Caracterización de proteínas con actividad antifúngica producidas

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INTRODUCCIÓN
I. INTRODUCCIÓN
I.1. POBLACIÓN FÚNGICA EN ALIMENTOS
Las condiciones ambientales en que se procesan o se almacenan diversos tipos de
alimentos favorecen la proliferación en su superficie de mohos que tienen la capacidad de
adaptarse y desarrollarse en sustratos muy diversos. Las especies de mohos aisladas con
mayor frecuencia en alimentos pertenecen a los géneros Penicillium, Aspergillus,
Fusarium y Eurotium, aunque las características intrínsecas de cada tipo de producto
condicionan el desarrollo de unas especies u otras (Filtenborg y col., 1996). Así, frutas y
verduras frescas suelen alterarse durante su almacenamiento por P. expansum, P.
crustosum, P. solitum, P. digitatum o P. italicum (Pitt y Hocking, 1997; Moss, 2008), y los
cereales en cambio pueden ser colonizados por P. polonicum, P. verrucosum, P.
aurantiogriseum, P. viridicatum, A. flavus, A. niger o A. versicolor (Filtenborg y col.,
1996).
En productos cárnicos madurados y quesos es frecuente el desarrollo una población
fúngica superficial durante la maduración que puede tener efectos deseables, ya que
contribuye en gran medida a la formación del sabor, aroma o textura del producto acabado
(Kinsella y Hwang, 1976; Martín y col., 2004, 2006), e incluso puede ejercer un efecto
protector frente a otros microorganismos no deseados (Geisen y col., 1992; Leistner, 1994;
Singh y Dincho, 1994; Berwal y Dincho, 1995; Molimard y col., 1995; Decker y Nielsen,
2005). Por ello, en quesos es frecuente el uso de determinadas cepas de mohos como P.
roqueforti o P. camemberti como cultivos iniciadores (Nielsen y col., 1998), y se ha
propuesto la elaboración de este tipo de cultivos para su utilización en productos cárnicos
madurados a partir de cepas no toxigénicas de P. chrysogenum (Benito y col., 2005; Martín
y col., 2006), Penicillium olsonii (López-Díaz y col., 2001) o Penicillium camemberti
(Bruna y col., 2003).
Pero el desarrollo de mohos no siempre es deseable en alimentos. En quesos se han
descrito cepas alterantes de P. commune y P. nalgiovense que pueden conducir a un
deterioro en las características sensoriales (Filtenborg y col., 1996), debido normalmente a
la liberación de exoenzimas durante su desarrollo. Estos enzimas pueden continuar su
actividad independientemente de que se produzca la eliminación del micelio, causando
desde olores o coloraciones desagradables hasta la completa desintegración de la estructura
del alimento. No obstante, el aspecto más preocupante respecto del desarrollo de una
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población fúngica incontrolada en alimentos es su capacidad para producir micotoxinas. El
desarrollo de mohos en quesos puede conllevar la producción de micotoxinas (Marth y
Yousef, 1991), muchas de las cuales han sido detectadas en estos productos (Lafont y col.,
1979; Le Bars, 1979; Northolt y col., 1980; Jarvis, 1983; Siemens y Zawistowski, 1993).
De manera similar, en productos cárnicos madurados la población microbiana superficial
está constituida fundamentalmente por mohos, muchos de los cuales son toxigénicos
(Bullerman y col., 1969; Escher y col., 1973; Halls y Ayres, 1973; Wu y col., 1974;
Leistner, 1984; Núñez y col., 1996; López-Díaz y col., 2001).
I.2. CONSECUENCIAS DEL DESARROLLO DE MOHOS EN PRODUCTOS CÁRNICOS
MADURADOS
Con respecto a la población fúngica que puede colonizar la superficie de productos
cárnicos, existen tanto mohos que contribuyen en gran medida a la formación del sabor,
aroma o textura del producto acabado, como especies que pueden deteriorar las
características sensoriales o producir micotoxinas.
I.2.1. EFECTOS BENEFICIOSOS
La población fúngica que coloniza los productos cárnicos madurados presenta
diversas actividades metabólicas que pueden mejorar las características sensoriales del
producto. Así, la presencia de mohos puede ofrecer actividad antioxidante, lipolítica,
proteolítica, contribuir al aroma y sabor del producto e incluso ejercer un efecto protector
por su actividad antimicrobiana (Figura I1).
El efecto antioxidante de los mohos se atribuye a que la mayoría de las cepas
aisladas de jamón poseen una actividad catalasa moderada (Núñez, 1995), destruyendo
peróxidos y reduciendo las reacciones oxidativas (Burgos, 1981), por lo que disminuye el
enranciamiento y contribuyen a la estabilización del color del producto (Liepe, 1982;
Smith y Palumbo, 1983; Geisen, 1993). Además, muchos mohos producen compuestos con
acción antioxidante (Ludemann y col., 2004).
Por otra parte, numerosas especies de mohos pertenecientes al género Penicillium
poseen exoenzimas con actividad lipolítica que pueden incrementar la concentración de
ácidos grasos libres y compuestos volátiles en jamón (Huerta y col., 1987; Molina y col.,
1991) y embutidos (Toledo y col., 1996), contribuyendo así al desarrollo del sabor y aroma
del producto. Se ha observado actividad lipolítica en mohos del género Penicillium
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aislados de jamón ibérico (Núñez, 1995; Alonso, 2004) y de embutidos (Selgas y col.,
1999; Ludemann y col., 2004).
Figura I1. Efectos beneficiosos por el desarrollo de mohos durante la maduración del
jamón (Núñez, 1995).
Adicionalmente, aunque la proteolisis en los productos cárnicos se ha asociado
generalmente a la actividad de los enzimas presentes en el tejido muscular (Toldrá y col.,
1992; Sárraga y col., 1993), algunas cepas aisladas de jamón curado producen una intensa
hidrólisis de proteínas miofibrilares y un gran incremento de aminoácidos libres cuando se
inoculan en carne de cerdo estéril y productos cárnicos madurados (Rodríguez y col., 1998;
Martín y col., 2001, 2002, 2004). En este sentido, se ha demostrado una intensa actividad
proteolítica de una cepa de P. chrysogenum (Rodríguez y col., 1998; Martín y col., 2001,
2002; Benito y col., 2003; Alonso, 2004), a partir de la cual se ha purificado una proteasa
muy activa en las condiciones de maduración de diferentes productos cárnicos (Benito y
col., 2002). También se han aislado de embutidos mohos muy proteolíticos de los géneros
Mucor y Penicillium (Trigueros y col., 1995), capaces de incrementar la concentración de
aminoácidos libres en estos productos (Bruna y col., 2001).
Los mohos pueden favorecer la formación del aroma característico de los productos
curados por diversas vías. En primer lugar, los aminoácidos liberados, además de potenciar
el sabor de los alimentos (Suyama y Shimuzu, 1982; Nishimura y Kato, 1988), pueden
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intervenir en reacciones de condensación de Maillard (Shahidi y col., 1986; Mottram y
Whitfield, 1995) y de degradación de Strecker (Barbieri y col., 1992), en las que se pueden
formar compuestos responsables del aroma del jamón curado (García y col., 1991; Ruiz y
col., 1998; Whitfield, 1992).
Finalmente, los mohos pueden tener un efecto protector frente a microorganismos
patógenos o alterantes (Geisen y col., 1992; Leistner, 1994; Singh y Dincho, 1994; Berwal
y Dincho, 1995). Así, el 75% de los mohos aislados en jamón ibérico inhiben el
crecimiento de Staphylococcus aureus (Núñez y col., 1996), y diversas especies no
toxigénicas de mohos tienen la capacidad de inhibir la producción de micotoxinas e incluso
de destruir las formadas por otras especies toxigénicas (Mislivec y col. 1988). No obstante,
este efecto podría deberse a actividades tan poco deseables como la producción de toxinas
o de antibióticos, o bien a mecanismos más deseables como a interferencias metabólicas o
a la liberación de proteínas antimicrobianas.
I.2.2. EFECTOS PERJUDICIALES
El desarrollo de ciertos mohos, como los géneros Mucor y Rhizopus, puede
provocar un aspecto no deseable al jamón o embutidos (Figura I2). Por otro lado, los
mohos con gran capacidad lipolítica pueden ocasionar olores anómalos (Sanz, 1986). De
hecho, se ha identificado a P. commune como el responsable del “defecto del ácido
fénico”, caracterizado por conferir al jamón un fuerte olor a fenol (Spotti y col., 1988).
Figura I2. Contaminación fúngica superficial en jamón curado.
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No obstante, el aspecto más preocupante respecto al desarrollo de una población
fúngica superficial en estos productos es su capacidad para producir micotoxinas. Las
micotoxinas son metabolitos secundarios tóxicos que al ser ingeridos en pequeñas
cantidades, pueden ocasionar diferentes patologías relacionadas con inmunosupresión,
carcinogénesis, neurotoxicidad, etc. (Samson y col., 1995; Richard, 2007). Existen
micotoxinas que sólo son producidas por un grupo reducido de mohos, y otras generadas
por especies muy diversas incluso de géneros diferentes (Frisvad y Thrane, 1995). En este
sentido, se han descrito especies como Penicillium griseofulvum capaces de producir gran
variedad de micotoxinas (Frisvad y Filtenborg, 1990). No obstante, el perfil y la
concentración de micotoxinas que puede producir un moho, así como la posterior difusión
al interior del alimento, no sólo dependen de la naturaleza del microorganismo sino
también de la composición del alimento y del procesado al que esté sometido. Además,
estos metabolitos son muy resistentes a los tratamientos físicos y químicos, por lo que una
vez que las micotoxinas están en el alimento pueden permanecer en él incluso después del
procesado (Osborne y col., 1996; Castellá y col., 1998; Katta y col., 1999). Muchos mohos
colonizadores de alimentos son capaces además de producir antibióticos como la penicilina
(Andersen y Frisvad, 1994), lo que supone un riesgo para los consumidores alérgicos a este
antibiótico (Bremmelgaard, 1998).
El potencial toxigénico de los mohos aislados de productos cárnicos se ha
establecido en distintos trabajos. Se ha comprobado que el 80% de 1481 aislados del
género Penicillium a partir de embutidos son potencialmente toxigénicos (Eckardt y col.,
1979). Más del 75% de los mohos aislados en jamón ibérico muestran toxicidad o
mutagenicidad en ensayos biológicos (Núñez y col., 1996). Entre estos mohos, el más
frecuente es P. commune, productor de ácido ciclopiazónico (Núñez y col., 1996; Sosa y
col., 2002), destacando también P. restrictum, A. flavus y A. niger. Además, todos los
aislados de P. commune obtenidos de chorizo de Cantimpalos mostraron una alta toxicidad
para larvas de Artemia salina (López-Díaz y col., 2001). Estas especies pueden producir
micotoxinas en los productos cárnicos sobre los que se desarrollan (Núñez y col., 2007),
habiéndose comprobado en embutidos y jamones curados la presencia de citrinina,
patulina, rugulosina, ocratoxina A, ácido ciclopiazónico, citreoviridina, breviamida A,
fumitremorgina B, griseofulvina y verruculógeno TR1 (Leistner, 1984). Además, las
micotoxinas pueden difundir al interior de los productos, donde pueden mantenerse incluso
cuando el micelio se ha eliminado (Leistner, 1984).
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Por todo esto, resulta preocupante el crecimiento incontrolado de mohos
potencialmente toxigénicos en la superficie de los productos cárnicos madurados (Núñez y
col., 1996; Sabater-Vilar y col., 2003) y sería recomendable controlar su desarrollo en este
tipo de productos.
I.3. CONTROL DE MOHOS EN PRODUCTOS CÁRNICOS MADURADOS
Existen numerosos mecanismos para controlar el desarrollo microbiano en distintos
tipos de alimentos. Entre los métodos físicos de control se encuentra la utilización de
tratamientos térmicos, radiaciones ionizantes o ultravioleta, altas presiones, pulsos
eléctricos o de luz, o campos magnéticos. También pueden emplearse agentes químicos,
incluyendo ácidos orgánicos, aceites esenciales, peróxido de hidrógeno, etc., así como el
envasado en atmósferas controladas o modificadas. Estos tratamientos resultan eficaces
para evitar el desarrollo de mohos en determinados alimentos, como frutas, verduras, pan,
etc. No obstante, en los productos cárnicos madurados está limitada la aplicación directa de
fungicidas. Además, las medidas higiénicas para prevenir la contaminación durante la
elaboración se ven desbordadas en los secaderos y bodegas, donde resulta muy difícil
eliminar totalmente la contaminación ambiental. Por otra parte, la inespecificidad de estos
tratamientos evitaría la posible contribución beneficiosa de los mohos a la maduración de
estos productos.
Por todo ello, en productos cárnicos madurados es necesario adoptar estrategias
alternativas que garanticen el desarrollo de los microorganismos deseables y la seguridad
de estos alimentos.
I.3.1. CULTIVOS PROTECTORES
Para controlar la población fúngica que se desarrolla en la superficie de los
productos cárnicos madurados podrían utilizarse cultivos protectores formados por mohos
capaces de inhibir por competición a los mohos no deseables, siempre teniendo en cuenta
que deben ser no toxigénicos y mostrar una actividad metabólica adecuada. No obstante,
esta opción se ve limitada por la evolución de las condiciones ecológicas durante la
maduración, resultando muy poco probable que los mohos seleccionados superen a todas
las especies habituales en la amplísima gama de temperatura y actividad de agua que tiene
lugar durante la maduración (Rodríguez y col., 1994). Una posibilidad radicaría en el
empleo de mohos que dispusieran de algún elemento adicional que favoreciese su
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desarrollo, como la producción de antibióticos. Lógicamente, esta opción se ha descartado
por las posibles reacciones alérgicas en el consumidor o la inducción de resistencia en
otros microorganismos como consecuencia de su empleo continuado.
Otra posible estrategia consiste en el empleo de proteínas antibacterianas como las
bacteriocinas, que están siendo ampliamente utilizadas en otro tipo de alimentos, o de los
microorganismos que las producen. Así, la nisina es una bacteriocina autorizada como
conservante alimentario (Gálvez y col., 2007). Pero este tipo de proteínas, a pesar de tener
un espectro de inhibición muy amplio, no inhibe a los mohos. Por ello, ni las bacteriocinas
ni los microorganismos que las producen pueden ser empleados para evitar el crecimiento
fúngico en alimentos.
Sin embargo, la reciente descripción de proteínas antifúngicas naturales elaboradas
por mohos abre nuevas perspectivas en este sentido. La utilización de cepas de mohos
productoras de proteínas antifúngicas como cultivos protectores en productos cárnicos
madurados, permitiría limitar el desarrollo de mohos toxigénicos sin afectar a los
deseables, contribuyendo así a estandarizar las características del producto.
I.3.2. PROTEÍNAS ANTIFÚNGICAS
La investigación de nuevos agentes antifúngicos ha experimentado un notable
avance en los últimos años como consecuencia del incremento de presentación de micosis
en pacientes inmunodeprimidos por el SIDA o por tratamientos anticancerígenos,
especialmente infecciones por distintas especies de Candida o en aspergilosis invasivas
(Anaissie, 2008; de la Torre y col., 2008; Pemán y Almirante, 2008). Esta búsqueda de
nuevos compuestos antifúngicos se ha extendido también a otros campos como la
agricultura, donde la proliferación de mohos patógenos en los cultivos ocasiona grandes
pérdidas económicas. Así, se han caracterizado nuevas proteínas con actividad antifúngica
(Upadhyay y Srivastava., 2008; Yan y col., 2008) y se han producido plantas transgénicas
que las expresen (Girlanda y col., 2008; Swathi-Anyradha y col., 2008).
En los últimos años se han purificado y caracterizado proteínas con efecto
antifúngico a partir de plantas (Chu y col., 2003; Ng y col., 2003), bacterias (Yang y col.,
2002), mohos (Marx y col., 2007; Meyer, 2008), artrópodos (Tomie y col., 2003), o
reptiles (Gomes y col., 2005), así como también se han diseñado péptidos sintéticos
(Patrzykaty col., 2003).
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El mecanismo de acción de las proteínas antifúngicas es tan variado como los
organismos que las producen (Selitrennikoff, 2001; Theis y Stahl, 2004). Algunos
atraviesan la membrana e interfieren en la síntesis de ADN, ARN o de proteínas, inhiben la
cadena respiratoria celular (Cociancich y col., 1993), disminuyen el ATP (Cociancich y
col., 1993), inactivan los ribosomas fúngicos (Hwu y col., 2000) o provocan la
desporalización de la membrana (Thevissen y col., 1997). Otros en cambio forman canales
y poros en la membrana alterando su permeabilidad y provocando la pérdida de iones y
otros solutos (Thevissen y col., 1999; Lee y col., 2001) o pueden interferir en la síntesis de
componentes esenciales de la pared celular como el -1,3-glucano (Odds y col., 2003), la
quitina (Yang y Gong, 2002; Von der Weid y col., 2003) o el quitosán (Shimosaka y col.,
2005), pudiendo incluso hidrolizarlos (Sela-Buurlage y col., 1993; Graham y Sticklen,
1994). Esto debilita la pared y origina la lisis de la célula.
No obstante, las proteínas antifúngicas de mayor utilidad son las que presentan
mecanismos de acción específicos para la cubierta externa de los hongos (Groll y col.,
1998), ya que suelen ser poco tóxicas para las células de mamíferos. En la interacción de la
proteína con la membrana desempeñan un papel destacado la carga (Hoover y col., 2003) y
el carácter catiónico o hidrofóbico del compuesto (Friedrich y col., 1999; Hancock y
Chapple, 1999). Así, la hidrofobicidad y la estructura en -hélice, se correlacionan con la
toxicidad para las células de mamíferos, mientras que la carga positiva está más
relacionada con la actividad antimicrobiana (Hong y col., 2001). Esto se debe a que las
membranas celulares de bacterias y mohos se componen principalmente de fosfolípidos
aniónicos que favorecen la unión de la proteína antifúngica y su actividad, mientras que
estos fosfolípidos no están presentes en muchos eucariotas superiores o están confinados
en la cara interna de la membrana plasmática, no estando en contacto con el medio
extracelular (Rivas y Andreu, 2003; Theis y Stahl, 2004).
Por otra parte, existe una gran variabilidad en cuanto a la composición de
aminoácidos y estructura secundaria y terciaria de las proteínas antifúngicas, aunque la
mayoría se caracterizan por ser catiónicas y por formar estructuras anfipáticas (Hwang y
Vogel, 1998). Se han descrito cuatro proteínas antifúngicas catiónicas y de bajo peso
molecular producidas por mohos: AFP de Aspergillus giganteus (Nakaya y col., 1990;
Lacadena y col., 1995), PAF de Penicillium chrysogenum (Marx y col., 1995), Anafp de
Aspergillus niger (Lee y col., 1999) y AcAFP de A. clavatus (Skouri-Gargouri y Gargouri,
2008). Todas ellas se caracterizan por su pequeño tamaño (51-58 aminoácidos), su alto
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contenido en residuos de cisteína y su fuerte carácter básico. Además, sus estructuras
terciarias están estabilizadas por tres o cuatro puentes disulfuro y forman estructuras con
un marcado carácter anfipático (Nakaya y col., 1990; Lacadena y col., 1995).
Sin embargo, la aplicación de los mohos productores como cultivos protectores en
alimentos madurados en general y en productos cárnicos en particular exige que dichos
mohos estén adaptados a las condiciones ecológicas de estos alimentos, que no los alteren
y que no sean productores de micotoxinas. Ninguna de las cepas a partir de las que se han
purificado proteínas antifúngicas ha sido obtenida a partir de productos madurados. Por
ello, es esperable que no se adapten a las condiciones ecológicas particulares de este tipo
de productos de manera que puedan competir con los mohos que crecen de forma
espontánea. Además, algunos de estos mohos pertenecen al género Aspergillus, que sólo es
predominante en estos alimentos en las últimas fases de su elaboración/maduración,
cuando la aw es muy baja. Por lo tanto, la utilización de mohos requiere encontrar cepas
adaptadas a este tipo de alimentos y que sean capaces de elaborar las proteínas antifúngicas
en estos productos. En este caso, las obtenidas de especies del género Penicillium,
empleadas como cultivos iniciadores, plantearían menos objeciones que las de Aspergillus,
debido a que este último género se relaciona más frecuentemente con la alteración de
alimentos y la producción de micotoxinas (Perrone y col., 2007).
I.3.3. PROTEÍNAS ANTIFÚNGICAS DE Penicillium chrysogenum
En estudios previos se seleccionaron los aislados no toxigénicos de P. chrysogenum
RP42C y AS51D, procedentes de jamón curado, por su capacidad para inhibir el desarrollo
de mohos toxigénicos frecuentes en alimentos. Adicionalmente, se purificaron dos
proteínas con actividad antifúngica, PgAFP y PgChP, a partir de RP42C y AS51D,
respectivamente (Acosta y col., 2009a).
PgAFP y PgChP se obtuvieron como consecuencia de una búsqueda exhaustiva de
proteínas antifúngicas con carga neta positiva, debido a que el carácter catiónico de este
tipo de proteínas se asocia con una menor toxicidad para las células de mamíferos. Así,
PgAFP y PgChP fueron obtenidas mediante cromatografía líquida para la purificación
rápida de proteínas (FPLC) empleando inicialmente columnas de intercambio catiónico y
posteriormente de filtración en gel (Acosta y col., 2009a).
La proteína PgAFP mostró un amplio espectro de inhibición frente a mohos
toxigénicos aislados de jamón curado, tales como A. flavus, A. niger, A. parasiticus, P.
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griseolfulvum, P. solitum o P. restrictum. En cambio, la proteína PgChP sólo afectó a P.
commune, aunque no se descarta que tenga actividad antifúngica frente a otros mohos no
estudiados (Acosta y col., 2009a, b).
Las proteínas PgAFP y PgChP mostraron un peso molecular muy diferente por
SDS-PAGE, siendo de aproximadamente 9 y 37 kDa, respectivamente.
El punto isoeléctrico de PgAFP fue de 9,2, por lo que la proteína resultó ser muy
catiónica. Por otra parte, el hecho de que PgAFP perdiera actividad frente a algunas cepas
del género Aspergillus tras el tratamiento con α-amilasa, sugirió la posibilidad de algún
tipo de glicosilación en la proteína, hecho que se confirmó mediante una tinción de ácido
periódico-Schiff (PAS; Zacharius y col., 1969) siguiendo el método descrito por Deepak y
col. (2003). Además, la secuencia amino terminal parecía indicar que se trataba de una
proteína antifúngica no descrita hasta el momento. No obstante, el análisis mediante
MALDI-TOF de los fragmentos obtenidos de PgAFP tras la digestión con tripsina no
ofreció demasiada información sobre su composición aminoacídica, por lo que apenas se
obtuvieron datos relativos a la estructura primaria de esta proteína (Acosta y col., 2009b).
También se confirmó la producción de PgAFP en condiciones similares a las de las
etapas iniciales de la maduración. Esto podría suponer para P. chrysogenum RP42C una
ventaja adicional para competir frente al resto de mohos que se desarrollan en la superficie
de los productos cárnicos madurados, lo que sería de gran valor para su utilización como
cultivo protector. Además, PgAFP mostró sensibilidad al tratamiento con enzimas como
tripsina, pepsina y lisozima, lo cual permite ser optimistas respecto a su uso en alimentos,
ya que puede ser degradada en el tracto gastrointestinal (Acosta, 2006).
Por otra parte, aunque la información disponible de PgChP se limita a su peso
molecular y a su capacidad para inhibir a P. commune, dadas las diferencias en la masa y
en el espectro de inhibición entre PgAFP y PgChP, en este trabajo se aborda la
caracterización de ambas proteínas.
I.4. CARACTERIZACIÓN DE PROTEÍNAS
Para el estudio de las proteínas se pueden determinar distintas características, entre
las que destacan el peso molecular, el punto isoeléctrico, la secuencia de aminoácidos y la
del gen que la codifica, así como la sensibilidad a distintos tratamientos (térmicos, pHs,
iones, etc.).
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I.4.1. DETERMINACIÓN DEL PESO MOLECULAR
I.4.1.1. Electroforesis en gel de poliacrilamida
Uno de los métodos más habituales para estimar el peso molecular de una proteína
es la electroforesis en gel de poliacrilamida, que puede utilizarse para separar proteínas en
función de su carga y tamaño en condiciones no desnaturalizantes o nativas (Bonnerjea y
col., 1986), o bien sólo en función del tamaño de sus subunidades, en condiciones
desnaturalizantes (Weber y Osborn, 1969). Muchos autores han utilizado la electroforesis
desnaturalizante tanto para comprobar la pureza como para calcular el peso molecular de
proteínas antifúngicas (Lacadena y col., 1995; Marx y col., 1995; Martínez-Ruiz y col.,
1997; Lee y col., 1999).
La electroforesis en gel de poliacrilamida tiene la ventaja de que puede llevarse a
cabo en cualquier laboratorio ya que la inversión en material e instrumentos es mínima. Sin
embargo, este método puede llevar a errores en el peso molecular estimado. De hecho,
proteínas con modificaciones post-traduccionales presentan una movilidad anómala ya que
estas modificaciones provocan que no se una adecuadamente el SDS en electroforesis
desnaturalizantes (Russ y Poláková, 1973; Bagger y col., 2007). Igualmente, proteínas con
puntos isoeléctricos extremos pueden repeler las cargas negativas del SDS y no migrar
como lo deberían hacer teóricamente en función de su peso molecular (Mondstadt y
Holldorf, 1990).
I.4.1.2. Espectrometría de masas
En los últimos años se ha aplicado una técnica muy conocida en otros campos,
aunque novedosa en el estudio de proteínas: la espectrometría de masas. La mayor parte de
los análisis bioquímicos por espectrometría de masas utilizan la ionización por electrospray
(ESI) o por desorción con láser asistida por matriz (MALDI), junto a un analizador de
tiempo de vuelo (TOF) (Ashcroft, 2005). ESI y MALDI son métodos de ionización
“suave” que producen moléculas cargadas con poca fragmentación, aún tratándose de
muestras de muy alta masa molecular.
Tanto en ESI como en MALDI se protona la muestra a analizar y como resultado se
genera un espectro masa/carga (m/z) que tiene una amplia distribución de múltiples iones
cargados. Este espectro m/z puede ser convertido a un espectro con las masas reales
(Figura I3). Manualmente, la masa real puede ser calculada con la fórmula m/z= (M +
nH+)/n, donde M es la masa molecular de la proteína y n es el número de protones
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asociados a ella. El error asociado a esta técnica suele rondar el 0,01% de la masa
(Ashcroft, 2005).
Figura I3. Determinación del peso molecular mediante espectrometría de masas con
ionización por ESI y MALDI.
Tanto MALDI como ESI son técnicas ampliamente utilizadas para la determinación
del peso molecular de las proteínas, aunque requieren una fuerte inversión en instrumentos,
así como tener amplios conocimientos en su manejo y en la interpretación de los
resultados. A efectos prácticos, la ionización MALDI suele ser más atractiva porque tiene
mayor tolerancia a sustancias químicas o sales presentes en la muestra, reduciendo así la
necesidad de purificación que requiere la ionización ESI (Standing, 2003). Por lo demás,
ambas son técnicas muy valiosas y precisas para determinar el peso molecular de una
proteína.
I.4.2. DETERMINACIÓN DEL PUNTO ISOELÉCTRICO
Las técnicas más frecuentes para el cálculo del punto isoeléctrico de una proteína
suelen basarse en el isoelectroenfoque, que puede llevarse a cabo mediante electroforesis
en gel en una o dos dimensiones, así como mediante electroforesis capilar.
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I.4.2.1. Isoelectroenfoque
El isoelectroenfoque es un tipo de electroforesis que permite separar los
componentes de una muestra en función de su punto isoeléctrico. Se basa en la capacidad
de las moléculas de variar su carga neta en función del pH del medio. La electroforesis se
lleva a cabo añadiendo una mezcla de anfolitos con diferentes puntos isoeléctricos en el
medio, para crear un gradiente de pH entre dos electrodos con carga eléctrica opuesta. Al
aplicar una diferencia de potencial cada anfolito se mueve hacia uno u otro electrodo en
función de su propia carga y se detiene cuando el pH del medio que le rodea es igual a su
punto isoeléctrico. Esta técnica se caracteriza por una gran repetibilidad y resolución
(Fuentes Arderiu y col., 1998).
I.4.2.2. Electroforesis bidimensional
Este tipo de electroforesis consiste en realizar dos separaciones electroforéticas
consecutivas de una misma muestra en función de propiedades diferentes en dos
direcciones distintas. Lo más frecuente es realizar la primera separación por
isoelectroenfoque según el punto isoeléctrico, y la segunda en gel de poliacrilamida con
SDS (PAGE-SDS) según la masa molar (Figura I4). Esta técnica se suele emplear para la
separación de muestras complejas de proteínas, pudiendo calcular puntos isoeléctricos,
masas molares, cuantificar proteínas, etc.
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Figura I4. Electroforesis bidimensional.
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I.4.2.3. Electroforesis capilar por isoelectroenfoque
La electroforesis capilar es una técnica desarrollada en los últimos años que
consiste en realizar técnicas electroforéticas convencionales (electroforesis en gel,
isoelectroenfoque, etc.) en capilares de sílice fundida con un diámetro interno de entre 25 y
75 µm y una longitud que puede oscilar entre 7 y 100 cm. Tanto uno de los extremos del
capilar como los electrodos se encuentran en una disolución tampón. La muestra se añade
cerca del extremo anódico y se aplica una elevada diferencia de potencial al sistema, que
hace que los componentes de la muestra se muevan a lo largo del capilar y pasen a través
de un detector colocado cerca del cátodo (Holland y Sepaniak, 1993; Berg y col., 2008).
Estudios comparativos entre isoelectroenfoque, electroforesis bidimensional y
electroforesis capilar revelan que las tres técnicas proporcionan una información fiable
acerca del punto isoeléctrico (Valenzuela y col., 1999). Sin embargo, la electroforesis
capilar aporta ventajas como la necesidad de una mínima cantidad de muestra, alta
reproducibilidad y rapidez en los análisis comparativos. A pesar de esto, la electroforesis
bidimensional es una de las técnicas más utilizadas en laboratorios de proteómica para la
separación de mezclas de proteínas y la determinación del punto isoeléctrico.
I.4.3. IDENTIFICACIÓN DE PROTEÍNAS
En la actualidad, en la identificación de proteínas se utilizan diversos
procedimientos, entre los que se incluyen la secuenciación del extremo N-terminal,
detección con anticuerpos específicos, composición de aminoácidos, etc. Todos estos
métodos generalmente son lentos, laboriosos o caros (Gil, 2004).
Hasta hace poco tiempo, para obtener la secuencia aminoacídica de una proteína la
técnica más utilizada era la degradación de Edman. Pero la espectrometría de masas,
debido a su rapidez y elevada sensibilidad, se ha convertido en el método de elección para
la identificación de proteínas a gran escala, que es el primer paso para el estudio de
proteínas de distintos organismos (Russeth y col., 2006; Samyn y col., 2006). Además, esta
técnica también permite la caracterización de modificaciones post-traduccionales que
presentan relevancia fisiológica, tales como la glicosilación y la fosforilación (Claverol y
col., 2003; Morelle y col., 2006). El análisis de las proteínas mediante espectrometría de
masas ha sido posible gracias al desarrollo de varios métodos de ionización suave como
ESI o MALDI para convertir biomoléculas grandes, polares y no volátiles, en iones en fase
gaseosa.
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Esta técnica implica varios pasos: la conversión de los péptidos en fase gaseosa
mediante técnicas de ionización suave como MALDI o ESI; la separación de los iones
según su m/z en un analizador de masas (por ejemplo un analizador de tiempo de vuelo); la
fragmentación opcional de los iones seleccionados, por ejemplo mediante una disociación
inducida por colisión (CID) en un espectrómetro de masas en tándem, combinando dos
analizadores diferentes; y por último la medida de las masas en un detector obteniendo un
espectro de masas que refleja la abundancia de los iones frente a su valor m/z.
Para la identificación de proteínas se han desarrollado dos estrategias que se
encuentran representadas esquemáticamente en la Figura I5: identificación mediante
mapeo peptídico o huella peptídica (PMF: peptide mass fingerprinting) e identificación
mediante etiqueta de secuencia o peptídica, que conlleva la fragmentación de péptidos y la
obtención de la secuencia aminoacídica total o parcial utilizando un espectrómetro de
masas en tándem.
I.4.3.1. Mapeo peptídico o huella peptídica.
El mapeo peptídico o huella peptídica es una técnica utilizada rutinariamente para
identificar proteínas de forma rápida, normalmente a partir de geles de SDS-PAGE o 2DPAGE, que se realiza en un espectrómetro de masas tipo MALDI-TOF o ESI-TOF. La
huella peptídica de una determinada proteína es un conjunto de péptidos generados
mediante la digestión con una proteasa específica. Estas masas peptídicas experimentales
son comparadas con las masas peptídicas teóricas de proteínas presentes en bases de datos.
Para ello se han desarrollado diversos algoritmos (PEPSEA, PROFOUND, MS-FIT,
PEPTIDENT, etc) disponibles en Internet. Para la identificación correcta de la proteína se
requiere que las masas de un gran número de péptidos coincidan con sus masas teóricas, y
que cubran parte de la secuencia de la proteína de la base de datos. Además, si la cantidad
de proteína es pequeña, el número de péptidos observados puede ser reducido y por tanto la
proteína no se puede identificar con fiabilidad.
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Figura I5. Identificación de proteínas mediante espectrometría de masas.
I.4.3.2. Secuencia peptídica o etiqueta peptídica.
La secuencia peptídica es una estrategia para la identificación de proteínas no
incluidas en las bases de datos o que presentan identificaciones ambiguas mediante
MALDI/ESI-TOF.
Los espectrómetros de masas en tándem MS/MS permiten también la determinación
de la secuencia de aminoácidos. Para ello se selecciona un ión por la masa en un primer
espectrómetro y se fragmenta por colisión con un gas. Posteriormente los fragmentos
obtenidos se analizan en un segundo espectrómetro. Puede utilizarse con una fuente de
ionización tipo MALDI o ESI. A partir de la protonación de los péptidos, estos iones
pueden ser seleccionados y sometidos a una disociación inducida por colisión (CID), en la
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cual interaccionan con el gas de colisión (normalmente nitrógeno o argón), y tiene lugar la
fragmentación a lo largo del esqueleto peptídico. Los espectros de fragmentación MS/MS
proporcionan información sobre la identidad y posición del aminoácido en el péptido. Por
tanto es posible adquirir la secuencia completa del péptido si se parte de una fragmentación
de buena calidad.
No obstante a veces sólo se obtiene una secuenciación parcial del péptido. La
información sobre la secuencia parcial, la masa del péptido, y la localización exacta de los
aminoácidos, pueden permitir la identificación de la proteína en las bases de datos. A esta
aproximación se le denomina identificación por etiqueta peptídica. Existen diversas
herramientas bioinformáticas que se pueden utilizar para la interpretación de la
fragmentación de un péptido (PEPSEA, SEQUEST, PEPFRAG, MSTAG y MASCOT).
El método de la secuenciación de novo es la estrategia de elección para aquellas
proteínas de las que se desconozca la secuencia del gen que las codifica. El objetivo de esta
aproximación es obtener secuencias completas de aminoácidos que se deduzcan de novo de
los espectros MS/MS (bien mediante interpretación manual o con ayuda de programas
informáticos) y buscar homologías con proteínas presentes en bases de datos. Esta
estrategia permite encontrar proteínas homólogas en otras especies utilizando programas de
búsqueda como BLAST o FASTA.
Finalmente, aunque la proteína de interés pueda no ser identificada o sea una
proteína desconocida, la información sobre las secuencias obtenidas se puede utilizar para
diseñar cebadores que permitan la obtención y clonación del gen que la codifica.
I.5. CARACTERIZACIÓN DEL GEN QUE CODIFICA UNA PROTEÍNA
Caracterizar el gen que codifica una proteína es de gran utilidad porque permite en
muchos casos completar la secuencia de aminoácidos. Adicionalmente, el conocimiento de
la secuencia genética que codifica una proteína permite su clonación en un vector de
expresión y una síntesis más eficiente de la proteína en un organismo distinto del que
normalmente la produce.
I.5.1. BIBLIOTECAS DE ADN COMPLEMENTARIO
La obtención de bibliotecas de ADN complementario o ADNc es una de las
estrategias que se han utilizado con proteínas antifúngicas (Marx y col., 1995). Para ello es
necesario elegir un vector o plásmido que se utilizará como herramienta para la
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construcción de la biblioteca. Los vectores plasmídicos son capaces de clonar fragmentos
de ADNc desde cientos hasta unas 9000 pares de bases, ya que tamaños mayores necesitan
otros vectores como los cósmidos. La elección del plásmido depende de los sitios de
clonación, de restricción y de los lugares de resistencia a antibióticos. Una vez elegido el
vector se une al ADNc sintetizado. Posteriormente se introducen en el microorganismo
competente elegido, formándose los transformantes que constituirán la biblioteca. Los
clones relacionados con la síntesis de la proteína se detectan mediante una sonda de ADN
(Redkar y col., 1996) que puede elaborarse a partir de la secuencia del extremo amino
terminal de la proteína purificada.
Otras estrategias encaminadas a encontrar los genes relacionados con la síntesis de
la proteína están basadas en la realización de PCRs directamente sobre el ADN genómico
(Geisen, 2000) o el ARN mensajero (RT-PCR) del microorganismo productor de la
proteína. Para ello se pueden utilizar cebadores diseñados a partir de las secuencias amino
y carboxilo terminal de la proteína purificada.
I.5.2. AMPLIFICACIÓN RÁPIDA DE EXTREMOS DE ADNc (RACE-PCR)
Existen técnicas que permiten la amplificación de un gen partiendo de un fragmento
conocido de la proteína sin que ésta corresponda necesariamente a los extremos amino o
carboxilo terminal. Una de estas técnicas se basa en la amplificación de extremos de un
gen y es denominada amplificación rápida de extremos de ADNc mediante PCR o RACEPCR (“ Rapid Amplification of cDNA Ends - Polymerase Chain Reaction” ).
Para llevar a cabo esta técnica se requiere conocer una parte del gen que queremos
caracterizar. Esta región se puede obtener mediante el diseño de cebadores a partir de
secuencias de aminoácidos obtenidas previamente y la posterior amplificación por PCR
utilizando ADN genómico del microorganismo.
