobial treatments can thus be an environmentally-friendly

alternative to disinfection protocols traditionally used in the food industry.

Keywords: Staphylococcus aureus; biofilm; electrolyzed water; benzalkonium

chloride; peracetic acid.

Chapter 4

156

4.1. Introduction

The ingestion of food containing staphylococcal enterotoxins is a major cause of

foodborne intoxications in humans worldwide (EFSA, 2012; Hennekinne et al., 2012).

Nonetheless, staphylococcal food poisoning (SFP) usually resolves within 24-48 h after

onset, so most cases are not reported to healthcare services. As a result, the actual

incidence of SFP is known to be much higher than reported (Argudín et al., 2010;

Lawrynowicz-Paciorek et al., 2007).

In a recent study, a high incidence of Staphylococcus aureus (~ 25%) was found in

fishery products marketed in Spain (Vázquez-Sánchez et al., 2012). Meanwhile,

changes in Spanish national regulations (RD 135/2010), following Commission

Regulation (EC) No 2073/2005, revoked the use of S. aureus as a microbiological

criterion for a number of foods, including several types of fishery products. Being the

largest producer and the second largest consumer of seafood in the European Union

(Eurostat, 2010; FAO, 2012), the safety of seafood produced or consumed in Spain are

of outmost importance.

Because humans are natural reservoirs of S. aureus, most of the concern has focused

on preventing the spread from food handlers to food products. However, S. aureus can

also attach and form biofilms on food-contact surfaces (DeVita et al., 2007; Gutiérrez et

al., 2012; Herrera et al., 2007; Simon and Sanjeev, 2007; Sospedra et al., 2012). The

formation of biofilms increases the likelihood of long-term presence of S. aureus in

food-related environments due to an increased tolerance to adverse conditions,

including the application of biocides (Bridier et al., 2011a; Van-Houdt and Michiels,

2010; Vázquez-Sánchez et al., 2014).

Many disinfectants are used to kill undesirable microorganisms in the food industry,

but most have a high environmental impact and produce harmful effects on workers,

and many are also corrosive to metal surfaces (Zabala et al., 2011). Electrolyzed water

(EW) is a promising alternative to traditional disinfectants applied in the food industry.

It is prepared on-site by electrolysis of a saturated salt solution, so that it is less

dangerous for workers, more environmentally-friendly and less expensive (Ayebah and

Hung, 2005; Huang et al., 2008; Issa-Zacharia et al., 2010; Ozer and Demirci, 2006).

The bactericidal activity of EW derives from the combined action of a relatively high


157

available chlorine concentration (ACC) and a high positive oxidation-reduction

potential (ORP). Numerous studies have shown the effectiveness of EW against

different foodborne pathogens, including S. aureus, but only a few studies have been

conducted to determine the potential of EW to eliminate S. aureus from food contact

surfaces (Deza et al., 2005; Guentzel et al., 2008; Park et al., 2002; Sun et al., 2012).

The present study was therefore aimed to examine the potential of EW against

biofilms formed on stainless steel surfaces by several strains of S. aureus isolated from

fishery products. With this aim, the resistance of such biofilms to a single application of

different types of EW was firstly determined. Subsequently, the efficacy of the

sequential application of EW, either on its own or combined with classical disinfectants

such as benzalkonium chloride and peracetic acid, was assessed.


4.2.1. Bacterial strains

Four se-positive strains of S. aureus (St.1.01, St.1.04, St.1.07 and St.1.08) isolated

from different commercial fishery products were used (Vázquez-Sánchez et al., 2012).

The strains were formerly identified as S. aureus by specific biochemical (coagulase,

DNAse and mannitol fermentation) and genetic tests (23s rDNA), and subsequently

characterized as different strains by RAPD-PCR (Vázquez-Sánchez et al., 2012).

Bacterial stocks of each strain were maintained at ─80°C in tryptic soy broth (TSB)

(Panreac Química, Spain) containing 20% glycerol (v/v). A stock culture of each strain

was thawed and then subcultured twice in TSB at 37°C for 24 h under static conditions

prior to each experiment.

4.2.2. Antibacterial agents

Electrolyzed water (EW) was generated using an Envirolyte® EL-400 Unit

(membrane electrolytic cell, model R-40, Envirolyte Industries International Ltd.,

Estonia) following manufacturer´s recommendations. A saturated sodium chloride

solution and tap water were simultaneously pumped into the equipment at room

temperature and at a current intensity of 20-25 A. Acidic (AEW), neutral (NEW) and

basic electrolyzed water (BEW) were produced by redirecting appropriate amounts of

Chapter 4

158

acidic anolyte solution (pH = 2.0-3.0, ORP ~ 1200 mV) to the cathode chamber

containing the alkaline catholyte solution (pH = 11.0-12.0, ORP ~ ─900 mV) after

electrolysis. The oxidation-reduction potential (ORP) and the pH of each EW were

measured using a portable pH & REDOX 26 multimeter (Crison Instruments S.A,

Spain), and the available chlorine concentration (ACC) was determined by iodometric

titration (APHA-AWWA-WPCF, 1992). The physicochemical properties of each EW

are indicated in Table 4.1.

Table 4.1. Physicochemical properties of acidic (AEW), neutral (NEW) and basic electrolyzed

water (BEW) generated. ORP: oxidation-reduction potential. ACC: available chlorine

concentration.

Electrolyzed water pH ORP (mV) ACC (mg/L)

AEW 2.99 ± 0.19 1171.75 ± 53.32 565.00 ± 35.22

NEW 6.06 ± 0.12 967.17 ± 36.52 856.33 ± 19.63

BEW 7.95 ± 0.13 843.00 ± 52.53 541.00 ± 39.64

Benzalkonium chloride (50% (v/v) solution, Guinama Absoluta Calidad, Spain) and

peracetic acid (40% (v/v) solution in acetic acid:water, Fluka, Sigma-Aldrich, Spain)

were also used in sequential applications.

All disinfectants were diluted in ultrapure water to working concentrations just

before each assay.


Stainless steel coupons (AISI 304, 2B finish; Markim Galicia S.L., Spain) of


surfaces. Coupons were soaked in 2 M NaOH to remove residues, and then rinsed

several times with distilled water, air-dried in a biosafety cabinet and autoclaved before

use. One sterile coupon was placed into each well of a sterile 24-well flat-bottom

microtiter plate (Falcon®, Becton Dickinson Labware, USA).

Overnight cultures of each strain were adjusted to an absorbance value at 700 nm of

0.100 ± 0.01 with phosphate buffer saline (PBS, composed by 7.6 g/L NaCl, 0.2 g/L

KCl and 0.245 g/L Na2HPO4 (BDH Prolabo, VWR International Eurolab, Spain); and

0.71 g/L K2HPO4 (Panreac Química, Spain)). This value corresponds to a cell

concentration of 108 CFU/mL for all strains. PBS-suspended cells were then serially


159

diluted in TSB, and a 700 μL aliquot (containing approximately 7×105 CFU) was added

into each microtiter well. Inoculum size was checked in all cases by plating on tryptic

soy agar (TSA) (Cultimed, Panreac Química, Spain). A negative control with no

inoculum was included in all assays. Microplates were incubated at 25°C for 48 h under

static conditions.

4.2.4. Single application of electrolyzed water (EW)

The resistance of biofilms to each EW was assessed in terms of the logarithmic

reduction (LR) in the number of viable biofilm cells per square centimetre under the

experimental conditions tested. After biofilms being formed for 48 h at 25°C, coupons

were placed into a new microplate and washed with 1 mL of PBS for 10 s to remove

non-adhered cells. Coupons were then exposed to 1.5 mL of EW for 30 min (unless

otherwise indicated). Three coupons were used for each treatment. Three positive

controls (i.e., coupons exposed to sterile water) were also included in each assay.

Subsequently, coupons were placed into sterile glass vials and 9 ml of neutralizing broth

(0.34 g/L KH2PO4 and 5 g/L Na2S2O3 (Probus, Spain); and 3 g/L soy lecithin, 1 g/L L-

histidine and 3% (v/v) polysorbate 80 (Fagron Iberica, Spain)) were added and left to

stand for 10 min at room temperature, according to Luppens et al. (2002). Surviving

viable cells were collected by thoroughly rubbing the surface of coupons with two

sterile swabs (Deltalab, Spain). Cells were resuspended by vigorously vortexing swabs

for 1 min in 9 ml of peptone water (10 g/L triptone (Cultimed, Panreac Química, Spain)

and 5 g/L sodium chloride (BDH Prolabo, VWR International Eurolab, Spain)). Ten-

fold serial dilutions of resuspended cells were made in peptone water and aliquots of 0.1

mL of appropriate dilutions were spread on TSA plates. Also, the number of viable

biofilm cells washed out during neutralization was quantified by plating 0.1 mL aliquots

of appropriate dilutions of neutralizing broth on TSA. The total number of surviving

biofilm cells was determined by adding up both counts after incubation at 37°C for 24

h. LR was then calculated as the difference between the logarithm of the total number of

viable cells in non-EW-exposed biofilms and the logarithm of the number of surviving

viable cells in EW-exposed biofilms. All assays were repeated twice using independent

bacterial cultures.

Chapter 4

160

The resistance of planktonic cells was assessed in terms of the minimal bactericidal

concentration (MBC), which was considered to be the lowest disinfectant concentration

necessary to kill all free-living bacterial cells under the experimental conditions tested.

Planktonic counterparts (i.e. non-adhered cells) were diluted in TSB to achieve a similar

cell concentration to that in biofilms. Planktonic cells (0.1 mL) were exposed to 0.1 mL

of EW for 30 min and then immediately neutralized (2.0 mL). Eleven different

concentrations were tested for each EW (ranging between 50-550 mg/L for AEW and

BEW, and 300-800 mg/L for NEW). Each concentration was tested in triplicate in two

independent experiments. An aliquot of 0.3 mL of each neutralized disinfectant-treated

bacterial culture was added to 1.7 mL of TSB and incubated at 37°C for 24 h. Bacterial

growth was monitored visually. In addition, visually undetectable growth was checked

by plating a 0.1 mL aliquot of culture medium on TSA and searching for the presence of

colonies after 24 h at 37°C. Bacterial growth indicated the presence of viable cells in

disinfectant-treated cultures.

4.2.5. Sequential application treatments

Sequential treatments consisted in the application of NEW either twice or in

combination with benzalkonium chloride (BAC) or peracetic acid (PAA). A second

order rotatable factorial design was carried out for each sequential application assay,

according to Box et al. (1989). Each assay consisted of nine combinations of variables

with five replicates in the centre of the domain. Biocide concentrations applied during

the first (NEW1, BAC1, PAA1) and the second disinfection (NEW2, BAC2, PAA2) were

used as independent variables. Ranges and codifications for each variable are shown in

Table 4.2.

The resistance of biofilms was also assessed in terms of logarithmic reduction. Once

non-adhered cells were removed, coupons were exposed to 1.5 mL of the first biocide

for 5 min. To prevent cross-reactions between different disinfectants, each coupon was

washed with 1 mL of PBS for 10 s before being exposed to 1.5 mL of the second

biocide for another 5 min. The number of viable biofilm cells washed out was

determined by plating a 0.1 mL aliquot of the washing PBS on TSA followed by

incubation at 37°C for 24 h. Coupons were then neutralized (9 mL) for 10 min and the

number of surviving biofilm cells were determined as aforementioned. Counts of cells


161

rubbed from coupons and washed out by PBS or by neutralizing broth were added up

and LR was calculated as aforementioned. LR data were fitted to a second-order

polynomial model by means of a least-squares method. Each combination of

concentrations of biocides was tested in triplicate (i.e. three coupons) in two

independent experiments.

Table 4.2. Natural and coded values of independent variables used in central composite

rotatable experimental designs.

Codified values

Natural values

NEW1

(mg/L ACC) BAC1

(mg/L)

PAA1

(mg/L) NEW2

(mg/L ACC) BAC2

(mg/L) PAA2

(mg/L)

1 1 500 500 250 500 500 250

1 ‒1 500 500 250 100 100 50

‒1 1 100 100 50 500 500 250

‒1 ‒1 100 100 50 100 100 50

1.41 0 583 583 291 300 300 150

‒1.41 0 17 17 9 300 300 150

0 1.41 300 300 150 583 583 291

0 ‒1.41 300 300 150 17 17 9

0 0 300 300 150 300 300 150






Statistical significance was accepted at a confidence level greater than 95% (P < 0.05).

Occasionally, a level greater than 99% (P < 0.01) is considered to remark differences

between variables.

In the case of sequential application assays, a Student´s t-test was done to determine

if there were statistical differences between the coefficients of the equations obtained in

the factorial design. Additionally, the model consistency was tested by the Fisher´s F-

test applied to the following mean squares ratios: model / total error; (model + lack of

fitting) / model; total error / experimental error; lack of fitting / experimental error.

Chapter 4

162

4.3. Results

4.3.1. Single application of electrolyzed water (EW)

4.3.1.1. Effects of pH

Biofilms containing between 7.0-7.2×107 CFU/cm

2 were exposed to 500 mg/L ACC

of acidic (AEW), neutral (NEW) or basic electrolyzed water (BEW) during 30 min to

evaluate how the pH of EW affected the antimicrobial activity. Logarithmic reductions

in the number of viable biofilm cells (LR) higher than 4 log CFU/cm2

were obtained in

all cases (Figure 4.1). However, no significant differences were observed in the

resistance of biofilms to the different types of EWs.

Figure 4.1. Logarithmic reduction (LR) in the number of viable cells of 48-h-old biofilms of S.

aureus after exposure to 500 mg/L ACC of acidic (AEW), neutral (NEW) and basic

electrolyzed water (BEW) for 30 min.

The effects of the pH of EW on the resistance of 48-h-old planktonic counterpart

cells were assessed in terms of the minimum bactericidal concentration (MBC). AEW

showed the highest effectiveness against planktonic cells, with MBC values ranging

from 500 (S. aureus St.1.07) to 550 mg/L ACC (St.1.01, St.1.04 and St.1.08).

Meanwhile, the MBC of NEW ranged from 700 (St.1.07) to 750 mg/L ACC (St.1.01,

St.1.04 and St.1.08). Unfortunately, the MBC of BEW could not be determined for any

of the strains, because the highest ACC concentration generated by the EW production

unit at pH 8.0 (~ 540 mg/L ACC) was not high enough to kill all cells.

Although AEW showed the highest efficacy against planktonic cells, no differences

against biofilms were observed between the different types of EW. Consequently, NEW

was selected for the following studies bearing in mind that the yield rate of the

production unit was much higher when conditions to produce NEW were set (Table

4.1).


163

4.3.1.2. Effects of active chlorine concentration

The antimicrobial effectiveness of NEW was firstly assessed by exposing 48-h-old

biofilms of each strain to a number of ACC during 30 min. As expected, the effect of

NEW increased significantly (P < 0.01) with increasing ACC (Figure 4.2). Thus,

average LR values of 3.84, 4.44 and 5.51 log CFU/cm2 were reached by applying 200,

500 and 800 mg/L ACC, respectively. However, no significant differences were

detected among strains at each dose applied. S. aureus St.1.01 was therefore chosen for

the following studies because it forms the most resistant biofilms to BAC and PAA

among the strains tested (Vázquez-Sánchez et al., 2014).


aureus strains after exposure to different doses of neutral electrolyzed water (NEW) for 30 min.

4.3.1.3. Effects of exposure time

The effects of the exposure time on the resistance to NEW of biofilms formed by S.

aureus St.1.01 was examined after 5, 10, 15, 20, 25 and 30 min of exposure to 200, 500

and 800 mg/L ACC.

As expected, the number of viable biofilm cells decreased as either ACC or exposure

time increased (Figure 4.3). No significant differences were thus found, for instance,

between the effects of 200 mg/L ACC for 30 min and 500 mg/L ACC for 5 min, neither

between 500 mg/L ACC for 30 min and 800 mg/L ACC for 5 min. The effects of both

ACC and exposure time were clearly non-linear within the ranges of study, with the net

effect increasing noticeably after 10-15 min of exposure at 800 mg/L ACC.

Chapter 4

164


aureus St.1.01 caused by different doses of NEW at several exposure times.

According to the European standard EN 13697 (CEN, 2002), a disinfectant has to be

able to kill over 4 log CFU/cm2 after 5 min of exposure. However, this effect could only

be achieved when 800 mg/L ACC of NEW were applied. It seemed thus clear that

another strategy was needed for NEW to be effective enough against biofilms.

4.3.2. Double sequential application of NEW

As could be inferred from Figure 4.3, the application of NEW produced a high

reduction in the number of biofilm viable cells initially. Thus, a reduction of

approximately 3 log CFU/cm2 was reached after only 5 min of exposure to 200 mg/L

ACC. The resistance of 48-h-old biofilms of S. aureus St.1.01 to a double sequential

application of NEW during 5 min each was thus examined next. A central composite

rotatable experimental design with two independent variables (the ACC applied firstly

[NEW1] and secondly [NEW2]) was used.

The logarithmic reduction (LR) in the number of viable biofilm cells was adequately

described by the following second order polynomial model (statistical analyses are

specified in Appendix):


165

This model is composed by a positive first order term and a lower negative second

order term for each application. These terms reflect that the antimicrobial activity of

NEW increases sub-linearly with concentration for both applications. Hardly differences

were found in the values of the parameters and therefore in the bactericidal efficacies as

a function of the order of application. A low negative interaction term between

applications was also found. This term shows that the net effect of NEW2 decreased

when high doses of NEW1 had been applied and, on the contrary, it increased when

NEW1 was used at low doses.

As shown in Figure 4.4, a double sequential application of NEW killed over 4 log

CFU/cm2 of biofilm cells after only 5 min of exposure in most of the experimental

range. Although this strategy was highly effective, the repeated application of a same

disinfectant would involve a risk of adaptation. Therefore, it seemed convenient that

sequential applications were based on the use of different biocides, preferably with

different cellular targets. The sequential application of NEW with traditional industrial

disinfectants such as benzalkonium chloride (BAC) or peracetic acid (PAA) was

subsequently tested by means of a similar approach, based on a central composite

rotatable experimental design for two variables (the concentration of each biocide).


aureus St.1.01 after a double sequential application of NEW for 5 min each.

Chapter 4

166

4.3.3. Sequential application of NEW and BAC

When biofilms were exposed to the sequential applications of NEW1-BAC2 and

BAC1-NEW2 during 5 min each biocide, LR was described by the following polynomial

models (statistical analyses are showed in Appendix):

Likewise for the double sequential application of NEW, the effects of NEW and

BAC on LR were described by a positive first order term and a negative second order

term each, independently of the order of application. A negative interaction term was

also noted for both combinations. Significant (P < 0.01) differences were observed

between the first and second order terms of NEW1 and BAC1. Accordingly, BAC was

found to be more effective and less reactive than NEW. In contrast, interaction terms

were similar in both sequences.

LR values higher than 4 log CFU/cm2 were reached in most of the experimental

range by both NEW1-BAC2 (Figure 4.5A) and BAC1-NEW2 (Figure 4.5B) sequences.

Differences in the bactericidal efficacy of both sequences hardly existed, except in two

cases. Thus, when high concentrations of BAC were used in the first application, the

killing effect was slightly higher than when NEW was applied instead. Similarly, when

high concentrations of BAC were applied following low-medium concentrations of

NEW, the effect was slightly higher than that of the reverse sequence. Both sequences

showed a first order term for the first biocide significantly (P < 0.01) higher than that of

NEW1-NEW2, but no differences were detected for the other terms.


167

Figure 4.5A-B. Logarithmic reduction (LR) in the number of viable cells of 48-h-old biofilms

of S. aureus st.1.01 after sequential application of NEW and BAC for 5 min each. Sequences

were NEW1-BAC2 (A) and BAC1-NEW2 (B).

Chapter 4

168

4.3.4. Sequential application of NEW and PAA

The effectiveness of the sequential application of NEW and PAA was also assessed

in both possible forms: NEW1-PAA2 and PAA1-NEW2. Like previous models, the

effects of each NEW and PAA were described by a positive first order term and a

negative second order term, and a negative interaction term also appeared, as shown in

the following equations (statistical analyses are specified in Appendix):

Both first and second order terms of PAA1 were significantly (P < 0.01) higher than

those of NEW1. The effect and the reactivity of PAA were therefore higher than of

NEW. Meanwhile, the interaction term was higher (P < 0.01) in the case of NEW1-

PAA2.

LR values higher than 4 log CFU/cm2 were also achieved in most of the

experimental range by both NEW1-PAA2 (Figure 4.6A) and PAA1-NEW2 (Figure 4.6B)

sequences. Unlike previously, differences between sequences were found in a large part

of the experimental range, so the order of application seemed to have some influence on

the bactericidal effect. Thus, the effect of PAA1-NEW2 was generally higher than

NEW1-PAA2 when low-medium concentrations of the second biocide were applied,

whereas the opposite was found when low-intermediate concentrations of the first

biocide were followed by medium-high concentrations of the second.

In comparison with previous sequence NEW1-NEW2, the single terms of PAA1-

NEW2 and NEW1-PAA2 sequences were significantly (P < 0.01) higher, whereas the

interaction term was lower (P < 0.01). In contrast, no significant differences were

observed between the first order terms of BAC1 and PAA1, but the second order term of

PAA1 was significantly (P < 0.01) higher than that of BAC1. In addition, the interaction

terms of the sequences applying PAA were significantly (P < 0.01) lower than those of

BAC-based sequences.


169

Figure 4.6A-B. Logarithmic reduction (LR) in the number of viable cells of 48-h-old biofilms

of S. aureus St.1.01 after sequential application of NEW and PAA for 5 min each. Sequences

were NEW1-PAA2 (A) and PAA1-NEW2 (B).

Chapter 4

170

4.4. Discussion

In the present study, the effects of several parameters potentially affecting the

antimicrobial activity of EW (pH, dose and exposure time) on biofilms of S. aureus

were examined in a relatively wide experimental range. In contrast, most studies dealing

with the antimicrobial effect of EW have been carried out in liquid cultures (i.e.

planktonic cells) and only a few have been made on biofilms, despite they are a

common source of microbial contamination in the food industry. Actually, most works

have been focused on assessing the efficacy of EW generated at a specific pH (Chen et

al., 2013; Deza et al., 2005; Guentzel et al., 2008; Liu et al., 2006; Monnin et al., 2012;

Park et al., 2002; Phuvasate and Su, 2010). Often too, only one dose has been applied

during a particular exposure time against a small number of strains. In the case of S.

aureus, the strain ATCC 6538 is the only one used in these studies as well as in

standard bactericidal tests. However, the strains tested in this study had previously

shown a higher biofilm-forming ability on stainless steel and a higher resistance to

disinfectants commonly used in the food industry than S. aureus ATCC 6538 (Vázquez-

Sánchez et al., 2014). Therefore, it could be expected that they would have a higher

likelihood of long-term presence in food-related environments (clearly a worse-case

scenario than S. aureus ATCC 6538), which was considered to make them more

suitable for the study.

The results obtained in this study have shown that the variation in the pH of

production affects the effectiveness of EW against S. aureus in the planktonic state.

Thus, the effectiveness of EW increased with decreasing pH. These results are in

concordance with previous studies in which the effects of several types of EWs against

planktonic cells of different foodborne pathogens were assessed (Cao et al., 2009;

Rahman et al., 2010a), but the range of pH tested in such studies was lower than in this

study.

The higher efficacy of AEW and NEW in comparison with BEW is at least partially

accounted for the presence of a much higher proportion of hypochlorous acid (HClO)

(Len et al., 2000; Vorobjeva et al., 2004), which has a higher oxidizing ability than

hypochlorite ions (ClO─), predominantly present at pH above 7.5. Meanwhile,

differences between the bactericidal activities of AEW and NEW are presumably due to


171

a much higher amount of dissolved Cl2 gas, hydrogen peroxide (H2O2) and reactive

oxygen species (ROS) in the former, and of ClO─ in the latter (Len et al., 2000;

Vorobjeva et al., 2004). The low pH and the high oxidation-reduction potential (ORP)

of AEW (Table 4.1) could additionally enhance the entrance of HClO into bacterial

cells by disruption of membranes (Huang et al., 2008; McPherson, 1993).

However, the variation of pH did not affect the effectiveness of EW against S. aureus

biofilms. This lack of effect is considered to be due to the role of the extracellular

matrix of biofilms, which acts as a protective barrier that limits molecular diffusion

towards cellular targets and also reacts with active chlorine compounds (Chen and

Stewart, 1996; Oomori et al., 2000; White, 1999). Therefore, other factors were taken

into account to decide which type of EW was most appropriate for biofilm removal. In

this sense, AEW would enhance the corrosion of food-processing equipments and could

cause health issues in handlers as a consequence of Cl2 off-gassing (Ayebah and Hung,

2005; Guentzel et al., 2008), so NEW seemed to have a higher potential than AEW for

long-term application in the food industry. In addition, the yield rate of the production

unit used in the study was much higher when conditions to produce NEW were set (with

respect to AEW or BEW). Consequently, the study focused on the application of NEW

subsequently.

