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Facultad de Ciencias y Tecnologías Químicas
Departamento de Química Analítica y Tecnología de los Alimentos
BIOTECNOLOGÍA MICROBIANA:
PRODUCCIÓN, CARACTERIZACIÓN E
INMOVILIZACIÓN DE ENZIMAS DE INTERÉS
INDUSTRIAL
TESIS DOCTORAL
SHEILA ROMO SÁNCHEZ
CIUDAD REAL, 2013
Departamento de Química Analítica y Tecnología de los Alimentos Facultad de Ciencias y Tecnologías Químicas
BIOTECNOLOGÍA MICROBIANA: PRODUCCIÓN,
CARACTERIZACIÓN E INMOVILIZACIÓN DE
ENZIMAS DE INTERÉS INDUSTRIAL
SHEILA ROMO SÁNCHEZ
Visado en Ciudad Real, 21 de Diciembre de 2012
Trabajo presentado para optar al grado de Doctor por la Universidad de Castilla-La Mancha,
Fdo: Sheila Romo Sánchez
Fdo.: María Arévalo Villena
Profesor Contratado Doctor de Tecnología de los Alimentos Universidad de Castilla-La Mancha
Fdo.: Ana Isabel Briones Pérez Catedrática de Tecnología de los Alimentos
Universidad de Castilla-La Mancha
Departamento de Química Analítica y Tecnología de los Alimentos Facultad de Ciencias y Tecnologías Químicas
Dª. Juana Rodríguez Flores, Catedrática de Universidad y Secretaria del Departamento de Química Analítica y Tecnología de los Alimentos de la Universidad de Castilla-La Mancha. CERTIFICA: Que el presente trabajo de investigación titulado “Biotecnología microbiana: producción, caracterización e inmovilización de enzimas de interés industrial” constituye la Tesis Doctoral que presenta Dña. Sheila Romo Sánchez, para aspirar al Grado de Doctor en Investigación Básica y Aplicada en Recursos Cinegéticos en Química, y que ha sido realizada en el Departamento de Química Analítica y Tecnología de los Alimentos, cumpliendo todos los requisitos necesarios, bajo la dirección de la Dra. Dª Ana Briones Pérez y la Dra. Dª. María Arévalo Villena. Y para que así conste, expido y firmo el presente certificado en Ciudad Real a veintiuno de diciembre de dos mil doce.
VºBº Fdo.: Ana Isabel Briones Pérez Fdo.: Juana Rodríguez Flores Directora del Departamento Secretaria del Departamento
“Soy de las que piensan que la ciencia tiene una gran belleza.
Un científico en su laboratorio no es sólo un técnico: es
también un niño colocado ante fenómenos naturales que le
impresionan como un cuento de hadas”
- Marie Curie -
A los que más quiero.
Mis padres, mi hermana y Pedro
Este es uno de esos momentos en el que se cierra una etapa y comienza otra.
Etapa que para mí realmente comenzó en 2005 cuando entré a formar parte de este
grupo de investigación. En esta trayectoria, muchas son las personas que han formado
parte de mi vida y a las que tengo que agradecer que hayan estado conmigo en los
buenos momentos pero sobre todo en los malos.
GRACIAS A MIS DIRECTORAS DE TESIS. ANA, confiaste en mí desde el primer día
y me diste la oportunidad de conocer un mundo que ignoraba por completo y que día
tras día creaba adicción. Gracias por tu paciencia, comprensión y flexibilidad en todo, y
porque sin ti no lo habría podido realizar. MARÍA, no sólo has sido mi directora de
tesis, sino también mi confidente, un gran apoyo dentro y fuera del laboratorio, me
has transmitido alegría y serenidad en numerosas ocasiones, eres una gran amiga y sé
que te tengo para siempre. Os quiero y siempre formareis parte de mi vida.
GRACIAS JUAN. Por compartir tus conocimientos y darnos sabios consejos. Y
porque junto con Ana habéis hecho de BIOLEV una gran “familia”.
GRACIAS MARÍA, CHUS y NURIA. Mis amigas de Ciudad Real. Nos conocimos en
el laboratorio y nos hicimos inseparables. Vuestras risas, ánimos y tantos momentos
irrepetibles en el trabajo y fuera de él han hecho que los años hayan pasado
demasiado rápido.
GRACIAS A TODAS LAS PERSONAS QUE HAN PASADO POR EL LABORATORIO.
Patricia, Mónica y María M. por vuestra ayuda y porque los buenos momentos siempre
quedan. A Héctor, Conrado y Milla por traer al laboratorio tanta ciencia y aportarme
tanto personal como profesionalmente. A Gamze porque hiciste que cada segundo
fuera “very delicious”. A Irene, Ana, Cristina, Estefanía, Raquel, Cristiane, Inés, Rubén,
Ellen, Patti y tantos otros con los que compartir pipetas ha sido una tarea fácil.
GRACIAS A TODOS MIS AMIGOS, especialmente a LAURA, REBECA, ANI,
LORENA, GEMA, MIRIAM, ANAS Y CRISTINAS. Por demostrarme que las amigas “del
alma” existen.
GRACIAS MAMÁ Y GRACIAS PAPÁ. Sin vosotros nada sería posible. Tengo la gran
suerte de tener unos padres maravillosos ¡que me habéis dado tanto!, especialmente
amor y eso puede con todo.
GRACIAS PALOMA. Porque creo que sin ti, no soy nada. Eres fundamental en mi
vida.
GRACIAS PEDRO. Cada día que pasa estoy más segura de que te necesito. Tus
consejos, generosidad, paz, optimismo, apoyo incondicional y cariño hacen que todo
sea posible. Y esto también, ha sido posible.
GRACIAS A MIS ABUELOS. Porque cerca o lejos siempre estáis ahí.
GRACIAS A MIS PRIMOS Y TIOS. Por esas comilonas que hacen que me olvide de
todo. Y a ALEJANDRO porque eres “mi niño” y siempre me haces feliz.
GRACIAS A MI FAMILIA POLÍTICA. Por dejarme conoceros cada día mejor y por
compartir momentos entrañables. A PABLO, JORGE y BLANCA porque vuestra
inocencia y alegría me evaden del mundo real.
GRACIAS FÁTIMA. Porque no todo era Máster, también había muchas risas y
mucha complicidad.
GRACIAS NOELIA, MARIO, EDUARDO. Por vuestras aportaciones en el trabajo.
Y GRACIAS A TODOS los que han hecho de esta Tesis Doctoral una realidad
PRÓLOGO
La enzimología es actualmente uno de los campos de interés en el ámbito
de la biotecnología. La pasión por las enzimas de Arthur Kornerbg quedó
patente a lo largo de su vida, “Jamás me he encontrado con una enzima que
carezca de interés”, y sirvió a muchos investigadores para confiar cada vez más
en estos catalizadores biológicos.
Esta memoria de Tesis Doctoral recoge estudios, en los que se abordan
aspectos relacionados con las enzimas. Se agrupan en tres grandes bloques:
identificación de especies microbianas procedentes del ecosistema oleico y
estudio de sus propiedades biotecnológicas; obtención de enzimas de interés
mediante crecimiento de mohos por fermentación en fase sólida sobre
diversos subproductos; y caracterización e inmovilización de enzimas
industriales.
Está estructurada en siete capítulos, así tras la justificación del tema
(capítulo 1), se hace un recorrido acerca del estado del arte revisando los
aspectos más relevantes objeto de estudio (capítulo 2). En el capítulo 3 se
recogen los objetivos planteados y en el cuatro y cinco, las publicaciones
derivadas del trabajo y sus resúmenes. Los dos últimos capítulos se dedican a
las conclusiones generales y a la bibliografía (capítulos 6 y 7 respectivamente).
ÍNDICE
JUSTIFICACIÓN
1
1. ANTECEDENTES BIBLIOGRÁFICOS
5
1.1. PRODUCTOS Y SUBPRODUCTOS AGROINDUSTRIALES
5
1.2. BIODIVERSIDAD MICROBIANA EN AMBIENTES OLEICOS
7
1.3. FERMENTACIÓN EN FASE SÓLIDA
12
1.4. ENZIMAS
17
1.4.1. Enzimas lignocelulósicas
18
1.4.2. Inmovilización de enzimas
25
2. OBJETIVOS
35
3. ARTÍCULOS CIENTÍFICOS
39
3.1. Yeast biodiversity from oleic ecosystems: study of their biotechnological properties
43
3.2. Fungi isolated from olive ecosystems and screening of their potential biotechnological use
51
3.3. Production and immobilization of enzymes by solid-state fermentation of agroindustrial waste
59
3.4. Immobilization of commercial cellulase and xylanase on different polymer supports (alginate-chitin and chitosan-chitin) by different supports
81
3.5. Immobilization of β-glucosidase and its application for enhancement of aroma precursors in Muscat wine
103
4. RESUMEN DE ARTÍCULOS
137
4.1. Biodiversidad de levaduras procedentes de ecosistemas oleicos: Estudio de sus propiedades biotecnológicas
137
4.2. Mohos aislados de ecosistemas oleicos y su uso en biotecnología
139
4.3. Producción e inmovilización de enzimas mediante fermentación en fase sólida de residuos agroindustriales
141
4.4. Inmovilización de celulasa y xilanasa comerciales sobre diferentes soportes poliméricos (quitina-alginato y quitina-quitosano) y mediante diferentes métodos
144
4.5. Inmovilización de una β-glucosidasa y su aplicación para la liberación de precursores del aroma en un vino Moscatel
146
5. CONCLUSIONES GENERALES
153
6. BIBLIOGRAFÍA
157
Justificación
Sheila Romo Sánchez, 2013 1
Los residuos generados durante los procesos de elaboración del aceite de
oliva y del vino, dos de las grandes fuentes económicas de Castilla-La Mancha,
aunque suponen un problema medio ambiental se podrían emplear para la
obtención de compuestos con alto valor añadido por sus interesantes
propiedades tecnológicas y nutricionales.
Para poder explotar sus beneficios, reutilizarlos y/o revalorizarlos es
necesario aplicar ciertos pre-tratamientos más o menos complejos, que
facilitan su biotransformación, biorremediación, o detoxificación biológica,
procesos llevados a cabo normalmente mediante fermentación en fase sólida.
Una de las aplicaciones con más potencial y de mayor interés en la
industria, es la búsqueda y producción de enzimas de origen microbiano,
utilizando sustratos económicos que reduzcan los costes de producción a
escala industrial.
En la actualidad, la biotecnología de enzimas es un núcleo común entre
sectores de la agricultura, medio ambiente, medicina, e industrias
farmacéuticas, químicas o de alimentos. Una excelente alternativa para su uso
es la inmovilización, que supone entre otras ventajas, el aumento de la
estabilidad y la posibilidad de reúso.
Antecedentes Bibliográficos
Sheila Romo Sánchez, 2013 5
1.1. SUBPRODUCTOS AGROINDUSTRIALES
La industria de alimentos genera residuos en el proceso de elaboración y
preparación, a través de las aguas residuales y efluentes y por los alimentos
alterados. Producen grandes volúmenes de subproductos, tanto sólidos como
líquidos, a consecuencia de la producción, preparación y consumo de
alimentos. Éstos son caros de recoger, tratar y eliminar, y representan una
pérdida de materiales valiosos. De modo que es importante convertirlos
rápidamente en productos inocuos sin causar daño al entorno (Lee, 2000).
España presenta la mayor superficie dedicada al cultivo de olivos (Olea
europea L.) y es el primer productor y exportador de aceite de oliva y de
aceitunas de mesa. A nivel nacional y en términos de superficie, este sector
ocupa la segunda posición, después de los cereales, y se encuentra repartido
en 34 de las 50 provincias (Agencia para el Aceite de Oliva-AAO).
Castilla-La Mancha representa la segunda región olivarera (16% de la
superficie total), siendo Toledo y Ciudad Real las dos provincias más
representativas. (Figura 1). De ese porcentaje, el 96% corresponden a
variedades de aceituna para almazara (Castellana, Cornicabra y Picual) y el 4%
restante a variedades de mesa.
Figura 1. Distribución geográfica de la superficie olivarera en España (AAO)
Antecedentes Bibliográficos
6 Sheila Romo Sánchez, 2013
En el proceso de molturación de la aceituna y dependiendo del sistema
de centrifugación empleado, además del aceite se obtienen subproductos
como las aguas de vegetación y los orujos o alpeorujos (alpechines). Estos
residuos sólidos, formados por celulosa, lignina, grasa, hemicelulosa y pectina,
son altamente fitotóxicos, lo que supone un problema medioambiental.
Con el fin de minimizar este impacto se han desarrollado dos procesos de
extracción alternativos al método tradicional de presión. La diferencia entre
ambos radica en que se sustituye la centrifugación de tres fases (proporción
de agua 1:1), por un sistema de decantación de dos fases que requiere menos
agua pero genera orujos más húmedos y de difícil manejo. Este “proceso
ecológico” reduce casi el 75% el volumen de residuos en las almazaras y es el
más usado hoy en día.
Ya Giannoutsou y col. (2004) sugirieron que el alpeorujo es un buen
sustrato para el crecimiento de levaduras, y que tras su fermentación se
puede utilizar como fertilizante, aditivo alimentario o crecimiento de hongos
comestibles.
Otro de los cultivos importantes de la economía española (cuarto en
producción a nivel mundial) es el de la vid (Vitis vinífera L.), siendo Castilla-La
Mancha la región con más viñedos (más de 600.000 Ha de superficie) y la
mayor productora de vino, superando el 50% del producto total nacional.
Además, un menor porcentaje de uva se comercializa como uva de mesa o
desecada (Ali y col., 2010).
En el proceso de vinificación, los residuos que se generan son
básicamente orujos constituídos por hollejos, semillas y raspones, que
suponen un 13% del peso de la uva (Ruberto y col., 2008). Su composición es
variable, y son ricos en azúcares, alcoholes, compuestos carbonílicos,
Antecedentes Bibliográficos
Sheila Romo Sánchez, 2013 7
polifenoles, sustancias minerales, pectinas, celulosa, etc. Además, su bajo pH y
su contenido en sustancias fenólicas de carácter fitotóxico y antibacteriano,
hacen que su degradación biológica sea difícil (Bustamante y col., 2008).
Estos subproductos se suelen emplear para alimentación animal,
fertilización del suelo, crecimiento de hongos comestibles, obtención de
biocombustibles, producción de energía o extracción de aceite a partir de sus
semillas (Molero y col., 1995). Recientemente, se trabaja en el aislamiento de
sustancias antioxidantes con fines nutracéuticos.
A pesar de que las industrias agroalimentarias, se han adecuado a las
exigencias legislativas, algunas de ellas carecen de un plan de gestión de
residuos, y se limitan a ubicarlos en vertederos o a usarlos de forma
ineficiente, lo que supone un problema medioambiental, que obliga a la
búsqueda de procesos para su tratamiento y reutilización.
Para un desarrollo sostenible, es necesario conocer en profundidad la
materia prima y estudiar sus propiedades químicas y biotecnológicas.
1.2. BIODIVERSIDAD MICROBIANA EN ECOSISTEMAS OLEICOS
La microbiota espontánea de las aceitunas está formada
mayoritariamente por levaduras, bacterias ácido lácticas (LAB),
enterobacterias y hongos filamentosos. En su biodiversidad y concentración
influyen factores como la variedad, temperatura, pluviosidad, tipo de suelo,
fertilización, regadío, prácticas de cultivo, podredumbres o tipo de
recolección, entre otros.
Las aceitunas de mesa se elaboran mediante fermentaciones
espontáneas por levaduras y LAB. Estas últimas son objeto de estudio por la
Antecedentes Bibliográficos
8 Sheila Romo Sánchez, 2013
producción de bacteriocinas y ácido láctico (Sánchez y col., 2001; Delgado y
col., 2007). Sin embargo, últimamente se ha reconsiderado la influencia
positiva que ejercen las levaduras en el proceso de elaboración (Arroyo-López
y col., 2008). Así, Alves y col. (2012) demostraron que al final de dicha
fermentación hay presencia de especies de Zigosaccharomyces mrakii y
Saccharomyces cerevisiae y ausencia de Escherichia coli y enterobacterias.
Con respecto a las aceitunas destinadas a producción de aceite de oliva,
pocas son las referencias de su biodiversidad. Romo-Sánchez y col. (2010) y
Alves-Baffi y col. (2012) estudiaron la variabilidad de especies de levaduras y
mohos respectivamente, tanto en los frutos frescos como en las pastas y
orujos.
1.2.1. Identificación de especies
La identificación y clasificación de especies microbianas por métodos
clásicos se lleva a cabo utilizando criterios morfológicos y fisiológicos. Entre los
primeros se estudian aspectos macroscópicos (forma, color, aspecto y
consistencia de las colonias; área, longitud o anchura de hifas) y microscópicos
(reproducción sexual, morfología celular o formación de pseudomicelio). Los
fisiológicos valoran la capacidad de fermentación y asimilación de hidratos de
carbono y compuestos nitrogenados, la resistencia a altas presiones osmóticas
o la hidrólisis de la urea, entre otros (Samson y col., 1995; Barnett y col.,
2000).
Otros métodos fenotípicos usados para la caracterización son el análisis
del perfil de ácidos grasos de las membranas plasmáticas (Malfeito-Ferreira y
col., 1989), la caracterización de las proteínas intracelulares (Van Vuuren y
Antecedentes Bibliográficos
Sheila Romo Sánchez, 2013 9
Van der Meer, 1987), o la detección de isoenzimas en Saccharomyces sensu
stricto (Duarte y col., 1999).
En las últimas décadas se han desarrollado numerosas técnicas de
biología molecular, independientes del estado fisiológico de la célula y que
suponen una buena alternativa a las metodologías tradicionales, ya que tienen
un poder discriminante superior, pudiendo llegar a caracterizar a nivel de
cepa. Entre los métodos moleculares más frecuentes se encuentran:
i. Análisis de cariotipos
Schawrtz y Carton (1984) desarrollaron un método electroforético (PFGE-
Pulsed Field Gel Electrophoresis), que consistía en dos campos eléctricos
alternativamente pulsantes (de los cuáles, al menos uno no era homogéneo) y
orientados perpendicularmente, permitiendo así la separación de moléculas
de ADN con un tamaño máximo de 2Mbp en una matriz de agarosa. Sin
embargo, la migración de las moléculas de ADN es inestable, por lo que
surgieron otras alternativas como OFAGE (Orthogonal Field Alternation Gel
Electrophoresis) o FIGE (Fiel Inversion Gel Electrophoresis), destacando esta
última por diferenciar fragmentos de ADN de más de 2 Mbp.
Otros métodos más sofisticados y resolutivos son el CHEF (Contour-
clamped homogeneous electric field), TAFE (Transverse Alternating Field
Electrophoresis) y PACE (Programmable autonomously controlled electrode
gel electrophoresis).
Independientemente del sistema utilizado, suelen ser herramientas
valiosas para analizar genomas fúngicos. Vallejo y col. (1996) mediante esta
técnica demostraron la existencia de polimorfismo cromosómico en Botrytis
cinérea. Algunos autores la han empleado en estudios poblacionales de
Antecedentes Bibliográficos
10 Sheila Romo Sánchez, 2013
levaduras durante la fermentación vínica (Briones y col., 1996; Izquierdo y col.,
1997).
ii. Análisis de restricción del ADN mitocondrial(ADNmt)
El ADN mitocondrial posee un alto polimorfismo intraespecífico y es muy
estable durante los procesos de multiplicación celular (Ribéreau-Gayon y col.,
2000). Lo que hace que sea de las más utilizadas para diferenciar a nivel de
cepa en eucariotas.
Se basa en la digestión del ADN total con endonucleasas de restricción
del tipo GCAT, que no reconocen las secuencias ricas en GC ni ricas en AT,
típicas del ADNmt. El alto número de cortes del ADN nuclear respecto al de la
mitocondria hace que aquel se rompa en fragmentos muy pequeños que
migrarán más rápido durante la electroforesis y, por tanto, no interferirán en
la visualización de las bandas correspondientes del ADNmt (Fernández-Espinar
y col., 2005).
Al igual que la técnica anterior, se ha empleado para caracterizar cepas o
para comprobar la implatación de levaduras seco-activas en mostos en
fermentación (Barrajón y col., 2009).
iii. Técnicas de hibridación
Se basan en utilizar sondas marcadas complementarias a la secuencia de
ADN a identificar que se separan por electroforesis y se transfieren a
membranas de nylon o nitrocelulosa mediante el método Southern. La
reacción se monitoriza espectrofotométricamente midiendo la cinética de
formación de híbridos.
