Oxifuncionalización selectiva de compuestos alifáticos por peroxigensas fúngicas

Esteban Daniel Babot Sevilla, 2016 Oxifuncionalización selectiva de compuestos alifáticos por peroxigenasas fúngicas  OXIFUNCIONALIZACIÓN SELECTIVA DE COMPUESTOS ALIFÁTICOS POR PEROXIGENASAS FÚNGICAS Memoria que presenta: Esteban Daniel Babot para optar al título de Doctor en Ciencias Químicas por la Universidad de Sevilla.  Oxifuncionalización selectiva de compuestos alifáticos por peroxigenasas fúngicas Visado en Sevilla, a 16 de junio de 2016 LOS DIRECTORES EL TUTOR Dr. D. Fernando de Pablos Pons Catedrático de la Universidad de Sevilla Memoria que presenta: Esteban Daniel Babot Para optar al grado de Doctor en Ciencias Químicas por la Universidad de Sevilla. Dr. D. José C. del Río Andrade Profesor de Investigación del CSIC IRNAS - CSIC Dra. Dña. Ana Gutiérrez Suárez Investigador Científico del CSIC IRNAS - CSIC  DOCTOR D. JOSÉ ENRIQUE FERNÁNDEZ LUQUE, DIRECTOR DEL INSTITUTO DE RECURSOS NATURALES Y AGROBIOLOGÍA DE SEVILLA DEL CONSEJO SUPERIOR DE INVESTIGACIONES CIENTÍFICAS. CERTIFICA: Que la presente Memoria de Investigación titulada “Oxifuncionalización selectiva de compuestos alifáticos por peroxigenasas fúngicas”, presentada por Esteban Daniel Babot para optar al grado de Doctor en Ciencias Químicas, ha sido realizada en el Departamento de Biotecnología Vegetal, bajo la dirección de los Drs. Dña. Ana Gutiérrez Suárez y D. José C. del Río Andrade, reuniendo todas las condiciones exigidas a los trabajos de Tesis Doctorales. En Sevilla, a 16 de junio de 2016 AGRADECIMIENTOS Este trabajo se ha llevado a cabo en el Instituto de Recursos Naturales y Agrobiología de Sevilla (IRNAS) del Consejo Superior de Investigaciones Científicas (CSIC). Ha sido financiado por una beca JAE-Predoc del CSIC y por los proyectos europeos “Novel and more robust fungal peroxidases as industrial biocatalyst” (PEROXICATS, KBBE-2010-4-265397) e “Industrial oxidoreductases” (INDOX, GA-KBBE-2013-7-613549). Quiero expresar mi sincero agradecimiento a las personas que tanto directa como indirectamente me han acompañado durante el transcurso de esta Tesis: A los Dres. Ana Gutiérrez y José Carlos del Río, directores de esta Tesis, por haberme dado la oportunidad de realizar este trabajo de investigación en su laboratorio, por todo lo que me han aportado tanto a nivel científico como a nivel personal y por la confianza en su trato personal. Al Prof. Ángel Martínez, del Centro de Investigaciones Biológicas (CIBCSIC), por ofrecerme la posibilidad de realizar una estancia en su laboratorio, aportándome numerosos y valiosos conocimientos, y por haber seguido el desarrollo de esta Tesis. Al Prof. Fernando de Pablos Pons, Catedrático de la Universidad de Sevilla, tutor de esta Tesis, por la ayuda en la parte burocrática. A los Dres. Henrik Lund y Lisbeth Kalum, de la empresa Novozymes (Bagsvaerd, Dinamarca), por suministrarnos la peroxigenasa recombinante de Coprinopsis cinerea. Al Prof. Martin Hofrichter y al Dr. Rene Ullrich, de la Universidad de Zittau- Dresden (Zittau-Alemania), por suministrarnos la peroxigenasa de Agrocybe aegerita. A los Dres. Katrin Scheibner y Glenn Gröbe, de la empresa JenaBios (Jena, Alemania), por suministrarnos la peroxigenasa de Marasmius rotula. A los Dres. Victor Guallar y Fátima Lucas y a los doctorandos Marina Cañellas y Ferrán Sancho, por la realización e interpretación de los estudios computacionales presentados en esta Tesis. Al Dr. Manuel Angulo, del servicio de Resonancia Magnética Nuclear del CITIUS (Sevilla, España), por la realización e interpretación de los espectros de RMN de la Vitamina D.  i  ÍNDICE RESUMEN …………………………………………………………….. 1 1. INTRODUCCIÓN ..……………………………………………. 7 1.1 Reacciones de oxifuncionalización ………..…….…………….. 7 1.2 Oxidorreductasas …………………………….….…………….. 8 1.2.1 Clasificación ………………………………………………. 8 1.2.1.1 Oxidasas ……………………………………………….. 11 1.2.1.2 Deshidrogenasas .……………………………………… 11 1.2.1.3 Oxigenasas/hidroxilasas .………………………………. 11 1.2.1.4 Peroxidasas ………………………………………..… 12 1.2.2 Reacciones enzimáticas de Citocromo P450 y Peroxigenasas ……………………………………………... 18 1.2.2.1 Citocromo P450 monooxigenasas …..………………… 18 1.2.2.2 Peroxigenasas ………………………………………….. 21 2. OBJETIVOS ..