As expected, the effectiveness of NEW against S. aureus biofilms increased with

increasing ACC and exposure time. In fact, the high reactivity of NEW caused a high

initial reduction of viable biofilm cells, presumably by killing cells located in the outer

layers of biofilms, which are more exposed to biocide molecules. Ayebah et al. (2005)

had also shown that acidic electrolyzed water (pH ~ 2.4) produced a reduction of 4.3 to

5.2 log CFU/coupon in the number of viable cells of L. monocytogenes biofilms after

only 30-120 s. Afterwards, the inactivation rate of viable biofilm cells was much lower.

Once easily accessible cells are killed, the chemical species of EW need to diffuse

through the extracellular matrix of the biofilm to reach bacterial cells, but this matrix

acts as a barrier that physically hinders diffusion and present many reactive sites for

biocide molecules (Chen and Stewart, 1996; Oomori et al., 2000; White, 1999). When

800 mg/L ACC were applied, the antimicrobial effect of NEW increased over that

linearly expected, particularly after 15 min, which is thought to be due to a higher

exposition of bacterial cells as a result of the disruption of the biofilm matrix. Similarly,

Chapter 4

172

Kim et al. (2001) described that acidic electrolyzed water (pH ~ 2.5) decreased the

number of viable cells in Listeria monocytogenes biofilms noticeably for the first 30 s,

followed by a considerable deceleration of the antimicrobial activity and a final increase

in the inactivation rate of biofilm cells.

According to the European quantitative surface test of bactericidal activity EN

13697, a disinfectant has to be able to kill at least 4 log CFU/cm2 biofilm cells after 5

min of exposure with the aim of achieving an effective disinfection of food-contact

surfaces in a short time. High concentrations of NEW (800 mg/L ACC) were needed to

achieve this effect on S. aureus biofilms, but this would be costly and not

environmentally-friendly. Nevertheless, logarithmic reductions higher than 3 log

CFU/cm2 were obtained by applying only 200 mg/L of NEW during 5 min, which

suggested that consecutive applications of low concentrations of NEW could increase

significantly the bactericidal effect against biofilms, with clear advantages in terms of

both dose and exposure time.

As expected, a double sequential application of NEW at low concentrations was

enough to achieve logarithmic reductions higher than 4 log CFU/cm2 after 5 min of

exposure each. However, this procedure could involve a risk of adaptation to EW, as

previously observed after the repeated use of a same chlorinated disinfectant (Braoudaki

and Hilton, 2004; Bridier et al., 2011a; Lundén et al., 2003; Maalej et al., 2006;

Sanderson and Stewart, 1997). Therefore, it seemed convenient that sequential

applications were based on the use of different biocides, preferably with different

cellular targets.

The potential of combining NEW with benzalkonium chloride (BAC) or peracetic

acid (PAA) was thus examined. Logarithmic reductions demanded by the European

standard test EN 13697 were also reached in all cases by applying low concentrations of

each biocide. Comparatively, PAA was more effective than NEW and BAC, as the

concentrations of PAA used in the study were about 2-fold lower than those of NEW

and BAC. A high oxidative potential, non-specificity, a high reactivity (higher than that

of BAC, as shown by the magnitude of the second order terms) and a small size

enabling diffusion through the biofilm matrix have been indicated to account for a

higher antimicrobial activity of PAA against biofilms than BAC and chlorine-based

compounds such as NEW (Bessems, 1998; Bridier et al., 2011b; Denyer and Stewart,


173

1998; Kitis, 2004; Saá-Ibusquiza et al., 2011). However, when sequences were based on

the use of different disinfectants, washing was mandatory to prevent cross-reaction

between them. As a result, operation time increased and approximately 10% of cells

surviving after the first disinfection were washed out. Therefore, wash water would

require to be collected and subjected to additional biocidal processes (e.g., heat, UV

radiation, chlorination, ozonation) to prevent dissemination and reattachment of

dispersed cells (Casani and Knøchel, 2002; Fähnrich et al., 1998). In contrast, no

intermediate washing was needed when NEW was applied twice.

Various combined treatments have also shown an enhanced antimicrobial effect

against S. aureus biofilms respect to single disinfection (Caballero-Gómez et al., 2013;

Hendry et al., 2009; Xing et al., 2012). However, in these studies, biofilms were formed

under conditions rarely found in the food industry such as a growth temperature of 37°C

(Hendry et al., 2009; Xing et al., 2012) or exposed to antimicrobials for a longer time

(60 min) than that specified in EN 13697 (Caballero-Gómez et al., 2013), so they hardly

have any application in the food industry. In the case of EW-based combined

treatments, many studies have dealt with food products (Chen et al., 2013; Kim et al.,

2003; Koseki et al., 2004; Mahmoud et al., 2006; Rahman et al., 2010b, 2011; Wang et

al., 2004, 2006), whereas studies performed on food-contact surfaces have been

sporadic (Ayebah et al., 2005; Chen et al., 2013), despite biofilms are a common source

of food contamination. However, these two studies were focused on combining acidic

EW with alkaline EW or ultrasounds, respectively. Therefore, the NEW-based

combined treatments proposed in the present study represent an effective, safe,

environmentally-friendly and less expensive alternative to be applied in the food

industry.

4.5. Conclusions

Electrolyzed water (EW) is considered a promising alternative for bacterial

disinfection in the food industry as it is environmentally-friendly, safe-to-use and

relatively inexpensive. In contrast to planktonic cells, the bactericidal activity of EW

against biofilms was not affected by variations in the pH of production. Neutral EW

(NEW) seems thus to show a higher potential for long-term application in the food

industry, as it is less corrosive to metal surfaces and less toxic to handlers than acidic

Chapter 4

174

EW. However, a high ACC of NEW was required to comply with the specifications set

in the European quantitative surface test of bactericidal activity (LR ≥ 4 log CFU/cm2

after 5 min of exposure). Nevertheless, the combination of NEW and other disinfectants

resulted in a promising alternative to control the contamination of food-processing

facilities by foodborne pathogens in terms of dose and exposure time. This strategy can

be also extended to almost any other biocide combination, although working-safe and

environmentally-friendly options such as NEW should be preferentially selected to be in

concordance with present and future regulatory landscapes. Furthermore, the

combination of biocides should be addressed to search intelligent solutions of

disinfection in the sense described by the principles of hurdle technology for food

products. Thus, biocides should be chosen on a mechanistic basis (with the aim to

obtain some synergistic effect), so a much greater knowledge on the mechanisms of

action of biocides is required, and disinfection treatments (dose, time of exposure and

type of sequence) should be optimized by a predictive microbiology-based approach.

This strategy could reduce considerably adverse effects on working, environmental and

economic aspects.


175

Appendix

Statistical data of factorial design and test of significance for the different second order

polynomial models showed in this work.

Table Ap.1. Sequence NEW1-NEW2.

C N Y Ŷ Coefficients t Model

1 1 6.056 5.990 5.191 124.510 5.191

1 ‒1 5.254 5.114 0.573 17.373 0.573 NEW1

‒1 1 5.278 5.162 0.597 18.079 0.597 NEW2

‒1 ‒1 3.842 3.652 ‒0.159 3.401 ‒0.159 NEW1 NEW2

1.41 0 5.688 5.781 ‒0.110 3.110 ‒0.110

‒1.41 0 3.998 4.161 ‒0.101 2.854 ‒0.101

0 1.41 5.755 5.831

0 ‒1.41 3.966 4.148 Average value 5.0610

0 0 5.191 5.191 Expected average value 5.1911

0 0 5.278 5.191

0 0 5.278 5.191 Var (Ee) 0.0087

0 0 5.056 5.191 t (α < 0.05; ʋ = 4) 2.7760

0 0 5.153 5.191

SS ʋ MS MSMd/MSE 43.737 (α = 0.05) 3.860

Md 5.701 5 1.1402 MSMdLF/MSM 0.641 (α = 0.05) 9.120

E 0.182 7 0.0261 MSE/MSEe 3.000 (α = 0.05) 6.000

Ee 0.035 4 0.0087 MSLF/MSEe 5.666 (α = 0.05) 6.260

LF 0.148 3 0.0492 r2 0.969

Total 5.884 12 r2

adjusted 0.947

Y, observed response; Ŷ, expected response; SS, sum of squares; ʋ, degrees of freedom; MS,

mean squares; Md, Model; E, total error; Ee, experimental error; LF, lack of fitting; MSE, mean

squares for total error; MSEe, mean squares for experimental error; MSLF, mean squares for

lack of fitting; MSMd, mean squares for model; MSMdLF, mean squares for (model + lack of

fitting).

Chapter 4

176

Table Ap.2. Sequence NEW1-BAC2.


1 1 6.040 5.994 5.145 376.660 5.145

1 ‒1 5.040 5.029 0.608 56.313 0.608 NEW1

‒1 1 5.102 5.074 0.630 58.296 0.630 BAC2

‒1 ‒1 3.510 3.517 ‒0.148 9.690 ‒0.148 NEW1 BAC2

1.41 0 5.704 5.736 ‒0.134 11.610 ‒0.134

‒1.41 0 4.009 4.016 ‒0.107 9.197 ‒0.107

0 1.41 5.776 5.821

0 ‒1.41 4.048 4.043 Average value 4.9953


0 0 5.137 5.145

0 0 5.174 5.145 Var (Ee) 0.0009

0 0 5.102 5.145 t (α < 0.05; ʋ = 4) 2.7760

0 0 5.174 5.145

SS ʋ MS MSMd/MSE 907.777 (α = 0.05) 3.860

Md 6.398 5 1.2795 MSMdLF/MSM 0.626 (α = 0.05) 9.120

E 0.010 7 0.0014 MSE/MSEe 1.511 (α = 0.05) 6.000

Ee 0.004 4 0.0009 MSLF/MSEe 2.193 (α = 0.05) 6.260

LF 0.006 3 0.0020 r2 0.998

Total 6.408 12 r2adjusted 0.997





fitting).


177

Table Ap.3. Sequence BAC1-NEW2.


1 1 6.037 6.034 5.149 556.271 5.149

1 ‒1 5.116 5.073 0.643 87.888 0.643 BAC1

‒1 1 4.996 5.017 0.615 83.904 0.615 NEW2

‒1 ‒1 3.537 3.518 ‒0.134 12.984 ‒0.134 BAC1 NEW2

1.41 0 5.815 5.843 ‒0.108 13.737 ‒0.108

‒1.41 0 4.030 4.024 ‒0.131 16.576 ‒0.131

0 1.41 5.774 5.756

0 ‒1.41 3.983 4.022 Average value 5.0025


0 0 5.134 5.149

0 0 5.134 5.149 Var (Ee) 0.0004

0 0 5.134 5.149 t (α < 0.05; ʋ = 4) 2.7760

0 0 5.172 5.149

SS ʋ MS MSMd/MSE 1302.425 (α = 0.05) 3.860

Md 6.573 5 1.3147 MSMdLF/MSM 0.626 (α = 0.05) 9.120

E 0.007 7 0.0010 MSE/MSEe 2.356 (α = 0.05) 6.000

Ee 0.002 4 0.0004 MSLF/MSEe 4.165 (α = 0.05) 6.260

LF 0.005 3 0.0018 r2 0.999

Total 6.580 12 r2

adjusted 0.998





fitting).

Chapter 4

178

Table Ap.4. Sequence NEW1-PAA2.


1 1 6.037 6.033 5.149 556.276 5.149

1 ‒1 4.881 4.886 0.606 82.752 0.606 NEW1

‒1 1 5.009 5.023 0.674 92.016 0.674 PAA2

‒1 ‒1 3.450 3.473 ‒0.101 9.734 ‒0.101 NEW1 PAA2

1.41 0 5.701 5.705 ‒0.150 19.175 ‒0.150

‒1.41 0 4.014 3.992 ‒0.145 18.377 ‒0.145

0 1.41 5.815 5.812

0 ‒1.41 3.926 3.910 Average value 4.9676


0 0 5.172 5.149

0 0 5.134 5.149 Var (Ee) 0.0004

0 0 5.172 5.149 t (α < 0.05; ʋ = 4) 2.7760

0 0 5.134 5.149

SS ʋ MS MSMd/MSE 2908.016 (α = 0.05) 3.860

Md 6.869 5 1.3738 MSMdLF/MSM 0.625 (α = 0.05) 9.120

E 0.003 7 0.0005 MSE/MSEe 1.103 (α = 0.05) 6.000

Ee 0.002 4 0.0004 MSLF/MSEe 1.240 (α = 0.05) 6.260

LF 0.002 3 0.0005 r2 0.9995

Total 6.872 12 r2

adjusted 0.9992





fitting).


179

Table Ap.5. Sequence PAA1-NEW2.


1 1 6.037 6.045 5.206 321.421 5.206

1 ‒1 4.923 4.939 0.634 49.485 0.634 PAA1

‒1 1 4.851 4.889 0.613 47.825 0.613 NEW2

‒1 ‒1 3.495 3.551 ‒0.060 3.341 ‒0.060 PAA1 NEW2

1.41 0 5.736 5.732 ‒0.185 13.469 ‒0.185

‒1.41 0 4.000 3.940 ‒0.162 11.771 ‒0.162

0 1.41 5.774 5.748

0 ‒1.41 4.056 4.018 Average value 4.9924


0 0 5.213 5.206

0 0 5.259 5.206 Var (Ee) 0.0013

0 0 5.213 5.206 t (α < 0.05; ʋ = 4) 2.7760

0 0 5.172 5.206

SS ʋ MS MSMd/MSE 556.104 (α = 0.05) 3.860

Md 6.598 5 1.3197 MSMdLF/MSM 0.626 (α = 0.05) 9.120

E 0.017 7 0.0024 MSE/MSEe 1.809 (α = 0.05) 6.000

Ee 0.005 4 0.0013 MSLF/MSEe 2.889 (α = 0.05) 6.260

LF 0.011 3 0.0038 r2 0.997

Total 6.615 12 r2adjusted 0.996





fitting).

181

Chapter 5. Antimicrobial activity of essential

oils against Staphylococcus aureus biofilms

Antimicrobial activity of essential oils against S. aureus biofilms

183

Antimicrobial activity of essential oils against


Daniel Vázquez-Sánchez, Marta López Cabo, Juan José Rodríguez-Herrera

Seafood Microbiology and Technology Section, Marine Research Institute (IIM), Spanish

National Research Council (CSIC), Eduardo Cabello 6, 36208, Vigo (Spain).

Abstract

The effectiveness of nineteen essential oils (EOs) against planktonic cells of S.

aureus was firstly assessed by minimal inhibitory concentration (MIC). Planktonic cells

showed a wide variability in resistance to EOs, with thyme oil as the most effective,

followed by lemongrass oil and then vetiver oil. The eight EOs most effective against

planktonic cells were subsequently tested against 48-h-old biofilms formed on stainless

steel. All EOs reduced significantly (P < 0.01) the number of viable biofilm cells, but

none of them could remove biofilms completely. Thyme and patchouli oils were the

most effective, but high concentrations were needed to achieve logarithmic reductions

over 4 log CFU/cm2 after 30 min exposure.

The use of sub-lethal doses of thyme oil prevented biofilm formation and enhanced

the efficiency of thyme oil and benzalkonium chloride against biofilms. However, some

cellular adaptation to thyme oil was detected. Combined thyme oil-based treatments

were thus considered as a suitable alternative for the removal of S. aureus biofilms in

food-processing facilities. But EO-based treatments should be based on the rotation and

combination of different EOs or with other biocides to prevent the emergence of

antimicrobial-resistant strains.

Keywords: Staphylococcus aureus; biofilm; essential oil; thyme; benzalkonium

chloride.

Chapter 5

184

5.1. Introduction

One of the major causes of foodborne intoxications in humans worldwide is the

ingestion of food containing staphylococcal enterotoxins (Bhatia and Zahoor, 2007;

EFSA, 2012; Hennekinne et al., 2012). Nonetheless, it is known that the actual

incidence of staphylococcal food poisoning is underestimated, because it usually

resolves 24-48 h after onset (Argudín et al., 2010; Lawrynowicz-Paciorek et al., 2007).

Most strategies aimed to prevent the contamination of food with S. aureus are

focused on the importance of food handlers as natural reservoirs. However, this

pathogen has also a high ability to form biofilms on food-contact surfaces (DeVita et

al., 2007; Gutiérrez et al., 2012; Herrera et al., 2007; Simon and Sanjeev, 2007;

Sospedra et al., 2012). The formation of biofilms increases the resistance to adverse

conditions, such as the application of disinfectants (Bridier et al., 2011a; Van-Houdt and

Michiels, 2010; Vázquez-Sánchez et al., 2014). Consequently, forming biofilms

increases the likelihood of long-term presence in food-related environments and,

therefore, the risk of food contamination as well as the spread of the bacterium.

Many disinfectants are applied to remove undesirable microorganisms from food-

contact surfaces, but most are corrosive to metal surfaces, harmful for workers or have a

high environmental impact (Zabala et al., 2011). Thus, scientific interest in the

antimicrobial properties of plant essential oils (EOs) has emerged. EOs comprise a wide

variety of aromatic oily liquids (approximately 3000 EOs are known nowadays) with a

versatile composition, which are usually extracted from different plant materials such as

flowers, fruits, herbs, leaves, roots and seeds (Bakkali et al., 2008; Burt, 2004) EOs

have shown to have antioxidant (Brenes and Roura, 2010), antibacterial (Oussalah et al.,

2007), antiparasitic (George et al., 2009), insecticidal (Nerio et al., 2010), antiviral

(Astani et al., 2011) and antifungal (Tserennadmid et al., 2011) properties. Nonetheless,

few studies have evaluated the potential of EOs to remove S. aureus from food-contact

surfaces (Adukwu et al., 2012; Lebert et al., 2007; Millezi et al., 2012). Also, the poor

solubility and partial volatility of EOs have limited their practical application (Delaquis

et al., 2002; Kalemba and Kunicka, 2003).


185

The present study was therefore aimed to evaluate the potential of essential oils

against biofilms formed by S. aureus on stainless steel surfaces. With this aim, the

resistance of such biofilms to a single application of an array of EOs was firstly

determined. Subsequently, it was investigated the potential of applying thyme oil (the

EO with the highest effect) on the formation and removal of biofilms (on its own and in

combination with benzalkonium chloride), as well as the risk of bacterial adaptation to

thyme oil.


5.2.1. Bacterial strain

The sea-positive strain S. aureus St.1.01 was used. It was identified as S. aureus by

specific biochemical (coagulase, DNAse and mannitol fermentation) and genetic tests

(23s rDNA sequencing), and characterized by RAPD-PCR with three primers

(Vázquez-Sánchez et al., 2012). The RAPD pattern of St.1.01 was also shown by

different S. aureus isolates from several commercial fishery products. Bacterial stocks

were maintained at ─80°C in tryptic soy broth (TSB) (Panreac Química, Spain)

containing 20% glycerol (v/v). A stock culture of the strain was thawed and sub-

cultured twice in TSB at 37°C for 24 h under static conditions prior to each experiment.

5.2.2. Antibacterial agents

Nineteen pure essential oils (EOs) industrially produced by steam distillation of

different plant parts (Table 5.1) were tested in the present study. All EOs were diluted in

ultrapure water with 0.15% (w/v) bacteriological agar (Panreac Química, Spain) before

each experiment. Benzalkonium chloride was purchased as a 50% (v/v) solution

(Guinama Absoluta Calidad, Spain), which was diluted in ultrapure water to working

concentrations prior to each assay.

Chapter 5

186

Table 5.1. Origin of pure essential oils used in this study

Common name Plant species Distilled part

Anise1

Pimpinella anisum Seeds

Carrot2

Daucus carota Plant and flowers

Citronella1

Cymbopogon nardus Herbs

Coriander2

Coriandrum sativum Plant and flowers

Cumin1

Cuminum cyminum Seeds

Eucalyptus globulus

2 Eucalyptus globulus Leaves

Eucalyptus radiata3

Eucalyptus radiata Leaves

Fennel1

Foeniculum vulgare Plants

Geranium2

Pelargonium graveolens Plant and flowers

Ginger2

Zingiber officinale Plant and flowers

Hyssop1

Hyssopus officinalis Plant with flowers

Lemongrass2

Cymbopogon citratus Plant and flowers

Marjoram2

Origanum majorana Plant and flowers

Palmarosa2

Cymbopogon martinii Plant and flowers

Patchouli2

Pogostemon patchouli Leaves

Sage2

Salvia officinalis Plant and flowers

Tea tree2

Melaleuca alternifolia Leaves

Thyme1

Thymus vulgaris Plant with flowers

Vetiver2

Vetiveria zizanioides Plant and flowers

1Esential Arôms, Dietéticos Intersa S.A. (Spain)

2Mon Deconatur S.L. (Spain)

3Pranaróm International (Belgium)


Stainless steel coupons (AISI 304, 2B finish) (Markim Galicia S.L., Spain) of


surfaces. Coupons were soaked in 2 M NaOH to remove residues, rinsed several times

with distilled water, air-dried in a biosafety cabinet and autoclaved before use. One

sterile coupon was placed into each well of a sterile 24-well flat-bottom microtiter plate

(Falcon®, Becton Dickinson Labware, USA).

Overnight cultures of S. aureus St.1.01 were adjusted to an absorbance value at 700

nm of 0.100 ± 0.01 with phosphate buffer saline (PBS, composed by 7.6 g/L NaCl, 0.2

g/L KCl and 0.245 g/L Na2HPO4 (BDH Prolabo, VWR International Eurolab, Spain);

and 0.71 g/L K2HPO4 (Panreac Química, Spain)), which corresponds to a cell


187

concentration of 108 CFU/mL. PBS-suspended cells were then 100-fold serially diluted

in TSB and 700 μL aliquots (containing approximately 7×105 CFU) were added into

each well with a coupon. Inoculum size was checked in all cases by plating on tryptic

soy agar (TSA) (Cultimed, Panreac Química, Spain). A negative control with no

inoculum and a blank with medium only were included in all assays. Microplates were

incubated at 25°C under static conditions until analysis.

5.2.4. Biocide resistance assays

5.2.4.1. Resistance of planktonic cells

The resistance of planktonic cells to EOs was evaluated in terms of minimal

inhibitory concentration (MIC), which was defined as the lowest concentration at which

no bacterial growth was detected under experimental conditions. The method proposed

by Mann and Markham (1998) was followed, but with some modifications.

Aliquots of 95 μL of bacterial cultures adjusted to an absorbance of 0.100 ± 0.01 as

aforementioned (containing thus approximately 105 CFU) were exposed to 95 μL of a

specific series of concentrations of each EO in a sterilized 96-well U-bottom microtiter

plates (Falcon®, Becton Dickinson Labware, USA). For each EO, sixteen different

concentrations (0.01%, 0.02%, 0.03%, 0.04%, 0.05%, 0.06%, 0.07%, 0.08%, 0.09%,

0.10%, 0.50%, 1%, 1.5%, 2%, 2.5% and 3% (v/v)) were tested in triplicate in two

independent experiments. A positive control with no EOs, a negative control with no

inoculum and a blank with medium only were included in all assays.

Microplates were incubated for 24 h at 37°C under static conditions. Wells were then

stained with 10 µl of 0.01% (w/v) resazurin sodium salt solution (Sigma-Aldrich

Química, Spain) and incubation at 37°C continued for further 2 h. Bacterial growth was

monitored visually as colour change from blue to pink, which indicates the presence of

viable cells in cultures. Visually undetectable growth was also checked by plating 0.1

mL aliquots of cultures on TSA and searching for the presence of colonies after 24 h at

37°C.

5.2.4.2. Resistance of biofilms

The resistance of biofilms to EOs was determined in terms of the logarithmic

reduction of viable biofilm cells (LR) caused by treatment with EO.

Chapter 5

188

The EOs with the highest effectiveness against planktonic cells were assessed against

48-h-old biofilms (i.e., biofilms representing an example of worst-case scenario in the

food industry). After incubation time, coupons were placed into a new microplate and

washed with 1 mL of PBS to remove non-adhered cells. Coupons were then exposed to

1.5 mL of EO for 30 min. For each EO, twelve concentrations (0.10%, 0.25%, 0.50%,

0.75%, 1%, 1.5%, 2%, 2.5%, 3%, 4%, 6% and 8% (v/v)) were tested in triplicate. Three

positive controls consisting of biofilms exposed to sterile water were also included in

each assay. Subsequently, coupons were placed into sterile glass vials and 9 mL of

neutralizing broth (0.34 g/L KH2PO4 and 5 g/L Na2S2O3 (Probus, Spain); and 3 g/L soy

lecithin, 1 g/L L-histidine and 3% (v/v) polysorbate 80 (Fagron Iberica, Spain)) were

added and left to stand for 10 min at room temperature, following Luppens et al. (2002).

Biofilm cells were then collected by thoroughly rubbing the surface of coupons with

two sterile swabs (Deltalab, Spain). Swabs were immersed in 9 mL of peptone water

(10 g/L triptone (Cultimed, Panreac Química, Spain) with 5 g/L sodium chloride (BDH

Prolabo, VWR International Eurolab, Spain)) and vigorously vortexed for 1 min for

resuspension of cells. Ten-fold serial dilutions of resuspended cells were made in

peptone water and 0.1 mL aliquots of appropriate dilutions were spread on TSA plates.