A pesar de que diversos autores (Vaughan-Martini, 1995) han empleado
diferentes secuencias génicas para caracterizar cepas de S. cerevisiae, Querol
Antecedentes Bibliográficos
Sheila Romo Sánchez, 2013 11
y col. (1992) mostró que el análisis de restricción del ADNmt o la electroforesis
de cromosomas son más discriminantes que las técnicas de hibridación.
iv. Métodos basados en las técnicas de PCR (Polymerase Chain Reaction)
Son los más rápidos para identificar especies y cepas. Consisten en
amplificar una secuencia conocida del material genético por acción de una
ADN polimerasa, tras la unión de cebadores complementarios a zonas diana
de ambas hebras de la cadena de ADN. Existen estudios en los que se
amplifica todo el ADNr nuclear, proceso conocido como ribotipado (Smole
Mozina y col., 1997) y en otros sólo se amplifican las regiones ribosómicas NTS
(non-transcribed spacer), seguidas de una restricción con enzimas (RFLP-
Restriction Fragment Length Polimorphism) (Caruso y col., 2002) realizando
así una identificación a nivel de especie.
Para la diferenciación de cepas, las técnicas más utilizadas son RAPD
(Random Amplified Polymorphic) y microsatélites. La primera utiliza un único
cebador de cadena muy corta y secuencia arbitraria (Williams y col., 1990) y
una temperatura de hibridación baja (37ºC). Los microsatélites (< 10 pb) o
minisatélites (10 – 100 pb), emplean diversos cebadores cuya secuencia
complementaria se repite con frecuencia a lo largo del genoma, anillando a
una temperatura superior (60 - 65ºC), lo que hace que los productos de PCR
sean más estables que en la RAPD-PCR (Marinangeli y col., 2004).
v. Secuenciación nucleotídica
Esta técnica permite conocer la composición y orden de los nucleótidos
en una determinada secuencia de material genético.
Antecedentes Bibliográficos
12 Sheila Romo Sánchez, 2013
El Método de Terminación de la Cadena o Método de Sanger, en el que
se usan didesoxinucleótidos trifosfato como terminadores de la cadena de
ADN, es el más eficiente y menos radiactivo.
Mediante un programa informático de alineamiento (BLAST), se compara
la secuencia problema (genes ribosomales 5.8S, 18S o 26S) con otras
depositadas en bases de datos. De esta forma, se pueden establecer
relaciones entre los distintos niveles taxonómicos.
La secuenciación del gen 18S del ADNr de Saccharomyces demostró que
este género es muy heterogéneo y que sus especies se encuentran
entremezcladas con algunas de Zygosaccharomyces, Torulaspora y
Kluyveromyces. No obstante, las seis especies pertenecientes al grupo
Saccharomyces sensu stricto (S. cerevisiae, S. bayanus, S. paradoxus, S.
pastorianus, S. krudiavzevii, S. cariocanus y S. mikatae), están muy cercanas
entre sí, y constituyen un grupo diferenciado de otras especies del mismo y de
otros géneros (James y col., 1997).
La tecnología actual permite realizar el proceso a alta velocidad, lo que ha
resultado crucial para proyectos de gran envergadura como el conocimiento
del Genoma Humano, de animales (Drosophila melanogaster), de plantas
(Arabidopsis thaliana) y de microorganismos (Saccharomyces cerevisiae).
1.3. FERMENTACIÓN EN FASE SÓLIDA
La fermentación en fase sólida (FFS) es un proceso de transformación del
material no soluble en casi ausencia de agua libre. Este material actúa como
soporte físico y fuente de nutrientes de los microorganismos y posee la
Antecedentes Bibliográficos
Sheila Romo Sánchez, 2013 13
suficiente humedad para permitir su crecimiento y metabolismo (Singhania y
col., 2009).
En la Figura 2 se observa el modelo propuesto por Moo-Young y col.
(1983), en el que se disponen las partículas sólidas húmedas y una fase
gaseosa continua alrededor de un hongo filamentoso (lado izquierdo) o un
organismo unicelular (lado derecho).
Figura 2. Disposición espacial de la FFS
De los diferentes microorganismos utilizados en la FFS destacan, debido a
su eficacia y competitividad, los hongos filamentosos que representan un 50%
(da Silva y col., 2005). Sus ventajas principales son que sus hifas penetran con
facilidad en el sustrato sólido, excretan enzimas hidrolíticas y son tolerantes a
la baja actividad de agua, resistiendo niveles de alta presión osmótica
(Gutiérrez-Rojas y col., 1995). En menor medida, se emplean levaduras en un
30% (Alfani y col., 2000) y en menor medida actinomicetos (15%) y bacterias
(5%) (Amin, 1992).
Antecedentes Bibliográficos
14 Sheila Romo Sánchez, 2013
En la Figura 3, se observa la disposición de las hifas en el sustrato sólido:
en el interior de la matriz (capa 3), sobre su superficie (capa 2) o hacia el
exterior (capa 1).
Figura 3. Distribución de las hifas fúngicas en un sustrato sólido (Rahardjo y col.,
2006).
La velocidad de penetración de las hifas en el sustrato depende del
oxígeno disponible. Así, a medida que el moho se desarrolla, la capa 1 se hace
más densa y húmeda y se transforma en la capa 2, cuyo grosor aumenta hasta
el punto de convertir en anaerobia su parte más interna, por lo que el oxígeno
se agota con facilidad en el interior del sustrato. Bajo estas condiciones de
anoxia, el micelio de las capas 2 y 3 cesa su crecimiento o comienza a
fermentar (Rahardjo y col., 2006). Se establece un estrecho contacto entre las
hifas y la superficie del sustrato facilitando el transporte de nutrientes a través
de la membrana celular. La excreción de metabolitos, se realiza en la porción
apical de las hifas minimizándose el efecto de la dilución que se da en la
fermentación en cultivo sumergido (SmF).
Por otra parte, también existen parámetros extrínsecos como la
temperatura, humedad, aireación, agitación, diseño del reactor, entre otros,
que influyen en el rendimiento del proceso.
Micelio aéreo (1)
Micelio húmedo (2)
Micelio penetrante (3)
Interfase aire-líquido
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Sheila Romo Sánchez, 2013 15
En la elección del sustrato, hay que considerar ciertos factores como el
tamaño de partícula, la porosidad y la composición química, así como la
disponibilidad en el mercado y el precio. Los subproductos generados en la
industria agroalimentaria, serían potencialmente aptos para este proceso y
además su uso solventaría problemas económicos y medioambientales
causados por su acumulación (Pandey y col., 2000a).
1.3.1. Aplicaciones industriales de la FFS
Las diferentes características reológicas, cinéticas y termodinámicas que
existen entre los sistemas de FFS y SmF hacen que los microorganismos
prefieran para su crecimiento a la primera, debido a la similitud con su hábitat
natural (Singhania y col., 2009).
Se ha observado que el empleo de cultivos mixtos en FFS aumenta la
producción de enzimas y en ocasiones provoca un mayor efecto sinérgico
(Gutiérrez-Correa y col., 1999).
Se ha aplicado para la producción de alimentos o de metabolitos de
interés con importante valor añadido (dos Santos y col., 2004; Saqib y col.,
2010) y es un proceso exclusivo para la obtención de glucoamilasa (Ishida y
col., 2000) y de esporas fúngicas usadas en biocontrol (de Vrije y col., 2001).
Uno de sus inconvenientes reside en la baja estabilidad de los
metabolitos frente a los factores ambientales y en que resulta difícil de
controlar en procesos a gran escala (Hölker y Lenz, 2005).
Se han desarrollado nuevas aplicaciones en distintos sectores como el
agrícola, medioambiental y fermentativo (Pérez-Guerra y col., 2003). Uno de
los más importantes es la biotransformación de residuos de cosechas. Se ha
utilizado también, en el tratamiento de subproductos agrícolas y
Antecedentes Bibliográficos
16 Sheila Romo Sánchez, 2013
revalorización de cultivos tropicales (Soccol y Vandenberghe, 2003); para
mejorar la calidad nutricional de la cassava (Pandey y col., 2000a) o en la
producción de alimentos para rumiantes (Villas-Bôas y col., 2002).
En la producción de bioetanol existen numerosos estudios centrados
principalmente en la sacarificación de los sustratos, mediante el crecimiento
de mohos (Soccol y col., 2010; Bon y Ferrara, 2007).
Otra de sus aplicaciones es la biorremediación y detoxificación biológica
de compuestos. Ejemplos claros son la biodegradación de herbicidas, como la
atrazina (utilizado para controlar el crecimiento de las malas hierbas que
añadido a una mezcla de algodón con paja y trigo se inocula con Pleurotus
pulmonarius) (Masaphy y col., 1996); o la detoxificación de compuestos
antifisiológicos generados durante el procesado del café (cafeína, taninos y
polifenoles obtenidos de la pulpa y cáscara del café) (Pandey y col., 2000b).
En la industria de alimentos la FFS se aplica profusamente para la
elaboración de productos orientales, como el Kimchi (LAB), el Miso (A. niger,
lactobacillus), Tempeh (Rhizopus sp.), o Torani (Candida sp., Saccharomyces
sp.) (Christen, 1995). En nuestra cultura otros ejemplos son la maduración de
embutidos o de quesos tipo Brie y azules mediante Penicillium camemberti o
roquefortii (Couto y Sanromán, 2006).
En otros sectores biotecnológicos, también se emplea para la producción
de compuestos aromáticos (aceites esenciales y saborizantes), misceláneos
(pigmentos, surfactantes, vitaminas, xantano), bioactivos (micotoxinas,
giberelinas, antibióticos, hormona) y ácidos orgánicos (ácido cítrico, fumárico
y láctico) (Pandey y col., 2000a).
De entre el amplio número de posibilidades que ofrece en diversos
sectores de la industria, es sin duda la producción de enzimas la aplicación
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Sheila Romo Sánchez, 2013 17
más importante. Entre los trabajos más recientes, Dhillon y col. (2011)
utilizaron residuos agrícolas para producir celulasas y hemicelulasas
excretadas de Aspergillus niger y Trichoderma reseei y, Brijwani y Vadlani se
centraron en la producción de enzimas celulolíticas a partir de soja.
1.4. ENZIMAS
Las enzimas “son catalizadores biológicos, que aumentan la rapidez o
velocidad de una reacción química, sin verse alterada ella misma en el proceso
global” (Mathews y col., 2003). Se descubrieron en la segunda mitad del siglo
XIX y los avances en biotecnología de las últimas décadas, especialmente en
genética e ingeniería de proteínas, les han otorgado un papel fundamental
dentro de los procesos industriales.
Poseen alto poder catalítico, elevada especificidad y son activas a
temperatura ambiente y presión atmosférica.
Según la Comisión Enzimática y, dependiendo de la reacción que
catalicen, se clasifican en oxidorreductasas (EC 1), transferasas (EC 2),
hidrolasas (EC 3), liasas (EC 4), isomerasas (EC 5) y ligasas (EC 6).
Las enzimas comerciales utilizadas por la industria proceden de fuentes
animales, vegetales o microbianas. En este sentido, los microorganismos, de
status GRAS (Generally Recognized as Safe), son la fuente más importante de
producción gracias a su alto rendimiento, que por otra parte, se puede
mejorar por variación de los parámetros físico-químicos durante su
crecimiento o por manipulación genética. Otras propiedades que los hacen
deseables son su diversidad metabólica, estabilidad genética, crecimiento a
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18 Sheila Romo Sánchez, 2013
gran escala y el empleo de tecnologías limpias que disminuyen el coste de
producción.
La tendencia actual es reemplazar los catalizadores convencionales no
biológicos por procesos biotecnológicos que impliquen el uso de
microorganismos y/o enzimas.
El mercado de enzimas supone más de 500.000 toneladas/año; y aunque
existen más de 3800 enzimas reconocidas oficialmente, la mayoría se
comercializan como extractos crudos, no demasiados puros, a excepción de
los usados en clínica, análisis o diagnóstico. Las industrias que hacen uso de
ellos son las de detergentes, alimentos, alimentación animal, y de tratamiento
de residuos.
Se comercializan en función a su actividad, más que por su peso o
volumen, de forma que es esencial mantener la estabilidad de la preparación
enzimática durante el almacenamiento. Para ello, en ocasiones se mejoran
mediante técnicas de biología molecular o de inmovilización y estos hechos
hacen que la ingeniería de enzimas constituya uno de los retos más
apasionantes, complejos e interdisciplinares de la biotecnología actual.
(Guisan, 2006).
1.4.1. Enzimas lignocelulósicas
Los residuos agrícolas están constituidos principalmente por compuestos
lignocelulósicos, formados por celulosa, hemicelulosa, lignina y en menor
proporción y sólo en algunos casos por pectinas. En la Figura 4 se muestra su
distribución en la pared de una célula vegetal.
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Sheila Romo Sánchez, 2013 19
Figura 4. Estructura de la pared celular de los vegetales
A continuación se comentan las enzimas más interesantes en la
degradación de estos compuestos:
Celulasas
La celulosa, biomolécula orgánica más abundante en la naturaleza, es un
polisacárido compuesto por moléculas de glucosa anhidro unidas mediante
enlaces β-(1,4)-glicosídicos, siendo su unidad básica la celobiosa (Figura 5), la
unidad básica repetida es el disacárido celobiosa. Su compleja conformación,
así como su unión con otros polímeros como la lignina, hemicelulosa, almidón,
proteínas y elementos minerales, la hacen una molécula resistente a la
hidrólisis (Zaldivar y col., 2001).
Figura 5. Estructura de la celulosa
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20 Sheila Romo Sánchez, 2013
La sacarificación de este polisacárido requiere la acción conjunta de
endoglucanasas (endo-1,4-β-glucanasa, EC 3.2.1.4) que hidrolizan el polímero
de celulosa desde el interior, originando extremos reductores y no-reductores;
exoglucanasas o celobiohidrolasas (exo-1,4-β-glucanasa EC 3.2.1.91), las
cuáles actúan sobre esas terminaciones liberando celobiosa y oligosacáridos; y
β-glucosidasas o celobiasas (EC 3.2.1.21), que hidrolizan la celobiosa, liberan
azúcares fermentables e inhiben la acción de exo- y endoglucanasas (Kaur y
col., 2012).
El complejo enzimático de la celulasa además de utilizarse en la hidrólisis
de materiales celulósicos, se emplea en la mejora de la textura y palatabilidad
de vegetales de baja calidad, y para acelerar el secado de los vegetales
(Prescott y Dunn’s, 1982).
Xilanasas
El xilano (Figura 6), el mayor componente de las hemicelulosas de la
pared de las células vegetales, es un polisacárido heterogéneo formado por
cadenas de xilosa unidas mediante enlaces β-1,4. Dependiendo del tipo de
materia prima puede contener residuos de arabinosa, ácido glucurónico y
ácido arabino-glucurónico (Gupta y Kar, 2008).
Figura 6. Estructura del xilano
La hidrólisis del xilano mediante xilanasas supone una alternativa a la
hidrólisis química. El sistema de enzimas xilanolíticas se compone de β-1,4-
endoxilanasa, β-xiloxidasa, α-1-arabinofuranosidasa, α-glucuronidasa, acetil
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Sheila Romo Sánchez, 2013 21
xilano esterasa y ácido fenol esterasa (Beg y col., 2001). De todas ellas, la β-
1,4-endoxilanasa (E.C. 3.2.1.8) y β-xiloxidasa son las más importantes; la
primera ataca los enlaces internos β-1,4 de la cadena principal y la segunda
libera residuos de xilosa (Bakir y col., 2001). El resto de enzimas degradan las
ramificaciones de este heteropolímero.
Se utilizan en el blanqueado del papel y la pulpa, disminuyendo así la
cantidad de cloro necesaria en el proceso; en la industria textil para tratar las
fibras; en la alimentación animal o en la industria farmaceútica como
suplemento de dietas. Pero es sin duda en el sector de los alimentos donde se
aplican con mayor frecuencia: en mejora de propiedades nutricionales;
recuperación de aromas, aceites esenciales, vitaminas, sales minerales o
pigmentos; en la digestión de las semillas del café; recuperación de azúcares
fermentables a partir de hemicelulosas; clarificación de zumos de frutas, junto
con pectinasas y celulasas; o en aditivos de la harina de trigo para mejorar la
manipulación de la pasta (Polizeli y col., 2005).
Ligninasas
La lignina, el segundo biopolímero más abundante en la naturaleza, es el
único sintetizado de forma natural que posee una cadena principal aromática.
Está formada por subunidades de guaiacol, siringol y p-hidroxifenol en distinta
proporción dependiendo de su procedencia (Figura 7). El acoplamiento
oxidativo de estos alcoholes monoméricos forma una estructura compleja de
difícil degradación (Wong, 2009).
Antecedentes Bibliográficos
22 Sheila Romo Sánchez, 2013
Figura 7. Estructura de la lignina
La lignina se degrada mediante ligninasas, clasificadas en dos grupos:
fenol-oxidasas (lacasas) y peroxidasas (lignina-peroxidasa, manganeso-
peroxidasa y peroxidasa versátil) (Dashtban y col., 2010).
Este sistema enzimático posee prometedoras aplicaciones en la industria
del papel y la pulpa, la biodegradación de químicos tóxicos y especialmente en
la síntesis química y en la industria textil (Alam y col., 2005).
Pectinasas
La pectina se encuentra en la mitad de la lamela y en la pared celular
primaria de las plantas superiores (Kashyap y col., 2001). Como muestra la
Figura 8, es un polisacárido complejo compuesto por cadenas de ácido D-
galacturónico unidos por enlace α-1,4. Éste se presenta en forma de cadenas
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Sheila Romo Sánchez, 2013 23
en zig-zag con ramificaciones cortas de azúcares neutros (L-ramnosa, D-
glucosa, L-arabinosa, D-xilosa, D-galactosa). Los grupos carboxil del ácido
galacturónico están parcialmente esterificados mediante grupos metilo y
parcial o completamente neutralizados por iones sodio, potasio o amonio.
Dependiendo del tipo de modificación en la cadena principal, las sustancias
pécticas se clasifican en protopectina, ácido péctico, ácido pectínico y pectina
(Kashyap y col., 2001).
Figura 8. Estructura de la pectina
En función del mecanismo de acción, las enzimas que degradan la pectina
se clasifican en tres grupos. Protopectinasas: solubilizan la protopectina dando
lugar a pectina soluble altamente polimerizada; pectinesterasas, catalizan la
de-esterificación del grupo metoxil de la pectina dando lugar a ácido péctico; y
depolimerasas, que hidrolizan el enlace α–(1,4)-glicosídico
(polimetilgalacturonasas y poligalacturonasas) o lo rompen por trans-
eliminación (liasas) (Kashyap y col., 2001).
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24 Sheila Romo Sánchez, 2013
Se utilizan en la industria de alimentos para clarificar zumos de frutas o
vinos, mejorar los rendimientos de extracción y la textura de algunos frutos,
eliminar la piel de los cítricos, facilitar la degradación de las cubiertas de
algunas semillas como el café; en la industria del papel y textil; en el
tratamiento de aguas residuales; y en la purificación de plantas virosas (Jayani
y col., 2005).
Los microorganismos con mayor capacidad de biosíntesis y excreción de
estas enzimas mediante fermentación en fase sólida son los mohos,
empleados en numerosas investigaciones.
Para la obtención de enzimas lignocelulósicas, en numerosas
investigaciones se ha empleado la fermentación en fase sólida llevada a cabo
por mohos. Destacando algunas de ellas, Ncube et al. (2012) obtuvieron altos
niveles de celulasa con Aspergillus niger sobre Jatropha; Thermoascus
auranticus Miehe es un buen productor de xilanasa cuando se crece sobre
maíz y hierba (da Silva et al., 2005), mientras que Aspergillus niger lo es sobre
cebada (Soliman et al., 2012).
En otro estudio y utilizando Phanerochaete chrysosporium y la biomasa
del aceite de palma se obtuvo una elevada concentración de ligninasa (Alam y
col., 2005). Rodriguez-Fernández y col. (2011) optimizaron la producción de
pectinasas de Aspergillus sp., consiguiendo un buen rendimiento a las 72 h del
proceso.
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Sheila Romo Sánchez, 2013 25
1.4.2. Inmovilización de enzimas
Tanto por razones económicas como técnicas muchos de los procesos
químicos catalizados por enzimas requieren el re-uso o el uso continuo de los
biocatalizadores durante un período de tiempo.
La mayoría de enzimas pierden estabilidad en las condiciones de trabajo
o se inactivan por la inhibición de sustratos/productos, por lo que una
alternativa sería su inmovilización permitiendo además, que el proceso
biotecnológico sea económicamente más rentable (Arroyo, 1998).
La inmovilización de enzimas se define como un “proceso en el que se
confina o localiza la enzima en una región definida del espacio para dar lugar
a formas insolubles que retienen su actividad catalítica y que pueden ser
reutilizadas repetidamente” (Wingard, 1972). Posteriormente, esta definición
se amplió a “aquel proceso por el que se restringen, completa o parcialmente,
los grados de libertad de movimiento de enzimas, por su unión a un soporte”
(Taylor, 1991).