…………………………………………………… 35 3. MATERIALES Y MÉTODOS ..………………………………. 39 3.1 Enzimas ..……………………………………………………... 39 3.1.1 Peroxigenasa de Agrocybe aegerita ..……………………….. 39 3.1.2 Peroxigenasa recombinante de Coprinopsis cinerea ..…….... 40 ii  3.1.3 Peroxigenasa de Marasmius rotula …….….…..…………… 40 3.2 Sustratos ..…………..……….………………………………… 41 3.3 Reacciones enzimáticas ..……….……………………………... 44 3.4 Recuperación de los productos de reacción …………………… 45 3.5 Análisis mediante GC-MS .…………………………………... 45 3.5.1 Alcanos y alquenos …………..………………….………... 45 3.5.2 Ácidos grasos y ésteres (≤ C14) ……………………………. 45 3.5.3 Ácidos grasos y ésteres (≥ C14), esteroides y secosteroides .. 45 3.5.4 Identificación y cuantificación de los productos de reacción 46 3.6 Análisis mediante Resonancia Magnética Nuclear (NMR) …. 46 3.7 Análisis computacional (PELE) ...……………………………. 46 4. RESULTADOS GENERALES Y DISCUSIÓN ..…………… 51 4.1 Oxifuncionalización selectiva de compuestos alifáticos con peroxigenasas fúngicas ..………………………………………. 51 4.1.1 Reacciones de oxifuncionalización de compuestos alifáticos lineales …………………………………………………….. 51 4.1.1.1 Ácidos grasos ………………………………………….. 51 4.1.1.2 Alcoholes grasos ………………………………………. 54 4.1.13 Alcanos y alquenos ……………………………………. 56 iii  4.1.2 Reacciones de oxifuncionalización de compuestos alifáticos cíclicos …………………………………………………….. 58 4.1.2.1 Esteroides …………..………………………………… 58 4.1.2.2 Secosteroides ……………………………..…………... 63 4.2 Mecanismo de oxigenación/oxidación de los compuestos alifáticos con peroxigenasas …………………………………. 65 4.3 Estudios computacionales ……………………………………. 65 4.3.1 Esteroides ………………………………………………… 67 4.3.2 Secosteroides ……………………………………………... 70 5. REFERENCIAS ...………………………………………………. 77 6. PUBLICACIONES ..…………………………………………… 93 6.1 Artículos científicos ………………………………………….. 97 Publicación I: Gutiérrez A., Babot E.D., Ullrich R., Hofrichter M., Martínez A.T., del Río J.C. (2011). Regioselective oxygenation of fatty acids, fatty alcohols and other aliphatic compounds by a basidiomycete hemo-thiolate peroxidase. Archives of Biochemistry and Biophysics, 514, 33-43. 97 Publicación II: Babot E.D., del Río J.C., Kalum L., Martínez A.T. and Gutiérrez A. (2012). Oxyfunctionalization of aliphatic compounds by a recombinant peroxygenase from Coprinopsis cinerea. Biotechnology and Bioengineering, 110(9), 2323-2332. ..…………………………………………….…….. 133 iv  Publicación III: Babot E.D., del Río J.C., Canellas M., Sancho F., Lucas F., Guallar V., Kalum L., Lund H., Gröbe G., Scheibner K., Ullrich R., Hofrichter M., Martínez A.T. and Gutiérrez A. (2015) Steroid hydroxylation by basidiomycete peroxygenases: A combined experimental and computational study. Applied and Environmental Microbiology, 81: 4130-4142. 163 Publicación IV: Babot E.D., del Río J.C., Kalum L., Martínez A.T. and Gutiérrez A. (2015). Regioselective hydroxylation in the production of 25-hydroxyvitamin D by Coprinopsis cinerea peroxygenase. ChemCatChem. 7, 283-290. ……………....……. 209 Publicación V: Lucas F., Babot E.D., Cañellas M., del Río J.C., Kalum L., Ullrich R., Hofrichter M., Guallar V., Martínez A.T. and Gutiérrez A. 2016. Molecular determinants for selective C25-hydroxylation of vitamins D2 and D3 by fungal peroxygenases. Catalysis Science & Technology. 6, 288-295 .….………………………………………………… 233 6.2 Patente ………………………..……………………………… 261 Patente I: Lund H., Brask J., Kalum L., Gutiérrez Suárez A., Babot E. D., Ullrich R., Hofrichter M, Martínez Ferrer A. T., del Río Andrade J. C. (2013). Enzymatic preparations of diols. WO 2013/004639. PCT/EP2012/062763. ………………….. 261 7. CONCLUSIONES ...……………………………………………. 295   1  RESUMEN La presente Tesis aborda el estudio de la oxifuncionalización enzimática de diferentes compuestos alifáticos lineales (ácidos grasos, alcoholes grasos y alcanos/alquenos) y cíclicos (esteroides y secosteroides), catalizada por diferentes peroxigenasas fúngicas (en presencia de H2O2 como co-sustrato) incluyendo las peroxigenasas salvajes de Agrocybe aegerita (AaeUPO) y Marasmius rotula (MroUPO) y la peroxigenasa recombinante de Coprinopsis cinerea (rCciUPO). La evaluación de la conversión de los distintos sustratos así como la identificación de los productos de reacción se llevó a cabo mediante Cromatografía de Gases acoplada a Espectrometría de Masas (GC-MS) y en