Existen dos tipos de RACE-PCR en función del extremo que se vaya a amplificar:
5’-RACE y 3’-RACE (Scotto-Lavino y col., 2006a y b). En la Figura I6 se representa un
esquema general de la amplificación de un gen completo mediante RACE-PCR. En primer
lugar es necesaria la extracción de ARN a partir de un cultivo del microorganismo en el
cual se ha inducido la transcripción del gen que codifica la proteína. Seguidamente, se
realiza una transcripción reversa empleando unos adaptadores o secuencias conocidas que
hibridan con los extremos 5’ y 3’ del ARN inicial (Figura I7). Por lo tanto, el ADNc
resultante llevará consigo estas secuencias conocidas, generando dos poblaciones de ADNc
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5’ y 3’ para las amplificaciones de los extremos 5’ y 3’, respectivamente. Para la
amplificación final de los extremos del gen a caracterizar se llevan a cabo dos reacciones
de PCR, una para el extremo 5’ y otra para el extremo 3’, que emplearán respectivamente
las dos poblaciones de ADNc 5’ y 3’ obtenidas anteriormente por retrotranscripción. Se
utilizan además dos cebadores por cada reacción: uno complementario a la secuencia
conocida del adaptador que se unió a los extremos 5’ o 3’ (LUP o SUP), y otro diseñado a
partir de la parte conocida del gen que codifica la proteína (5’-RACE GSP o 3’-RACE
GSP). Para su confirmación, los fragmentos son clonados y secuenciados. Una vez
conocidos los extremos 5’ y 3’, la obtención del gen completo es una simple reacción de
PCR empleando dos cebadores diseñados a partir de los extremos 5’ y 3’ conocidos del
gen.
La determinación de la secuencia de ADN que codifica la proteína antifúngica
puede ser de gran interés para la transformación de otros microorganismos, con el fin de
expresar la proteína con una alta eficacia para su producción a nivel industrial. Esto
facilitaría su utilización como aditivo en alimentos en los que la presencia de mohos no sea
deseable. Por otra parte, también podría servir para expresar la proteína en otros
microorganismos aislados de productos cárnicos madurados que presenten unas
características deseables, pero que sean incapaces de competir con el resto de la población
fúngica.
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INTRODUCCIÓN
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21
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INTRODUCCIÓN
I.6. EVALUACIÓN DE TOXICIDAD EN PROTEÍNAS ANTIFÚNGICAS
Un importante prerrequisito para cualquier aplicación de proteínas antifúngicas es
confirmar su inocuidad mediante ensayos de toxicidad. En este tipo de ensayos se usan
células, tejidos vivos, líneas celulares u organismos, para evaluar los efectos de cualquier
sustancia. Básicamente consisten en su exposición a determinadas concentraciones de la
proteína a evaluar por un tiempo determinado.
El material biológico empleado para los ensayos, ya sean células u organismos,
debe tener alta sensibilidad a los tóxicos, ya que al establecer las concentraciones seguras
para este tipo de material, se espera proteger a los organismos superiores sobre los que se
pretenden emplear las proteínas antifúngicas.
Uno de los métodos comúnmente empleados para demostrar la seguridad y
selectividad de una proteína es el ensayo de actividad hemolítica (Hickey y col., 2003; Jin
y col., 2009). En este método, la evaluación de la posible toxicidad de la proteína objeto de
estudio se realiza empleando glóbulos rojos procedentes de mamíferos y comprobando la
hemólisis producida. Si se produce hemólisis, la proteína tiene efectos citotóxicos y no es
apta para su utilización; si por el contrario la proteína no ocasiona la lisis de los glóbulos
rojos, merecerá seguir siendo estudiada. La ausencia de actividad hemolítica generalmente
se acepta como prueba de que la proteína es segura (Van’t Hof y col., 2001). Algunas
ventajas de la evaluación de la actividad hemolítica con respecto a otros métodos son que
se evita el sacrificio de animales de experimentación y es un método que no requiere
equipos o instrumentos específicos y puede ser realizado en cualquier laboratorio de
investigación. Sin embargo, los ensayos in vitro a menudo no reproducen las condiciones
fisiológicas en los consumidores, donde la proteína puede sufrir cambios que afecten a su
toxicidad (Theis y Stahl, 2004). Por lo tanto, conviene complementar esta prueba con otro
tipo de evaluaciones.
Un método adicional para la comprobación de efectos citotóxicos en proteínas son
los ensayos con líneas celulares, procedentes de células humanas que han perdido la
capacidad de controlar su división celular. La ventaja de las líneas celulares es que tienen
características similares a las células del mismo tipo en el organismo vivo, y a su vez
evitan el sacrificio de animales de experimentación para la obtención de cultivos primarios.
No obstante, ya que las líneas celulares han sufrido ciertas modificaciones en factores
como los del control de la división celular, los cultivos primarios normalmente procedentes
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INTRODUCCIÓN
de ratas o ratones de laboratorio siguen siendo ampliamente utilizados para la evaluación
de la toxicidad (Szappanos y col., 2005, 2006).
Entre los ensayos que emplean organismos vivos para la evaluación de la toxicidad,
uno de los más útiles y sencillos de desarrollar es el empleo de larvas de Artemia salina.
En este ensayo, las larvas son expuestas a la sustancia a evaluar y se calcula el porcentaje
de mortalidad de las mismas tras un tiempo determinado. La principal ventaja de este
método es que las larvas son especialmente susceptibles a sustancias tóxicas, junto con la
fácil disponibilidad de este organismo. Además, las larvas de Artemia salina no requieren
condiciones especiales de cultivo, simplemente necesitan ser incubadas en un medio que
simula al agua de mar, donde normalmente se desarrolla este organismo. Por todo ello, este
método ha sido ampliamente empleado en múltiples estudios, incluyendo la evaluación de
la presencia de micotoxinas en alimentos (Núñez y col., 1996, 2007; López-Díaz y col.,
2001; González y col., 2007), producción de micotoxinas de hongos patógenos de plantas
(Moretti y col., 2007), en hierbas medicinales (González y col., 2007) o la evaluación de
sustancias empleadas en odontología (Milhem y col., 2008) e incluso de nueva síntesis
(Chohan, 2009).
I.7. EFECTO DE DIFERENTES COMPUESTOS SOBRE LA ACTIVIDAD DE
PROTEÍNAS ANTIFÚNGICAS.
El efecto de diversos tratamientos o sustancias sobre la actividad antifúngica tiene
gran interés para la caracterización de estas proteínas. En primer lugar permitirá formular
hipótesis con respecto a los posibles mecanismos de acción implicados en la actividad que
muestran frente a distintos mohos sensibles. Por otra parte, la influencia de determinados
compuestos o procesos que puedan estar presentes o incorporarse en los alimentos
condicionará en gran medida sus posibles aplicaciones. Por lo tanto, para su
caracterización, se puede estudiar el efecto que ejercen en las proteínas diversos factores
como tratamientos térmicos (Munimbazi y Bulleman, 1998; von der Weid y col., 2003;
Skouri-Gargouri y Gargouri, 2008), diferentes valores de pH (Harighi y col., 2007), o
distintas concentraciones de cationes (Almeida y col., 2000; Kaiserer y col., 2003),
quelantes (Edlind y col., 2002), detergentes (Naeem y col., 2006) o fosfato (Liu y col.,
2002).
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I.7.1. TEMPERATURA
En diversos alimentos es frecuente la aplicación de tratamientos térmicos, por lo
que la estabilidad de la proteína antifúngica al calor será imprescindible para su posible
aplicación en este tipo de productos.
La termoestabilidad de una proteína está directamente relacionada con su
estructura. De hecho las proteínas resistentes a altas temperaturas poseen gran cantidad de
aminoácidos cargados y residuos aromáticos en su estructura primaria, son frecuentes las
interacciones iónicas o modificaciones post-traduccionales en las mismas, y además son
también muy estables al tratamiento con proteasas (Trivedi y col., 2006; Zhou y col.,
2007). Además, las proteínas plegadas o con gran cantidad de puentes disulfuro son más
resistentes a altas temperaturas.
Las proteínas antifúngicas para uso en alimentos no requieren una alta
termoestabilidad, ya que no irán destinadas a productos sometidos a un tratamiento de
esterilización en los que generalmente no van a sobrevivir microorganismos. Por el
contrario, sí puede ser de interés que resistan tratamientos equivalentes a la pasterización
que se pueden aplicar en productos lácteos o cárnicos.
I.7.2. pH
La evaluación de la estabilidad de las proteínas antifúngicas ante distintos valores
de pH permitiria evaluar su aplicación en alimentos ácidos o fermentados como embutidos
o quesos madurados y encurtidos, siendo el pH de estos últimos incluso menor de 3,7. Del
mismo modo que la temperatura, la resistencia de una proteína a diferentes pHs puede
indicar el grado de estabilidad de su estructura.
I.7.3. CATIONES
El estudio de la influencia de distintos cationes sobre las proteínas es de gran valor
para su caracterización, ya que muchas proteínas antimicrobianas, especialmente las que
actúan sobre membranas, son sensibles a distintas concentraciones de los mismos (Theis y
Stahl, 2004). No obstante existen proteínas antifúngicas, como algunas defensinas de
plantas, que mantienen su actividad incluso con altas concentraciones de iones en el medio
(Thevissen y col., 1999). La presencia de cationes puede además modificar la estructura o
conformación de la proteína (Parker y col., 1996; Sissi y Palumbo, 2009) e incluso
interferir en la unión con las membranas sobre las que actúan. En este tipo de proteínas los
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cationes pueden interferir en la unión tanto con los fosfolípidos (Osborn y col., 1995;
Thevissen y col., 1997, 1999), como con los receptores específicos sobre los que actúan en
las membranas de los microorganismos sensibles (Almeida y col., 2000; Kaiserer y col.,
2003; Theis y col., 2003). Además en algunos casos, el efecto sobre la actividad de la
proteína es diferente si están presentes cationes monovalentes o divalentes (Broekaert y
col., 1995). Por ello, la exposición de proteínas a diferentes cationes como Ca2+, Mg2+, Na+
o K+, puede ofrecer una valiosa información sobre el tipo de mecanismo de acción
implicado en la actividad antimicrobiana.
Por otro lado, la resistencia a cationes es esencial para la aplicación de proteínas
antimicrobianas en alimentos con alta concentración de sales, como es el caso de algunos
productos cárnicos madurados, ya que estas sales podrían provocar una disminución de la
actividad de la proteína e incluso llegar a inactivarla (Theis y Stahl, 2004).
De manera similar, una baja concentración de iones en el medio también puede
modificar la actividad de una proteína, tanto potenciándola como inhibiéndola (Edlind y
col., 2002). Por ello es necesario evaluar también el efecto sobre la proteína de
secuestradores de iones como EDTA y EGTA. Algunos como EDTA y fosfato suelen ser
empleados como quelantes de metales en alimentos no ácidos que requieren aditivos sin
sabor (Calvo, 1991), por lo que es necesario estudiar la influencia de estos compuestos en
la actividad antimicrobiana si se pretende aplicar la proteína en este tipo de productos.
I.7.4. DETERGENTES
Los detergentes, dependiendo de su carácter iónico, pueden influir en las proteínas
de distinta manera, modificando su estructura e incluso desnaturalizándolas, o potenciando
su actividad al favorecer la solubilidad de sus sustratos.
Los detergentes no aniónicos, como tween 20, tween 80 y Triton X-100, se utilizan
como suplementos en medios de cultivo para favorecer la dispersión de los
microorganismos inoculados. Este tipo de detergentes solubilizan los sustratos sin alterar la
estructura de las proteínas, por lo que no suelen ser muy agresivos contra microorganismos
o proteínas.
Por el contrario los detergentes iónicos, como docecil sulfato sódico (SDS) o ácido
deoxicólico, tienen gran actividad antimicrobiana, ya que ocasionan la disrupción de las
membranas y tienen efectos líticos, por lo que son conocidos como potentes
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desnaturalizantes de proteínas. Usualmente, estos detergentes son utilizados para el estudio
de proteínas ya que pueden alterar la conformación de las mismas. Así, la adición de
ciertos detergentes puede llevar a la inducción de una estructura en α–hélice en ciertas
proteínas (Naeem y col., 2006).
El uso de detergentes no iónicos como emulsionantes también está extendido en la
industria alimentaria. De hecho en España se emplean tween 20 y tween 80 como
emulsionantes en confitería, repostería y para la elaboración de galletas (Calvo, 1991). La
resistencia de la actividad antimicrobiana de las proteínas a estos detergentes haría que
fuesen unas buenas candidatas para su uso en este tipo de alimentos.
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RESULTADOS
Characterization of PgAFP, a novel antifungal protein from Penicillium
chrysogenum
Andrea Rodríguez-Martín, Susan Liddella, Raquel Acosta, Félix Núñez, Miguel A.
Asensio*
Higiene y Seguridad Alimentaria, Facultad de Veterinaria. Universidad de Extremadura, Avda. de la
Universidad, s/n. 10071- Cáceres, Spain.
a
Division of Animal Sciences, University of Nottingham. Sutton Bonington Campus. Loughborough,
Leicestershire. LE12 5RD. United Kingdom.
* Corresponding author: Telephone (+34) 927 257 125; fax: (+34) 927 257 110.
E-mail: masensio@unex.es
ABSTRACT
A new antifungal protein PgAFP from Penicillium chrysogenum is described. The protein
showed a molecular weight of 6.5 kDa by electrospray ionization mass spectrometry (ESIMS). Peptide sequences were obtained by digesting the protein with trypsin and
chymotrypsin and analyzing the resulting peptides by ESI-MS/MS. The obtained
sequences showed high similarity with the antifungal protein Anafp from Aspergillus
niger. No evidence for N- or O- deglycosylations was observed upon chemical and
enzymatic treatments of PgAFP. PgAFP is a new protein that belongs to a group of small,
cysteine-rich, and basic proteins with antifungal activity. However, the characterization of
PgAFP should be exhaustive to demonstrate that this protein is harmless to humans before
any potential use as preservative in foods.
KEY WORDS: antifungal protein, dry-fermented food, growth inhibition, posttranslational modifications.
INTRODUCTION
The antifungal protein PgAFP was purified in previous studies by fast protein liquid
chromatography (Acosta et al., 2009a) from the strain RP42C, which was isolated from
dry-cured ham and characterized as P. chrysogenum based on classical taxonomical
methods (Pitt and Hocking, 1997). PgAFP showed a strong antifungal activity against
selected toxigenic moulds (Acosta et al., 2009a).
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MANUSCRITO 1
Many fungal proteins have been reported including ribosome inactivating proteins,
antifungal proteins, ribonucleases, ubiquitin-like proteins, lectins, cellulases, xylanases,
laccases, invertases, and trehalose phosphorylases (Ng, 2004). There is only scant
information available about antifungal proteins of fungal origin. PgAFP showed some
similarity (Acosta et al., 2009b) to a group of antifungal peptides secreted filamentous
fungi of the group Ascomycetes, composed of proteins with common characteristics
including being cysteine rich, highly basic, and of low molecular weight. Only four
proteins have been reported in this group: PAF from Penicillium chrysogenum (Marx et
al., 1995), AFP from Aspergillus giganteus (Nakaya et al., 1990; Lacadena et al., 1995),
Anafp from Aspergillus niger (Lee et al., 1999) and AcAFP from Aspergillus clavatus
(Skouri-Gargouri and Gargouri, 2008). The amino acid sequence for all these proteins is
known, except for AcAFP.
In previous studies (Acosta et al., 2009b) PgAFP was positive for glycosylation by
Periodic Acid Schiff reaction (Deepak et al., 2003). No glycosylations have been described
for PAF, AFP, Anafp, and AcAFP, but this kind of post-translational modification has
been described in the antifungal protein from Urginea indica (Deepak et al., 2003) or
chitosanases from Mycobacter AL-1 and P. islandicum (Hedges and Wolfe, 1974; Fenton
and Eveleigh, 1981).
The aim of this work was to determine the amino acid sequence and molecular weight of
PgAFP. In addition, PgAFP was exposed to enzymatic and chemical deglycosylation
treatments, and analyzed for possible changes in the protein by SDS-PAGE and
electrospray ionization mass spectrometry. The genetic identity of the producer mould was
also confirmed.
MATERIALS AND METHODS
Microbial strain
The strain RP42C was isolated from dry-cured ham, and it was identified as Penicillium
chrysogenum according to classical morphological and biochemical studies (Acosta et al.,
2009a). This mould produces the antifungal protein PgAFP (Acosta et al., 2009a).
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RESULTADOS
Genetic identification of the producer mould
Genetic characterization of RP42C was carried out by amplifying the fragments 18S-28S
rRNA intergenic spacer (ITS) by polymerase chain reaction (White et al., 1990) and
determining the DNA sequence.
DNA isolation. The mould RP42C was grown in malt extract broth made of glucose 2%
(w/v), malt extract 2% (w/v), and peptone 0.1% (w/v), pH 4.5, at 25ºC for 5 days. After
freezing by adding liquid nitrogen, two grams of mycelium were ground up using a mortar
and pestle. Lysis was performed by homogenizing the obtained powder in 4 ml 0.05 M
Tris-HCl, pH 8, 0.005 M ethylenediamine tetraacetic acid (EDTA), 0.05 M NaCl (TES)
buffer and 1% (w/v) sodium dodecyl sulfate (SDS). Afterwards, 200 µl proteinase K (20
µg/µl) were added, and the mixture was incubated at 60ºC for 35 min, then cooled on ice
and extracted with phenol-chloroform-isoamyl alcohol (25:24:1). The suspension was
centrifuged at 3,000 ×g for 2 min at 4ºC. The upper phase was transferred to a fresh tube
and DNA was precipitated by the addition of 3 M sodium acetate, pH 5.2 to a final
concentration of 10% (w/v) and two volumes of cold ethanol. After centrifugation, the
pellet was cleaned with ethanol at 70%, centrifuged again, resuspended in sterile water and
treated with 50 µl RNase (10 µg/µl) at 37ºC for 60 min. Finally, DNA was extracted with
phenol-chloroform-isoamyl alcohol and precipitated again as indicated above, resuspended
in water and stored at -70ºC. The quantity and quality of the purified DNA was determined
using a Biophotometer (Eppendorf AG, Hamburg, Germany).
ITS amplification. DNA fragments for sequence analysis were obtained by PCR using
primers ITS1 (TCCGTAGGTGAACCTGCGG) and ITS4 (TCCTCCGCTTATTGATAT
GC). Amplified products comprised the internal transcribed spacer 1 (ITS1), 5.8S, and
ITS2 regions of the ribosomal DNA. For each 50 µl reaction, a mixture was prepared
containing 50 ng of genomic DNA, 0.1 vol. of 10× PCR buffer (10 mM Tris-HCl, pH 8.8,
50 mM KCl, 0.1% Triton X-100), 4 mM MgCl2, 1 mM of deoxynucleoside triphosphates
(Roche, Mannheim, Germany), 0.4 µM of each primer, and 2 U of DyNAzyme I DNA
Polymerase (Finnzymes, Espoo, Finland). Reactions were run on a programmable thermal
cycler Mastercycler epgradient (Eppendorf AG) with an initial denaturation of 3 min at
96°C, followed by 30 cycles at 94°C for 30 s, 60°C for 30 s, and 72°C for 40 s, with a final
extension at 72°C for 10 min. Aliquots of the PCR were electrophoresed on 1% (w/v)
agarose gels in 1× TAE buffer (40 mM Tris-acetate, 1 mM EDTA, pH 8) using a Sub-Cell
GT apparatus (Bio-rad Laboratories, Hercules, USA). Products were visualized using the
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MANUSCRITO 1
UV transillumination G-Box (Syngene, Fredericks, USA) and photographed with a
GeneSnap camera (Syngene). DNA fragments were gel-purified and sequenced at the
Institute of Biomedicine (CSIC, Valencia, Spain). The sequences obtained were searched
against the public databases using the Basic Local Alignment Search Tool (BLAST).
PgAFP purification
P. chrysogenum RP42C was grown in malt extract broth at 25ºC for 15 days. The medium
was filtered through a 0.45 µm diameter pore size cellulose acetate membrane and PgAFP
was purified using an ÄKTA fast protein liquid chromatography system (Amersham
Pharmacia Biotech AB, Uppsala, Sweden) according to Acosta et al. (2009a). For this,
filtered medium was loaded on a HiTrap SP HP 5 ml ion exchange column (Amersham
Pharmacia Biotech AB) equilibrated with 0.02 M sodium acetate buffer, pH 4.5. Bound
proteins were eluted with a gradient of NaCl to 1 M (0 to 25% in 15 column volumes
(CV), increasing to 100% in 2 CV, followed by 15 CV at 100%) in 0.02 M sodium acetate
buffer. Eluted proteins were detected at 214 nm. The fraction that showed antifungal
activity in previous studies (Acosta et al., 2009a) was further purified using a HiLoad
26/60 Superdex 75 prep grade gel filtration column (Amersham Pharmacia Biotech AB)
equilibrated with 50 mM sodium phosphate, 0.15 M NaCl buffer, pH 7, at a constant flow
rate of 2.5 ml/min. Data collection and processing were performed using the UNICORN
software 4.12 (Amersham Pharmacia Biotech AB). Finally, the fraction containing PgAFP
was desalted and concentrated with YM-3 Microcon Centrifugal Filter Units (Millipore
Iberica S.A.U., Madrid, Spain), before storage at -20ºC.
Protein characterization
Sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE). SDS-PAGE
was performed as described Laemmli (1970). The protein PgAFP was separated on 4%
(v/v) polyacrylamide stacking and 12% (v/v) polyacrylamide resolving gels. Gels were
stained with Imperial Protein Stain (Pierce Biotechnology, Rockford, USA). Two
molecular weight marker sets were used: Precision Plus Protein Standards with ten proteins
between 10 and 250 kDa (Bio-Rad) and Low Molecular Weight Electrophoresis calibration
kit with proteins between 14.4 and 97 kDa (Pharmacia, Uppsala, Sweden).
Molecular weight determination by ESI-MS. The molecular weight of PgAFP was
determined by electrospray mass spectrometry in a Q-TOF2 mass spectrometer (Waters
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RESULTADOS
Ltd, Manchester, UK). The protein was concentrated and dissolved in ammonium acetate.
PgAFP (0.1 µg/µl) with 1% (v/v) formic acid was desalted using C18 reverse phase pipette
Zip-Tips (Millipore) according to the manufacturer’s instructions. Samples were eluted
into 50% (v/v) methanol, 0.1% (v/v) formic acid, and were loaded into nanospray needles
(Waters) for analysis. The mass spectrometer was operated with a capillary voltage of 9001200 V and the sampling cone at 45-50 V. Data were acquired between 400 and 3000 m/z
with a cycle time of 1 second. Mass spectra resulted from combining typically 5 min of
data. Spectra were deconvoluted using the MaxEnt1 maximum entropy software (Waters)
with an output range of 4,000 and 11,000 Da, at a resolution of 1 Da and a peak width of
0.75 Da.
Reduction and alkylation of PgAFP. Reduction and alkylation of the protein prior to
deglycosylation or enzymatic digestion were performed with the protein both in
acrylamide gel pieces and in solution. After SDS-PAGE, PgAFP (25 µg per well) was
stained using Imperial Protein Stain. The band was excised from the gel and washed three
times by incubation in 100 µl of de-stain solution made of 50 mM ammonium bicarbonate
and 50% acetonitrile for 15 min at 40ºC with intermittent vortexing. Gel pieces were
incubated at 56ºC in 100 µl of reducing solution (10 mM dithiothreitol in 50 mM
ammonium bicarbonate) that was removed after 45 min. One hundred µl of alkylation
solution (55 mM iodoacetamide in 50 mM ammonium bicarbonate) was added and
incubation continued at room temperature in the dark for 45 min. The gel slice was then
washed in 100 µl of 50 mM ammonium bicarbonate for 5 min at 40ºC and the supernatant
was discarded. Gel pieces were washed twice in 100 µl of 50 mM ammonium bicarbonate
in 50% (v/v) acetonitrile for 1 min, and centrifuged. Sample was dehydrated in 100 µl of
acetonitrile and incubated for 1-2 min. Finally, the supernatant was discarded and the gel
pieces were dried at 40ºC for a few min. After addition of the digestive enzyme, the tube
containing the gel pieces was cooled at 4°C for 20 min and finally incubated at the optimal
temperature of each enzyme.
To digest PgAFP in solution, different quantities of the protein (12.5, 25 and 62.5 µg) were
added to tubes with 50 mM ammonium bicarbonate up to a volume of 5 µl. Dithiothreitol 5
mM (dissolved in 50 mM ammonium bicarbonate) was added (5 µl) and the final solution
was incubated at 50ºC for 30 min. After adding 10 µl of 20 mM iodoacetamide (dissolved
in 50 mM ammonium bicarbonate), sample was incubated at room temperature for 30 min
in the dark before the digestion with each deglycosidase or protease.
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Analysis for glucidic residues. Chemical deglycosylation was carried out with PgAFP in
solution using two different chemical methods: acid hydrolysis with HCl (Krapf et al.,
2006; Leis et al., 1997) and -elimination with NaOH (Geert and Thomas-Oates, 1998;
Tarelli, 2007). For acid hydrolysis with HCl, 1 µg of PgAFP was incubated in 20 mM HCl
at 50ºC for 3 h. -elimination with NaOH was performed incubating 1µg of PgAFP with
0.2 M NaOH at 45ºC for 16 h. Finally, both samples were analyzed by ESI-MS.
Enzymatic deglycosylation was carried out using endoglycosidase F1 (Northstar
BioProducts, Liverpool, UK), PNGase F (New England Biolabs, Ipswich, UK), and the EDEGLY commercial kit (Sigma, Madrid, Spain). PgAFP protein (200 µg) was treated with
4 µl of endoglycosidase F1 in 250 mM NaH2PO4 at pH 4.5, the sample was incubated at
37ºC for 1 h, and analyzed by ESI-MS. PNGase F was used following the manufacturer’s
instructions. Two additional assays were performed without detergent with PgAFP in-gel
and in-solution, one with a prior reduction and alkylation as above and another one heating
PgAFP at 100ºC for 5 minutes before adding the enzyme. All samples were analyzed by
SDS-PAGE and ESI-MS. The E-DEGLY deglycosylation kit (Sigma), containing PNGase
F,
-2(3,6,8,9)
neuraminidase,
O-glycosidase,
(1-4)-galactosidase,
and
−N-
acetylglucosaminidase, was also used. Reactions were carried out under native and
denaturing conditions according to the manufacturer’s instructions and analysed by SDSPAGE.
Amino acid sequencing
Digestion with enzymes
Trypsin digestion. PgAFP was digested with trypsin gold (Promega, Hampshire, UK).
Different treatments with and without reduction and alkylation steps were performed prior
to digestion. Gel piece samples were incubated with trypsin gold (10 ng/µl) in 50 mM
ammonium bicarbonate at 40ºC or 60ºC for 1.5, 3, and 16 h. PgAFP in-solution was
incubated at 37ºC for 16 h in 50 mM ammonium bicarbonate, 5% (v/v) acetonitrile and 10
ng/µl of trypsin gold.
Chymotrypsin digestion. Digestions with the reduced and alkylated protein were performed
with PgAFP in-gel and in-solution. Gel pieces containing PgAFP were digested at different
concentrations (1, 5, 10, and 15 ng/µl) of chymotrypsin (Roche, Welwyn Garden City,
UK) in buffer (100 mM Tris-HCl, 10 mM CaCl2, pH 7.8) at 25ºC and 30ºC for 16 h.
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PgAFP in-solution was digested by chymotrypsin at 0.1, 1, 5 or 10 ng/µl with 5%
acetonitrile and the same buffers detailed above. Temperatures and incubation time were
identical to the in-gel samples.
Electrospray ionization mass spectrometry (ESI-MSMS) analysis for peptide sequence
determination
Peptide samples were made to 0.1% (v/v) formic acid, then desalted using Zip-Tip C18
reverse phase pipette tips (Millipore) according to manufacturer’s instructions. Peptides
were eluted with 50% (v/v) acetonitrile, 0.1% (v/v) formic acid. Samples were loaded into
individual borosilicate nanospray needles (Waters) using 1-10 µl GELoader pipette tips
(Eppendorf) and analyzed in a Q-TOF2 mass spectrometer (Waters), a high resolution
hybrid orthogonal angle-Time of Flight tandem LC mass spectrometer, fitted with a
nanoflow ESI (electrospray ionization) source (Waters). The mass spectrometer was
operated with a capillary voltage of 900–1,200 V in positive ion mode, using argon as the
collision gas. Survey scans of peptide mass spectra were performed with the sampling cone
set at 45–50 V and data were acquired from 400–1,600 m/z with a cycle time of 2.4
seconds. Peptide mass spectra presented resulted from combining typically 5 min of data
acquisition. Individual peptide ions were selected manually for tandem MS analysis.
Collision voltages were optimised manually for individual peptides. Tandem MS
fragmentation spectra were collected typically from 50 to 1,600 m/z. Tandem MS spectra
were deconvoluted into singly charged, mono-isotopic masses using the MaxENT3
maximum entropy software, and peptide sequences were determined by manual
interpretation using the PepSeq software within the MassLynx V 4.0 package (Waters). De
novo peptide sequences were searched against the public database BLASTP using
parameter settings for “short and nearly exact matches”.
RESULTS
Molecular genetic characterisation of P. chrysogenum RP42C
Amplification of the 18S-28S rRNA intergenic spacer (ITS) sequence produced a band
with 394 bp. The DNA sequence of the amplified was determined and compared against
other sequences using BLAST. The sequence from RP42C mould shared 100% of identity
and showed a maximum score with the 18S-28S rRNA intergenic spacer of Penicillium
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chrysogenum strain L1 (Accession number in GenBank: FJ467371). Thus, the producer
mould RP42C was confirmed as Penicillium chrysogenum.
Molecular weight determination
The molecular mass of purified PgAFP was estimated from its migration on SDS-PAGE.
The protein migrated as a single band of approximately 10 kDa (Figure 1).
Figure 1. SDS-PAGE of PgAFP (5 µg). M: Pharmacia Low Molecular Weight Markers.
In ESI-MS, PgAFP resolved as a short series of multiply charged peaks, with the dominant
peak set of around 1,083 Da corresponding to the +6 charge state (Figure 2). The m/z
spectrum was deconvoluted and converted to the molecular mass profile using Maximum
Entropy processing (Figure 3). The mass profile shows a dominant peak of 6,494 Da,
corresponding to the mass of mature PgAFP.
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Figure 2. Electrospray mass spectra of PgAFP. A) Spectrum showing the major multiply
charged peaks. B) Expanded spectrum showing the +6 charge state peak set.
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6494.29
6592.22
mass
Figure 3. Molecular mass profile from Maximum Entropy processing. PgAFP resolves as a
single major peak with a mass of 6,494 Da.
Analysis for glucidic residues
Analysis of PgAFP for glycosylations did not show any evidence of N- or O-linkedoligosaccharides. Chemical deglycosylation with acid hydrolysis or -elimination revealed
no changes in the mass spectrum profiles obtained in ESI-MS compared with untreated
controls. No differences on gel-mobility of PgAFP were observed on SDS-PAGE after
attempting deglycosylation by enzymatic means using endoglycosidase F1 (data not
shown), PNGase F (Figure 4), or the E-DEGLY deglycosylation kit (Figure 4). ESI-MS
analysis showed no change in the mass spectrum profiles of PgAFP treated with
endoglycosidase F1 and PNGase F (data not shown).
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Figure 4. SDS-PAGE analysis of attempted enzymatic deglycosylation of PgAFP.
A) PNGase F-treated PgAFP (1) untreated PgAFP (2). B) E-DEGLY deglycosylation kit
under denaturing conditions (1, 2) or native conditions (3, 4). 1 and 3 are untreated controls.
M: Protein molecular weight standards.
Peptide sequences
The peptides obtained from PgAFP after digestion by trypsin and chymotrypsin were
analyzed by electrospray ionization tandem mass spectrometry and the amino acid
sequences deduced. Mass spectrum profiles of peptides obtained from PgAFP digestions
are shown in Figure 5. There were no major differences in the peptide profiles for in-gel
compared to in-solution digestions. Six de novo peptide sequences were obtained by
treatment with trypsin and chymotrypsin (Figure 5, Table 1). The survey spectrum and the
processed tandem MS spectrum of a representative peptide are shown in Figure 6.
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Figure 5. Mass spectra of PgAFP peptides after trypsin (A) and chymotrypsin (B) digestions.
Table 1. De novo obtained peptide sequences. Lower case letters indicate amino acids which were
tentative assignments. Dashes indicate the presence of additional amino acids which were
unassigned. Note that L and I are interchangeable here since these two residues are isobaric.
Sequence
Number
de novo sequence
Peptide
Charge
Digestive
Mass
state
enzyme
1
LSKFGGECSL
549.26
+2
Chymotrypsin
2
FGGECSLK
449.24
+2
Trypsin
3
HNTCTYLK
490.26
+2
Trypsin
4a
NHVVNCGSAANKK
466.88
+3
Trypsin
607.308
+2
Trypsin
4b
NHVVNCGSAANK
5
-yDEHHK
468.19
+2
Trypsin
6
-cqTPV
540.92
+3
Chymotrypsin
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Figure 6. ESI MS (A) and MS/MS spectrum (B) of a PgAFP tryptic peptide with a charge state
of +3 and precursor mass of 607.308 (peptide 4b NHVVNCGSAANK in Table 1).
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DISCUSSION
To confirm the previous identification of PgAFP based on morphology and biochemical
characteristics (Acosta et al., 2009a), the 18S-28S rRNA intergenic spacer was analyzed.
The obtained sequence confirmed that the producer mould RP42C belongs to P.
chrysogenum.
The molecular mass of PgAFP obtained by SDS-PAGE was estimated to be over 10 kDa
(Figure 1), whereas ESI-MS revealed a lower mass of 6.5 kDa. For large molecules, such
as intact proteins, ESI-MS is capable of determining the molecular weight to within 0.01%
of the total molecular weight. For a mass of 6,494 Da, this is 0.65 Da. Therefore, it seems
likely that PgAFP migrates anomalously in SDS-PAGE. Many characteristics of proteins
can cause them to migrate atypically on SDS-PAGE, in particular, lipoproteins and
glycoproteins migrate anomalously because SDS cannot bind with lipids or
oligosaccharides (Grefrath and Reynolds, 1974). In previous studies (Acosta et al., 2009b)
PgAFP showed evidences of glycosylations by Periodic Acid Schiff reaction (Deepak et
al., 2003). Of all protein postranslational modifications currently known, such as
acetylation, phosphorylation, formylation, or glycosylation, the last is by far the most
common. Moreover, glycosylated proteins are not uncommon among antifungal proteins,
such as an antifungal glycoprotein from Urginea indica (Deepak et al., 2003) or
chitosanases from Mycobacter AL-1 and P. islandicum (Hedges and Wolfe, 1974; Fenton
and Eveleigh, 1981). We therefore used a range of enzymatic and chemical
deglycosylation methods to assess possible N-linked and O-linked glycosylation of
PgAFP. We found no evidence of N- or O-linked oligosaccharides. Although in some
cases the assay products were assessed using only SDS-PAGE, in which the resolution
may have been insufficient to reveal relatively small mass changes, no evidence of mass
changes were found using highly resolving MS analysis. Atypical migration on SDSPAGE can be due to other causes that do not involve post-translational modifications. For
example, proteins with extreme isoelectric points can show a higher apparent molecular
weight (around a 25% higher) due to electrostatic repulsions causing less SDS to bind and
a consequent decrease in their migration velocity (Mondstadt and Holldorf, 1990).
Notably, the 42 residues obtained using de novo sequencing that together constitute around
72% of PgAFP (a calculated molecular weight of 4.7 kDa, compared with the 6.5 kDa
mass observed by ESI-MS), have a predicted pI of pH 8.8. In addition, in previous studies
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this protein showed an experimental isoelectric point of pH 9.2 by capillary isoelectric
focusing electrophoresis (Acosta et al. 2009b).
Amino acid sequences of PgAFP peptides were obtained using tandem MS and de novo
sequencing (Table 1). The sequences were searched against the public sequence databases
to find any similar proteins present. A multiple alignment was performed with the most
similar proteins (Figure 7).
PgAFP
Anafp
A.niger
PAF
AFP
A.clavatus
LSKFGGECSLKHNTCTYL
------------------------------------------LSKYGGECSLEHNTCTYR
---MQLTSIAIILFAAMGAIANPIAAE---ADNLVARE--AELSKYGGECSVEHNTCTYL
---MQITTVALFLFAAMGGVATPIES---VSNDLDARAEAGVLAKYTGKCTKSKNECKYK
MKFVSLASLGFALVAALGAVATPVEADSLTAGGLDARDESAVLATYNGKCYKKDNICKYK
MKVVSLASLGFALVAALGVVASPVDADSLAAGGLDARDESAVQATYDGKCYKKDNICKYK
PgAFP
Anafp
A.niger
PAF
AFP
A.clavatus
-K
HVVNCGSAANKK
yDEHHK
cqTPV
-KDGKNHVVSCPSAANLRCKTDRHHCRYDDHHKTVDCQTPV
-KGGKDHIVSCPSAANLRCKTERHHCEYDEHHKTVDCQTPV
NDAGKDTFIKCPKFDNKKCTKDNNKCTVDTYNNAVDCD--AQSGKTAICKCYV---KKCPRDGAKCEFDSYKGKCYC---AQSGKTAICKCYV---KVCPRDGAKCEFDSYKGKCYC----
Figure 7. Alignment of PgAFP amino acid sequences with the most similar proteins: Anafp
(Lee et al., 1999), XP_001391221 (hypothetical protein from Aspergillus niger CBS 513.88),
AAA92718 (PAF from P. chrysogenum), P17737 (AFP from A. giganteus), and XP_001267787
(hypothetical protein from A. clavatus).
The PgAFP amino acid sequences had the highest homology with Anafp (81%), and a
hypothetical protein (79 %), both from Aspergillus niger. Three other proteins had lower
levels of similarity with PgAFP; PAF (33%), AFP (26%), and a hypothetical protein from
Aspergillus clavatus (21%). Both PgAFP and PAF are produced by P. chrysogenum, but
they are clearly different proteins. The aligned proteins and PgAFP have common
characteristics: they are small (5.8-6.6 kDa), cysteine-rich, and basic, due to the presence
of a high amount of lysine residues (Marx, 2004). The mature proteins PAF and AFP are
cleavage products of precursor forms which carry a pre-pro-sequence, with a signal peptide
and a prosequence that keep the proteins in an inactive form until exported to the
extracellular medium (Marx, 2004). Since PgAFP may belong to this group of proteins, it
could also be produced as a precursor form. In contrast with other antifungal proteins, such
as some chitosanases, no glycosylations have been described in this group. The apparent
lack of glycosylation of PgAFP is consistent with it being a member of this group.
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Recently, the genome of P. chrysogenum Wisconsin 54-1255 has been sequenced (Van den
Berg et al., 2008). A putative protein sequence is present (accession number CAP80456)
which shows homology with the amino acid sequences obtained from PgAFP in this work.
However, the production of the protein by this mould has not been reported.
PgAFP is a new antifungal protein obtained from P. chrysogenum. Since PgAFP was
produced by a species generally recognized as safe by the United States Food and Drug
Administration (Marx, 2004) it may be permissible for use as preservative in foods.
However, the characterization of PgAFP should be exhaustive to demonstrate that this
protein is harmless to humans. For this, future objectives will include obtaining the coding
sequence of this novel and potent antifungal protein.
ACKNOWLEDGEMENTS
This work was supported by the Spanish Ministry of Education and Science (AGL200406546-ALI) and FEDER. Andrea Rodríguez and Raquel Acosta were recipient of FPI
grants from the Spanish Ministry of Education and Science.