Esta tecnología presenta una serie de ventajas como el aumento de la
estabilidad enzimática, reutilización del complejo, minimización de costes y
posibilidad de diseñar reactores de fácil manejo y control, adaptados a la
aplicación de la enzima inmovilizada, permitiendo cargas elevadas de enzima.
Sin embargo, también supone inconvenientes tales como su alteración
conformacional con respecto al estado nativo, gran heterogeneidad del
sistema enzima-soporte o posible pérdida de actividad.
Desde el punto de vista industrial, el desarrollo práctico de protocolos
para inmovilización de enzimas está relacionado con la simplicidad,
efectividad de costes y, estabilización enzimática (Guisan, 2006). Por otra
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26 Sheila Romo Sánchez, 2013
parte, el re-uso a largo plazo requiere la preparación de matrices con
derivados estables y propiedades funcionales deseables.
La inmovilización, por el mero hecho de unir las proteínas a soportes
sólidos, permite disponer de biocatalizadores insolubles fácilmente separables
del sustrato; sin embargo, no implica necesariamente estabilización, por lo
que es importante elegir el soporte más adecuado para mantener la
estructura terciaria de la proteína, evitando su desnaturalización y la pérdida
de actividad enzimática (Mozhaev, 1993).
Esta unión enzima-soporte puede ser reversible o irreversible. La primera,
conlleva menor estabilidad, debido a que la estructura terciaria no alcanza la
rigidez óptima, excepto en el caso de enzimas multiméricas, con estructuras
cuaternarias que la refuerzan (Torres y col., 2004).
1.4.2.1. Soportes
Como matrices se utilizan materiales orgánicos e inorgánicos, inertes,
disponibles y económicamente rentables, aunque el soporte ideal además
debe presentar alta estabilidad, regenerabilidad, capacidad para aumentar la
especificidad/actividad enzimática y reducir tanto la inhibición del producto
como la contaminación microbiana (Singh, 2009).
Éstos se clasifican en dos grupos:
- Inorgánicos: naturales (bentonita, piedra pómez, sílice, óxidos de
metales, vidrio no poroso, alúmina, cerámicas, sílice, entre otros) o
manufacturados (óxidos de metales, vidrio no poroso, alúmina, cerámicas o
gel de sílice).
- Orgánicos: polímeros naturales como polisacáridos (celulosa, almidón,
dextranos, agar-agar, quitina, alginatos, agarosa, quitosano entre otros),
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Sheila Romo Sánchez, 2013 27
polímeros sintéticos como poliolefinas (por ejemplo, poliestireno), polímeros
acrílicos (poliacrilatos, poliacrilamidas, polimetacrilatos, etc.) o proteínas
fibrosas (colágeno o queratina), alcohol polivinílico o poliamidas.
1.4.2.2. Métodos de inmovilización
Basándose en el tipo de unión entre enzima y soporte se dividen en:
retención física, donde las interacciones entre el soporte y la enzima son
débiles y unión química en la que se establecen enlaces iónicos o covalentes
entre ambos (Arroyo, 1998).
Retención física
Incluye el atrapamiento en capas o fibras y la inclusión en membranas
(microencapsulación y reactores de membrana) (Figura 9).
El atrapamiento consiste en la inclusión de la enzima en una matriz sólida
porosa formada generalmente por prepolímeros fotoentrecruzables o
polímeros del tipo poliacrilamida, colágeno, alginato, carragenato o resinas de
poliuretano. La enzima se suspende en la solución del monómero y se provoca
la polimerización por un cambio de temperatura o por la adición de un agente
polimerizante, pudiéndose dar en geles o fibras, siendo el último más
resistente.
Fan y col. (2011) introdujeron una variante, en la que se añadía un
reactivo bi-funcional (glutaraldehído) para reforzar la unión enzima-soporte
obteniéndose mejores resultados.
La inclusión en membranas, engloba a la microencapsulación y a los
reactores de membrana. Ambas técnicas se fundamentan en la inclusión de la
enzima en membranas semipermeables que permiten el intercambio de
metabolitos, pero que retienen la enzima. En el primer caso las microcápsulas
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28 Sheila Romo Sánchez, 2013
se producen en presencia del biocatalizador, que queda retenido en su
interior y en el segundo el biocatalizador se introduce en el interior de un
sistema con membranas previamente formadas (Alvero y col., 1998).
La microencapsulación puede ser permanente o no, dependiendo de si
son generadas por polimerización interfacial o por surfactantes.
Los reactores de membrana permiten trabajar en continuo, por lo que
resultan de gran interés en la industria. Otros son los de lecho fluidizado y los
agitados (éstos menos utilizados por la agitación severa a la que se someten
los biocatalizadores inmovilizados) (Alvero y col., 1998).
Figura 9. Métodos de inmovilización mediante retención física
Unión química
En esta tecnología destacan la unión a soportes y la reticulación (Figura
10).
Actualmente, el más utilizado en la industria es la unión a soportes. El
tipo de enlace y la elección de la matriz determinan el comportamiento.
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Sheila Romo Sánchez, 2013 29
Las enzimas se inmovilizan en los soportes mediante adsorción o unión
covalente. En el primero, la unión se produce por interacciones iónicas,
fuerzas de Van der Waals y puentes de hidrógeno, sin funcionalización de la
matriz. Influyen factores como el pH del medio, la concentración y tipo de
iones, o el diámetro del poro entre otros. Una variante dentro de esta técnica
es la utilización de resinas de intercambio iónico (aluminosilicatos), que se ha
empleado para la inmovilización de lipasas funcionalizando las resinas con
grupos amínicos y sulfónicos.
La unión covalente, se basa en activar grupos químicos del soporte para
que reaccionen con los grupos nucleofílicos de las proteínas.
Xu y col. (2011) inmovilizaron simultáneamente una celulasa y una
xilanasa sobre un polímero soluble reversible mostrando buenas actividades
enzimáticas; Jung y col. (2012) tras inmovilizar una β-glucosidasa en gel de
sílice mediante enlace covalente lograron mantener alto poder catalítico tras
20 ciclos de reúso. Por otra parte, novedosos estudios muestran que la
modificación covalente de la matriz es una herramienta esencial para mejorar
la estabilidad de las enzimas bajo condiciones de desnaturalización (Darias y
Villalonga, 2001; Gómez y Villalonga, 2000; Ramirez y col., 2002).
El reticulado (entrecruzamiento o cross-linking), consiste en el uso de
reactivos bi-funcionales (aldehídos, diiminoésteres, diisocianatos, diaminas,
sales de bisdiazonio) que actúan de puente entre las moléculas de enzima. El
resultado es la unión irreversible que permite a la enzima resistir condiciones
extremas de pH y temperatura.
El uso de procedimientos mixtos, consiste en inmovilizar la enzima por
adsorción sobre una resina de intercambio iónico o soporte polimérico y
después añadir el reactivo bi-funcional, normalmente glutaraldehído; o utilizar
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30 Sheila Romo Sánchez, 2013
este reactivo para activar el soporte, y después mediante adsorción
(crosslinking-adsorption) incorporar la enzima.
Siguiendo ambos métodos, Su y col. (2010) inmovilizaron una β-
glucosidasa comercial sobre dos soportes diferentes.
Un tipo de reticulación novedoso consiste en la cristalización de enzimas
y su posterior reticulado con glutaraldehído (Cross-Linked Enzyme Crystals o
CLECs). La estabilidad se debe al entramado cristalino formado, donde las
moléculas de enzima están rodeadas únicamente por otras moléculas de
proteína, actuando ellas mismas como soporte y quedando estabilizada su
estructura terciaria por las uniones covalentes intermoleculares. Roy y
Abraham (2006) cristalizaron una lacasa utilizando sulfato amónico y
mejoraron la estabilidad de la enzima nativa.
Figura 10. Métodos de inmovilización mediante unión química
Antecedentes Bibliográficos
Sheila Romo Sánchez, 2013 31
1.4.2.3. Aplicaciones
Los biosensores son una herramienta esencial en medicina, en el control
de calidad de las industrias o en medioambiente para el tratamiento de aguas
residuales (Durán y Marcato, 2012). Contienen una molécula biológica
inmovilizada próxima a un traductor que, en contacto con el analito,
transforma la señal química en señal eléctrica. La inmovilización se lleva a
cabo normalmente por inclusión en membranas semipermeables y unión
covalente a membranas. Diez y col. (2012) y Villalonga y col. (2012, 2011a,
2011b), han empleado uniones covalentes para crear biosensores
inmovilizados, mejorándose su estabilidad en todos los casos.
En enfermedades causadas por alteración o carencia de una enzima, su
inmovilización permitiría la administración de la misma consiguiendo su
liberación retardada. En la industria farmacéutica es un alternativa clave a la
síntesis química donde no es conveniente trabajar a altas temperaturas o se
requiere alta especificidad de sustrato.
Otra de las aplicaciones más importantes es en el procesado, preparación
y conservación de alimentos como la obtención de edulcorantes a partir de la
hidrólisis del almidón. Se han empleado soportes como el quitosano en la co-
inmovilización de pepsina y proteasa para la hidrólisis de proteínas del trigo
(Katchalski, 1993) y la inmovilización de lipasas ha facilitado la interestificación
de aceites y grasas. Anjani y col. (2007) y Akin y col. (2012) encapsulaban
ambas enzimas para mejorar las características organolépticas de quesos.
En alimentación animal, se utilizan preparados enzimáticos que no
resisten los procesos de elaboración y acondicionamiento de los piensos. Por
otra parte, al ser solubles en agua su separación de los sustratos y productos
es difícil, por lo que no es posible la reutilización. Además las condiciones
Antecedentes Bibliográficos
32 Sheila Romo Sánchez, 2013
metabólicas de los animales inhiben las enzimas incluidas en las raciones por
lo que su aprovechamiento se ve limitado (Johnson y col., 1993). La
inmovilización de enzimas solventaría estos problemas, ofreciendo sistemas
que soporten el pH del rumen y facilitando así la absorción total del alimento.
2.
Objetivos
Sheila Romo Sánchez, 2013 35
Los objetivos del presente trabajo son:
Conocer la biodiversidad de levaduras y mohos en pastas, orujos y
aceitunas de diferentes almazaras de Castilla-La Mancha, y estudiar
sus características biotecnológicas (Artículos I y II).
Producir enzimas lignocelulósicas mediante fermentación en fase
sólida (FFS), usando subproductos agroalimentarios (orujos de
aceituna y hollejos de uva) (Artículo III).
Adecuar los extractos multienzimáticos mediante técnicas de
purificación, liofilización e inmovilización para su posible uso en la
industria (Artículo III).
Aplicar diversas técnicas de inmovilización a enzimas comerciales
(celulasa y xilanasa) y estudiar sus características bioquímicas y
cinéticas comparándolas con las de las enzimas nativas (Artículo IV).
Inmovilizar una β-glucosidasa comercial de uso en Enología y
comparar el efecto de la enzima nativa y la inmovilizada en la hidrólisis
de precursores del aroma en mostos y vinos (Artículo V).
Artículos
Sheila Romo Sánchez, 2013 39
Esta tesis doctoral recoge las siguientes publicaciones:
I. Yeast biodiversity from oleic ecosistems: Study of their
biotechnological properties. Sheila Romo Sánchez, Milla Alves-Baffi,
María Arévalo-Villena, Juan Úbeda-Iranzo, Ana Briones-Pérez. Food
Microbiology 27: 487-492, 2010.
II. Fungi isolated from olive ecosystems and screening of their potential
biotechnological use. Milla Alves-Baffi, Sheila Romo-Sánchez, Juan
Úbeda-Iranzo, Ana Briones-Pérez. New Biotechnology 29: 451-456,
2012.
III. Production and immobilization of enzymes by solid-state fermentation
of agroindustrial waste. Sheila Romo Sánchez, Irene Gil Sánchez,
María Arévalo Villena, Ana Briones Pérez. Applied Microbiology and
Biotechnology (Enviado)
IV. Immobilization of commercial cellulase and xylanase on different
polymer supports (alginate-chitin and chitosan-chitin) by different
methods. Sheila Romo Sánchez, Conrado Camacho, Héctor L. Ramirez,
María Arévalo-Villena. New Biotechnology (Aceptado).
V. Immobilization of β-glucosidase and its application for enhancement
of aroma precursors in Muscat wine. Sheila Romo Sánchez, María
Arévalo Villena, Esteban García Romero, Héctor L. Ramirez, Ana
Briones Pérez. Food and Bioprocess Technology (Enviado).
Artículo I
Sheila Romo Sánchez, 2013 43
Artículo I
44 Sheila Romo Sánchez, 2013
Artículo I
Sheila Romo Sánchez, 2013 45
Artículo I
46 Sheila Romo Sánchez, 2013
Artículo I
Sheila Romo Sánchez, 2013 47
Artículo I
48 Sheila Romo Sánchez, 2013
Artículo II
Sheila Romo Sánchez, 2013 51
Artículo II
52 Sheila Romo Sánchez, 2013
Artículo II
Sheila Romo Sánchez, 2013 53
Artículo II
54 Sheila Romo Sánchez, 2013
Artículo II
Sheila Romo Sánchez, 2013 55
Artículo II
56 Sheila Romo Sánchez, 2013
Artículo III
Sheila Romo Sánchez, 2013 59
Artículo III
60 Sheila Romo Sánchez, 2013
PRODUCTION AND IMMOBILIZATION OF ENZYMES BY SOLID-STATE
FERMENTATION OF AGROINDUSTRIAL WASTE
Sheila Romo Sánchez, Irene Gil Sánchez, María Arévalo Villena, Ana Briones
Pérez
Food Science and Technology. University of Castilla-La Mancha (UCLM), Av.
Camilo José Cela, 13071 Ciudad Real, Spain
*Corresponding author: Sheila Romo Sánchez
Abstract The recovery of by-products from agri-food industry is currently
one of the major challenges of Biotechnology. Castilla-La Mancha produces
around three tons of waste coming from olive oil and wine industries, both of
which have a pivotal role in the economy of this region. For this reason, this
study reports on the exploitation of grape skins and olive pomaces for the
production of lignocellulosic enzymes through solid-state fermentation by
using two molds (Aspergillus niger 113N and Aspergillus fumigatus 3). In some
trials, a wheat supplementation with a 1:1 ratio was used to improve the
growth conditions, and the particle size of the substrates was altered through
milling. Separate fermentations were run and collected after 2, 4, 6, 8, 10 and
15 days to monitor enzymatic activity (xylanase, cellulase, β-glucosidase,
pectinase). The highest values were recorded after 10 and 15 days of
fermentation. The use of A. niger on unmilled grape skin yielded the best
outcomes (47.05 U xylanase / g by-product). The multi-enzymatic extracts
obtained were purified, freeze-dried, and immobilized on chitosan by
adsorption to assess the possible advantages provided by the different
techniques.
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Sheila Romo Sánchez, 2013 61
Keywords: enzyme · mold · solid-state fermentation · enzymatic activity ·
chitosan
Introduction
Considering the great deal
of green waste in nature, some
microorganisms show great
potential in biotechnology because
of their ability to metabolize
lignocellulosic compounds. Agro-
industrial waste, such as bagasse of
sugar cane, wheat, corn, rice husk,
fruit skin, and olive pomace
(concentrated waste coming from
alcohol and oil mill industries), is
on the increase because of
industrialization. This poses a
problem in terms of space available
and environmental pollution. In
Spain, olive oil and winemaking
companies figure prominently in
the agri-food sector as some of the
leading production and export
industries. This extensive
production results in
approximately three tons of waste
a year. Traditional methods for
olive oil and wine waste
management include chemical,
physical, and biological processes.
Hemicelluloses and
celluloses represent over 50% of
dry weight of by-products. These
may be transformed into soluble
sugars by acid or enzymatic
hydrolysis (Cara et al. 2008), which
would be an inexpensive and
abundant renewable energy
source. Solid-state fermentation
has been described as a process of
insoluble matter transformation
that serves both as a physical
support and as a source of
nutrients in the absence or near-
absence of free water. However,
the substrate should be moist
enough to allow for growth and
metabolism of the microorganisms
(Singhania et al. 2009). Because
solid-state fermentation shows
great potential in metabolite
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62 Sheila Romo Sánchez, 2013
production (Holker et al. 2004), this
process is particularly useful in the
production of microbial products,
such as food, fuel oil, and industrial
chemicals and pharmaceuticals.
Accordingly, the waste produced
would be used as substrate in
solid-state fermentation (SSF) for
enzyme production (Couto and
Sanromán 2006).
Lignocellulosic enzymes
include cellulases, xylanases, and
ligninases, which are able to break
down the structure of cellulose,
hemicellulose, and lignin of plant
materials. Enzymes of the
cellulolytic system that degrade
cellulose altogether are exo-β-1,4-
glucanase, endo-β-1,4-glucanase,
and β-glucosidase (da Silva et al.
2005). Hemicellulose, the second
most abundant renewable biomass
in nature, is largely made up of
xylan (25-30% of dry weight of
plant cell wall). Xylanases are the
enzymes that can hydrolize xylan,
and they are of considerable
interest for their industrial
application (Soliman et al., 2012).
Pectinases, another group of
enzymes, degrade pectins, which
are structural polysaccharides
formed by units of polygalacturonic
acid and inclusions of other
monosaccharides located in the
middle lamella and in the primary
wall of higher plants (Kashyap et al.
2001).
Both molds and yeasts are
able to thrive in this type of
substrate thanks to their enzymatic
systems. Molds are particularly
interesting because they have a set
of advantages: (i) they play a major
role in a number of food industries;
(ii) they excrete enzymes of a
different nature that allow molds
to metabolize complex mixtures of
organic compounds found in most
residues (Jin et al. 2002); (iii) their
filamentous morphology or pellet
guarantees enzyme separation and
production at a low cost (da Silva
et al. 2005; Salihu et al. 2012).
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Sheila Romo Sánchez, 2013 63
The aim of this study is to
produce β-glucosidase, cellulase,
xylanase, and pectinase by solid-
state fermentation of byproducts,
considering factors such as
composition of the medium, milling
degree, and microbial strain.
Enzymes were purified, lyophilized,
and immobilized for industrial
application of the enzymatic
extract obtained.
Materials and Methods
Microorganisms
After initial experiments for
strain screening, two local fungal
strains, Aspergillus niger (strain Nº
113N) and Aspergillus fumigatus
(strain Nº 3), were selected (both
deposited in the GenBank
Database with accession numbers
FJ499449 and FJ499462). These
strains were isolated from olive
paste and olives, respectively, in
the Yeast Biotechnology Laboratory
at the University of Castilla-La
Mancha (Spain) (Alves-Baffi et al.
2012). Molds were maintained
using the method suggested by
Castellani.
A. niger and A. fumigatus
were deposited in the Spanish Type
Culture Collection (CECT) whose
accession numbers are 20828 and
20827 respectively
(http://www.cect.org).
Natural substrates
Grape skin (S) and olive
pomace (P) were the byproducts
collected locally from agro-
industries, as well as the wheat (W)
used as a medium supplement. The
grape skins and olive pomace were
air dried at 40 ºC / 24 h, and
separated into two series, one of
which was milled with an ultra-
centrifugal mill (Retsch ZM200)
using a sieve with a pore size of 0.5
µm. The other series was not
milled.
The chemical composition
of S and P was analyzed,
quantifying moisture, starch, lignin,
cellulose, hemicellulose, pectin, fat,
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64 Sheila Romo Sánchez, 2013
and ash percentage (Van Soest;
PNT-LACC/FQ 002; Gravimetry;
AOAC 1997, 985.29; Regulation
152/2009 Annex II H Proceeding B;
Regulation (EC) 152/2009 Annex III
A).
Solid-state fermentation and
enzyme production
To obtain the precultures,
the Aspergillus niger and
Aspergillus fumigatus strains were
grown in 250 mL Erlenmeyer flasks
containing 50 mL PDA medium
(agar slant) at 30 ºC / 65 %
moisture for 7 days. Then, 100 mL
of the basal medium (0.3% KH2PO4;
0.35 % (NH4)2SO4; 0.05 %
MgSO4·7H2O; 0.05 % CaCl2) was
added to culture, and slightly
scratched to obtain mycelial
suspension.
The design of SSF medium
was prepared according to da Silva
et al. (2005) with slight
modifications. 5 mL of suspension
of each mycelium were transferred
to 100 mL Erlenmeyer flasks
containing 5 g of solid substrate to
be tested (S or P).
As many as 8 different
fermentations were run for each of
the molds, depending on whether
or not wheat was added and the
by-products were milled.