Also, the number of viable biofilm cells washed out during neutralization was

quantified by plating 0.1 mL aliquots of appropriate dilutions of neutralizing broth on

TSA. Surviving biofilm cells were counted by adding up both counts after incubation at

37°C for 24 h. LR was determined as the difference between the logarithm of the total

number of viable cells in non-disinfectant-exposed biofilms and the logarithm of the

number of surviving viable cells in disinfectant-exposed biofilms.

Subsequently, the effects of the presence of thyme oil at sub-lethal doses (0.020%,

0.025%, 0.030% and 0.040% (v/v)) during biofilm formation on the efficacy of thyme

oil and BAC against such biofilms was evaluated. Because those sub-lethal doses

inhibited the growth of S. aureus, biofilms were formed during 7, 10, 14 and 26 days,

respectively, to have a concentration of viable cells between 5-9×107 CFU/cm

2 in all

cases. Biofilms formed during 4 days in the absence of thyme oil were also included as

a control. After incubation, biofilms were exposed to thyme oil (the same sub-lethal

concentrations) or BAC (50, 200, 1000, 2500, 5000, 10000, 15000 and 20000 mg/L) for


189

30 min and then immediately neutralized. Finally, viable biofilm cells were counted and

LR was calculated as aforementioned.

All assays were repeated twice using independent bacterial cultures.

5.2.4.3. Determination of growth kinetics

A series of test tubes containing 5 mL TSB with 0.15% agar and different sub-lethal

doses of thyme oil (0.020%, 0.025%, 0.030% and 0.040% (v/v)) were inoculated with

100 μL of cultures of S. aureus containing approximately 105 CFU and subsequently

incubated at 25°C under static conditions for up to 30 days. Absorbance was read out at

700 nm in triplicate every 24 h of incubation. Each tube was used for only one reading

and then discarded. A positive control with no thyme oil, a negative control with no

inoculum and a blank with medium only were included in all assays. Growth kinetics

was determined using two independent bacterial cultures.

Experimental data were fitted to the Gompertz equation proposed by Zwietering et

al. (1990):

(

) { [

]}

where ODt is the optical density at 700 nm at time t (days), OD0 is the optical density at

the time of inoculation, A is a dimensionless asymptotic value, µmax is the maximum

specific growth rate (1/days), and λ is the detection time (days), i.e., the time at which

OD at 700 nm is firstly noted to increase.






Statistical significance was accepted at a confidence level greater than 95% (P < 0.05),

but a level greater than 99% (P < 0.01) is occasionally considered to remark differences

between variables. The variability among EOs was calculated by means of the

coefficient of variation (CV), which is defined as the ratio of the standard deviation to

the mean.

Chapter 5

190

5.3. Results

5.3.1. Effectiveness of essential oils (EOs) against planktonic cells

The resistance of S. aureus planktonic cells to 19 different EOs was determined in

terms of the minimum inhibitory concentration (MIC). As shown in Figure 5.1, a very

high degree of variability was found in the effectiveness of the different EOs tested (CV

= 88.35%). Thus, whereas thyme oil showed the highest efficacy (MIC = 0.04% (v/v)),

followed closely by lemongrass (MIC = 0.06%) and vetiver (MIC = 0.07%) oils, anise,

carrot and ginger oils were very much less effective, with a MIC over 50-fold higher

(MIC = 3%). In between, the application of citronella, cumin, geranium, palmarosa and

patchouli oils showed a MIC of 0.5%, whereas coriander, Eucalyptus globulus, E.

radiata, fennel, hyssop, marjoram, sage and tea tree oils showed a MIC of 1%. On the

basis of these results, the most effective EOs (i.e., thyme, lemongrass, vetiver,

citronella, cumin, geranium, palmarosa and patchouli oils) were tested against biofilms

subsequently.

Figure 5.1. Minimum inhibitory concentration (MIC) of essential oils against S. aureus St.1.01

planktonic cells.


191

5.3.2. Effectiveness of EOs against 48-h-old biofilms

The resistance of 48-h-old biofilms to the selected EOs was determined in terms of

the logarithmic reduction of viable biofilm cells (LR) caused by EO-treatments.

None of the EOs was able to eradicate completely biofilms. Likewise for planktonic

cells, a high variability was found in the effectiveness of EOs. The variability was

higher at high concentrations (CV ~ 24% at 8%) than at low concentrations (CV ~ 5.6%

at 0.10%).

Thyme and patchouli oils showed the highest antimicrobial efficacy against biofilms

in all cases, followed by citronella oil (Figure 5.2A-L). Although no significant

differences were found between these three EOs in the range 0.10-0.25%, citronella oil

was significantly (P < 0.05) less effective than thyme and patchouli oils at higher

concentrations. Thus, a LR ~ 3.5 log CFU/cm2 was achieved with the application of 8%

citronella oil, whereas the same dose of patchouli and thyme oils caused a LR of 4.2 and

4.3 log CFU/cm2, respectively.

The efficacy of the remaining EOs against biofilms was found to be dependent on the

dose. In general, palmarosa and vetiver were least effective, with LR significantly (P <

0.05) lower than those of lemongrass (in the range 0.75-8%), geranium (1.5-8%) and

cumin (2-8%) oils. In fact, only palmarosa showed a significantly (P < 0.05) higher

effect than cumin at 0.25% and 0.75%. Lemongrass was significantly (P < 0.05) more

effective than cumin and geranium at low doses (0.25-2.5% and 0.5-1.5% respectively),

whereas cumin showed a significantly (P < 0.05) higher efficacy than lemongrass in the

range 6-8%. Meanwhile, geranium was significantly (P < 0.05) more effective than

cumin between 1-2.5%.

Noteworthy differences were found between the ranking of EOs on the basis of the

effectiveness against biofilms and planktonic cells. In particular, lemongrass and vetiver

oils were highly efficacy against planktonic cells but poorly against biofilms, whereas

patchouli and citronella oils showed a high effect against biofilms but rather moderate

against planktonic cells. In contrast, thyme oil showed the highest efficacy against both

biofilms and planktonic cells, so this EO was chosen for use in subsequent studies.

Chapter 5

192

Figure 5.2A-L. Logarithmic reduction of viable cells (LR) in 48-h-old biofilms exposed to

different EOs for 30 min. Each figure (A-L) corresponds to each dose tested. Letters denote

statistically significant differences (P < 0.05) between EOs at each dose, being defined the most

effective EO with the letter a.

5.3.3. Effects of sub-lethal doses of thyme oil on bacterial growth

Cultures of S. aureus St.1.01 were exposed to four different sub-lethal doses of

thyme oil (0.020%, 0.025%, 0.030% and 0.040% (v/v)) for 30 days at 25°C to evaluate

whether it inhibited growth kinetics.

The application of sub-lethal doses of thyme oil slowed down the growth of St.1.01

(Figure 5.3A). The detection time (λ) increased significantly (P < 0.01) with the dose of

thyme oil (Table 5.2). Thus, λ was found to be 25-fold higher when the dose of thyme

oil was increased from 0.020 to 0.040%. The addition of thyme oil also produced a


193

significant (P < 0.01) reduction in µmax, which was approximately 2-fold lower between

0.025-0.040% than when no EO was added (µmax = 3.375 ± 0.014 1/days).

Consequently, whereas bacterial cells reached stationary phase after about 4 days in the

absence of thyme oil, it was delayed for approximately 3, 6, 10 and 22 days when

0.020%, 0.025%, 0.030% and 0.040% of thyme oil, respectively, were present.

Bacterial cells of stationary-phase cultures obtained after 26 days at 25°C in the

presence of 0.040% of thyme oil were subsequently used to find out if they had adapted

to thyme oil. The kinetics of growth of these cells showed slight differences in λ, with

values 0.3-0.9 days shorter than non-thyme oil-exposed cells (Table 5.2). It was also

observed a slight increase in µmax at 0.020-0.025% of thyme oil (0.14-0.25 1/days

faster), but no significant differences were found in µmax between 0.030-0.040%. As a

result, cultures of thyme oil-exposed cells reached stationary phase about 1 day earlier

(Figure 5.3B). Therefore, thyme oil-exposed cells would seem to show some adaptation

to sub-lethal doses of thyme oil.

Figure 5.3A-B. Growth kinetics of S. aureus in the presence of sub-lethal doses of thyme oil.

Non-thyme oil-exposed cells (A) and cells previously exposed to thyme oil (0.040% (v/v)) for

26 days (B) were cultured at 25°C. Experimental values are represented by symbols, whereas

corresponding expected values are shown as solid lines.

Chapter 5

194

Table 5.2. Detection time (λ) and maximum specific growth rate (µmax) of non-thyme oil-

exposed cells and thyme oil-exposed cells of S. aureus grown at 25°C in the presence of several

sub-lethal doses of thyme oil (0.020%, 0.025%, 0.030% and 0.040% (v/v)). Thyme oil-exposed

cells were obtained by previously culturing in the presence of 0.040% thyme oil for 26 days at

25°C.

Parameter Thyme oil (% v/v) Non-thyme oil-exposed cells Thyme oil-exposed cells

λ

(days)

0.020* 0.749 ± 0.012

0.416 ± 0.047

0.025* 3.719 ± 0.022

3.026 ± 0.021

0.030* 6.898 ± 0.043

6.618 ± 0.080

0.040* 18.550 ± 0.050

17.634 ± 0.089

µmax

(1/days)

0.020* 1.851 ± 0.013

2.097 ± 0.032

0.025* 1.575 ± 0.023

1.719 ± 0.012

0.030 1.500 ± 0.020

1.467 ± 0.043

0.040 1.310 ± 0.044

1.239 ± 0.031

*Statistically significant differences (P < 0.05) in λ or µmax between both types of cells at same

sub-lethal dose.

5.3.4. Effectiveness of thyme oil against biofilms formed under sub-lethal

doses of thyme oil

A study was carried out next to assess whether the resistance of biofilms to thyme oil

changed as a result of the presence of sub-lethal doses of thyme oil during biofilm

formation. Because of the inhibitory effect of sub-lethal doses of thyme oil on the

growth of S. aureus, biofilms obtained when cultures reached stationary phase were

tested. All biofilms contained between 5-9×107 CFU/cm

2.

As shown in Figure 5.4, the application of 0.040% of thyme oil produced a slight

decrease in the number of viable cells (LR ~ 0.5 log CFU/cm2) of biofilms formed in the

absence of this EO. However, the resistance of biofilms to thyme oil decreased

significantly (P < 0.01) when sub-lethal doses of thyme oil were present during biofilm

formation, with LR > 3 log CFU/cm2 being frequently achieved. Moreover, some

combinations of thyme oil caused LR > 4 log CFU/cm2 (0.020% + 0.040%, 0.030% +

0.030%, 0.025% + 0.040% and 0.040% + 0.030%) and even LR > 5 log CFU/cm2

(0.030% + 0.040% and 0.040% + 0.040%), where the former figure is the concentration

of thyme oil present during biofilm formation, and the latter is the concentration applied

for biofilm removal.


195

Figure 5.4. Logarithmic reductions of viable cells (LR) in biofilms formed under different sub-

lethal doses of thyme oil after treatment with thyme oil for 30 min.

Although combining the presence of a sub-lethal dose of thyme oil during biofilm

formation with the subsequent application of thyme oil for biofilm removal seems to be

a highly effective strategy, it involves some risk of adaptation to thyme oil, as reported

in the previous section. It was therefore considered convenient to examine the

effectiveness of applying a different biocide to remove biofilms, in particular

benzalkonium chloride, which is commonly used in the food industry.

5.3.5. Effectiveness of benzalkonium chloride (BAC) against biofilms

formed under sub-lethal doses of thyme oil

The effectiveness of BAC against biofilms formed under sub-lethal doses of thyme

oil (0.020%, 0.025%, 0.030% and 0.040% (v/v)) was therefore assessed subsequently.

As shown in Figure 5.5, the presence of sub-lethal doses of thyme oil during biofilm

formation also increased significantly (P < 0.01) the susceptibility of biofilms to BAC,

with LR > 3 log CFU/cm2 being achieved in most of the experimental range. Some

combinations of thyme oil and BAC also caused LR > 4 log CFU/cm2 (0.020% thyme

oil and 10000-20000 mg/L BAC, 0.025% thyme oil and 10000-15000 mg/L BAC,

0.030-0.040% thyme oil and 5000-10000 mg/L BAC) and even LR > 5 log CFU/cm2

(0.020-0.025% thyme oil and 20000 mg/L BAC, and 0.030-0.040% thyme oil and

15000-20000 mg/L BAC).

Chapter 5

196

Figure 5.5. Logarithmic reductions of viable cells (LR) in biofilms formed under different sub-

lethal doses of thyme oil after the application of BAC for 30 min.

5.4. Discussion

The results obtained in this study have shown a high variability in the effectiveness

against S. aureus planktonic cells among nineteen EOs. Variations in the nature and

concentration of active compounds, as well as synergistic interactions between some of

them, likely explain some variability in antimicrobial activity of EOs (Bakkali et al.,

2008; Burt, 2004; Hyldgaard et al., 2012). Thus, phenol compounds were found to be

most active, followed by aldehydes, ketones, alcohols, ethers and hydrocarbons

(Kalemba and Kunicka, 2003). A high variability in the efficacy of EOs against

planktonic cells of S. aureus had been previously reported (Ertürk, 2010; Hammer et al.,

1999; Mayaud et al., 2008; Prabuseenivasan et al., 2006; Silva et al., 2013). However,

the ranking of EOs based on the biocidal effect on S. aureus planktonic cells was

considerably different from those previous studies, probably due to the use of different

strains and assay conditions. Standard bactericidal tests used as a reference the clinical

strain S. aureus ATCC 6538. However, a food-related strain (St.1.01) was tested in the

present study because it has both a biofilm-forming ability on stainless steel and a

resistance to disinfectants commonly used in the food industry higher than S. aureus

ATCC 6538 (Vázquez-Sánchez et al., 2014). Thus, it was considered more suitable (at

least clearly a worse-case scenario than S. aureus ATCC 6538) to evaluate the potential

of the application of EOs against biofilms in the food industry.

As expected, the resistance of S. aureus to EOs increased notably as a result of

biofilm formation with respect to free-living counterparts. However, the majority of


197

studies about the antimicrobial effects of EOs have been carried out using suspension

cultures (i.e. planktonic cells), whereas only a few studies have been made on biofilms

despite they are a common source of microbial contamination in the food industry. In

fact, this is the first study that evaluates the activity of cumin, geranium, palmarosa,

patchouli and vetiver oils against S. aureus biofilms. It is well-known that the use of

suspension cultures as experimental models underestimates considerably the doses

needed for removal of bacterial cells. But more importantly, this study clearly shows

that the ranking of EOs by antimicrobial effectiveness against biofilms and planktonic

cells was dissimilar. It thus seems that factors such as the composition and architecture

of the biofilm as well as the reactivity, hydrophobicity and diffusion rate of EOs into the

biofilm matrix affect significantly the effectiveness of EOs.

The eight most effective EOs against planktonic cells also reduced the number of

biofilm viable cells significantly, but none of them was able to completely eradicate S.

aureus biofilms at doses tested (0.1-8% (v/v)). It seems thus clear that preventing

biofilm formation should be targeted rather than disruption and removal of biofilms, as

suggested by Kelly et al. (2012). In fact, Adukwu et al. (2012) also reported that

lemongrass and grapefruit oils did not remove 48-h-old biofilms of three MSSA and

two MRSA strains at any concentration tested (i.e., 0.06-4%), despite biofilms were

exposed for 24 h to EOs. Neither the exposure to 1% of lemon or citronella oils for 15

min was able to kill 240-h-old biofilms of S. aureus formed on polypropylene (Millezi

et al., 2012). Only in the case of the clinical strain S. aureus DMST 4745, 24-h-old

biofilms formed on polystyrene at 37ºC were totally eradicated with the application of

4% of lemongrass for 1 h (Aiemsaard et al., 2011).

Likewise for planktonic cells, thyme oil (from Thymus vulgaris) was found to be the

most effective EO against biofilms of S. aureus. Kavanaugh and Ribbeck (2012) also

observed a high efficacy of thyme oil against biofilms formed by a clinical MRSA

strain, with a minimum biofilm eradication concentration (MBEC) lower than tee tree

oil (0.15% and > 5%, respectively). The high efficacy of thyme oil has been allocated to

a high content in thymol and carvacrol (Hudaib et al., 2002; Rota et al., 2008). These

two phenolic compounds cause alterations in fluidity, permeability and composition of

membrane, depletion of intracellular ATP and ATPase inhibition, as well as

perturbation of different metabolic pathways and gene responses (Di-Pasqua et al.,

Chapter 5

198

2007; La-Storia et al., 2011; Trombetta et al., 2005; Walsh et al., 2003). In S. aureus,

thymol also inhibits the synthesis and secretion of α-hemolysin, SEA and SEB (Qiu et

al., 2010). The high hydrophilicity of thymol and carvacrol has been suggested to

account for the antimicrobial activity of thyme oil against biofilms, as it could enhance

the diffusion of thyme oil into the extracellular matrix (Griffin et al., 1999; Nostro et al.,

2007).

The presence of sub-lethal doses of thyme oil inhibited growth kinetics of planktonic

cultures of S. aureus St.1.01 (the detection time increased whereas the maximum

specific growth rate decreased). It was therefore considered that it could also affect

biofilm formation. Thus, when biofilms were formed in the presence of sub-lethal doses

of thyme oil, the resistance of biofilms to thyme oil and BAC decreased highly. This

decrease was proportional to the sub-lethal dose applied. As a result, the doses of

biocide needed to significantly reduce viable biofilm cells (i.e. LR ≥ 4 log CFU/cm2)

were much lower than in biofilms formed with no thyme oil present. The treatment of

food-contact surfaces with thyme oil would thus be an interesting alternative to inhibit

or at least slow down the formation of S. aureus biofilms as well as to reduce the doses

of disinfectant required to be eradicated.

Thyme oil also showed a higher antimicrobial potential against S. aureus biofilms

than BAC, a chemical widely used for disinfection of stainless steel surfaces in the food

industry. This is probably due to the high reactivity of BAC with extracellular matrix

components, which would diminished the concentration of active compounds that are

effective against biofilm cells (Bridier et al., 2011b). Thyme oil-based treatments can

thus be considered as an effective, safe, less corrosive and more environmentally-

friendly alternative in the food industry.

Nonetheless, growth kinetics of planktonic cultures of S. aureus St.1.01 has shown

that a long-term exposure to thyme oil can produce some adaptation (shown by a

decrease in λ and an increase in µmax). Previously, bacterial adaptation to individual

active compounds of EOs (e.g., carvacrol, thymol, camphor or cineole) had been

detected (Di-Pasqua et al., 2010; Turina et al., 2006; Ultee et al., 2000), but not to crude

EOs (including thyme oil). However, it is thought that adaptation was affected by the

volatility of some active compounds of thyme oil, so it actually occurred at a

concentration lower than 0.040%. Thus, the growth of thyme oil-adapted cells would


199

also start at a concentration lower than the initial dose, particularly at high initial sub-

lethal doses.

To prevent the emergence of antimicrobial-resistant strains, antimicrobial treatments

using EOs should be based on the rotation and combination of different EOs (or

combinations of different active compounds that produce a synergistic lethal effect) or

with other biocides. Several EO-based combined treatments have been proposed as an

antimicrobial strategy against S. aureus. Blends of oregano, basil and bergamot have

shown a synergistic bactericidal effect (Lv et al., 2011), whereas an additive bactericidal

effect was detected for mixtures of cinnamon and clove (Goñi et al., 2009).

Combinations of clove and rosemary (Fu et al., 2007) or thyme and anise (Al-Bayati,

2008) have been also tested, but the efficacy did not significantly increase with respect

to the application of these EOs individually. However, none of these studies has dealt

with S. aureus biofilms. In fact, only Oliveira et al. (2010) has evaluated the

effectiveness of an EO-based combined treatment (blends of citronella and lemongrass)

against biofilms of L. monocytogenes, but they were formed at a temperature rather

uncommon in the food industry (37°C). Therefore, the combined treatments using

thyme oil proposed in this study represent a novel strategy for prevent biofilm formation

on food-contact surfaces as well as a highly effective disinfectant procedure.

5.5. Conclusions

Many essential oils (EOs) are well-known to have antimicrobial properties, so they

are considered a promising alternative for disinfection in the food industry. However,

not all EOs are highly effective antimicrobials. Thus, a high variability in the

effectiveness against S. aureus biofilms and planktonic cells was found among EOs

tested. But more importantly, the rankings of EOs by antimicrobial effectiveness against

biofilms and planktonic cells were dissimilar. Therefore, biofilm-related factors should

be also taken into account to select the most effective EO to be applied in the food

industry.

Though several EOs showed a high antimicrobial effect against S. aureus biofilms,

with thyme oil as most effective, none of them was able to completely eradicate

biofilms at doses tested. The use of EOs should thus be targeted to the prevention of

Chapter 5

200

biofilm formation rather than disruption and removal of biofilms formed. The presence

of thyme oil at sub-lethal doses was found to slow down the development of S. aureus

biofilms, as well as to notably enhance their subsequent removal by exposure to

antimicrobials such as thyme oil or BAC. The treatment of food-contact surfaces with

thyme oil would be an interesting preventive strategy. However, loss of active

compounds due to volatility would have to be solved. Also, long-term exposure to

thyme oil could cause bacterial adaptation. Therefore, treatments against biofilms

should be based on the rotation and combination of different EOs (or active compounds)

or with other biocides to prevent the emergence of antimicrobial-resistant strains. And

the combination of biocides should be addressed to search intelligent solutions of

disinfection in a similar sense to hurdle technology. This approach could reduce adverse

effects on working, environmental and economic aspects. The choice of biocides should

thus be aimed to obtain synergistic effects so a greater knowledge of the mechanisms of

action of biocides would be required. Individual active compounds of EOs would be

used preferentially instead of crude EOs.

203

General Discussion

General Discussion

205

General Discussion

Despite the great importance of fishery products in Spain –it is the largest producer

and the second largest consumer in the European Union (Eurostat, 2010; FAO, 2012)–,

the first chapter of the present thesis represents the first incidence study that analyses

different categories of fishery products of different origin and type of processing

commercialized in Spain. In fact, previous information available on microbial safety of

fishery products made or sold in Spain is scarce, and only González-Rodríguez et al.

(2002) and Herrera et al. (2006) examined the presence of S. aureus in cold-smoked

salmon and 50 samples of fresh marine fish. A high incidence of S. aureus was detected

in commercialized fishery products (~ 25%), being most of them se-carriers (i.e.,

putative enterotoxigenic strains). In addition, a significant proportion of these products

(~ 11%) exceeded the regulatory limits in force at the time the study was conducted,

which involves a serious risk of staphylococcal food poisoning for the consumer. These

results also questioned the adequacy of recent changes in Spanish national regulations

(RD 135/2010), following Commission Regulation (EC) No 2073/2005, which revoked

the use of S. aureus as a microbiological criterion for a number of foods, including

several fishery products.

Moreover, the emergence of multidrug resistant pathogens is additionally generating

an environmental hazard to the food supply and human health, as it makes eradication

more difficult and incidence to increase (Livermore, 2000; Popovich et al., 2007;

Ribeiro et al., 2007). Particularly, methicillin-resistant S. aureus (MRSA) are being

increasingly found outside clinical settings (Popovich et al., 2007; Ribeiro et al., 2007;

Stankovic et al., 2007), including in fishery products (Beleneva, 2011). In the present

thesis, however, all S. aureus isolated from marketed fishery products were found to be

multidrug-resistant MSSA (Chapter 1). Although EFSA (2009b) reported that there is

no current evidence that eating food contaminated with MRSA may lead to an increased

risk of humans becoming healthy carriers or infected with MRSA strains, the risk of

multidrug-resistant MSSA, which are present in food more frequently than MRSA, has

not been examined yet. In fact, it has been recently underlined the need of incidence

studies of multidrug-resistant strains in food to clarify their public health relevance

(Waters et al., 2011). Meanwhile, some preventive control measures should be taken.

General Discussion

206

It is also well known that biofilm formation allows bacteria to tolerate adverse

environmental conditions (e.g. presence of biocides) much better than free-living

counterparts (Srey et al., 2013; Van-Houdt and Michiels, 2010). In this thesis, putative

enterotoxigenic S. aureus strains isolated from fishery products exhibited, generally, a

high biofilm-forming ability on two common materials of food-contact surfaces,

polystyrene and stainless steel (Chapters 2 and 3). As a result, these strains also showed

a high resistance to three disinfectants traditionally used in the food industry such as

benzalkonium chloride (BAC), peracetic acid (PAA) and sodium hypochlorite (NaClO).

Moreover, the minimum biofilm eradication concentrations (MBECs) determined in this

work under some conditions usual in the food industry were considerably higher than

doses often recommended by manufacturers for BAC (200-1000 mg/L), PAA (50-350

mg/L) and NaClO (50-800 mg/L) (Gaulin et al., 2011). Among these disinfectants, PAA

was found to be the most effective against both biofilms and planktonic cells, followed

by NaClO and then BAC, but latters are more frequently used in the food industry.

