assessment of control strategies...
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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
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)
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
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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
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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
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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. Material and Methods ....................................................................................... 157
4.2.1. Bacterial strains .......................................................................................... 157
4.2.2. Antibacterial agents .................................................................................... 157
4.2.3. Conditions for biofilm formation................................................................ 158
4.2.4. Single application of electrolyzed water (EW) ........................................... 159
4.2.5. Sequential application treatments ............................................................... 160
4.2.6. Statistical analysis ....................................................................................... 161
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. Material and Methods ....................................................................................... 185
5.2.1. Bacterial strain ............................................................................................ 185
5.2.2. Antibacterial agents .................................................................................... 185
5.2.3. Conditions for biofilm formation................................................................ 186
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
5.2.5. Statistical analysis ....................................................................................... 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
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Resumen
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Resumen
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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
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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
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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
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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.
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Abstract
16
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.
21
22
23
General Introduction
24
General Introduction
25
General Introduction
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).
General Introduction
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).
General Introduction
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).
General Introduction
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.
General Introduction
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
General Introduction
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).
General Introduction
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).
General Introduction
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.
General Introduction
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).
General Introduction
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
General Introduction
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).
General Introduction
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).
General Introduction
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
General Introduction
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.
General Introduction
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
General Introduction
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
General Introduction
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).
General Introduction
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
General Introduction
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
General Introduction
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).
General Introduction
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
General Introduction
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
General Introduction
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.
General Introduction
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)
General Introduction
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
General Introduction
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
General Introduction
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).
General Introduction
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)
General Introduction
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).
54
Justificación y Objetivos
56
Justificación y Objetivos
57
Justificación y Objetivos
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.
Justificación y Objetivos
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.
59
60
61
Justification and Objectives
62
Justification and Objectives
63
Justification and Objectives
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:
Justification and Objectives
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.
65
66
67
Chapter 1. Incidence and characterization of
Staphylococcus aureus in fishery products
marketed in Galicia (Northwest Spain)
68
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.,
Incidence and characterization of S. aureus in fishery products
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
Incidence and characterization of S. aureus in fishery products
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.
Incidence and characterization of S. aureus in fishery products
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).
Incidence and characterization of S. aureus in fishery products
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.
Incidence and characterization of S. aureus in fishery products
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.
Incidence and characterization of S. aureus in fishery products
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
Incidence and characterization of S. aureus in fishery products
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.
Incidence and characterization of S. aureus in fishery products
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.
Incidence and characterization of S. aureus in fishery products
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
Incidence and characterization of S. aureus in fishery products
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.
Incidence and characterization of S. aureus in fishery products
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
Incidence and characterization of S. aureus in fishery products
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.
94
95
Chapter 2. Impact of food-related
environmental factors on the adherence and
biofilm formation of natural Staphylococcus
aureus isolated from fishery products
96
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.
Impact of food-related environmental factors on the biofilm formation of S. aureus
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:
Impact of food-related environmental factors on the biofilm formation of S. aureus
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
Impact of food-related environmental factors on the biofilm formation of S. aureus
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
Impact of food-related environmental factors on the biofilm formation of S. aureus
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.
Impact of food-related environmental factors on the biofilm formation of S. aureus
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
Impact of food-related environmental factors on the biofilm formation of S. aureus
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
.
Impact of food-related environmental factors on the biofilm formation of S. aureus
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
Impact of food-related environmental factors on the biofilm formation of S. aureus
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 =
Impact of food-related environmental factors on the biofilm formation of S. aureus
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
Impact of food-related environmental factors on the biofilm formation of S. aureus
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.
Impact of food-related environmental factors on the biofilm formation of S. aureus
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.
Impact of food-related environmental factors on the biofilm formation of S. aureus
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).
123
124
125
Chapter 3. Biofilm-forming ability and
resistance to industrial disinfectants of
Staphylococcus aureus isolated from fishery
products
126
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.
Biofilm-forming ability and resistance to industrial disinfectants of S. aureus
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.
Biofilm-forming ability and resistance to industrial disinfectants of S. aureus
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
Biofilm-forming ability and resistance to industrial disinfectants of S. aureus
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
Biofilm-forming ability and resistance to industrial disinfectants of S. aureus
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).
Biofilm-forming ability and resistance to industrial disinfectants of S. aureus
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.
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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.
Biofilm-forming ability and resistance to industrial disinfectants of S. aureus
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.
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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
Biofilm-forming ability and resistance to industrial disinfectants of S. aureus
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.
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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
Biofilm-forming ability and resistance to industrial disinfectants of S. aureus
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
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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
Biofilm-forming ability and resistance to industrial disinfectants of S. aureus
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
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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
Biofilm-forming ability and resistance to industrial disinfectants of S. aureus
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
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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.
Biofilm-forming ability and resistance to industrial disinfectants of S. aureus
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
151
152
153
Chapter 4. Single and sequential application
of electrolyzed water with benzalkonium
chloride or peracetic acid for removal of
Staphylococcus aureus biofilms
154
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
Single and sequential application of EW with BAC or PAA for removal of S. aureus biofilms
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. Material and Methods
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.