S: unmilled grape skin
SW: unmilled grape skin (2.5 g) +
wheat (2.5 g)
mS: milled grape skin
mSW: milled grape skin (2.5 g) +
wheat (2.5 g)
P: olive pomace
PW: olive pomace (2.5 g) + wheat
(2.5g)
mP: milled olive pomace
mPW: milled olive pomace (2.5 g)
+ wheat (2.5g)
Enzyme production was
sampled after 2, 4, 6, 10, and 15
days for assessment over time. The
flasks were incubated at 25 ºC / 65
% moisture during the time period
set. Afterwards, 100 mL of distilled
water was added to the flasks and
they were maintained under
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Sheila Romo Sánchez, 2013 65
shaking for 30’ at room
temperature for extracellular
enzyme extraction. The content of
the flasks was centrifuged and the
supernatant was maintained at -20
ºC until analysis.
Enzyme assays
Activity of the enzymes
pectinase, xylanase, cellulase, and
β-glucosidase was assessed. 0.1 mL
of the extract was incubated at
37ºC / 30’ with 0.4 mL of the
corresponding substrate in 0.1 M
acetate buffer, pH 5: 0.5% xylan
from beechwood (Sigma) for
xylanase; 1%
carboxymethylcellulose (CMC –
Panreac) for cellulase; 0.5% pectin
from apple (Sigma) for pectinase;
and 1% D-(+)-cellobiose (Fluka) for
β-glucosidase. Following the
incubation period, the reducing
sugars released were measured by
using the 3,5-dinitrosalicylic acid
(DNSA) method (Miller 1959).
Assessment of β-glucosidase
activity was based on the protocol
suggested by Arévalo-Villena et al.
(2006a).
Parallelly, blank
absorbance was measured in the
same way as the samples but with
no incubation (at 0 time). For
reducing sugar measurement,
standard curves of galacturonic
acid, xylose, and glucose were
used, depending on the enzyme to
be assessed.
Assays were performed in
triplicate in all cases and outcomes
were calculated as enzyme units
(U) defined as µmol of released
product per minute under reaction
conditions.
Purification and lyophilization of
enzymes
A purified extract was
obtained after solid-state
fermentation of the mold-substrate
pair that yielded the best results at
the previous stages of the study.
The multi-enzymatic extract
underwent a variety of
technological processes, and its
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66 Sheila Romo Sánchez, 2013
activity was monitored at each of
the stages.
Supernatant was purified
by size exclusion using filter
cartridges designed for the
concentration and purification of
small peptides, oligonucleotides,
nucleic acids, enzymes, antibodies,
and other molecules (Pall, New
York, USA). The molecule size
selected was 100 KDa, considered
best suitable for excreted protein
(Palacios et al.; Ncube et al. 2002;
Krisana et al. 2005; Arévalo-Villena
et al. 2007). Samples were run at
4ºC for 1h at 4500 rpm, and the
enzymatic activity was monitored
(Arévalo-Villena et al. 2006b).
The purified extract, quick
frozen at -80ºC, was subsequently
lyophilized at pressures of 2*10-2
and -53ºC without cryoprotectant
and with 20% skimmed milk
(unpublished data by García-López
and Uruburu-Fernández).
The state of proteins of the
purified and lyophilized extract was
monitored by SDS-PAGE
electrophoresis in sulfate-
polyacrylamide gel (Bio-Rad
CriterionTM XT Precast Gel 12%
Bis-Tris) using a 8-220 Kda
molecular weight marker (MW
Sigma ColorBurst). The staining was
performed with Coomassie Blue.
Multi-enzymatic extract
immovilization
The purified and lyophilized
enzymes (with and without
cryoprotectant) were immobilized
on chitosan by physical adsorption.
The immobilization conditions,
which were established based on
previous research (Romo et al.
2012), involved an enzymatic
concentration of 0.048 g/mL in
acetate buffer pH 5, 50mM in a
final concentration of 50 mL. The
reaction was maintained for 2h at
10ºC under gentle shaking.
Afterwards, up to 4 washes were
performed with the same buffer to
eliminate the unintegrated protein
and cryoprotectant.
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Sheila Romo Sánchez, 2013 67
Xylanase activity of the
enzymatic extracts was monitored
in triplicate prior to and after each
biotechnological process following
the previously described protocol.
RESULTS
Natural substrates
Table 1 shows the
composition of each of the
lignocellulosic substrates used in
this study. The main constituents in
both cases were lignins and
celluloses, which are a
potential adequate source of
carbon for polysaccharide-
hydrolyzing microorganisms.
However, the values of
nitrogenized content were low,
which would a priori hinder the
development of the molds. For this
reason, some fermentations were
supplemented with wheat. The
differences between both by-
products lay in the high fat content
and the higher percentage of
pectins found in olive pomace.
Table 1. Raw material composition (%) in grape skins and olive pomace
Composition Grape skins Olive pomace
Humidity 5.58 3.43
Starch 2.52 1.35
Lignin 22.48 22.6
Cellulose 25.78 22.96
Hemicellulose 3.66 5.89
Pectin 1.27 5.82
Fat 9.84 15.92
Ash 6.48 3.3
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68 Sheila Romo Sánchez, 2013
Solid-state fermentation and
enzyme production
Once their composition
was determined, the by-products
were used for mold growth and
enzyme production by SSF. The
combinations assayed are shown
below.
S: unmilled grape skin
SW: unmilled grape skin (2.5 g) +
wheat (2.5 g)
mS: milled grape skin
mSW: milled grape skin (2.5 g) +
wheat (2.5 g)
P: olive pomace PW: olive pomace (2.5
g) + wheat (2.5g)
mP: milled olive pomace
mPW: milled olive pomace (2.5 g) +
wheat (2.5g)
Fig. 1 shows the values of the
enzymatic activities (cellulase,
pectinase, β-glucosidase, and
xylanase) obtained on grape skins
with or without wheat supplement
during SSF
Fermentation on unmilled
substrates was beneficial both for
A. niger and A. fumigatus. A
significant increase in activity was
observed from the sixth day of
fermentation, reaching its
maximum rate on the tenth and
thirteenth days. This is likely due to
the very metabolic development of
the microorganisms. Upon
comparison of the enzymatic
activities of both molds, A. niger
was found to yield the best results
in all cases, even showing a
significant activity in grape skins
with no wheat supplement.
Xylanase reached 47.05 U per g of
substrate on the fifteenth day of
fermentation on S. β-glucosidase
also showed a rich activity on the
tenth day (29.47 U / g), although
this only occurred when the grape
skins were supplemented with
wheat (SW).
The enzymatic activity of A.
fumigatus was significantly poorer
in all cases. Even though the
highest activity rate was reported
on β-glucosidase, a great
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Sheila Romo Sánchez, 2013 69
difference was found between this
rate and that reported on A. niger
(10.02 U versus 4.17 U). This only
occurred when wheat was added,
which might be indicative of a
greater need of A. fumigatus for
nitrogenized material.
Fig. 1. Enzymatic activity of A. fumigatus and A. niger in solid-state fermentation over time (days). S: grape skins; SW: grape skins and wheat; mS: milled grape skins; mSW: milled grape skins and wheat.
Artículo III
70 Sheila Romo Sánchez, 2013
Fig. 2 includes the figures
of fermentation with olive pomace
and wheat. The enzymatic activity
rate of both molds was higher than
that of grape skins, particularly in
the case of A. niger. The maximum
activity values were reported on A.
fumigatus on the fourth day of
growth for xylanase production (8
U). Virtually all cases required
wheat supplementation and
unmilled substrates, even for mold
growth (none of the molds was
able to grow during milled olive
pomace fermentation; data not
shown).
Fig. 2. Enzymatic activity of A. fumigatus and A. niger in solid-state fermentation over time (days). P: olive pomace; PW: olive pomace and wheat.
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Sheila Romo Sánchez, 2013 71
Enzyme purification, lyophilization
and immobilization
Based on the previous
results, the microorganism-
substrate pair selected was A.
niger-unmilled grape skins for SSF
for 15 days.
The enzymatic extract
obtained was purified and
lyophilized according to the
protocols described in the
materials and methods section.
Both protein size and
concentration and xylanase activity
were monitored because this was
the most frequently excreted
enzyme (Fig. 1).
Protein electrophoresis of
the raw and purified-lyophilized
enzymatic extracts of A. niger is
shown in Fig. 3. Although the
proteic patterns of both extracts
were similar, the pattern of the
purified-lyophilized extract shows a
lower activity of the enzymes in
both cases. This fact preluded
almost total loss of proteins, and
thus, of the enzymatic activity.
Fig. 3. Zymogram of the multi-enzymatic systems of A. niger, cultured on grape skins. Lane M: molecular weight marker; Lane 1: non-purified multi-enzymatic system; Lane 2: purified and lyophilized multi-enzymatic system.
For data confirmation, the
enzymes’ capacity for hydrolysis
was measured using xylan as
substrate (Table 2). Protein
purification (pore size 100 KDa)
held back 83% of enzymatic
activity, which proved the cut size
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72 Sheila Romo Sánchez, 2013
selected right. Nevertheless, the
process of lyophilization had a
devastating impact on the activity
(almost 90% of it was lost). The use
of the cryoprotectant could not
prevent denaturalization, as shown
by the low percentages reported
(16.4 and 13.0% with and without
cryoprotectant,
respectively).
Finally, immobilization of the
lyophil led to almost total loss of
the activity (less than 10% in both
cases, with and without
cryoprotectant).
Table 2. Xylanase activity percentage (%) of the enzymatic extracts of raw, purified,
lyphilized, and immobilized A. niger.
DISCUSSION
This study relied on two
molds in the genus Aspergillus
(A. niger and A. fumigatus),
which were isolated from oleic
environments. Solid-state
fermentations were run on two
substrate types, i.e. grape skins
and olive pomace, for industrial-
use enzyme production. The
analysis of the composition of
both substrates showed that their
main constituents are lignins and
celluloses, which is consistent
with the data reported in the
literature. According to
Georgieva and Ahring (2007),
olive pomace is largely made of
lignin (22.8%), a percentage that
is very close to that reported in
our study (22.6%). Cara et al.
(2008) determined the
composition of raw materials of
olive trees, reporting dry
Raw extract
Purified extract
Lyophilized extract Immobilized extract
100.0 ± 1.3
82.8 ± 0.3
With cryoprotectant
Without cryoprotectant
With cryoprotectant
Without cryoprotectant
16.4 ± 2.6 13.0 ± 0.1 5.4 ± 0.10 0.03 ± 0.0
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Sheila Romo Sánchez, 2013 73
biomass percentages slightly
higher than those reported by
our study (25% in cellulose and
15.8% in hemicellulose). This
difference in percentage may be
due to the mixture of substrates
with other types of materials
from the trees themselves. As for
grape skins, the data found in the
literature differ significantly
from those reported by the
present study (González-
Centeno et al. 2010).
Enzyme production using
filamentous molds for solid-state
fermentation or submerged
fermentation is a widely used
biotechnological method. Selection
of a fermentation type depends on
the physiological adaptation of the
organism. The growth pattern of
filamentous molds in submerged
culture usually ranges between
pellet-like and filamentous, which
may affect enzymatic production
(Mitchell and Lonsane 1992).
Previous studies conducted in our
laboratory with submerged culture
(Alves-Baffi et al. 2012) reported
no activity of pectinase and β-
glucosidase. In contrast, the same
molds cultured in SSF in the
present study did show activity
(Fig. 1). This is due to the fact that
SSF provides the molds with a
closed environment similar to their
natural habitat, which stimulates
them to produce more
hemicellulolytic enzymes (da Silva
et al. 2005).
For all four enzymes
examined, wheat supplement had
an impact both on A. niger and A.
fumigatus because of the easily
assimilable nitrogen that wheat
provides and the increase in
fermentation area during the
process. This increase is associated
with the particle size of substrates.
According to Salihu et al. (2012),
this is one of the factors that most
affect fermentation, together with
pre-treatment and water retention
capacity.
Artículo III
74 Sheila Romo Sánchez, 2013
In resume, the study
provides evidence of the
appropriateness of grape skins and
olive pomace for enzyme
production, and thus, the validity
of these byproducts for this
purpose is reinforced. The
combination A. niger-grape skin
(strain-substrate) yielded optimal
results, being xylanase the most
productive enzyme. Milling of both
substrates did not improve enzyme
production in any case. While not
necessary for grape skins, wheat
supplement stimulated microbial
growth and enzyme production in
olive pomace.
Selection of cut size for
purification (100 kDa) was based
on the size of enzymes under
examination: β-glucosidase, 100
kDa (Arévalo-Villena et al. 2007);
xylanase, 31 kDa (Ncube et al.
2002); pectinase, between 25 and
50 kDa (Palacios et al.); and
cellulase, 21 kDa (Krisana et al.
2005). The outcomes were
consistent with previous research,
since the purification process
yielded an enzymatic extract with
almost 83% of residual activity.
The lyophilization process,
however, did not meet
expectations, not even with the
help of a cryoprotectant. There are
many compounds that can be used
to this aim (glycerol, skimmed milk,
dimethyl sulfoxide, and
carbonhydrates, such as glucose,
lactose, saccharose, and inositol).
Further research with other
cryoprotectant agents is thus
needed for yield improvement.
The results yielded in our study for
adsorption-induced multi-
enzymatic extract immobilization
on chitosan are not representative,
nor do they prove this process right
or wrong. The reason for this is
that the residual activity of the
lyophil used was much too low
(activity dropped more than 80%).
Although there is no research
addressing immobilization of multi-
Artículo III
Sheila Romo Sánchez, 2013 75
enzymatic systems, there are
studies dealing with coupling
between two enzymes. The
objective of such studies is either
to obtain a final product or to
eliminate a by-product of one of
the two enzymes for being
contaminating or detrimental to
the process (van de Velde et al.
2000). Accordingly, some of these
biological catalysts would be able
to prevent others from coupling on
the substrate. Currently, there is an
interest in the development of
techniques that guarantee enzyme
immobilization on a support on a
specific area, revealing significant
zones for stabilization. All these
data pave the way for future
studies including lyophilization
assays with different
cryoprotectants and/or assays
involving immobilization of the
purified extract without prior
lyophilisation, which might be the
key to the biotechnological
advance of this topic.
Aknowledgements
The authors wish to express their
gratitude to Dr. Mario Canales for
protein electrophoresis assistant
and to Dr. Hector L. Ramirez for
matrix for immobilization transfer.
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Artículo IV
82 Sheila Romo Sánchez, 2013
Immobilization of commercial cellulase and xylanase on different polymer
supports (alginate-chitin and chitosan-chitin) by different methods
Sheila Romo-Sánchez1, Conrado Camacho2, Héctor L. Ramirez2 and María
Arévalo-Villena1 1 Food Science and Technology. University of Castilla la Mancha (UCLM), Av. Camilo
José Cela, 13071 Ciudad Real, Spain 2 Biotechnology Center. University of Matanzas, Cuba
*Corresponding author: María Arévalo Villena
Abstract. Industrial applications of cellulase and xylanase require enzymes that
are highly stable, able to function over a wide pH and temperature range, and
economically viable in terms of reusability. This study sought to optimize
immobilization conditions for two commercial enzymes (cellulase and
xylanase) on a range of polymers (alginate-chitin and chitosan-chitin) using
three chemical methods (adsorption, reticulation, and crosslinking-
adsorption). The best results were obtained using chitosan, a support that may
display a number of valuable features including biocompatibility,
biodegradability, lack of toxicity, antibacterial properties, chelation of heavy
metal ions, hydrophobicity and good protein binding. The optimum pH for
binding was 4.5 for cellulase and 5.0 for xylanase, while the optimal enzyme
concentrations were 170 µg/mL and 127.5 µg/mL, respectively. In some cases,
the use of a low concentration of crosslinking agent (glutaraldehyde) was
found to improve stability of the immobilization process. The most striking
finding was the reusability of enzymes, and particularly of immobilized
cellulase using glutaraldehyde, which after 19 cycles retained 64% activity.
These results confirm the economic and biotechnical advantages of enzyme
immobilization for a range of industrial applications.
Artículo IV
Sheila Romo Sánchez, 2013 83
Keywords: immobilization, cellulase, xylanase, crosslinking-adsorption,
glutaraldehyde, reticulation
Introduction
Enzymes have enormous potential
as industrial catalysts, largely
because they are substrate-specific
and easy to produce. Their use is
becoming increasingly widespread,
especially in the biological and
chemical industries. Hydrolytic
enzymes, for example, are widely
used in the textile, pulp and paper
industries [1]. The engineering of
enzymes for this purpose has been
hailed as one of the most exciting,
complex and interdisciplinary goals
of biotechnology. Despite their
large-scale commercial availability,
however, these biocatalysts still
have certain drawbacks, chief
among which is their extremely
limited reusability [2]. Enzyme
immobilization is a technology
aimed at enhancing the stability of
enzyme-related processes [3], with
a view to enabling continuous
processing through the reuse of
enzymes [4].
Immobilized enzymes can be
defined as “enzymes physically
confined or localized in a certain
defined region of space with
retention of their catalytic
activities, which can be used
repeatedly and continuously” [5].
In bioaffinity immobilization, the
enzyme/protein is immobilized via
bioaffinity interactions [6].
Immobilization enables continuous
economic operation, automation
and the recovery of product with a
high degree of purity [7]. For that
reason, there is a growing
industrial demand for immobilized
biocatalysts.
Immobilized cellulase and xylanase
are widely used in the
biotechnology industry, among
other things for clarifying juices
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84 Sheila Romo Sánchez, 2013
and wines, for extracting plant oils
and coffee, for the bioconversion
of agricultural waste [8], and for
improving the digestibility of
animal feed ingredients [9]. A
major application at present is in
the biodegradation or
bioconversion of cellulose- and
hemicellulose-containing materials
to monomeric sugars [10].
Agricultural waste rich in
lignocellulosic material could be
used in manufacturing a whole
range of commercial products
including ethanol [11], organic
acids [12] and, if the process were
economically competitive, other
chemical products [13].
The present study investigated the
immobilization of cellulase and
xylanase on a range of supports,
analyzing the behavior of various
biochemical parameters and
comparing it to that of the native
enzymes.
Materials and Methods
Materials
Chitin from lobster shells (degree
of deacetylation = 10% [14],
average particle size = 30 µm) was
obtained from Empresa Mario
Muñoz (Havana, Cuba). Chitosan
was obtained by alkaline
deacetylation of chitin [15]; the
degree of deacetylation was 90%
[14]. Sodium alginate extracted
from Laminaria hyperborea was
purchased from BDH (Poole, UK).
Molecular weight was 1.97×105.
Commercially-available soluble
xylanase and cellulase were
purchased from Novozymes.
Glutaraldehyde (50%, w/v) was
obtained from SIGMA (St.Louis,
MO, USA). All other chemicals were
of analytical grade.
Preparation of carriers
Alginate-Chitin: 600 mg of alginate
was dispersed in 60 ml of
potassium-phosphate buffer (pH
Artículo IV
Sheila Romo Sánchez, 2013 85
6.0) to which 150 mg of 1-ethyl-3-
(3-dimethylaminopropyl)
carbodiimide (EDAC) was then
added. The reaction was
maintained at room temperature
for 1 h with continuous stirring.
Activated alginate was mixed with
3 g of a chitin solution (w/v),
dissolved in 30 mL distilled water
and stirred for 16 h at 25 ºC.
Chitosan-Chitin: 1 g of chitin was
dispersed in 10 ml distilled water,
and glutaraldehyde was added to a
final concentration of 5% (v/v). The
reaction was maintained at 25 ◦C
for 4 h with continuous stirring.
The solid was collected by
centrifugation, washed several
times with distilled water until no
aldehyde was detected in waste,
and finally suspended in 25 ml of
distilled water. Activated chitin was
mixed with a 1% chitosan solution
(w/v) dissolved in 3% acetic acid
(v/v) and stirred for 4 - 6 h. NaBH4
was then added to a final
concentration of 200 mM, and the
solution was stirred for 16 h. The
solid was collected by filtration and
washed several times with distilled
water.
Both carriers were collected by
centrifugation, washed several
times with distilled water and dried
in vacuum drying apparatus until
their use for immobilization by
adsorption and reticulation
(adsorption-crosslinking). The
other immobilization method
(crosslinking-adsorption) required
additional preparation of carrier
using a bifunctional reagent
(glutaraldehyde), which can block
amino groups and render
polysaccharide structures (alginate
or chitosan) more inert and
resistant to the acid medium [16].
Preparation of crosslinked
chitosan-chitin carrier: one gram of
chitosan-chitin pre-treated carrier
was added to 20 mL of two chosen
glutaraldehyde concentrations
(15% (v/v) and 0.5% (v/v)) and
dissolved in 200 mM phosphate
Artículo IV
86 Sheila Romo Sánchez, 2013
buffer (pH 7.0). The mixture was
stirred in the dark at 25 ºC for 1
hour (support activated with 0.5 %
glutaraldehyde) or 15 h (support
activated with 15 %
glutaraldehyde). The crosslinked
chitosan carrier was centrifuged
and washed several times with
optimal immobilization buffer.