Accordingly, the entrance of these strains in food-processing facilities does not only

involve an immediate risk to food safety but more importantly the risk of long-term

presence (even persistence) unless appropriate measurements are applied to kill them.

The risk of the presence of S. aureus on food-contact surfaces was found to be highly

strain dependent. In fact, a high variation in the expression of genes involved in biofilm

formation (e.g. icaA, rbf, sarA and σB) was detected between different strains (Chapter

2), which could in turn produce differences in the composition and architecture of the

extracellular matrix and, therefore, in the stability and resistance of biofilms.

Nonetheless, the current European Union standard bactericidal tests EN 1040 (CEN,

2005), EN 1276 (CEN, 2009) and EN 13697 (CEN, 2002) use a small number of type

strains (and only one S. aureus, the strain ATCC 6538) to assess the efficacy of

disinfectants. In this regard, most strains isolated in this thesis from fishery products

showed a higher biofilm-forming ability and biocide resistance than S. aureus ATCC

6538. Thus, the use of a wide collection of strains for the assessment of the bactericidal

activity of disinfectants seems to be necessary to ensure that they are correctly applied

on surfaces.

Data obtained in this thesis also showed that different environmental factors present

in the food industry (e.g., temperature, ionic strength, availability of nutrients) can

General Discussion

207

affect the adhesion and biofilm formation of S. aureus on food-contact surfaces

(Chapters 2 and 3). Interestingly, a statistical trend was found between biofilm-forming

ability and the type of processing applied to the fishery products that strains were

isolated from, which seems to show a selective pressure of food-processing conditions

on S. aureus. Moreover, it was observed that temperature also affected the antimicrobial

resistance of 168-h-old biofilms and planktonic counterpart cells to BAC, being this

effect highly dependent to the adaptive response of each strain to low temperatures (as

defined by the growth kinetics). Therefore, it is also important that standard bactericidal

tests truly simulate conditions found in the food industry and clinical settings, as

previously recommended some authors (Briñez et al., 2006; Langsrud et al., 2003;

Meyer et al., 2010).

The introduction of innovative disinfection strategies could be an effective

alternative to avoid, or at least reduce, the risk of biofilm formation and antimicrobial

resistance. In the present thesis, electrolyzed water (EW) and several essential oils

(EOs) have been evaluated as alternatives to traditional disinfectants used in the food

industry, because they are less hazardous for the health of workers and more

environmentally-friendly.

In the case of EW (Chapter 4), data provided in this thesis showed that variations in

the pH of production not significantly affect the effectiveness of EW against S. aureus

biofilms, in contrast to planktonic cells. This lack of effect is considered to be due to the

role of the extracellular matrix of biofilms, which acts as a protective barrier that limits

molecular diffusion towards cellular targets and also reacts with active chlorine

compounds (Chen and Stewart, 1996; Oomori et al., 2000; White, 1999). Neutral EW

(NEW) was therefore used in subsequent studies as it has a higher potential for long-

term application than acidic EW (i.e., a lower corrosiveness and toxicity (Ayebah and

Hung, 2005; Guentzel et al., 2008)) and due to the higher yield rate of the production

unit at neutral pH.

The efficacy of NEW against S. aureus biofilms increased with increasing ACC and

exposure time. The application of NEW caused a high initial reduction in the number of

viable biofilm cells, presumably by killing cells located in the outer layers of biofilms,

which are more exposed to biocide molecules. Afterwards, the inactivation rate of

biofilm cells by NEW slowed down considerably, probably as the extracellular matrix

General Discussion

208

physically hinders diffusion and present many reactive sites for biocide molecules

(Chen and Stewart, 1996; White, 1999; Oomori et al., 2000), and it only increase with

the exposure to 800 mg/L ACC during at least 15 min as a result of the disruption of the

biofilm matrix. Similar effects were reported by Kim et al. (2001) on Listeria

monocytogenes biofilms with the application of AEW.

High concentrations of NEW (800 mg/L ACC) were thus needed to achieve the

logarithmic reductions (LR) demanded by the European quantitative surface test of

bactericidal activity EN 13697 (≥ 4 log CFU/cm2 after 5 min of exposure), but this

would be costly and not environmentally-friendly. Nevertheless, a noticeable effect (LR

> 3 log CFU/cm2) was obtained by applying only 200 mg/L of NEW during 5 min,

which suggested that consecutive applications of low concentrations of NEW could

increase significantly the bactericidal effect against biofilms, with clear advantages in

terms of both dose and exposure time.

As expected, a double sequential application of NEW at low concentrations was

enough to achieve the LR higher than 4 log CFU/cm2 after 5 min of exposure each.

However, this procedure could involve a risk of adaptation to EW, as previously

observed after the repeated use of a same chlorinated disinfectant (Braoudaki and

Hilton, 2004; Bridier et al., 2011a; Lundén et al., 2003; Maalej et al., 2006; Sanderson

and Stewart, 1997). Therefore, it seemed convenient that sequential applications were

based on the use of different biocides, preferably with different cellular targets.

The potential of combining NEW with benzalkonium chloride (BAC) or peracetic

acid (PAA) was thus examined. LR demanded by the European standard test EN 13697

were also reached in all cases by applying low concentrations of each biocide. But

washing was mandatory to prevent cross-reaction between them, so operation time

increased and wash water would require additional biocidal processes (e.g., heat, UV

radiation, chlorination, ozonation) to kill dispersed cells during washing (Casani and

Knøchel, 2002; Fähnrich et al., 1998).

Alternatively, the efficacy of commercial essential oils (EOs) against biofilms and

planktonic counterparts of S. aureus St.1.01 was also assessed in this thesis (Chapter 5).

A high variability in the biocidal effectiveness against planktonic cells was observed

between the nineteen commercial EOs tested (anise, carrot, citronella, coriander, cumin,

General Discussion

209

Eucalyptus globulus, Eucalyptus radiata, fennel, geranium, ginger, hyssop, lemongrass,

marjoram, palmarosa, patchouli, sage, tea-tree, thyme and vetiver). This variability

could be explained by variations in the nature and concentration of active compounds,

as well as synergistic interactions between some of them (Burt, 2004; Bakkali et al.,

2008; Hyldgaard et al., 2012). Thus, phenol compounds were found to be most active,

followed by aldehydes, ketones, alcohols, ethers and hydrocarbons (Kalemba and

Kunicka, 2003).

The resistance of S. aureus to EOs increased notably as a result of biofilm formation

with respect to free-living counterparts. However, the ranking of EOs by antimicrobial

effectiveness against biofilms and planktonic cells was dissimilar. It thus seems that

factors such as the composition and architecture of the biofilm as well as the reactivity,

hydrophobicity and diffusion rate of EOs into the biofilm matrix affect significantly the

effectiveness of EOs. The eight most effective EOs against planktonic cells (thyme,

lemongrass, vetiver, citronella, cumin, geranium, palmarosa and patchouli) also reduced

the number of biofilm viable cells significantly, but none of them was able to

completely eradicate S. aureus biofilms at doses tested (0.1-8% (v/v)). It seems thus

clear that preventing biofilm formation should be targeted rather than disruption and

removal of biofilms, as suggested by Kelly et al. (2012).

Thyme oil (from Thymus vulgaris) was found to be the most effective EO against

both biofilms and planktonic cells of S. aureus, presumably by its high content in the

phenolic compounds thymol and carvacrol (Hudaib et al., 2002; Rota et al., 2008). The

use of sub-lethal doses of thyme oil prevented biofilm formation and enhanced the

efficiency of thyme oil and BAC against biofilms. The treatment of food-contact

surfaces with thyme oil would be an interesting preventive strategy. Nonetheless, a

long-term exposure to thyme oil can produce some adaptation and problems of volatility

of active compounds. Therefore, antimicrobial treatments using EOs should be based on

the rotation and combination of different EOs (or active compounds) or with other

biocides to prevent the emergence of antimicrobial-resistant strains.

211

Conclusiones Generales


213


Los resultados obtenidos en esta tesis han permitido obtener las siguientes

conclusiones:

1. S. aureus fueron detectados en una significativa proporción (~ 25%) de productos

pesqueros comercializados en España en 2008 y 2009, sobre todo cepas con

capacidad enterotoxigénica. Una cantidad significativa de los productos regulados no

cumplían los límites vigentes, y una alta proporción de los no regulados también

estaban contaminados, lo cual revela una cierta eficacia de las políticas legales para

asegurar la higiene en la industria alimentaria. Por tanto, se recomienda revisar los

programas de prerrequisitos y mejorar las prácticas de higiene en las operaciones de

manipulación y procesado, desde la captura o cultivo del pescado hasta su punto de

venta, para certificar la seguridad de los productos pesqueros consumidos en España.

Además, estos resultados cuestionan que se haya eliminado a los estafilococos

coagulasa positivos (mayormente S. aureus) como criterio microbiológico en

productos listos para consumir y listos para cocinar.

2. Una considerable variabilidad en la capacidad de adhesión y formación de

biopelículas fue encontrada entre las cepas con capacidad enterotoxigénica de S.

aureus aisladas de productos pesqueros. Está variabilidad estaba claramente

influenciada por diferentes condiciones ambientales relevantes en la industria

alimentaria como temperatura, osmolaridad y disponibilidad de nutrientes, así como

propiedades intrínsecas de cada cepa. Así, se detectó que la adhesión inicial

incrementa al aumentar la fuerza iónica del medio, mientras que la formación de

biopelículas aumenta en presencia de glucosa y, en menor medida, de cloruro sódico

y magnesio. Además, se encontraron diferencias significativas en la expresión de

ciertos genes relacionados con la formación de biopelículas entre cepas y a distintas

condiciones ambientales, lo cual sugiere que la presencia de biopelículas en plantas

de procesado de alimentos esté condicionada por la adaptación a los estreses

ambientales. En este sentido, parece que el procesado del alimento genera una

presión selectiva, por lo que cepas con una alta capacidad de formación de

biopelículas se detectan más frecuentemente en productos altamente manipulados y

procesados.


214

3. La mayoría de las cepas con capacidad enterotoxigénica mostraron una capacidad de

formación de biopelículas sobre superficies de poliestireno y acero inoxidable, así

como una resistencia a cloruro de benzalconio (BAC), ácido peracético (PAA) e

hipoclorito sódico (NaClO) mayor que la de S. aureus ATCC 6538, la cepa de

referencia usada en los métodos bactericidas estándar. Como resultado, fueron

necesarias mayores dosis de estos desinfectantes que las recomendadas por los

fabricantes para eliminar biopelículas formadas por todas las cepas sobre superficies

de contacto con el alimento. No se encontró correlación entre la concentración

mínima necesaria para erradicar biopelículas y la concentración bactericida mínima,

y las cepas mostraron un orden de clasificación distinto para cada desinfectante.

Todos estos resultados cuestionan y sugieren una revisión de los actuales métodos

bactericidas estándar (la mayoría basados en cultivos líquidos usando unas pocas

cepas, y sólo un S. aureus). Por tanto, se recomienda tomar decisiones sobre la

aplicación de desinfectantes basándose en el uso de biopelículas como sistema

experimental, analizando un amplio número de cepas, y en la simulación de las

condiciones ambientales presentes en la industria alimentaria.

4. La evaluación de estrategias de desinfección más respetuosas con el medioambiente,

basadas en la aplicación de agua electrolizada (EW) y ciertos aceites esenciales

(EOs), ha permitido obtener las siguientes conclusiones:

4.1 Apenas hubo diferencias en la actividad bactericida del EW frente a biopelículas

debido a variaciones en el pH de producción. Esto reveló que el EW neutra

(NEW) puede tener un mayor potencial como desinfectante a largo plazo que el

EW ácida debido a su menor corrosión en superficies metálicas y a su menor

toxicidad sobre los manipuladores. Sin embargo, se necesitó una alta

concentración de cloro activo para cumplir con las especificaciones demandadas

por el método cuantitativo europeo de actividad bactericida en superficie. La

aplicación secuencial doble de NEW o la aplicación secuencial de NEW con

desinfectantes clásicos (BAC o PAA) resultó en una prometedora alternativa

para eliminar S. aureus de instalaciones de procesado de alimento en términos de

dosis y tiempo de exposición.


215

4.2 Se encontró una alta variabilidad en la eficacia de los EOs evaluados frente a

células planctónicas y biopelículas formadas en acero inoxidable, pero ninguno

fue capaz de erradicar completamente las biopelículas. El aceite de tomillo fue el

más efectivo en ambos casos, pero altas concentraciones fueron necesarias para

reducir significativamente el número de células viables en la biopelícula.

Alternativamente, la presencia de dosis sub-letales de aceite de tomillo ralentizó

el desarrollo de biopelículas, y mejoró la eficacia del aceite de tomillo y del

BAC frente a estas. Por tanto, el uso de EOs debe enfocarse a estrategias de

prevención de la formación de biopelículas más que a la erradicación de

biopelículas establecidas. Sin embargo, cierta adaptación al aceite de tomillo fue

detectada, lo cual podría ser un inconveniente este uso. Por tanto, estas

estrategias se deben basar en la rotación y combinación de diferentes EOs o con

otros biocidas para prevenir la emergencia de cepas resistentes a los

antimicrobianos.

217

General Conclusions

General Conclusions

219

General Conclusions

The results obtained in this thesis have permitted the following conclusions to be drawn:

1. A remarkable number (~ 25%) of fishery products collected from the retail sector in

Spain in 2008 and 2009 was found to be contaminated with S. aureus, mostly with

strains able to produce enterotoxins. A significant proportion of regulated products

did not comply with legal limits in force, but most contaminated products was not

subject to any regulation, which seems to reveal some effects of legal policies on the

efforts of the industry to ensure food hygiene. A revision of pre-requisite programs

leading to improve hygienic practices in handling and processing operations from

fishing or farming to retail is therefore recommended to ensure the safety of seafood

consumed in Spain. Also, these results call into question the following repeal of

coagulase positive staphylococci (mainly S. aureus) as a microbiological criterion for

ready-to-cook products and most ready-to-eat products.

2. A significant variability in adhesion and biofilm-forming properties was found

among putative enterotoxigenic S. aureus strains isolated from fishery products. This

variability was clearly influenced by environmental conditions relevant for the food

industry such as temperature, osmolarity and nutrient availability, as well as by

intrinsic properties of strains. Thus, initial adhesion was increased by high ionic

strength conditions, whereas biofilm formation was enhanced by glucose and, to a

lower extent, by sodium or magnesium chloride. Significant differences in the

expression of some biofilm-related genes were also found among strains and

environmental conditions, which suggest that the presence of biofilms in food-

processing facilities would be conditioned by the adaptation to environmental

stresses. In this sense, it seems that food processing could have applied some

selective pressure, so strains with a high biofilm-forming ability were more likely to

be found in highly handled and processed products.

3. Most putative enterotoxigenic strains showed a biofilm-forming ability on

polystyrene and stainless steel and a resistance to benzalkonium chloride (BAC),

peracetic acid (PAA) and sodium hypochlorite (NaClO) higher than that of S. aureus

ATCC 6538, the reference strain used in bactericidal standard tests. As a result,

General Conclusions

220

doses of BAC, PAA and NaClO higher than those recommended by manufacturers

were needed to remove biofilms of all strains from food-contact surfaces. No

correlation was found between minimum biofilm eradication concentration and

minimum bactericidal concentration either, and strains were ranked differently

according to biofilm resistance to each disinfectant. All these results question and

call for a revision of current bactericidal standard tests (mostly suspension-based

using a few strains, and only one S. aureus). Therefore, decision-making on the

application of disinfectants must be based on a widespread use of biofilms as

experimental systems, testing a relatively wide number of strains, and on the

simulation of environmental conditions found in the food industry.

4. The assessment of disinfection strategies more environmentally-friendly, based on

the application of electrolyzed water (EW) and some essential oils (EOs), have

allowed to obtain the following conclusions:

4.1. Hardly differences were noticed in the bactericidal activity of EW against

biofilms as a result of variations in the pH of production. This revealed that

neutral EW (NEW) would have a higher potential than acidic EW for long-term

use as an antimicrobial in the food industry due to a lower corrosiveness to metal

surfaces and a lower toxicity to handlers. However, a high available chlorine

concentration was needed to comply with specifications demanded by the

European quantitative surface test of bactericidal activity. A double sequential

application of NEW or a sequential application of NEW and a classical

disinfectant (either BAC or PAA) was found to be a promising alternative to

remove S. aureus from food-processing facilities in terms of dose and exposure

time.

4.2. A highly variable effectiveness against planktonic cells and biofilms formed on

stainless steel was found among EOs tested, but no EO removed biofilms

completely. Thyme oil was found to be the most effective in both cases, but high

concentrations were needed to reduce significantly the number of viable biofilm

cells. Alternatively, the presence of sub-lethal doses of thyme oil slowed down

biofilm formation, and enhanced the efficiency of thyme oil and BAC against

biofilms. Therefore, EOs should be aimed to be a preventive strategy against

General Conclusions

221

biofilms rather than to remove biofilms formed. However, some adaptation to

thyme oil was detected, which could be a drawback for this intended use. The

rotation and combination of different EOs or with other biocides should be

therefore employed to prevent the emergence of antimicrobial-resistant strains.

223

Bibliografía / References


225


Abee, T., Wouters, J.A., 1999. Microbial stress response in minimal processing. Int. J. Food

Microbiol. 50, 65–91.

Abrahim, A., Sergelidis, D., Kirkoudis, I., Anagnostou, V., Kaitsa-Tsiopoulou, E., Kazila, P.,

Papa, A., 2010. Isolation and antimicrobial resistance of Staphylococcus spp. in freshwater

fish and Greek marketplaces. J. Aquat. Food Prod. Technol. 19, 93–102.

Adukwu, E.C., Allen, S.C.H., Phillips, C.A., 2012. The anti-biofilm activity of lemongrass

(Cymbopogon flexuosus) and grapefruit (Citrus paradisi) essential oils against five strains

of Staphylococcus aureus. J. Appl. Microbiol. 113, 1217–1227.

Aiemsaard, J., Aiumlamai, S., Aromdee, C., Taweechaisupapong, S., Khunkitti, W., 2011. The

effect of lemongrass oil and its major components on clinical isolate mastitis pathogens

and their mechanisms of action on Staphylococcus aureus DMST 4745. Res. Vet. Sci. 91,

31–37.

Akpolat, N.O., Elçi, S., Atmaca, S., Akbayin, H., Gül, K., 2003. The effects of magnesium,

calcium and EDTA on slime production by Staphylococcus epidermidis strains. Folia

Microbiol. 48, 649–653.

Alarcón, B., Vicedo, B., Aznar, R., 2006. PCR-based procedures for detection and

quantification of Staphylococcus aureus and their application in food. J. Appl. Microbiol.

100, 352–364.

Al-Bayati, F.A., 2008. Synergistic antibacterial activity between Thymus vulgaris and

Pimpinella anisum essential oils and methanol extracts. J. Ethnopharmacol. 116, 403–406.

Al-Thawadi, S.I., Kessie, G., Dela Cruz, D., Al-Ahdal, M.N., 2003. A comparative study on the

application of 3 molecular methods in epidemiological typing of bacterial isolates using

MRSA as a prototype. Saudi Med. J. 24, 1317–1324.

Ammendolia, M.G., Di-Rosa, R., Montanaro, L., Arciola, C.R., Baldassarri, L., Di Rosa, R.,

1999. Slime production and expression of the slime-associated antigen by staphylococcal

clinical isolates. J. Clin. Microbiol. 37, 3235–3238.

Anderson, G.G., O’Toole, G. a, 2008. Innate and induced resistance mechanisms of bacterial

biofilms. Curr. Top. Microbiol. Inmunol. 322, 85–105.

Angioni, A., Barra, A., Coroneo, V., Dessi, S., Cabras, P., 2006. Chemical composition,

seasonal variability, and antifungal activity of Lavandula stoechas L. ssp. stoechas

essential oils from stem/leaves and flowers. J. Agric. Food Chem. 54, 4364–4370.

APHA-AWWA-WPCF, 1992. Standard methods for the examination of water and wastewater,

17th ed. Díaz de Santos, Madrid.

Aras, Z., Aydin, I., Kav, K., 2012. Isolation of methicillin-resistant Staphylococcus aureus from

caprine mastitis cases. Small Rumin. Res. 102, 68–73.

Arciola, C.R., Campoccia, D., Gamberini, S., Cervellati, M., Donati, E., Montanaro, L., 2002.

Detection of slime production by means of an optimised Congo red agar plate test based

on a colourimetric scale in Staphylococcus epidermidis clinical isolates genotyped for ica

locus. Biomaterials 23, 4233–4239.

Argudín, M.Á., Mendoza, M.C., Rodicio, M.R., 2010. Food poisoning and Staphylococcus

aureus enterotoxins. Toxins 2, 1751–1773.

Asao, T., Kumeda, Y., Kawai, T., Shibata, T., Oda, H., Haruki, K., Nakazawa, H., Kozaki, S.,

2003. An extensive outbreak of staphylococcal food poisoning due to low-fat milk in


226

Japan: estimation of enterotoxin A in the incriminated milk and powdered skim milk.

Epidemiol. Infect. 130, 33–40.

Astani, A., Reichling, J., Schnitzler, P., 2011. Screening for antiviral activities of isolated

compounds from essential oils. Evidence-Based Complement. Altern. Med. 2011, 253643.

Ayebah, B., Hung, Y.-C., 2005. Electrolyzed water and its corrosiveness on various surface

materials commonly found in food processing facilities. J. Food Process Eng. 28, 247–264.

Ayebah, B., Hung, Y.-C., Frank, J.F., 2005. Enhancing the bactericidal effect of electrolyzed

water on Listeria monocytogenes biofilms formed on stainless steel. J. Food Prot. 68,

1375–1380.

Baddour, M.M., Abu-El-Kheir, M.M., Fatani, A.J., 2007. Comparison of mecA polymerase

chain reaction with phenotypic methods for the detection of methicillin-resistant

Staphylococcus aureus. Curr. Microbiol. 55, 473–479.

Bagge-Ravn, D., Ng, Y., Hjelm, M., Christiansen, J., Johansen, C., Gram, L., 2003. The

microbial ecology of processing equipment in different fish industries—analysis of the

microflora during processing and following cleaning and disinfection. Int. J. Food

Microbiol. 87, 239–250.

Bakkali, F., Averbeck, S., Averbeck, D., Idaomar, M., 2008. Biological effects of essential oils-

a review. Food Chem. Toxicol. 46, 446–475.

Balaban, N., Rasooly, A., 2000. Staphylococcal enterotoxins. Int. J. Food Microbiol. 61, 1–10.

Barber, M.A., 1914. Milk poisoning due to a type of Staphylococcus albus occurring in the

udder of a cow. Philipine J. Sci. 89, 515–519.

Barnes, L.M., Lo, M.F., Adams, M.R., Chamberlain, a H., Chamberlain, H.L., 1999. Effect of

milk proteins on adhesion of bacteria to stainless steel surfaces. Appl. Environ. Microbiol.

65, 4543–4548.

Basti, A.A., Misaghi, A., Salehi, T.Z., Kamkar, A., 2006. Bacterial pathogens in fresh, smoked

and salted Iranian fish. Food Control 17, 183–188.

Becker, K., Roth, R., Peters, G., 1998. Rapid and specific detection of toxigenic Staphylococcus

aureus: use of two multiplex PCR enzyme immunoassays for amplification and

hybridization of staphylococcal enterotoxin genes, exfoliative toxin genes, and toxic shock

syndrome toxin 1 gene. J. Clin. Microbiol. 36, 2548–2553.

Beech, I.B., Sunner, J. a, Hiraoka, K., 2005. Microbe-surface interactions in biofouling and

biocorrosion processes. Int. Microbiol. 8, 157–168.

Beleneva, I.A., 2011. Incidence and characteristics of Staphylococcus aureus and Listeria

monocytogenes from the Japan and South China seas. Mar. Pollut. Bull. 62, 382–387.

Bellon-Fontaine, M.-N., Rault, J., Oss, C.J., 1996. Microbial adhesion to solvents: a novel

method to determine the electron-donor/electron-acceptor or Lewis acid-base properties of

microbial cells. Colloids Surfaces B Biointerfaces 7, 47–53.

Bergdoll, M.S., Wong, A.C.L., 2006. Staphylococcal intoxications, in: Foodborne Infections

and Intoxications. Elsevier Inc., pp. 524–552.

Bessems, E., 1998. The effect of practical conditions on the efficacy of disinfectants. Int.

Biodeterior. Biodegrad. 41, 177–183.

Bhatia, A., Zahoor, S., 2007. Staphylococcus aureus enterotoxins: A review. J. Clin. Diagnoses

Res. 1, 188–197.


227

Bien, J., Sokolova, O., Bozko, P., 2011. Characterization of virulence factors of Staphylococcus

aureus: Novel function of known virulence factors that are implicated in activation of

airway epithelial proinflammatory response. J. Pathog. 2011, 1–13.