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of antifungal protein-producing molds from dry-cured meat products. Int. J. Food
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Acosta, R., Rodríguez-Martín, A., Bermúdez, E., Núñez, F., and Asensio, M.A. Partial
characterization and antifungal activity of a protein from Penicillium chrysogenum. Food
Microbiol. Submitted (2009b).
Deepak, A.V., Thippeswamy, G., Shivakameshwari, M.N., and Salimath, B.P.
Isolation and characterization of a 29-kDa glycoprotein with antifungal activity from bulbs
of Urginea indica. Biochem. Biophys. Res. Comm. 311: 735-742 (2003).
Fenton, D. M. and Eveleigh, D. E. Purification and mode of action of a chitosanase from
Penicillium islandicum. J. Gen. Microbiol. 126: 151-165 (1981).
Geert, J. D. and Thomas-Oates, J. Analysis of glycoproteins and glycopeptides using
fast-atom bombardment. Methods Mol. Biol. 61: 231-241 (1998).
Grefrath, S.P. and Reynolds, J.A. The molecular weight of the major glycoprotein from
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Hedges, A. and Wolfe, R. S. Extracellular enzyme from Myxobacter Al-1 that exhibits
both -1,4 glucanase and chitosanase activities. J. Bact. 120: 844-853 (1974).
Krapf, D., Vidal, M., Arranz, S. E., and Cabada, M. O. Characterization and biological
properties of L-HGP, a glycoprotein from the amphibian oviduct with acrosome-stabilizing
effects. Biol. Cell. 98: 403–413 (2006).
Lacadena, J., Martínez del Pozo, A., Gasset, M., Patiño, B., Campos-Olivas, R.,
Vázquez, C., Martínez-Ruiz, A., Mancheño, J.M., Oñaderra, M., and Gavilanes, J.G.
Characterization of the antifungal protein secreted by the mould Aspergillus giganteus.
Arch. Biochem. Biophys. 20: 273-281 (1995).
Laemmli, U.K. Cleavage of structural proteins during the assembly of the head of
bacteriophage T4. Nature 227: 680–685 (1970).
Lee, D.G., Shin, S.Y., Maeng, C-Y., Jin, Z.Z., Kim, K.L., and Hahm, K-S. Isolation
and characterization of a novel antifungal peptide from Aspergillus niger. Biochem.
Biophy. Res. Commu. 263: 646-651 (1999).
Leis, O., Madrid, J.F., Ballesta, J., and Hernández, F. N- and O-linked oligosaccharides
in the secretory granules of rat paneth cells: an ultrastructural cytochemical study. J.
Histoch. Cytoch. 45: 285-294 (1997).
Marx, F., Haas, H., Reindl, M., Stöffler, G., Lottspeich, F., and Redl, B. Cloning,
structural organization and regulation of expression of the Penicillium chrysogenum paf
gene encoding an abundantly secreted protein with antifungal activity. Gene. 167: 167-171
(1995).
Marx, F. Small, basic antifungal proteins secreted from filamentous ascomycetes: a
comparative study regarding expression, structure, function and potential application. Appl.
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Mondstadt, G.M. and Holldorf, A.W. Arginine deaminase from Halobacterium.
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Nakaya, K., Omata, K., Okahashi, I., Nakamura, Y., Kolkenbrock, H., and Ulbrich,
N. Amino acid sequence and disulfide bridges of an antifungal protein isolated from
Aspergillus giganteus. Eur. J. Biochem. 193: 31-38 (1990).
Ng, T.B. Peptides and proteins from fungi. Peptides. 25: 1055-1073 (2004).
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Pitt, J.I. and Hocking, A.D. Fungi and food spoilage. Blackie Academic and Professional.
London, United Kingdom (1997).
Skouri-Gargouri, H. and Gargouri, A. First isolation of a novel thermostable antifungal
peptide secreted by Aspergillus clavatus. Peptides. 29: 1871-1877 (2008).
Tarelli, E. Resistance to deglycosylation by ammonia of IgAl O-glycopeptides:
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A.J., Garcia-Estrada, C., Fedorova, N.D., Harris, D.M., Heijne, W.H., Joardar, V.,
Kiel, J.A., Kovalchuk, A., Martin, J.F., Nierman, W.C., Nijland, J.G., Pronk, J.T.,
Roubos, J.A., van der Klei, I.J., van Peij, N.N., Veenhuis, M., von Dohren, H.,
Wagner, C., Wortman, J., and Bovenberg, R.A. Genome sequencing and analysis of the
filamentous fungus Penicillium chrysogenum. Nat. Biotechnol. 26: 1161-1168 (2008).
White, T.J., Bruns, T., Lee, S., and Taylor, J. Amplification and direct sequencing of
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Academic Press, Inc., San Diego. Pages 315-322 (1990).
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Manuscrito 2: Genetic characterization of
the gene encoding the novel antifungal protein PgAFP
from Penicillium chrysogenum
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Genetic characterization of the gene encoding the novel antifungal
protein PgAFP from Penicillium chrysogenum
Andrea Rodríguez-Martín, Raquel Acosta, Félix Núñez, Mª José Benitoa, Miguel A.
Asensio*
Higiene y Seguridad Alimentaria, Facultad de Veterinaria. Universidad de Extremadura, Avda. de la
Universidad, s/n. 10071- Cáceres, Spain.
a
Nutrición y Bromatología, Escuela de Ingenierías Agrarias, Universidad de Extremadura, Carretera de
Cáceres s/n, 06071- Badajoz, Spain
* Corresponding author: Telephone (+34) 927 257 125; fax: (+34) 927 257 110.
E-mail: masensio@unex.es
ABSTRACT
The strain RP42C from Penicillium chrysogenum produces a small protein PgAFP that
inhibits the growth of some toxigenic moulds. The gene encoding the precursor of this
antifungal protein has been characterized. cDNA was prepared from isolated total RNA by
RT-PCR. The cDNA fragment encoding the pgafp gene was obtained by PCR using
degenerate primers based on two known amino acid sequences of the purified PgAFP.
Full-length cDNA of pgafp was obtained by RACE-PCR. Comparison of nucleotide
sequence from genomic fragment and cDNA of the gene, revealed the presence of a 279 bp
coding region which is interrupted by two introns of 63 and 62 bp in length. The precursor
of the antifungal protein consists of 92 amino acids and appears to be processed to the
mature 58 amino acids PgAFP by two-step process. The deduced amino acid sequence of
the preproprotein revealed 75 % identity to an antifungal protein predicted from the
Aspergillus niger CBS 513.88 genome.
KEY WORDS: preproprotein, protective cultures, dry fermented foods.
INTRODUCTION
The strain RP42C of the mould Penicillium chrysogenum secretes a small protein PgAFP
that has been purified for its ability to inhibit toxigenic moulds isolated from dry-cured
meat products (Acosta et al., 2009a,b). Several amino acid sequences of this protein were
obtained by electrospray ionization mass spectrometry (Rodríguez-Martín et al., 2009)
revealing a high homology with the antifungal protein Anafp (Lee et al., 1999). Anafp
protein belongs to a new group of antimicrobial proteins named small, basic and cysteine51
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rich antifungal proteins. The recent sequencing of the genome of different moulds has
shown several genes that could encode proteins with common characteristics to this group.
However, only four proteins have been reported: Anafp from Aspergillus niger (Lee et al.,
1999), PAF from Penicillium chrysogenum (Marx et al., 1995), AFP from Aspergillus
giganteus (Nakaya et al., 1990; Lacadena et al., 1995), and AcAFP from Aspergillus
clavatus (Skouri-Gargouri and Gargouri, 2008). Only AFP and PAF have been genetically
characterizated. In addition a gene homologous to paf gene has been described from
Penicillium nalgiovense (Geisen, 2000).
The genes encoding PAF and AFP have open reading frames interrupted by two introns
with conserved splice sites. They encode products of 92 and 94 amino acids, respectively,
and these proteins are synthesized as a long precursor with a peptide signal and a
presequence that are removed to obtain mature proteins.
Genetic characterization of the gene encoding PgAFP will allow to clon it in other
microorganisms, to increase PgAFP production and to obtain protective cultures for future
applications in foods, environment or medicine. In addition, the genetic sequence will give
valuable information for a better characterization of the protein.
MATERIALS AND METHODS
Microbial strain
P. chrysogenum RP42C was isolated from dry-cured ham, and belongs to the fungal
collection of Food Hygiene, University of Extremadura.
DNA isolation
P. chrysogenum RP42C was grown in malt extract broth made with 2% (w/v) glucose, 2%
(w/v) malt extract, and 0.1% (w/v) peptone, pH 4.5 at 25ºC under continuous shaking for 5
days. The mycelium was obtained by filtering the culture through Whatman paper nº2.
Two grams of mycelium were broken with mortar and pestle after freezing by adding
liquid nitrogen. Fungal lysis was performed by homogenizing the obtained powder in 4 ml
0.05 M Tris-HCl, pH 8; 0.005 M ethylenediamine tetraacetic acid (EDTA); 0.05 M NaCl
(TES) buffer and 1% (w/v) sodium dodecyl sulfate (SDS). Then, 200 µl proteinase K (20
µg/µl) were added, the mix was incubated at 60ºC for 35 min, cooled on ice, and extracted
with phenol-chloroform-isoamyl alcohol (25:24:1). The suspension was centrifuged at
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3000×g for 2 min at 4 ºC. The upper phase containing DNA was transferred to a fresh tube
and precipitated by adding 3 M sodium acetate pH 5.2 to a final concentration of 10% (v/v)
and two volumes of cold ethanol. After centrifugation, the pellet was cleaned with 70%
(v/v) ethanol, centrifuged again in the conditions above mentioned, resuspended in sterile
water, and treated with 50 µl RNase (10 µg/µl) at 37ºC for 60 min. Finally, DNA was
extrated with phenol-chloroform-isoamyl alcohol, precipitated again as indicated above,
resuspended in water, and stored at -70ºC.
Quantity and quality of purified DNA was determined spectrophotometrically in a
Biophotometer (Eppendorf AG, Hamburg, Germany).
Genomic DNA amplification
Polymerase chain reactions were performed with genomic DNA (100 ng) as template in 50
µl reaction mixtures containing 50 pmol each of forward and reverse primers, 0.5 mM each
of the deoxyribonucleotide triphosphates (dNTPs), 0.1 vol of 10× PCR buffer (22.5 mM
MgCl2, 500 mM Tris-HCl pH 9.2, 140 mM ammonium sulfate), 1.8 mM MgCl2 and 2.5
units pfu turbo polymerase (Stratagene, La Jolla, USA). The degenerated primers, pgafpDPF1 and pgafp-DPR1A (Table 1), were designed from amino acid sequences previously
obtained by mass spectrometry (Rodríguez-Martín et al., 2009). Three reactions were
done: one with both forward and reverse primers, another only with forward, and another
only with reverse primer.
Table 1. Degenerated primers used to amplify the pgafp gene.
Primer names
Amino acid sequences and designed primers*
L
pgafp-DPF1
H
N
T
C
T
5’-TN AAR CAY AAY ACN TGN AC-3’
C
pgafp-DPR1A
K
G
S
A
A
N
3’-CR CCN AGN CGN CGN TT-5’
* R for purine, Y for pirimidine, N for A, C, T, or G.
DNA amplification was carried out in a Px2 thermocycler (Thermo Scientific,
Cramlington, UK). The PCR program consisted of initial denaturation (94ºC for 5 min), 35
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cycles of denaturation (94ºC for 5 s), annealing (45-57ºC range for 5 s, gradient 12), and
extension (72ºC for 1 s) steps, followed by a final extension at 72 ºC for 4 min. PCR
products were electrophoresed on 2% (w/v) agarose gels with MetaPhor agarose (BMA,
Rockland, ME, USA). Two different markers were used for agarose gels: DNA marker
from Gibco (0.15-12kb bands) (Gibco, GrandIsland, NY, USA) and DNA size standard
from Bio-Rad (0.05-2kb) (Bio-rad Laboratories, Madrid, Spain). PCR products detected
only in the reactions with both reverse and forward primers were gel-purified and cloned
into the pCR2.1-TOPO vector (Invitrogen, Barcelona, Spain). One Shot TOP10
Chemically Competent E. coli (Invitrogen) were transformed with that vector. To confirm
the insert in the vector, digestions with restriction enzyme EcoRI or PCR with M13
forward and reverse primers (Invitrogen) were carried out. Finally, vectors were sequenced
in the Institute of Biomedicine (CSIC, Valencia, Spain).
RNA isolation
P. chrysogenum RP42C was grown in malt extract broth as indicated for DNA isolation.
Subsequently, mycelium was quickly frozen with liquid nitrogen and stored at -80ºC until
RNA extraction. Mycelium was broken with mortar and pestle after adding liquid nitrogen.
RNA extraction was performed with RNeasy Plant Mini Kit (Qiagen) following the
manufacturer’s instructions. The final extracted RNA was resuspended in 40 µl of RNasefree deionized water.
Rapid amplification of cDNA ends-Polymerase Chain Reaction (RACE-PCR)
Amplification of 5’ and 3’ ends of pgafp gene was carried out using the SMART RACE
cDNA
Amplification
Kit
(Clontech
Laboratories,
Saint
Germain
en
Laye,
France).
cDNA synthesis by reverse transcriptase-polymerase chain reaction (RT-PCR). Two
separate populations of cDNA were synthesized according to the amplification kit
instructions: 5’-RACE-Ready cDNA and 3’-RACE-Ready cDNA. After incorporating the
SMART sequence into both the 5’- and 3’-RACE-Ready cDNA populations, RACE PCR
reactions were performed using the Universal Primer A Mix (UPM), in conjunction with
distinct gene-specific primers.
Gene-specific primers design. A gene-specific primer (GSP) was designed for amplifying
the 3’- end (3’-RACE GSP: 5’-GGGTGGAAAGAACCATGTAGTCAATTGCG-3’) from
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the sequence previously obtained by PCR with degenerate primers pgafp-DPF1 and pgafpDPR1A. Amplification of the 5’- end was carried out designing a gene-specific primer (5’RACE GSP: 5’-AACTGGGGTCTGGCAGTCAACCCTC-3’) from the 3’-end also
previously obtained.
Polymerase chain reactions were performed with 5’- or 3’-RACE-Ready cDNAs (50 ng) as
template in 50 µl reaction mixtures containing 1 mM of each of the dNTPs (Roche,
Madrid, Spain), 0.5 µM of the 5’- or 3’-RACE GSP primers, 5 µl of Universal Primer A
Mix (10×), 0.1 vol of 10× PCR buffer (10 mM Tris-HCl, pH 8.8, 50 mM KCl, 0.1 %
Triton X-100), 4 mM MgCl2 and 2.5 units pfu turbo polymerase (Stratagene). The PCR
program for obtaining the 3’-end consisted of 35 cycles of denaturation (94ºC for 30s),
annealing (68ºC for 30s), and extension (72ºC for 3 min).
The PCR program for 5’-end amplification was: 5 cycles of 2 steps (94ºC for 30 s and
72ºC for 3 min), 5 cycles of 3 steps (94ºC for 30 s, 70ºC for 30 s and 72ºC for 3 min) and
27 cycles of 3 steps (94ºC for 30 s, 68ºC for 30 s and 72ºC for 3 min). Finally, 5’- and 3’RACE-PCR products were gel-purified and cloned into pCR2.1-TOPO vectors, which
were used to transform One Shot TOP10 Chemically Competent E. coli (Invitrogen) before
purifying and sequencing the plasmid.
Digestions with restriction enzyme Eco RI (Roche) or PCR with M13 forward primer
(Invitrogen) were performed to confirm the insert in the vector.
Full-length cDNA amplification. The full cDNA fragment of the gene pgafp was obtained
by PCR with specific primers (Fwd-PrePro pgafp: 5’-ATGCAGATCACCAGCATTGCC3’ and Rev-PrePro/Mature pgafp: 5’-TCAAACTGGGGTCTGGCAGTC-3’) designed
from 5’ and 3’ ends. PCR reaction mixtures at a final volume of 50 µl contained 5µl of 5’RACE-Ready cDNA, 0.5 µM each of forward and reverse primers, 0.5 mM each of the
dNTPs, 0.1 vol of 10× PCR buffer (10 mM Tris-HCl, pH 8.8, 50 mM KCl, 0.1% Triton X100), 4 mM MgCl2 and 2.5 units pfu turbo polymerase (Stratagene). The PCR program
consisted of 35 cycles of denaturation (94ºC for 30 s), annealing (65ºC for 30 s), and
extension (72ºC for 3 min). PCR products were gel-purified and cloned into pCR2.1TOPO vectors, which were used to transform One Shot TOP10 Chemically Competent E.
coli (Invitrogen) before purifying and sequencing the plasmid. Digestions with restriction
enzyme Eco RI (Roche) or PCR with M13 forward primer (Invitrogen) were performed to
confirm if the insert was in the vector.
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Amplification of the whole pgafp gene
The gene sequence was amplified by PCR using Fwd-PrePro pgafp and RevPrePro/Mature pgafp primers. PCR mixtures were composed by 100 ng of genomic DNA,
0.5 µM each of forward and reverse primers, 0.5 mM each of the dNTPs, 0.1 vol of 10×
PCR buffer (10 mM Tris-HCl, pH 8.8, 50 mM KCl, 0.1% Triton X-100), 4 mM MgCl2,
and 2.5 units pfu turbo polymerase (Stratagene) and deionized water until a final volume of
50µl. The PCR program consisted of 30 cycles of denaturation (94ºC for 30 s), annealing
(65ºC for 30 s), and extension (72ºC for 3 min). PCR products were cloned in pCR2.1TOPO vector that later was used to transform cells of One Shot TOP10 Chemically
Competent E. coli. The plasmid was purified and sequenced. Introns sequences were
confirmed by GenScan software.
RESULTS
Partial amplification of pgafp gene using degenerated primers
The pgafp gene partial amplification was done using the two degenerate primers, pgafpDPF1 and pgafp-DPR2A (Table 1). After different reactions, the optimal annealing
temperature was 45.1ºC. PCR products run in agarose gel showed one small band of
around 75 bp that was amplified only in reactions with both the forward and the reverse
primers (Figure 1). This band was selected for cloning and sequencing.
Figure 1. Products obtained by PCR using DPF1 primer (F), DPF1+DPR2A primers (F+R),
and DPR2A (R) primer. M: DNA standards. The arrow indicates the selected band for
cloning and sequencing.
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After sequencing, the fragment found was 70 bp length (5’-tgaagcataatacgtgcacatacctaaag
ggtggaaagaaccatgtagtcaattgcggttctgccgctaa-3’), and the translated sequence (KHNTCTYL
KGGKNHVVNCGSAA) was compared in Basic Local Alignment Search Tool (BLAST)
showing common sequences with Anafp from Aspergillus niger.
Sequence of pgafp gene by RACE-PCR
Both 3’ and 5’ -ends of the gene pgafp were obtained by amplification from cDNA. mRNA
was extracted from a 5 days-old culture of P. chrysogenum RP42C and was used to obtain
the two different populations of 5’- and 3’-RACE-Ready cDNA by RT-PCR. The gene
specific primer 3’-RACE GSP (5’-GGGTGGAAAGAACCATGTAGTCAATTGCG-3’),
designed from the known pgafp sequence of genomic DNA, allowed to amplify the 3’-end
by RACE-PCR. The resulting product of about 450 bp included 121 bp of the 3’-end of
pgafp (Figure 2). Amplification of the 5’-end was carried out with a new gene specific
primer, 5’-RACE GSP (5’-AACTGGGGTCTGGCAGTCAACCCTC-3’), designed from
3’-end sequence. The obtained band, of approximately 440 bp, included the full pgafp
cDNA gene.
A)
B)
Figure 2. cDNA fragments of 5’- and 3’-RACE PCR (A and B, respectively).
M: DNA standards.
Amplifications of complete gene in genomic DNA and cDNA were performed by
designing two new primers Fwd-full pgafp (5’-ATGCAGATCACCAGCATTGCC-3’) and
Rev-full pgafp (5’-TCAAACTGGGGTCTGGCAGTC-3’). The sequence of genomic DNA
was 404 bp, whereas the amplified cDNA was 279 bp (Figure 3). The difference between
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the sequences revealed the existence of two introns of 63 and 62 bp (Figure 4), absent in
the cDNA, which were confirmed by GenScan software.
Figure 3. pgafp gene amplification of the with genomic DNA (1) and cDNA (2). M: DNA
standards.
5’-atgcagatcaccagcattgccattgtcttcttcgccgcaatgggtgcggttgctaacccc
M Q I T S I A I V F F A A M G A V A N P
atcgcgagggagtcggacgatcttgatgcccgagacgtacagcttagtaaattcggagga
I A R E S D D L D A R D V Q L S K F G G
gtaagttcttcttacaagacgtctatatagaaatagcactaacctttctgaaccacttta
<<
Intron 1
caggaatgcagcttgaaacacaacacgtgcacatacctaaagggtggaaagaaccatgta
>> E C S L K H N T C T Y L K G G K N H V
gtcaattgcggttcggccgccaacaagaaggtaggttccgattcgattcggggccaattg
V N C G S A A N K K <<
Intron 2
atttgttcttatcatttaatcttcatctacagtgcaagtctgatcgccaccactgtgaat
>> C K S D R H H C E
acgatgagcaccacaagagggttgactgccagaccccagtttga-3’
Y D E H H K R V D C Q T P V -
Figure 4. Full-length DNA sequence of pgafp gene. The open reading frame has been translated
to amino acids. Consensus splice sequences and internal marker of fungal introns are underlined.
The open reading frame was translated to protein by ExPASy proteomics server at the
Swiss Institute of Bioinformatics (http://www.expasy.org). Analysis of the signal peptidase
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cleavage site predicted by the program SignalP (Nielsen et al., 1997) suggested that the
pre-sequence consisted of the first 18 amino acids (MQITSIAIVFFAAMGAVA).
DISCUSSION
P. chrysogenum RP42C produces a high amount of the PgAFP in the first week of
incubation (Acosta, 2006). Thus, to obtain appropriate cDNA populations, a 5 days-old
culture was used to make sure that the mRNA from gene encoding PgAFP was actively
being produced. The RACE-PCR was effective to obtain pgafp gene. Thus, the designed
primers and obtained cDNA populations were adequate to amplify the 3’ and 5’ –ends of
pgafp. When the complete sequence was obtained, a difference between the pgafp gene
from genomic DNA (404 bp) and from cDNA (279 bp) could be observed (Figure 3).
Thus, the existence of two introns of 63 and 62 bp in the gene, which are absent in the
cDNA, was confirmed by GenScan software. The small size of these introns compared to
those of mammalian introns is a typical feature of fungal genes (Gurr et al., 1987). The 5’and 3’-ends of the two introns are quite similar (Figure 4) and coincide with the consensus
splice sequences for fungal introns: 5’-splice donor site GTNNGT (N= A, G, T or C) and
the consensus 3’-splice acceptor site DYAG (D= A, G or T; Y= C or T). In addition, the
first intron has the internal CTAAC sequence, very common in fungal introns (Wiesner et
al., 1988).
The pgafp open reading frame encodes a 92 amino acid-long precursor of the PgAFP. The
first 18 amino acids correspond with a secretion signal sequence according to the SignalP
program (Nielsen et al., 1997). The next residues (19-34) were not found in the amino acid
sequence obtained from the purified protein (Rodríguez-Martín et al., 2009). These 16
amino acids constitute a prosequence that would be removed before or during the release
of mature PgAFP. Prosequences play an important role by preventing protein activity
before secretion. Only after the prosequence is cleaved off, the mature protein adopts its
correct tertiary structure and gains its final activity. Antifungal proteins produced as
preproproteins, such as PAF and AFP (Nakaya et al., 1990; Marx et al., 1995), have been
described from other moulds.
The deduced mature PgAFP has 58 amino acid residues. It shows a high percentage (27.6
%) of basic amino acids. The isoelectric point was estimated to be 8.8 using the Compute
pI tool from Expasy (Gasteiger et al., 2005). The isoelectric point of PgAFP estimated by
isoelectrofocusing in previous studies (Acosta et al., 2009b) was 9.2, higher than the
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predicted value with Compute pI tool. However, the prediction can be not exact for highly
basic and small proteins (Gasteiger et al., 2005). A pI of 9.2 results in a net positive charge
under physiological conditions, which is a common characteristic among antifungal
proteins from moulds.
The accessibility of the whole genome sequence of an increasing number of
microorganisms facilitates the search for genes encoding potential antifungal proteins.
Three of these genes, those from Aspergillus niger, Neosartorya fischeri, and Gibberella
zeae, show partial homology with PgAFP protein. The sequences of the hypothetical
proteins from these genes as well as other reported antifungal proteins are aligned in Figure
5. The protein showing the highest homology (75%) with the precursor of PgAFP was the
hypothetical protein from A. niger. Despite both PAF and PgAFP precursors are produced
by Penicillium chrysogenum, they only share a similarity of 43%.
There are six cysteines conserved in all the similar proteins (Figure 5). Those residues
would play an important role on stabilizing of the tertiary structure of PgAFP by formation
of disulfide bridges (Nakaya et al. 1990; Marx et al., 1995; Lee et al., 1999).
PgAFP
Anafp
A.niger
N.fischeri
G.zeae
PAF
AFP
A.clavatus
---MQITSIAIVFFAAMGAVANPIARE---SDDLDARD--VQLSKFGGECSLKHNTCTYL
------------------------------------------LSKYGGECSLEHNTCTYR
---MQLTSIAIILFAAMGAIANPIAAE---ADNLVARE--AELSKYGGECSVEHNTCTYL
---MQITKISLFLFVGIGVVASPIHAE---SDGLNARAVNAADLEYKGECFTKDNTCKYK
---MQFSTIIPLFVAAMGVVATPVNSP---AQELDARGNLFPRLEYWGKCTKAENRCKYK
---MQITTVALFLFAAMGGVATPIESV---SNDLDARAEAGVLAKYTGKCTKSKNECKYK
MKFVSLASLGFALVAALGAVATPVEADSLTAGGLDARDESAVLATYNGKCYKKDNICKYK
MKVVSLASLGFALVAALGVVASPVDADSLAAGGLDARDESAVQATYDGKCYKKDNICKYK
PgAFP
Anafp
A.niger
N.fischeri
G.zeae
PAF
AFP
A.clavatus
K-GGKNHVVNCGSAANKKCKSDRHHCEYDEHHKRVDCQTPV
K-DGKNHVVSCPSAANLRCKTDRHHCRYDDHHKTVDCQTPV
K-GGKDHIVSCPSAANLRCKTERHHCEYDEHHKTVDCQTPV
ID-GKTYLAKCPSAANTKCEKDGNKCTYDSYNRKVKCDFRH
NDKGKDVLQNCPKFDNKKCTKDGNSCKWDSASKALTCY--NDAGKDTFIKCPKFDNKKCTKDNNKCTVDTYNNAVDCD--AQSGKTAICKCYV---KKCPRDGAKCEFDSYKGKCYC---AQSGKTAICKCYV---KVCPRDGAKCEFDSYKGKCYC----
Figure 5. Alignment of PgAFP amino acid sequences (Rodríguez-Martín et al., 2009) with
other antifungal proteins. Accession numbers in Genbank are XP_001391221 (hypothetical
protein from Aspergillus niger CBS 513.88), XP_001262586 (hypothetical protein from
Neosartorya fischeri NRRL 181), XP_384921 (hypothetical protein from Gibberella zeae PH1), AAA92718 (PAF from P. chrysogenum), P17737 (AFP from A. giganteus) and
XP_001267787 (hypothetical protein from Aspergillus clavatus). Common amino acids are
denoted by shaded background letters.
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In previous studies PgAFP showed a molecular weight of 6,494 Da by mass spectrometry
(Rodríguez-Martín et al., 2009). Translating the genetic sequence according to Gasteiger et
al. (2005) the predicted mass of the mature protein was 6,500 Da, very close to the weight
determined by mass spectrometry. In previous works, PgAFP protein was tested for having
glycosylations by treatments with deglycosilases and chemical compounds, and no linkedoligosaccharides were found. The minimal difference found between predicted and
experimental molecular weights confirms that this protein is not glycosylated.
Recently, the genome of P. chrysogenum Wisconsin 54-1255 has been sequenced (Van den
Berg et al., 2008). A putative protein can be found in BLAST (accession number
CAP80456) that differs from the precursor of PgAFP in only three amino acids. However,
the production of the protein by this mould has not been reported.
In conclusion, the gene enconding the antifungal protein PgAFP has been sequenced. Such
as similar proteins, PgAFP is synthesized as a long precursor with a peptide signal and a
presequence that are removed to obtain mature proteins. The characterization of pgafp
gene will permit its cloning for overproduction or heterologous expression of the protein in
other microorganisms.
ACKNOWLEDGEMENTS
This work was supported by the Spanish Ministry of Education and Science (AGL200406546-ALI) and FEDER. Andrea Rodríguez and Raquel Acosta were the recipients of FPI
grants from the Spanish Ministry of Education and Science. Authors would like to thank
Dr. Susan Liddell (Division of Animal Sciences) and Prof. Ian Connerton (Division of
Food Sciences) for their support during Andrea Rodriguez stay’s at Nottingham
University.
REFERENCES
Acosta, R. Doctoral Thesis: “Selección de Penicillium productores de péptidos
antifúngicos para su utilización en productos cárnicos madurados”. University of
Extremadura. Spain. 2006.
Acosta, R., Rodríguez-Martín, A., Martín, A., Núñez, F., and Asensio, M.A. Selection
of antifungal protein-producing molds from dry-cured meat products. Int. J. Food
Microbiol. Submitted (2009a).
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Acosta, R., Rodríguez-Martín, A., Bermúdez, E., Núñez, F., and Asensio, M.A. Partial
characterization and antifungal activity of a protein from Penicillium chrysogenum. Food
Microbiol. Submitted (2009b).
Gasteiger, E., Hoogland, C., Gattiker, A., Duvaud, S., Wilkins, M.R., Appel, R.D.,
and Bairoch, A. Protein Identification and Analysis Tools on the ExPASy Server;
(In) John M. Walker (ed): The Proteomics Protocols Handbook, Humana Press.
Full text - Copyright Humana Press (2005).
Geisen, R. P. nalgiovense carries a gene which is homologous to the paf gene of P.
chrysogenum which codes for an antifungal peptide. Int. J. Food Microbiol. 62: 95-101
(2000).
Gurr, S.J., Uncles, S.E., and Kinghorn, J.R. The structure and organization of nuclear
genes of filamentous fungi. In: Kinghorn, J.R. (Ed.). Gene structure in Eukaryotic
Microbes. IRL Press, Oxford, pp. 93-193 (1987).
Lacadena, J., Martínez del Pozo, A., Gasset, M., Patiño, B., Campos-Olivas, R.,
Vázquez, C., Martínez-Ruiz, A., Mancheño, J.M., Oñaderra, M., Gavilanes, J.G.
Characterization of the antifungal protein secreted by the mould Aspergillus giganteus.
Arch. Biochem. Biophys. 20: 273-281 (1995).
Lee, D.G., Shin, S.Y., Maeng, C-Y., Jin, Z.Z., Kim, K.L. and Hahm, K-S. Isolation and
characterization of a novel antifungal peptide from Aspergillus niger. Biochem. Biophy.
Res. Commu. 263: 646-651 (1999).
Marx, F., Haas, H., Reindl, M., Stöffler, G., Lottspeich, F. and Redl, B. Cloning,
structural organization and regulation of expression of the Penicillium chrysogenum paf
gene encoding an abundantly secreted protein with antifungal activity. Gene. 167: 167-171
(1995).
Nakaya, K., Omata, K., Okahashi, I., Nakamura, Y., Kolkenbrock, H., and Ulbrich,
N. Amino acid sequence and disulfide bridges of an antifungal protein isolated from
Aspergillus giganteus. Eur. J. Biochemistry. 193: 31-38 (1990).
Nielsen, H., Engelbrecht, J., Brunak, S., and Von Heijne, G. Identification of
prokaryotic and eukaryotic signal peptides and prediction of their cleavage sites. Protein
Eng. 10: 1-6. (1997).
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Rodríguez-Martín, A., Liddell, S., Acosta, R., Nuñez, F., and Asensio, M.A.
Characterization of PgAFP, a novel antifungal protein from Penicillium chrysogenum.
Manuscrito 1 (2009).
Skouri-Gargouri, H. and Gargouri, A. First isolation of a novel thermostable antifungal
peptide secreted by Aspergillus clavatus. Peptides. 29: 1871-1877 (2008).
Van den Berg, M.A., Albang, R., Albermann, K., Badger, J.H., Daran, J.M., Driessen,
A.J., Garcia-Estrada, C., Fedorova, N.D., Harris, D.M., Heijne, W.H., Joardar, V.,
Kiel, J.A., Kovalchuk, A., Martin, J.F., Nierman, W.C., Nijland, J.G., Pronk, J.T.,
Roubos, J.A., van der Klei, I.J., van Peij,N.N., Veenhuis,M., von Dohren, H., Wagner,
C., Wortman, J. and Bovenberg, R.A. Genome sequencing and analysis of the
filamentous fungus Penicillium chrysogenum. Nat. Biotechnol. 26: 1161-1168 (2008).
Wiesner, P., Beck, J., Beck, K.F., Ripka, S., Müller, G., Lücke, S. and Schweizer, E.
Isolation and sequence analysis of the fatty acid synthetase FAS2 gene from Penicillium
patulum. Eur. J. Biochem. 177: 69-79 (1988).
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Manuscrito 3: Preliminary evaluation of the toxicity and
sensitivity to different treatments of the antifungal protein
PgAFP from Penicillium chrysogenum
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Preliminary evaluation of the toxicity and sensitivity to different
treatments
of
the
antifungal
protein
PgAFP
from
Penicillium
chrysogenum
Andrea Rodríguez-Martín, Raquel Acosta, Félix Núñez, Miguel A. Asensio*
Higiene y Seguridad Alimentaria, Facultad de Veterinaria. Universidad de Extremadura, Avda. de la
Universidad, s/n. 10071- Cáceres, Spain.
* Corresponding author: Telephone (+34) 927 257 125; fax: (+34) 927 257 110.
E-mail: masensio@unex.es
ABSTRACT
The protein PgAFP obtained from Penicillium chrysogenum RP42C inhibits toxigenic
moulds found in foods. The aim of this work was to evaluate the potential toxicity of
PgAFP and to study basic characteristics of its antifungal activity, including the effect of
treatments and compounds common in foods. No haemolytic activity or toxicity on
Artemia salina larvae was detected. PgAFP showed neither chitinase nor chitosanase
activities. The protein showed a high stability to heat treatments up to 100ºC for 15 min,
and at pH values from 1 to 12. Some detergents potentially present in food as indirect
additives interfered with the antifungal activity, but tween 20 had no impact on the protein.
Only high concentration of other compounds, such as EDTA and sodium phosphate,
affected PgAFP activity when low levels of the protein were present. Monovalent ions
showed very little impact, whereas Ca2+ blocked the antifungal activity even at low
concentrations. These results indicate that PgAFP could be used to prevent mould growth
in foods with high salt levels or where thermal or acid stability are essential.
KEY WORDS: antifungal activity, food preservative, protective culture.
INTRODUCTION
Penicillium chrysogenum has been proposed as starter culture for mould fermented foods
(Martín et al., 2002; Benito et al., 2005), mainly due to its ability to produce volatile
compounds that contribute to flavour development (Ramírez et al., 1995; Jacobsen and
Hinrichsen, 1997; Martín et al., 2006). However, the continuous evolution of the physicochemical characteristics along ripening can favour the development of undesirable moulds,
which would not only lead to a negative flavour of the product (Lund et al., 1995), but also
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to mycotoxin production (Núñez et al., 1996, 2000; Nielsen et al., 1998; López-Díaz et al.,
2001; Sosa et al., 2002; Sabater-Vilar et al., 2003). Some mould can produce different
antifungal compounds, including proteins. The use of natural antifungal proteins could be
attractive to satisfy consumer's demand. Therefore, fungal cultures with antagonistic
activity against spoilage and toxigenic moulds could be of interest to improve the
microbial safety of the product.
In previous studies, P. chrysogenum RP42C isolated from dry-cured ham showed a broadspectrum inhibition against toxigenic moulds, including Aspergillus niger, A. flavus, and P.
commune due to the protein PgAFP (Acosta et al., 2009a, b).
Various mechanisms of action have been described for antifungal proteins, including
interference in synthesis of DNA, RNA, or proteins, damage to cellular ribosome (Hwu et
al., 2000), and membrane channel and pore formation (Thevissen et al., 1999).
Nevertheless, the most useful antifungal proteins are those specific against fungal
membrane and cell wall, because of their lesser toxic effects on mammalian cells (Groll et
al., 1998). Cationic antifungal proteins, such as AFP from A. giganteus and PAF from P.
chrysogenum, have proved to be specific for fungal membranes (Marx et al., 2008; Meyer
et al., 2008) and not for mammalian cells (Szappanos et al., 2005 and 2006). Interestingly,
PgAFP shows similar characteristics to AFP and PAF, both in the amino acid sequence and
the isoelectric point (Marx, 2004; Acosta et al., 2009b; Rodríguez-Martín et al., 2009a, b).
A prerequisite for any application of antifungal proteins or the producer mould on foods is
the lack of toxicity for consumers. The use of antifungal proteins from ascomycetes can be
limited by chemical compounds present and physical treatments applied to foods. Many
antifungal proteins, especially membrane-acting peptides, are sensitive to cations, which
can be a crucial point in the application to food products with high salt concentrations
(Theis and Stahl, 2004). Given that PgAFP is produced by a mould isolated from a
substrate with high salt concentration, its antifungal activity could withstand high-cation
levels. Moreover, antifungal proteins should also be active under the singular conditions of
foods where moulds can grow, particularly in a wide range of pH values. Other compounds
that can be present on foods, such as detergents and chelators, may also interfere with
antifungal proteins. In addition, stability to heat treatments can be of utmost interest for the
potential use of antifungal proteins as food preservative.
The aim of this work was to carry out a preliminary evaluation of the potential toxicity of
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PgAFP, as well as the impact of salts, heat treatments, different pH values, and detergents
on its antifungal activity.
MATERIALS AND METHODS
Microbial strains
The PgAFP producing mould, P. chrysogenum RP42C, and the toxigenic moulds
Penicillium restrictum Pr341 and Aspergillus niger An261, were isolated from dry-cured
ham and belong to the fungal collection of Food Hygiene (University of Extremadura,
Spain).
PgAFP purification
P. chrysogenum RP42C was grown at 25ºC for 15 days in malt extract broth made of
glucose 2% (w/v), malt extract 2% (w/v), and peptone 0.1% (w/v), pH 4.5. Then, the
medium was filtered through a 0.45 µm diameter pore size cellulose acetate membrane and
PgAFP was purified using an ÄKTA fast protein liquid chromatography system
(Amersham Pharmacia Biotech AB, Uppsala, Sweden) according to Acosta et al. (2009a).