Optimal conditions for immobilized
enzymes
The methods employed for enzyme
immobilization are outlined below.
The first step was to optimize
enzyme-support conditions.
Adsorption
For this method, 10 mL of support
(alginate-chitin and chitosan-chitin)
at a concentration of 0.03 g/mL
were suspended in the relevant
buffer (depending on the pH used)
to a final volume of 40 mL. The
following variables were optimized:
- pH: a fixed enzyme
concentration was tested (170
µg/mL) with pH values ranging
from 2.5 to 5.5 [17], using 50
mM citric acid/Na2HPO4 (pH 2.5)
and 50 mM sodium
acetate/acetic acid (pH 3.0-5.5).
A total of 7 immobilization
reactions were obtained per
support and enzyme.
- Enzyme concentration: different
concentrations of enzyme were
tested (8.5 µg/mL, 17 µg/mL,
42.5 µg/mL, 84 µg/mL, 127.5
µg/mL, 170 µg/mL and 340
µg/mL), obtaining 7
immobilization reactions per
enzyme at the optimal pH.
Both assays were kept in darkness
at 10 ºC for 16 h, with continuous
gentle stirring.
- Binding time: using the two
optimized parameters (pH and
enzyme concentration), the two
enzymes were then tested over
a range of enzyme-support
binding times (20, 40, 60, 90,
120, 150, 180, 210, 210 and 240
min), yielding a total of 9
reactions for each enzyme.
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Sheila Romo Sánchez, 2013 87
Reticulation (Adsorption-
crosslinking)
After optimizing adsorption
conditions for the chitin-chitosan
support, immobilization was
performed by reticulation, by
adding glutaraldehyde to the
chitosan-enzyme system at 5
different concentrations (from
0.125% to 1.5%) for 0.5 h. [18].
Crosslinking-adsorption
Having prepared the crosslinked
chitosan carrier, immobilization by
adsorption was repeated under
previously-optimized conditions.
In all cases, after immobilization
the suspension was collected by
centrifugation and repeatedly
washed with 50 mM of the buffer
used for immobilization. Excess
glutaraldehyde (reticulation
method) or protein (all methods)
was removed.
Measurement of cellulase /
xylanase activity, and retained
protein
Once the enzyme was bound to the
support, enzyme activity was
checked. The reaction mixture
comprised 100 μl of enzyme
(cellulase or xylanase) solution and
400 μl of 1% (w/v)
carboxymethylcellulose (CMC) or
xylan substrate, respectively, in 50
mM acetate buffer (pH 5.5). The
mixture was incubated for 30 min
at 37 ºC and reducing sugars were
measured using the dinitrosalicylic
acid method [19], using glucose
and xylose, respectively, as
standards. Assays were performed
in triplicate. One unit of enzyme
activity (U) was defined as the
amount of enzyme required to
release one mol of glucose per
minute under these assay
conditions.
Native and immobilized enzyme
activity recovery was calculated as:
activity recovery (%) = (total
activity of enzyme / maximum total
activity of enzyme) *100%.
Artículo IV
88 Sheila Romo Sánchez, 2013
Absorbed protein was calculated as
the difference between added
protein and unbound protein
detected in washing solutions after
immobilization. Concentrations
were calculated using the Bradford
method [20], with bovine serum
albumin (BSA) as standard.
Biotechnological characterization
of free and immobilized enzymes
Having established the optimal
conditions for immobilization, the
influence of pH, temperature and
storage stability on enzyme activity
was determined; kinetic constants
and degree of reusability were also
investigated. To determine pH
stability, cellulase and xylanase
were incubated with CMC and
xylan, respectively at 37 ºC for 30
minutes in different buffers (100
mM citric acid/Na2HPO4 pH 2.0 -
2.5; 100 mM sodium acetate/acetic
acid pH 3.0 - 6.0; 100 mM
Na2HPO4/ NaH2PO4 pH 6.5 – 8.0).
using 50 mM (pH 2.5) and 50 mM
(pH 3.0-5.5). Heat stability was
determined by measuring residual
activity after preincubating the
enzymes for 10 minutes at
different temperatures (from 40 to
90 ºC at 5 ºC intervals). Aliquots
were chilled quickly and cellulase
and xylanase activity was
measured [17].
The kinetic properties Vmax
( mol∙min-1∙mg-1) and Km (mM)
were determined by measuring
cellulase and xylanase activity
using various CMC and xylan
concentrations (0.025 % to 0.5%).
Values for Vmax and Km were
calculated using Michaelis-Menten
plots [21].
Finally, the reusability of
immobilized enzymes was
determined by running consecutive
cycles on CMC and xylan (cellulase
and xylanase, respectively). All
reactions were maintained at 37 ºC
for 30 minutes, applying in all cases
the previously-identified optimal
Artículo IV
Sheila Romo Sánchez, 2013 89
conditions. After each reaction,
cellulase and xylanase were
washed with the appropriate
buffer. The activity of the
immobilized enzyme was
expressed as percentage residual
activity.
Statistical analysis
The statistical significance of the
effect of free and immobilized
enzymes in each assay, obtained in
triplicate, was determined by one-
way analysis of variance (ANOVA,
version 12.9). Significant
differences in results for each
variable (pH, T, etc) and each type
of enzyme (unbound and
immobilized) were also analyzed.
Results
In all cases, results were expressed
as percentage relative activity,
taking 100% as the maximum
capacity for each assay.
Selection of immobilization method
Chitosan-chitin proved to be the
best carrier for immobilizing both
cellulase and xylanase. Results
obtained on chitin-alginate were
unsatisfactory, although xylanase
kept linked at pH 2.5 to 4, the
protein concentration of
immobilized enzyme was very low;
cellulase immobilization only was
possible at pH 3, so this support
was discarded in both cases. The
best immobilization methods were
adsorption and reticulation, while
crosslinking-adsorption proved less
effective with both enzymes (data
not shown).
Optimal conditions for enzyme
immobilization
Conditions for the immobilization
of commercial cellulase and
xylanase are shown in Figure 1.
Testing the pH range from 2.5 to
5.5, maximum relative activity
(100%) was achieved at pH 5.0 for
xylanase and pH 4.5 for cellulase
Artículo IV
90 Sheila Romo Sánchez, 2013
(in 50 mM sodium acetate/acetic
acid buffer; Figure 1a).
The optimal enzyme concentration
(tested over the range 8.5 to 340
µg/mL) was found to be 127.5 for
xylanase and 170 µg/mL for
cellulase (Figure 1b). The optimal
enzyme-support binding time for
immobilization proved to be 2 h for
xylanase and 2.5 h for cellulase
(data not shown). Once these
parameters had been optimized,
they were applied for
immobilization by reticulation with
glutaraldehyde. The most effective
glutaraldehyde concentration was
0.125; higher values prompted
reduced activity, especially for
cellulose (Figure 1c).
Figure 1. Effect of different conditions on the activity of immobilized cellulose (C) and xylanase (X). (a) optimum pH; (b) optimum enzyme concentration; (c) optimum glutaraldehyde concentration. Relative activity was calculated by taking maximum enzyme activity as 100% in each case.
Biochemical characterization of
free and immobilized enzymes
The pH stability of free and
immobilized enzymes was
determined by carrying out the
enzyme assay at different pH
values (pH 2.0 – 8.0). The optimum
pH curves for both enzymes are
(a)
(b)
(c)
Artículo IV
Sheila Romo Sánchez, 2013 91
shown in Table 1a. Xylanase
displayed good stability over the
pH range from 3.0 to 8.0; both
native and immobilized enzyme
achieved 100% activity at pH 6.0.
By contrast, cellulase displayed
good activity only at acidic pH
values (from 2.0 to 4.0), with
maximum activity at pH 3.0
(immobilized cellulase) and pH 4.0
(native cellulase). Higher pH values
reduced activity in all three cases.
Enzyme heat stability is charted in
Table 1b. Immobilization of
cellulase, both by adsorption
and by reticulation, prompted
greater resistance to temperature
increases. After 10 minutes at 75
ºC, the activity of the native
enzyme had dropped to 3.7%,
whereas immobilized enzyme
retained around 50% of its original
activity. By contrast, native
xylanase displayed greater heat
resistance than immobilized
enzyme at 75ºC, while the reverse
was true at 55 ºC (97.8%
adsorption; 91.6% reticulation;
88.7% free).
Table 1(a). Effect of pH on xylanase and cellulase activity (%) using different buffers (pH 2.0-8.0)
pH FX AX RX FC AC RC
2.0 1.5 ± 0 a
0 ± 0 a
0 ± 0 a
87.7 ± 5 a 87.9 ± 9
a 39.2 ± 0
b
3.0 75.3 ± 1 a
76.6 ± 3 a
76.3 ± 5 a
91.9 ± 5 a
100 ± 7 a
100 ± 11 a
4.0 87.9 ± 3 a
76.7 ± 10 a 79.0 ± 3
a 100 ± 9
a 88.1 ± 5
a 88.5 ± 10
a
5.0 95.3 ± 3 a
95.5 ± 4 a
97.0 ± 1 a
68.8 ± 7 a
55.5 ± 3 a
63.9 ±9 a
6.0 100 ± 1 a
100 ± 3 a
100 ± 4 a
42.9 ± 5 a
62.7 ± 0 b
0 ± 0 c
7.0 86.9 ± 0 a
84.7 ± 4 a
80.6 ± 2 a
40.6 ± 1 a
11.9 ± 2 b
0 ± 0 c
8.0 77.6 ± 5 a
78.3 ± 2 a
78.8 ± 1 a
25.8 ± 1 a
4.3 ± 6 b
0 ± 0 b
Table 1(b). Effect of temperature (40ºC – 90ºC) on xylanase and cellulase activity (%)
Artículo IV
92 Sheila Romo Sánchez, 2013
Temp FX XA XR FC CA CR
40 100 ± 2 a
100 ± 0 a
99.9 ± 0 a
97.0 ± 0 a
98.5 ± 1 a
89.2 ± 0 b
45 98.9 ± 2
a 99.6 ± 1
a 100 ± 2
a 98.9 ± 1
a 99.6 ± 1
a 90 ± 2
b
50 99.3 ± 1 a
99.9 ± 1 a
99.2 ± 1 a
100 ± 5a 100 ± 4
a 90.2 ± 15
a
55 88.7 ± 1 a
97.8 ± 0 b
91.6 ± 3 a
96.1 ± 3 a
99.6 ± 3 a
87.3 ± 4 b
60 86.7 ± 6 a
83.1 ± 8 a
84.8 ± 1 a
96.3 ± 4 a
95.3 ± 7 a
100 ± 13 a
65 77.2 ± 12 a
79.2 ± 2 a
83.0 ± 1 a
92.1 ± 0 a
88.2 ± 3 a
90.3 ± 9 a
70 76.2 ± 3 b
73.8 ± 2 a, b
69.0 ± 4 a
70.2 ± 5 a
69.6 ± 7 a
78.3 ± 0 b
75 68.1 ± 1 a
39.3 ± 1 b
26.3 ± 2 c 3.7 ± 2
b 64.3 ± 8
a 46.8 ± 6
a
80 68.1 ± 1 b
19.9 ± 3 a
15.0 ± 3 a
0 ± 0 b
41.7 ± 6 a
51.1 ± 9 a
85 61.5 ± 3 b
14.2 ± 1 a
11.1 ± 1 a
0 ± 0 a
0 ± 1 a
0 ± 1 a
90 59.9 ± 1 a
10.6 ± 1 b
8.0 ± 1 c 0 ± 0
a 0 ± 1
a 0 ± 2
a
Relative activity was calculated by taking the maximum activity of each free or immobilized enzyme as 100%. FX: free xylanase; XA: xylanase immobilized by adsorption; XR: xylanase immobilized by reticulation; FC: free cellulase; CA: cellulase immobilized by adsorption: CR: cellulase immobilized by reticulation. Different letters indicate significant differences (95% confidence) among three enzymes (free, adsorbed and reticulated).
The kinetic constant (Km) and the
maximum reaction rate (Vmax) were
obtained from Michaelis Menten
plots. The kinetic behavior of
cellulase and xylanase was
modified by immobilization. Km, a
measure of the substrate’s affinity
for the enzyme, was higher for the
immobilized enzymes than for their
native counterparts (Table 2), while
values for Vmax were similar for
both types.
The relative activity of the two
immobilized enzymes over
consecutive cycles of use is shown
in Figure 2. In both cases, activity
diminished with reuse. Xylanase
could only be reused 8 times with
both types of immobilization,
achieving with the reticulated
enzyme a maximum of 25% on the
eighth cycle. By contrast,
reticulation-immobilized cellulase
retained up to 64% activity after 19
cycles, while the adsorption-
immobilized enzyme displayed only
32% of its initial activity after the
same number of cycles.
Artículo IV
Sheila Romo Sánchez, 2013 93
Table 2. Kinetic constants of native and immobilized cellulase and xylanase enzymes
Cellulase Xylanase
Vmáxb Km
a Vmáx
b Km
a
Free 1.99 6.56 1.595 0.083
Immobilized by adsorption 1.74 12 1.66 0.115
Immobilized by reticulation 1.94 11.5 1.27 0.095
a Michaelis constant, Km, was defined as concentration (mM) of substrate
b Maximum velocity, Vmáx refers to the substrate decomposition rate ( molmin
-1mg
-1)
Figure 2. Reusability of cellulase and xylanase immobilized by adsorption and reticulation. (a) XA: xylanase immobilized by adsorption; XR: xylanase immobilized by reticulation; (b) CA: cellulase immobilized by adsorption; CR: cellulase immobilized by reticulation. Data were expressed as means of three independent trials. Relative activities were calculated by taking maximum immobilized-enzyme activity as 100% in each case.
(a)
(b)
Artículo IV
94 Sheila Romo Sánchez, 2013
Discussion
In this study, two enzymes –
cellulase and xylanase – were
immobilized separately with a view
to improving their performance in
comparison with their native
counterparts. Optimal conditions
for the two enzymes were very
similar. The most effective support
proved to be chitosan, which has
additional advantages including
low cost, biocompatibility, good
hydrophobicity, high porosity, and
a large adhesion area. Moreover,
its structure ensures minimal steric
hindrance during immobilization
[22]. Also the optimal methods
used for immobilization in both
enzymes were adsorption and
reticulation, while crosslinking-
adsorption was. The glutaraldehide
concentration used to activate the
support (0.5% and 15% (v/v)) could
inhibit the enzyme, maybe in
future works it could by tried the
crosslinking-adsorption using lower
concentration of glutaraldehide.
Optimal pH was found to be 4.5 for
cellulase and 5.0 for xylanase,
reflecting the fact that polyanionic
matrices give rise to the
partitioning of protons between
the bulk phase and the enzyme
microenvironment, prompting
changes in the optimal pH value.
Changes depend on the
immobilization method used, and
on the structure and charge of the
matrix [23].
With regard to biochemical
properties, neither of the enzymes
was affected by changes in pH, and
there was no change in the pH
response of the enzymes after
immobilization. Xylanase, both free
and immobilized, displayed good
stability over the pH range from 3.0
to 8.0 and there was no significant
differences between the adsorbed,
reticulated and free xylanases as a
function of pH. The same fact was
observed by Pal and Khanum that
reported that xylanase covalently
immobilized on glutaraldehyde-
Artículo IV
Sheila Romo Sánchez, 2013 95
alginate beads displayed behavior
identical to that of the native
enzyme at the same pH values [24].
Cellulase, both free and
immobilized, proved stable only at
acidic pHs. One-way ANOVA
revealed significant differences
between the three forms of
cellulase at all pH values: at pH 3.0,
free cellulase displayed the least
hydrolysis, though it retained its
activity slightly better than
cellulase immobilized at pH 4.0. At
neutral values (pH 6.0),
reticulation-immobilized cellulase
lost all activity, whilst free and
adsorption-immobilized cellulase
retained around 50% and 60%,
respectively, of its initial activity.
Similar findings are reported by
Zhou [25] for cellulase immobilized
on N-succinyl-chitosan. In all cases,
changes in behavior as a function
of pH may depend on the charge
both of the enzyme and of the solid
support.
Both xylanase and cellulase proved
stable over the temperature range
from 40ºC to 70 ºC, and although
statistical analysis revealed
significant differences between
formats over that range, for
practical purposes these variations
were negligible. For cellulase, both
immobilization methods enhanced
heat stability, with results
significantly better than those of
the native enzyme at temperatures
of over 75ºC.
This enhanced thermostability is
attributed to the covalent binding
of cellulase to the copolymer [26].
In research using other matrices for
immobilization [27, 28], enzymes
did not remain stable over such a
wide temperature range. Akkaya et
al., have reported that optimal
temperatures for immobilized
enzyme may be higher, lower or
the same as for the native enzyme
[29].
With regard to kinetic parameters,
immobilization prompted an
Artículo IV
96 Sheila Romo Sánchez, 2013
increase in the value of Km, which
might be due to changes in the
accessibility of the substrate to the
active sites of the enzyme caused
by diffusional limitations, steric
effects and enzyme structural
changes following immobilization
[30]. A similar increase has been
noted in other studies [28].
Buchholz suggested that the
increase in Km upon immobilization
of xylanase could be due to a
conformational change in the
enzyme resulting in lower affinity
for the substrate [31].
Reusability is a major requirement
for industrial enzyme applications.
Xylanase displayed good activity up
to its eighth reuse; after six
consecutive cycles, the adsorption-
immobilized and reticulation-
immobilized enzyme retained 91%
and 81%, respectively, of its initial
activity. These findings are slightly
better than those reported by
Kapoor and Kuhad [32], who found
that immobilized xylanase retained
70% of its initial activity.
Cellulase was reused up to 19
times, retaining good activity
(32.09 % for adsorbed enzyme,
63.8% for reticulated enzyme).
These values are higher than those
obtained by Wu et al. [33], who
recorded only 36% residual activity
after 6 reuses.
Conclusions
Two commercial enzymes, xylanase
and cellulase, were immobilized on
different supports using diverse
chemical binding methods, in order
to optimize their use in an
industrial setting. The best results
were obtained using chitosan as
support and adsorption and
crosslinking with glutaraldehyde
for immobilization.
While biochemical analysis showed
that both enzymes could be
successfully reused, results for
immobilized cellulase using
glutaraldehyde as crosslinking
Artículo IV
Sheila Romo Sánchez, 2013 97
agent were particularly striking:
after 19 reuses, the enzyme
retained 64% of its initial activity.
These results confirm the economic
and biotechnical advantages of
enzyme immobilization, especially
regarding to the number of reuses,
which open the possibility of
different industrial applications.
Acknowledgements
The authors wish to express their
gratitude to Spanish Ministerio
Educación y Ciencia and INIA for
funding this research, under the
project “Inmovilización de enzimas
para su aplicaciónen la industria
agroalimentaria” (2010-COB-3763);
and to Castilla-La Mancha
University (Spain) for the grant to
C. Camacho and H Ramirez during
their studies at UCLM. On the other
hand, authors appreciate
discussions with Dra Ana Isabel
Briones Pérez and her help during
the work development.
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Artículo V
104 Sheila Romo Sánchez, 2013
Immobilization of β-glucosidase and its application for enhancement of
aroma precursors in Muscat wine
Short running head: -β-glucosidase immobilized-
Sheila Romo Sánchez1, María Arévalo Villena1*, Esteban García Romero2,
Héctor L. Ramirez3, Ana Briones Pérez1
1 Food Science and Technology. University of Castilla la Mancha (UCLM), Av. Camilo
José Cela, 13071 Ciudad Real, Spain
2 Instituto de la Vid y el Vino de Castilla la Mancha, Carretera de Albacete s/n, 13700
Tomelloso, Spain
3 Biotechnology Center. University of Matanzas, Cuba
Corresponding author: María Arévalo Villena
e-mail address: maria.arevalo@uclm.es
Abstract. Enzyme immobilization is becoming more widely practised in
biotechnology because of the advantages that this method brings. In this
study, commercial β-glucosidase was immobilized on diverse supports by using
different methods. It was found that the most appropriate matrix was chitosan
by adsorption and reticulation. The optimal immobilization conditions were pH
3.5, immobilization time 120 min, and concentration of crosslinker
glutaraldehyde 0.25%. Stability and resistance of the immobilized enzymes
were assessed, which revealed a number of advantages, such as a lower
enzyme dose required for immobilization (367 times lower than the free
enzyme dose recommended by the manufacturer), high stability over time,
and reusability. In vitro studies of cellobiose and in vivo studies of wine and
Artículo V
Sheila Romo Sánchez, 2013 105
aroma precursors isolated from grape must yielded similar outcomes with
respect to enzyme hydrolysis of free and immobilized proteins.