Bokarewa, M.I., Jin, T., Tarkowski, A., 2006. Staphylococcus aureus: staphylokinase. Int. J.

Biochem. Cell Biol. 38, 504–509.

Bone, F.J., Bogie, D., Morgan-Jones, S.C., 1989. Staphylococcal food poisoning from sheep

milk cheese. Epidemiol. Infect. 103, 449–458.

Bore, E., Langsrud, S., Langsrud, Ø., Rode, T.M., Holck, A., 2007. Acid-shock responses in

Staphylococcus aureus investigated by global gene expression analysis. Microbiology 153,

2289–2303.

Bos, R., Mei, H.C., Busscher, H.J., 1999. Physico-chemistry of initial microbial adhesive

interactions-its mechanisms and methods for study. FEMS Microbiol. Rev. 23, 179–230.

Box, G.E.P., Hunter, W.G., Hunter, J.S., 1989. Estadística para investigadores: Diseño,

innovación y descubrimiento, 1st ed. Reverté, Barcelona.

Branda, S.S., Vik, S., Friedman, L., Kolter, R., 2005. Biofilms: the matrix revisited. Trends


Braoudaki, M., Hilton, A.C., 2004. Adaptive resistance to biocides in Salmonella enterica and

Escherichia coli O157 and cross-resistance to antimicrobial agents. J. Clin. Microbiol. 42,

73–78.

Brenes, A., Roura, E., 2010. Essential oils in poultry nutrition: Main effects and modes of

action. Anim. Feed Sci. Technol. 158, 1–14.

Bresee, J., Maier, K.E., Boncella, A.E., Melander, C., Feldheim, D.L., 2011. Growth inhibition

of Staphylococcus aureus by mixed monolayer gold nanoparticles. Small 7, 2027–2031.

Bridier, A., Briandet, R., Thomas, V., Dubois-Brissonnet, F., 2011a. Resistance of bacterial

biofilms to disinfectants: a review. Biofouling 27, 1017–1032.

Bridier, A., Briandet, R., Thomas, V., Dubois-Brissonnet, F., 2011b. Comparative biocidal

activity of peracetic acid, benzalkonium chloride and ortho-phthalaldehyde on 77 bacterial

strains. J. Hosp. Infect. 78, 208–213.

Bridier, A., Dubois-Brissonnet, F., Greub, G., Thomas, V., Briandet, R., Brissonnet, F.D., 2011.

Dynamics of the action of biocides in Pseudomonas aeruginosa biofilms. Antimicrob.

Agents Chemother. 55, 2648–2654.

Briñez, W.J., Roig-Sagués, A.X., Herrero, M.H., López-Pedemonte, T., Guamis, B., 2006.

Bactericidal efficacy of peracetic acid in combination with hydrogen peroxide against

pathogenic and non pathogenic strains of Staphylococcus spp., Listeria spp. and

Escherichia coli. Food Control 17, 516–521.

Budzynska, A., Wieckowska-Szakiel, M., Sadowska, B., Kalemba, D., Rózalska, B., 2011.

Antibiofilm activity of selected plant essential oils and their major components. Polish J.


Bukowski, M., Wladyka, B., Dubin, G., 2010. Exfoliative toxins of Staphylococcus aureus.

Toxins (Basel). 2, 1148–1165.

Burt, S., 2004. Essential oils: their antibacterial properties and potential applications in foods-a

review. Int. J. Food Microbiol. 94, 223–253.

Byun, D.E., Kim, S.H., Shin, J.H., Suh, S.P., Ryang, D.W., 1997. Molecular epidemiologic

analysis of Staphylococcus aureus isolated from clinical specimens. J. Korean Med. Sci.

12, 190–198.


228

Byun, M.W., Kim, J.H., Kim, D.H., Kim, H.J., Jo, C., 2007. Effects of irradiation and sodium

hypochlorite on the micro-organisms attached to a commercial food container. Food

Microbiol. 24, 544–548.

Caballero-Gómez, N., Abriouel, H., Grande, M.J., Pérez-Pulido, R., Gálvez, A., 2013.

Combined treatments of enterocin AS-48 with biocides to improve the inactivation of

methicillin-sensitive and methicillin-resistant Staphylococcus aureus planktonic and

sessile cells. Int. J. Food Microbiol. 163, 96–100.

Cao, W., Zhu, Z.W., Shi, Z.X., Wang, C.Y., Li, B.M., 2009. Efficiency of slightly acidic

electrolyzed water for inactivation of Salmonella enteritidis and its contaminated shell

eggs. Int. J. Food Microbiol. 130, 88–93.

Carmo, S.L., Dias, R.S., Linardi, V.R., Sena, J.M., Santos, A.D., Faria, E.M., Pena, E.C., Jett,

M., Heneine, L.G., 2002. Food poisoning due to enterotoxigenic strains of Staphylococcus

present in Minas cheese and raw milk in Brazil. Food Microbiol. 19, 9–14.

Casani, S., Knøchel, S., 2002. Application of HACCP to water reuse in the food industry. Food

Control 13, 315–327.

CDC, 1968. Morbidity and mortality weekly report 17, 109–110.



Cebrián, G., Sagarzazu, N., Pagán, R., Condón, S., Mañas, P., 2007. Heat and pulsed electric

field resistance of pigmented and non-pigmented enterotoxigenic strains of Staphylococcus

aureus in exponential and stationary phase of growth. Int. J. Food Microbiol. 118, 304–

311.

CEN, 2002. European standard EN 13697: Chemical disinfectants and antiseptics - Quantitative

non-porous surface test for evaluation of bactericidal and/or fungicidal activity of chemical

disinfectants used in food, industrial, domestic and institutional areas. Test method Requir.

without Mech. action (phase 2, step 1).


suspension test for evaluation of basic bactericidal activity of chemical disinfectants and

antiseptics. Test method Requir. (phase 1).


suspension test for evaluation of bactericidal activity of chemical disinfectants and

antiseptics used in food, industrial, domestic and institutional areas. Test method Requir.

(phase 2, step 1).

Cerca, N., Brooks, J.L., Jefferson, K.K., 2008. Regulation of the intercellular adhesin locus

regulator (icaR) by SarA, sigmaB, and IcaR in Staphylococcus aureus. J. Bacteriol. 190,

6530–6533.

Cha, J.O., Lee, J.K., Jung, Y.H., Yoo, J.I., Park, Y.K., Kim, B.S., Lee, Y.S., 2006. Molecular

analysis of Staphylococcus aureus isolates associated with staphylococcal food poisoning

in South Korea. J. Appl. Microbiol. 101, 864–871.

Chambers, H.F., DeLeo, F.R., 2009. Waves of resistance: Staphylococcus aureus in the

antibiotic era. Nat. Rev. Microbiol. 7, 629–641.

Chandra, J., Patel, J.D., Li, J., Zhou, G., Mukherjee, P.K., McCormick, T.S., Anderson, J.M.,

Ghannoum, M.A., 2005. Modification of surface properties of biomaterials influences the

ability of Candida albicans to form biofilms. Appl. Environ. Microbiol. 71, 8795–8801.


229

Chapman, J.S., 2003. Disinfectant resistance mechanisms, cross-resistance, and co-resistance.

Int. Biodeterior. Biodegrad. 51, 271–276.

Characklis, W.G., Marshall, K.C., 1990. Biofilms. John Wiley & Sons, Inc., New York, NY.

Chemat, F., Zill-e-Huma, Khan, M.K., 2011. Applications of ultrasound in food technology:

Processing, preservation and extraction. Ultrason. Sonochem. 18, 813–835.

Chen, G., 2003. Escherichia coli adhesion to abiotic surfaces in the presence of non-ionic

surfactants. J. Adhes. Sci. Technol. 17, 2131–2139.

Chen, T.-R., Chiou, C.-S., Tsen, H.-Y., 2004. Use of novel PCR primers specific to the genes of

staphylococcal enterotoxin G, H, I for the survey of Staphylococcus aureus strains isolated

from food-poisoning cases and food samples in Taiwan. Int. J. Food Microbiol. 92, 189–

197.

Chen, X., Li, P., Wang, X., Gu, M., Zhao, C., Sloan, A.J., Lv, H., Yu, Q., 2013. Ex vivo

antimicrobial efficacy of strong acid electrolytic water against Enterococcus faecalis

biofilm. Int. Endod. J. 46, 938–946.

Chen, X., Stewart, P.S., 1996. Chlorine penetration into artificial biofilm is limited by a

reaction−diffusion interaction. Environ. Sci. Technol. 30, 2078–2083.

Cheung, A.L., Bayer, A.S., Zhang, G., Gresham, H., Xiong, Y.-Q., 2004. Regulation of

virulence determinants in vitro and in vivo in Staphylococcus aureus. FEMS Immunol.

Med. Microbiol. 40, 1–9.

Chmielewski, R.A.N., Frank, J.F., 2006. A predictive model for heat inactivation of Listeria

monocytogenes biofilm on buna-N rubber. LWT - Food Sci. Technol. 39, 11–19.

Choi, N.-C., Park, S.-J., Lee, C.-G., Park, J.-A., Kim, S.-B., 2011. Influence of surfactants on

bacterial adhesion to metal oxide-coated surfaces. Environ. Eng. Res. 16, 219–225.

Ciftci, A., Findik, A., Onuk, E.E., Savasan, S., 2009. Detection of methicillin resistance and

slime factor production of Staphylococcus aureus in bovine mastitis. Brazilian J.

Microbiol. 40, 254–261.

CLSI, 2011. Performance standards for antimicrobial susceptibility testing; twenty-first

informational supplement. CLSI 31.

Colombari, V., Mayer, M.D.B., Laicini, Z.M., Mamizuka, E., Franco, B.D.G.M., Destro, M.T.,

Landgraf, M., 2007. Foodborne outbreak caused by Staphylococcus aureus: phenotypic

and genotypic characterization of strains of food and human sources. J. Food Prot. 70,

489–493.

Corrigan, R.M., Rigby, D., Handley, P., Foster, T.J., 2007. The role of Staphylococcus aureus

surface protein SasG in adherence and biofilm formation. Microbiology 153, 2435–2446.

Costerton, J.W., Cheng, K.J., Geesey, G.G., Ladd, T.I., Nickel, J.C., Dasgupta, M., Marrie, T.J.,

1987. Bacterial biofilms in nature and disease. Annu. Rev. Microbiol. 41, 435–464.

Costerton, J.W., Geesey, G.G., Cheng, K.J., 1978. How bacteria stick. Sci. Am. 238, 86–95.

Costerton, J.W., Lappin-Scott, H.M., 1995. Bacterial biofilms in nature and disease, in: Lappin-

Scott, H.M., Costerton, J.W. (Eds.), Microbial Biofilms. Cambridge University Press,

Cambridge, pp. 1–11.

Costerton, J.W., Lewandowski, Z., Caldwell, D.E., Korber, D.R., Lappin-scott, H.M., 1995.

Microbial biofilms. Annu. Rev. Microbiol. 49, 711–745.


230

Cramton, S., Ulrich, M., Götz, F., Döring, G., 2001. Anaerobic conditions induce expression of

polysaccharide intercellular adhesin in Staphylococcus aureus and Staphylococcus

epidermidis. Infect. Immun. 69, 4079–4085.

Cramton, S.E., Gerke, C., Schnell, N.F., Nichols, W.W., Götz, F., 1999. The intercellular

adhesion (ica) locus is present in Staphylococcus aureus and is required for biofilm

formation. Infect. Immun. 67, 5427–5433.

Cremonesi, P., Luzzana, M., Brasca, M., Morandi, S., Lodi, R., Vimercati, C., Agnellini, D.,

Caramenti, G., Moroni, P., Castiglioni, B., 2005. Development of a multiplex PCR assay

for the identification of Staphylococcus aureus enterotoxigenic strains isolated from milk

and dairy products. Mol. Cell. Probes 19, 299–305.

Cretenet, M., Even, S., Le-Loir, Y., 2011. Unveiling Staphylococcus aureus enterotoxin

production in dairy products: a review of recent advances to face new challenges. Dairy

Sci. Technol. 91, 127–150.

Cucarella, C., Solano, C., Valle, J., Amorena, B., Lasa, Í., Penadés, J.R., 2001. Bap, a

Staphylococcus aureus surface protein involved in biofilm formation. J. Bacteriol. 183,

2888–2896.

Cucarella, C., Tormo, M.Á., Knecht, E., Amorena, B., Lasa, Í., Foster, T.J., Penadés, J.R., 2002.

Expression of the biofilm-associated protein interferes with host protein receptors of

Staphylococcus aureus and alters the infective process. Infect. Immun. 70, 3180–3186.

Cucarella, C., Tormo, M.Á., Úbeda, C., Trotonda, M.P., Monzón, M., Peris, C., Amorena, B.,

Lasa, I., Penadés, J.R., 2004. Role of biofilm-associated protein bap in the pathogenesis of

bovine Staphylococcus aureus. Infect. Immun. 72, 2177–2185.

Cue, D., Lei, M.G., Luong, T.T., Kuechenmeister, L., Dunman, P.M., O’Donnell, S., Rowe, S.,

O’Gara, J.P., Lee, C.Y., 2009. Rbf promotes biofilm formation by Staphylococcus aureus

via repression of icaR, a negative regulator of icaADBC. J. Bacteriol. 191, 6363–6373.

Dack, G.M., Gary, W.E., Woolpert, O., Wiggers, H., 1930. An outbreak of food poisoning

proved to be due to a yellow hemolytic Staphylococcus. J. Prev. Med. 4, 167–175.

Daniels, R., Vanderleyden, J., Michiels, J., 2004. Quorum sensing and swarming migration in

bacteria. FEMS Microbiol. Rev. 28, 261–289.

Da-Silva, M.L., Rogério Matté, G., Germano, P.M.L., Matté, M.H., 2010. Occurrence of

pathogenic microorganisms in fish sold in São Paulo, Brazil. J. Food Saf. 30, 94–110.

Davey, M.E., O´Toole, G.A., 2000. Microbial biofilms: from ecology to molecular genetics.

Microbiol. Mol. Biol. Rev. 64, 847–867.

Davies, D.G., Geesey, G.G., 1995. Regulation of the alginate biosynthesis gene algC in

Pseudomonas aeruginosa during biofilm development in continuous culture. Appl.

Environ. Microbiol. 61, 860–867.

De-Beer, D., Srinivasan, R., Stewart, P.S., 1994. Direct measurement of chlorine penetration

into biofilms during disinfection. Appl. Environ. Microbiol. 60, 4339–4344.

De-Buyser, M.L., Janin, F., Dilasser, F., 1985. Contamination of ewe cheese with

Staphylococcus aureus: study of an outbreak of food poisoning, in: Jelkaszewicz, J. (Ed.),

The Staphylococci. Gustav Fisher Verlag, Stuttgart, pp. 677–678.

Delaquis, P.J., Stanich, K., Girard, B., Mazza, G., 2002. Antimicrobial activity of individual and

mixed fractions of dill, cilantro, coriander and eucalyptus essential oils. Int. J. Food

Microbiol. 74, 101–109.


231

DeLeo, F.R., Diep, B.A., Otto, M., 2009. Host defense and pathogenesis in Staphylococcus

aureus infections. Infect. Dis. Clin. North Am. 23, 1–17.

Denyer, S.P., Stewart, G.S.A., 1998. Mechanisms of action of disinfectants. Int. Biodeterior.

Biodegrad. 41, 261–268.

Deplano, A., Mendonça, R., Ryck, R., Struelens, M.J., 2006. External quality assessment of

molecular typing of Staphylococcus aureus isolates by a network of laboratories. J. Clin.

Microbiol. 44, 3236–3244.

Deplano, A., Schuermans, A., Eldere, J. V, Witte, W., Meugnier, H., Etienne, J., Grundmann,

H., Jonas, D., Noordhoek, G.T., Dijkstra, J., Belkum, A., Leeuwen, W., Tassios, P.T.,

Legakis, N.J., Zee, A., Bergmans, A., Blanc, D.S., Tenover, F.C., Cookson, B.C., O´Neil,

G., Struelens, M.J., 2000. Multicenter evaluation of epidemiological typing of methicillin-

resistant Staphylococcus aureus strains by repetitive-element PCR analysis. J. Clin.

Microbiol. 38, 3527–3533.

DeVita, M.D., Wadhera, R.K., Theis, M.L., Ingham, S.C., 2007. Assessing the potential of

Streptococcus pyogenes and Staphylococcus aureus transfer to foods and customers via a

survey of hands, hand-contact surfaces and food-contact surfaces at foodservice facilities.

J. Foodserv. 18, 76–79.

Deza, M.A., Araujo, M., Garrido, M.J., 2005. Inactivation of Escherichia coli, Listeria

monocytogenes, Pseudomonas aeruginosa and Staphylococcus aureus on stainless steel

and glass surfaces by neutral electrolysed water. Lett. Appl. Microbiol. 40, 341–346.

Dinges, M.M., Orwin, P.M., Schlievert, P.M., 2000. Exotoxins of Staphylococcus aureus. Clin.

Microbiol. Rev. 13, 16–34.

Di-Pasqua, R., Betts, G., Hoskins, N., Edwards, M., Ercolini, D., Mauriello, G., 2007.

Membrane toxicity of antimicrobial compounds from essential oils. J. Agric. Food Chem.

55, 4863–4870.

Di-Pasqua, R., Mamone, G., Ferranti, P., Ercolini, D., Mauriello, G., 2010. Changes in the

proteome of Salmonella enterica serovar Thompson as stress adaptation to sublethal

concentrations of thymol. Proteomics 10, 1040–1049.

Donlan, R., Costerton, J., 2002. Biofilms: survival mechanisms of clinically relevant

microorganisms. Clin. Microbiol. Rev. 15, 167–193.

Donlan, R.M., 2002. Biofilms: microbial life on surfaces. Emerg. Infect. Dis. 8, 881–890.

Dufour, D., Leung, V., Lévesque, C.M., 2012. Bacterial biofilm: structure, function, and

antimicrobial resistance. Endod. Top. 22, 2–16.

Dunne, W.M., 2002. Bacterial Adhesion : Seen Any Good Biofilms Lately ? Clin. Microbiol.

Rev. 15, 155–166.

EFSA, 2006. The Community summary reports on trends and sources of zoonoses, zoonotic

agents, antimicrobial resistance and foodborne outbreaks in the European Union in 2005.

EFSA J. 94, 1–288.

EFSA, 2007. The Community summary report on trends and sources of zoonoses, zoonotic

agents, antimicrobial resistance and foodborne outbreaks in the European Union in 2006.

EFSA J. 130, 1–352.

EFSA, 2009a. The Community summary report on food-borne outbreaks in the European Union

in 2007. EFSA J. 271, 1–102.


232

EFSA, 2009b. Joint scientific report of ECDC, EFSA and EMEA on meticillin resistant

Staphylococcus aureus (MRSA) in livestock, companion animals and food. EFSA Sci.

Rep. 301, 1–10.

EFSA, 2010. The Community summary report on trends and sources of zoonoses and zoonotic

agents and food-borne outbreaks in the European Union in 2008. EFSA J. 8, 1–368.

EFSA, 2011. The European Union summary report on trends and sources of zoonoses and

zoonotic agents and food-borne outbreaks in 2009. EFSA J. 9, 1–378.

EFSA, 2012. The European Union summary report on trends and sources of zoonoses, zoonotic

agents and food-borne outbreaks in 2010. EFSA J. 10, 1–442.

Ehlbeck, J., Schnabel, U., Polak, M., Winter, J., Woedtke, T., Brandenburg, R., Hagen, T.,

Weltmann, K.-D., 2011. Low temperature atmospheric pressure plasma sources for

microbial decontamination. J. Phys. D. Appl. Phys. 44, 013002.

Eisenberg, M.S., Gaarslev, K., Brown, W., Horwitz, D., 1975. Hill staphylococcal food

poisoning aboard a commercial aircraft. Lancet 2, 595–599.

Elias, S., Banin, E., 2012. Multi-species biofilms: living with friendly neighbors. FEMS


Ennajar, M., Bouajila, J., Lebrihi, A., Mathieu, F., Savagnac, A., Abderraba, M., Raies, A.,

Romdhane, M., 2010. The influence of organ, season and drying method on chemical

composition and antioxidant and antimicrobial activities of Juniperus phoenicea L.

essential oils. J. Sci. Food Agric. 90, 462–470.

Ertürk, Ö., 2010. Antibacterial and antifungal effects of alcoholic extracts of 41 medicinal

plants growing in Turkey. Czech J. Food Sci. 28, 53–60.

EUCAST, 2003. Determination of minimum inhibitory concentrations (MICs) of antibacterial

agents by broth dilution. Eur. J. Clin. Microbiol. Infect. Dis. 1–7.

EUCAST, 2011. Expert rules in antimicrobial susceptibility testing. Eur. Soc. Clin. Microbiol.

Infect. Dis. 1–34.

Eurostat, 2007. Fishery statistics: Data 1990-2006, 2007th ed. Eurostat Pocketbooks,

Luxembourg.

Eurostat, 2010. Fishery statistics: Data 1995-2008, 2009th ed. Eurostat Pocketbooks,

Luxembourg.

Evenson, M.L., Hinds, M.W., Bernstein, R.S., Bergdoll, M.S., 1988. Estimation of human dose

of staphylococcal enterotoxin A from a large outbreak of staphylococcal food poisoning

involving chocolate milk. Int. J. Food Microbiol. 7, 311–316.

Fähnrich, A., Mavrov, V., Chmiel, H., 1998. Membrane processes for water reuse in the food

industry. Desalination 119, 213–216.

FAO, 2012. Fishery and aquaculture statistics, 2010th ed. Food and Agriculture Organization of

the United Nations, Rome.

FDA, 1992. Staphylococcus aureus, in: Lampel, K.A., Al-Khaldi, S., Cahill, S.M. (Eds.), Bad

Bug Book. Foodborne Pathogenic Microorganisms and Natural Toxins Handbook. Center

for Food Safety and Applied Nutrition, pp. 87–92.

FDA, 2012. Staphylococcus aureus, in: Lampel, K.A., Al-Khaldi, S., Cahill, S.M. (Eds.), Bad

Bug Book. Foodborne Pathogenic Microorganisms and Natural Toxins Handbook. Second

Edition. Center for Food Safety and Applied Nutrition, pp. 87–92.


233

Fenner, D.C., Bürge, B., Kayser, H.P., Wittenbrink, M.M., 2006. The anti-microbial activity of

electrolysed oxidizing water against microorganisms relevant in veterinary medicine. J.

Vet. Med. 53, 133–137.

Fitz-James, I., Botteldoorn, N., Veld, P., Dierick, C., 2008. Joined investigation of a large

outbreak involving Staphylococcus aureus, in: Proceeding FoodMicro 2008. Aberdeen

(UK), september 2.

Fitzpatrick, F., Humphreys, H., O’Gara, J.P., 2005. The genetics of staphylococcal biofilm

formation--will a greater understanding of pathogenesis lead to better management of

device-related infection? Clin. Microbiol. Infect. 11, 967–973.

Flemming, H.-C., Neu, T.R., Wozniak, D.J., 2007. The EPS matrix: the “house of biofilm

cells”. J. Bacteriol. 189, 7945–7947.

Forsythe, S.J., Hayes, P.R., 1998. Food hygiene, microbiology and HACCP, 3rd ed. Aspen.

Foster, T.J., Höök, M., 1998. Surface protein adhesins of Staphylococcus aureus. Trends


Freeman, D.J., Falkiner, F.R., Keane, C.T., 1989. New method for detecting slime production

by coagulase negative staphylococci. J. Clin. Pathol. 42, 872–874.

Fu, Y., Zu, Y., Chen, L., Shi, X., Wang, Z., Sun, S., Efferth, T., 2007. Antimicrobial activity of

clove and rosemary essential oils alone and in combination. Phyther. Res. 21, 989–994.

Fueyo, J.M., Martín, M.C., González-Hevia, M.A., Mendoza, M.C., 2001. Enterotoxin

production and DNA fingerprinting in Staphylococcus aureus isolated from human and

food samples. Relations between genetic types and enterotoxins. Int. J. Food Microbiol.

67, 139–145.

Fux, C.A., Costerton, J.W., Stewart, P.S., Stoodley, P., 2005. Survival strategies of infectious

biofilms. Trends Microbiol. 13, 34–40.

Fux, C.A., Wilson, S., Stoodley, P., 2004. Detachment characteristics and oxacillin resistance of

Staphyloccocus aureus biofilm emboli in an in vitro catheter infection model. Infecti. J.

Bacteriol. 186, 4486–4491.

García, P., Madera, C., Martínez, B., Rodríguez, A., 2007. Biocontrol of Staphylococcus aureus

in curd manufacturing processes using bacteriophages. Int. Dairy J. 17, 1232–1239.

García, P., Martínez, B., Obeso, J.M., Rodríguez, A., 2008. Bacteriophages and their application

in food safety. Lett. Appl. Microbiol. 47, 479–85.

García, P., Rodríguez, L., Rodríguez, A., Martínez, B., 2010. Food biopreservation: promising

strategies using bacteriocins, bacteriophages and endolysins. Trends Food Sci. Technol.