For this, filtered medium was loaded on a HiTrap SP HP 5 ml ion exchange column
(Amersham Pharmacia Biotech AB) equilibrated with 0.02 M sodium acetate buffer, pH
4.5. Bound proteins were eluted with the following gradient of 0.02 M sodium acetate, 1 M
NaCl buffer, pH 4.5: 0% to 25% in 15 column volumes (CV), increase to 100% in 2 CV,
and finally keeping 100% for 15 CV. Eluted proteins were detected at 214 nm. The
fraction that showed antifungal activity in previous studies (Acosta et al., 2009a) was
purified using a HiLoad 26/60 Superdex 75 prep grade gel filtration column (Amersham
Pharmacia Biotech AB) equilibrated with 50 mM sodium phosphate, 0.15 M NaCl buffer,
pH 7, at a constant flow rate of 2.5 ml/min. Data collection and processing were done
using the UNICORN software 4.12 (Amersham Pharmacia Biotech AB). Finally, the
fraction containing PgAFP was desalted and concentrated with YM-3 microcon centrifugal
filter units (Millipore Iberica S.A.U., Madrid, Spain) and kept at -20ºC.
Protein was quantified by Johnson method according to Córdoba (1990). An aliquot of the
purified protein (100 µl) was dried at 100 ºC. The sample was digested by adding 200 µl of
sulfuric acid at 120 ºC. Then, 4.8 ml of distilled water, 3 ml of 4 N NaOH, and 2 ml of
Nessler reactive made of KI 0.4% (w/v), 0.4% HgI2 (w/v), and 0.175% (w/v) arabic gum
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were added. After 10 min at room temperature, the absorbance was measured at 490 nm in
a spectrophotometer. A calibration curve was obtained with different concentrations of
ammonium sulfate.
Sodium dodecyl sulphate-polyacrylamide gel eletrophoresis (SDS-PAGE)
SDS-PAGE was performed as described Laemmli (1970). The protein PgAFP was
separated on a 4% (v/v) polyacrylamide stacking and 8% (w/v) polyacrylamide resolving
gels. Gels were stained with Imperial Protein Stain (Pierce Biotechnology, Inc., Rockford,
U.S.A.). The molecular weight was determined by comparison of its electrophoretic
mobility with proteins of the molecular mass marker (6.5-66 kDa) from Sigma (SigmaAldrich Química S.A., Madrid, Spain).
Haemolytic activity assay
The haemolytic activity of the protein was determined using sheep defibrinated blood
(Oxoid). The erythrocytes were isolated by centrifugation (1,000 ×g for 6 min at room
temperature) and washed three times with 50 mM sodium phosphate, 0.15 M NaCl buffer,
pH 7. The cell concentration in the stock suspension was adjusted to 1×109 cells/ml. A
volume of 50 µl of cells suspension was added to 200 µl of sterile concentrated protein
(300 µg/ml) and incubated at 37ºC. After 0, 8, and 24 h, the samples were placed on ice,
immediately centrifuged at 4ºC, and the supernatant was carefully collected. The
haemolysis was determined by measuring the absorbance of the supernatant at 570 nm
(Moerman et al., 2002) and 541 nm (Lai et al., 2005). Controls for 0% and 100%
haemolysis were obtained suspending sheep red blood cells in the buffer or distilled water,
respectively. The haemolysis percentage was calculated using the following equation:
Percentage haemolysis of supernatants = [(Abs in the peptide solution − Abs in buffer)/Abs
in water solution] × 100.
Three independent experiments were conducted with triplicate readings for each one.
Toxicity assay using brine shrimp larvae (Artemia salina)
The toxicity of PgAFP was evaluated using brine shrimp larvae, following the Harwig and
Scott method (1971) with modifications. Brine shrimp medium (BSM) was made of 0.3%
(w/v) dipotassium glycerophosphate, 3% (w/v) NaCl, 0.03% (w/v) CaCl2 • 2H2O, 0.05%
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(w/v) MgSO4 • 7H2O, 0.15% (w/v) MgCl2 • 6H2O, 0.08% (w/v) KCl, 0.01% (w/v)
MgBr2 • 6H2O, and 0.6% (w/v) glycine. The medium was adjusted to pH 6.5. Brine
shrimp larvae (100-200 mg) were grown in 100 ml BSM under continuous shaking (140
rpm) for 30 h at 30ºC. The test was carried out in Erlenmeyer flasks containing 20 to 40
larvae in 10 ml BSM. PgAFP was added at final concentrations of 3, 16, 33, 57, and 75
µg/ml. Buffer without PgAFP was used in negative controls. Dead larvae were counted just
before incubation and after 24 h at 30ºC in a shaker at 80 rpm. Finally, the mortality
percentage was calculated. PgAFP was indicated as nontoxic when death rate of larvae was
not significantly different (p <0.05) from spontaneous mortality in controls.
Evaluation of chitinase and chitosanase activities
Chitinase and chitosanase activities were determined with a modification of the 3,5dinitrosalicylic acid method (Su et al., 2006) by measuring the reducing sugars liberated
during the hydrolysis of chitin or chitosan. One ml of the 1% (w/v) chitin or chitosan
(Sigma-Aldrich) in 1% (v/v) acetic acid was added to 1 ml of PgAFP (300 µg/ml) and
incubated for 2-3 h at 37ºC. Then, to detect the hydrolisis reaction, 2 ml of dinitrosalicylic
acid reagent, made of 1% (w/v) 3,5-dinitrosalicylic acid, 0.05% (w/v) sodium sulphite and
1% (w/v) phenol dissolved, were added and the sample was incubated at 100ºC for 45 min.
Finally, 1 ml of 40% (w/v) potassium sodium tartrate was added and the solution was
centrifuged. The supernatant was measured at 520 and 575 nm. To quantify the reducing
sugars liberated a calibration curve was obtained with different concentrations of D-(+)Glucosamine hydrochloride (Sigma-Aldrich). A chitinase from Streptomyces griseus
(Sigma-Aldrich) with both chitinase and chitosanase activities was used as positive control.
Preparation of colloidal chitin: Colloidal chitin was obtained following the method of
Roberts and Selitrennikott with minor modifications (Harighi et al., 2007). Ten grams of
chitin powder were added slowly into 100 ml of H3PO4 (85%) at 25ºC under vigorous
shaking for 2 h. This solution was carefully added over 1 liter of absolute ethanol at -20ºC.
The mix was shaken for 30 min and stored until use. Ten ml of suspension were
centrifuged, the pellet was washed three times with 50 mM sodium phosphate buffer pH 7,
and finally resuspended in distilled water. The final concentration of colloidal chitin was
around 1%.
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Chitinase activity after gel electrophoresis
The chitinase activity assay was carried out as described by Trudel and Asselin (1989).
PAGE-SDS was performed in 12% (v/v) polyacrylamide gradient gel containing 0.01%
(w/v) glycol chitin and 0.1% (w/v) SDS. A stacking gel with 4% (v/v) polyacrylamide
containing 0.1% (w/v) SDS was used. Samples were boiled for 5 min with 20% (v/v)
glycerol, 0.025% (w/v) bromophenol blue, 2% (w/v) SDS and 5% (v/v) -mercaptoethanol
in 62.5 mM Tris-HCl (pH 6.8). The gel was run at 100 V for 1 h.
After electrophoresis, the gel was incubated for 2 h at 37 ºC under continuous shaking in
100 mM sodium acetate buffer (pH 5.0) containing 1% (v/v) Triton X-100. Then, the gel
was stained with a 0.01% (w/v) calcofluor white M2R (Sigma-Aldrich) in 500 mM TrisHCl (pH 8.9). After 5 min, this solution was removed and the gel was incubated in distilled
water at room temperature for about 1 h. Lytic zones were visualized in the UV
transilluminator Syngene G-Box (Synoptics, Fredericks, USA). The gel was photographed
and analyzed with the GeneSnap software (Synoptics). An additional gel was stained only
with Imperial Protein Stain (Pierce Biotechnology, Rockford, U.S.A.). The chitinase from
S. griseus (Sigma) was used as positive control. The molecular mass marker used was 666.5 kDa (Sigma).
Glycol chitin synthesis: 0.2 g glycol chitosan was dissolved in 4 ml of 10 % (v/v) acetic
acid and the mixture was shaken overnight at 22ºC. Methanol (18 ml) was slowly added
and the solution was filtered through 0.45 µm pore size cellulose acetate membrane filters.
Acetic anhydride (0.3 ml) was added and kept for 30 min. The resulting gel was cut into
small pieces and after adding 1 vol of methanol was shaken for 4 min. This suspension was
centrifuged at 27000 ×g at 4ºC for 15 min and the pellet was resuspended in 20 ml of
0.02% (w/v) sodium azide. The final concentration of the glycol chitin was 1% (w/v).
Evaluation of antifungal activity
Quantitative assay for fungal growth inhibition was performed following the protocol
developed by Broekaert et al. (1990) with some modifications (Acosta et al., 2009a). The
toxigenic strains P. restrictum Pr341 and A. niger An261 (Núñez et al., 1996; Díaz, 1999)
were used as reference strains. To perform the inhibition test in microtitter plates, 100 µl of
the fraction obtained by gel filtration chromatography were dispensed in triplicate wells.
Each well was loaded with 100 µl of concentrated malt extract broth (4% malt extract, 4%
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glucose and 0.2% peptone, pH 4.5) inoculated with the reference strain to c.a. 106
spores/ml. The plates were incubated for up to 96 h at 25ºC. Fungal growth was
determined by measuring the optical density variation at 595 nm after 96 h.
Effect of heat-treatments
The study of the protein stability after heat treatment was carried out according to Okkers
et al. (1999). Aliquots of PgAFP protein (300 µg/ml) were exposed to different
combinations of temperatures (60, 80, and 100ºC) and times (10, 20, and 30 min), as well
as to sterilization at 121ºC for 15 min. After the heat-treatment, the samples were cooled
on ice and tested for antifungal activity in microtiter plates.
PgAFP antifungal activity after exposure at different pH values
PgAFP (300 µg/ml) was retained in microcon centrifugal filter units YM-3 (Millipore) and
then dissolved in different buffers at the following values: pH 1, 2 (HCl/KCl); pH 3
(glycine/HCl); pH 4, 5, 6 (citric acid/sodium phosphate); pH 7 (sodium phosphate/NaCl);
pH 8 (Tris/HCl); pH 9, 10 (Tris/NaOH); and pH 12 (KCl/NaOH). Samples were incubated
at 25ºC for 2 h and the pH adjusted to 4.5 before testing the antifungal activity.
Effect of different chemicals on PgAFP antifungal activity
To study the influence of some chemicals on the antifungal activity, several assays were
carried out with 3, 30, and 300 µg PgAFP/ml in presence of the following compounds:
NaCl, CaCl2, KCl, or MgCl2 (0.1, 1, 10, and 100 mM); EDTA or EGTA at 0.01, 0.1, and 1
mM; 1% of Triton X-100, tween-20, tween-80, SDS, or deoxycholic acid; NaH2PO4 at 0,
0.1, 1, 10, and 100 mM.
Statistical analysis
Data were statistically analyzed by SPSS software package (version 13.0. SPSS Inc.,
Chicago, USA) for a variance analysis (one way ANOVA) and the means were separated
by the Tukey honest significant difference test.
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RESULTS
Evaluation of PgAFP toxicity
None of the readings at either 570 or 541 nm during the incubation time revealed
haemolytic activity. The concentration of PgAFP tested (240 µg/ml) showed 0%
haemolysis after 24 h.
In an additional assay with Artemia salina, the larvae mortality was 0% even at the highest
concentration of the protein 75 µg/ml after 24 h incubation. Therefore, PgAFP showed no
toxicity at any concentration tested.
Chitosanase and chitinase activities
No chitosanase or chitinase activity was observed by the 3,5-dinitrosalicylic acid method,
as no glucosamine was liberated by PgAFP in the conditions assayed.
To further evaluate the chitinase activity, chitin hydrolysis was tested by electrophoresis in
a glycol chitin gel. No activity was observed loading the polyacrylamide gel with 50 µg of
PgAFP (Figure 1). Lytic zones by chitinase from S. griseus used as positive control were
observed as black bands after staining with calcofluor white M2R.
Figure 1. Chitinase activity by gel electrophoresis. SDS-PAGE with glycol chitin was
stained with Imperial Protein Stain (A) and Calcofluor White M2R (B). M: protein marker,
Q: chitinase from S. griseus (100 µg).
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Effect of heat-treatments and different pH values
PgAFP protein showed a remarkable stability at high temperatures. PgAFP kept antifungal
activity against P. restrictum and A. niger even after being heat-treated at 80ºC for 30 min
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or 100ºC for 15 min (Figure 2).
10 20 30 10 20 30 3 10 15 20 30 15 15a
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10 20 30 10 20 30 3 10 15 20 30 15 15a
60ºC
80ºC
100ºC
121ºC
Figure 2. Antifungal activity of heat-treated PgAFP expressed as mean and standard deviation of
reference moulds growth after 96 h. Asterisks indicate statistically significant differences to
positive control with non heat-treated PgAFP (***p<0.001). C+: positive control with non heattreated PgAFP; C-: negative control without PgAFP. a: Autoclave cycle.
Incubation in buffers at pH values from 1 to 12 had no effect on the stability of PgAFP and
the protein remained active against the reference moulds (Figure 3).
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Figure 3. Antifungal activity of PgAFP after treatment at different pH values expressed as mean
and standard deviation of reference moulds growth after 96 h. C-: negative control without PgAFP.
Effect of cations
The impact of different monovalent and divalent cations at 0.1, 1, 10, and 100 mM on the
inhibitory activity of PgAFP (3, 30, and 300 µg/ml) over P. restrictum and A. niger was
tested (Figure 4). Monovalent cations did not seem to exert a strong impact on the
antifungal activity. The antifungal activity on A niger decreased only with at the highest
concentration of K+ or Na+ (100 mM) and the lowest level of PgAFP. The remaining
conditions tested did not alter the activity of PgAFP against P. restrictum and A. niger.
Divalent cations showed a deeper impact on PgAFP activity against both sensitive moulds.
The highest level (100 mM) of Ca2+ decreased PgAFP activity, irrespectively of the protein
concentration tested. Even 10 mM Ca2+ reduced the inhibition with 3 µg/ml of PgAFP.
Mg2+ showed a lower effect than Ca2+, given that only 100 mM decreased the activity with
low PgAFP concentrations.
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Figure 4. Effect of monovalent (A) and divalent (B) cations on antifungal activity of PgAFP
expressed as mean and standard deviation of reference moulds growth after 96 h. Asterisks indicate
statistically significant differences to the negative control without PgAFP (*p<0.05; **p<0.01;
***p<0.001).
Effect of calcium chelators
EDTA and EGTA showed a different effect on PgAFP activity. EGTA had no influence on
PgAFP antifungal activity, even at the highest concentration tested (1 mM). EDTA at
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levels up to 0.01 mM had no impact on PgAFP antifungal activity against either sensitive
mould, whereas 0.1 mM limited the activity of lower levels of PgAFP just on P. restrictum
(Figure 5). The highest concentration of EDTA (1 mM) prevented growth of P. restrictum
and A. niger (data not shown), and no further effect could be established.
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Figure 5. Effect of EDTA on the antifungal activity of PgAFP expressed as mean and standard
deviation of reference moulds growth after 96 h. Asterisks indicate statistically significant
differences to the negative control without PgAFP (***p<0.001).
Effect of detergents
Even though non-ionic detergents limited the growth of the reference moulds, it was
possible to evaluate their effect on PgAFP activity. However, SDS totally prevented
growth of reference moulds, and no effect on PgAFP could be established. For both
sensitive strains, tween 20 had no effect, tween 80 only reduced the activity with 3 µg/ml
of PgAFP, and Triton X-100 decreased the activity at all PgAFP concentrations (Figure 6).
With deoxycholic acid PgAFP did not show antifungal activity against P. restrictum,
whereas the effect of PgAFP on A. niger could not be observed due to the extremely poor
growth of this mould.
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Figure 6. Effect of Triton X-100 (T100), tween 20 (T20), tween 80 (T80), and deoxycholic acid
(DAc) on the antifungal activity of PgAFP expressed as mean and standard deviation of reference
moulds growth after 96 h. Asterisks indicate statistically significant differences to the negative
control without PgAFP (**p<0.01; ***p<0.001).
Effect of sodium phosphate
Sodium phosphate influenced the antifungal activity against both reference moulds,
particularly on A. niger (Figure 7). With P. restrictum, the effect was limited to high levels
of phosphate (10-100 mM) with the lowest PgAFP concentration. However, the lowest
phosphate concentration (0.1 mM) produced a significant effect on the activity against A.
niger with 3 µg/ml of PgAFP. This effect increased gradually with the phosphate
concentration.
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Figure 7. Effect of phosphate on the antifungal activity of PgAFP expressed as mean and standard
deviation of reference moulds growth after 96 h. Asterisks indicate statistically significant
differences to the negative control without PgAFP (**p<0.01; ***p<0.001).
DISCUSSION
P. chrysogenum RP42C has been proposed as protective culture on mould fermented foods
(Acosta et al., 2009b), and the use of PgAFP as additive could also be of interest to prevent
unwanted moulds.
Common methods to evaluate safety are the haemolytic assay (Theis et al, 2004) and the
brine shrimp larvae (Artemia salina) test (Harwig and Scott, 1971; Milhem et al., 2008;
Chohan, 2009). The absence of cytolytic activity to red blood cells is generally accepted as
proof that the protein can be regarded as safe (Van’t et al., 2001). Brine shrimp larvae
(Artemia salina) test has been also used to screen fungal toxins (Harwig and Scott, 1971)
or synthetic compounds (Milhem et al., 2008; Chohan, 2009) for toxicity. The results here
presented revealed neither haemolytic activity against red blood cells nor lethal effects on
brine shrimp larvae when PgAFP was tested up to 240 and 75 µg/ml, respectively.
The information available on other antifungal proteins from moulds reveals that neither
PAF nor AFP was toxic up to 100 µg/ml to human umbilical vein endothelial cells
(Szappanos et al., 2005, 2006). There are also bacterial antifungal proteins non-toxic to
blood red cells, such as PPEBL21 from Escherichia coli at 1,250 µg/ml (Yadav et al.,
2007). On the other hand, some antifungal proteins from bacteria have shown toxic effects.
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SAP protein isolated from Streptomyces sp. strain AP77 was non-toxic up to 250 µg/ml to
human dermal fibroblasts but toxic at all higher doses (Woo et al., 2002). A more potent
toxicity was found for syringomycin, a protein that has shown 100% haemolysis at 20
µg/ml (Sorensen et al., 1996). Antifungal peptides purified from Bacillus cereus were also
reported to be highly toxic to human erythrocytes (Latoud et al., 1986). The toxicity of
most reported antifungal proteins to mammalian cells has become a major limitation for
their use to develop new antifungal drugs or preparing improved antifungal formulations.
Therefore, the lack of toxicity showed by PgAFP in these preliminary tests supports its
potential for food use, although a more complete evaluation is required.
It is known that most chitin containing organisms produce chitinases, which are
presumably required for morphogenesis of cell walls and exoskeleton (Gooday, 1971).
Apart from these functions, these enzymes have also been found to be involved in the host
defence against pathogens (Somashekar and Joseph, 1996). Chitinases from moulds have
been described to have antifungal activity, such as from Thermomyces lanuginosus (Guo et
al., 2005) and from Fusarium chlamydosporum (Mathivanan et al., 1998). In addition, in a
parallel work an antifungal protein from P. chrysogenum AS51D that hydrolyses
deacetylated chitin has been described (Rodríguez-Martín et al., 2009c). However, PgAFP
did not show either chitinase or chitosanase activities when tested in solution or in gel
(Figure 1). Therefore, the antifungal activity of this protein cannot be based on the
degradation of these compounds commonly found in fungal cell wall.
Some detergents can be present on various foods as a consequence of their use as direct or
indirect additives. Detergents are unique among protein denaturants, and some of them can
alter protein conformation at low concentrations. The addition of detergents can either
increase or decrease the helical conformation, and even convert
forms into helices
(Naeem et al., 2006). The effect of ionic and non-ionic detergents on PgAFP varied
according to the sensitive mould and the protein concentration tested (Figure 6). Triton X100 and deoxycholic acid inhibited the antifungal activity, irrespectively of PgAFP
concentration. However, the inhibitory effect of both tweens was limited to tween 80 at the
lowest PgAFP concentration tested. Ionic detergents tend to denature proteins by altering
native three dimensional structures. Some ionic detergents, such as deoxycholic acid, can
also act as open-channel blockers, as it has been shown for porins (Duret and Delcour,
2006). No further information is available to suggest whether the effect of deoxycholic
acid is mainly due to changes in the tertiary structure of PgAFP or in the fungal membrane.
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In this study, the non-ionic detergent Triton X-100 seems to counteract the antifungal
activity of PgAFP. However, the partial inhibition on growth of reference moulds made it
difficult to evaluate the particular impact of Triton X-100 on PgAFP activity.
Finally, the activity of PgAFP decreased as sodium phosphate concentration increased,
although the protein remained active up to 100 mM of phosphate.
Among the main factors that can interfere with the antifungal activity in foods are salts, pH
and heat treatments. PgAFP was highly resistant to a wide range of temperature and pH
values. As it is shown in figure 2, PgAFP displayed antifungal activity after treatment at
80ºC for 30 min, and even at 100ºC for 15 min. Similarly, incubation in buffers at pH
values from 1 to 12 had no effect on PgAFP antifungal activity. Additionally, PgAFP
showed a high stability to proteases, such as pepsin and papain (Acosta, 2006). These
characteristics are similar to those of other antifungal proteins from moulds. AFP from A.
giganteus is stable at high temperature, protease degradation, and a broad pH range due to
a specific folding pattern (Lacadena et al., 1995). The mature form of AFP folds into five
antiparallel
strands which define a small and compact -barrel, stabilized by internal
disulfide bridges formed by eight cysteine residues (Campos-Olivas et al., 1995). It has
been assumed that Anafp, NAF, and PAF have the same tertiary structure, although only
three disulfide bridges can be formed by the six cysteine residues present in these proteins
(Marx, 2004). PgAFP has common characteristics to these antifungal proteins, including a
high stability, six cysteine residues, and a quite high similarity in the amino acid sequence
(Rodríguez-Martín et al., 2009a, b). Thus, PgAFP could also have a tertiary structure based
on a compact -barrel.
On the other hand, cation sensitivity has been described for basic antimicrobial proteins,
including AFP and PAF. In fact, AFP activity decreases in the presence of Ca2+, K+, and
Na+ ions (Theis et al., 2003), and PAF activity is sensitive to Mg2+, K+ and Na+ (Kaiserer
et al., 2003). Cations seem to disturb the interaction of antifungal proteins with negatively
charged membrane components (Barkai-Golan et al., 1978) or specific receptors
(Broekaert et al., 1992). The latter has been suggested for AFP (Theis et al., 2003) and
PAF (Kaiserer et al., 2003).
The influence of high cation levels on PgAFP depended on the ion charge, protein
concentration and the sensitive mould. The highest effect was obtained with the divalent
cations (Figure 4). Defensins are also more sensitive to divalent than to monovalent cations
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(Terras et al., 1992), particularly to Ca2+ (Lehrer et al., 1989; Cociancich et al., 1993;
Osborn et al., 1995). This indicates that the effect of monovalent cations depended on the
reference mould, with no effect on the activity against P. restrictum. The antagonistic
effect of ions over plant defensins is dependent on the fungus and thus on the conformation
of the putative target site (Terras et al., 1992; Osborn et al., 1995). Thus, the distinct effect
of cations on PgAFP activity against each sensitive mould tested can be explained by
differences on receptor conformation. In addition, the results obtained for PgAFP suggest
that the ratio of protein to ion concentration plays a decisive role on the interaction with
protein activity.
EDTA can also be present in foods due to its use as metal sequestrant additive. The ability
of EDTA and EGTA for chelating ions is well known, being EDTA the most potent of
them (Choucair et al., 2006). However, EDTA chelates both Ca2+ and Mg2+, whereas
EGTA is more specific for Ca2+ ions. EGTA did not influence PgAFP activity, whereas
EDTA inhibited this activity only against P. restrictum at the lower concentrations of the
protein tested (Figure 5). Thus, an excess of divalent cations has a stronger effect on
PgAFP activity than the lack of those ions.
In summary, PgAFP protein is stable over a wide range of pH values and active in
presence of high concentrations of ions and at high temperatures. All this information
revealed that PgAFP has potential to prevent mould growth in a wide range of foods with
high salt levels or where thermal or acid stability are essential. However, the use of this
protein as additive requires an in-depth toxicity evaluation.
ACKNOWLEDGEMENTS
This work was supported by the Spanish Ministry of Education and Science (AGL200406546-ALI) and FEDER. Andrea Rodríguez and Raquel Acosta were the recipients of a
FPI grant from the Spanish Ministry of Education and Science.
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Acosta, R. Doctoral Thesis: “Selección de Penicillium productores de péptidos
antifúngicos para su utilización en productos cárnicos madurados”. University of
Extremadura (2006).
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Acosta, R., Rodríguez-Martín, A., Martín, A., Núñez, F., and Asensio, M.A. Selection
of antifungal protein-producing molds from dry-cured meat products. Int. J. Food
Microbiol. Submitted (2009a).
Acosta, R., Rodríguez-Martín, A., Bermúdez, E., Núñez, F., and Asensio, M.A. Partial
characterization and antifungal activity of a protein from Penicillium chrysogenum. Food
Microbiol. Submitted (2009b).
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Manuscrito 4: Characterization of PgChP, a novel chitosanase
with antifungal activity from Penicillium chrysogenum
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Characterization of PgChP, a novel chitosanase with antifungal activity
from Penicillium chrysogenum
Andrea Rodríguez-Martín, Susan Liddella, Raquel Acosta, Félix Núñez, Miguel A.
Asensio*
Higiene y Seguridad Alimentaria, Facultad de Veterinaria. Universidad de Extremadura, Avda. de la
Universidad, s/n. 10071- Cáceres, Spain.
a
Division of Animal Sciences, University of Nottingham. Sutton Bonington Campus. Loughborough,
Leicestershire. LE12 5RD. United Kingdom.
* Corresponding author: Telephone (+34) 927 257 125; fax: (+34) 927 257 110.
E-mail: masensio@unex.es
ABSTRACT
A new fungal chitosanase with antifungal activity, PgChP from Penicillium chrysogenum,
is described. Two isoforms were found by SDS-PAGE in the purified extract of PgChP.
After enzymatic deglycosylation only the smaller isoform was observed by SDS-PAGE.
Identical amino acid sequences were obtained from the two proteins and these showed a
high similarity with fungal chitosanases. Analysis of the molecular mass by electrospray
ionization mass spectrometry (ESI-MS) revealed the existence of six peaks from 30 to 31
kDa differentiated by a mass multiple of 162 Da, the typical mass of galactose or mannose
residues. This could be due to different levels of glycosylation of the same protein PgChP.
The isoelectric point was aproximately pH 5 estimated by two-dimensional polyacrylamide
gel electrophoresis. PgChP is the first chitosanase to be described from P. chrysogenum.
KEY
WORDS:
antifungal
protein,
food
preservative,
glycosylation,
N-linked
oligosaccharides, protective culture, toxigenic mould.
INTRODUCTION
The antifungal protein PgChP was selected in previous studies (Acosta et al., 2009a) from
the strain AS51D isolated from dry-cured ham and characterized as Penicillium
chrysogenum according to classical taxonomical methods (Pitt and Hocking, 1997). PgChP
was purified by fast protein liquid chomatography and showed antifungal activity against
selected toxigenic moulds (Acosta et al., 2009a).
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Antifungal proteins have been described from diverse organisms including plants and
animals, but only scant information is available about antifungal proteins of fungal origin.
Many antifungal proteins have interesting applications because they attack compounds of
fungal walls that are not present in mammalian cells, such as glucanases, chitinases and
chitosanases (Adams, 2004). Fungal infections and contaminations have led to an
increasing demand for antifungal drugs in diverse fields including agriculture, medicine
and food protection. Many antifungal drugs have low efficacy rates, show severe side
effects and cause resistance on the microorganisms, as antibiotics (Meyer, 2008). This
makes it necessary to study new compounds, including fungal proteins such as PgChP.
However, before any application of PgChP, an exhaustive characterization of the protein is
required.
In previous studies, another antifungal protein from P. chrysogenum, named PgAFP,
showed evidence of glycosylation by periodic acid-Schiff staining (Acosta et al., 2009b).
Glycan chains in glycoproteins participate in many biological processes, such as protein
interactions, or confer stability on the protein against proteases or heat treatments (Morelle
et al., 2006). These modifications introduce variations in the molecular mass and net
charge of the protein. Enzymatic and chemical deglycosylation treatments followed by
SDS-PAGE and electrospray ionization mass spectrometry are commonly used to evaluate
this sort of post-translational modification (Claverol et al., 2003). Additional information
on the molecular mass and amino acid sequences is typically obtained by mass
spectrometry after digestion in-gel or in-solution with proteolytic enzymes (Habermann et
al., 2004).
This paper describes the amino acid sequencing, molecular mass determination and
isoelectric point estimation of the antifungal protein PgChP.
MATERIALS AND METHODS
Microbial strain
P. chrysogenum AS51D was isolated from dry-cured ham, being identified according to
classical morphological and biochemical studies (Acosta et al., 2009a).
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Genetic identification of the mould
Genetic characterization of AS51D was carried out by amplifying the fragments 18S-28S
rRNA intergenic spacer (ITS) by polymerase chain reaction (White et al., 1990).
DNA isolation. The mould AS51D was grown in malt extract broth, made of glucose 2%
(w/v), malt extract 2% (w/v), and peptone 0.1% (w/v), pH 4.5, at 25ºC for 5 days. Two
grams of mycelium were broken with morter and pestle after freezing by adding liquid
nitrogen. Lysis was performed by homogenizing the powder in 4 ml 0.05 M Tris-HCl, pH
8, 0.005 M ethylenediamine tetraacetic acid (EDTA), 0.05 M NaCl (TES) buffer and 1%
(w/v) sodium dodecyl sulfate (SDS). Afterwards, 200 µl proteinase K (20 µg/µl) was
added, and the mixture was incubated at 60ºC for 35 min, cooled on ice and extracted with
phenol-chloroform-isoamyl alcohol (25:24:1). The suspension was centrifuged at 3,000 ×g
for 2 min at 4ºC. The upper phase containing DNA was transferred to a fresh tube and
precipitated by the addition of 3 M sodium acetate pH 5.2 to a final concentration of 10%
(w/v) and two volumes of cold ethanol. After centrifugation, the pellet was washed with
70% (v/v) ethanol, centrifuged again in the conditions mentioned above, resuspended in
sterile water and treated with 50 µl RNase (10 µg/µl) at 37ºC for 60 min. Finally, DNA
was extracted with phenol-chloroform-isoamyl alcohol and precipitated again as indicated
above, resuspended in water and stored at -70ºC. The quantity and quality of purified DNA
was determined spectrophotometrically in a Biophotometer (Eppendorf AG, Hamburg,
Germany).
ITS amplification. PCR fragments for sequence analysis were obtained using primers ITS1
(TCCGTAGGTGAACCTGCGG) and ITS4 (TCCTCCGCTTATTGATATGC) covering
the internal transcribed spacer 1 (ITS1) region, 5.8S, and ITS2 region of the ribosomal
DNA. For each 50 µl reaction, a mixture was prepared containing 50 ng of genomic DNA,
0.1 vol. of 10× PCR buffer (10 mM Tris-HCl, pH 8.8, 50 mM KCl, 0.1% Triton X-100), 4
mM MgCl2, 1 mM of deoxynucleoside triphosphates (Roche, Mannheim, Germany), 0.4
µM of each primer, and 2 U of DyNAzyme I DNA Polymerase (Finnzymes, Espoo,
Finland). Reactions were run on a programmable thermal cycler Mastercycler epgradient
(Eppendorf AG) with an initial denaturation of 3 min at 96°C, followed by 30 cycles at
94°C for 30 s, 60°C for 30 s, and 72°C for 40 s, with a final extension at 72°C for 10 min.
Aliquots of the PCR were electrophoresed on 2% (w/v) agarose gels in 1× TAE buffer (40
mM Tris-acetate, 1 mM EDTA) pH 8, using a Sub-Cell GT apparatus (Bio-rad
Laboratories, Hercules, USA). Products were visualized using the UV transillumination G91
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Box (Syngene, Fredericks, USA) and photographed with a GeneSnap camera (Syngene).
DNA fragments were gel-purified and sequenced at the Institute of Biomedicine (CSIC,
Valencia, Spain). The sequences obtained were searched against the public databases using
the Basic Local Alignment Search Tool (BLAST).
PgChP purification
P. chrysogenum AS51D was grown in malt extract broth at 25ºC for 15 days. The medium
was filtered through a 0.45 µm diameter pore size cellulose acetate membrane and PgChP
was purified using an ÄKTA fast protein liquid chromatography system (Amersham
Bioscience AB, Uppsala, Sweden) according to Acosta et al. (2009a). For this, filtered
medium was loaded on a HiTrap SP HP 5 ml ion exchange column (Amersham Pharmacia
Biotech AB) equilibrated with 0.02 M sodium acetate buffer, pH 4.5. Bound proteins were
eluted with a gradient of NaCl to 1 M (0 to 25% in 15 column volumes (CV), increasing to
100% in 2 CV, followed by 15 CV at 100%) in 0.02 M sodium acetate buffer. Eluted
proteins were detected at 214 nm. The fraction that showed antifungal activity in previous
studies (Acosta et al., 2009a) was purified using a HiLoad 26/60 Superdex 75 prep grade
gel filtration column (Amersham Pharmacia Biotech AB) equilibrated with 50 mM sodium
phosphate, 0.15 M NaCl buffer pH 7, at a constant flow rate of 2.5 ml/min. Data collection
and processing were done using the UNICORN software 4.12 (Amersham Pharmacia
Biotech AB). Finally, the fraction containing PgChP was desalted and concentrated with
YM-10 Microcon Centrifugal Filter Units (Millipore Iberica, Madrid, Spain) and stored at 20ºC.
Sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE)
SDS-PAGE was performed as described Laemmli (1970). The protein PgChP was
separated on a 4% (v/v) polyacrylamide stacking and 8% (v/v) polyacrylamide resolving
gel. Gels were stained with Imperial Protein Stain (Pierce Biotechnology, Rockford, USA).
Two preparations of molecular weight markers were used: Precision Plus Protein
Standards with ten proteins between 10 and 250 kDa (Bio-Rad) and the Low Molecular
Weight Electrophoresis calibration kit with proteins between 14.4 and 97 kDa (Pharmacia,
Uppsala, Sweden).
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Reduction and alkylation of PgChP
Reduction and alkylation of the protein prior to deglycosylation or enzymatic digestion
were performed with the protein in SDS-PAGE gel pieces or in solution. After SDSPAGE, PgChP (25 µg per well) was stained using Imperial Protein Stain. The band was
excised from the gel and washed three times by incubation in 100 µl of de-stain solution
(50 mM ammonium bicarbonate, 50% acetonitrile) for 15 min at 40ºC with intermittent
vortexing. Gel pieces were incubated at 56ºC in 100 µl of reducing solution (10 mM
dithiothreitol in 50 mM ammonium bicarbonate) which was discarded after 45 min and
replaced with 100 µl of alkylation solution (55 mM iodoacetamide in 50 mM ammonium
bicarbonate) and incubation continued at room temperature in the dark for 45 min. The gel
pieces were then washed in 100 µl of 50 mM ammonium bicarbonate for 5 min at 40ºC
and the supernatant was discarded. The gel was washed twice in 100 µl of 50 mM
ammonium bicarbonate in 50% acetonitrile for 1 min and centrifuged. Sample was
dehydrated in 100 µl of acetonitrile and incubated for 1-2 min. Finally, the supernatant was
discarded and the gel pieces were dried at 40ºC for a few min. After addition of the
digestive enzyme, the tube containing the gel pieces was cooled at 4°C for 20 min and
finally incubated at the optimal temperature of each enzyme.
For digestion of PgChP in-solution, different quantities of the protein (1.25, 2.5, and 6.25
µg) were added into tubes containing 50mM ammonium bicarbonate up to a final volume
of 5 µl. Dithiothreitol 5 mM dissolved in 50 mM ammonium bicarbonate was added (5 µl)
and the final solution was incubated at 50ºC for 30 min. After adding 10 µl of 20 mM
iodoacetamide dissolved in 50 mM ammonium bicarbonate, sample was incubated at room
temperature for 30 min in the dark before digestion with each enzyme.
Digestion with enzymes for amino acid sequencing
Trypsin digestion. Digestion with the reduced and alkylated protein was performed with
PgChP in-gel and in-solution. Gel pieces containing PgChP were incubated with 10 ng/µl
trypsin gold (Promega, Hampshire, United Kingdom) in 50 mM ammonium bicarbonate at
40 or 60ºC for 1.5, 3, or 16 h. Additionally, further automated in-gel digestions were
carried out. For these, samples of PgChP in-gel pieces were de-stained, dehydrated,
reduced, alkylated, washed, and digested using the ProteomeWorks MassPREP robotic
liquid handling station (Waters, Manchester, UK), essentially as described above. For the
digestion step, the microtiter plate containing the gel pieces was cooled to 6°C for 10 min
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before addition of 25 µl per well of trypsin gold at 10 ng/µl. The plate was incubated at
6°C for 20 min and incubated at 37°C for 4 h. PgChP was digested in-solution by
incubating at 37ºC for 16 h after adding 50 mM ammonium bicarbonate, containing 5%
acetonitrile and 10 ng/µl of trypsin gold. All digested protein samples were stored at 4°C
until ESI-MS analysis.
Chymotrypsin digestion. Digestion with the reduced and alkylated protein was performed
with PgChP in-gel and in-solution. Gel pieces were digested at different concentrations (1,
5, 10, and 15 ng/µl) of chymotrypsin (Roche, Welwyn Garden City, UK) in buffer (100
mM Tris-HCl, 10 mM CaCl2 buffer pH 7.8), at 25ºC and 30ºC for 16 h. PgChP in-solution
was digested by chymotrypsin at 0.1, 1, 5, or 10 ng/µl with 5% (v/v) acetonitrile and the
same buffer mentioned above. Temperatures and incubation time were identical to those
used for in-gel sample digestions.
Analysis for glucidic residues
Chemical deglycosylation was carried out with PgChP in solution and using two different
chemical methods: acid hydrolysis with HCl (Leis et al., 1997; Krapf et al., 2006) and elimination with NaOH (Geert and Thomas-Oates, 1998; Tarelli, 2007). For acid
hydrolysis with HCl, 1 µg of PgChP was incubated in 20 mM HCl at 50ºC for 3 h. elimination with NaOH was performed by incubating 1µg of PgChP in 0.2 M NaOH at
45ºC for 16 h. Finally, both samples were analyzed by ESI-MS.
Enzymatic deglycosylation
PNGase F (New England Biolabs, Ipswich, UK) was used following the manufacturer’s
instructions and samples were analyzed by SDS-PAGE.
E-DEGLY deglycosylation kit (Sigma, Madrid, Spain) contained the following
deglycosylases: PNGase F, -2(3,6,8,9) neuraminidase, O-glycosidase, (1-4)-galactosidase,
and −N-acetylglucosaminidase. Reactions were carried out under native and denaturing
conditions according to manufacturer’s instructions and products were subjected to SDSPAGE analysis.