4.2.3. 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, 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
Single and sequential application of EW with BAC or PAA for removal of S. aureus biofilms
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.
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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
Single and sequential application of EW with BAC or PAA for removal of S. aureus biofilms
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
4.2.6. 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.
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.
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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).
Single and sequential application of EW with BAC or PAA for removal of S. aureus biofilms
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).
Figure 4.2. Logarithmic reduction (LR) in the number of viable cells of 48-h-old biofilms of S.
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.
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164
Figure 4.3. Logarithmic reduction (LR) in the number of viable cells of 48-h-old biofilms of S.
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):
Single and sequential application of EW with BAC or PAA for removal of S. aureus biofilms
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).
Figure 4.4. Logarithmic reduction (LR) in the number of viable cells of 48-h-old biofilms of S.
aureus St.1.01 after a double sequential application of NEW for 5 min each.
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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.
Single and sequential application of EW with BAC or PAA for removal of S. aureus biofilms
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).
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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.
Single and sequential application of EW with BAC or PAA for removal of S. aureus biofilms
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
Single and sequential application of EW with BAC or PAA for removal of S. aureus biofilms
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,
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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,
Single and sequential application of EW with BAC or PAA for removal of S. aureus biofilms
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.
Single and sequential application of EW with BAC or PAA for removal of S. aureus biofilms
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.
C N Y Ŷ Coefficients t Model
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 Expected average value 5.1447
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
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).
Single and sequential application of EW with BAC or PAA for removal of S. aureus biofilms
177
Table Ap.3. Sequence BAC1-NEW2.
C N Y Ŷ Coefficients t Model
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 5.172 5.149 Expected average value 5.1490
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
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
178
Table Ap.4. Sequence NEW1-PAA2.
C N Y Ŷ Coefficients t Model
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 5.134 5.149 Expected average value 5.1490
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
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).
Single and sequential application of EW with BAC or PAA for removal of S. aureus biofilms
179
Table Ap.5. Sequence PAA1-NEW2.
C N Y Ŷ Coefficients t Model
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 5.172 5.206 Expected average value 5.2057
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
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).
180
181
Chapter 5. Antimicrobial activity of essential
oils against Staphylococcus aureus biofilms
182
Antimicrobial activity of essential oils against S. aureus biofilms
183
Antimicrobial activity of essential oils against
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 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).
Antimicrobial activity of essential oils against S. aureus biofilms
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. Material and Methods
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)
5.2.3. 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 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
Antimicrobial activity of essential oils against S. aureus biofilms
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
Antimicrobial activity of essential oils against S. aureus biofilms
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.
5.2.5. 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.
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.
Antimicrobial activity of essential oils against S. aureus biofilms
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
Antimicrobial activity of essential oils against S. aureus biofilms
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.
Antimicrobial activity of essential oils against S. aureus biofilms
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
Antimicrobial activity of essential oils against S. aureus biofilms
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
Antimicrobial activity of essential oils against S. aureus biofilms
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.
201
202
203
General Discussion
204
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.
210
211
Conclusiones Generales
212
Conclusiones Generales
213
Conclusiones Generales
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.
Conclusiones Generales
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.
Conclusiones Generales
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.
216
217
General Conclusions
218
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.
222
223
Bibliografía / References
224
Bibliografía / References
225
Bibliografía / References
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
Bibliografía / References
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.
Bibliografía / References
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
Microbiol. 13, 20–26.
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.
Microbiol. 60, 35–42.
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.
Bibliografía / References
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.
CDC, 1976. Morbidity and mortality weekly report 25, 317–318.
CDC, 1983. Morbidity and mortality weekly report 32, 183–184.
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).
CEN, 2005. European standard EN 1040: Chemical disinfectants and antiseptics - Quantitative
suspension test for evaluation of basic bactericidal activity of chemical disinfectants and
antiseptics. Test method Requir. (phase 1).
CEN, 2009. European standard EN 1276: Chemical disinfectants and antiseptics - Quantitative
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.
Bibliografía / References
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.
Bibliografía / References
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.
Bibliografía / References
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.
Bibliografía / References
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
Microbiol. Rev. 36, 990–1004.
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.
Bibliografía / References
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
Microbiol. 6, 484–488.
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.
Bibliografía / References
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.
Bibliografía / References
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.
Bibliografía / References
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.
Bibliografía / References
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
Bibliografía / References
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.
Microbiol. 8, 317–327.
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.
Bibliografía / References
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.
Bibliografía / References
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.
Bibliografía / References
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.
Bibliografía / References
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.
Microbiol. Rev. 17, 218–234.
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.
Bibliografía / References
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.
Bibliografía / References
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.
Bibliografía / References
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.
Food Microbiol. 28, 418–425.
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.
Bibliografía / References
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.
Food Control 26, 98–106.
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.
Bibliografía / References
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.
Food Control 31, 572–585.
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.
Bibliografía / References
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.
Bibliografía / References
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.
Bibliografía / References
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.
Food Microbiol. 125, 279–285.
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
Agradecimientos / Acknowledgements
255
Agradecimientos / Acknowledgements
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,
Agradecimientos / Acknowledgements
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.