Keywords β-glucosidase; immobilization; chitosan; adsorption; reticulation;
aroma precursors
1. Introduction
The use of β-glucosidase has
gained momentum owing to its the
ability of this enzyme to catalyze
transglycosylation reactions. These
reactions are of interest to the
wine industry since they improve
the aroma of young wines (Jatinder
et al., 2007). Nowadays, the
aromatic profile of a wine is one of
the essential parameters for
quality determination.
Many potential aroma compounds
present in grapes are in the form of
nonvolatile, flavorless glycosides.
These conjugates have attracted
much interest as precursors of
aroma and flavor compounds in
wine. Numerous studies report the
presence of glycoside fractions of
terpenic, norisoprenoids, C6
alcohols or phenolic compounds in
wines and grapes (Ibarz et al.,
2006; Sánchez-Palomo et al.,
2007).
Reports indicate that not all
glycosides are present in all grape
varieties, and that concentrations
vary across varieties (Bayonove et
al., 1993). Major precursors include
structures such as β-D-
glucopyranoside, 6-O-α-L-
arabinosyl-β-D-glucopyranoside, 6-
O-α-L-arabinosyl- β-D-
glucopyranoside, 6-O-β-D-
apiofuranosyl-β-D-glucopyranoside
apiosylglycosides (Gunata et al.,
1988; Voirin et al., 1992).
Terpenes (linalool, geraniol,
nerol, citronellol, α-terpineol,
linalool oxide, etc.) are some of the
Artículo V
106 Sheila Romo Sánchez, 2013
major components that contribute
to the variety character of wine
aroma. In addition, grapes also
include compounds called “aroma
precursors”, the most active of
which are glycosides—mostly
linalool, nerol, and geraniol
(Palmeri and Spagna, 2007). These
compounds are usually present in
two fractions: (i) a free fraction,
which contributes directly to must
aroma; and (ii) a bound fraction,
which forms non aromatic
glycosides (these glycosides may
later release their aromatic load).
The bound fraction is quantitatively
more significant than the free
fraction so glycosidase-catalysed
hydrolysis releases the terpenes
responsible for the so highly prized
fruity aroma in white wines
(Arévalo-Villena et al., 2006b).
Immobilized enzymes are generally
not used in winemaking, and they
are directly added by the enologist.
These enzymes must be drawn
from the wine after taking effect.
On the other hand, enzyme reuse
through multiple cycles of batch
fermentation processes results in
significant cost savings. Moreover
immobilized enzymes can easily be
separated from the wine. A further
benefit when compared with free
enzymes in solution is that
immobilized enzymes are more
robust, and are often more stable
and resistant to environmental
changes (Krajewska, 2004). In
addition, immobilisation of β-
glucosidase enables an accurate
management of the conversion
degree, achieving a rapid and
controlled liberation of terpenes.
This favours a quick sale, while
preserving a fraction of bound
aromas to be released over time
(González-Pombo et al., 2011).
As a result of it all, the aim of
this study was to immobilize
commercial β-glucosidase on
diverse supports by using different
methods. Optimal conditions and
maximum activity were analyzed
Artículo V
Sheila Romo Sánchez, 2013 107
and subsequently used to optimize
the aroma-increasing effects of
isolated aroma precursors of
muscat grape must and of wines of
the same variety.
2. Materials and Methods
2.1. Materials
Sodium alginate and chitosan,
both mixed with chitin, were the
supports used for the
immobilization process. Chitin from
lobster shells (degree of
deacetylation 10% (Baxter et al.,
1992); mean particle size 30 µm)
was obtained from Empresa Mario
Muñoz (Havana, Cuba). Chitosan
was prepared by alkaline
deacetylation of chitin (Kurita et
al., 1977), which was 90% (Baxter
et al., 1992). Sodium alginate from
Laminaria hyperborea was
purchased from BDH (Poole, UK).
Other compounds used were
commercially available soluble β-
glucosidase (Lallemand) because it
is one of the most important
enzymes used in winemaking and
glutaraldehyde (50%) (w/v)
(SIGMA) as crosslinking agent. All
other chemicals were of analytical
grade.
2.2 Synthesis of supports
Chitosan and alginate-coated
chitin supports were used due to
the good references in previous
works (Darias & Villalonga, 2001;
Gómez et al., 2006a)
Alginate-Chitin: 600 mg of
alginate were dispersed in 60 mL of
potassium-phosphate buffer (pH
6.0) and 150 mg of 1-etil-3-(3-
dimetilaminopropil) carboiimide
(EDAC) were added. The reaction
was maintained at room
temperature for 1 h with
continuous stirring. Activated
alginate was mixed with 3 g of a
chitin solution (w/v) and was
dissolved in 30 mL distilled water
and stirred for 16 h at 25 ºC.
Chitosan-Chitin: 1 g of chitin
was dispersed in 10 mL distilled
water and glutaraldehyde was
Artículo V
108 Sheila Romo Sánchez, 2013
added to a final 5% concentration
(v/v). The reaction was maintained
at 25 ºC for 4 h with continuous
stirring. The solid was collected by
centrifugation, washed several
times with distilled water until no
aldehyde was detected in waste,
and finally suspended in 25 ml of
distilled water. Activated chitin was
mixed with a 1% chitosan solution
(w/v) dissolved in 3% acetic acid
(v/v) and stirred for 4-6 h. NaBH4
was then added to a final
concentration of 200 mM, and the
solution was stirred for 16 h.
After that, both supports were
collected by centrifugation, washed
several times with distilled water
and dried in vacuum drying
apparatus until their use for
immobilization by adsorption and
crosslinking (adsorption-
crosslinking). Other immobilization
method (crosslinking-adsorption)
required an additional preparation
of support.
Preparation of crosslinked chitosan
carrier: 1 g of pretreated chitosan
above was added into 20 mL of two
chosen glutaraldehyde solutions
(15% (v/v) and 0.5% (v/v)) and they
were dissolved in 200 mM
phosphate buffer (pH 7.0). The
mixture was stirred in the dark at
25 ºC for 1 hour (support activated
with 0.5 % glutaraldehyde) or 15 h
(support activated with 15 %
glutaraldehyde). The crosslinked
chitosan carrier was centrifuged
and washed several times with
optimal immobilization buffer
(Erzheng et al., 2010)
2.3. Immobilization of β-glucosidase
The methods employed for
enzyme immobilization are
described below. For optimization
of the conditions for enzyme-
support linked, an initial amount of
10 mL of each matrix at
concentration of 0.03 g/mL was
prepared. β-glucosidase was then
added to a final volume of 40 mL.
Artículo V
Sheila Romo Sánchez, 2013 109
2.3.1. Adsorption
- Optimum pH: for this assay a
fixed concentration of enzyme (170
µg/mL) was immobilized at
different pH values (from 2.5 to
6.5) (Gómez et al., 2006b) by using
50 mM citric acid/Na2HPO4 (pH
2.5); 50 mM sodium acetate/acetic
acid (pH 3.0-6.0); and 50 mM
Na2HPO4/ NaH2PO4 (pH 6.5). 10
immobilization reactions per
support were so obtained.
- Optimum enzyme
concentration: different
concentrations of enzyme were
used (8.5 µg/mL, 17 µg/mL, 42.5
µg/mL, 84 µg/mL, 127.5 µg/mL,
170 µg/mL, 340 µg/mL, and 680
µg/mL) at optimum pH values. 8
immobilization reactions per
support were obtained.
Both assays were maintained at 10
ºC for 16 h.
- Optimum linked time: the
reactions were maintained for
20, 40, 60, 90, 120, 150, 180,
210 and 240 min at 10 ºC at the
optimum pH values and enzyme
concentration used for the
previous assays.
In any case, the suspension
was collected following
immobilization by centrifugation,
and was repeatedly washed with
50 mM corresponding buffer.
2.3.2. Adsorption-crosslinking
(reticulation)
This type of immobilization
was performed under the same
optimum conditions established for
adsorption. To this aim,
glutaraldehyde was added to the
chitosan-enzyme system at 5
different concentrations (from
0.125% to 1.5%) for 30 minutes
(Bernath and Venkatasubramanian,
1986).
2.3.3. Crosslinking-adsorption
Once the matrix was prepared
(see synthesis of supports section),
enzyme immobilization was
achieved based on the conditions
previously described for the
adsorption process.
Artículo V
110 Sheila Romo Sánchez, 2013
After immobilization was
completed for each assay, the
suspension was collected by
centrifugation and repeatedly
washed with the corresponding
buffer, so excess of glutaraldehyde
(crosslinking method) or protein (in
all methods) was removed.
Enzyme activity in each
preparation as well as the amount
of protein immobilized were
quantified as discussed below.
2.4. Determination of β-glucosidase
activity and adsorbed protein.
β-glucosidase activity of native
and immobilized enzymes was
determined by hydrolysis of
cellobiose and spectrophotometric
quantification of released glucose
by using an enzymatic test (Glucose
Go, Arévalo-Villena et al., 2006a).
The reaction mixture (100 μL β-
glucosidase solution and 400 μL of
1% (w/v) D-(+)-cellobiose in 50 mM
citrate-phosphate buffer (pH 5.5))
was incubated for 30 min at 37 ºC.
Each assay was conducted by
triplicate to determine the units of
enzyme (U). One U is defined as
amount of enzyme required to
release one mol of glucose per
minute under the above
mentioned assay conditions.
The amount of protein
adsorbed was quantified by
difference between the adsorbed
and free protein after
immobilization. Protein
concentration was calculated with
the Bradford method (Bradford,
1976) using BSA (bovine serum
albumin) as a standard.
2.5. Biotechnological
characterization of free and
immobilized β-glucosidase
Once the optimum
immobilization conditions were
established, the influence of
different variables such as pH,
temperature, and storage stability
on the activity was determined.
Kinetic constants and reuse
number were measured as well.
For that, the optimum enzyme
Artículo V
Sheila Romo Sánchez, 2013 111
concentration for immobilization
was used. In contrast, for free
enzyme, the dose recommended
by the manufacturer was
employed, except for the assay
determining stability over time,
which was carried out with the
same amount as that used for
immobilization.
pH stability: native and
immobilized β-glucosidase were
incubated with cellobiose at 37 ºC
for 30 min at different pH values
(2.0 to 8.0 at intervals of 0.5) using
different buffers (100 mM citric
acid/Na2HPO4, pH 2.0-2.5; 100 mM
sodium acetate/acetic acid, pH 3.0-
6.0; and 100 mM Na2HPO4/
NaH2PO4, pH 6.5-8.0).
Temperature stability: the
thermal stability of the free and
immobilized enzyme was tested by
incubation for 10 min at 45 – 90 ºC
at 5º intervals. The aliquots were
chilled quickly and assayed for
hydrolytic activity.
Storage stability: The stability
of enzymes over time was
measured by maintaining both
(free and immobilized) enzymes at
4 ºC and 28 ºC for 3 months at
optimal pH. The β-glucosidase
activity was assayed at regular
intervals (once week).
Kinetic properties: Vmax
( molmin-1mg-1) and kinetic
constant, Km (mM), were
determined from Michaelis-
Menten plots of specific activities
at 0.025% to 1% concentrations of
substrate, and the rates were
measured, ranging from 0.2 to five
times the value of Km. The values
of Vmax and Km were determined
by nonlinear regression (Arévalo-
Villena et al., 2006c).
Reusability of immobilized β-
glucosidase: Immobilized β-
glucosidase activity was assessed
by hydrolysis of cellobiose in
consecutive cycles, reusing the
enzyme in the same conditions of
all previous assays. After each
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112 Sheila Romo Sánchez, 2013
reaction, the enzyme was washed
with acetate buffer (pH 3.5; 50
mM) to remove any residual
substrate.
2.6. Enzymatic hydrolysis of aroma
precursors in grape must.
The glycosylated compounds
of a Muscat must were extracted
by retention and subsequent
elution in C18 columns following
the method proposed by Arévalo-
Villena et al. (2006a). 400 µL of the
resulting extract was mixed with
100 µL of free and immobilized
enzyme and maintained for 12 days
at 37 ºC. Every day, the glucose
liberated was checked. The amount
of free precursors was compared
with the total amount of
precursors obtained after acid
hydrolysis.
2.7. Application of immobilized and
free β-glucosidase in white
wine.
2.7.1. Microvinification.
Must from Muscat variety
harvested from Castilla-La Mancha
cultivars with pH of 3.5 was
microvinificated by duplicated in
dark bottles with 50 ppm of SO2,
and inoculated with 106 cells/mL of
BCS 103, a commercial
Saccharomyces bayanus strain.
Fermentation was carried out at 18
ºC, monitored for weight lost until
sugars were consumed. Glucose
and fructose contents were
measured by HPLC. Wines were
subsequently decanted and
fractionated for the next assay.
2.7.2. Aroma precursors
hydrolyzed by free and
immobilized β-glucosidase.
Wine fractions of 30 mL were
added with the optimum
immobilization concentration and
the concentration recommended
by the manufacturer for
immobilized and free enzyme,
respectively. All flasks were
maintained at 20 ºC for 16 days
due to it is the temperature used in
white vinifications. Enzymes were
removed by adding bentonite (20
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Sheila Romo Sánchez, 2013 113
g/hL). Samples were centrifugated
and kept at 4 ºC until analyzing the
volatile compounds. Each
enzymatic treatment was
performed in triplicate.
2.8. Quantification of Volatile
Compounds by Gas
Chromatography.
2.8.1. Solid phase extraction.
Volatile compounds were
extracted using the method
developed by Ibarz, et al. (2006).
Twenty five millilitres of wine were
passed through a preconditioned
polypropylene-divinylbenzene
cartridge (0.2 g of Lichrolut EN (40-
120 μM), Merck) using 4-nonanol
as internal standard. The column
was rinsed with 25 mL of water to
eliminate sugars, acids, and other
polar compounds. The free fraction
was eluted with 15 mL of pentane-
dichloromethane (2:1 v/v). Extracts
were concentrated to 100 μL by
distillation in a Vigreux column at
40ºC under nitrogen stream, and
then, kept at –20ºC until analysis.
2.8.2. GC-MS analysis.
Samples were analysed by
GC/MS using a ThermoQuest mod.
TraceGC gas chromatograph and a
DSQII mass detector with a
quadrupole analyser. All masses
were obtained in electronic impact
mode at 70 eV. 2 μL of the extract
were injected in a BP-21 (SGE)
column [FFAP phase (a
nitroterephthalic acid (TPA)
modified polyethylene glycol), 60
m x 0.32 mm and 0.25 μm film
thickness]. The chromatographic
conditions were the following:
oven temperature 43ºC (15 min) -
2ºC/min - 125ºC – 1ºC/min – 150ºC
- 4ºC/min - 200ºC (45 min) and
carrier gas helium (1.4 mL/min,
split 1/15, splitless time 0.5 min).
Separated compounds were
identified by comparing their mass
spectra and their chromatographic
retention indices, using commercial
products as a standard.
Quantification was performed by
analyzing the characteristic m/z
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114 Sheila Romo Sánchez, 2013
fragment for each compound using
the internal standard method.
Results for non-available products
were shown as the relationship
between the area of each
compound and that of the internal
standard.
2.9. Statistical analysis.
The statistical significance of
the effect of free and immobilized
β -glucosidase of each assay
obtained in triplicate analysis was
determined by one-way analysis of
variance (ANOVA, version 12.9).
Significant differences were also
analyzed between the different
results for one same variable (pH,
temp, etc.) in each type of enzyme
(immobilized and free).
3. Results
3.1. Effects of supports and
methods of immobilizatio.
Two carriers (alginate-chitin
and chitosan-chitin) and three
different immobilization methods
were used (adsorption,
reticulation, and crosslinking-
adsorption). The results obtained
showed that the alginate-chitin
support failed to retain the enzyme
at all pH values set (data not
shown). Chitosan was the most
appropriate polysaccharide for
immobilization both by adsorption
and by reticulation. Crosslinking
adsorption did not result an
adequate method (data no shown).
Martino et al. (1996b) in a similar
study, observed a considerable
affinity on chitosan too, obtaining
immobilization yields of 55-85%)
3.2. Optimization of immobilization
methods
The glucosidase activity of the
immobilized enzymes on chitosan-
chitin was measured. All results
were expressed as relative activity
in the form of (%) activity = activity
of enzyme / maximum activity of
enzyme) * 100%. An exception was
the assay determining stability over
time, in which enzyme units (U)
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Sheila Romo Sánchez, 2013 115
were adopted because the dose
used in this method was the same
for all cases.
The results of optimization of
the different variables are shown in
Fig. 1. Optimum immobilization pH
value (1a), immobilization time
(1b), and concentration of β-
glucosidase (1c) were 3.5, 120 min,
and 170 μg/mL respectively for
adsorption immobilization
process.
Under these conditions,
immobilization by crosslinking was
carried out with an optimum
glutaraldehyde concentration of
0.25% (Fig. 1d).
Fig. 1. Effect of optimum conditions on the activity of immobilized β-glucosidase. a: immobilization pH; b: immobilization time (min); c: enzyme concentration (μg/mL); d: glutaraldehyde concentration (%)
a b
c d
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116 Sheila Romo Sánchez, 2013
3.3. Biotechnological
characterization of free and
immobilized β-glucosidase
For the assay determining
stability over time, free enzyme
was carried out with the same
amount as that used for
immobilization (170 ug/mL in the
final reaction). In contrast, for the
rest of assays the dose
recommended by the
manufacturer for free enzyme was
employed (0.1 g/mL).
The biotechnological
characterization of enzymes was
carry out by studying the following
parameters:
pH stability: the influence of
pH values on free and immobilized
β-glucosidase was studied in a
range of 2.0-8.0 (Fig. 2a). The three
enzymes were stable in the same
pH range, from 4.0 to 6.0, whereas
it decreased sharply when the pH
value was out of this range.
Temperature stability: the
rates of thermal stability of the
free and immobilized β-glucosidase
were studied at the temperature
range of 45 to 90ºC (Fig. 2b). All
cases showed an increased activity
as temperature rose, reaching a
maximum at 70ºC for the enzyme
immobilized by adsorption and at
75ºC for the cross-linked and free
enzymes. The activity rate dropped
from 80ºC, which affected the
enzyme immobilized by reticulation
the most. Accordingly, this enzyme
only retained 45.52% of its relative
activity at 90 ºC.
a
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Sheila Romo Sánchez, 2013 117
Fig. 2. Effect of pH (a) and temperature (b) values on the activity of free and immobilized β-glucosidase. F: free enzyme; A: immobilized enzyme by adsorption; R: immobilized enzyme by reticulation.
Storage stability: Fig. 3 shows
the storage stability of free and
immobilized β-glucosidase at 28 ºC
and 4 ºC for 3 months. No decrease
in the activity of the immobilized
enzyme was observed. In contrast,
an increase was observed after 15
and 30 days. As regards free β-
glucosidase, its activity rate was
reported as virtually zero when
used in the same dose as that of
the immobilized enzyme. In similar
studies, the immobilized enzyme
showed lower activity than the free
one at 24ºC (Martino et al., 1996a).
Fig. 3. Storage stability of free and immobilized β-glucosidase at 4 ºC and 28 ºC. F28: free enzyme at 28 ºC; A28: immobilized enzyme by adsorption at 28 ºC; R28: immobilized enzyme by reticulation at 28 ºC; F4: free enzyme at 4ºC; A4: immobilized enzyme by adsorption at 4 ºC; R4: immobilized enzyme by reticulation at 4 ºC.
Kinetic properties: Table 1
shows the Km and Vmax values of
free and immobilized β-
glucosidase. The kinetic behaviour
of β-glucosidase was changed via
immobilization. The Km value was
lower for the free enzyme than for
the immobilized enzyme. In
contrast, the Vmax value was higher
for native β-glucosidase.
b
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118 Sheila Romo Sánchez, 2013
Table 1. Apparent kinetic constants of native and immobilized by adsorption and reticulation β-glucosidase
Vmáxa Km
b
Free 3.6 0.18
Immobilized by adsorption
3.36 0.2
Immobilized by reticulation
3.48 0.21
a: In molmin
-1mg
-1
b: In mM
Reusability of immobilized β-
glucosidase: As shown in Fig. 4, the
immobilized β-glucosidases were
used until the activity fell. Both
enzymes maintained 21% of their
activity after 7 reuses. However,
the enzyme immobilized by
crosslinking showed higher values
of activity during the 7 first assays.
Fig. 4. Reusability of immobilized β-glucosidase by adsorption and crosslinking. A: enzyme immobilized by adsorption; R: enzyme immobilized by reticulation.