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Electrospray ionization mass spectrometry (ESI-MS) analysis for peptide sequence
determination
After deglycosylation and enzymatic digestion, formic acid was added to a final
concentration of 0.1% (v/v); then the sample was desalted using Zip-Tip C18 reverse phase
pipette tips (Millipore) according to manufacturer’s instructions. Peptides were eluted with
an equal volume of 50% (v/v) acetonitrile and 0.1% (v/v) formic acid. Samples were
loaded into individual borosilicate nanospray needles (Waters) using 1-10 µl GELoader
pipette tips (Eppendorf) and analyzed in a Q-TOF2 mass spectrometer (Waters). ESI-MS
was carried out with a capillary voltage of 900–1200 V in positive ion mode, using argon
as the collision gas. Survey scans of peptide mass spectra were performed with the
sampling cone set at 45–50 V and data were acquired from 400–1500 m/z with a cycle
time of 2.4 seconds. Peptide mass spectra presented resulted from combining typically 5 –
15 min of data acquisition. Individual peptide ions were selected manually for tandem MS
analysis. Collision voltages were optimised manually for individual peptides. Tandem MS
fragmentation spectra were collected typically from 50 to 1600 m/z. Tandem MS spectra
were deconvoluted into singly charged, mono-isotopic masses using the MaxENT3
maximum entropy software and peptide sequences were determined by manual
interpretation using the PepSeq software within the MassLynx V 4.0 package (Waters,
UK). De novo peptide sequences were searched against the public databases using
BLASTP with parameter settings for “short and nearly exact matches". A multiple
sequence alignment of fungal chitosanases was produced using the program T-Coffee at
www.ebi.ac.uk.
Molecular weight determination by ESI-MS
The molecular weight of PgChP was determined by electrospray mass spectrometry in a QTOF2 mass spectrometer (Waters, UK). PgChP (0.125 µg/µl) in 1% (v/v) formic acid was
desalted using C18 reverse phase pipette Zip-Tips (Millipore) according to the
manufacturer’s instructions. Protein was eluted into 50% (v/v) methanol, 1% (v/v) formic
acid, and loaded into nanospray needles for analysis. The mass spectrometer was operated
with a capillary voltage of 900-1200 V and the sampling cone at 45-50 V. Data were
acquired between 400 and 3000 m/z with a cycle time of 1 second. Mass spectra resulted
from combining typically 5 min of data. Spectra were deconvoluted using the MaxEnt1
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maximum entropy software with an output range of 20,000 and 40,000 Da, at a resolution
of 1 Da and a peak width of 0.75 Da.
Isoelectric point estimation by 2D-PAGE
Isoelectric focusing was performed on 7cm IPG ReadyStrips pH 3–10NL (Bio-Rad).
Concentrated protein (20 µg) was dissolved in a sample buffer containing 7 M urea, 2 M
thiourea, 4% (w/v) CHAPS, 0.005% (w/v) bromophenol blue, 0.5% (v/v) destreak reagent
(GE Healthcare, Uppsala, Sweden) and 0.5% ampholyte and incubated at room
temperature for 60 min. The first-dimension separation was performed on a Protean IEF
Cell System (Bio-Rad). The IPG strip was loaded using active in-gel rehydration.
Electrofocusing was performed as follows: 250 V for 15 min, then at a linear gradient from
250-4,000 V for 2 h, followed by 4,000 V until a total of 18,000 V/hr was reached. The
voltage decreased to a holding voltage of 500 V. After IEF, the strip was equilibrated with
a sample equilibration buffer (SEB) containing 6 M urea, 375 mM Tris-HCl (pH 8.8), 2%
(w/v) Sodium Dodecyl Sulphate (SDS), 30% (v/v) glycerol, and 0.0125% (w/v)
bromophenol blue. In the first incubation, SEB contained 1% (w/v) dithiothreitol, after 8
min this solution was replaced with SEB containing 25% (w/v) iodoacetamide and the strip
was incubated for a further 8 min. The IEF strip was placed onto 8% (v/v) polyacrylamide
gels and sealed in with agarose. SDS-PAGE was then carried out at 125 V for 50 min.
Proteins were visualized by staining with Imperial Protein Stain (Pierce Biotechnology)
and gel images were scanned on a GS800 scanner (BioRad). Precision Plus Protein
Standards (Bio-Rad) were used as molecular weight markers. A separate 2D-gel was
performed with 2-D SDS-PAGE Standards (15 µl) (Bio-Rad) to provide calibrated pI
points to allow estimation of the pI of PgChP.
RESULTS
Genetic characterisation of P. chrysogenum AS51D
Amplification of the 18S-28S rRNA intergenic spacer (ITS) sequence revealed a band of
585 bp, which was sequenced and compared against other sequences in BLAST. The strain
AS51D displayed 100% of identity and the maximum score with the 18S-28S rRNA
intergenic spacer of Penicillium chrysogenum strain FRR 807, accession number in
GenBank: AY373839. Thus, AS51D was confirmed as P. chrysogenum.
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Analysis of PgChP protein by SDS-PAGE
SDS-PAGE of purified PgChP to estimate the molecular mass was revealed two discrete
bands (Figure 1): an intensely stained band showing a molecular mass of 37 kDa and a
fainter one of around 40 kDa.
Figure 1. SDS-PAGE of PgChP protein, showing two bands of 40 kDa (1) and 37 kDa (2). M:
Pharmacia low molecular weight standard.
Mass spectrometric analysis of PgChP digestions
Figure 2 shows typical survey spectrum peptide profiles obtained by mass spectometry for
the 37 kDa protein band digested with trypsin and chymotrypsin. The 40 kDa band was
also analysed in a similar manner. Overall, the peptide mass spectrum profiles obtained for
the upper and lower bands showed a high degree of similarity (Figure 3).
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Figure 2. Mass spectra showing typical peptide profiles obtained from trypsin (A) and
chymotrypsin (B) digestions of PgChP (37 kDa band).
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Figure 3. Mass spectra showing the highly similar peptide profiles obtained from trypsin
digestion of the 37 kDa (A) and 40 kDa (B) bands from SDS-PAGE of PgChP.
De novo amino acid sequences
Tandem MS was also performed on peptides from PgChP trypsin and chymotrypsin
digestions to obtain amino acid sequences. All of the 40 kDa band peptide sequences
obtained were also present in the 37 kDa band. Together with the mass spectrum profiles,
the data indicated that the two bands observed on SDS-PAGE analysis were likely to
represent the same protein or two very similar proteins. The 37 kDa band was subjected to
extensive tandem MS analysis to provide increased amino acid sequence coverage of the
protein.
A total of twelve de novo peptide sequences were obtained from trypsin and chymotrypsin
digestions from the 37 kDa band and four from the 40 kDa band (Table 1).
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Table 1. PgChP-de novo peptide sequences obtained from trypsin and chymotrypsin digestions
from 37 kDa and 40 kDa. L and I are interchangeable. Italic letters indicate amino acids which
were tentative assignments.
Sequence
Number
Peptide
de novo sequence
mass
Charge
Digestive
enzyme
1
NKPDGGPPGSYF
618.29
+2
Chymotrypsin
2a
-FAAGSSLPVAALQSAAAK
959.81
+3
Trypsin
2b*
YFAAGSSLPVAALQSAAAK
911.48
+2
Trypsin
2c*
PVAALQSAAAK
1026.61
+1
Trypsin
3
-PDATYPLDGDNGA-
866.40
+2
Trypsin
4
-TIHSDWAK
528.79
+2
Trypsin
5
-WLADMDVDCDGLD-
1139.03
+2
Trypsin
6
GNPDGQHQTNFGAL-
728.33
+2
Trypsin
7
-AAYEVPFFVIPDR
988.21
+3
Trypsin
8a
GALPGNNVGAVICDGK
771.40
+2
Trypsin
8b
GALPGNNVGAVICDGK
720.89
+2
Chymotrypsin
9a
-PQVLGEASWLMAR
1053.50
+2
Trypsin
9b*
MFYGIYGDSDGDTPQVLGEASRFFAR
972.43
+3
Trypsin
10a
FTGDDSVLPSSALNK
775.87
+2
Trypsin
10b*
-TYILFTGDDSVLPSSALNK
1352.95
+3
Trypsin
11
NYVTNFTTLR
614.82
+2
Trypsin
12
RSMGDKLM
469.22
+2
Chymotrypsin
*: Indicates sequences from the 40 kDa band.
-: indicates the presence of additional unassigned amino acids.
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Analysis for glucidic residues
After enzymatic deglycosylation of PgChP with the PNGase F or the E-DEGLY
deglycosylation kit, only the band of 37 kDa was detected (Figure 4). However, chemical
deglycosylation with HCl and NaOH revealed no changes in spectra with untreated and
treated PgChP by ESI-MS.
gChP
Figure 4. SDS-PAGE of purified PgChP untreated (lane 1), and treated with PNGase F (lane
2). The arrow indicates the 37 kDa band which remains after deglycosylation. M: Precision
Plus Protein Standard.
Molecular mass determination of PgChP protein by electrospray ionization mass
spectrometry
ESI-MS was used to establish the accurate mass of intact PgChP. The m/z spectrum was
deconvoluted and converted to the molecular mass profile using Maximum Entropy
processing (Figure 5A). The mass profile shows a dominant peak of 30,771.9 Da with a
series of surrounding peaks differing from each other by approximately 162 Da (Figure
5B).
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Figure 5. Molecular mass profile from Maximum Entropy processing of PgChP. The most
abundant peak has a mass of 30771.9 Da (A). Detailed view revealing a series of peaks
separated by approximately 162 Da (B). Additional peaks surrounding the major peaks
represent sodium adducted forms (+/- 22 Da).
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Isoelectric point estimation
2D-PAGE analysis showed that PgChP migrates on a broad pH range gel in a discrete
region of the gel at approximately pH 5 (Figure 6).
A)
B)
Figure 6. Isoelectric point (pI) determination of PgChP by 2D SDS-PAGE. A: PgChP is
indicated with an arrow. B: 2D SDS-PAGE standards. Numbers within figure B indicate the pIs
of 4 standard proteins that migrate as a series of spots.
DISCUSSION
In previous studies the strain AS51D was characterized as P. chrysogenum using methods
based on morphology and biochemical characteristics, including the production of
secondary metabolites by high performance liquid chromatography (Acosta et al., 2009a).
In the present study, the sequence of the 18S-28S rRNA intergenic spacer confirmed the
previous characterization. P. chrysogenum is regarded as a safe mould of starter cultures
(Núñez et al., 1996; Benito et al., 2003) together with other close species, such as
Penicillium nalgiovense (Geisen, 2000). To fulfill this role, AS51D could be used either as
a source for antifungal proteins or as protective culture.
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Two bands were found in the concentrated PgChP fraction by SDS-PAGE analysis with
estimated molecular weights of 37 and 40 kDa. The comparison between the amino acid
sequences obtained from both bands showed a high degree of similarity (Table 1). All
sequences obtained from the 40 kDa band were derived from three different regions of the
37 kDa protein: sequences 2a, 2b, and 2c in one region; 9a and 9b in a second; and 10a and
10b in a third region. Where they overlapped, peptide sequences from the 40 kDa band
were identical to those from the 37 kDa band, with the only exception being for peptide 9b
for which 3 of the 5 C-terminal residues could be only tentatively assigned from the
tandem MS spectrum. The higher number of sequences obtained from the 37 kDa band
result from the more in-depth analysis carried out with this protein, including digestion
with chymotrypsin, and in-gel and in-solution digestions.
After exposing the PgChP protein to N-Glycosidase F (PNGase F) only the protein of 37
kDa was observed (Figure 4). This deglycosylation enzyme cleaves between the innermost
GlcNAc and asparagine residues of high mannose hybrid and complex oligosaccharides
from N-linked glycoproteins (Maley et al., 1989; Plummer and Tarentino, 1991). Thus, the
protein of 40 kDa is glycosylated, and its peptidic core migrates at the same molecular
mass as the 37 kDa protein. Chemical deglycosylation with HCl and NaOH produced no
changes in mass spectra when treated and untreated PgChP were compared by ESI-MS.
Thus, neither sialic acid residues nor O-linked oligosaccharides are present (Leis et al.,
1997; Geert and Thomas-Oates, 1998; Krapf et al., 2006; Tarelli, 2007).
The analysis of the molecular mass of the protein by mass spectrometry revealed the
existence of six major peaks centring a dominant peak of around 30.8 kDa, differentiated
by a mass multiple of 162 Da, the mass of a galactose or mannose residue. It is therefore
possible that the purified fraction of PgChP has multiple forms of the protein each with a
different number of mannose or galactose residues. The glycosylation of the protein likely
explains the difference in the mass estimated by SDS-PAGE and ESI-MS. It is known that
glycoproteins can migrate atypically by SDS-PAGE because SDS does not bind
oligosaccharides (Nelson, 1971; Russ and Poláková, 1973; Bagger et al., 2007). Given that
the ESI-MS provides an exact molecular mass to within 0.01%, it can be deduced that the
protein has an atypical migration by SDS-PAGE.
Glycosylated proteins are not uncommon among antifungal enzymes, such as an antifungal
glycoprotein from Urginea indica (Deepak et al., 2003) or chitosanases from Mycobacter
AL-1 and P. islandicum (Hedges and Wolfe, 1974; Fenton and Eveleigh, 1981).
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Glycosylation represents the most common form of protein post-translational
modifications. Carbohydrates can be found in both eukaryotes and prokaryotes and they
participate in many biological processes including inter- and extra-cellular signalling,
protein regulations and interactions or molecular recognition. Glycan chains are added to
proteins as a set of variations on a core structure, and can vastly increase the complexity of
protein molecules. Glycans can determine the localization, activity and function of proteins
and can influence physicochemical characteristics of proteins such as folding, solubility,
resistance to degradation by proteases or increase in thermal stability (Hildegard and
Geyer, 2006; Morelle et al., 2006).
In N-glycosylation, glycans are attached to the protein via N-acetylglucosamine (GlcNAc)
to the amide group of asparagine within an Asn-X-Ser/Thr (sometimes also Cys) motif,
where X is any amino acid apart from proline (Morelle et al., 2006). Although the
complete amino acid sequence of PgChP is unknown, at least one consensus site for Nglycosylation is present in the protein (Table 1, peptide 11 -NFT-). This potential site for
glycosylation in PgChP supports the interpretation that the 37 kDa band represents the
deglycosylated core of the 40 kDa band.
The de novo amino acid sequences obtained from PgChP (Table 1) showed the highest
similarity with sequences of seven described or predicted fungal chitosanases from A.
fumigatus (accession number: XP_754126), N. fischeri (XP_001262949), A. niger
(XP_001390407), A. oryzae-1 (BAD08218), A. oryzae-2 (XP_001824257), A. terreus
(XP_001209034), and A. clavatus (XP_001274664). The twelve peptide sequences from
PgChP were placed onto a multiple sequence alignment of these protein sequences (Figure
7).
Three highly conserved amino acid sequences have been described in all of the fungal
chitosanases. These three regions are located in the vicinity of the centre of the open
reading frames, and are considered to be important for catalytic function (Shimosaka et al.,
2005). In each of these three regions there is one conserved carboxylic amino acid (Glu or
Asp) considered to be essential or important for the catalytic function as a proton donor in
a large number of glycosyl hydrolases (Monzingo et al., 1996; Robertus et al., 1998). At
least one of these conserved regions corresponds with the de novo peptide sequence 9 of
PgChP, which has the conserved glutamic acid (underlined): PQVLGEASWLMAR.
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PgChP
A.fumigatus
N.fischeri
A.niger
A.oryzae-1
A.oryzae-2
A.terreus
A.clavatus
* ***
* *
***** * *
NKPDGGPPGSYFAAGSSLPVAALQSAAAK
MPSTTIIRQLAISLALCN-SALGQVVNGADYNKPNGGPPASFFAAASTMPVAALQAAAAK
MPSKTIIRQLAISVALCN-SALGQVVNGADYNKPNGGPPASFFAAASTMPVAALQAAAAK
MAFKTTAG--LAFLALAG-SVKAQSVDGSKYNSPSNGPPASYFAAATTLPVAALQSAAAK
MPIKSFASRLALSLAICG-TAMGQKVNGADYNKPDGGPPAKFFQASSSIPVAAIQAAAAK
MPIKSFASRLALSLAICG-TAMGQKVNGADYNKPDGGPPAKFFQASSSIPVAAIQAAAAK
MVFKKAAIGLTLPFALFSSVALGQTVDGSDYDSPNGGPPGSYFAAASTMPVAALQAAAAK
MIFKSSLSHAAASLLLLT-PALAQKVQGPEYNKPSAGPPASFFAAAPTMPVAALKSAVAR
PgChP
A.fumigatus
N.fischeri
A.niger
A.oryzae-1
A.oryzae-2
A.terreus
A.clavatus
* **
*** **
**********
PDATYPLDGDNGA
TIHSDWAK
WLADMDVDCDGLD
ATKVPSLATYPVSQDKGAAKSTIHTDWASFSEGASISWVADMDVDCDGLNSGC------ASKVPSLATYPVSQDSGAAKSTIHTDWASFSEGASISWVADMDVDCDGLNSGC------ASSVPSKATYPVNTDDDSPKSTIHSDWVKFNQGAALSWVADMDVDCDGIDYKC------ASKVPSHATYPIGQ--GSTKSTIHSDWAGFSEGAAFSFIADMDVDCDGLNHGC------ASKVPSHATYPIGQ--GSTKSTIHSDWAGFSEGAAFSFIADMDVDCDGLNHGC------ATKVPSYATYPVSQDDNAKKSTIHSDWASFSQGAAISWVADMDVDCDGIDSGCEHEIKTS
ASVVPKNAAYPVNQD-GGPTATIHADWASLPTAAAYVYTADMDVDCDGLDHNC-------
PgChP
A.fumigatus
N.fischeri
A.niger
A.oryzae-1
A.oryzae-2
A.terreus
A.clavatus
** **
** *** ***** ***
* **
****
**
GNPDGQHQTNFGALAAYEVPFFVIPDR
AGALPGNNVGAVIC----DGK
QGNPDGQPQTNWGALSAYEVPFIVIPDKYLSANSGALPGNNIAAVIC----NGKMFYGIL
QGNPDGQPQTNWGALSAYEVPFIVIPDKYLSANTGALPGNNIAAVIC----NGKMFYGIL
KGNGDGLPETNWGALSAYEVPWIVIPDQFLTANEDLLPGNNVAAVIC----NGKMYYGIL
KGNPDGQKETNWGALSAYEVPFIVIPQEFLDANKGTLKGNAVAAVIC----NGKMFYGIF
KGNPDGQKETNWGALSAYEVPFIVIPQEFLDANKGTLKGNAVAAVICATSSNGKMFYGIF
QGNPDGQDATNWGALAAYEVPFIVIPQKYLDHNGNALKGNNIAAVIC----NGKMFYGIL
KGNPDGQPDTNFGALAAYEVPFVVIPDRFATTYASALPGNNIVAVIC----DGKMFYGIF
PgChP
A.fumigatus
N.fischeri
A.niger
A.oryzae-1
A.oryzae-2
A.terreus
A.clavatus
* * *********
*
**
**
PQVLGEASWLMAR
LLFTGDDSVLPSSALNKNY
GDSNGDSPQVTGEASWLMARTCFPNEGLNGNNGHTGVDVTYIVFTGKNAVLPSSALTKNY
GDSNGDSPQVTGEASWLMARTCFPNEGLNGNNGHTGVDVTYIVFTGKDAVLPSSALTKNY
GDSNGDDPEVTGEASWLMARTCFPDDDLNGAEGHAEADVTCKPYT-------RSALNKNY
GDSNGDSPQVTGEASWLMARTCFPKEDLNGNKGHTAADVTYIVFTGDKAVLPSSALNKNY
GDSNGDSPQVTGEASWLMARTCFPEEDLNGNKGHTAADVTYIVFTGDKAVLPSSALNKNY
GDANGDEPQVTGEASWLMARTCFPNEGLNGNKGHTAADVTYILFTGDESVLPSSALNENY
GDTDGDHPQVIGEASWLMARTCFPNDNLNGDSGHVPADVTYIFFTGKDSVLPSSAVNKNY
PgChP
A.fumigatus
N.fischeri
A.niger
A.oryzae-1
A.oryzae-2
A.terreus
A.clavatus
* * ****** *
VTNFTTLRSMGDKLM
ITNFTTLRSMGDKLVNALVSNLGLSGTPPTK---------TTTRVTTTTTKPT-SAASCS
ITNFTTLRSMGDKLVNALASKLGLSGTPPTK---------TTTLVTTTTTKPT-STASCS
ITNFSTLRSMGDKLVGALASNLGLTSSA------------------------SGSTATCS
ITNFDTLRSMGDSLVGALAKNLNLGGGGGNP---------PTTLTTTSIPEPTGGSGSCS
ITNFDTLRSMGDSLVGALAKNLNLGGGGGNP---------PTTLTTTSIPEPTGGSGSCS
ITNFSTLRSMGDKLVNALASNLGLSGGSGGT---------PTT--TTTATQPT-STGTCS
ITDFTKLRSMGDSLVNAFASQLGISSGGGNGGGDGGHTTLHTTTATRTATAPT-STATCD
PgChP
A.fumigatus
N.fischeri
A.niger
A.oryzae-1
A.oryzae-2
A.terreus
A.clavatus
WAGHCL-----GASCSSDDDCADALVCTAGKCSVDGAA-TCSWEGHCEGASCSSDDDCSD
WAGHCL-----GASCSSNDDCADALVCAAGKCSVDGAA-TCSWEGHCEG----------WQGHCE-----GAICSTEDDCSDDLVCDSGKCSSPDED---------------------WPGHCA-----GATCSSNDDCSDDLTCQNGKCASDGSAETCSWEGHCKGATCSSNDDCSD
WPGHCAGFKNKGATCSSNDDCSDDLACQNGKCASDGSAETCSWEGHCKGATCSSNDDCSD
WAGHCE-----GASCSTNDDCSDQLACKNGVCSTDGEV-VCSWEGHCEGATCSSNDDCSD
WEGHCL-----GTACSNNGECSDPFGCINGFCNYDPTL-TCQWAGHCVGASCSSNDQCSD
PgChP
A.fumigatus
N.fischeri
A.niger
A.oryzae-1
A.oryzae-2
A.terreus
A.clavatus
DLYCKSGSCTA------------------------------------P----------------------------VYS-------------------------------------DGDDEDDEDEEENDEDDEDDDDEDDEDDE-----ELACISGICSVDNGVETCEWEGHCEGASCSSHDDCDGNLACKNGKCSA--ELACISGICSVDNGVETCEWEGHCEGASCSSHDDCDGNLACKNGKCSA--ELYCAGGACTSA--------------------------------------PFACIDGACAVDT-SLDCSRKGHCAGTTCLSDSDCSRPLSCILGVCANQSG
Figure 7. Alignment of PgChP amino acid sequences with the most similar proteins in the public
sequence databases. Shaded-background letters show similarities of the proteins with the obtained
sequences of PgChP. Accession numbers in Genbank are XP_754126 (A. fumigatus),
XP_001262949 (N. fischeri), XP_001390407 (A. niger), BAD08218 (A. oryzae-1), XP_001824257
(A. oryzae-2), XP_001209034 (A. terreus), and XP_001274664 (A. clavatus). Asterisks indicate
amino acid residues completely conserved in all sequences. Numbered segments correspond to de
novo obtained peptide sequences for PgChP (see Table 1).
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A protein sequence homology analysis was performed calculating the percentages of
amino acid identity between the sequences obtained from PgChP protein shown in table 1
and the aligned sequences from the seven fungal chitosanases (Table 2).
Table 2. Similarity between the amino acid sequences obtained from PgChP and the aligned amino
acid sequences of the similar fungal chitosanases described in Figure 7. Percentages higher than
85% are background shaded.
Similarity (%)
PgChP
A. fumigatus
N. fischeri
A. niger
A. oryzae-1
A. oryzae-2
A. terreus
A. clavatus
1
75
75
67
75
75
75
67
2a
78
78
83
83
83
78
61
2b*
74
74
84
79
79
79
53
3
61
54
38
46
46
46
38
4
75
75
75
75
75
75
75
5
85
85
92
85
85
92
85
6
86
86
64
79
79
79
86
7
77
77
69
69
69
77
92
8a,b
76
76
71
65
65
65
82
9a
92
92
85
92
92
92
92
9b*
73
73
69
73
73
69
73
10a
72
78
67
83
83
83
78
10b*
76
81
43
86
86
86
81
11
90
90
80
80
80
80
70
12
87
87
87
75
75
87
75
Sequences
All peptide sequences from PgChP showed high similarity (from 38 to 92%) with the
fungal chitosanases (Table 2). In addition to sequence 9, already discussed as a conserved
region in fungal chitosanases, sequence 5 had a similarity higher than 85% in all the
compared proteins.
The high similarity with fungal chitosanases and the conserved region identified in
sequence 9, confirm that PgChP belong to this kind of proteins. Chitosanases (E.C.
3.2.1.132) are the glycoside hydrolases that cleave the glycosidic bonds of chitosan. These
enzymes have been reported from viruses, bacteria and fungi (Wang et al., 2008).
Chitosanases are classified into five glycoside hydrolase families, GH-5, GH-8, GH-46,
GH-75, and GH-80, according to amino acid sequence homology. Fungal chitosanases
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belong to the GH-75 family (Cheng et al., 2005), which is composed mainly by
chitosanases from Aspergillus (Cheng and Li, 2000; Zhang et al., 2000) and Fusarium
(Shimosaka et al., 1996; Cantarel et al., 2008). Most chitosanases are characterized by a
low molecular mass in the range of 10-50 kDa (Fenton and Eveleigh, 1981; Somashekar
and Joseph, 1992; Alfonso et al., 1992).
The isoelectric point (pI) estimated from 2D-PAGE and isoelectrofocusing was close to pH
5, which is within the range of isoelectric points, from pH 4 to 5, described for fungal
chitosanases (Fenton and Eveleigh, 1981; Alfonso et al., 1992; Shimosaka et al., 1993).
The range of predicted isoelectric points (pH 3.83-4.97) and molecular masses (33-40 kDa)
of the selected fungal chitosanases are quite similar to the experimental values obtained for
PgChP (Table 3).
Table 3. Isoelectric points (pI) and molecular masses of PgChP and selected fungal chitosanases.
Isoelectric Point
Molecular Mass (kDa)
Protein/producer mould
2D-PAGE/IEF
Predicted*
ESI-MS
Theoretical*
PgChP/ P. chrysogenum
5
-
30-31
-
XP_001274664/ A. fumigatus
-
4.64
-
35.1
XP_001262949/ N. fischeri
-
4.97
-
33.1
XP_001390407/ A. niger
-
3.83
-
33.4
BAD08218/ A. oryzae-1
-
4.69
-
38.9
XP_001824257/ A. oryzae-2
-
4.71
-
39.8
XP_001209034/ A. terreus
-
4.25
-
36.0
XP_001274664/ A. clavatus
-
4.64
-
40.1
* Predicted pI values and theoretical molecular masses were calculated using the ProtParam tool in
the ExPASy proteomics server at the Swiss Institute of Bioinformatics (http://www.expasy.org).
Recently, the genome of P. chrysogenum Wisconsin 54-1255 has been sequenced (Van den
Berg et al., 2008). A putative protein can be found in BLAST (accession number
CAP80409) that shows high homology with the amino acid sequences obtained from
PgChP in this work. However, the production of the protein by this mould has not been
reported.
Currently, only two chitosanases have been reported from the genus Penicillium, one from
P. spinulosum and another from P. islandicum (Fenton et al., 1981; Ak et al., 1998), but
none from P. chrysogenum. Given that P. chrysogenum is used to obtain generally
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recognized as safe (GRAS) enzymes, PgChP could be used as a food preservative against
toxigenic moulds. However, before using PgChP for food protection, an in-depth
toxicological evaluation of the antifungal protein is a prerequisite for its application. Thus,
a full characterization should be done for this novel chitosanase.
ACKNOWLEDGEMENTS
This work was supported by the Spanish Ministry of Education and Science (AGL200406546-ALI) and FEDER. Andrea Rodríguez was the recipient of a FPI grant from the
Spanish Ministry of Education and Science. Thanks to Alex Rathbone, University of
Nottingham, for tuition in 2D gels.
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Manuscrito 5: Genetic characterization of
the gene enconding the novel chitosanase PgChP
from Penicillium chrysogenum
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Genetic characterization of the gene encoding the novel chitosanase
PgChP from Penicillium chrysogenum
Andrea Rodríguez-Martín, Raquel Acosta, Félix Núñez, Mª José Benitoa, Miguel A.
Asensio*
Higiene y Seguridad Alimentaria, Facultad de Veterinaria. Universidad de Extremadura, Avda. de la
Universidad, s/n. 10071- Cáceres, Spain.
a
Nutrición y Bromatología, Escuela de Ingenierías Agrarias, Universidad de Extremadura, Carretera de
Cáceres s/n, 06071- Badajoz, Spain
* Corresponding author: Telephone (+34) 927 257 125; fax: (+34) 927 257 110.
E-mail: masensio@unex.es
ABSTRACT
The protein PgChP is a new chitosanase with antifungal activity produced by Penicillium
chrysogenum AS51D. The gene encoding the chitosanase PgChP has been sequenced. A
DNA fragment of the gene encoding pgchp was obtained by PCR using degenerate primers
based on two known amino acid sequences of the purified PgChP. cDNA was prepared for
RACE-PCR from total RNA of P. chrysogenum by RT-PCR. Full-length cDNA and
complete genomic DNA of pgchp were obtained using primers designed from 5’ and 3’ –
ends. Comparing the cDNA and genomic DNA, four introns of 60, 53, 59, and 59 bp were
observed. Full-length cDNA has 915 bp, corresponding to an open reading frame encoding
a protein of 304 amino acid residues. The first 21 amino acids matched a signal peptide.
The predicted mass of the translated nucleotidic sequence was 32 kDa and the predicted
isoelectric point was 4.8. The translated sequence of pgchp revealed that the protein had
two carboxylic amino acids (Glu and Asp) described as essential for the catalytic function
of fungal chitosanases. Two N-glycosylation consensus sequences were observed in the
translated pgchp gene. The putative amino acid sequence showed high homology with
fungal chitosanases.
KEY WORDS: antifungal activity, glycosylation, N-linked oligosaccharides, dryfermented foods.
INTRODUCTION
P. chrysogenum AS51D was isolated from meat products in previous studies (Acosta et al.,
2009a). This mould produces the protein PgChP, selected for its ability of inhibiting the
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growth of some toxigenic moulds. In previous works, different de novo amino acid
sequences obtained from PgChP showed significant similarities with the respective
sequences of seven fungal chitosanases, belonging to the GH-75 family of fungal
chitosanases (Rodríguez-Martín et al., 2009). Chitosanases (E.C. 3.2.1.132) are produced
by a large number of microorganisms including bacteria and fungi. Numerous bacterial
chitosanases are able to degrade exogenously added chitosan (Shimosaka et al., 2000;
Yoon et al., 2002; Lee et al., 2006; Zhang et al., 2007). On the contrary, fungal
chitosanases have been reported from only a limited number of strains and their
physiological roles are unclear (Fenton and Eveleigh, 1981; Zhang et al., 2000). The
synthesis of some chitosanases can be induced by adding different substrates such as
glucosamine (Zhang et al., 2001) and chitosan (Zhu et al., 2007).
Analysis of the molecular weight of PgChP by ESI-MS revealed six isoforms differing
from each other by approximately 162 Da, the mass of one residue of mannose. These
evidences indicated that the chitosanase PgChP was a glycoprotein with different degrees
of glycosylation. Various deglycosylation treatments revealed the presence of N-linked
oligosaccharides in PgChP (Rodríguez-Martín et al., 2009).
The aim of this paper was to study the genetic sequence encoding the chitosanase PgChP.
This sequence will allow studying the existence of introns, the complete putative amino
acid sequence, and glycosylation consensus sequences.
MATERIALS AND METHODS
Microbial strain
P. chrysogenum AS51D was isolated from dry-cured ham and belongs to the fungal
collection of Food Hygiene, University of Extremadura.
DNA isolation
P. chrysogenum AS51D was grown in malt extract broth made with 2% (w/v) glucose, 2%
(w/v) malt extract, and 0.1% (w/v) peptone, pH 4.5 at 25ºC under continuous shaking, for
5 days. The mycelium was obtained by filtering the culture through paper Whatman nº2.
Two grams of mycelium were broken with mortar and pestle after freezing by adding
liquid nitrogen. Fungal lysis was performed by homogenizing the obtained powder in 4 ml
0.05 M Tris-HCl, pH 8; 0.005 M ethylenediamine tetraacetic acid (EDTA); 0.05 M NaCl
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(TES) buffer and 1% (w/v) sodium dodecyl sulfate (SDS). Next, 200 µl proteinase K (20
µg/µl) were added, and the mix was incubated at 60ºC for 35 min, cooled on ice and
extracted with phenol-chloroform-isoamyl alcohol (25:24:1). The suspension was
centrifuged at 3000×g for 2 min at 4ºC. The upper phase containing DNA was transferred
to a fresh tube and precipitated by adding 3 M sodium acetate pH 5.2 to a final
concentration of 10% (w/v) and two volumes of cold ethanol. After centrifugation, the
pellet was cleaned with 70% (v/v) ethanol, centrifuged again, resuspended in sterile water,
and treated with 50 µl RNase (10 µg/µl) at 37ºC for 60 min. Finally, DNA was extracted
with phenol-chloroform-isoamyl alcohol, and precipitated again as indicated above,
resuspended in water, and stored at -70ºC. Quantity and quality of purified DNA was
determined spectrophotometrically in a Biophotometer (Eppendorf AG, Hamburg,
Germany).
Genomic DNA amplification
Polymerase chain reactions were performed with genomic DNA (100 ng) as template in 50
µl reaction mixtures containing 50 pmol each of forward and reverse degenerated primers,
0.5 mM each of deoxyribonucleotide triphosphates, (dNTPs, Roche, Madrid, Spain), 0.1
vol of 10× PCR buffer (22.5 mM MgCl2, 500 mM Tris-HCl pH 9.2, 140 mM ammonium
sulfate), 1.8 mM MgCl2 and 2.5 units of pfu turbo polymerase (Stratagene, La Jolla, USA).
The degenerated primers pgchp-DPF1 and pgchp-DPR2A (Table 1) were designed from
amino acid sequences previously obtained by mass spectrometry (Rodríguez-Martín et al.,
2009). Additionally, for pgchp-DPR2A the contiguous residues conserved in fungal
chitosanases (Shimosaka et al., 2005) were also included.
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Table 1. Degenerated primers used to amplify the pgchp gene.
Primer names
Amino acid sequences and designed primers*
N
pgchp-DPF1
P
D
G
G
P
5’-AAY AAR CCN GAY GGN GGN CC-3’
M
pgchp-DPR2A
K
A
R
T
C
F
P
3’-AC CGN GCN TGN ACR AAR GG-5’
* R for purine, Y for pirimidine, N for A, C, T, or G.
Conserved amino acids used for designing the primer pgchp-DPR2A are underlined.
DNA amplification was carried out in a Px2 thermocycler (Thermo Scientific,
Cramlington, UK). Three reactions were done, one with both forward and reverse primers,
and two with either forward or reverse primers. The PCR program consisted of initial
denaturation (94ºC for 5 min), 35 cycles of denaturation (94ºC for 1min), annealing (5660ºC range for 30 s, gradient 12), and extension (72ºC for 30 s), followed by a final
extension at 72ºC for 4 min. PCR products were electrophoresed on 2% (w/v) agarose gels
in 1× TAE buffer (40 mM Tris-acetate, 1 mM EDTA), pH 8 ussing a Sub-Cell GT
apparatus (Bio-rad Laboratories, Madrid, Spain). Two different markers were used for
agarose gels: 1kb DNA ladder (0.25-12 kb bands) (Promega, Hampshire, United Kingdom)
and DNA size standard (0.05-2kb) (Bio-Rad). Then, products detected only in the reactions
with both reverse and forward primers were gel-purified and cloned into the pCR2.1TOPO vector (Invitrogen, Barcelona, Spain). One Shot TOP10 Chemically Competent E.
coli cells (Invitrogen) were transformed with that vector. To confirm the insert in the
vector, digestions with restriction enzyme EcoRI (Roche) were carried out. Finally, vectors
were sequenced in the Institute of Biomedicine (CSIC, Valencia, Spain).
RNA isolation
P. chrysogenum AS51D was grown in malt extract broth, pH 4.5 at 25ºC under continuous
shaking for 4 days. Chitosanase production was induced in a medium with glucosamine
(Zhang et al., 2001) made with 0.3% (w/v) NaNO3, 0.05% (w/v) KCl, 0.05% (w/v)
MgSO4•7H2O, 0.01% (w/v) FeSO4•7H2O, and 2% (w/v) D(+)-glucosamine hydrochloride
(Sigma, Dorset, United Kingdom), dissolved in 25 mM phosphate buffer pH 5.8. Mould
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was cultured in glucosamine medium at 25ºC under continuous shaking for 17 h.
Subsequently, mycelium was quickly frozen with liquid nitrogen and stored at -80ºC until
RNA extraction. Frozen mycelium was broken with mortar and pestle after adding liquid
nitrogen. RNA extraction was performed with RNeasy Plant Mini Kit (Qiagen, Crawley,
UK) following the manufacturer’s instructions. Total extracted RNA was resuspended in
40 µl of RNase-free deionized water.
Rapid amplification of cDNA ends by Polymerase Chain Reaction (RACE-PCR)
Amplification of 5’ and 3’ ends of the pgchp gene was carried out using the SMART
RACE cDNA Amplification Kit (Clontech Laboratories, Saint Germain en Laye,
France).
cDNA synthesis by reverse transcriptase-polymerase chain reaction (RT-PCR). Two
separate populations of cDNA were synthesized according to the amplification kit
instructions: 5’-RACE-Ready cDNA and 3’-RACE-Ready cDNA. After incorporating the
SMART sequence into both the 5’- and 3’-RACE-Ready cDNA populations, RACE PCR
reactions were performed using the Universal Primer A Mix (UPM), in conjunction with
distinct gene-specific primers.
Gene-specific primers design. Two gene-specific primers (GSP) were designed for
amplifying the 5’ and 3’- ends (5’-RACE GSP: CCATTGCAGATAACGGCACCAACG
and 3’-RACE GSP: GTTCGACAAGGGTGCGGCATATG). Polymerase chain reactions
were performed with 5’- or 3’-RACE-Ready cDNAs (50 ng) as template in 50 µl reaction
mixtures containing 1 mM of dNTPs (Roche), 0.5 µM of the 5’- or 3’-RACE GSP primers,
5 µl of Universal Primer A Mix (10×), 0.1 vol of 10× PCR buffer (10 mM Tris-HCl, pH
8.8, 50 mM KCl, 0.1% Triton X-100), 4 mM MgCl2, and 2.5 units pfu turbo polymerase
(Stratagene). A program for touchdown PCR was carried out for 5’-end amplification: 5
cycles of 2 steps (94ºC for 30 s and 72ºC for 3 min), 5 cycles of 3 steps (94ºC for 30 s,
70ºC for 30 s and 72ºC for 3 min) and 27 cycles of 3 steps (94ºC for 30 s, 68ºC for 30 s
and 72ºC for 3 min). The PCR program to obtain the 3’-end consisted of 30 cycles of
denaturation (94ºC for 30 s), annealing (68ºC for 30 s), and extension (72ºC for 3 min).