21, 373–382.

Garrido, V., Vitas, A.I., García-Jalón, I., 2009. Survey of Listeria monocytogenes in ready-to-

eat products: Prevalence by brands and retail establishments for exposure assessment of

listeriosis in Northern Spain. Food Control 20, 986–991.

Garzoni, C., Kelley, W.L., 2009. Staphylococcus aureus: new evidence for intracellular

persistence. Trends Microbiol. 17, 59–65.

Gaulin, C., Lê, M., Shum, M., Fong, D., 2011. Disinfectants and sanitizers for use on food

contact surfaces. Natl. Collab. Cent. Environ. Heal. 1–15.

Geoghegan, J.A., Corrigan, R.M., Gruszka, D.T., Speziale, P., Gara, J.P.O., Potts, J.R., Foster,

T.J., 2010. Role of surface protein SasG in biofilm formation by Staphylococcus aureus. J.

Bacteriol. 192, 5663–5673.


234

George, D.R., Smith, T.J., Shiel, R.S., Sparagano, O.A.E., Guy, J.H., 2009. Mode of action and

variability in efficacy of plant essential oils showing toxicity against the poultry red mite,

Dermanyssus gallinae. Vet. Parasitol. 161, 276–282.

Gerke, C., Kraft, A., Sübmuth, R., Schweitzer, O., Götz, F., 1998. Characterization of the N-

acetylglucosaminyltransferase activity involved in the biosynthesis of the Staphylococcus

epidermidis polysaccharide intercellular adhesin. J. Biol. Chem. 273, 18586–18593.

Gertz, S., Engelmann, S., Schmid, R., Ziebandt, a K., Tischer, K., Scharf, C., Hacker, J.,

Hecker, M., 2000. Characterization of the sigma(B) regulon in Staphylococcus aureus. J.

Bacteriol. 182, 6983–6991.

Giaouris, E., Chapot-Chartier, M.-P., Briandet, R., 2009. Surface physicochemical analysis of

natural Lactococcus lactis strains reveals the existence of hydrophobic and low charged

strains with altered adhesive properties. Int. J. Food Microbiol. 131, 2–9.

Gibson, H., Taylor, J.H., Hall, K.E., Holah, J.T., 1999. Effectiveness of cleaning techniques

used in the food industry in terms of the removal of bacterial biofilms. J. Appl. Microbiol.

87, 41–48.

Gilbert, P., Allison, D., McBain, A., 2002. Biofilms in vitro and in vivo: do singular

mechanisms imply cross-resistance? J. Appl. Microbiol. Symp. Suppl. 92, 98–110.

Gilbert, P., Moore, L.E., 2005. Cationic antiseptics: diversity of action under a common epithet.

J. Appl. Microbiol. 99, 703–715.

Gómez-López, V.M., Ragaert, P., Debevere, J., Devlieghere, F., 2007. Pulsed light for food

decontamination: a review. Trends Food Sci. Technol. 18, 464–473.

Goñi, P., López, P., Sánchez, C., Gómez-Lus, R., Becerril, R., Nerín, C., 2009. Antimicrobial

activity in the vapour phase of a combination of cinnamon and clove essential oils. Food

Chem. 116, 982–989.

González-Rodríguez, M.-N., Sanz, J.-J., Santos, J.-A., Otero, A., García-López, M.-L., 2002.

Numbers and types of microorganisms in vacuum-packed cold-smoked freshwater fish at

the retail level. Int. J. Food Microbiol. 77, 161–168.

Gormley, F.J., Little, C.L., Rawal, N., Gillespie, I.A., Lebaigue, S., Adak, G.K., 2011. A 17-

year review of foodborne outbreaks: describing the continuing decline in England and

Wales (1992-2008). Epidemiol. Infect. 139, 688–699.

Gottenbos, B., van der Mei, H.C., Klatter, F., Nieuwenhuis, P., Busscher, H.J., 2002. In vitro

and in vivo antimicrobial activity of covalently coupled quaternary ammonium silane

coatings on silicone rubber. Biomaterials 23, 1417–1423.

Götz, F., Bannerman, T., Schleifer, K., 2006. The genera Staphylococcus and Macrococcus.

Prokaryotes 4, 5–75.

Graham, D.M., Pariza, M.W., Glaze, W.H., Erdman, J.W., Newell, G.W., Borzelleca, J.F.,

1997. Use of ozone for food processing. Food Technol. 51, 72–76.

Griffin, S.G., Wyllie, S.G., Markham, J.L., Leach, D.N., 1999. The role of structure and

molecular properties of terpenoids in determining their antimicrobial activity. Flavour

Fragr. J. 14, 322–332.

Gross, M., Cramton, S.E., Go, F., Peschel, A., 2001. Key role of teichoic acid net charge in

Staphylococcus aureus colonization of artificial surfaces. Infect. Immun. 69, 3423–3426.

Guan, W., Li, S., Yan, R., Tang, S., Quan, C., 2007. Comparison of essential oils of clove buds

extracted with supercritical carbon dioxide and other three traditional extraction methods.

Food Chem. 101, 1558–1564.


235

Gündoğan, N., Citak, S., Turan, E., 2006. Slime production, DNase activity and antibiotic

resistance of Staphylococcus aureus isolated from raw milk, pasteurised milk and ice

cream samples. Food Control 17, 389–392.

Guentzel, J.L., Liang Lam, K., Callan, M.A., Emmons, S.A., Dunham, V.L., 2008. Reduction of

bacteria on spinach, lettuce, and surfaces in food service areas using neutral electrolyzed

oxidizing water. Food Microbiol. 25, 36–41.

Gutiérrez, D., Delgado, S., Vázquez-Sánchez, D., Martínez, B., Cabo, M.L., Rodríguez, A.,

Herrera, J.J., García, P., 2012. Incidence of Staphylococcus aureus and analysis of

associated bacterial communities on food industry surfaces. Appl. Environ. Microbiol. 78,

8547–8554.

Guzel-Seydim, Z.B., Greene, A.K., Seydim, A.C., 2004. Use of ozone in the food industry.

LWT - Food Sci. Technol. 37, 453–460.

Haas, C.J.C., Veldkamp, K.E., Peschel, A., Weerkamp, F., Wamel, W.J.B., Heezius, E.C.J.M.,

Poppelier, M.J.J.G., Kessel, K.P.M., Strijp, J.A.G., 2004. Chemotaxis inhibitory protein of

Staphylococcus aureus, a bacterial antiinflammatory agent. J. Exp. Med. 199, 687–695.

Habimana, O., Le-Goff, C., Juillard, V., Bellon-Fontaine, M.-N., Buist, G., Kulakauskas, S.,

Briandet, R., 2007. Positive role of cell wall anchored proteinase PrtP in adhesion of

lactococci. BMC Microbiol. 7, 36.

Haggar, A., Ehrnfelt, C., Holgersson, J., Flock, J.-I., 2004. The extracellular adherence protein

from Staphylococcus aureus inhibits neutrophil binding to endothelial cells. Infect.

Immun. 72, 6164–6167.

Hall-Stoodley, L., Costerton, J.W., Stoodley, P., 2004. Bacterial biofilms: from the natural

environment to infectious diseases. Nat. Rev. 2, 95–108.

Hall-Stoodley, L., Stoodley, P., 2005. Biofilm formation and dispersal and the transmission of

human pathogens. Trends Microbiol. 13, 7–10.

Hammer, K.A., Carson, C.F., Riley, T. V, 1999. Antimicrobial activity of essential oils and

other plant extracts. J. Appl. Microbiol. 86, 985–990.

Handke, L.D., Conlon, K.M., Slater, S.R., Elbaruni, S., Fitzpatrick, F., Humphreys, H., Giles,

W.P., Rupp, M.E., Fey, P.D., O´Gara, J.P., 2004. Genetic and phenotypic analysis of

biofilm phenotypic variation in multiple Staphylococcus epidermidis isolates. J. Med.

Microbiol. 53, 367–374.

Hendry, E.R., Worthington, T., Conway, B.R., Lambert, P.A., 2009. Antimicrobial efficacy of

eucalyptus oil and 1,8-cineole alone and in combination with chlorhexidine digluconate

against microorganisms grown in planktonic and biofilm cultures. J. Antimicrob.

Chemother. 64, 1219–1225.

Hennekinne, J.-A., Brun, V., De Buyser, M.-L., Dupuis, A., Ostyn, A., Dragacci, S., 2009.

Innovative application of mass spectrometry for the characterization of staphylococcal

enterotoxins involved in food poisoning outbreaks. Appl. Environ. Microbiol. 75, 882–

884.

Hennekinne, J.-A., De-Buyser, M.-L., Dragacci, S., 2012. Staphylococcus aureus and its food

poisoning toxins: characterization and outbreak investigation. FEMS Microbiol. Rev. 36,

815–836.

Herrera, F.C., Santos, J.A., Otero, A., García-López, M.-L., 2006. Occurrence of foodborne

pathogenic bacteria in retail prepackaged portions of marine fish in Spain. J. Appl.

Microbiol. 100, 527–536.


236

Herrera, J.J.R., Cabo, M.L., González, A., Pazos, I., Pastoriza, L., 2007. Adhesion and

detachment kinetics of several strains of Staphylococcus aureus subsp. aureus under three

different experimental conditions. Food Microbiol. 24, 585–591.

Høiby, N., Bjarnsholt, T., Givskov, M., Molin, S., Ciofu, O., 2010. Antibiotic resistance of

bacterial biofilms. Int. J. Antimicrob. Agents 35, 322–332.

Horiba, N., Hiratsuka, K., Onoe, T., Yoshida, T., Suzuki, K., Matsumoto, T., Nakamura, H.,

1999. Bactericidal effect of electrolyzed neutral water on bacteria isolated from infected

root canals. Oral surgery, Oral Med. Oral Pathol. 87, 83–87.

Houston, P., Rowe, S.E., Pozzi, C., Waters, E.M., Gara, J.P.O., 2011. Essential role for the

major autolysin in the fibronectin-binding protein-mediated Staphylococcus aureus

biofilm phenotype. Infect. Immun. 79, 1153–1165.

Huang, Y.-R., Hung, Y.-C., Hsu, S.-Y., Huang, Y.-W., Hwang, D.-F., 2008. Application of

electrolyzed water in the food industry. Food Control 19, 329–345.

Hudaib, M., Speroni, E., Di Pietra, A.M., Cavrini, V., 2002. GC/MS evaluation of thyme

(Thymus vulgaris L.) oil composition and variations during the vegetative cycle. J. Pharm.

Biomed. Anal. 29, 691–700.

Hunt, S.M., Werner, E.M., Huang, B., Hamilton, M.A., Stewart, P.S., 2004. Hypothesis for the

role of nutrient starvation in biofilm detachment hypothesis for the role of nutrient

starvation in biofilm detachment. Appl. Environ. Microbiol. 70, 7418–7425.

Hussain, A.I., Anwar, F., Nigam, P.S., Ashraf, M., Gilani, A.H., 2010. Seasonal variation in

content, chemical composition and antimicrobial and cytotoxic activities of essential oils

from four Mentha species. J. Sci. Food Agric. 90, 1827–1836.

Hyldgaard, M., Mygind, T., Meyer, R.L., 2012. Essential oils in food preservation: mode of

action, synergies, and interactions with food matrix components. Front. Microbiol. 3, 12.

Ikeda, T., Tamate, N., Yamaguchi, K., Makino, S., 2005. Mass outbreak of food poisoning

disease caused by small amounts of staphylococcal enterotoxins A and H. Appl. Environ.

Microbiol. 71, 2793–2795.

Issa-Zacharia, A., Kamitani, Y., Morita, K., Iwasaki, K., 2010. Sanitization potency of slightly

acidic electrolyzed water against pure cultures of Escherichia coli and Staphylococcus

aureus, in comparison with that of other food sanitizers. Food Control 21, 740–745.

Issa-Zacharia, A., Kamitani, Y., Tiisekwa, A., Morita, K., Iwasaki, K., 2010. In vitro

inactivation of Escherichia coli, Staphylococcus aureus and Salmonella spp. using slightly

acidic electrolyzed water. J. Biosci. Bioeng. 110, 308–313.

Izano, E.A., Amarante, M.A., Kher, W.B., Kaplan, J.B., 2008. Differential roles of poly-N-

acetylglucosamine surface polysaccharide and extracellular DNA in Staphylococcus

aureus and Staphylococcus epidermidis biofilms. Appl. Environ. Microbiol. 74, 470–476.

Jablonski, L.M., Bohach, G.A., 2001. Staphylococcus aureus, in: Beuchat, L., Doyle, M.,

Montville, T. (Eds.), Fundamentals of Food Microbiology. American Society for

Microbiology, Washington DC, pp. 410–434.

Jain, A., Agarwal, A., 2009. Biofilm production, a marker of pathogenic potential of colonizing

and commensal staphylococci. J. Microbiol. Methods 76, 88–92.

Jay, J.M., Loessner, M.J., Golden, D.A., 2005. Staphylococcal gastroenteritis, in: Jay, J.M.,

Loessner, M.J., Golden, D.A. (Eds.), Modern Food Microbiology. Springer Science, New

York, NY, pp. 545–566.


237

Jefferson, K.K., Cramton, S.E., Götz, F., Pier, G.B., 2003. Identification of a 5-nucleotide

sequence that controls expression of the ica locus in Staphylococcus aureus and

characterization of the DNA-binding properties of IcaR. Mol. Microbiol. 48, 889–899.

Johnson, E.A., Nelson, J.H., Johnson, M., 1990. Microbiological safety of cheese made from

heat-treated milk II. J. Food Prot. 53, 519–540.

Jongerius, I., Köhl, J., Pandey, M.K., Ruyken, M., Kessel, K.P.M., Strijp, J.A.G., Rooijakkers,

S.H.M., 2007. Staphylococcal complement evasion by various convertase-blocking

molecules. J. Exp. Med. 204, 2461–2471.

Jongerius, I., Puister, M., Wu, J., Ruyken, M., Strijp, J.A.G., Rooijakkers, S.H.M., 2010.

Staphylococcal complement inhibitor modulates phagocyte responses by dimerization of

convertases. J. Immunol. 184, 420–425.

Kalemba, D., Kunicka, A., 2003. Antibacterial and antifungal properties of essential oils. Curr.

Med. Chem. 10, 813–829.

Kaplan, J.B., Izano, E.A., Gopal, P., Karwacki, M.T., Kim, S., Bose, J.L., Bayles, K.W., 2012.

Low levels of β-lactam antibiotics induce extracellular DNA release and biofilm formation

in Staphylococcus aureus. MBio 3, e00198–12.

Kavanaugh, N.L., Ribbeck, K., 2012. Selected antimicrobial essential oils eradicate

Pseudomonas spp. and Staphylococcus aureus biofilms. Appl. Environ. Microbiol. 78,

4057–4061.

Kelly, D., McAuliffe, O., Ross, R.P., Coffey, A., 2012. Prevention of Staphylococcus aureus

biofilm formation and reduction in established biofilm density using a combination of

phage K and modified derivatives. Lett. Appl. Microbiol. 54, 286–291.

Kérouanton, A., Hennekinne, J.A., Letertre, C., Petit, L., Chesneau, O., Brisabois, A., De

Buyser, M.L., 2007. Characterization of Staphylococcus aureus strains associated with

food poisoning outbreaks in France. Int. J. Food Microbiol. 115, 369–375.

Khadre, M., Yousef, A., Kim, J., 2001. Microbiological aspects of ozone applications in food: a

review. J. Food Sci. 66, 1242–1252.

Kim, C., Hung, Y.-C., Brackett, R.E., Frank, J.F., 2001. Inactivation of Listeria monocytogenes

biofilms by electrolyzed oxidizing water. J. Food Process. Preserv. 25, 91–100.

Kim, C., Hung, Y.-C., Brackett, R.E., Lin, C.-S., 2003. Efficacy of electrolyzed oxidizing water

in inactivating Salmonella on alfalfa seeds and sprouts. J. Food Prot. 66, 208–214.

Kitamoto, M., Kito, K., Niimi, Y., Shoda, S., Takamura, A., Hiramatsu, T., Akashi, T., Yokoi,

Y., Hirano, H., Hosokawa, M., Yamamoto, A., Agata, N., Hamajima, N., 2009. Food

poisoning by Staphylococcus aureus at a university festival. Jpn. J. Infect. Dis. 62, 242–

243.

Kitis, M., 2004. Disinfection of wastewater with peracetic acid: a review. Environ. Int. 30, 47–

55.

Knetsch, M.L.W., Koole, L.H., 2011. New strategies in the development of antimicrobial

coatings: the example of increasing usage of silver and silver nanoparticles. Polymers

(Basel). 3, 340–366.

Kogan, G., Sadovskaya, I., Chaignon, P., Chokr, A., Jabbouri, S., 2006. Biofilms of clinical

strains of Staphylococcus that do not contain polysaccharide intercellular adhesin. FEMS

Microbiol. Lett. 255, 11–16.

Koseki, S., Yoshida, K., Kamitani, Y., Isobe, S., Itoh, K., 2004. Effect of mild heat pre-

treatment with alkaline electrolyzed water on the efficacy of acidic electrolyzed water


238

against Escherichia coli O157:H7 and Salmonella on Lettuce. Food Microbiol. 21, 559–

566.

Kumar, C.G., Anand, S.K., 1998. Significance of microbial biofilms in food industry: a review.

Int. J. Food Microbiol. 42, 9–27.

Kumar, R., Surendran, P.K., Thampuran, N., 2009. Detection and characterization of virulence

factors in lactose positive and lactose negative Salmonella serovars isolated from seafood.

Food Control 20, 376–380.

Labrie, S.J., Samson, J.E., Moineau, S., 2010. Bacteriophage resistance mechanisms. Nat. Rev.


Lamers, R.P., Muthukrishnan, G., Castoe, T.A., Tafur, S., Cole, A.M., Parkinson, C.L., 2012.

Phylogenetic relationships among Staphylococcus species and refinement of cluster groups

based on multilocus data. BMC Evol. Biol. 12, 171.

Langsrud, S., Sidhu, M.S., Heir, E., Holck, A.L., 2003. Bacterial disinfectant resistance—a

challenge for the food industry. Int. Biodeterior. Biodegrad. 51, 283–290.

Langsrud, S., Sundheim, G., Holck, A.L., 2004. Cross-resistance to antibiotics of Escherichia

coli adapted to benzalkonium chloride or exposed to stress-inducers. J. Appl. Microbiol.

96, 201–208.

Lappin, E., Ferguson, A.J., 2009. Gram-positive toxic shock syndromes. Lancet Infect. Dis. 9,

281–290.

Lasa, I., Penadés, J.R., 2006. Bap: a family of surface proteins involved in biofilm formation.

Res. Microbiol. 157, 99–107.

La-Storia, A., Ercolini, D., Marinello, F., Di-Pasqua, R., Villani, F., Mauriello, G., 2011.

Atomic force microscopy analysis shows surface structure changes in carvacrol-treated

bacterial cells. Res. Microbiol. 162, 164–172.

Lawrynowicz-Paciorek, M., Kochman, M., Piekarska, K., Grochowska, A., Windyga, B., 2007.

The distribution of enterotoxin and enterotoxin-like genes in Staphylococcus aureus

strains isolated from nasal carriers and food samples. Int. J. Food Microbiol. 117, 319–

323.

Lebert, I., Leroy, S., Talon, R., 2007. Effect of industrial and natural biocides on spoilage,

pathogenic and technological strains grown in biofilm. Food Microbiol. 24, 281–287.

Lee, J.H., 2003. Methicillin (oxacillin)-resistant Staphylococcus aureus strains isolated from

major food animals and their potential transmission to humans. Appl. Environ. Microbiol.

69, 6489–6494.

Le-Loir, Y., Baron, F., Gautier, M., 2003. Staphylococcus aureus and food poisoning. Genet.

Mol. Res. 2, 63–76.

Len, S. V, Hung, Y.C., Erickson, M., Kim, C., 2000. Ultraviolet spectrophotometric

characterization and bacterial properties of electrolyzed oxidizing water as influenced by

amperage and pH. J. Food Prot. 63, 1534–1537.

Lequette, Y., Boels, G., Clarisse, M., Faille, C., 2010. Using enzymes to remove biofilms of

bacterial isolates sampled in the food-industry. Biofouling 26, 421–431.

Levine, W.C., Bennett, R.W., Choi, Y., Henning, K.J., Rager, J.R., Hendricks, K. a, Hopkins,

D.P., Gunn, R.A., Griffin, P.M., 1996. Staphylococcal food poisoning caused by imported

canned mushrooms. J. Infect. Dis. 173, 1263–1267.

Lewis, K., 2010. Persister cells. Annu. Rev. Microbiol. 64, 357–372.


239

Li, W.-R., Xie, X.-B., Shi, Q.-S., Duan, S.-S., Ouyang, Y.-S., Chen, Y.-B., 2011. Antibacterial

effect of silver nanoparticles on Staphylococcus aureus. BioMetals 24, 135–141.

Liao, L.B., Chen, W.M., Xiao, X.M., 2007. The generation and inactivation mechanism of

oxidation–reduction potential of electrolyzed oxidizing water. J. Food Eng. 78, 1326–

1332.

Lim, Y., Jana, M., Luong, T.T., Lee, C.Y., 2004. Control of glucose-and NaCl-induced biofilm

formation by rbf in Staphylococcus aureus. J. Bacteriol. 186, 722–729.

Liu, C., Duan, J., Su, Y.-C., 2006. Effects of electrolyzed oxidizing water on reducing Listeria

monocytogenes contamination on seafood processing surfaces. Int. J. Food Microbiol. 106,

248–253.

Livermore, D.M., 2000. Antibiotic resistance in staphylococci. Int. J. Antimicrob. Agents 16, 3–

10.

Loc-Carrillo, C., Abedon, S.T., 2011. Pros and cons of phage therapy. Bacteriophage 1, 111–

114.

Lowy, F.D., 1998. Staphylococcus aureus infections. N. Engl. J. Med. 339, 520–532.

Lu, T.K., Collins, J.J., 2007. Dispersing biofilms with engineered enzymatic bacteriophage.

Proc. Natl. Acad. Sci. U. S. A. 104, 11197–11202.

Lu, T.K., Koeris, M.S., 2011. The next generation of bacteriophage therapy. Curr. Opin.

Microbiol. 14, 524–531.

Lundén, J., Autio, T., Markkula, A., Hellström, S., Korkeala, H., 2003. Adaptive and cross-

adaptive responses of persistent and non-persistent Listeria monocytogenes strains to

disinfectants. Int. J. Food Microbiol. 82, 265–272.

Luppens, S.B.I., Reij, M.W., Heijden, R.W.L. Van Der, Rombouts, F.M., Abee, T., 2002.

Development of a standard test to assess the resistance of Staphylococcus aureus biofilm

cells to disinfectants. Appl. Environ. Microbiol. 68, 4194–4200.

Lv, F., Liang, H., Yuan, Q., Li, C., 2011. In vitro antimicrobial effects and mechanism of action

of selected plant essential oil combinations against four food-related microorganisms.

Food Res. Int. 44, 3057–3064.

Maalej, S., Dammak, I., Dukan, S., 2006. The impairment of superoxide dismutase coordinates

the derepression of the PerR regulon in the response of Staphylococcus aureus to HOCl

stress. Microbiology 152, 855–861.

Madsen, J.S., Burmølle, M., Hansen, L.H., Sørensen, S.J., 2012. The interconnection between

biofilm formation and horizontal gene transfer. FEMS Immunol. Med. Microbiol. 65, 183–

195.

Mafu, A.A., Plumety, C., Deschênes, L., Goulet, J., 2011. Adhesion of pathogenic bacteria to

food contact surfaces: influence of pH of culture. Int. J. Microbiol. 2011, 1–10.

Mahmoud, B.S.M., Yamazaki, K., Miyashita, K., Shin, I.I., Suzuki, T., 2006. A new technology

for fish preservation by combined treatment with electrolyzed NaCl solutions and essential

oil compounds. Food Chem. 99, 656–662.

Maillard, J.Y., 2002. Bacterial target sites for biocide action. J. Appl. Microbiol. Symp. Suppl.

92, 16S–27S.

Maira-Litrán, T., Kropec, A., Abeygunawardana, C., Joyce, J., Mark III, G., Goldmann, D.A.,

Pier, G.B., 2002. Immunochemical properties of the staphylococcal poly-N-

acetylglucosamine surface polysaccharide. Infect. Immun. 70, 4433–4440.


240

Mann, C.M., Markham, J.L., 1998. A new method for determining the minimum inhibitory

concentration of essential oils. J. Appl. Microbiol. 84, 538–544.

Mann, E.E., Rice, K.C., Boles, B.R., Endres, J.L., Ranjit, D., Chandramohan, L., Tsang, L.H.,

Smeltzer, M.S., Horswill, A.R., Bayles, K.W., 2009. Modulation of eDNA release and

degradation affects Staphylococcus aureus biofilm maturation. PLoS One 4, e5822.