3.4. Enzymatic hydrolysis of aroma
precursors in grape must
The aroma precursors isolated
from the grape must were brought
into contact with the enzymes
under examination. The results of
the hydrolysis of bonded
compounds by free and
immobilized β-glucosidase are
shown in Fig. 5, where 100%
corresponds to total hydrolysis of
precursors (acid hydrolysis). It is
shown that the enzyme
immobilized by adsorption
hydrolyzed around 5% of
precursors after 48 hours of
treatment only, whereas the free
enzyme started activity from the
fifth day. The maximum hydrolysis
rates obtained were 34%, 29%, and
25% for the free enzyme, the
enzyme immobilized by adsorption,
and the enzyme immobilized by
Artículo V
Sheila Romo Sánchez, 2013 119
reticulation, respectively. It should
be noted that the latter enzyme
reached its maximum activity rate
after one week in contact with the
substrate.
Fig. 5. Percentage of aroma precursors hydrolyzed by enzymatic treatment of a glycosidic extract from Muscat must. F: free enzyme; A: enzyme immobilized by adsorption; R: enzyme immobilized by reticulation. 170 µg/mL is the concentration final of all enzymes.
3.5. Application of immobilized and
free β-glucosidase to white
wine
Table 2 shows the contents of
volatile terpenes, alcohols,
lactones, norisoprenoids, and other
compounds of interest in wine
aroma that are prone to be
released from glycosides following
treatment with immobilized
enzymes at the optimum
immobilization concentration and
with native enzymes at the
concentration recommended by
the manufacturer.
After the enzymatic
treatment, the group of alcohols
showed a very significant increase
in benzyl alcohol, an increase
which was more significant in the
immobilized enzyme than in the
free enzyme. By contrast, the free
enzyme treatment released higher
amounts of 2-phenylethanol and 3-
methyl-2-buten-1-ol.
With respect to C6 alcohols,
the outcomes showed that the
immobilized enzyme by
crosslinking caused a slight
increase in 1-hexanol, (E), and (Z)-
3-hexenol.
Treatment with immobilized
enzyme generally led to a greater
release of terpenic compounds,
except for the isomers of pyranoid
oxides of linalool and of 6-methyl-
5-hepten-2-ol, whose glycosides
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120 Sheila Romo Sánchez, 2013
were more intensively hydrolyzed
by the free enzyme.
Wine treatment with enzyme
immobilized by reticulation was
more effective in the release of the
norisoprenoids 3-hydroxy-β-
damascone and 3-oxo-α-ionol.
Damascenone, the third
norisoprenoid analyzed, did not
undergo any changes in
concentration after the enzymatic
treatments, which may be due to
the fact that this compound is not
present in the form of glycoside.
Regarding to the group of
phenols involved a significant
increase in concentration of these
compounds in the majority of cases
was observed. This aroma
enhancement was generally more
significant with immobilized
enzymes, except for the
compounds 4-methyl-2,6-di-tert-
butylphenol, methyl vanillate, and
acetovanillone.
Finally, wines treated with
free enzymes showed higher
concentrations of 4-
ethoxycarbonil-γ-butyrolactone,
whereas wines treated with
enzymes immobilized by
reticulation showed higher
concentrations of γ-butyrolactone,
furfural, and N(2-phenylethyl)-
acetamide.
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Sheila Romo Sánchez, 2013 121
Table 2. Volatile compounds in the Muscat wine (µg/L) released following treatment with immobilized and free enzymes (170 µg/mL and 0.1 g/mL respectively)
Volatile compounds in wine
Immobilized by adsorption
Immobilized by reticulation
Free enzyme Control
Alcohols
1-Pentanol 46.3 ± 1.6 40.6 ± 7.6 49.6 ± 26.2 34.3 ± 5.0
3-Methyl-2-buten-1ol 13.0 ± 1.4
b 13.0 ± 0.8
b 21.29 ± 5.92
c 2.4 ± 0.3
a
c-2-Penten-1-ol 17.3 ± 2.2 17.6 ± 3.9 15.45 ± 4.62 14.9 ± 1.3
Benzyl alcohol 288.9 ± 11.8 b
304.1 ± 8.4 b
183.4 ± 92.9 b
20.7 ± 1.1 a
2-Phenyletanol 18318.9 ± 293.9 a
21407.6 ± 3357.3 a
29639.4 ± 2441.4 b
22035.3 ± 791.2 a
Alcohols C6
1-Hexanol 688.7 ± 31.2 b. c. d
771.2 ± 86.4 d
513.7 ± 128.2 a. c
533.4 ± 39.6 a. b
(E)-3-Hexenol 15.9 ± 2.3 a
20.2 ± 0.6 b
14.9 ± 1.4 a
15.2 ± 0.8 a
(Z)-3-Hexen-1-ol 15.4 ± 1.7 c
16.3 ± 1.7 c
10.8 ± 0.4 b
5.4 ± 0.5
a
(E)-2-Hexenol 12.9 ± 1.1 a
18.2 ± 1.7 a
22.7 ± 1.0 a.b
37.0 ± 19.1
b
Terpenes
Linalool 358.2 ± 12.5 b
348.5 ± 8.5 b
272.3 ± 40.9
a 239.8
± 11.9
a
α-Terpineol 106.0 ± 3.1 c 101.6 ± 5.5
b. c 89.2 ± 7.7
b 69.9
± 0.7
a
Citronellol 20.2 ± 5.7 16.5 ± 4.0 25.2 ± 10.8 14.0 ± 1.8
Nerol 140 ± 1.3 c 129.4 ± 11.6
c 71.9 ± 7.2
b 9.0
± 0.5
a
Geraniol 110.2 ± 8.1 b
92.4 ± 18.7 b
97.7 ± 31.2 b
29.8 ± 1.0
a
t-Pyran-linalool oxide 155.0 ± 1.9 146.5 ± 22.0 162.6 ± 46.1 101.2 ± 3.2
c-Pyran-linalool oxide 34.5 ± 0.6
a 34.0 ± 7.9
a.b 51.1
± 5.2
c 37.0
± 1.9
b
c-Rose oxide 3.6 ± 0.2 b
3.5 ± 0.1 b
2.0 ± 1.2 a
0.5 ± 0.06 a
t-Rose oxide 1.0 ± 0.1 b
1.2 ± 0.1 b
0.6 ± 0.3 a
0.1 ± 0.00 c
3,7-Dimethyl-1,5,7-octatrien-3-ol 43.7 ± 0.9
c 40.8 ± 4.6
c 9.8
± 7.3
a 29.8
± 0.2
b
3,7-Dimethyl-1,5-octadien-3,7-diol 650.1 ± 137.0 551.1 ± 348.8 760.8 ± 224.4 637 ± 2.8
3,7-dimethyl-1,7-octadien-3,6-diol 99.2 ± 23.4
c.b.d 125.6 ± 42.4
d 49.4 ± 22.7
a. b 49.8 ± 0.5
a. c
2,7-Dimethyl-4,5-octanodiol 9.1 ± 0.4
a 6.6 ± 0.5
a 18.6
± 3.7
b 16.5
± 0.8
b
6-Methyl-5-hepten-2-ol 9.7 ± 1.6
a 11.7 ± 0.4
a 32.6 ± 15.9
b 2.4 ± 0.41
a
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122 Sheila Romo Sánchez, 2013
Volatile compounds in wine
Immobilized by adsorption
Immobilized by reticulation
Free enzyme Control
Norisoprenoids
Damascenone 2.6 ± 0.4 2.2 ± 0.6 1.4 ± 1.1 1.6 ± 0.1
3-Hydroxy-β-Damascone 3.5
± 0.7
a 12.1 ± 4.3
b 2.5 ± 0.9
a 1.4
± 1.9
a
3-Oxo-α-ionol 14 ± 6.3 b
37.8 ± 6.6 c 9.5 ± 2.8
a. b 1.8 ± 0.1
a
Phenols
4-Methyl-2,6-di-tert-butyl-phenol 49.4 ± 23.5
a 63.8 ± 11.4
a 281.6
± 39.2
b 112.7 ± 27.2
a
Phenol 5.5 ± 1.1 b
4.4 ± 0.2 a. b
4.1 ± 0.8 a. b
3.2 ± 0.1
a
Eugenol 2.1 ± 0.4 b
2.2 ± 0.2 b
2.0 ± 1.1
b 0.4
± 0.1
a
Siringol 5.5 ± 0.9 c
2.3 ± 0.2
a 4.2
± 0.1
b 1.9
± 0.0
a
Vanillin 14.3 ± 1.8 16.0 ± 4.2 12.8 ± 6.7 4.7 ± 0.9
Methyl vanillate 2.7 ± 0.5
a 4.2 ± 0.3
a.b 5.3
± 1.5
b 2.4
± 0.2
a
Ethyl vanillate 2.6 ± 0.3 b
1.9 ± 0.7
b 0.6
± 0.3
a 0.3
± 0.0
a
Acetovainillone 6.3 ± 0.4 a
12.9 ± 2.0
b 13.5 ± 1.4
b 4.8
± 0.4
a
Homovanillyl_alcohol 22.1 ± 3.8
a 76.2
± 29.4
b 12.2 ± 3.4
a 7.9 ± 1.2
a
Lactones and others
γ-butirolactone 106.4 ± 40.1 a.b
151.6 ± 5.5 a.b
91.8 ± 13.8 b
82.7 ± 5.3
a
4-Ethoxycarbonyl-γ-butyrolactone 359.2 ± 56.9
a 230.3 ± 53.1
a 513.5
± 145.6
b 303.3 ± 8.3
a
4(1-hydroxy-ethyl)-γ-butirolactone 44.8 ± 4.0 44.8 ± 31.9 36.8 ± 2.1 35.7 ± 1.6
Furfural 9.3 ± 0.4
b 9.7
± 1.3
b 3.2
± 2.7
a 1.5
± 0.4
a
N-(2-Phenylethyl)-acetamide 22.1
± 3.4
a 80.3
± 11.5
b 11.4
± 2.9
a 18.5
± 0.8
a
Different letters indicate significant differences (95% confidence) between free and immobilized enzymes
Artículo V
Sheila Romo Sánchez, 2013 123
4. Discussion
The aim of this study was to
improve the stability of the enzyme
β-glucosidase by immobilization.
This enzyme is currently used in
enology to release volatile
compounds and its immobilization
could be interesting due to the
possibility of reusability and
because could be required in lower
doses than the free enzymes
thanks to the enhancement of
enzymatic stability.
The most appropriate matrix
for both immobilization types was
chitosan, which ensures economic
benefits because chitosan is a
polyamino saccharide obtained at a
relatively low cost. For this reason,
it has been increasingly used in
food, pharmaceutical, medical, and
agricultural applications
(Krajewska, 2004).
Concerning optimization of
the immobilization process, the
optimum crosslinking pH value was
3.5. This value results from the
activity of the polyionic matrixes,
which causes the partitioning of
protons between the bulk phase
and the enzyme
microenvironment, and eventually,
a change in the optimum pH value.
This change depends on the
immobilization method as well as
on the structure and charge of the
matrix (Arica et al., 1999; Busto et
al., 1997).
Knowing the biotechnological
characteristics of both free and
immobilized enzymes is crucial to
assess the efficacy of the
immobilization method. The results
showed the same trend for all
enzymes inasmuch as they were
stable in a range of 4.0 to 6.0. In
spite of this, the statistical analysis
showed significant differences
between the enzymes at different
pH values. At pH 4.0, the enzyme
immobilized by reticulation
showed a lower activity than that
of the other two enzyme types. In
contrast, at pH 5.5 it was the
Artículo V
124 Sheila Romo Sánchez, 2013
enzyme immobilized by adsorption
that hydrolyzed to a lesser extent.
Similar findings were reported in
other studies (Fan et al., 2011;
Gómez et al., 2008). These
perturbations on pH-profiles could
be attributed to pH changes in the
region of the immobilized enzyme
particles, resulting from polymer
characteristics and product
accumulation (Arica et al., 2000).
However, these differences are not
economically significant, especially
because the free enzyme dose
used—which is the one
recommended by the
manufacturer—is 367 times higher
than the doses used for
immobilization throughout the
study.
All enzymes showed a stable
behaviour at different
temperatures in the pH range
studied. Romo et al. (2012)
observed a similar behaviour in a
commercial cellulase. The
enzymatic activity clearly increased
from 60ºC, and began to decrease
from 80ºC. A number of studies
show that the optimum
temperature of an immobilized β-
glucosidase could be higher, lower
or the same as the free β-
glucosidase and its stability vary
considerably depending on the
strain producer (Busto et al., 1997).
This fact supports the variance
observed in our study. The ANOVA
results for a factor showed values
significantly lower for the
immobilized enzymes in the
temperature range of 45 to 60ºC.
The enzyme immobilized by
reticulation was the least active,
which stands to reason, since the
enzyme is more protected, and
thus, its active centre is less
accessible to the substrate.
Stability during storage is one
of the most important
characteristics of enzymes. As the
results show, immobilized enzymes
were stable at both storage
temperatures (4ºC and 28ºC).
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Sheila Romo Sánchez, 2013 125
Chang and Juang (2007) and Fan et
al. (2011) observed a similar
behavior in a dried-composite
immobilized β-glucosidase stored
at 4 ºC and in an enzyme
immobilized after storage for 20
days, respectively. The increase in
activity after 15 days might be a
mechanism to prevent enzymatic
denaturalization after
immobilization (Erzeng et al.,
2010). The fact that the free
enzyme shows virtually no activity
in the same dose as the
immobilized enzyme is one of the
most relevant findings of the
present study because it is
indicative of the potential
economic savings that technique
would guarantee.
The kinetic constants
observed in our study were similar
to those reported by Busto et al.
(1995). Generally, immobilization
increases hindrance between the
enzyme and its substrate, and thus,
increases the Km value (Su et al.,
2010). Few immobilization studies
have reported a decrease in Km
values for the immobilized
enzymes (Fan et al., 2011).
The immobilization process
also facilitates the efficient
recovery and reuses of enzymes,
which is the sine qua non of
economic viability in many
applications, and enables their use
in continuous process (Sheldon,
2007), which ensures substantial
economic benefits. Jung et al.
(2012) observed the immobilized β-
glucosidases on silica gel
maintained 80% of its relative
activity after 20 reuses. Su et al.
(2010) used an immobilized β-
glucosidase on alginate for 50
times and the residual activity was
about 93.6% of its initial activity. In
our study, both immobilized
enzymes maintained 21% of their
initial activity after 7 reuses.
Enzymatic hydrolysis is
required to enrich wine flavour by
releasing free aromatic compounds
Artículo V
126 Sheila Romo Sánchez, 2013
from natural glycoside precursors.
As a preliminary step to release
volatile compounds in wine,
hydrolysis of the different enzymes
was performed with actual
substrates (glycosylated
precursors)—not with cellobiose,
as had always been done. The
glycosidic extracts obtained from
the Muscat must were treated with
free enzymes and enzymes
immobilized by adsorption and
reticulation. The immobilized
enzymes needed less time to start
activity than free β-glucosidase,
which would provide a
biotechnological and economic
advantage to winemaking. Arévalo-
Villena et al. (2006b) reported
similar findings.
Concerning direct volatile
compound release on the wine, it
was observed that the enzymatic
treatments generally caused an
expected increase in compounds
from glycosidic precursors. A
significant increase in benzyl
alcohol contributed nice blackberry
(Latrasse, 1991), floral, and sweet
notes to the wine aroma. The use
of immobilized enzymes would be
particularly beneficial because
these enzymes released a higher
amount of volatile compounds.
The aldehydes and C6 alcohols
provide wine with green, greasy
aromas (López-Tamames et al.,
1997), which contribute aromatic
complexity if added in moderate
concentrations, as in our study.
The family of terpenic compounds
is the basic aromatic constituent of
varieties such as Muscat, among
others. These compounds add
floral, sweet, and citric aromas
(Guth, 1997). The amount of most
of the terpenic compounds
analyzed in this study increased
significantly following the
enzymatic treatment. Particularly
remarkable was the release of
large amounts of nerol, and to a
lesser extent, of geraniol, amounts
Artículo V
Sheila Romo Sánchez, 2013 127
which were larger when it came to
immobilized enzymes.
Finally, null increase of
damascenone correlates with
findings reported by other authors
(Sánchez-Palomo et al., 2007) and
with data yet to be published by
our research group. These data
show that β-ionone, another
compound of the damascenone
group—but not found in the wines
analyzed in this study—, is only
present in free form in grapes.
5. Conclusions
This study shows that the
alginate-chitin support is not
appropriate for immobilization of
the commercial β-glucosidase
analyzed. The matrix chitosan-
chitin was shown to be the optimal
support for immobilization both by
adsorption and reticulation.
Optimal immobilization
variables were consistent with
winemaking in all cases, which
makes the process suitable for
enology. The doses and time
required for hydrolysis for
immobilized enzymes were lower
and shorter, respectively than
those needed by the free one.
Moreover, enzyme reusability
guarantees great economic
benefits.
On the other hand, both types
of immobilized enzyme acted with
the same or even more intensity
than the free enzyme in the release
of some compounds desirable for
wine aroma.
In conclusion, the present
study shows that the use of
immobilized enzymes in enology is
a valid alternative for winemaking,
and could provide substantial
advantages to the sector.
Aknowledgements
The authors wish to express
their gratitude to Junta de
Comunicades de Castilla La Mancha
for funding this research, which
was performed in the framework
Artículo V
128 Sheila Romo Sánchez, 2013
of project “Inmovilización de
enzimas para su aplicaciónen la
industria agroalimentaria” (Ref:
2010-COB-3763). And the
International Foundation for
Science, Stockholm, Sweden and
the Organization for the
Prohibition of Chemical Weapons,
The Hague, The Netherlands,
through a Grant to Héctor L.
Ramirez (Grant F/3004-67).
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ARTÍCULO I. Biodiversidad de levaduras procedentes de ecosistemas
oleicos: Estudio de sus propiedades biotecnológicas
El objetivo del trabajo fue la identificación de levaduras en frutos, pastas y
orujos de dos variedades de aceituna (Arbequina y Cornicabra) procedentes de
Castilla-La Mancha; así como el estudio de sus propiedades biotecnológicas
con interés industrial.
Durante la campaña 2007-2008, se recogieron muestras de distintos
olivares y almazaras. Tras realizar el aislamiento de levaduras mediante
siembra en agar YPD, se escogieron al azar un total de 108 colonias para su
posterior identificación a nivel de especie mediante Reacción en Cadena de la
Polimerasa y posterior Análisis de Restricción (PCR – RFLP). Para asignar la
especie, los fragmentos de restricción obtenidos se compararon con los
publicados en las base de datos (Yeast-id (www.yeast-id.org) y Quidy ‘03).
Algunos individuos no se pudieron identificar por este método, por lo que
hubo que recurrir a la secuenciación de las regiones comprendidas entre los
genes 18S y 28S del ADNr y en ocasiones, a la 26S (datos no publicados). El
alineamiento de secuencias proporcionado por el programa informático BLAST
(http://blast.ncbi.nlm.nih.gov/Blast.cgi) y la comparación de los resultados en
bases de datos públicas permitió identificarlos.
Se obtuvieron 14 especies pertenecientes a 7 géneros
(Zygosaccharomyces, Pichia, Lachancea, Kluyveromyces, Saccharomyces,
Candida, Torulaspora), de ellas 5 se pudieron identificar por PCR-RFLP y el
resto mediante secuenciación. Un total de 20 individuos se depositaron en la
base de datos de GenBank (www.ncbi.nlm.nih.gov/genbank/), con los números
de acceso GQ340429-48 (identidad ≥ 88%).
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La distribución de la microbiota por variedades de aceituna puso de
manifiesto que en Cornicabra la especie mayoritaria fue Pichia holstii (39%)
seguida por Zygosaccharomyes fermentati (25%), mientras que en Arbequina
fueron Pichia caribbica (59%) y Z. fermentati (23%). La biodiversidad resultó
mayor en Cornicabra que en Arbequina (11 especies vs. 6). Por otra parte, se
encontró que sólo Z. fermentati, P. caribbica y Lachancea. sp se aislaron en los
tres sustratos estudiados.
Tras la identificación se procedió a la caracterización biotecnológica de las
cepas más representativas. Para ello, el 30% de los individuos de cada especie
(un total de 42 aislados), se sometieron a las siguientes pruebas enzimáticas:
celulasa sobre carboximetilcelulosa (CMC), poligalacturonasa (PGA) mediante
ácido poligalacturónico, β-glucosidasa sobre celobiosa, peroxidada con H2O2,
lipasa mediante CaCl2/Tween 80 y glucanasa sobre β-glucano.
Todas las actividades se chequearon de forma cualitativa mediante
observación de un halo de hidrólisis (PGA), de un precipitado (lipasa), de
crecimiento (celulasa y β-glucosidasa) y de formación de burbujas
(peroxidada). La glucanásica se cuantificó estudiando los azúcares reductores
liberados (ácido dinitrosalicílico – DNSA).