Finally, 5’- and 3’-RACE-PCR products were gel-purified and cloned into pCR2.1-TOPO
vectors, which were used to transform One Shot TOP10 Chemically Competent E. coli
(Invitrogen). Then plasmids were purified and sequenced. Digestions with restriction
enzyme EcoRI (Roche) were performed to confirm the insert in the vector.
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Full-length cDNA amplification. The full cDNA fragment of the gene pgchp was obtained
by PCR with specific primers Fwd-pgchp (ATGATGACCTACAGCCGCTTAATCCC)
and Rev-pgchp (TCAATCACTGGGGCATTTTCCAC) designed from 5’ and 3’ ends.
PCR reaction mixtures at a final volume of 50 µl contained 5’-RACE-Ready cDNA (50
ng), 0.5 µM each of forward and reverse primers, 0.5 mM each of the dNTPs, 0.1 vol of
10× PCR buffer (10 mM Tris-HCl, pH 8.8, 50 mM KCl, 0.1% Triton X-100), 4 mM
MgCl2 and 2.5 units pfu turbo polymerase (Stratagene). The PCR program consisted of 30
cycles of denaturation (94ºC for 30 s), annealing (65ºC for 30 s), and extension (72ºC for 3
min). PCR products were gel-purified and cloned into pCR2.1-TOPO vectors. These
plasmids were used to transform One Shot TOP10 Chemically Competent E. coli cells
(Invitrogen). Then vectors were purified and sequenced. Digestions with restriction
enzyme EcoRI (Roche) were performed to confirm the wanted insert in the vector.
Amplification of the whole genomic gene
The genomic gene sequence was amplified by PCR using Fwd-pgchp and Rev-pgchp
primers. PCR reactions contained 100 ng of genomic DNA, 0.5 µM each of forward and
reverse primers, 0.5 mM each of the dNTPs, 0.1 vol of 10× PCR buffer (10 mM Tris-HCl,
pH 8.8, 50 mM KCl, 0.1% Triton X-100), 4 mM MgCl2 and 2.5 units pfu turbo polymerase
(Stratagene), and deionized water until a final volume of 50µl. The PCR program consisted
of 30 cycles of denaturation (94ºC for 30 s), annealing (65ºC for 30 s), and extension (72ºC
for 3 min). PCR products were gel-purified and cloned into pCR2.1-TOPO vectors. These
plasmids were used to transform One Shot TOP10 Chemically Competent E. coli cells
(Invitrogen). Then vectors were purified and sequenced. Alignment between gene
sequences obtained from genomic DNA and cDNA was performed. Introns sequences
were confirmed by GenScan software.
RESULTS
Partial amplification of pgchp gene using degenerated primers
The pgchp gene partial amplification from genomic DNA was done using two degenerated
primers: pgchp-DPF1 and pgchp-DPR2A (Table 1). After different reactions, the optimal
annealing temperature was 57.7ºC. PCR products obtained with these primers were run in
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agarose gel. Just one band with high intensity was amplified in reactions with both forward
and reverse primers (Figure 1). This band was cloned and sequenced.
Figure 1. PCR products using forward DPF1 (F), both forward DPF1 and reverse DPR2A (F+R),
and reverse DPR2A (R) primers. M: DNA standards. The arrow indicates the selected band.
After sequencing, the selected fragment was found to consist of 657 bp. The obtained
sequence was translated and compared in Basic Local Alignment Search Tool (BLAST)
with known sequences. A high degree of similarity with several fungal chitosanases was
found, as it is discussed later. The common sequences included three introns described for
genes encoding fungal chitosanases.
Sequence of the pgchp gene by RACE PCR
After induction with glucosamine, mRNA was extracted and used to obtain both 5’- and
3’-RACE-Ready cDNA populations by RT-PCR. Two gene specific primers were
designed from the obtained genomic DNA sequence without introns to get 5’- and 3’-ends:
5’-RACE GSP (5’-CCATTGCAGATAACGGCACCAACG-3’) and 3’-RACE GSP (5’GTTCGACAAGGGTGCGGCATATG-3’), respectively. After 5’ and 3’-RACE reactions,
two bands with the expected size (Figure 2) were gel-purified from 5’- and 3’-RACE
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PCRs, cloned, and sequenced. The cDNA ends of the gene 5’ and 3’ were 559 bp and 796
bp, respectively.
Figure 2. cDNA fragments of 5’- and 3’-RACE PCR (A and B, respectively). M: DNA
standards.
Next, two new primers were designed to obtain the full encoding cDNA (Fwd-pgchp: 5’ATGATGACCTACAGCCGCTTAATCCC-3’
and
Rev-pgchp:
5’-
TCAATCACTGGGGCATTTTCCAC-3’). The full-length cDNA contained a 915 bp open
reading frame encoding a protein of 304 amino acid residues (Figure 3) that included the
known PgChP sequences.
Genomic DNA of pgchp gene, amplified using the same primers for cDNA, was 1146 bp
length. Comparison of cDNA and genomic sequences revealed that pgchp gene had four
introns with a length of 60, 53, 59, and 59 bp (Figure 3). These introns were confirmed by
GenScan program.
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5’-atgatgacctacagccgcttaatccccttcgctctctttgccttgtttggcacagggctc
M M T Y S R L I P F A L F A L F G T G L
gcgcaaacagtcgatggctccaaattcaacaagccagacggcggtccaccaggaagctac
A Q T V D G S K F N K P D G G P P G S Y
ttcgctgctggctcgtctattcccgtcgctgctttgcagagcgctgctgcaaaggctcgc
F A A G S S I P V A A L Q S A A A K A R
actcccgtgccagatgcaacctatcctattaatggcgataacggtgctaagaaagttacc
T P V P D A T Y P I N G D N G A K K V T
atccacagcgactgggctaagttcgacaaggtagatactctcactggtcggttaattgca
I H S D W A K F D K <<
Intron 1
aggtccgcatgcttattaaatggatcatagggtgcggcatatgtttggattgcggatatg
>> G A A Y V W I A D M
gacgtcgactgcgacggcattgactacaagtgcaaggtacggtcactattgttgtctgtc
D V D C D G I D Y K C K <<
Intron 2
tgagatgttattgctgacccgatgttcagggaaacccggatggtcagcaccaaaccaact
>> G N P D G Q H Q T N
tcggagctttggctgcgtatgaagtgccgttctttgtgattcccgacaggtttggaacca
F G A L A A Y E V P F F V I P D R F G T
agtacgcgaagcagcttcctggaaacaacgttggtgccgttatctggtatggtttcctac
K Y A K Q L P G N N V G A V I C<<
ctttgtaagatatctcagcggcgtcttgttctaaccgaaagctagcaatggaaagatgtt
Intron 3
>> N G K M F
ctacggaatttacggagactccgatggcgatacccctcaggtcatcggcgaggcctcatg
Y G I Y G D S D G D T P Q V I G E A S W
gttaatggcccggacctgcttccctaatgatgacttgaatggcaatagtggccatggtga
L M A R T C F P N D D L N G N S G H G D
tgttgatgtgacctgtaagtttgcagccttggcacatctagatccgtattctgtcaacta
V D V T <<
Intron 4
ctaatctaaccagatatcctcttcactggcgacgactcggtgctccccagcagcgctctc
>>Y I L F T G D D S V L P S S A L
aacaagaattatgtcaccaacttcaccactctgcgctctatgggagacaaactcatgact
N K N Y V T N F T T L R S M G D K L M T
gctcttgcgaagaacctcaagttggtagatggaggtgatggaggttctccaacaactacg
A L A K N L K L V D G G D G G S P T T T
gctgggtccaaccctactagcgggtcttgtgagtgggagggacactgtgctggtgcgtct
A G S N P T S G S C E W E G H C A G A S
tgcaaagatgaaaatgattgctccgatcaactggtctgcaaaagtggaaaatgccccagt
C K D E N D C S D Q L V C K S G K C P S
gattga-3’
D -
Figure 3. Full-length DNA sequence of pgchp gene. The open reading frame has been
translated to amino acids. Typical sequences of fungal introns are underlined.
The open reading frame was translated to protein by ExPASy proteomics server at the
Swiss Institute of Bioinformatics (http://www.expasy.org). Analysis of the signal peptidase
cleavage site predicted by the SignalP program (Nielsen et al., 1997) suggested that the
pre-sequence consisted of the first 21 amino acids (MMTYSRLIPFALFALFGTGLA).
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MANUSCRITO 5
DISCUSSION
RACE-PCR is a method widely used in the characterization of genes. The successful of
this technique is highly depended on the amount of mRNA from the gene enconding the
studied protein. Thus, the production of PgChP was induced by glucosamine, as it has been
reported for other chitosanases (Zhang et al., 2001). After the mRNA extraction, both
populations 5’- and 3’-RACE-Ready cDNAs were satisfactorily obtained by RT-PCR.
Using gene specific primers designed from the sequence amplified with degenerated
primers, the 5’- and 3’-ends were obtained by RACE-PCR.
When the complete sequence was obtained, a difference between the pgchp gene from
genomic DNA (1146 bp) and from cDNA (915 bp) could be observed (Figure 3). Thus, the
existence of four introns of 60, 53, 59, and 59 bp in the gene, which are absent in the
cDNA, was confirmed by GenScan software. The small size of these introns compared to
mammalian introns is a typical feature from fungal genes (Gurr et al., 1987). The 5’- and
3’-ends of the four introns are very similar (Figure 3) and coincide with the consensus
splice sequences for fungal introns 5’-splice donor site (GT), the 3’-splice acceptor site
(AG). In addition, three of them have the internal CTRAC sequence (Figure 3), typical of
fungal introns (Wiesner et al., 1988).
The translated sequence of the gene pgchp encodes a chitosanase of 304 amino acids of the
PgChP protein. Comparing the putative amino acid sequence of pgchp gene with seven
fungal chitosanases, the homology was about 60 %. Several common sequences were
found among all these proteins (Figure 4). Three conserved regions have been described in
fungal chitosanases as important for catalytic function (Shimosaka et al., 2005). In two of
these regions there are one aspartic and one glutamic acid residues, essentials for catalytic
function of the chitosanase from F. solani. The predicted amino acid sequence of PgChP
also shows these two residues (Figure 4). Carboxylic amino acids (Glu or Asp) have been
identified as necessary residues for the catalytic function as proton donors in a large
number of glycosyl hydrolases (Monzingo et al., 1996; Robertus et al., 1998). PgChP and
the other seven fungal chitosanases show eleven common residues of aspartic acid as well
as two of glutamic acid (Figure 4). Moreover, 10 cysteines of PgChP are found in all these
fungal chitosanases (Figure 4). Cysteine residues are known to play an important role on
the stabilization of the tertiary structure by the formation of disulfide bridges in fungal
proteins (Nakaya et al., 1990; Lacadena et al., 1995; Marx et al., 1995; Lee et al., 1999).
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RESULTADOS
PgChP
A.fumigatus
N.fischeri
A.niger
A.oryzae-1
A.oryzae-2
A.terreus
A.clavatus
--MMTYSRLIPFALFALFG-TGLAQTVDGSKFNKPDGGPPGSYFAAGSSIPVAALQSAAA
MPSTTIIRQLAIS-LALCN-SALGQVVNGADYNKPNGGPPASFFAAASTMPVAALQAAAA
MPSKTIIRQLAIS-VALCN-SALGQVVNGADYNKPNGGPPASFFAAASTMPVAALQAAAA
MAFKTTAG--LAF-LALAG-SVKAQSVDGSKYNSPSNGPPASYFAAATTLPVAALQSAAA
MPIKSFASRLALS-LAICG-TAMGQKVNGADYNKPDGGPPAKFFQASSSIPVAAIQAAAA
MPIKSFASRLALS-LAICG-TAMGQKVNGADYNKPDGGPPAKFFQASSSIPVAAIQAAAA
MVFKKAAIGLTLP-FALFSSVALGQTVDGSDYDSPNGGPPGSYFAAASTMPVAALQAAAA
MIFKSSLSHAAAS-LLLLT-PALAQKVQGPEYNKPSAGPPASFFAAAPTMPVAALKSAVA
57
58
58
56
58
58
59
58
PgChP
A.fumigatus
N.fischeri
A.niger
A.oryzae-1
A.oryzae-2
A.terreus
A.clavatus
KARTPVPDATYPINGDNGAKKVTIHSDWAKFDKGAAYVWIADMDVDCDGIDYKC-----KATKVPSLATYPVSQDKGAAKSTIHTDWASFSEGASISWVADMDVDCDGLNSGC-----KASKVPSLATYPVSQDSGAAKSTIHTDWASFSEGASISWVADMDVDCDGLNSGC-----KASSVPSKATYPVNTDDDSPKSTIHSDWVKFNQGAALSWVADMDVDCDGIDYKC-----KASKVPSHATYPIGQ--GSTKSTIHSDWAGFSEGAAFSFIADMDVDCDGLNHGC-----KASKVPSHATYPIGQ--GSTKSTIHSDWAGFSEGAAFSFIADMDVDCDGLNHGC-----KATKVPSYATYPVSQDDNAKKSTIHSDWASFSQGAAISWVADMDVDCDGIDSGCEHEIKT
RASVVPKNAAYPVNQD-GGPTATIHADWASLPTAAAYVYTADMDVDCDGLDHNC------
111
112
112
110
110
110
119
111
PgChP
A.fumigatus
N.fischeri
A.niger
A.oryzae-1
A.oryzae-2
A.terreus
A.clavatus
-KGNPDGQHQTNFGALAAYEVPFFVIPDRFGTKYAKQLPGNNVGAVIC----NGKMFYGI
-QGNPDGQPQTNWGALSAYEVPFIVIPDKYLSANSGALPGNNIAAVIC----NGKMFYGI
-QGNPDGQPQTNWGALSAYEVPFIVIPDKYLSANTGALPGNNIAAVIC----NGKMFYGI
-KGNGDGLPETNWGALSAYEVPWIVIPDQFLTANEDLLPGNNVAAVIC----NGKMYYGI
-KGNPDGQKETNWGALSAYEVPFIVIPQEFLDANKGTLKGNAVAAVIC----NGKMFYGI
-KGNPDGQKETNWGALSAYEVPFIVIPQEFLDANKGTLKGNAVAAVICATSSNGKMFYGI
SQGNPDGQDATNWGALAAYEVPFIVIPQKYLDHNGNALKGNNIAAVIC----NGKMFYGI
-KGNPDGQPDTNFGALAAYEVPFVVIPDRFATTYASALPGNNIVAVIC----DGKMFYGI
166
167
167
165
165
169
175
166
PgChP
A.fumigatus
N.fischeri
A.niger
A.oryzae-1
A.oryzae-2
A.terreus
A.clavatus
226
227
227
218
225
229
235
226
PgChP
A.fumigatus
N.fischeri
A.niger
A.oryzae-1
A.oryzae-2
A.terreus
A.clavatus
YGDSDGDTPQVIGEASWLMARTCFPNDDLNGNSGHGDVDVTYILFTGDDSVLPSSALNKN
LGDSNGDSPQVTGEASWLMARTCFPNEGLNGNNGHTGVDVTYIVFTGKNAVLPSSALTKN
LGDSNGDSPQVTGEASWLMARTCFPNEGLNGNNGHTGVDVTYIVFTGKDAVLPSSALTKN
LGDSNGDDPEVTGEASWLMARTCFPDDDLNGAEGHAEADVTCKPYT-------RSALNKN
FGDSNGDSPQVTGEASWLMARTCFPKEDLNGNKGHTAADVTYIVFTGDKAVLPSSALNKN
FGDSNGDSPQVTGEASWLMARTCFPEEDLNGNKGHTAADVTYIVFTGDKAVLPSSALNKN
LGDANGDEPQVTGEASWLMARTCFPNEGLNGNKGHTAADVTYILFTGDESVLPSSALNEN
FGDTDGDHPQVIGEASWLMARTCFPNDNLNGDSGHVPADVTYIFFTGKDSVLPSSAVNKN
*
*
YVTNFTTLRSMGDKLMTALAKNLKLVDGGDGG---------SPT--TTAGSNPT--SGSC
YITNFTTLRSMGDKLVNALVSNLGLSGTPPTK---------TTTRVTTTTTKPT-SAASC
YITNFTTLRSMGDKLVNALASKLGLSGTPPTK---------TTTLVTTTTTKPT-STASC
YITNFSTLRSMGDKLVGALASNLGLTSSA------------------------SGSTATC
YITNFDTLRSMGDSLVGALAKNLNLGGGGGNP---------PTTLTTTSIPEPTGGSGSC
YITNFDTLRSMGDSLVGALAKNLNLGGGGGNP---------PTTLTTTSIPEPTGGSGSC
YITNFSTLRSMGDKLVNALASNLGLSGGSGGT---------PTT--TTTATQPT-STGTC
YITDFTKLRSMGDSLVNAFASQLGISSGGGNGGGDGGHTTLHTTTATRTATAPT-STATC
PgChP
A.fumigatus
N.fischeri
A.niger
A.oryzae-1
A.oryzae-2
A.terreus
A.clavatus
EWEGHCA-----GASCKDENDCSDQLVCKSGKCPSD-----------------------SWAGHCL-----GASCSSDDDCADALVCTAGKCSVDGAA-TCSWEGHCEGASCSSDDDCS
SWAGHCL-----GASCSSNDDCADALVCAAGKCSVDGAA-TCSWEGHCEGVYS------SWQGHCE-----GAICSTEDDCSDDLVCDSGKCSSPDED--------------------SWPGHCA-----GATCSSNDDCSDDLTCQNGKCASDGSAETCSWEGHCKGATCSSNDDCS
SWPGHCAGFKNKGATCSSNDDCSDDLACQNGKCASDGSAETCSWEGHCKGATCSSNDDCS
SWAGHCE-----GASCSTNDDCSDQLACKNGVCSTDGEV-VCSWEGHCEGATCSSNDDCS
DWEGHCL-----GTACSNNGECSDPFGCINGFCNYDPTL-TCQWAGHCVGASCSSNDQCS
304
331
324
288
331
340
337
339
PgChP
A.fumigatus
N.fischeri
A.niger
A.oryzae-1
A.oryzae-2
A.terreus
A.clavatus
---------------------------------------------------DDLYCKSGSCTA------------------------------------P----------------------------------------------------------------------DGDDEDDEDEEENDEDDEDDDDEDDEDDE-----DELACISGICSVDNGVETCEWEGHCEGASCSSHDDCDGNLACKNGKCSA--DELACISGICSVDNGVETCEWEGHCEGASCSSHDDCDGNLACKNGKCSA--DELYCAGGACTSA--------------------------------------DPFACIDGACAVDT-SLDCSRKGHCAGTTCLSDSDCSRPLSCILGVCANQSG
273
277
277
254
276
280
283
285
344
317
380
389
350
390
Figure 4. Alignment of PgChP with other fungal chitosanases. Common amino acids are denoted
by background shaded letters. Asterisks indicate essential amino acid residues for catalytic activity
of fungal chitosanases. Accession numbers in Genbank are XM_754126 (A. fumigatus),
XM_001262949
(N.
fischeri),
XM_001390407
(A.
niger),
BAD08218
(A.
oryzae-1),
XP_001824257 (A. oryzae-2), XM_001209034 (A. terreus), XM_001274664 (A. clavatus).
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Six glycosylated isoforms of PgChP with N-linked oligosaccharides have been reported
(Rodríguez-Martín et al., 2009). In N-glycosylations, glycans are attached to the protein
via N-acetylglucosamine (GlcNAc) to the amide group of asparagine within an Asn-XSer/Thr/Cys motif, where X is any amino acid apart from proline (Morelle et al., 2006).
Two N-glycosylation consensus sequences were found in the translated pgchp gene: NFT
(residues 230-232) and NDC (residues 288-290).
Glycosylation is the most common post-translational modification in proteins.
Carbohydrates participate in important biological processes, including inter- and
extracellular signalling, protein regulations and interactions, and molecular recognition.
Glycans can determine the localization, activity and function of proteins, as well as
influence on physicochemical characteristics of proteins such as folding, solubility,
resistance to proteases or thermal stability (Geyer et al., 2006; Morelle et al., 2006). The
potential role of the N-linked oligosaccharides of PgChP on intra- or extracellular
signalling could be essential for the antifungal activity of this protein.
The predicted mass of the translated nucleotidic sequence, according to the Compute
pI/Mw tool (Gasteiger et al., 2005), was 31,991 Da, which is close to the 30 to 31 kDa
determined for the six excreted isoforms of the protein by ESI-MS (Rodríguez-Martín et
al, 2009). The difference between the predicted mass and the determined weight can be
explained by the lack of the signal peptide (2,300 Da) in the excreted protein and the
presence of N-linked oligosaccharides. The predicted isoelectric point (Gasteiger et al.,
2005) was 4.8, which is close to the value of 5 determined by 2D-PAGE—IEF (RodríguezMartín et al., 2009).
Recently, the genome of P. chrysogenum Wisconsin 54-1255 has been sequenced (Van den
Berg et al., 2008). A putative protein can be found in BLAST (accession number
CAP80409) that differs from the precursor of PgChP in only four amino acids. However,
the production of the protein by this mould has not been reported.
In conclusion, the gene encoding the chitosanase PgChP with antifungal activity has been
sequenced. The putative amino acid sequence of PgChP shows two consensus sites for
glycosylations. The role of N-linked oligosaccharides from PgChP on antifungal activity
deserves further investigation.
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ACKNOWLEDGEMENTS
This work was supported by the Spanish Ministry of Education and Science (AGL200406546-ALI) and FEDER. Andrea Rodríguez and Raquel Acosta were recipients of a FPI
grants from the Spanish Ministry of Education and Science. Authors would like to thank
Dr. Susan Liddell (Division of Animal Sciences) and Prof. Ian Connerton (Division of
Food Sciences) for their support during Andrea Rodriguez stay’s at Nottingham
University.
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Lee, Y.S., Yoo, J.S., Chung, S.Y., Lee, Y.C., Cho, Y.S., and Choi, Y.L. Cloning,
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Morelle, W., Canis, K., Chirat, F., Faid, V., and Michalski, J-C. The use of mass
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Nakaya, N., Omata, K., Okahashi, I., Nakamura, Y., Kolkenbrock, H.J., and Ulbrich,
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Wagner, C., Wortman, J., and Bovenberg, R.A. Genome sequencing and analysis of the
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Sphingomonas sp. CJ-5. J. Zhejiang. Univ. Sci. B. 8(11): 831-838 (2007).
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Manuscrito 6: Effect of different treatments
on chitosanase and antifungal activity of PgChP
from Penicillium chrysogenum
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RESULTADOS
Effect of different treatments on chitosanase and antifungal activity of
PgChP from Penicillium chrysogenum
Andrea Rodríguez-Martín, Raquel Acosta, Félix Núñez, Miguel A. Asensio*
Higiene y Seguridad Alimentaria, Facultad de Veterinaria. Universidad de Extremadura, Avda. de la
Universidad, s/n. 10071- Cáceres, Spain.
* Corresponding author: Telephone (+34) 927 257 125; fax: (+34) 927 257 110.
E-mail: masensio@unex.es
ABSTRACT
The protein PgChP showed antifungal and chitosanase activities with potential application
in food industry as protective additive and production of chitosan oligomers, respectively.
Haemolytic assay was negative, essential characteristic for using it for human applications.
PgChP was resistant at heat treatments even higher than 60ºC. Different ions influenced on
chitosanase and antifungal activities of the protein; thus, Na+ and Mg2+ affected to the
chitosanase activity, whereas the antifungal activity was only affected for Mg2+ and Ca2+
and not for monovalent ions. Non-ionic detergents remained or improved the chitosanase
action, although decreased the antifungal activity. Ionic detergents inhibited both
antifungal and chitosanase activities. EGTA (0.1 mM), Triton X-100 (1%) and phosphate
(1 mM) increased the chitosanase activity of PgChP. The antifungal activity does not seem
to be related to chitosan degradation in sensitive moulds.
KEY WORDS: chitosan oligomer, chitosanase, antifungal activity, food preservative,
haemolytic activity.
INTRODUCTION
Natural antifungal proteins are a new source of clinically useful therapeutics. Antifungal
proteins have different action mechanisms. Some of them interfere on DNA, RNA, and
protein synthesis, or they can act on fungal walls in diverse ways. Some of them decrease
the ATP level (Cociancich et al.., 1993) or depolarize the membrane (Thevissen et al..,
1996). Other antifungal proteins interfere with essential elements of fungal walls, such as
-1,3-glucan (Odds et al.., 2003), chitin (Yang and Gong, 2002; Von der Weid et al.,
2003), or chitosan (Davis and Eveleigh, 1984). These compounds can be even hydrolized
by some antifungal proteins like glucanases, chitinases, and chitosanases.
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MANUSCRITO 6
The most useful antifungal proteins are those that act specifically against fungal
membranes because of their lesser toxic effects on mammalian cells (Groll et al., 1998).
However, none of these antifungal proteins is currently utilized because they have shown
to be cation-sensitive. Sensitivity to high-ionic-strength conditions may be a crucial point
in the application of antifungal proteins under physiological conditions, or in many food
products with high salt concentration. The protein PgChP was previously purified from the
strain P. chrysogenum AS51D for its ability to inhibit the growth of toxigenic moulds
(Acosta et al., 2009). Given that this mould was isolated from dry-cured ham, a substrate
with a high salt concentration, PgChP could withstand cations and have adequate
characteristics for the use in foods.
PgChP showed high homology with fungal chitosanases according to its amino acid
sequence (Rodríguez-Martín et al., 2009a and 2009b). Thus, PgChP could have
chitosanase as well as antifungal activity. Chitosanases can degrade a variety of chitosans
with different degrees of acetylation (Ohtakara et al., 1988; Osswald et al., 1993) to yield
chitosan oligomers, which have interesting biological properties, including antifungal and
antibacterial activity (Allan and Hadwiger, 1979; Hirano and Nagano, 1989), as food
preservatives (Rivas et al., 2000).
Chitosan is a polysaccharide found in fungal walls (Zhang et al., 2000; Kumiko et al.,
2002; Di Mario, 2008) and its degradation by chitosanases can be responsible for inhibiting
target moulds (Saito et al., 2009). Thus, the antifungal activity of PgChP could be related
to the chitosanase activity.
To design potential applications, the impact of environmental factors on PgChP activity
should be known. The aim of the present work was to study the influence of factors such as
temperature, different substrates, ions, and detergents on both antifungal and chitosanase
activities.
MATERIALS AND METHODS
Microbial strain
The PgChP producing mould, P. chrysogenum AS51D, and the toxigenic mould
Penicillium commune Pc332 were isolated from dry-cured ham and belong to the fungal
collection of Food Hygiene (University of Extremadura, Spain).
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PgChP purification
P. chrysogenum AS51D was grown at 25ºC for 15 days in malt extract broth made of
glucose 2% (w/v), malt extract 2% (w/v) and peptone 0.1% (w/v), pH 4.5. Then, the
medium was filtered though a 0.45 µm diameter pore size cellulose acetate membrane and
PgChP was purified using an ÄKTA fast protein liquid chromatography system
(Amersham Pharmacia Biotech AB, Uppsala, Sweden) according to Acosta et al. (2009).
For this, filtered medium was loaded on a HiTrap SP HP 5 ml ion exchange column
(Amersham Pharmacia Biotech AB) equilibrated with 0.02 M sodium acetate buffer, pH
4.5. Bound proteins were eluted with a gradient of 0.02 M sodium acetate, 1 M NaCl
buffer, pH 4.5: 0% to 25% in 15 column volumes (CV), increase to 100% in 2 CV, and
finally keeping 100% for 15 CV. Eluted proteins were detected at 214 nm. The fraction
that showed antifungal activity in previous studies (Acosta et al., 2009) was purified using
a HiLoad 26/60 Superdex 75 prep grade gel filtration column (Amersham Pharmacia
Biotech AB) equilibrated with 50 mM sodium phosphate, 0.15 M NaCl buffer, pH 7, at a
constant flow rate of 2.5 ml/min. Data collection and processing were done using the
UNICORN software 4.12 (Amersham Pharmacia Biotech AB). Finally, the fraction
containing PgChP was desalted and concentrated with YM-10 microcon centrifugal filter
units (Millipore Iberica S.A.U., Madrid, Spain) and kept at -20ºC.
Protein was quantified by Johnson method according to Córdoba (1990). An aliquot of the
purified protein (100µl) was dried in a heat cabinet at 100 ºC. The sample was digested by
adding 200 µl of sulfuric acid in a sand bath at 120 ºC. Then, 4.8 ml of distilled water, 3 ml
of 4N NaOH and 2 ml of Nessler reactive made of KI 0.4% (w/v), 0.4% HgI2 (w/v) and
0.175% (w/v) arabic gum were added. After 10 min at room temperature the absorbance
was measured at 490 nm in a spectrophotometer. A calibration curve was obtained with
different concentrations of ammonium sulfate.
Sodium dodecyl sulphate-polyacrylamide gel eletrophoresis (SDS-PAGE)
SDS-PAGE was performed as described Laemmli (1970). The protein PgChP was
separated on a 4% (v/v) polyacrylamide stacking and 8% (v/v) polyacrylamide resolving
gels. Gels were stained with Imperial Protein Stain (Pierce Biotechnology, Inc., Rockford,
U.S.A.). The molecular weight was determined by comparison of its electrophoretic
mobility with proteins of the molecular mass marker (66-6.5 kDa) from Sigma (SigmaAldrich Química S.A., Madrid, Spain).
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Haemolytic activity assay
The haemolytic activity of the protein was determined using sheep defibrinated blood
(Oxoid). The erythrocytes were isolated by centrifugation (1000 ×g for 6 min at room
temperature) and washed three times with 50 mM sodium phosphate, 0.15 M NaCl buffer,
pH 7. The cell concentration in the stock suspension was adjusted to 1×109 cells/ml. A
volume of 50 µl of cells suspension was added to 200 µl of sterile concentrated protein (10
µg/ml) and incubated at 37ºC. After 0, 8, and 24 h, the samples were placed on ice,
immediately centrifuged at 4ºC, and the supernatant was carefully collected. The
haemolysis was determined by measuring absorbance of the supernatant at 570 nm
(Moerman et al., 2002) and 541 nm (Lai et al., 2005). Controls for 0% and 100%
haemolysis were obtained suspending sheep red blood cells in the buffer or distilled water,
respectively. The haemolysis percentage was calculated using the following equation:
Percentage haemolysis of supernatants = [(Abs in the peptide solution − Abs in buffer)/Abs
in water solution] × 100.
Three independent assays were conducted with triplicated readings for each one.
Evaluation of chitinase and chitosanase activities
Chitinase and chitosanase activities were determined with a modification of the 3,5dinitrosalicylic acid method (Su et al., 2006) by measuring the reducing sugars liberated
during the hydrolysis of chitin or chitosan. One ml of the 1% (w/v) chitin or chitosan
(Sigma-Aldrich) in 1% (v/v) acetic acid was added to 1 ml of the PgChP solution
(10µg/ml) and incubated for 2-4 h at 37ºC. Then, to detect the hydrolisis reaction, 2 ml of
dinitrosalicylic acid reagent, made of 1% (w/v) 3,5-dinitrosalicylic acid (Sigma-Aldrich),
0.05% (w/v) sodium sulfite, and 1% (w/v) phenol were added and the sample was
incubated at 100ºC for 45 min. Finally, 1 ml of 40% (w/v) potassium sodium tartrate was
added and the solution was centrifuged. The supernatant was measured at 520 and 575 nm.
To quantify the reducing sugars liberated, a calibration curve was obtained with different
concentrations of D-(+)-Glucosamine hydrochloride (Sigma-Aldrich). A chitinase from
Streptomyces griseus (Sigma-Aldrich) with both chitinase and chitosanase activities was
used as positive control.
Preparation of colloidal chitin: The colloidal chitin was obtained following the method of
Roberts and Selitrennikott with minor modifications (Harighi et al., 2007). Ten grams of
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chitin powder were added slowly into 100 ml of H3PO4 (85%) at 25ºC under vigorous
shaking for 2 h. This solution was carefully added over 1 liter of absolute ethanol at -20ºC.
The mix was shaken for 30 min and stored until use. Ten ml of suspension were
centrifuged, the pellet was washed three times with 50 mM sodium phosphate buffer pH 7,
and resuspended in distilled water. The final concentration of colloidal chitin was around
1%.
Chitinase activity assay by gel electrophoresis
The chitinase activity assay was carried out as described by Trudel and Asselin (1989).
PAGE-SDS was performed in 4% (v/v) polyacrylamide stacking gel and 8% (v/v)
separating polyacrylamide gradient gel containing 0.01% (w/v) glycol chitin and 0.1%
(w/v) SDS. Samples were boiled for 5 min with 20% (v/v) glycerol, 0.025% (w/v)
bromophenol blue, 2% (w/v) SDS and 5% (v/v) -mercaptoethanol in 62.5 mM Tris-HCl
(pH 6.8). The gel was run at 100 V for 1 h.
After electrophoresis, the gel was incubated for 2 h at 37 ºC under continuous shaking in
100 mM sodium acetate buffer (pH 5.0) containing 1% (v/v) Triton X-100. Then, the gel
was stained with a 0.01% (w/v) calcofluor white M2R (Sigma-Aldrich) in 500 mM TrisHCl (pH 8.9). After 5 min, this solution was removed and the gel was incubated in distilled
water at room temperature for about 1 h. Lytic zones were visualized in the UV
transilluminator Syngene G-Box (Synoptics, Fredericks, USA). The gel was photographed
and analyzed with the GeneSnap software (Synoptics). An additional gel was stained only
with Imperial Protein Stain (Pierce Biotechnology, Inc., Rockford, U.S.A.). The chitinase
from S. griseus (Sigma) was used as positive control. The molecular mass marker used was
66-6.5 kDa (Sigma).
Glycol chitin synthesis: 0.2 g glycol chitosan was dissolved in 4 ml of 10% acetic acid and
the mixture was shaken overnight at 22ºC. Methanol (18 ml) was slowly added and the
solution was filtered through 0.45 µm pore size cellulose acetate membrane filters. Acetic
anhydride (0.3 ml) was added and kept for 30 min. The resulting gel was cut into small
pieces and after adding 1 vol of methanol was shaken for 4 min. This suspension was
centrifuged at 27000 ×g at 4ºC for 15 min and the pellet was resuspended in 20 ml of
0.02% (w/v) sodium azide. The final concentration of the glycol chitin was 1% (w/v).
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Evaluation of Antifungal activity
Quantitative assay for fungal growth inhibition was performed following the protocol
developed by Broekaert et al. (1990) with some modifications (Acosta et al., 2009). P.
commune Pc332 (Núñez et al., 1996) was used as reference strain. To perform the
inhibition test in microtitter plates, 100 µl of the fraction obtained by gel filtration
chromatography were dispensed in triplicate wells. Each well was loaded with 100 µl of
concentrated malt extract broth (4% malt extract, 4% glucose and 0.2% peptone, pH 4.5)
inoculated with the reference strain to c.a. 106 spores/ml. The plates were incubated for up
to 72 h at 25 ºC. Fungal growth was determined by measuring the optical density variation
at 595 nm.
Effect of heat-treatments
The study of the protein stability after heat treatment was carried out according to Okkers
et al. (1999). Aliquots of PgChP protein (10 µg/ml) were exposed to different
combinations of temperatures (60, 80, and 100 ºC) and times (10, 20, and 30 min), as well
as to sterilization at 121 ºC for 15 min. After the heat-treatment, the samples were cooled
on ice. Then, antifungal and chitosanase activities were evaluated as described above.
Influence of different chemicals on chitosanase and antifungal activity
To study the influence of some chemicals on the protein, the activity of PgChP (10 µg /ml)
was tested with different compounds: NaCl, CaCl2, KCl, or MgCl2 (0.1, 1, 10, and 100
mM); EDTA or EGTA at 0.01, 0.1, and 1 mM; 0.5% and 1% of Triton X-100, tween-20,
tween-80, SDS, or deoxycholic acid; NaH2PO4 at 0, 0.1, 1, 10, and 100 mM.
Statistical analysis
Data were statistically analyzed using SPSS software package (version 13.0. SPSS Inc.,
Chicago, USA) by a variance analysis (one way ANOVA) and the means were separated
by the Tukey honest significant difference test.
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RESULTS
Haemolytic activity
To study the haemolytic activity of PgChP on mammalian cells, the effect of the protein
was examined on sheep red blood cells. The protein PgChP did not show haemolytic
activity, being the value obtained of 0%, even after 24 h.
Chitosanase and chitinase activities
The protein PgChP showed chitosanase activity, but no chitinase activity was found by the
3,5-dinitrosalicylic acid method (Figure 1).
Figure 1. Chitinase and chitosanase activities of PgChP expressed as liberated D-glucosamine after
4 h. Asterisks indicate statistically significant differences to the negative control without protein
(***p<0.001). Chitinase from S. griseus (50 µg/ml) was used as positive control.
To further evaluate the chitinase activity, the hydrolysis of chitin was tested by
electrophoresis in a glycol chitin gel. No activity was observed loading the polyacrilamide
gel with 18 µg of PgChP (Figure 2). Lytic zones by chitinase from S. griseus, used as
positive control, were observed as black bands after staining with calcofluor white M2R.
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&%
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Figure 2. Chitinase activity of PgChP evaluated by gel electrophoresis. SDS-PAGE gels
with glycol chitin were stained with Imperial Protein Stain (A) and Calcofluor White M2R
(B). M: protein marker, Q: Chitinase from S. griseus (100 µg).
Protein stability to different temperatures
PgChP protein showed a remarkable stability at high temperatures. The chitosanase
activity did not show a significant reduction after heat treatments even at 60ºC for 30 min
(Figure 3).
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Figure 3. Effect of heat-treatment on PgChP chitosanase activity expressed as mean and standard
deviation of liberated glucosamine after 2 h. Asterisks indicate statistically significant differences
(p<0.05) to the positive control without heat-treatment (C+).
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The antifungal activity seems to withstand even treatments of 80ºC up to 20 min (Figure
4), whereas higher temperatures inactivated PgChP antifungal activity allowing an
%'%
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unrestricted growth of the sensitive mould.
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Figure 4. Antifungal activity of heat-treated PgChP expressed as mean and standard deviation of P.
commune Pc332 growth after 48 h. Asterisks indicate statistically significant differences (p<0.05)
to the positive control with non heat-treated PgChP. C+: positive control with non heat-treated
PgChP; C-: negative control without PgChP.
Effect of ions on PgChP chitosanase and antifungal activities
PgChP chitosanase activity was not affected by the presence of up to 100 mM CaCl2 or
KCl ions whereas it decreased at the highest NaCl or MgCl2 concentration (Figure 5).