Marino, M., Frigo, F., Bartolomeoli, I., Maifreni, M., 2011. Safety-related properties of

staphylococci isolated from food and food environments. J. Appl. Microbiol. 110, 550–

561.

Marriott, N., Gravani, R.B., 2006. Principles of food sanitation, 5th ed. Springer, New York,

NY.

Marshall, K.C., 1976. Interfaces in microbial ecology. Harvard University Press, Cambridge,

Mass, pp. 44–47.

Martín, M.C., Fueyo, J.M., González-Hevia, M.A., Mendoza, M.C., 2004. Genetic procedures

for identification of enterotoxigenic strains of Staphylococcus aureus from three food

poisoning outbreaks. Int. J. Food Microbiol. 94, 279–286.

Martínez, O., Rodríguez-Calleja, J.M., Santos, J.A., Otero, A., García-López, M.L., 2009.

Foodborne and indicator bacteria in farmed molluscan shellfish before and after

depuration. J. Food Prot. 72, 1443–1449.

Mauermann, M., Eschenhagen, U., Bley, T., Majschak, J.-P., 2009. Surface modifications –

Application potential for the reduction of cleaning costs in the food processing industry.

Trends Food Sci. Technol. 20, Supple, S9 – S15.

Maukonen, J., Mättö, J., Wirtanen, G., Raaska, L., Mattila-Sandholm, T., Saarela, M., 2003.

Methodologies for the characterization of microbes in industrial environments: a review. J.

Ind. Microbiol. Biotechnol. 30, 327–356.

Mayaud, L., Carricajo, A., Zhiri, A., Aubert, G., 2008. Comparison of bacteriostatic and

bactericidal activity of 13 essential oils against strains with varying sensitivity to

antibiotics. Lett. Appl. Microbiol. 47, 167–173.

McPherson, L.L., 1993. Understanding ORP’s in the disinfection process. Water Eng. Manag.

140, 29–31.

Meira, Q.G.S., Barbosa, I.M., Athayde, A.J.A.A., Siqueira-Júnior, J.P., Souza, E.L., 2012.

Influence of temperature and surface kind on biofilm formation by Staphylococcus aureus

from food-contact surfaces and sensitivity to sanitizers. Food Control 25, 469–475.

Melchior, M.B., Fink-Gremmels, J., Gaastra, W., 2007. Extended antimicrobial susceptibility

assay for Staphylococcus aureus isolates from bovine mastitis growing in biofilms. Vet.

Microbiol. 125, 141–149.

Merino, N., Toledo-arana, A., Valle, J., Solano, C., Lopez, J.A., Foster, T.J., José, R., Vergara-

irigaray, M., Calvo, E., Penade, R., 2009. Protein A-mediated multicellular behavior in

Staphylococcus aureus. J. Bacteriol. 191, 832–843.

Meyer, B., Morin, V.N., Rödger, H.-J., Holah, J., Bird, C., 2010. Do European Standard

Disinfectant tests truly simulate in-use microbial and organic soiling conditions on food

preparation surfaces? J. Appl. Microbiol. 108, 1344–1351.

MHLW, 2011. Food poisoning statistics 2009. Japan.

Milanov, D., Lazic, S., Vidic, B., Petrovic, J., Bugarski, D., Seguljev, Z., 2010. Slime

production and biofilm forming ability by Staphylococcus aureus bovine mastitis isolates.

Acta Vet. Brno. 60, 217–226.


241

Millezi, F.M., Pereira, M.O., Batista, N.N., Camargos, N., Auad, I., Cardoso, M.D.G., Piccoli,

R.H., 2012. Susceptibility of monospecies and dual-species biofilms of Staphylococcus

aureus and Escherichia coli to essential oils. J. Food Saf. 32, 351–359.

Monnin, A., Lee, J., Pascall, M.A., 2012. Efficacy of neutral electrolyzed water for sanitization

of cutting boards used in the preparation of foods. J. Food Eng. 110, 541–546.

Morandi, S., Brasca, M., Lodi, R., Brusetti, L., Andrighetto, C., Lombardi, A., 2010.

Biochemical profiles, restriction fragment length polymorphism (RFLP), random

amplified polymorphic DNA (RAPD) and multilocus variable number tandem repeat

analysis (MLVA) for typing Staphylococcus aureus isolated from dairy products. Res.

Vet. Sci. 88, 427–435.

Moreillon, P., Que, Y.-A., 2004. Infective endocarditis. Lancet 363, 139–149.

Møretrø, T., Hermansen, L., Holck, A.L., Sidhu, M.S., Rudi, K., Langsrud, S., 2003. Biofilm

formation and the presence of the intercellular adhesion locus ica among staphylococci

from food and food processing environments. Appl. Environ. Microbiol. 69, 5648–5655.

Morris, C.A., Conway, H.D., Everall, P.H., 1972. Food-poisoning due to staphylococcal

enterotoxin E. Lancet 300, 1375–1376.

Nema, V., Agrawal, R., Kamboj, D.V., Goel, A.K., Singh, L., 2007. Isolation and

characterization of heat resistant enterotoxigenic Staphylococcus aureus from a food

poisoning outbreak in Indian subcontinent. Int. J. Food Microbiol. 117, 29–35.

Nerio, L.S., Olivero-Verbel, J., Stashenko, E., 2010. Repellent activity of essential oils: a

review. Bioresour. Technol. 101, 372–378.

Niemira, B.A., 2008. Irradiation sensitivity of planktonic and biofilm-associated Listeria

monocytogenes and L. innocua as influenced by temperature of biofilm formation. Food

Bioprocess Technol. 3, 257–264.

Niemira, B.A., Solomon, E.B., 2005. Sensitivity of planktonic and biofilm-associated

Salmonella spp . to ionizing radiation. Appl. Environ. Microbiol. 71, 2732–2736.

Nikbakht, M., Nahaei, M.R., Akhi, M.T., Asgharzadeh, M., Nikvash, S., 2008. Molecular

fingerprinting of meticillin-resistant Staphylococcus aureus strains isolated from patients

and staff of two Iranian hospitals. J. Hosp. Infect. 69, 46–55.

Normanno, G., Dambrosio, A., Quaglia, N.C., Celano, G. V, Germinario, G.L., Parisi, A., 2003.

Hygienic-sanitary evaluations about typically raw-eaten seafoods (Apulia). Ind. Aliment.

42, 961–964.

Normanno, G., Firinu, A., Virgilio, S., Mula, G., Dambrosio, A., Poggiu, A., Decastelli, L.,

Mioni, R., Scuota, S., Bolzoni, G., Di-Giannatale, E., Salinetti, A.P., La Salandra, G.,

Bartoli, M., Zuccon, F., Pirino, T., Sias, S., Parisi, A., Quaglia, N.C., Celano, G. V, 2005.

Coagulase-positive staphylococci and Staphylococcus aureus in food products marketed in

Italy. Int. J. Food Microbiol. 98, 73–79.

Normanno, G., La Salandra, G., Dambrosio, A., Quaglia, N.C., Corrente, M., Parisi, A.,

Santagada, G., Firinu, A., Crisetti, E., Celano, G. V., 2007. Occurrence, characterization

and antimicrobial resistance of enterotoxigenic Staphylococcus aureus isolated from meat

and dairy products. Int. J. Food Microbiol. 115, 290–296.

Nostro, A., Roccaro, A.S., Bisignano, G., Marino, A., Cannatelli, M. a, Pizzimenti, F.C., Cioni,

P.L., Procopio, F., Blanco, A.R., 2007. Effects of oregano, carvacrol and thymol on

Staphylococcus aureus and Staphylococcus epidermidis biofilms. J. Med. Microbiol. 56,

519–523.


242

Novotny, L., Dvorska, L., Lorencova, A., Beran, V., Pavlik, I., 2004. Fish: a potential source of

bacterial pathogens for human beings. Vet. Med. 2004, 343–358.

O´Riordan, K., Lee, J.C., 2004. Staphylococcus aureus capsular polysaccharides. Clin.


O’Donnell, C., Tiwari, B., Cullen, P., Rice, R., 2012. Ozone in food processing. John Wiley &

Sons Ltd., Chichester, United Kingdom.

O’Gara, J.P., 2007. Ica and beyond: biofilm mechanisms and regulation in Staphylococcus

epidermidis and Staphylococcus aureus. FEMS Microbiol. Lett. 270, 179–188.

O’Neill, E., Pozzi, C., Houston, P., Humphreys, H., Robinson, D.A., Loughman, A., Foster,

T.J., O’Gara, J.P., 2008. A novel Staphylococcus aureus biofilm phenotype mediated by

the fibronectin-binding proteins, FnBPA and FnBPB. J. Bacteriol. 190, 3835–3850.

Obeso, J.M., Martínez, B., Rodríguez, A., García, P., 2008. Lytic activity of the recombinant

staphylococcal bacteriophage ΦH5 endolysin active against Staphylococcus aureus in

milk. Int. J. Food Microbiol. 128, 212–218.

Ogston, A., 1882. Micrococcus poisoning. J. Anat. Physiol. 17, 24–58.

Oh, S.K., Lee, N., Cho, Y.S., Shin, D.-B., Choi, S.Y., Koo, M., 2007. Occurrence of toxigenic

Staphylococcus aureus in ready-to-eat food in Korea. J. Food Prot. 70, 1153–1158.

Oliveira, M., Bexiga, R., Nunes, S.F., Carneiro, C., Cavaco, L.M., Bernardo, F., Vilela, C.L.,

2006. Biofilm-forming ability profiling of Staphylococcus aureus and Staphylococcus

epidermidis mastitis isolates. Vet. Microbiol. 118, 133–140.

Oliveira, M.M.M., Brugnera, D.F., Cardoso, M.D.G., Alves, E., Piccoli, R.H., 2010.

Disinfectant action of Cymbopogon sp. essential oils in different phases of biofilm

formation by Listeria monocytogenes on stainless steel surface. Food Control 21, 549–553.

Olsen, J.E., Christensen, H., Aarestrup, F.M., 2006. Diversity and evolution of blaZ from

Staphylococcus aureus and coagulase-negative staphylococci. J. Antimicrob. Chemother.

57, 450–460.

Omoe, K., Hu, D.-L., Takahashi-Omoe, H., Nakane, A., Shinagawa, K., 2005. Comprehensive

analysis of classical and newly described staphylococcal superantigenic toxin genes in

Staphylococcus aureus isolates. FEMS Microbiol. Lett. 246, 191–198.

Omoe, K., Ishikawa, M., Shimoda, Y., Hu, D., Ueda, S., Shinagawa, K., 2002. Detection of seg,

seh and sei in Staphylococcus aureus isolates and determination of the enterotoxin

productivities of S . aureus isolates harboring seg , seh , or sei genes. J. Clin. Microbiol.

40, 857–862.

Oomori, T., Oka, T., Inuta, T., Arata, Y., 2000. The efficiency of disinfection of acidic

electrolyzed water in the presence of organic materials. Anal. Sci. 16, 365–369.

Ortega, E., Abriouel, H., Lucas, R., Gálvez, A., 2010. Multiple roles of Staphylococcus aureus

enterotoxins: pathogenicity, superantigenic activity, and correlation to antibiotic

resistance. Toxins 2, 2117–31.

Ostyn, A., De-Buyser, M.L., Guillier, F., Groult, J., Felix, B., Salah, S., Delmas, G.,

Hennekinne, J.A., 2010. First evidence of a food poisoning outbreak due to staphylococcal

enterotoxin type E, France, 2009. Euro Surveill. 15, 1–4.

Otto, C., Zahn, S., Rost, F., Zahn, P., Jaros, D., Rohm, H., 2011. Physical methods for cleaning

and disinfection of surfaces. Food Eng. Rev. 3, 171–188.

Otto, M., 2008. Staphylococcal biofilms. Curr. Top. Microbiol. Inmunol. 322, 207–228.


243

Otto, M., 2013. Staphylococcal infections: mechanisms of biofilm maturation and detachment

as critical determinants of pathogenicity. Annu. Rev. Med. 64, 175–188.

Oulahal-Lagsir, N., Martial-Gros, a., Boistier, E., Blum, L.J., Bonneau, M., 2000. The

development of an ultrasonic apparatus for the non-invasive and repeatable removal of

fouling in food processing equipment. Lett. Appl. Microbiol. 30, 47–52.

Oussalah, M., Caillet, S., Saucier, L., Lacroix, M., 2007. Inhibitory effects of selected plant

essential oils on the growth of four pathogenic bacteria: E. coli O157:H7, Salmonella

Typhimurium, Staphylococcus aureus and Listeria monocytogenes. Food Control 18, 414–

420.

Ozer, N.P., Demirci, A., 2006. Electrolyzed oxidizing water treatment for decontamination of

raw salmon inoculated with Escherichia coli O157:H7 and Listeria monocytogenes Scott

A and response surface modeling. J. Food Eng. 72, 234–241.

Pagedar, A., Singh, J., Batish, V.K., 2010. Surface hydrophobicity, nutritional contents affect

Staphylococcus aureus biofilms and temperature influences its survival in preformed

biofilms. J. Basic Microbiol. 50, S98–S106.

Paolini, J., Barboni, T., Desjobert, J.-M., Djabou, N., Muselli, A., Costa, J., 2010. Chemical

composition, intraspecies variation and seasonal variation in essential oils of Calendula

arvensis L. Biochem. Syst. Ecol. 38, 865–874.

Papadopoulou, C., Economou, E., Zakas, G., Salamoura, C., Dontorou, C., Apostolou, J., 2007.

Microbiological and pathogenic contaminants of seafood in Greece. J. Food Qual. 30, 28–

42.

Park, E.S., Kim, H.S., Kim, M.N., Yoon, J.S., 2004. Antibacterial activities of polystyrene-

block-poly(4-vinyl pyridine) and poly(styrene-random-4-vinyl pyridine). Eur. Polym. J.

40, 2819–2822.

Park, H., Hung, Y.-C., Kim, C., 2002. Effectiveness of electrolyzed water as a sanitizer for

treating different surfaces. J. Food Prot. 65, 1276–1280.

Parsek, M.R., Greenberg, E.P., 2005. Sociomicrobiology: the connections between quorum

sensing and biofilms. Trends Microbiol. 13, 27–33.

Pascual, A., Llorca, I., Canut, A., 2007. Use of ozone in food industries for reducing the

environmental impact of cleaning and disinfection activities. Trends Food Sci. Technol.

18, S29–S35.

Pedro, S., Albuquerque, M.M., Nunes, M.L., Bernardo, M.F., 2004. Pathogenic bacteria and

indicators in salted cod (Gadus morhua) and desalted products at low and high

temperatures. J. Aquat. Food Prod. Technol. 13, 39–48.

Peeters, E., Nelis, H.J., Coenye, T., 2008. Comparison of multiple methods for quantification of

microbial biofilms grown in microtiter plates. J. Microbiol. Methods 72, 157–165.

Peles, F., Wagner, M., Varga, L., Hein, I., Rieck, P., Gutser, K., Keresztúri, P., Kardos, G.,

Turcsányi, I., Béri, B., Szabó, A., 2007. Characterization of Staphylococcus aureus strains

isolated from bovine milk in Hungary. Int. J. Food Microbiol. 118, 186–193.

Pereira, A., Mendes, J., Melo, L.F., 2008. Using nanovibrations to monitor biofouling.

Biotechnol. Bioeng. 99, 1407–1415.

Pereira, V., Lopes, C., Castro, A., Silva, J., Gibbs, P., Teixeira, P., 2009. Characterization for

enterotoxin production, virulence factors, and antibiotic susceptibility of Staphylococcus

aureus isolates from various foods in Portugal. Food Microbiol. 26, 278–82.


244

Periasamy, S., Joo, H.-S., Duong, A.C., Bach, T.-H.L., Tan, V.Y., Chatterjee, S.S., Cheung,

G.Y.C., Otto, M., 2012. How Staphylococcus aureus biofilms develop their characteristic

structure. PNAS 109, 1281–1286.

Pesavento, G., Ducci, B., Comodo, N., Nostro, A. Lo, 2007. Antimicrobial resistance profile of

Staphylococcus aureus isolated from raw meat: A research for methicillin resistant

Staphylococcus aureus (MRSA). Food Control 18, 196–200.

Philip-Chandy, R., Scully, P.J., Eldridge, P., Kadim, H.J., Grapin, M.G., Jonca, M.G.,

D’Ambrosio, M.G., Colin, F., 2000. An optical fiber sensor for biofilm measurement using

intensity modulation and image analysis. IEEE J. Sel. Top. Quantum Electron. 6, 764–772.

Phuvasate, S., Su, Y.-C., 2010. Effects of electrolyzed oxidizing water and ice treatments on

reducing histamine-producing bacteria on fish skin and food contact surface. Food Control

21, 286–291.

Pinchuk, I.V., Beswick, E.J., Reyes, V.E., 2010. Staphylococcal enterotoxins. Toxins (Basel). 2,

2177–2197.

Planchon, S., Gaillard-Martinie, B., Dordet-Frisoni, E., Bellon-Fontaine, M.N., Leroy, S.,

Labadie, J., Hébraud, M., Talon, R., 2006. Formation of biofilm by Staphylococcus

xylosus. Int. J. Food Microbiol. 109, 88–96.

Popovich, K., Hota, B., Weinstein, R., 2007. Treatment of community-associated methicillin-

resistant Staphylococcus aureus. Curr. Infect. Dis. Rep. 9, 398–407.

Poulsen, L., 1999. Microbial biofilm in food processing. LWT-Food Sci. Technol. 32, 321–326.

Prabuseenivasan, S., Jayakumar, M., Ignacimuthu, S., 2006. In vitro antibacterial activity of

some plant essential oils. BMC Complement. Altern. Med. 6, 39.

Prat, C., Bestebroer, J., Haas, C.J.C., Strijp, J.A.G., Kessel, K.P.M., 2006. A new

staphylococcal anti-inflammatory protein that antagonizes the formyl peptide receptor-like

1. J. Immunol. 177, 8017–8026.

Prigent-Combaret, C., Lejeune, P., 1999. Monitoring gene expression in biofilms. Methods

Enzymol. 310, 56–79.

Qiu, J., Wang, D., Xiang, H., Feng, H., Jiang, Y., Xia, L., Dong, J., Lu, J., Yu, L., Deng, X.,

2010. Subinhibitory concentrations of thymol reduce enterotoxins A and B and alpha-

hemolysin production in Staphylococcus aureus isolates. PLoS One 5, e9736.

Quave, C.L., Plano, L.R.W., Pantuso, T., Bennett, B.C., 2008. Effects of extracts from Italian

medicinal plants on planktonic growth, biofilm formation and adherence of methicillin-

resistant Staphylococcus aureus. J. Ethnopharmacol. 118, 418–428.

Rachid, S., Ohlsen, K., Wallner, U., Hacker, J., Hecker, M., Ziebuhr, W., 2000. Alternative

transcription factor sigma(B) is involved in regulation of biofilm expression in a

Staphylococcus aureus mucosal isolate. J. Bacteriol. 182, 6824–6826.

Rahman, S.M.E., Ding, T., Oh, D.-H., 2010. Effectiveness of low concentration electrolyzed

water to inactivate foodborne pathogens under different environmental conditions. Int. J.

Food Microbiol. 139, 147–153.

Rahman, S.M.E., Jin, Y.-G., Oh, D.-H., 2010. Combined effects of alkaline electrolyzed water

and citric acid with mild heat to control microorganisms on cabbage. J. Food Sci. 75,

M111–115.

Rahman, S.M.E., Jin, Y.-G., Oh, D.-H., 2011. Combination treatment of alkaline electrolyzed

water and citric acid with mild heat to ensure microbial safety, shelf-life and sensory

quality of shredded carrots. Food Microbiol. 28, 484–491.


245

Rendueles, E., Omer, M.K., Alvseike, O., Alonso-Calleja, C., Capita, R., Prieto, M., 2011.

Microbiological food safety assessment of high hydrostatic pressure processing: A review.

LWT - Food Sci. Technol. 44, 1251–1260.

Renner, L.D., Weibel, D.B., 2011. Physicochemical regulation of biofilm formation. MRS Bull.

36, 347–355.

Restaino, L., Frampton, E., Hemphill, J., Palnikar, P., 1995. Efficacy of ozonated water against

various food-related microorganisms. Appl. Environ. Microbiol. 61, 3471–3475.

Ribeiro, A., Coronado, A.Z., Silva-Carvalho, M.C., Ferreira-Carvalho, B.T., Dias, C.,

Rozenbaum, R., Del-Peloso, P.F., Leite, C.C.F., Teixeira, L.A., Figueiredo, A.M.S., 2007.

Detection and characterization of international community-acquired infections by

methicillin-resistant Staphylococcus aureus clones in Rio de Janeiro and Porto Alegre

cities causing both community- and hospital-associated diseases. Diagn. Microbiol. Infect.

Dis. 59, 339–345.

Rice, K.C., Mann, E.E., Endres, J.L., Weiss, E.C., Cassat, J.E., Smeltzer, M.S., Bayles, K.W.,

2007. The cidA murein hydrolase regulator contributes to DNA release and biofilm

development in Staphylococcus aureus. Proc. Natl. Acad. Sci. U. S. A. 104, 8113–8118.

Richards, M.S., Rittman, M., Gilbert, T.T., Opal, S.M., DeBuono, B.A., Neill, R.J., Gemski, P.,

1993. Investigation of a staphylococcal food poisoning outbreak in a centralized school

lunch program. Public Health Rep. 108, 765–71.

Roberts, T., 2007. WTP estimates of the societal costs of U.S. food-borne illness. Am. J. Agric.

Econ. 89, 1183–1188.

Rode, T.M., Langsrud, S., Holck, A., Møretrø, T., 2007. Different patterns of biofilm formation

in Staphylococcus aureus under food-related stress conditions. Int. J. Food Microbiol. 116,

372–83.

Rosenbach, F.J., 1884. Mikro-organismen bei den wund-infections-krankheiten des menschen.

Wiesbaden J. F. Bergmann, Göttingen.

Rosenberg, M., 1981. Bacterial adherence to polystyrene: a replica method of screening for

bacterial hydrophobicity. Appl. Environ. Microbiol. 42, 375–377.

Rosmaninho, R., Santos, O., Nylander, T., Paulsson, M., Beuf, M., Benezech, T., Yiantsios, S.,

Andritsos, N., Karabelas, A., Rizzo, G., Müller-Steinhagen, H., Melo, L.F., 2007.

Modified stainless steel surfaces targeted to reduce fouling – Evaluation of fouling by milk

components. J. Food Eng. 80, 1176–1187.

Rota, M.C., Herrera, A., Martínez, R.M., Sotomayor, J.A., Jordán, M.J., 2008. Antimicrobial

activity and chemical composition of Thymus vulgaris, Thymus zygis and Thymus hyemalis

essential oils. Food Control 19, 681–687.

Russell, A.D., 2003. Similarities and differences in the responses of microorganisms to

biocides. J. Antimicrob. Chemother. 52, 750–763.

Saá-Ibusquiza, P., Herrera, J.J.R., Cabo, M.L., 2011. Resistance to benzalkonium chloride,

peracetic acid and nisin during formation of mature biofilms by Listeria monocytogenes.


Sakoulas, G., Moellering, R.C., 2008. Increasing antibiotic resistance among methicillin-

resistant Staphylococcus aureus strains. Clin. Infect. Dis. 46, 360–367.

Sandel, M.K., McKillip, J.L., 2004. Virulence and recovery of Staphylococcus aureus relevant

to the food industry using improvements on traditional approaches. Food Control 15, 5–10.


246

Sanderson, S.S., Stewart, P.S., 1997. Evidence of bacterial adaptation to monochloramine in

Pseudomonas aeruginosa biofilms and evaluation of biocide action model. Biotechnol.

Bioeng. 56, 201–209.

Sass, P., Bierbaum, G., 2007. Lytic activity of recombinant bacteriophage phi11 and phi12

endolysins on whole cells and biofilms of Staphylococcus aureus. Appl. Environ.

Microbiol. 73, 347–352.

Sattar, S.A., Springthorpe, S., Mani, S., Gallant, M., Nair, R.C., Scott, E., Kain, J., 2001.

Transfer of bacteria from fabrics to hands and other fabrics: development and application

of a quantitative method using Staphylococcus aureus as a model. J. Appl. Microbiol. 90,

962–970.

Schelin, J., Wallin-Carlquist, N., Cohn, M.T., Lindqvist, R., Barker, G.C., Radstrom, P., 2011.

The formation of Staphylococcus aureus enterotoxin in food environments and advances

in risk assessment. Virulence 2, 580–592.

Schillaci, D., Arizza, V., Dayton, T., Camarda, L., Di Stefano, V., 2008. In vitro anti-biofilm

activity of Boswellia spp. oleogum resin essential oils. Lett. Appl. Microbiol. 47, 433–438.

Schleifer, K.H., Bell, J.A., 2009. Family VIII. Staphylococcaceae fam. nov.., in: Vos, P.D.,

Garrity, G., Jones, D., Krieg, N.R., Ludwig, W., Rainey, F.A., Schleifer, K.-H., Whitman,

W.B. (Eds.), Bergey´s Manual of Systematic Bacteriology, Volume 3: The Firmicutes.