Los resultados mostraron que ningún aislado presentaba actividad
lipásica, hecho positivo ya que su presencia en aceite afecta a su calidad,
debido al aumento de diacilgliceroles y ácidos grasos. Sólo cuatro hidrolizaron
la CMC (P. caribbica, P. holstii, P. mississippiensis y Z. florentinus) y otros
cuatro, (Pichia caribbica, Pichia mississippiensis, Candida diddensiae y Candida
thermophila), mostraron actividad PGA. Estas dos enzimas aumentan los
compuestos fenólicos de carácter antioxidante prolongando la vida útil del
producto, y además incrementan el rendimiento de la extracción del aceite de
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Sheila Romo Sánchez, 2013 139
oliva. Por el contrario, la actividad peroxidasa se detectó en 38 de las 42 cepas
estudiadas; y la β–glucosidasa, enzima capaz de hidrolizar la oleuropeína
(principal compuesto fenólico de las aceitunas) en 29 de las cepas. Casi la
mitad de los aislados estudiados excretaron la enzima β–glucanasa en mayor o
menor medida, lo cual es interesante para su posible aplicación en
alimentación animal o enología.
Este estudio reflejó la biodiversidad levaduriforme del ecosistema
olivarero en función de las variedades y del sustrato estudiado. Los
enzimogramas mostraron que las actividades enzimáticas son cepa
dependiente, por lo que habría que seleccionar individuos concretos para su
posible aplicación biotecnológica.
Aunque el uso de enzimas en este sector no está permitido, la presencia
de levaduras no fitopatógenas con alta o moderada actividad enzimática
podría mejorar la calidad y el rendimiento de la extracción de aceite.
ARTÍCULO II. Mohos aislados de ecosistemas oleicos y su uso en
biotecnología
Puesto que la microbiota de un ecosistema es compleja y no se limita a
individuos de un solo grupo, se continuó identificando y caracterizando a los
mohos presentes en los sustratos del trabajo anterior.
Se obtuvieron un total de 53 cultivos puros de mohos que se identificaron
morfológicamente (color, textura del micelio, formación de esporas) y por
secuenciación de la región ITS1-5.8S-ITS2 del ADNr.
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Los aislados se agruparon en 14 especies pertenecientes a 7 géneros
(Aspergillus, Penicillium, Rhizomucor, Mucor, Rhizopus, Lichtheimia y
Galactomyces). De todas las secuencias sólo 41 se publicaron en la base de
datos de GenBank (www.ncbi.nlm.nih.gov/genbank/) con números de acceso
FJ499432-FJ499462 y FJ227888-FJ227897 (identidad ≥ 97%).
La especie predominante fue A. fumigatus con un 30% de frecuencia
seguida por Penicillium commune, Galactomyces geotrichum y Rhizomucor
variabilis var. regulatior (11%, 10%, 10% respectivamente). Algunos individuos
sólo se pudieron identificar a nivel de género (Aspergillus sp., Penicillium sp. y
Galactomyces sp.).
Comparando la biodiversidad entre cada uno de los sustratos, las pastas
de las aceitunas ofrecieron la mayor variabilidad (10 especies diferentes),
seguidas a partes iguales por las aceitunas y los orujos, donde se identificaron
5 distintas.
En cuanto al estudio de las actividades enzimáticas, se chequearon
cualitativamente sobre placa la carboximetilcelulasa (CMCasa),
poligalacturonasa (PGA), β-glucosidasa y lipasa, utilizando para ello, los
mismos sustratos que en el estudio anterior (carboximetilcelulosa (CMC), ácido
poligalacturónico, celobiosa y CaCl2/Tween 80 respectivamente). La presencia
o no de actividad se detectó por crecimiento (CMCasa y β-glucosidasa), halos
de hidrólisis y de precipitación en PGA y lipasa respectivamente.
Los resultados ofrecieron una alta variabilidad. Casi todas las especies
tuvieron actividad β-glucosidásica con distinta intensidad, la CMCasa fue
excretada en 43 cepas, destacando Aspergillus niger, Aspergillus fumigatus,
Rhizomucor variabilis y Mucor fragilis por sus altos valores. PGA y lipasa se
liberaron en menos de la mitad de las cepas estudiadas.
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Se cuantificaron también algunas de las actividades enzimáticas más
interesantes para el sector agroalimentario (β-glucosidasa, pectinasa, celulasa
y xilanasa). Para ello, se utilizaron las cepas que mejores resultados
presentaron en los ensayos anteriores. Los mohos se crecieron en orujos de
aceituna y hollejos de uva (ricos en compuestos lignocelulósicos). La actividad
se evaluó en el sobrenadante a los 3, 5, 7, 8 y 10 días de fermentación
cuantificando la liberación de azúcares reductores (DNSA).
Una cepa de A. fumigatus 3 y otra de A. niger 113N, ambas depositadas
en la Colección Española de Cultivos Tipo (CECT) con los números de acceso
20827 y 20828 respectivamente, liberaron cantidades importantes de celulasas
cuando crecían sobre hollejos y de xilanasas sobre orujos, especialmente A.
niger. No se detectaron β-glucosidasa ni pectinasa.
Estas cepas podrían usarse en fermentación en fase sólida, tanto para la
producción de enzimas como para el pre-tratamiento de subproductos
agrícolas.
Por esta razón, se decidió avanzar en una nueva línea de investigación en
la que se plantearon nuevos retos que dieron continuidad al trabajo.
ARTÍCULO III. Producción e inmovilización de enzimas mediante
fermentación en fase sólida de residuos agroindustriales
Este estudio trató de aunar las buenas propiedades enzimáticas de las
cepas de Aspergillus fumigatus 3 y Aspergillus niger 113N y la necesidad de
ofrecer alternativas a los subproductos de sectores agroalimentarios.
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142 Sheila Romo Sánchez, 2013
Para ello, se produjo β-glucosidasa, celulasa, xilanasa, y pectinasa
mediante fermentación en fase sólida (FFS) sobre hollejos de uva y orujos de
aceituna, teniendo en cuenta factores como la composición del medio y el
grado de molienda. Las enzimas obtenidas se purificaron, liofilizaron e
inmovilizaron.
La composición química bruta de los subproductos, mostró que los
principales componentes en ambos eran ligninas y celulosas, y en menor
medida pectinas. En los orujos se hallaron también cantidades apreciables de
grasa.
Los sustratos se secaron y una fracción se molió utilizando un tamiz con
un tamaño de poro de 0.5 µm. Combinando las variables de suplementación
de nitrógeno (trigo) y tamaño de partícula, resultaron un total de 8
fermentaciones para cada uno de los mohos:
S: hollejos de uva no molidos
SW: hollejos de uva no molidos (2.5 g) + trigo (2.5 g)
mS: hollejos de uva molidos
mSW: hollejos de uva molidos (2.5 g) + trigo (2.5 g)
P: orujos de aceituna sin moler
PW: orujos de aceituna sin moler (2.5 g) + trigo (2.5g)
mP: orujos de aceituna molidos
mPW: orujos de aceituna molidos (2.5 g) + trigo (2.5g)
Las distintas fermentaciones se llevaron a cabo en cámara climática a
25ºC y 65% de humedad. Los sustratos se inocularon con 5 mL de una
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Sheila Romo Sánchez, 2013 143
suspensión de micelio procedente de un precultivo en agar patata (PDA) (30 ºC
/ 65 % humedad / 7 días).
A los 2, 4, 6, 10 y 15 días se obtuvo el extracto enzimático bruto de cada
cultivo. En él se cuantificaron las actividades enzimáticas xilanasa, pectinasa,
celulasa y β-glucosidasa sobre sus respectivos sustratos (artículos 1 y 2). En los
tres primeros casos se determinaron los azúcares reductores con DNSA y para
la β-glucosidasa se midió la glucosa libre mediante un kit enzimático específico
debido a interferencias en la cuantificación.
Los mejores resultados, tanto para el crecimiento como para la excreción
de enzimas, se obtuvieron a los 10 y 15 días de fermentación de los hollejos sin
moler. En los ensayos realizados con los orujos molidos, los mohos no se
desarrollaron, y en la mayoría de los casos, la suplementación con trigo resultó
conveniente. Bajo estas condiciones (15 días de fermentación con hollejos sin
moler y ausencia de trigo), A. niger (113N) presentó una actividad xilanásica de
47.05 U/g sustrato.
Su extracto enzimático se purificó parcialmente mediante exclusión
molecular (100 Kda) y se liofilizó con y sin crioprotector. El rendimiento de
cada proceso se calculó cuantificando la actividad xilanásica y se visualizó por
electroforesis de proteínas. Los resultados mostraron que la purificación
parcial retuvo en torno al 83 % de la actividad inicial; sin embargo, el proceso
de liofilización no resultó adecuado, perdiéndose casi el 90% de la capacidad
hidrolítica.
No obstante, con el fin de obtener un sistema multienzimático soportado
en una matriz inerte, dicho extracto se inmovilizó por adsorción sobre quitina-
quitosano. Aunque los resultados no fueron satisfactorios, este estudio
ofrecería la posibilidad de inmovilizar extractos brutos complejos.
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144 Sheila Romo Sánchez, 2013
ARTÍCULO IV. Inmovilización de celulasa y xylanasa comerciales sobre
varios soportes poliméricos (quitina-alginato y quitina-quitosano) y mediante
diferentes métodos
En los dos últimos artículos que integran esta Tesis Doctoral, y con la
colaboración del Centro de Tecnología Enzimática de la Universidad de
Matanzas, se trabajó en la inmovilización de enzimas.
En este trabajo, se inmovilizaron una celulasa y una xilanasa comerciales
sobre quitina-alginato y quitina-quitosano mediante tres métodos de unión
química: adsorción, reticulación y un método mixto, entrecruzamiento-
adsorción. En el último, antes de la inmovilización, el soporte se activó con
distintas concentraciones de glutaraldehído (0.5% y 15%).
Las variables que afectan a la unión se optimizaron utilizando una
concentración de 0.03 g/mL de cada soporte. Todas las reacciones se llevaban
a cabo a 10ºC, estudiando variables como el pH, la concentración óptima de
enzima, el tiempo de unión y la concentración óptima de glutaraldehído
(método de reticulación):
i. pH de unión enzima-soporte: manteniendo una concentración
constante de enzima (170 µg/mL) se modificaron los valores de pH de
cada reacción (de 2.5 a 5.5). Los resultados mostraron que el soporte
más adecuado para ambas enzimas era la quitina-quitosano. La unión
fue óptima a pH 5.0 para la xilanasa y 4.5 para la celulasa.
ii. Concentración óptima de enzima: se estudiaron siete reacciones de
inmovilización con diferente concentración de enzima (de 8.5 a 340
µg/mL), manteniendo un pH de 5.0 y 4.5 (xilanasa y celulasa
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Sheila Romo Sánchez, 2013 145
respectivamente). Los mejores valores de actividad fueron 127.5
µg/mL para la xilanasa y 170 µg/mL para la celulasa.
iii. Tiempo de unión enzima-soporte: se emplearon las condiciones
óptimas de pH y concentración de enzima, manteniendo las enzimas
con el soporte entre 20 y 240 minutos y cuantificando la actividad
cada 20. La unión fue máxima a los 120 minutos de contacto para la
xilanasa y 150 para la celulasa.
iv. Concentración óptima de glutaraldehído: una vez fijados el pH,
concentración de enzima y tiempo óptimo de inmovilización se añadió
glutaraldehído a 5 concentraciones diferentes resultando la óptima la
de 0.125% en ambos casos.
El método mixto (entrecruzamiento-adsorción) se descartó por ofrecer
pobres resultados, siendo la adsorción y la reticulación los seleccionados. Una
vez fijadas las mejores condiciones de inmovilización (celulasa: pH 4.5, 170
µg/mL de enzima, 2.5h de unión, 0.125% glutaraldehído; xilanasa: pH 5, 127.5
µg/mL , 120 minutos de unión, 0.125% glutaraldehído) se procedió a la
caracterización bioquímica de las enzimas libres e inmovilizadas por adsorción
y reticulación. Se estudió el pH óptimo de hidrólisis, estabilidad térmica,
constantes cinéticas, y número de reusos:
i. Estabilidad a cambios de pH: se llevó a cabo modificando el pH (de 2.0
a 8.0) de la CMC y el xilano. La xilanasa mostró buena estabilidad en un
amplio rango (de 3.0 a 8.0), tanto en su forma libre como inmovilizada.
Sin embargo, la celulasa únicamente fue estable a pHs ácidos,
alcanzando su máxima actividad a pH 3.0 y pH 4.0 (enzimas
inmovilizadas y nativa respectivamente).
Resúmen de articulos
146 Sheila Romo Sánchez, 2013
ii. Estabilidad térmica: tras mantener las enzimas durante 10 minutos a
diferentes temperaturas (de 40ºC a 90ºC) se observó que a 75ºC, la
actividad de la celulasa nativa cayó drásticamente, mientras que las
inmovilizadas mantuvieron alrededor de un 50% de su actividad inicial.
En el caso de la xilanasa, la inmovilización no supuso una ventaja tan
definida, ya que la enzima libre presentó buena actividad en todo el
rango de temperaturas.
iii. Constantes cinéticas: los valores de Km fueron menores en las enzimas
nativas que en las inmovilizadas, mientras que Vmáx fue similar en
ambos casos.
iv. Ciclos de reuso: la celulasa, se pudo utilizar 19 veces consecutivas,
presentado un actividad residual del 64% y 32% para la reticulada y la
adsorbida respectivamente en su último uso. En cuanto a la xilanasa
inmovilizada por ambos métodos, soportó 8 ciclos de reuso (25% la
reticulada y 14% la adsorbida).
De los resultados se dedujo que el proceso de inmovilización mejoró la
estabilidad de la celulasa y la xilanasa comerciales frente a cambios de pH y
temperatura, y que ambas presentaron buena capacidad de reutilización.
ARTÍCULO V. Inmovilización de una β-glucosidasa y su aplicación para
la liberación de precursores del aroma en un vino Moscatel
En este último artículo se llevó a cabo la inmovilización de una β-
glucosidasa enológica evaluándose su capacidad para hidrolizar los precursores
del aroma de mostos y vinos de la variedad Moscatel.
Resúmen de articulos
Sheila Romo Sánchez, 2013 147
Los soportes utilizados fueron quitina-alginato y quitina-quitosano y los
métodos empleados adsorción, reticulación y entrecruzamiento-adsorción (en
éste el soporte se activó con glutaraldehído al 0.5% y al 15%). La quitina-
quitosano fue la matriz más adecuada para las dos mejores técnicas de
inmovilización (adsorción y reticulación), descartándose el proceso de unión
mixta.
Realizando el mismo estudio que en el trabajo anterior, las condiciones
óptimas de unión enzima-soporte resultaron: pH 3.5; concentración de
enzima: 170 µg/mL; 120 minutos de tiempo de unión; y 0.25% la cantidad
óptima de glutaraldehído, en el caso de la reticulación. Bajo estas condiciones,
se realizó la caracterización bioquímica de las enzimas libre, inmovilizada por
adsorción e inmovilizada por reticulación, utilizando la dosis recomendada por
el fabricante en el caso de la libre excepto para el estudio de estabilidad
durante el almacenamiento, en el que se empleó la misma dosis para las tres
enzimas:
i. Estabilidad a cambios de pH: se modificó el pH de la celobiosa desde
4.0 hasta 8.0. Se observó que las tres enzimas eran estables en el
mismo rango (4.0-6.9) y su actividad fue nula fuera de él.
ii. Estabilidad térmica: las enzimas se mantuvieron durante 10 minutos a
distintas temperaturas (entre 45ºC y 90ºC). El máximo de actividad se
alcanzó a los 70ºC, en la inmovilizada por adsorción, y a los 75ºC en la
libre y la reticulada.
iii. Estabilidad durante el almacenamiento (a 4º y 28ºC): las enzimas se
mantuvieron durante tres meses a ambas temperaturas sin variación
de actividad.
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148 Sheila Romo Sánchez, 2013
iv. Constantes cinéticas: los valores Km para las enzimas inmovilizadas
fueron más altos que para la libre, comportamiento inverso al
observado para Vmáx
v. Ciclos de reúso: ambas enzimas inmovilizadas mantuvieron una
actividad del 21% aproximadamente tras 7 usos consecutivos.
Optimizados los parámetros de inmovilización y estudiadas las
características bioquímicas, se evaluó la hidrólisis de los precursores del aroma
de un mosto Moscatel, extraídos por retención y posterior elución con
columnas C18. 400 µL de precursores se mantuvieron durante 12 días con 100
µL de enzima (170 µg/mL) a 37ºC. Cada 24 h se cuantificó la glucosa liberada.
Los precursores del aroma se hidrolizaron en un 34%, 29% y 25% al
emplear la β-glucosidasa libre, inmovilizada por adsorción y por reticulación,
respectivamente.
Las enzimas inmovilizadas y libre se aplicaron a un vino Moscatel
elaborado en el laboratorio. Las microvinificaciones se llevaron a cabo a 18ºC
con una cepa comercial de Saccharomyces bayanus. Los vinos se decantaron y
fraccionaron en alícuotas de 30 mL, a las que se añadieron las enzimas a las
concentración óptima de inmovilización y en el caso de la enzima libre a la
concentración recomendada por el fabricante. Las muestras se mantuvieron a
20ºC durante 16 días y finalizado el tiempo estimado, se les adicionó bentonita
(20 g/hL). Como control se utilizó un vino sin adición de extracto.
El estudio de los compuestos volátiles liberados se determinó por
cromatografía de gases y espectrometría de masas.
Los vinos tratados con ambas β-glucosidasa inmovilizadas contenían una
mayor concentración de terpenos a excepción de los isómeros de los óxidos de
Resúmen de articulos
Sheila Romo Sánchez, 2013 149
piranoico, de los isómeros de los óxidos de linalol y de 6-metil-1,5-hepten-2-ol;
de fenoles como tirosol, 4-vinil guayacol y alcohol homovainíllico y de alcohol
bencílico. Aquéllos tratados con la inmovilizada por reticulación presentaron
un aumento en alcoholes C6 (1-hexanol (E), (Z)-3-hexenol), norisoprenoides
como 3-hydroxy-β-damascenona y 3-oxo-α-ionol o γ-butyrolactona, furfural, y
N(2-phenylethyl)-acetamida.
A falta de corroborar mediante Análisis Sensorial estos resultados, el uso
de la β-glucosidasa inmovilizada podría ser una realidad en Enología en un
futuro cercano, ya que además de ser estable y requerir dosis inferiores, liberó
los precursores del aroma en mostos y vinos de forma controlada.
Conclusiones generales
Sheila Romo Sánchez, 2013 153
1. Existe una biodiversidad considerable de levaduras no-Saccharomyces y
mohos tanto en aceitunas como en orujos de las variedades Cornicabra y
Arbequina. La presencia de β-glucosidasa y la ausencia de lipasas son
características de las levaduras encontradas, por otra parte A. fumigatus 3
(CECT 20827) y A. niger 113N (CECT 20828) liberaron cantidades
importantes de celulasas y xilanasas.
2. Aunque el uso de enzimas en este sector no está permitido, la presencia
de levaduras no fitopatógenas con alta o moderada actividad enzimática
mejoraría la calidad y el rendimiento de la extracción de aceite. En cuanto
a los mohos, las cepas con mejores propiedades, podrían usarse tanto
para la producción de extractos multienzimáticos como para el
aprovechamiento de subproductos agrícolas.
3. La Fermentación en Fase Sólida (FFS) de hollejos de uva y orujos de
aceituna, con o sin molienda y en presencia o no de material nitrogenado,
resulta interesante desde el punto de vista biotecnológico y
medioambiental.
4. Como es habitual en estos procesos, existe una estrecha relación entre el
binomio cepa-sustrato: A. niger (CECT 20828) crecido sobre hollejos sin
moler y en ausencia de trigo, produce cantidades importantes de xilanasa.
5. La inmovilización de extractos fúngicos multienzimáticos, purificados
parcialmente, supondría una alternativa al uso de enzimas libres en la
industria.
6. El estudio de los distintos métodos de inmovilización muestra que el
soporte más idóneo es quitina-quitosano para celulasas, xilanasas y beta-
Conclusiones generales
154 Sheila Romo Sánchez, 2013
glucosidasas comerciales, siendo la adsorción y el entrecruzamiento con
glutaraldehído las técnicas más adecuadas.
7. Entre las propiedades bioquímicas y funcionales, lo más destacable para la
celulasa es su capacidad de reutilización, y para la β-glucosidasa, la
disminución de la dosis requerida.
8. La inmovilización de la β-glucosidasa ofrece un sistema enzimático estable
que libera compuestos volátiles en mostos y vinos con dosis inferiores a la
nativa y con una excelente actividad residual tras repetidos usos.
9. En la innovación del sector enológico se podría contemplar la posibilidad
de emplear estos sistemas, resultando la aplicación de la β-glucosidasa
inmovilizada, una relación en un futuro cercano.
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