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.
0
/
1
"
"
Figure 5. Effect of monovalent (A) and divalent (B) ions on the chitosanase activity of PgChP
expressed as mean and standard deviation of liberated glucosamine after 3 h. Asterisks indicate
statistically significant differences to the positive control without ions (closed bars) (**p<0.01;
***p<0.001).
On the other hand, the inhibition of the reference mould was unaffected by monovalent
ions, K+ and Na+, but significantly decreased with high concentrations of divalent ions
(Figure 6).
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.
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Figure 6. Effect of monovalent (A) and divalent (B) ions on antifungal activity of PgChP against
P. commune Pc332 expressed as mean and standard deviation of optical density after 72 h.
Asterisks indicate statistically significant differences to the negative control (C-) without PgChP (*
p<0.05; ** p<0.01; *** p<0.001).
Effect of cation chelators
Activity of PgChP was also assayed in presence of EDTA and EGTA. The chitosanase
activity was strongly affected by high levels of chelators. Thus, 1 mM EDTA inhibited
chitosanase activity, whilst even low levels of EGTA increased chitosan hydrolysis by
PgChP (Table 1).
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Table 1. Effect of EDTA and EGTA on chitosanase activity of PgChP after 2 h incubation.
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Results are expressed as mean ± SD of liberated glucosamine. Asterisks indicate statistically
significant differences to the positive control without chelators (*p<0.05; ***p<0.001).
The antifungal activity of PgChP was observed at the lowest EDTA concentration (0.01
mM) but not at 0.1 mM (Table 2). The highest level of EDTA tested inhibited growth of
the reference mould, and no further effect on the antifungal activity could be observed.
EGTA did not seem to inhibit growth of the reference mould even at the highest
concentration tested. No effect of EGTA was observed on the antifungal activity against P.
commune Pc332, irrespectively of the concentration used (Table 2).
Table 2. Effect of EDTA and EGTA on the antifungal activity of PgChP against P. commune
Pc332 after 48 h incubation.
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Results are expressed as mean ± SD of optical density. Asterisks indicate statistically significant
differences to the negative control without PgChP (C-) (*p<0.05; **p<0.01; ***p<0.001).
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Resistance to protein-denaturing detergents
Detergents displayed different effects on the chitosanase activity of PgChP (Figure 7).
Non-ionic detergents showed no inhibition on the activity. In fact, tween 20, tween 80, and
Triton X-100 even increased the chitosanase activity. In contrast, the ionic detergents
sodium dodecil sulphate (SDS) and deoxycholic acid partially or completely inhibited the
chitosanase activity, respectively, for the 2 h incubation time.
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Figure 7. Effect of different detergents on chitosanase activity of PgChP expressed as mean and
standard deviation of liberated glucosamine after 2 h. Asterisks indicate statistically significant
differences to the positive control without detergents (*p<0.05; ***p<0.001).
On the other hand, PgChP showed no antifungal activity on P. commune Pc332 with 0.5 or
1% (w/v) of the tested detergents.
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Figure 8. Effect of different detergents on antifungal activity of PgChP against P. commune Pc332
expressed as mean and standard deviation of optical density after 72 h. Asterisks indicate
statistically significant differences to the negative control (C-) without PgChP (*** p<0.001).
Effect of sodium phosphate on the PgChP activity
The chitosanase activity decreased at the highest concentration of sodium phosphate tested,
whereas levels below 100 mM favoured this activity (Table 3).
Table 3. Influence of phosphate on the chitosanase activity of PgChP after 2 h of incubation.
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Results are expressed as mean ± SD of liberated glucosamine. Asterisks indicate statistically
significant differences to the positive control without phosphate (** p<0.01; *** p<0.001).
Phosphate also seems to affect the antifungal activity of PgChP, but the uneven growth of
the reference mould in negative controls did not allow to obtain more consistent results
(Table 4).
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Table 4. Influence of phosphate on the antifungal activity of PgChP against P. commune Pc332
after 72 h.
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Results are expressed as mean ± SD of optical density. Asterisks indicate statistically significant
differences to the negative control without PgChP (** p<0.01; *** p<0.001).
DISCUSSION
As it has been shown in a previous work, the protein PgChP showed antifungal activity
against P. commune Pc332 (Acosta et al., 2009), a toxigenic mould present in foods (Sosa
et al., 2002). PgChP was obtained by ion exchange chromatography looking for cationic
proteins (Acosta et al., 2009), because the cationic character in proteins is related to a
higher antifungal activity, but to a lower toxicity for mammalian cells (Hong et al., 2001).
Given that PgChP shows an isoelectric point close to 5 (Rodríguez-Martín et al., 2009a),
the protein is not cationic under physiological conditions and its potential as toxic for
humans has to be assessed. The haemolysis assay to test for cytolytic activity is generally
accepted as proof that the protein can be regarded as safe (Van’t Hof et al., 2001). PgChP
did not show any haemolytic activity even though in depth tests are needed to confirm the
safety of this protein.
The amino acid sequence of PgChP revealed a high homology to fungal chitosanases
(Rodríguez-Martín et al., 2009a, 2009b). Thus, the chitosanase activity of PgChP was
evaluated. As it is shown in figure 1, the protein showed a significant chitosanase activity.
Some microbial chitosanases, such us the chitosanase II from Acinetobacter sp. strain
CHB101, hydrolyze colloidal chitin and glycol chitin besides chitosan (Shimosaka et al.,
1995). However, PgChP did not show activity on either of these substrates, which agrees
with the fact that most chitosanases do not have chitinase activity (Su et al., 2006;
Fukamizo and Brzezinski, 1997).
Fungal chitosanases differ in their thermostability. There are chitosanases such as Csn2
from Gongronella sp. JG (Wang et al., 2008) that resist 50 and 55ºC for 30 and 11 min,
respectively. Bacterial chitosanases have shown stability within a similar or lower range of
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temperature, such as those from Acinetobacter sp. C-17 (Zhu et al., 2003) and
Sphingomonas sp. CJ.5, that are stable at 30-40ºC and 45ºC for one hour, respectively (Zhu
et al., 2007). Given that PgChP kept chitosanase activity after heating at 60ºC for 30 min
(Figure 3), it can be regarded as more stable than other chitosanases.
On the other hand, the antifungal activity of PgChP was more heat-resistant than
chitosanase activity. PgChP antifungal activity resisted even at 80 ºC for up to 20 min. The
fact that chitosanase and antifungal activities of PgChP differ in thermostability indicates
that chitosan-hydrolysis is not essential for the antifungal activity, as it will be discussed
later.
The presence of K+ and Ca2+ did not affect the chitosanase activity of PgChP at the highest
tested concentration, but 100 mM Mg2+ and Na+ inhibited that activity. However, K+ and
Ca2+ activate several fungal chitosanases (Alfonso et al., 1992; Fenton and Eveleigh,
1981). As a consequence, metal ions can have different impact on fungal chitosanases.
This effect can be related to the influence of different concentrations of particular ions on
the structure of chitosanases (Wang et al., 2008). Regarding the antifungal activity of
PgChP, over 1 mM Mg2+ or Ca2+ reduced the inhibition on P. commune, whereas
monovalent ions Na+ or K+ did not affect the antifungal activity (Figure 6). The fact that
divalent ions repressed the antifungal activity could mean that these ions compete with
PgChP for potential target sites in the fungal wall, as it has been described for metal ions
on other antifungal proteins (Broekaert et al., 1992; Kaiserer et al., 2003; Theis et al.,
2003). In addition, the divergent effect of Na+ and Ca2+ on PgChP chitosan hydrolysis and
fungal inhibition supports that different mechanisms of action are responsible for these
activities.
The activity of proteins can also be influenced by low concentrations of metal ions. The
addition 1 mM of EDTA, a chelator of Mg2+ and Ca2+ (Choucair et al., 2006), decreased
the chitosanase activity. Then, the absence of Mg2+ seems to inhibit the chitosanase activity
of PgChP. In contrast, EGTA inhibited neither chitosanase nor antifungal activity of
PgChP, and even notably increased the chitosanase activity (Table 1). EGTA is a quite
specific chelator of Ca2+ (Choucair et al., 2006). Thus, the chitosanase activity of PgChP
was not affected by the absence of this ion.
On the contrary, the antifungal activity was not affected by EGTA or low levels of EDTA
(Table 2), whereas high levels of EDTA strongly limited the development of the reference
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mould. Once more, the differences on chitosanase and antifungal activities of PgChP are
evidenced, due to the impact of low levels of metal ions.
Ionic and non-ionic detergents had a distinct effect on PgChP activities. Non-ionic
detergents (tween 20, tween 80, and Triton X-100) led to an increase of chitosanase
activity (Figure 7), probably due to an improved protein-substrate interaction. However, no
clear effect of these detergents could be observed on the antifungal activity, due to the low
growth of P. commune Pc332 even when added at 0.5%.
The ionic detergents SDS and deoxycholic acid caused loss of both chitosanase and
antifungal activities. The effect of these detergents can be attributed to protein denaturation
that prevents PgChP from being active.
Finally, sodium phosphate increased the chitosanase activity of PgChP, except at the
highest concentration (100 mM) which caused loss of activity. These results are somehow
similar to those obtained with identical levels of NaCl. Thus, the main impact of sodium
phosphate on this activity can be attributed to Na+ concentration. The evaluation of
phosphate effect on the antifungal activity was hampered by an uneven growth of Pc332 in
control wells without PgChP. As it can be observed in table 4, 10 mM phosphate PgChP
exerted a significant inhibition (p<0.01) on P. commune, but this was not observed at lower
concentrations. The poorer growth reached by P. commune in the negative controls can
mask statistically significant differences due to the antifungal activity.
As it has been evidenced mainly by the residual activity after heat treatments and under
high Na+ or Ca+ concentrations, antifungal and chitosanase activities of PgChP are
independent from each other. This is in contrast to the antifungal activity of the protein
CtoA from Amycolatopsis sp. that has been attributed to the hydrolysis of chitosan in the
cell wall of the sensitive Rhizopus oryzae (Saito et al., 2009). In addition, it is unlike that
the antifungal activity of PgChP is related to the degradation of chitosan in the fungal wall
because chitosan has not been described in P. commune.
Applications of PgChP for chitosan-hydrolysis could take advantage of the thermal
stability to 60ºC and the increased activity of the protein by adding EGTA (0.1 mM),
Triton X-100 (1%) or phosphate (10 mM). Nontheless, the conditions of pH and
temperature for an optimum chitosanase activity should be determined.
The antifungal activity of the PgChP protein was not affected by high temperatures and
monovalent cations, indicating that PgChP have potential applications as a preservative in
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the food industry, particularly on substrates with a high concentration of salts. Thus, the
addition of PgChP could prevent the growth of unwanted moulds in dry-cured meat
products, where Na+ and K+ are abundant. Even though a complete study about the safety
of PgChP is necessary, the information here reported is promising for a potential use of this
protein as food preservative.
ACKNOWLEDGEMENTS
This work was supported by the Spanish Ministry of Education and Science (AGL200406546-ALI) and FEDER. Andrea Rodríguez and Raquel Acosta were the recipients of a
FPI grant from the Spanish Ministry of Education and Science.
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Allan, C.R. and Hadwiger, L.A. The fungicidal effect of chitosan on fungi of varying cell
wall composition. Exp. Mycol. 3: 285-287 (1979).
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Broekaert, W.F., Mariën, W., Terras, F.R.G., De Bolle, M.F.C., Proost, P., Van
Damme, J., Dillen, L., Claeys, M., Rees, S.B., Vanderleyden, J., and Cammue, B.P.A.
Antimicrobial peptides from Amaranthus caudatus seeds with sequence homology to the
cysteine/glycine-rich domain of chitin-binding proteins. Biochemistry. 31: 4308-4314
(1992).
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V. DISCUSIÓN
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V. DISCUSIÓN
Las proteínas PgAFP y PgChP, purificadas a partir de dos cepas de Penicillium
chrysogenum aisladas de jamón curado, mostraron actividad antifúngica en estudios
previos. Debido a la potencialidad que esta actividad presenta para su aplicación en el
control de mohos indeseables en alimentos se planteó la caracterización de dichas
proteínas.
V. 1. CARACTERIZACIÓN BIOQUÍMICA
La masa molecular de la proteína PgAFP se había estimado en torno a 9 kDa
mediante SDS-PAGE. Sin embargo, el análisis de la proteína purificada mediante
espectrometría de masas SELDI-TOF (Acosta y col., 2009b) y ESI-MS (Manuscrito 1)
reveló un peso molecular de únicamente 6,5 kDa (Figura V.1).
10000
Figura V.1. Peso molecular de PgAFP en SDS-PAGE (A) y análisis por SELDI-TOF (B;
Acosta y col., 2009b).
Esta discrepancia entre los resultados de las dos técnicas podría deberse a la
existencia de modificaciones post-traduccionales en la proteína (Pitt-Rivers y Ambesi
Impiombato, 1968; Tozzini y col., 1994; Murayama y col., 2003). Así se han descrito
migraciones anómalas de proteínas glicosiladas en SDS-PAGE atribuidas a una menor
capacidad de las mismas para unirse al SDS (Nelson, 1971; Russ y Poláková, 1973; Bagger
y col., 2007). PgAFP mostró además una reacción positiva a la tinción con ácido
periódico-Schiff (Acosta y col., 2009b), el cual oxida los grupos 1,2-glicol presentes en los
polisacáridos a aldehídos, que a su vez reaccionan con la fucsina para formar bandas de
color magenta o rosado (Zacharius y col., 1969). Por lo tanto, este resultado es indicativo
de la presencia de cadenas de glicano unidas a la proteína. Un posterior análisis de la
proteína mediante espectrometría de masas MALDI-TOF reveló la presencia de picos
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atípicos de 215 Da que no pudieron interpretarse como secuencias de aminoácidos, lo que
apoyó la hipótesis de que PgAFP tuviera algún tipo de glicosilación unida a su núcleo
proteico (Acosta, 2006). No obstante en el presente estudio, el tratamiento con una amplia
variedad de deglicosidasas no reveló ningún indicio de glicosilaciones, hecho que fue
confirmado posteriormente con la secuencia deducida a partir del gen que codifica dicha
proteína (Manuscritos 1 y 2). En consecuencia, la migración anómala mostrada por PgAFP
en los geles de poliacrilamida podría ser explicada por su alto punto isoeléctrico, estimado
en 9,2 (Acosta, 2006), que puede originar repulsiones electrostáticas que dificulten la
unión con el SDS impidiendo una correcta migración en los geles (Mondstadt y Halldorf.,
1990).
De manera similar, la proteína PgChP también mostró diferencias en el peso
molecular obtenido mediante SDS-PAGE y espectrometría de masas, aunque en esta
proteína sí pudo confirmarse la existencia de glicosilaciones (Manuscrito 4). Durante la
purificación de la proteína se procedió a su concentración, lo que permitió visualizar dos
bandas de 37 y 40 kDa en gel de poliacrilamida (Manuscrito 4, figura 1), la segunda de
ellas no detectada anteriormente (Acosta y col., 2009a). Después de tratamientos de
deglicosilación con PNGasa F se observó una única banda de 37 kDa en el gel. Estos
resultados indican que la cadena proteica posee esta masa molecular y posteriormente es
glicosilada, formándose una molécula de 40 kDa. Adicionalmente, la secuenciación
aminoacídica de ambas proteínas de 37 y 40 kDa reveló secuencias de aminoácidos
idénticas (Tabla 3, manuscrito 4), ratificando que las dos bandas observadas en los geles
correspondían a dos isoformas de la misma proteína, una de ellas glicosilada.
Posteriormente, un análisis más exhaustivo mediante espectrometría de masas con
fuente de ionización por electrospray reveló la existencia de glicosilaciones en PgChP, ya
que se detectaron seis isoformas de entre 30 y 31 kDa (Figura V.2) cuyo peso molecular
difería en múltiplos de 162 Da, masa que se corresponde con un residuo de galactosa o
manosa.
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Figura V.2. Peso molecular de las seis isoformas de PgChP determinado por ESI-MS.
La glicosilación de la proteína se confirmó posteriormente mediante el análisis de la
secuencia de aminoácidos que se deduce del gen que codifica PgChP (Manuscrito 5). Así,
la diferencia entre el peso molecular de la proteína deducida (29691 Da) y el valor
experimental de cada isoforma (30285, 30447, 30610, 30772, 30934 y 31098 Da),
corresponde a la parte glicídica de cada una de ellas, lo cual supone un nivel de
glicosilación en PgChP que va desde un 1,6 a un 4,5 % dependiendo de la isoforma.
Adicionalmente, el análisis tanto de los péptidos obtenidos tras la digestión con tripsina y
quimotripsina, como de la secuencia de aminoácidos deducida del gen que codifica PgChP,
reveló la presencia de dos secuencias consenso de glicosilación (Morelle y col., 2006) en
esta proteína: Asn-Phe-Thr (aminoácidos 230-232) y Asn-Asp-Cys (aminoácidos 288290).
La glicosilación representa la forma más común de las modificaciones posttraduccionales en proteínas tanto de eucariotas como de procariotas, donde participan en
numerosos procesos biológicos, tales como señalización inter- e intracelular, regulación de
proteínas o reconocimiento celular. Los glicanos pueden determinar la localización,
actividad y función de las proteínas que los contienen, así como influir en las
características fisicoquímicas de las mismas, tales como plegamiento, solubilidad,
resistencia a proteasas o aumento de la estabilidad a altas temperaturas (Hildegard y Geyer,
2006; Morelle y col., 2006). De hecho estos grupos glicídicos parecen estar implicados en
la actividad antifúngica de algunas proteínas purificadas a partir de plantas (Islam y col.,
2002; Deepak y col., 2003, Yamunarani y col., 2005; Kuo y col., 2008).
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Profundizando en el estudio de las secuencias de aminoácidos deducidas a partir de
los genes que codifican la producción de las proteínas PgChP y PgAFP, el análisis con el
programa SignalP (Nielsen y col., 1997a) permitió predecir la existencia de péptidos señal
o presecuencias en ambas proteínas. Estas secuencias comprenden los primeros 21
(MMTYSRLIPFALFALFGTGLA) y 18 (MQITSIAIVFFAAMGAVA) aminoácidos del
extremo N-terminal de PgChP y PgAFP, respectivamente (Manuscritos 2 y 4).
En el caso de PgAFP pudo confirmarse además la presencia de una prosecuencia
comprendida por los aminoácidos 19 a 34. Este fragmento, que difiere del péptido señal,
no está presente en la proteína madura, por lo que no pudo obtenerse mediante la
secuenciación aminoacídica de la proteína purificada a partir del medio de cultivo. Se han
descrito prosecuencias similares en proteínas con actividad antimicrobiana producidas
tanto por mohos (Oka y col., 1990; Wnendt y col., 1994; Marx y col., 1995), como por
levaduras (Tao y col., 1990; Schmitt y Breinig, 2002) y bacterias (Bormann y col., 1999),
que mantienen inactiva a la proteína en el interior de la célula. De hecho, es necesaria la
eliminación de estos fragmentos durante el proceso de secreción al medio para que la
proteína adopte una correcta estructura terciaria y adquiera su actividad (Eder y Fersht,
1995).
El análisis de las secuencias de aminoácidos deducidas permitió además estimar
tanto la masa como el pI de ambas proteínas (Gasteiger y col., 2005). Así, las masas
moleculares estimadas de PgChP y PgAFP una vez eliminados los respectivos péptidos
señal y la preprosecuencia fueron de 29691 y 6500 Da, respectivamente. Los pIs estimados
para PgChP (pH 4,8) y PgAFP (pH 8,8) fueron muy próximos a los de 5,0 y 9,2
respectivamente, determinados por electroforesis en gel de dos dimensiones (Manuscrito 4,
figura 5) y mediante electroforesis capilar de isoelectroenfoque (Acosta y col., 2009b).
Numerosos autores han relacionado el carácter catiónico con la actividad antifúngica
(Broekaert y col., 1992; Nielsen y col., 1997b; Koo y col., 1998; Hong y col., 2001), ya
que la presencia de aminoácidos de carga neta positiva sugiere la capacidad de
interaccionar con los fosfolípidos aniónicos de membrana de mohos (Sitaram y Nagaraj,
1999) pudiendo provocar alteraciones en la permeabilidad de la misma. Por lo tanto, el
punto isoeléctrico de ambas proteínas puede tener una gran relación con la actividad de las
mismas, fundamentalmente en el caso de PgChP, ya que presentan una carga neta negativa
a pH fisiológico. No obstante, la posible implicación de la carga en la actividad de ambas
proteínas se comentará más adelante.
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V. 2. CARACTERIZACIÓN GENÉTICA
Los genes que codifican la producción de PgAFP y de PgChP fueron secuenciados
mediante RACE-PCR. Los genes pgafp y pgchp, compuestos por 276 y 912 nucleótidos,
codifican los precursores de 92 y 304 aminoácidos, respectivamente (Tabla V.1). En
ambos casos el marco de lectura abierto (ORF) está interrumpido por pequeños intrones de
63 y 62 pb en pgafp, y de 60, 53, 59 y 59 pb en pgchp, típicos de genes fúngicos (Gurr y
col., 1987). Todos los intrones incluyen en los extremos 5’ y 3’ las secuencias consenso de
ensamblamiento o “consensus splice sequences” (Manuscrito 2, figura 5; Manuscrito 5,
figura 4), y muchos de ellos además contienen la secuencia interna CTRAC, siendo todas
estas secuencias típicas de genes fúngicos (Wiesner y col., 1988).
Tabla V.1. Características de los genes pgafp y pgchp.
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Uno de los factores más importantes para emplear una proteína a nivel industrial es
el bajo coste para producirla. Para identificar las condiciones de cultivo ideales y los
factores que estimulan su producción es necesario conocer los mecanismos que regulan la
expresión de los genes que la codifican. En este sentido, el estudio de la región promotora
de los genes proporciona valiosa información sobre las condiciones de expresión de las
proteínas que codifican. Así, la caracterización del promotor del gen que codifica para la
proteína PAF de P. chrysogenum permitió confirmar la presencia de cuatro sitios de unión
a CreA de represión de catabolitos de carbono y dos sitios de unión a AreA de represión de
metabolitos de nitrógeno (Marx, 2004). En este tipo de regulación, la transcripción del gen
se inhibe en presencia ciertos hidratos de carbono, fundamentalmente glucosa y fructosa
(Marx y col., 1995; Vollebregt y col., 1994), y algunos compuestos nitrogenados pueden
tanto inhibir como intensificar la producción, como es el caso de glutamina y amonio
respectivamente (Marx y col., 1995). Por otra parte, la expresión de la proteína AFP
producida por A. giganteus está regulada por otro tipo de factores ambientales como el pH
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y la presencia de fosfatos (Meyer, 2008). De hecho, el gen afp posee dos secuencias
consenso para un factor de regulación por pH (Tilburn y col., 1995) que hacen que la
producción de esta proteína se incremente cuando el moho productor se incuba en medios
alcalinos (Meyer y Stahl, 2002). La presencia de fosfatos por el contrario provoca una gran
disminución de la producción (Meyer y Stahl, 2002).
Debido a la influencia que numerosos factores ambientales pueden ejercer sobre la
expresión de PgAFP y PgChP, se hace necesario plantear nuevas investigaciones
encaminadas a la obtención de la región no codificante de pgafp y pgchp y al estudio de los
elementos de regulación presentes en los promotores, lo que permitirá optimizar las
condiciones de obtención de ambas proteínas. Por otra parte, ya se están llevando a cabo
diferentes ensayos para la expresión de ambos genes en microorganismos con mayor
velocidad de crecimiento como Escherichia coli o Pichia pastoris lo que facilitará la
obtención rápida de grandes cantidades de proteína, y de esta forma se posibilitará la
evaluación para una posible aplicación a nivel industrial. La clonación en E. coli permitirá,
en el caso de PgChP, la expresión de la proteína en un microorganismo procariota
obteniéndose una proteína recombinante sin glicosilaciones. Esta PgChP recombinante
puede resultar de gran interés para evaluar el papel que desempeñan las glicosilaciones en
la proteína y si este tipo de modificaciones post-traduccionales son necesarias para su
actividad antifúngica.
V. 3. COMPARACIÓN DE LA SECUENCIAS AMINOACÍDICAS DE PgAFP Y PgChP
CON OTRAS PROTEÍNAS
La caracterización a nivel bioquímico de PgChP y PgAFP reveló la existencia de
grandes diferencias entre ellas, aún tratándose ambas de proteínas con actividad
antifúngica producidas por P. chrysogenum. Así, en el caso de PgChP se trata de una
glicoproteína aniónica de unos 30 kDa que en principio muestra escasas características
comunes a PgAFP, de sólo 6,5 kDa y marcado carácter catiónico. El análisis de las
secuencias tanto aminoacídica como genética y su posterior comparación en bases de datos
permitió corroborar la existencia de tales diferencias. De hecho PgChP y PgAFP mostraron
semejanzas con diversas proteínas producidas por mohos con actividad quitosanasa o
antifúngica respectivamente.
PgChP reveló homologías tanto con quitosanasas fúngicas descritas como con
secuencias deducidas del genoma de distintos mohos, aunque la mayor similitud se
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encontró entre PgChP y siete quitosanasas fúngicas con los siguientes números de
adquisición en Genbank: XP_754126 (Aspergillus fumigatus), XM_001262949 (N.
fischeri), XM_001390407 (Aspergillus niger), BAD08218 (Aspergillus oryzae-1),
XP_001824257 (A. oryzae-2), XM_001209034 (Aspergillus terreus) y XP_001274664
(Aspergillus clavatus). PgChP presentó cerca del 60% de homología con las siete
proteínas, siendo la más parecida la quitosanasa CSN de A. fumigatus.
Las quitosanasas (E.C. 3.2.1.132) son glicosil hidrolasas (glicosidasas) que cortan
enlaces glicosídicos del quitosán, y son producidas por virus, bacterias y hongos (Wang y
col., 2008a, Wang y col., 2008b). En función de sus secuencias aminoacídicas se han
clasificado en cinco familias: GH-5, GH-8, GH-46, GH-75 y GH-80. Las quitosanasas
fúngicas están incluidas en la familia GH-75 (Cheng y col., 2005), compuesta
principalmente por glicosidasas producidas por mohos de los géneros Aspergillus y
Fusarium (Shimosaka y col., 1996; Cantarel y col., 2008).
Como era de esperar, la secuencia aminoacídica deducida del gen pgchp presentó
numerosas regiones comunes con las siete proteínas que mostraron mayor homología
(Manuscrito 5, figura 5), incluyendo tres regiones conservadas en quitosanasas muy
relacionadas con su actividad y dos aminoácidos, glutámico y aspártico, esenciales para la
función catalítica de este tipo de enzimas, tal como se ha descrito para la quitosanasa de F.
solani (Shimosaka y col., 2005). De la misma manera, los diez residuos de cisteína
presentes en PgChP están conservados en las siete quitosanasas del alineamiento, y
probablemente confieren estabilidad mediante la formación de enlaces disulfuro (Nakaya y
col., 1990; Marx y col., 1995; Lee y col., 1999).
Por otra parte, en la familia GH-75 existen varias hidrolasas producidas por
bacterias del género Streptomyces. PgChP mostró una homología del 25% con
quitosanasas deducidas de los genes de S. griseus y S. avermitilis. Dado que se ha descrito
que las quitosanasas bacterianas y fúngicas tienen un origen evolutivo diferente (Zhang y
col., 2000; Shimosaka y col., 2005), esa similitud se debería a la presencia de una serie de
aminoácidos de gran importancia para la estructura y la actividad catalítica del enzima
(Monzingo y col., 1996). De hecho los aminoácidos glutámico y aspártico, esenciales para
la actividad de las quitosanasas fúngicas (Shimosaka y col., 2005), están presentes también
en estas dos proteínas bacterianas (Figura V.3). Por el contrario, las tres secuencias
altamente conservadas en quitosanasas fúngicas no parecen tener relación con las
homólogas en las secuencias de las proteínas bacterianas (Figura V.3).
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PgChP
A.niger
A.oryzae-1
A.oryzae-2
A.fumigatus
S.avermitilis
S.griseus
--MMTYSRLIPFALFALFGTGLAQTVDGSKFNKPDGGPPGSYFAAGSSIPVAALQSAAAK
MAFKTTAGLA---FLALAGSVKAQSVDGSKYNSPSNGPPASYFAAATTLPVAALQSAAAK
MPIKSFASRLALS-LAICGTAMGQKVNGADYNKPDGGPPAKFFQASSSIPVAAIQAAAAK
MPIKSFASRLALS-LAICGTAMGQKVNGADYNKPDGGPPAKFFQASSSIPVAAIQAAAAK
MPSTTIIRQLAIS-LALCNSALGQVVNGADYNKPNGGPPASFFAAASTMPVAALQAAAAK
MRTRTLTLAAAAGAALLATAALPATASPSGTKAPASA------QEGS-VSAASLLAKVTS
MRTRVLAL----TSAACCAALLGAAALPASATGPGAGQEPLAAEAGAPATAAELLAEVRA
PgChP
A.niger
A.oryzae-1
A.oryzae-2
A.fumigatus
S.avermitilis
S.griseus
ARTPVPDATYPINGDNGAKKVTIHSDWAKFDKGAAYVWIADMDVDCDGIDYK-CKGNPDASSVPSKATYPVNTDDDSPKSTIHSDWVKFNQGAALSWVADMDVDCDGIDYK-CKGNGDASKVPSHATYPIG--QGSTKSTIHSDWAGFSEGAAFSFIADMDVDCDGLNHG-CKGNPDASKVPSHATYPIG--QGSTKSTIHSDWAGFSEGAAFSFIADMDVDCDGLNHG-CKGNPDATKVPSLATYPVSQDKGAAKSTIHTDWASFSEGASISWVADMDVDCDGLNSG-CQGNPDCSQI-SNGKY--KTDDETS-ATI----PVCGKNGAVFWKADMDIDCDGQVTGKCNGTTDP
CSRI-SKGAY--RTDSGASRATV----PVCGTGDAVFWKADMDIDCDGRETKACSRKTDP
PgChP
A.niger
A.oryzae-1
A.oryzae-2
A.fumigatus
S.avermitilis
S.griseus
-GQHQTNFGA-----LAAYEVPFFVIPD--RFGTKYAKQLPGNNVGAVIC----NGKMFY
-GLPETNWGA-----LSAYEVPWIVIPD--QFLTANEDLLPGNNVAAVIC----NGKMYY
-GQKETNWGA-----LSAYEVPFIVIPQ--EFLDANKGTLKGNAVAAVIC----NGKMFY
-GQKETNWGA-----LSAYEVPFIVIPQ--EFLDANKGTLKGNAVAAVICATSSNGKMFY
-GQPQTNWGA-----LSAYEVPFIVIPD--KYLSANSGALPGNNIAAVIC----NGKMFY
WFQDDTAFHQSDGKPLRADSLPYVVVPSSSSIWNYASAGIKGGGVVAVIY----NNKVEY
YFLPETAFPSSRGEPLDSAVLPHVVVPGASKVWDYRRSGLTGGSVVAVVY----RDRVRY
PgChP
A.niger
A.oryzae-1
A.oryzae-2
A.fumigatus
S.avermitilis
S.griseus
GIYGDSDGDTPQVIGEASWLMARTCFPNDDLNGNSGHGDVDVTYILFTGDDSVLPSSALN
GILGDSNGDDPEVTGEASWLMARTCFPDDDLNGAEGHAEADVTCKPYT-------RSALN
GIFGDSNGDSPQVTGEASWLMARTCFPKEDLNGNKGHTAADVTYIVFTGDKAVLPSSALN
GIFGDSNGDSPQVTGEASWLMARTCFPEEDLNGNKGHTAADVTYIVFTGDKAVLPSSALN
GILGDSNGDSPQVTGEASWLMARTCFPNEGLNGNNGHTGVDVTYIVFTGKNAVLPSSALT
AVVGDTG--PTKIIGEASYATAQALGIDPD--PETGGTDSGVTYILFKNS-QVSP----GVIGDTG--PTGIIGEASYAMAKTLGIDPD--PSTGGAESGVTYIAFKDS-RVSP-----
PgChP
A.niger
A.oryzae-1
A.oryzae-2
A.fumigatus
S.avermitilis
S.griseus
KNYVTNFTTLRSMGDKLMTALAKNLKLVDGGDGGSP--TTTAGSNPT--SGSCEWEGHCA
KNYITNFSTLRSMGDKLVGALASNLGLTSSA---------------SGSTATCSWQGHCE
KNYITNFDTLRSMGDSLVGALAKNLNLGGGGGNPPTTLTTTSIPEPTGGSGSCSWPGHCA
KNYITNFDTLRSMGDSLVGALAKNLNLGGGGGNPPTTLTTTSIPEPTGGSGSCSWPGHCA
KNYITNFTTLRSMGDKLVNALVSNLGLSGTPPTKTTTRVTTTTTKPT-SAASCSWAGHCL
---IESHSAAVSLGDSLAKQFLQNN-------------------------------------IESRARARSRGAERAREFVGR------------------------------------
PgChP
A.niger
A.oryzae-1
A.oryzae-2
A.fumigatus
S.avermitilis
S.griseus
-----GASCKDE----------------------------------------------------GAICSTEDDCSDDLVCDSGKCSSPDEDDGDDEDDEDEEENDEDDEDDDDE---------GATCSSNDDCSDDLTCQNGKCASDGSAETCSWEGHCKGATCSSNDDCSDELACIS
GFKNKGATCSSNDDCSDDLACQNGKCASDGSAETCSWEGHCKGATCSSNDDCSDELACIS
-----GASCSSDDDCADALVCTAGKCSVDGA-ATCSWEGHCEGASCSSD---------------------------------------------------------------------------------------------------------------------------------
PgChP
A.niger
A.oryzae-1
A.oryzae-2
A.fumigatus
S.avermitilis
S.griseus
--------------------------NDCSDQLVCKSGKCPSD
--------------------------DDEDDE----------GICSVDNGVETCEWEGHCEGASCSSHDDCDGNLACKNGKCSAGICSVDNGVETCEWEGHCEGASCSSHDDCDGNLACKNGKCSA--------------------------DDCSDDLYCKSGSCTAP
-------------------------------------------------------------------------------------
00
22
00
$1
22
00
$1 ,
$1
22
Figura V.3. Alineamiento de PgChP con otras quitosanasas fúngicas y bacterianas. Los
aminoácidos sombreados son comunes a todas las quitosanasas del alineamiento. Las tres
zonas altamente conservadas en quitosanasas fúngicas están englobadas en cuadros.
*: Residuos de ácido aspártico y glutámico esenciales para la función catalítica.
Números de adquisición de las proteínas: XP_001390407 (A. niger), BAD08218 (A. oryzae1), XP_001824257 (A. oryzae-2), XP_754126 (A. fumigatus), NP_823026 (S. avermitilis) y
YP_001822750 (S. griseus).
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De las quitosanasas fúngicas conocidas, sólo dos están producidas por mohos del
género Penicillium, concretamente P. islandicum y P. spinulosum (Fenton y Eveleigh,
1981; Ak y col., 1998), no habiéndose descrito hasta el momento ninguna quitosanasa
producida por P. chrysogenum. Por otra parte, aunque son frecuentes las glicosilaciones en
proteínas con actividad antifúngica purificadas a partir de plantas (Islam y col., 2002;
Deepak y col., 2003; Yamunarani y col., 2005), hasta el momento sólo se ha descrito este
tipo de modificación postraduccional en las quitosanasas de Mycobacter AL-1 (Hedges y
Wolfe, 1974) y P. islandicum (Fenton y Eveleigh, 1981).
Adicionalmente, tanto el peso molecular en torno a 30 kDa como el pI de 5
determinados para PgChP (Manuscrito 4), se encuentran dentro de los rangos descritos
para otras quitosanasas, de entre 10 y 50 kDa y pIs entre 4 y 10 (Somashekar y Joseph,
1996). La quitosanasa glicosilada con 30 kDa y pI 4,2 de P. islandicum (Fenton y
Eveleigh, 1981) es la más similar a PgChP desde el punto de vista bioquímico. No
obstante, hasta el momento se desconoce su secuencia aminoacídica, por lo que no es
posible establecer una homología entre ambas proteínas.
Por todo ello se puede afirmar que PgChP es una nueva quitosanasa de 30 kDa
producida por P. chrysogenum incluida en la familia GH-75. Además, esta proteína ha
mostrado actividad antifúngica frente a cepas de interés en alimentos (Acosta y col.,
2009a), lo cual no ha sido comprobado en otras quitosanasas.
En relación con PgAFP, el análisis de las secuencias tanto aminoacídica como
genética y su posterior comparación en bases de datos, permitió descartar la similitud con
PgChP. Así, PgAFP mostró homología con un conjunto de proteínas con actividad
antifúngica secretadas por mohos, concretamente: PAF de P. chrysogenum (Marx y col.,
1995), AFP de A. giganteus (Nakaya y col., 1990; Lacadena y col., 1995), Anafp de A.
niger (Lee y col., 1999) y AcAFP de A. clavatus (Skouri-Gargouri y Gargouri, 2008).
Estas proteínas tienen una estructura primaria diferente a la descrita en otras proteínas
antifúngicas producidas por plantas, invertebrados o anfibios, pero comparten
características comunes con ellas, como su pequeño tamaño (51-60 aminoácidos; 5,8-6,7
kDa), alto contenido en residuos de cisteína y un marcado carácter básico debido a la
presencia de lisina y arginina (Theis y Stahl, 2004). Adicionalmente, aunque hasta el
momento no se han descrito modificaciones post-traduccionales en este tipo de proteínas,
sí se han encontrado discrepancias en el peso molecular determinado mediante distintas
técnicas. Concretamente PAF corresponde a una banda de unos 12 kDa en SDS-PAGE
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(Marx y col., 1995), pero a partir de la secuencia aminoacídica deducida del gen su peso
sólo se estima en 6,3 kDa (Marx, 2004); esta diferencia de masa también se aprecia en
Anafp y AcAFP (Lee y col., 1999; Skouri-Gargouri y Gargouri, 2008). Estos resultados
son similares a los expuestos anteriormente para PgAFP, si bien estas diferencias podrían
atribuirse, como se indicó anteriormente, a su marcado carácter catiónico (Mondstadt y
Holldorf 1990).
Por lo tanto, PgAFP no sólo muestra una gran similitud en sus secuencias genética
y aminoacídica con este tipo de proteínas antifúngicas producidas por mohos, sino que
también comparte otra serie de características comunes, ya que se trata de una proteína de
58 aminoácidos, con una masa molecular de 6,5 kDa y pI de 9,2. Además, tal como se
recoge en la tabla V.2, esta proteína presenta otras características comunes, como un alto
contenido de aminoácidos básicos y seis residuos de cisteína.
Tabla V.2. Parámetros de las formas maduras de las proteínas PgAFP, Anafp, PAF, AFP, y las
deducidas de Penicillium nalgiovense (NAF), A. niger, Neosartorya fischeri y Giberella zeae.