Springer, pp. 392–426.

Schmid, D., Fretz, R., Winter, P., Mann, M., Höger, G., Stöger, A., Ruppitsch, W., Ladstätter,

J., Mayer, N., de Martin, A., Allerberger, F., 2009. Outbreak of staphylococcal food

intoxication after consumption of pasteurized milk products, June 2007, Austria. Wien.

Klin. Wochenschr. 121, 125–131.

Schroeder, K., Jularic, M., Horsburgh, S.M., Hirschhausen, N., Neumann, C., Bertling, A.,

Schulte, A., Foster, S., Kehrel, B.E., Peters, G., Heilmann, C., 2009. Molecular

characterization of a novel Staphylococcus aureus surface protein (SasC) involved in cell

aggregation and biofilm accumulation. PLoS One 4, e7567.

Schweizer, H.P., 2001. Triclosan: a widely used biocide and its link to antibiotics. FEMS

Microbiol. Lett. 202, 1–7.

Sharma, M., Anand, S.., 2002. Biofilms evaluation as an essential component of HACCP for

food/dairy processing industry – a case. Food Control 13, 469–477.

Shehata, A., 2008. Phylogenetic diversity of Staphylococcus aureus by random amplification of

polymorphic DNA. Aust. J. Basic Appl. Sci. 2, 858–863.

Sheridan, À., Lenahan, M., Duffy, G., Fanning, S., Burgess, C., 2012. The potential for biocide

tolerance in Escherichia coli and its impact on the response to food processing stresses.


Shimizu, A., Fujita, M., Igarashi, H., Takagi, M., Nagase, N., Sasaki, A., Kawano, J., 2000.

Characterization of Staphylococcus aureus coagulase type VII isolates from

staphylococcal food poisoning outbreaks (1980-1995) in Tokyo, Japan, by pulsed-field gel

electrophoresis. J. Clin. Microbiol. 38, 3746–3749.

Silva, N., Alves, S., Gonçalves, A., Amaral, J.S., Poeta, P., 2013. Antimicrobial activity of

essential oils from mediterranean aromatic plants against several foodborne and spoilage

bacteria. Food Sci. Technol. Int. 19, 503–510.

Simões, M., Simões, L.C., Vieira, M.J., 2010. A review of current and emergent biofilm control

strategies. LWT - Food Sci. Technol. 43, 573–583.


247

Simon, S.S., Sanjeev, S., 2007. Prevalence of enterotoxigenic Staphylococcus aureus in fishery

products and fish processing factory workers. Food Control 18, 1565–1568.

Singh, R., Ray, P., Das, A., Sharma, M., 2010. Penetration of antibiotics through

Staphylococcus aureus and Staphylococcus epidermidis biofilms. J. Antimicrob.

Chemother. 65, 1955–1958.

Smyth, C.J., Smyth, D.S., Kennedy, J., Twohig, J., Bolton, D., 2004. Staphylococcus aureus:

from man or animals—an enterotoxin iceberg?, in: Maunsell, B., Sheridan, J., Bolton, D.J.

(Eds.), Food Pathogen Epidemiology: Microbes, Maladies and Methods, Proceedings of an

International EU-RAIN Conference, 3–4 December Padua (Italy). Teagasc - The National

Food Centre, Dublin, pp. 85–102.

Sneath, P.H.A., Sokal, R.R., 1973. Numerical taxonomy: the principles and practice of

numerical classification. Freeman, San Francisco.

Solano, R., Lafuente, S., Sabate, S., Tortajada, C., García de Olalla, P., Hernando, A.V., Caylà,

J., 2013. Enterotoxin production by Staphylococcus aureus: An outbreak at a Barcelona

sports club in July 2011. Food Control 33, 114–118.

Song, B., Leff, L.G., 2006. Influence of magnesium ions on biofilm formation by Pseudomonas

fluorescens. Microbiol. Res. 161, 355–361.

Sospedra, I., Mañes, J., Soriano, J.M., 2012. Report of toxic shock syndrome toxin 1 (TSST-1)

from Staphylococcus aureus isolated in food handlers and surfaces from foodservice

establishments. Ecotoxicol. Environ. Saf. 80, 288–290.

Srey, S., Jahid, I.K., Ha, S., 2013. Biofilm formation in food industries: A food safety concern.


Stankovic, C., Mahajan, P. V, Asmar, B.I., 2007. Methicillin-resistant Staphylococcus aureus as

a cause of community-acquired pneumonia. Curr. Infect. Dis. Rep. 9, 223–227.

Straub, J.A., Hertel, C., Hammes, W.P., 1999. A 23S rDNA-targeted polymerase chain reaction-

based system for detection of Staphylococcus aureus in meat starter cultures and dairy

products. J. Food Prot. 62, 1150–1156.

Strommenger, B., Kettlitz, C., Werner, G., Witte, W., 2003. Multiplex PCR assay for

simultaneous detection of nine clinically relevant antibiotic resistance genes in

Staphylococcus aureus. J. Clin. Microbiol. 41, 4089–4094.

Struelens, M.J., Bauernfeind, A., Van-Belkum, A., Blanc, D., Cookson, B.D., Dijkshoorn, L.,

El-Solh, N., Etienne, J., Garaizar, J., Gerner-Smidh, P., Legakis, N., De-Lencastre, H.,

Nicolas, M.H., Pitt, T.L., Römling, U., Rosdahl, V., Witte, W., 1996. Consensus

guidelines for appropriate use and evaluation of microbial epidemiologic typing systems.

Clin. Microbiol. Infect. 2, 2–11.

Sun, J.-L., Zhang, S.-K., Chen, J.-Y., Han, B.-Z., 2012. Efficacy of acidic and basic electrolyzed

water in eradicating Staphylococcus aureus biofilm. Can. J. Microbiol. 58, 448–454.

Sutherland, I.W., Hughes, K.A., Skillman, L.C., Tait, K., 2004. The interaction of phage and

biofilms. FEMS Microbiol. Lett. 232, 1–6.

Taylor, J.H., Rogers, S.J., Holah, J.T., 1999. A comparison of the bactericidal efficacy of 18

disinfectants used in the food industry against Escherichia coli O157:H7 and

Pseudomonas aeruginosa at 10 and 20 degrees C. J. Appl. Microbiol. 87, 718–25.

Thaikruea, L., Pataraarechachai, J., Savanpunyalert, P., Naluponjiragul, U., 1995. An unusual

outbreak of food poisoning. Southeast Asian J. Trop. Med. Public Health 26, 78–85.


248

Thallinger, B., Prasetyo, E.N., Nyanhongo, G.S., Guebitz, G.M., 2013. Antimicrobial enzymes:

an emerging strategy to fight microbes and microbial biofilms. Biotechnol. J. 8, 97–109.

Thouvenin, M., Langlois, V., Briandet, R., Langlois, J.Y., Guerin, P.H., Peron, J.J., Haras, D.,

Vallee-Rehel, K., 2003. Study of erodable paint properties involved in antifouling activity.

Biofouling 19, 177–86.

Todd, E., Szabo, R., Gardiner, M.A., Aktar, M., Delorme, L., Tourillon, P., Rochefort, J., Roy,

D., Loit, T., Lamontagne, Y., Gosselin, L., Martineau, G., Breton, J.P., 1981. Intoxication

staphylococcique liée à du caillé de fromagerie – Québec. Rapp. Hebd. des Mal. au

Canada 7, 171–172.

Trampuz, A., Widmer, A.F., 2006. Infections associated with orthopedic implants. Curr. Opin.

Infect. Dis. 19, 349–356.

Trombetta, D., Castelli, F., Sarpietro, M.G., Venuti, V., Cristani, M., Daniele, C., Saija, A.,

Mazzanti, G., Bisignano, G., 2005. Mechanisms of antibacterial action of three

monoterpenes. Antimicrob. Agents Chemother. 49, 2474–2478.

Tserennadmid, R., Takó, M., Galgóczy, L., Papp, T., Pesti, M., Vágvölgyi, C., Almássy, K.,

Krisch, J., 2011. Anti yeast activities of some essential oils in growth medium, fruit juices

and milk. Int. J. Food Microbiol. 144, 480–486.

Turina, A. V, Nolan, M. V, Zygadlo, J.A., Perillo, M.A., 2006. Natural terpenes: self-assembly

and membrane partitioning. Biophys. Chem. 122, 101–113.

Ultee, A., Kets, E.P., Alberda, M., Hoekstra, F.A., Smid, E.J., 2000. Adaptation of the food-

borne pathogen Bacillus cereus to carvacrol. Arch. Microbiol. 174, 233–238.

Valle, J., Toledo-Arana, A., Berasain, C., Ghigo, J.-M., Amorena, B., Penadés, J.R., Lasa, I.,

2003. SarA and not sigmaB is essential for biofilm development by Staphylococcus

aureus. Mol. Microbiol. 48, 1075–1087.

Van-Belkum, A., Kluytmans, J., Van-Leeuwen, W., Bax, R., Quint, W., Peters, E., Fluit, A.,

Vandenbroucke-Grauls, C., Brule, A., Koeleman, H., 1995. Multicenter evaluation of

arbitrarily primed PCR for typing of Staphylococcus aureus strains. J. Clin. Microbiol. 33,

1537–1547.

Van-Belkum, A., Melles, D.C., 2005. Not all Staphylococcus aureus strains are equally

pathogenic. Discov. Med. 5, 148–152.

Van-Houdt, R., Michiels, C.W., 2010. Biofilm formation and the food industry, a focus on the

bacterial outer surface. J. Appl. Microbiol. 109, 1117–1131.

Vasudevan, P., Nair, M.K.M., Annamalai, T., Venkitanarayanan, K.S., 2003. Phenotypic and

genotypic characterization of bovine mastitis isolates of Staphylococcus aureus for biofilm

formation. Vet. Microbiol. 92, 179–185.

Vautor, E., Abadie, G., Pont, A., Thiery, R., 2008. Evaluation of the presence of the bap gene in

Staphylococcus aureus isolates recovered from human and animals species. Vet.

Microbiol. 127, 407–411.

Vázquez-Sánchez, D., Cabo, M.L., Ibusquiza, P.S., Rodríguez-Herrera, J.J., 2014. Biofilm-

forming ability and resistance to industrial disinfectants of Staphylococcus aureus isolated

from fishery products. Food Control 39, 8–16.

Vázquez-Sánchez, D., Habimana, O., Holck, A., 2013. Impact of food-related environmental

factors on the adherence and biofilm formation of natural Staphylococcus aureus isolates.

Curr. Microbiol. 66, 110–121.


249

Vázquez-Sánchez, D., López-Cabo, M., Saá-Ibusquiza, P., Rodríguez-Herrera, J.J., 2012.

Incidence and characterization of Staphylococcus aureus in fishery products marketed in

Galicia (Northwest Spain). Int. J. Food Microbiol. 157, 286–296.

Venkitanarayanan, K.S., Ezeike, G.O., Hung, Y.C., Doyle, M.P., 1999. Efficacy of electrolyzed

oxidizing water for inactivating Escherichia coli O157:H7, Salmonella enteritidis, and

Listeria monocytogenes. Appl. Environ. Microbiol. 65, 4276–4279.

Vesterholm-Nielsen, M., Olholm-Larsen, M., Olsen, J.E., Aarestrup, F.M., 1999. Occurrence of

the blaZ gene in penicillin resistant Staphylococcus aureus isolated from bovine mastitis in

Denmark. Acta Vet. Scand. 40, 279–286.

Victorin, K., 1992. Review of the genotoxicity of ozone. Mutat. Res. Genet. Toxicol. 277, 221–

238.

Vorobjeva, N. V, Vorobjeva, L.I., Khodjaev, E.Y., 2004. The bactericidal effects of electrolyzed

oxidizing water on bacterial strains involved in hospital infections. Artif. Organs 28, 590–

592.

Vuong, C., Kocianova, S., Voyich, J.M., Yao, Y., Fischer, E.R., DeLeo, F.R., Otto, M., 2004. A

crucial role for exopolysaccharide modification in bacterial biofilm formation, immune

evasion, and virulence. J. Biol. Chem. 279, 54881–54886.

Walker, S.P., Demirci, A., Graves, R.E., Spencer, S.B., Roberts, R.F., 2005. Cleaning milking

systems using electrolyzed oxidizing water. Trans. Am. Soc. Agric. Eng. 48, 1827–1833.

Walsh, S.E., Maillard, J.-Y., Russell, A.D., Catrenich, C.E., Charbonneau, D.L., Bartolo, R.G.,

2003. Activity and mechanisms of action of selected biocidal agents on Gram-positive and

-negative bacteria. J. Appl. Microbiol. 94, 240–247.

Wang, H., Feng, H., Luo, Y., 2004. Microbial reduction and storage quality of fresh-cut cilantro

washed with acidic electrolyzed water and aqueous ozone. Food Res. Int. 37, 949–956.

Wang, H., Feng, H., Luo, Y., 2006. Dual-phasic inactivation of Escherichia coli O157: H7 with

peroxyacetic acid, acidic electrolyzed water and chlorine on cantaloupes and fresh-cut

apples. J. Food Saf. 26, 335–347.

Wang, R., Braughton, K.R., Kretschmer, D., Bach, T.L., Queck, S.Y., Li, M., Kennedy, A.D.,

Dorward, D.W., Klebanoff, S.J., Peschel, A., DeLeo, F.R., Otto, M., 2007. Identification

of novel cytolytic peptides as key virulence determinants for community-associated

MRSA. Nat. Med. 13, 1510–1514.

Wang, S., Duan, H., Zhang, W., Li, J.-W., 2007. Analysis of bacterial foodborne disease

outbreaks in China between 1994 and 2005. FEMS Immunol. Med. Microbiol. 51, 8–13.

Waterman, S.H., Demarcus, T.A., Wells, J.G., Blake, P.A., 1987. Staphylococcal food

poisoning on a cruise ship. Epidemiol. Infect. 99, 349–353.

Waters, A.E., Contente-Cuomo, T., Buchhagen, J., Liu, C.M., Watson, L., Pearce, K., Foster,

J.T., Bowers, J., Driebe, E.M., Engelthaler, D.M., Keim, P.S., Price, L.B., 2011.

Multidrug-Resistant Staphylococcus aureus in US Meat and Poultry. Clin. Infect. Dis. 52,

1227–1230.

Watnick, P., Kolter, R., 2000. Biofilm, city of microbes. J. Bacteriol. 182, 2675–2679.

Weinrick, B., Dunman, P.M., Mcaleese, F., Murphy, E., Projan, S.J., Fang, Y., Novick, R.P.,

2004. Effect of mild acid on gene expression in Staphylococcus aureus. J. Bacteriol. 186,

8407–8423.

Weng, Y.-M., Chen, M.-J., Chen, W., 1999. Antimicrobial food packaging materials from

poly(ethylene-co-methacrylic acid). LWT - Food Sci. Technol. 32, 191–195.


250

Wertheim, H.F.L., Melles, D.C., Vos, M.C., van Leeuwen, W., van Belkum, A., Verbrugh,

H.A., Nouwen, J.L., 2005. The role of nasal carriage in Staphylococcus aureus infections.

Lancet Infect. Dis. 5, 751–762.

White, G.C., 1999. Handbook of chlorination and alternative disinfectants, 4th ed. John Wiley

and Sons, New York.

Wieneke, a a, Roberts, D., Gilbert, R.J., 1993. Staphylococcal food poisoning in the United

Kingdom , 1969-90. Epidemiol. Infect. 110, 519–531.

Wirtanen, G., Salo, S., 2003. Disinfection in food processing – efficacy testing of disinfectants.

Rev. Environ. Sci. Bio/Technology 2, 293–306.

Woolaway, M.C., Bartlett, C.L., Wieneke, A.A., Gilbert, R.J., Murrell, H.C., Aureli, P., 1986.

International outbreak of staphylococcal food poisoning caused by contaminated lasagne.

J. Hyg. (Lond). 96, 67–73.

Xing, K., Chen, X.G., Kong, M., Liu, C.S., Cha, D.S., Park, H.J., 2009. Effect of oleoyl-

chitosan nanoparticles as a novel antibacterial dispersion system on viability, membrane

permeability and cell morphology of Escherichia coli and Staphylococcus aureus.

Carbohydr. Polym. 76, 17–22.

Xing, M., Shen, F., Liu, L., Chen, Z., Guo, N., Wang, X., Wang, W., Zhang, K., Wu, X., Li, Y.,

Sun, S., Yu, L., 2012. Antimicrobial efficacy of the alkaloid harmaline alone and in

combination with chlorhexidine digluconate against clinical isolates of Staphylococcus

aureus grown in planktonic and biofilm cultures. Lett. Appl. Microbiol. 54, 475–482.

Xu, H., Zou, Y., Lee, H.-Y., Ahn, J., 2010. Effect of NaCl on the biofilm formation by

foodborne pathogens. J. Food Sci. 75, M580–585.

Yang, Z.-Q., Jiao, X.-A., Zhou, X.-H., Cao, G.-X., Fang, W.-M., Gu, R.-X., 2008. Isolation and

molecular characterization of Vibrio parahaemolyticus from fresh, low-temperature

preserved, dried, and salted seafood products in two coastal areas of eastern China. Int. J.


Yu, D., Zhao, L., Xue, T., Sun, B., 2012. Staphylococcus aureus autoinducer-2 quorum sensing

decreases biofilm formation in an icaR-dependent manner. BMC Microbiol. 12, 288.

Zabala, A.J., Font, M.O., Gallastegi, P.M., Ibarbia, E.S., Juaristi, A., Santa, L., Rodríguez, M.,

2011. Situación de los desinfectantes de uso ambiental y en industria alimentaria

registrados en España tras la publicación de la Directiva 98/8/CE. Rev. Esp. Salud Publica

85, 175–188.

Zeng, X., Tang, W., Ye, G., Ouyang, T., Tian, L., Ni, Y., Li, P., 2010. Studies on disinfection

mechanism of electrolyzed oxidizing water on E. coli and Staphylococcus aureus. J. Food

Sci. 75, M253–260.

Zhang, L., Pornpattananangku, D., Hu, C.-M.J., Huang, C.-M., 2010. Development of

nanoparticles for antimicrobial drug delivery. Curr. Med. Chem. 17, 585–594.

Zmantar, T., Kouidhi, B., Miladi, H., Mahdouani, K., Bakhrouf, A., 2010. A microtiter plate

assay for Staphylococcus aureus biofilm quantification at various pH levels and hydrogen

peroxide supplementation. New Microbiol. 33, 137–145.

Zwietering, M.H., Jongenburger, I., Rombouts, F.M., Riet, K., 1990. Modeling of the bacterial

growth curve. Appl. Environ. Microbiol. 56, 1875–81.

List of original publications

251

List of original publications

The thesis is based on the following original publications:

1. Vázquez-Sánchez, D., López-Cabo, M., Saá-Ibusquiza, P., Rodríguez-Herrera, J.J.,

2012. Incidence and characterization of Staphylococcus aureus in fishery products

marketed in Galicia (Northwest Spain). Int. J. Food Microbiol. (IF: 3.425; Q: Q1;

Year: 2012, JCR) 157, 286-296. ISSN: 0168-1605

2. Vázquez-Sánchez, D., Habimana, O., Holck, A., 2013. Impact of food-related

environmental factors on the adherence and biofilm formation of natural

Staphylococcus aureus isolates. Curr. Microbiol. (IF: 1.520; Q: Q4; Year: 2012,

JCR) 66, 110-121. ISSN: 0343-8651

3. Vázquez-Sánchez, D., Cabo, M.L., Ibusquiza, P.S., Rodríguez-Herrera, J.J., 2014.

Biofilm-forming ability and resistance to industrial disinfectants of Staphylococcus

aureus isolated from fishery products. Food Control (IF: 2.738; Q: Q1; Year: 2012,

JCR) 39, 8-16. ISSN: 0956-7135

4. Vázquez-Sánchez, D., Cabo, M.L., Rodríguez-Herrera, J.J. Single and sequential

application of electrolyzed water with benzalkonium chloride or peracetic acid for

removal of Staphylococcus aureus biofilms. J. Food Saf. (under review).

5. Vázquez-Sánchez, D., Cabo, M.L., Rodríguez-Herrera, J.J. Antimicrobial activity of

essential oils against Staphylococcus aureus biofilms. J. Food Saf. (under review).

253

Agradecimientos / Acknowledgements


255


Primero, quiero agradecer la labor del Dr. Juan José Rodríguez Herrera y de la Dra.

Marta López Cabo en la supervisión de esta tesis, así como en mi propia formación

científica en el ámbito de la Microbiología y Seguridad Alimentaria. Hacéis un tándem

perfecto del que tomé buena nota. Agradecer también a la Dra. Paloma Morán Martínez

su colaboración y ayuda en la publicación de esta tesis en la Universidad de Vigo.

Además, quiero darles mi más sincera gratitud a todos los compañeros y compañeras

que conocí en el IIM durante la realización de esta tesis. Gracias por vuestro apoyo

durante todos estos años. Sonia, Eva e Vanessa, foi un pracer traballar ó voso carón,

compartindo campá, chisqueiro, pipetas e emisora de radio: sempre facedes máis

sinxelo o traballo dos demais. Paula, gracias por todos los momentos inolvidables en el

laboratorio. Marta, Alberto, Teresa, Graciela y Ana, fue una suerte trabajar juntos y

espero tomar más cafés con vosotros. Pedro, ànim amb la teva tesis, que als propers

pintxos convides tu. Gabriel, Laura, Lola, Javier y Pilar, gracias por vuestra agradable

(aunque breve) compañía. Diana, Natalia, Adelaida, Lidia, Noemí, Maider y Jesús,

aprendimos mucho juntos y vuestra presencia en el laboratorio fue muy gratificante.

Gracias también al resto del personal del IIM por vuestra ayuda durante mi estancia,

especialmente a los departamentos de Ecología y Biodiversidad Marina y Biología y

Fisiología Larvaria de Peces, por dejarnos usar amablemente vuestro transiluminador; a

Juan Luis, por su excelente asesoramiento informático; a Luisa, por dejarme participar

en actividades de divulgación científica tan satisfactorias; a Frank y demás compañeros

de las clases de inglés; a Estrella, por su compañía y generosidad, y por mantener el

laboratorio siempre limpio y ordenado; y a todos los colegas de pachanga que siempre

me dieron su mejor pase (Elsi, Quitos, Alhambra, Miguel, Javi, Fer, Waldo, Jorge,

Gabriel, Mar, etc.).

Special thanks to Dr. Askild Holck and Dr. Olivier Habimana for their contributions

to the present doctoral thesis. It was a pleasure work together and I will always

appreciate your excellent advices. Thanks also to all people that I met at Nofima,


256

particularly to Monika, Diego, Nebojsa, Natthorn, Anne, Ulrike, etc., for all the

unforgettable moments that you have given me, and to my laboratory colleagues Tone

Mari, Signe, Birgitte, etc., for their inestimable assistance during my experiments.

Por otra parte, agradecer la inmensa paciencia y cariño de mi familia durante esta

larga travesía. Especialmente a mis padres Miguel y Fevi, a los que nunca seré capaz de

recompensar por su esfuerzo en darme unos principios y una educación digna, y por su

infinita paciencia. También a mi hermano Isaac, que siempre me sirvió de referencia

para crecer como persona. A mis abuelos Toncho y Lila, porque de mayor quisiera

llegar a ser como vosotros, gracias por vuestro apoyo y generosidad. A mi abuela María,

a la que siempre recordaré con cariño cuando vaya por Entrimo. Ó meu tío Xurxo, por

tódalas revistas do National Geographic e coleccións de A Nosa Terra das que tanto

gocei aprendendo. A mi madrina-bióloga Yadira, que me dio una infancia inolvidable.

A toda la primada coruñesa, que no son pocos y que a cada cual mejor. A mi familia de

Mallorca, especialmente a mi ahijada Ángela, que tanto eché de menos durante estos

años de tesis. A Liliana, por regalarme esta maravillosa portada para mi tesis. Y también

a la familia de Vero, por acogerme como uno más y tratarme siempre tan bien.

Muchas gracias a mis amigos y amigas de Vigo (y comarca) por todas las comidas y

cenas, fiestas y conciertos, excursiones y viajes, o simples paseos y quedadas. Óscar,

Paula, Raquel, Marta, Nuria, José, Plato, Carmen, Alba, Sandra, Roi, Adri, Gene, Almu,

María... ¡¡sois los mejores!! Siempre sabéis como hacerme pasar momentos

legen...darios. Y Óscar, ve preparando la mochila y los variformes que pronto saldremos

a hacer el Camino de nuevo.

Y finalmente quería agradecerle eternamente a la persona responsable de que tenga

hoy esta Tesis Doctoral entre mis manos. Vero, mi amor, gracias por quererme todos

estos años, por animarme, por apoyarme, por hacer que cada día a tu lado sea

inolvidable. Sin ti, esto nunca hubiese sucedido. Gracias, preciosa.

Assessment of control strategies against Staphylococcus aureus ...

Documents

Transcript of Assessment of control strategies against Staphylococcus aureus ...