BIOTECNOLOGÍA MICROBIANA: PRODUCCIÓN, CARACTERIZACIÓN E

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


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)


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,



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



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



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



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



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.



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



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).



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



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



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



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



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.



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



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



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).



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



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).



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.



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



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),



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



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.



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



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



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



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


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


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


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Artículo II


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Artículo III


Artículo 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

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.

Artículo III


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

Artículo III


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).

http://www.merriam-webster.com/medical/pectinolytic

Artículo III


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,

http://www.cect.org/

Artículo III


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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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

http://www.webmd.com/drugs/drug-18521-Carboxymethylcellulose+Sodium+Opht.aspx?drugid=18521&drugname=Carboxymethylcellulose+Sodium+Opht

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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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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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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

Artículo III


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.

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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.

Artículo III


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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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

Artículo III


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.

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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-

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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.

References

- Alves-Baffi M, Romo-Sánchez S,

Úbeda-Iranzo J, Briones-Pérez A

(2012) Fungi isolated from olive

ecosystems and screening of their

potential biotechnological use.

New Biotechnol 29:451-456.

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Artículo IV


Artículo IV


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


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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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

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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


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

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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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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%.

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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


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

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(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


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 (%)

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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


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)

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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


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


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


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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Recovery of hydrolytic enzymes

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[2] Datta S, Christena LR, Rajaram

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[3] Cao L, Schmid RD. Carrier-

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[4] Arya SK, Srivastava SK. Kinetics

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[8] Butt MS, Tahir-Nadeem M,

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[11] Qu Y, Zhu M, Liu K, Bao X, Lin J

(2006). Studies on cellulosic

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CG, Santana CC. Crosslinking of

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G, Simpson BK, Villalonga R.

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[21] Arévalo-Villena M, Úbeda-

Iranzo JF, Gundllapalli SB, Cordero

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[22] Chang MY, Juang RS. Use of

chitosan-clay composite as

immobilization support for

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Mateos M. Effect of immobilization

on the stability of bacterial and

fungal β–D-glucosidase. Process

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[24] Pal A, Khanum F. Covalent

immobilization of xylanase on

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Wang SL. Reversible immobilization

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Kumar L, Gupta VK. Immobilization

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Tümtü’rk H. Functional polymeric

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Sarhan AA. Immobilization of

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1992;32:1–13.

[32] Kapoor M, Kuhad RC.

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http://www.springerlink.com/content/vm817m5j42315117/




Artículo V


Artículo V


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: [email protected]

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

mailto:[email protected]

Artículo V


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


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


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


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


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.

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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


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


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


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


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

Artículo V


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

Artículo V


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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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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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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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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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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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

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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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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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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



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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Volatile compounds in wine



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

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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

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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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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

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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

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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

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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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Cabrera, G., Simpson, B.K., &

Villalonga, R. (2008).

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chitosan conjugate on hyaluronic-

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modified invertase on alginate-

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Artículo V


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Artículo V


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Artículo V


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-

Resúmen de articulos


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%).

http://blast.ncbi.nlm.nih.gov/Blast.cgi



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



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.



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.



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.



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



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.



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



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).



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.



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.



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



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


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


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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BIOTECNOLOGÍA MICROBIANA: PRODUCCIÓN, CARACTERIZACIÓN E