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Los seis residuos de cisteína presentes en PgAFP están conservados en todas las
proteínas antifúngicas de mohos (Marx, 2004), aunque AFP que posee dos residuos
adicionales (Wnendt y col., 1994). De hecho, la gran estabilidad mostrada por este tipo de
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proteínas frente a proteasas, altas temperaturas o pHs extremos (Lacadena y col., 1995) se
ha atribuido a la presencia de estos residuos, que estabilizan la estructura terciaria mediante
la formación de puentes disulfuro (Campos-Olivas y col., 1995).
Con respecto a las proteínas antifúngicas producidas por mohos, sólo han sido
caracterizados los genes que codifican AFP y PAF (Wnendt y col., 1994; Marx y col.,
1995). Asimismo, se ha descrito un gen homólogo al que codifica PAF en P. nalgiovense,
denominándose la proteína deducida NAF, aunque ésta no se ha purificado (Geisen, 2000).
El análisis de las secuencias de aminoácidos revela homologías del 31 al 42% entre PAF,
AFP y Anafp, mientras que PAF es 100 % idéntica a NAF. De las proteínas descritas y
purificadas, Anafp mostró la mayor similitud con PgAFP, concretamente un 79%.
Tal como ya se ha indicado en el apartado anterior para PgAFP, tanto AFP como
PAF son sintetizadas como preproproteínas, es decir contienen una secuencia péptido señal
para su secreción y una prosecuencia que es eliminada antes o durante su liberación al
medio (Marx, 2004). Con respecto a Anafp, aunque sólo se ha determinado la secuencia
aminoacídica de la proteína madura (Lee y col., 1999), se asume por su gran similitud con
el resto que también está codificada como preproproteína (Marx, 2004). En relación a la
proteína AcAFP sólo se han descrito los 21 primeros aminoácidos, de los cuales 20 y 12
residuos coinciden con la región amino terminal de AFP y PAF respectivamente.
En los últimos años, gracias a la secuenciación del genoma de diferentes mohos, se
han ido recogiendo en bases de datos proteínas deducidas que muestran similitud con este
grupo de proteínas antifúngicas. De hecho PgAFP, al igual que PAF, AFP y Anafp, ha
mostrado homologías con proteínas deducidas de N. fischeri y G. zeae (Manuscrito 2,
figura 6). Una de estas proteínas deducidas, concretamente del genoma de A. niger, ha
mostrado junto a Anafp una alta similitud con PgAFP, con un 75 y 79 % de homología,
respectivamente. Por otra parte, a pesar de que tanto PgAFP como PAF son producidas por
cepas de P. chrysogenum sólo mostraron un 43 % de homología, por lo que se trata de
proteínas diferentes. Por último, las secuencias que codifican las proteínas PgAFP y PgChP
se han localizado en el genoma de P. chrysogenum Wisconsin 54-1255 cuya secuencia
completa ha sido determinada recientemente (Van den Berg et al., 2008).
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V. 4. MECANISMO DE ACCIÓN DE LA ACTIVIDAD ANTIFÚNGICA DE PgAFP Y
PgChP
Conocer el mecanismo de acción de proteínas antifúngicas como PgChP y PgAFP
puede ser de gran utilidad para el desarrollo de nuevas estrategias antimicrobianas y para
determinar su posible aplicación en determinados productos.
Los mecanismos de acción de estas proteínas son tan variados como los organismos
que las producen, e incluyen degradación de polímeros de las paredes celulares fúngicas,
formación de canales o poros en la membrana, inactivación de ribosomas, alteración de la
síntesis de ADN o inhibición del ciclo celular (Selitrennikoff, 2001). No obstante, aún se
desconoce el modelo de acción de muchos de estos compuestos.
En cualquier caso, las proteínas antifúngicas de mayor utilidad son las que
presentan mecanismos de acción específicos para las membranas de los hongos (Groll y
col., 1998), ya que suelen ser poco tóxicas para las células de mamíferos. Los factores que
desempeñan un papel más destacado en la interacción del péptido con la membrana son la
carga (Hoover y col., 2003) y la hidrofobicidad del compuesto (Friedrich y col., 1999;
Hancock y Chapple, 1999). De hecho, el carácter catiónico se ha relacionado de manera
directa con la actividad antifúngica (Broekaert y col., 1992; Nielsen y col., 1997b; Koo y
col., 1998; Hong y col., 2001). La base de la especificidad de este tipo de proteínas
catiónicas es la composición lipídica de la membrana, ya que al estar cargadas
positivamente pueden interaccionar con los fosfolípidos aniónicos (Sitaram y Nagaraj,
1999) presentes en las membranas de bacterias y hongos. Por el contrario, en el caso de los
eucariotas superiores, esta unión es muy débil, ya que sus membranas están compuestas
principalmente por fosfolípidos sin cargas negativas (Theis y Stahl, 2004). El hecho de que
tanto PgChP como PgAFP muestren actividad antifúngica pero no produzcan hemólisis de
glóbulos rojos de mamíferos, puede apuntar a un mecanismo de acción específico para las
membranas de mohos.
Aunque la composición de la pared celular fúngica varía en función del género y la
especie, los polímeros más frecuentes son quitina, 1,3-glucano y 1,6-glucano. Otros
polímeros como el quitosán sólo se han descrito en un limitado número de hongos de los
géneros Rhizopus, Absidia, Fusarium o Mucor (Price y Storck, 1975; Alfonso y col., 1995;
Gao y col., 1995). Se ha atribuido un papel de defensa a las quitosanasas por su capacidad
de hidrolizar quitosán desestabilizando la pared celular (Somashekar y Joseph, 1996), y en
algunos casos se ha relacionado directamente la actividad antifúngica con la actividad
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quitosanasa (Saito y col., 2009). Sin embargo, distintos compuestos y tratamientos
térmicos provocaron efectos diferentes sobre la actividad quitosanasa y antifúngica de la
proteína PgChP, lo que podría significar que la actividad antifúngica no se debe a la
degradación del quitosán. Además, PgChP mostró actividad frente a P. commune, especie
en la que hasta el momento no se ha descrito la presencia de quitosán.
Por otra parte, tanto PgChP como PgAFP poseen carga neta positiva al pH de 4,5
empleado para evaluar la actividad antifúngica. Esta característica, junto a la capacidad de
formar estructuras anfipáticas, es común a todas las proteínas que actúan sobre las
membranas microbianas, aun presentando éstas estructuras primarias muy diversas y
secundarias tanto en hoja- como en α-hélice (Tossi y col., 2000). Se ha descrito un
mecanismo de acción para algunas de estas proteínas, el modelo Shai-Mat-suzaki-Huang
(SMH), que propone la interacción de éstas con los fosfolípidos de carga neta negativa
presentes en la membrana plasmática de bacterias y hongos provocando la formación de
poros en la misma (Matsuaki, 1999; Shai, 1999; Yang y col., 2000). Entre estas proteínas
se encuentran algunas defensinas, que son un grupo de proteínas y péptidos
antimicrobianos producidos por mamíferos (Ganz y col., 1990), insectos (Lamberty y col.,
1999) o plantas (Broekaert y col., 1995), muy similares a las proteínas antifúngicas
producidas por mohos en cuanto a su pequeño tamaño, carácter catiónico y presencia de
puentes disulfuro en su estructura. Sin embargo las defensinas producidas por mamíferos e
insectos, de menor tamaño (3 a 5 kDa), forman canales iónicos en la membrana plasmática
provocando la pérdida de minerales y metabolitos esenciales (Lehrer y col., 1989; White y
col., 1995). Adicionalmente, algunas defensinas de plantas producen una desporalización
de la membrana como consecuencia de su unión a los fosfolípidos aniónicos (Thevissen y
col., 1999). No obstante, PgAFP no presenta actividad frente a bacterias y levaduras
(Acosta y col., 2009b). Estos datos coinciden con lo descrito hasta ahora para las proteínas
antifúngicas más similares a PgAFP, como PAF (Kaiserer y col., 2003), AFP (Lacadena y
col., 1995; Theis y col., 2003) y AcAFP (Skouri-Gargouri y Gargouri, 2008). Anafp en
cambio, es la única proteína de este grupo activa frente a levaduras (Lee y col., 1999), si
bien hasta el momento se desconoce su mecanismo de acción. La mayoría de estas
proteínas además de no mostrar actividad frente a levaduras y bacterias, manifiestan
diferentes espectros de inhibición frente a mohos (Marx, 2004). Por ello, tal como se ha
descrito para algunas defensinas de plantas (Thevissen y col., 1997; Thomma y col., 2002),
se ha propuesto un mecanismo de acción basado en la existencia de receptores específicos
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para cada proteína en las membranas plasmáticas de los mohos sensibles (Marx y col.,
2007; Meyer, 2008). Esta idea se ve apoyada por el hecho de que la actividad de PAF y
AFP es fuertemente inhibida en presencia de Mg2+ y K+ (Theis y col., 2003) y Ca2+, Na+ y
K+ (Kaiserer y col., 2003), respectivamente, ya que estos cationes podrían saturar los
receptores existentes en las membranas de los mohos sensibles impidiendo la unión de las
proteínas antifúngicas y con ello su actuación (Theis y col., 2003).
Al igual que ha sido descrito para PAF y AFP, tanto PgChP como PgAFP han
mostrado diferente sensibilidad a los cationes presentes en el medio. Así, aunque altas
concentraciones de Mg2+ y Ca2+ inhiben la actividad antifúngica de ambas proteínas, los
cationes monovalentes Na+ y K+ únicamente afectan a la actividad de PgAFP (Manuscritos
3 y 6). Concretamente, al combinar una pequeña cantidad de proteína (3µg/ml) con una
alta concentración de iones (100 mM) se aprecia una pérdida de actividad, aunque sólo
frente a A. niger. Por lo tanto, el mecanismo de acción de PgChP y PgAFP podría basarse
en la existencia de receptores específicos en las membranas de los mohos sensibles,
ausentes en levaduras o bacterias. Estos receptores tendrían mayor afinidad por los
cationes divalentes y además diferirían en función del moho sensible, siendo por ejemplo
los receptores para PgAFP en A. niger más afines a Na+ y K+ que los presentes en P.
restrictum.
Por otra parte, aunque el mecanismo de acción de PAF y AFP podría estar mediado
por la presencia de receptores específicos en la membrana, ambas proteínas ejercen efectos
diferentes sobre los mohos sensibles. Concretamente, la unión de PAF con el receptor
específico incrementa la permeabilidad al K+, provocando una hiperpolarización de la
membrana que conlleva la entrada de la proteína mediante endocitosis. A partir de ese
momento se activan distintos mecanismos, como la producción de especies reactivas de
oxígeno o mitoptosis (formación de poros en las membranas mitocondriales), que
ocasionan la muerte celular programada del moho sensible (Marx y col., 2007). A pesar de
las similitudes con PAF, el efecto de AFP es diferente, ya que sólo los mohos resistentes
son capaces de introducirla por endocitosis posiblemente para una degradación con fines
nutricionales. La unión de AFP a los receptores específicos en los mohos sensibles,
provoca tanto alteraciones en la membrana como la inhibición de la enzima quitín sintasa
que interviene en la biosíntesis de la pared celular fúngica (Hagen y col., 2007; Meyer,
2008). La proteína AFP posee además en su extremo amino terminal la secuencia de unión
a la quitina CKYKAQ (Hagen y col., 2007), no descrita en PAF, Anafp, PgAFP, ni PgChP.
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Por todo ello, aunque la información disponible sugiere que el mecanismo de
acción de PgAFP y PgChP está basado en la interacción específica con receptores
presentes en las membranas fúngicas, sería necesario confirmarlo, y en su caso estudiar
tanto los receptores implicados, como el efecto que las proteínas ejercen tras la unión con
los mismos.
V. 5. APLICACIONES DE LAS CEPAS DE P. chrysogenum Y LAS PROTEÍNAS PgAFP Y
PgChP
Como ya se ha descrito en esta memoria, las condiciones ecológicas que ofrecen los
productos cárnicos madurados favorecen el desarrollo de una población fúngica
superficial, que aunque puede contribuir a la maduración (Martín y col., 2004, 2006),
también puede originar defectos o alteraciones (Sanz, 1986; Spotti y col., 1988) e incluso
producir micotoxinas (Núñez y col., 1996, 2007; López-Díaz y col., 2001; Sabater-Vilar y
col., 2003). Para controlar la población fúngica podrían utilizarse cultivos iniciadores que
al implantarse en el producto fuesen capaces de inhibir por competición a los mohos no
deseados. No obstante, la evolución de las condiciones ecológicas a lo largo de la
maduración puede hacer que los cultivos terminen siendo desplazados por otras cepas
mejor adaptadas. En cambio para los aislamientos de P. chrysogenum RP42C y AS51D, la
producción de las proteínas PgAFP y PgChP con actividad antifúngica, puede suponer una
ventaja adicional para competir frente al resto de mohos. De hecho, tanto PgAFP como
PgChP se purificaron como resultado de la búsqueda de mohos productores de proteínas
antifúngicas para su utilización como cultivos protectores en productos cárnicos
madurados (Acosta, 2006). Además, ambas cepas han sido aisladas de jamón curado, por
lo que se trata de mohos adaptados a las condiciones de temperatura, actividad de agua y
pH que se dan a lo largo del proceso de maduración. Adicionalmente, P. chrysogenum ha
sido propuesto como cultivo iniciador para la elaboración de productos madurados por su
contribución a la formación de sabor y aroma en el producto acabado (Martín y col., 2004,
2006; Benito y col., 2005). Dado que P. chrysogenum RP42C produce proteína activa en
condiciones similares a las de las etapas iniciales de la maduración (Acosta, 2006), será de
gran interés confirmar la producción tanto de PgAFP como de PgChP en productos
madurados. Además, el amplio espectro de inhibición mostrado fundamentalmente por P.
chrysogenum RP42C frente a mohos toxigénicos de interés en estos productos, como A.
flavus, A. parasiticus o P. commune, permite ser optimistas respecto a su utilización como
cultivo protector para garantizar la seguridad alimentaria en este tipo de alimentos.
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No obstante, también podrían emplearse ambas proteínas purificadas en productos
en los que no sea deseable el desarrollo de mohos. Además PgChP, debido a su actividad
quitosanasa, podría también emplearse para la producción de oligómeros de quitosán a
nivel industrial.
De forma general y como paso previo a la aplicación de una proteína a nivel
industrial, resulta de gran importancia garantizar que la actividad antifúngica se mantiene
en presencia de determinados compuestos que puedan estar presentes en el producto donde
se vaya a aplicar (Theis y Stahl, 2004). Adicionalmente el efecto de ciertos tratamientos a
temperaturas o pHs extremos puede condicionar también sus posibles aplicaciones. Por
otra parte, la posibilidad de incrementar la actividad de estas proteínas mediante la adición
de determinados sustratos es otro factor a tener en cuenta antes de proponer futuras
aplicaciones.
V. 5. 1. APLICACIÓN DE PgChP Y PgAFP COMO AGENTES ANTIFÚNGICOS
Como se ha comentado anteriormente, no sólo resulta de gran interés la posibilidad
de utilizar mohos productores de proteínas antifúngicas como cultivos protectores en
productos madurados, sino que también puede ser útil el uso de PgAFP y PgChP como
aditivo en alimentos en los que no sea deseable el desarrollo del moho productor.
El uso de proteínas antimicrobianas como protectores en alimentos no es un
concepto nuevo. Concretamente la nisina, bacteriocina producida por varias cepas de
Lactococcus lactis, ha sido aprobada como GRAS (“Generally Regarded as Safe”) en ocho
países de la Unión Europea. Esta proteína es ampliamente utilizada en productos lácteos,
vegetales, carne o pescado (Delves-Broughton y col., 1996) para evitar el desarrollo de
bacterias patógenas gram positivas como Bacillus cereus, Listeria monocytogenes o
determinadas especies de Enterococcus, Streptococcus y Staphylococcus (Lubelski y col.,
2008). Además, la nisina ha sido comercializada para el tratamiento de infecciones
gástricas causadas por Helicobacter spp. y algunas de sus variantes están siendo probadas
en ensayos clínicos para tratar enfermedades producidas por enterococos resistentes a la
vancomicina (Reddy y col., 2004).
No obstante, antes de proponer a PgAFP y PgChP para su utilización como
compuestos antifúngicos en alimentos se deben evaluar los posibles efectos indeseables
que pudieran ocasionar en los consumidores para evitar cualquier peligro de tipo sanitario.
Por otra parte, la estabilidad de las mismas tras el tratamiento a diferentes temperaturas y
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pHs o en presencia de determinados compuestos que se encuentren en los sustratos donde
se pretendan utilizar, condicionará en gran medida sus posibles aplicaciones.
Como se ha mencionado, tanto PgChP como PgAFP fueron inicialmente
purificadas mediante cromatografía de intercambio catiónico (Acosta y col, 2009a) con el
objeto de obtener proteínas con carácter básico, ya que esta característica está relacionada
con una mayor actividad antifúngica y una menor toxicidad para las células de mamíferos
(Hong y col., 2001). No obstante, si bien PgAFP mostró un pI de 9,2 (Acosta y col.,
2009b), PgChP mostró un pI de 5 (Manuscrito 4, figura 6), lo que supone que esta proteína
no es catiónica a pH fisiológico.
La ausencia de actividad hemolítica de PgChP y PgAFP frente a glóbulos rojos
(Manuscritos 3 y 6), y en el caso de PgAFP también de toxicidad para larvas de Artemia
salina (Manuscrito 3), permite ser optimistas hasta el momento con respecto a la inocuidad
de estas proteínas, ya que estas técnicas son comúnmente empleadas para chequear la
toxicidad de nuevas sustancias (Van’t Hof y col., 2001; Theis y Stahl, 2004; Milhem y col.,
2008; Chohan, 2009).
La ausencia de efectos negativos podría permitir la utilización de PgAFP y PgChP
en diversos campos de aplicación, si bien el presente estudio se ha enfocado hacia una
posible aplicación en alimentos. Por ello, se evaluó el efecto sobre la actividad antifúngica
tanto de diferentes tratamientos como de diversos compuestos comúnmente encontrados en
alimentos, como sales, detergentes o fosfatos (Tabla V.3; Manuscritos 3 y 6).
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Tabla V.3. Resistencia de PgAFP y PgChP a tratamientos térmicos, distintos pHs, cationes, fosfato
sódico, quelantes y detergentes en las proteínas, expresada como los valores máximos a los que se
mantiene actividad antifúngica.
D
E: /
F
G
$
= E: /
$
'
HI
7
I
I
:
G
I
G
>
/
/
>
7 F
G
J
J
=
"$!
@G
L L
M& '
5$&14$&
G
K
GGG
K
GGG
GGG'
GGG
&
GGG
GGG
GGG
a
: Máxima concentración empleada 100 mM.
: Máxima concentración empleada 1 mM.
c
: Máxima concentración empleada 1 %.
d
---: Pérdida de la actividad antifúngica de la proteína.
b
n.c.: Afectó al crecimiento del moho y su influencia no pudo determinarse.
n.d.: No determinado.
Muchos productos alimentarios se someten de forma rutinaria a tratamientos
térmicos, ya sea como consecuencia del propio proceso de elaboración o con el objeto de
garantizar su estabilidad microbiológica, lo que condiciona en gran medida la aplicación de
ambas proteínas. En este sentido, tanto PgChP como PgAFP mostraron actividad
antifúngica incluso tras tratamientos térmicos a 80 y 100ºC, respectivamente.
Dado que concentraciones de Ca2+ de 10 a 100 mM pueden inhibir la actividad de
estas proteínas, su uso se vería limitado en alimentos ricos en calcio como el queso. Si
embargo, la actividad de PgAFP y PgChP se mantuvo en presencia de altas
concentraciones de Na+ y K+, lo que puede ser de gran interés en productos cárnicos
madurados, donde abunda el cloruro sódico y puede haber otras sales de sodio y potasio.
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Además, estas proteínas muestran una mayor resistencia a estos iones que otras como PAF
o AFP, cuya actividad ya se reduce en presencia de 40-60 mM de Na+ (Kaiserer y col.,
2003) o Na+ y K+ (Theis y col., 2003), respectivamente.
Otras sustancias como el EDTA y los fosfatos se utilizan como aditivos, entre otras
razones, por su capacidad de secuestrar metales en alimentos no ácidos que requieren
aditivos sin sabor (Calvo, 1991). Si bien los quelantes no tuvieron un gran impacto en la
actividad antifúngica de PgChP y PgAFP, ésta se redujo en presencia de una alta
concentración (100 mM) de fosfato, aunque esta disminución, en el caso de PgAFP,
depende de la concentración de la proteína y del moho sensible utilizado.
El uso de detergentes no iónicos como emulsionantes también está extendido en la
industria alimentaria. De hecho, en España se emplean en confitería, repostería y para la
elaboración de galletas (Calvo, 1991), si bien de los detergentes evaluados en este estudio
sólo tween 20 y tween 80 están autorizados en alimentos (E-432 y E-433,
respectivamente). Los detergentes inhibieron la actividad antifúngica de PgChP, pero
PgAFP se mantuvo activa con un 1% de tween 20 o de tween 80. Dado que esta
concentración supera los niveles habituales en los alimentos de 0,1-0,3% (R.D. 142/2002),
la presencia de estos aditivos no afectaría a la actividad de PgAFP.
Además de la utilización de PgAFP y PgChP de forma individual, la combinación
de ambas o con algún tratamiento físico o químico podría suponer un efecto sinérgico que
incrementara la actividad antifúngica en los alimentos. Adicionalmente, aunque PgAFP
haya mostrado un espectro de inhibición muy amplio (Acosta y col, 2009b), el uso
combinado con PgChP podría ampliar la actividad a mohos que posean quitosán en la
pared celular como Mucor o Rhizopus (Shimosaka y col., 2005; Somashekar y Joseph,
1996), géneros alterantes en productos cárnicos madurados.
Por otra parte, la estabilidad mostrada por ambas proteínas al tratamiento con
temperaturas y pHs extremos abre un gran abanico de posibilidades respecto de su
combinación con diferentes tratamientos físicos o químicos. La combinación de
compuestos antimicrobianos con tratamientos térmicos, químicos o físicos como las altas
presiones o los campos eléctricos pulsados, está siendo ampliamente estudiada. Así, la
nisina ha sido combinada con lisozima y campos eléctricos pulsados (Sobrino-López y
Martín-Belloso, 2008), con tratamientos de frío (Cao-Hoang y col., 2008), con calor y
acidificación (Naim y col., 2008), o con ácido láctico y monolaurina (Tokarsky y Marshall,
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2008) para controlar diversos patógenos en alimentos. Combinaciones similares no sólo
pueden suponer un incremento en la actividad de PgAFP y PgChP, sino que también
podrían modificar su especificidad. De hecho la proteína antifúngica cecropina B, que por
sí sola no tiene efecto frente a E. coli, en combinación con lisozima es eficaz para combatir
este microorganismo (Casteels y col., 1993).
Por todo lo anterior, PgChP y PgAFP parecen tener características idóneas para ser
empleadas como agentes protectores en diversos alimentos que requieran un control frente
a mohos alterantes o toxigénicos. No es previsible que la utilización de estas proteínas
como aditivo provoque alteraciones en la flora intestinal beneficiosa del consumidor ya
que ambas poseen un mecanismo de acción específico frente a mohos. Por otra parte,
estudios realizados con PgAFP muestran la sensibilidad de esta proteína a tripsina, pepsina
y lisozima (Acosta, 2006), por lo que podría ser degradada en el tracto gastrointestinal.
No obstante, la aplicación de PgAFP y PgChP no tiene por qué estar limitada a un
uso en alimentos. De hecho se ha propuesto la utilización de proteínas antifúngicas
similares en otros campos como la agricultura o en terapéutica humana (Marx y col., 2007;
Meyer, 2008). En este sentido, la incorporación de genes que codifican proteínas
antifúngicas ha permitido la obtención de cultivos transgénicos resistentes a plagas de
hongos (Campo y col., 2008; Swathi-Anyradha y col., 2008) y ciertas proteínas
antifúngicas están siendo ya estudiadas para su uso como agentes farmacológicos
(Cappelletty y Eiselstein-McHitrick, 2007; Wiederhold y Lewis, 2007; Fortún y col.,
2009).
V. 5. 2. APLICACIÓN DE PgChP COMO QUITOSANASA
Dado que la caracterización aminoacídica de PgChP reveló que pertenecía a la
familia G-75 de quitosanasas, fue necesario comprobar su capacidad para hidrolizar
quitosán. Aunque algunas quitosanasas pueden hidrolizar polisacáridos distintos al
quitosán (Pelletier y Sygusch, 1990; Shimosaka y col., 1995; Kimoto y col., 2002), PgChP
no presentó indicios de degradación de la quitina (Manuscrito 6), como suele ser habitual
entre las quitosanasas (Fukamizo y Brzezinski, 1997; Su y col., 2006).
La quitina es el componente estructural mayoritario del exoesqueleto de
invertebrados y de las paredes celulares fúngicas (Lower, 1984; Ornum, 1992). Debido a
que la degradación de la quitina es muy lenta, la acumulación de desechos resultantes del
procesado de crustáceos es un problema emergente en la industria de productos de la pesca
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(Shahidi y Synowiecki, 1991). No obstante, estos desechos pueden ser empleados para la
producción de sustancias muy beneficiosas a nivel industrial, como son la quitina, el
quitosán, o los productos de la degradación de los mismos. De hecho, los oligómeros
resultantes de la degradación del quitosán tienen múltiples aplicaciones: como inhibidores
de la producción de micotoxinas (Cuero y col., 1991), purificadores de agua (Jeuniaux,
1986; Muzzarelli y col., 1989), neutralizantes de acidez en zumos de fruta y vino (Chen y
Li, 1996; Rwan y Wu, 1996; Spagna y col., 1996), e incluso como agentes antitumorales y
potenciadores del sistema inmune (Pae y col., 2001). Adicionalmente, y como resultado de
la búsqueda de nuevos antimicrobianos naturales, los oligómeros del quitosán han
mostrado actividad antimicrobiana frente a bacterias, levaduras y mohos (Yalpani y col.,
1992).
El quitosán puede ser degradado mediante hidrólisis química o enzimática,
obteniéndose oligómeros
de menor
(monómeros-trímeros) o
mayor
grado
de
polimerización (pentámeros-heptámeros), respectivamente. No obstante, los oligómeros
resultantes de una hidrólisis enzimática con quitosanasas muestran mejores características
funcionales (Kendra y Hadwiger, 1984; Uchida y col., 1988), lo que ha incrementado el
interés por este tipo de enzimas.
Dado que las quitosanasas difieren en sus propiedades enzimáticas (Wang y col.,
2008b), es necesario conocer el efecto sobre PgChP de diversos factores como iones o
detergentes para potenciar su actividad.
Los iones Ca2+ y K+ no tuvieron efecto alguno sobre la actividad quitosanasa de
PgChP a concentraciones de 100 mM, mientras que Mg2+ y Na+ sí influyeron en la
actividad por encima de 10 mM. En el caso de los quelantes, aunque la actividad se redujo
en presencia de una alta concentración (1 mM) de EDTA, con EGTA se llegó a triplicar la
cantidad de glucosamina liberada (Manuscrito 6). La influencia de altas concentraciones de
Mg2+ y Na+ sobre PgChP puede deberse a que los iones afecten a la estructura de la
proteína impidiendo su actividad, tal y como ha sido descrito para otras quitosanasas de
mohos (Wang y col., 2008b). De hecho, el tratamiento con SDS y ácido deoxicólico,
detergentes iónicos con conocido efecto desnaturalizante, produjo una inhibición de la
actividad de PgChP. Por el contrario los detergentes no iónicos, fundamentalmente el
Tritón X-100, favorecieron la actividad de la proteína. De forma similar, en presencia de
fosfato (10 mM) se duplicó la cantidad de glucosamina liberada.
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En conclusión, la producción de oligómeros de quitosán mediante hidrólisis con
PgChP puede incrementarse en presencia de EGTA, Tritón X-100 y fosfato, siendo las
concentraciones más apropiadas 0,1 mM, 1 % y 10 mM, respectivamente. La proteína
podría además ser empleada en presencia de altos niveles Ca2+ y K+.
No obstante, la actividad quitosanasa de PgChP debe ser evaluada a diferentes
temperaturas y pHs con el fin de encontrar las condiciones óptimas para la degradación del
quitosán.
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VI. CONCLUSIONES
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CONCLUSIONES
VI. CONCLUSIONES
1. PgAFP es una nueva proteína antifúngica de 6,5 kDa, en la que no se han detectado
glicosilaciones. De acuerdo con su secuencia primaria, pertenece a un grupo de
proteínas antifúngicas de pequeño tamaño, básicas y con alto contenido en cisteína,
que son producidas por mohos.
2. La proteína antifúngica PgChP presenta diversas isoformas glicosiladas con pesos
moleculares comprendidos entre 30 y 31 kDa y, de acuerdo con su secuencia
aminoacídica y su actividad enzimática, es una nueva quitosanasa perteneciente a la
familia GH-75, mayoritariamente compuesta por quitosanasas fúngicas.
3. Las secuencias aminoacídicas deducidas de los genes que codifican estas dos proteínas
revelan la existencia de un péptido señal en ambas y una prosecuencia en PgAFP que
no están presentes en las proteínas purificadas.
4. La ausencia de actividad hemolítica y de toxicidad frente a larvas de Artemia salina,
junto con la reconocida inocuidad de P. chrysogenum, permite plantear la aplicación de
estas dos proteínas en alimentos.
5. Ambas proteínas antifúngicas muestran una gran estabilidad a tratamientos térmicos, y
mantienen su actividad en presencia de quelantes o fosfato sódico a las concentraciones
que pueden encontrarse en alimentos.
6. La actividad antifúngica de las dos proteínas sólo se ve limitada en presencia de altas
concentraciones de iones divalentes o de detergentes iónicos, si bien los detergentes no
iónicos afectan a la proteína PgChP. Por lo tanto, estas proteínas pueden ser de utilidad
en un amplio rango de alimentos.
7. La actividad quitosanasa de PgChP es independiente de la actividad antifúngica y no se
ve limitada por altas concentraciones de iones de calcio y detergentes no iónicos.
8. Las proteínas PgAFP y PgChP muestran características adecuadas para controlar el
crecimiento de mohos indeseables en alimentos.
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CONCLUSIONES
VI. CONCLUSIONS
1. PgAFP is a novel antifungal protein of 6.5 kDa with no glycosylations. According to its
primary sequence, it belongs to a group of small, basic, and cystein-rich antifungal
proteins produced by moulds.
2. The antifungal protein PgChP has several glycosylated isoforms with molecular
weights ranging from 30 to 31 kDa. According to primary sequence and enzymatic
activity, it is a novel chitosanase belonging to the GH-75 family, mainly composed by
fungal chitosanases.
3. The putative amino acid sequences reveal a signal peptide in both proteins and one
prosequence in PgAFP that are not found in the purified proteins.
4. The lack of haemolytic activity and toxicity against Artemia salina larvaes together
with the safety granted to P. chrysogenum support the potential application of these
two proteins in foods.
5. Both antifungal proteins show a high thermal stability and keep their activity with
chelators or sodium phosphate at the concentrations found in foods.
6. The antifungal activity of the two proteins is only restricted by high concentrations of
divalent ions or ionic detergents, whereas non-ionic detergents only affect to PgChP.
Therefore, these two proteins can be of use in a wide range of foods.
7. The chitosanase activity of PgChP is not related to the antifungal activity, and it is
restricted by neither high calcium ion concentrations nor non-ionic detergents.
8. Both proteins show appropriate characteristics to control growth of unwanted moulds
in foods.
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BIBLIOGRAFÍA
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VIII. RESUMEN
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RESUMEN
VIII. RESUMEN
Las condiciones ambientales en que se procesan o se almacenan diversos tipos de
alimentos favorecen el desarrollo en su superficie de mohos que tienen la capacidad de
adaptarse y desarrollarse en sustratos muy diversos. En productos cárnicos madurados y
quesos es frecuente el desarrollo una población fúngica superficial que puede tener efectos
deseables, ya que contribuyen en gran medida a la formación del sabor, aroma o textura del
producto acabado. Pero el desarrollo de mohos no siempre es deseable en alimentos.
Muchos mohos son alterantes, llevando al deterioro de las características sensoriales de los
alimentos. No obstante, el aspecto más preocupante respecto del desarrollo de una
población fúngica en alimentos es su capacidad para producir micotoxinas.
Existen numerosos mecanismos para controlar el desarrollo microbiano en distintos
tipos de alimentos, incluyendo métodos físicos y químicos. Estos tratamientos resultan
eficaces para evitar el desarrollo de mohos en determinados alimentos, pero no son
adecuados para otros productos como los madurados, donde la presencia de ciertos mohos
es beneficiosa y además la aplicación de fungicidas está limitada. Una posible solución en
este tipo de alimentos sería el empleo de proteínas antifúngicas que evitaran el desarrollo
de mohos indeseables y que a la vez permitieran el crecimiento de mohos beneficiosos en
los productos madurados.
En estudios previos se seleccionaron dos cepas no toxigénicas de P. chrysogenum,
RP42C y AS51D, procedentes de jamón curado, por su capacidad para inhibir el desarrollo
de mohos toxigénicos frecuentes en alimentos. Además, se purificaron dos proteínas con
actividad antifúngica, PgAFP y PgChP a partir de RP42C y AS51D, respectivamente.
El objetivo fundamental de este trabajo ha sido la caracterización de las dos
proteínas PgAFP y PgChP. Esta caracterización se ha basado en: la determinación de sus
secuencias aminoacídicas, la obtención de las secuencias de los genes que las codifican; la
evaluación preliminar de su toxicidad; el estudio de la influencia de diversos tratamientos
sobre la estabilidad de las proteínas y el efecto sobre la actividad antifúngica de
compuestos comúnmente encontrados en alimentos.
Pese a que tanto PgAFP como PgChP son proteínas con efecto antifúngico, han
resultado ser muy diferentes en cuanto a sus pesos moleculares, puntos isoeléctricos y
secuencias aminoacídicas. PgAFP tiene un peso molecular de 6,5 kDa, un punto
isoeléctrico determinado previamente de 9,2, y no posee glicosilaciones pese a que
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estudios preliminares apuntaban a que presentaba esta modificación. Este hecho se
confirmó con la obtención de la secuencia del gen que codifica la proteína y la
consiguiente determinación de la secuencia aminoacídica deducida. PgChP sin embargo,
mostró tener seis isoformas glicosiladas con pesos moleculares comprendidos entre 30 y 31
kDa, separados por una masa de 162 Da que podría corresponder a un residuo de manosa o
galactosa. Además, la secuencia deducida del gen permitió encontrar dos posibles sitios de
glicosilación (Asn-Phe-Thr y Asn-Asp-Cys) y la presencia de cadenas glucídicas se
demostró mediante deglicosilaciones enzimáticas. PgChP además presentó un punto
isoeléctrico cercano a 5, por lo que la proteína no es catiónica.
Las secuencias obtenidas de los genes pgafp y pgchp que codifican PgAFP y
PgChP tuvieron un tamaño de 276 y 912 pb, que codifican 92 y 304 aminoácidos,
respectivamente. Además, estos genes presentaron varios intrones pequeños con
marcadores típicos de intrones fúngicos.
La obtención de las secuencias aminoacídicas permitió encontrar proteínas con
cierta similitud en su estructura primaria. Así, PgAFP presentó un 79% de homología con
la proteína Anafp de A. niger y PgChP resultó tener cerca del 60% de similitud con la
familia de hidrolasas GH-75 compuesta por quitosanasas principalmente fúngicas. Tanto
PgAFP como PgChP presentan en su secuencia aminoacídica deducida una parte que
corresponde a un péptido señal necesario para que ambas proteínas sean secretadas al
medio extracelular. Además PgAFP, al igual que Anafp y otras proteínas con estructura
primaria parecida, presenta una prosecuencia, que parece mantener la proteína inactiva
para no causar daño a su moho productor, hasta que este fragmento es eliminado durante su
secreción.
La actividad antifúngica de PgAFP y de PgChP se inhibe por altas concentraciones
(100 mM) de ciertos cationes, especialmente los divalentes Ca2+ y Mg2+. Al igual que lo
propuesto para otras proteínas como las defensinas, el mecanismo de acción de PgChP y
PgAFP podría basarse en la existencia de receptores específicos en las membranas de los
mohos sensibles y ausentes en los mohos resistentes. Los cationes podrían saturar estos
receptores específicos e impedir que las proteínas se unieran a ellos y ejercieran su acción
antifúngica.
PgChP muestra actividad hidrolítica sobre quitosán, lo que junto a la homología de
su secuencia aminoacídica con otras quitosanasas fúngicas confirma que pertenece a este
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grupo de enzimas. Esta capacidad de PgChP para degradar quitosán puede hacer que su
espectro de inhibición incluya a hongos con quitosán en su pared celular, como los géneros
Rhizopus y Mucor, alterantes habituales en alimentos madurados. Además, las
quitosanasas son utilizadas en la industria para la degradación de caparazones de moluscos,
así como para la obtención de polímeros de quitosán, de alto valor como protectores frente
a patógenos en alimentos, inhibidores de micotoxinas, purificadores de agua, etc.
Por otro lado, es necesario demostrar la inocuidad antes de proponer la aplicación
de cualquier nueva proteína en alimentos. Aunque será necesario ampliar estos estudios,
PgAFP y PgChP no mostraron evidencias de toxicidad sobre glóbulos rojos y larvas de
Artemia salina a concentraciones superiores a las que poseen actividad antifúngica.
De forma general y como paso previo a la aplicación de una proteína a nivel
industrial, resulta de gran importancia garantizar que la actividad antifúngica se mantiene
en presencia de determinados compuestos que puedan estar presentes en el producto donde
se vaya a aplicar. Adicionalmente el efecto de ciertos tratamientos a temperaturas o pHs
extremos puede condicionar también sus posibles aplicaciones.
Ambas proteínas PgAFP y PgChP mostraron ser estables a altas temperaturas,
permaneciendo activas tras ser sometidas a 100 ºC 15 minutos y 80 ºC 20 minutos,
respectivamente. PgAFP además mostró ser estable a valores de pH entre 1 y 12.
La actividad antifúngica de PgAFP y PgChP se mantuvo en presencia de altas
concentraciones (100 mM) de sodio y potasio, lo que puede ser de gran interés en
alimentos madurados, donde abunda el cloruro sódico y puede haber otras sales de sodio y
potasio. Los quelantes EDTA y EGTA no tuvieron un gran impacto en la actividad
antifúngica de PgChP y PgAFP, mientras que el fosfato sódico a alta concentración (100
mM) sí pudo inhibir esta actividad. Los detergentes también inhibieron la actividad
antifúngica de PgChP, pero PgAFP se mantuvo activa a concentraciones de tween 20 o de
tween 80 más altas a las empleadas en los alimentos.
En definitiva, PgAFP y PgChP han mostrado alta estabilidad, han resultado ser
inocuas en los ensayos realizados y resisten componentes comúnmente encontrados en
alimentos. Aunque es necesario profundizar en su estudio, estas proteínas y las cepas de P.
chrysogenum que las producen muestran características adecuadas para su utilización
frente a mohos alterantes y toxigénicos en alimentos crudos madurados.
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