algunos casos Resonancia Magnética Nuclear (RMN). Por otro lado, el mecanismo de reacción de dichas oxifuncionalizaciones enzimáticas se estudió utilizando H218O2 como cosustrato. Finalmente, con el objeto de investigar diferentes aspectos de las relaciones estructura-función de las peroxigenasas se realizaron estudios computacionales utilizando el programa PELE (Protein Energy Landscape Exploration). En términos generales, las peroxigenasas fúngicas AaeUPO y rCciUPO catalizaron la oxifuncionalización selectiva de los diferentes compuestos alifáticos lineales estudiados. Se observó que rCciUPO resultó ser más eficiente que AaeUPO hidroxilando los diferentes ácidos grasos (saturados e insaturados) ya que consiguió mayor grado de conversión de los sustratos (en las mismas condiciones de reacción) aunque con ambas enzimas se observó que dicho grado de conversión dependía de la longitud de la cadena del ácido graso. Por otro lado, ambas enzimas dieron lugar a los mismos productos de reacción que fueron principalmente derivados monohidroxilados (en las posiciones -1 y -2) de los ácidos grasos, tanto saturados como insaturados. Sin embargo, con el ácido miristoleico, se observó una estricta regioselectividad en las reacciones de ambas enzimas formándose exclusivamente el isómero monohidroxilado en la posición -2. Se estudiaron además, las reacciones de varios ésteres de ácidos grasos con las enzimas AaeUPO y rCciUPO observándose también la formación de derivados monohidroxilados en las posiciones -1 y -2. En cuanto a los alcoholes grasos, ambas enzimas fueron capaces de transformar estos compuestos en igual medida. Como productos principales de la reacción se obtuvieron los ácidos grasos correspondientes, así como derivados hidroxilados en las posiciones -1 y -2 de los alcoholes y de los ácidos. En las reacciones enzimáticas 2  realizadas con los alcanos se obtuvieron diferentes productos monohidroxilados en las posiciones -1 y -2, y dihidroxilados en las posiciones 2,(-1); 2,(-2) y 3,(-2). Se observó que la predominancia de los derivados mono- o dihidroxilados está relacionada con la concentración de acetona presente en el medio de reacción. Cabe mencionar que la obtención enzimática de dioles a partir de alcanos reviste gran interés industrial y se depositó una patente internacional (WO 2013/004639 A2) que se ha incluido en la Tesis. Por otro lado, también se estudió la oxifuncionalización selectiva de diferentes compuestos esteroidales (esteroles libres y esterificados, y cetonas e hidrocarburos esteroidales) por las peroxigenasas rCciUPO, AaeUPO y MroUPO. En general se observó una mayor reactividad de la rCciUPO (seguida de AaeUPO) que de la MroUPO frente la mayoría de estos compuestos esteroidales, excepto con el pregnano (que sólo fue hidroxilado por la MroUPO). La hidroxilación catalizada por estas peroxigenasas tuvo lugar en la cadena lateral (principalmente en la posición C-25) en la mayoría de los compuestos estudiados. Sólo se observó hidroxilación en el anillo (además de en la cadena lateral) en los compuestos esteroidales con dobles enlaces conjugados en C-3 y C-5 (además de en el pregnano). En esta Tesis se ha demostrado la hidroxilación regioselectiva de la vitamina D (vitamina D3 y D2) por las peroxigenasas rCciUPO y AaeUPO, en la posición C-25. La hidroxilación de estos secosteroides reviste gran interés industrial ya que además de que la reacción trascurre con una alta regioselectividad en todos los casos (100% con rCciUPO) la 25-hidroxitamina D se considera mejor agente terapéutico para diferentes enfermedades que la propia vitamina D y también tiene aplicación en alimentación animal. Con el fin de estudiar el mecanismo de la oxifuncionalización enzimática de los diferentes compuestos alifáticos mencionados, se llevaron a cabo reacciones de los mismos en presencia de H218O2, y se observó que el átomo de oxígeno introducido por las enzimas en los diferentes sustratos proviene del H2O2 (y no del O2) demostrándose la actividad peroxigenasa de las enzimas en estas reacciones de hidroxilación. Finalmente, un enfoque combinado mediante estudios experimentales y computacionales permitió demostrar que no existe un único factor que determine el perfil de hidroxilación del sustrato por la enzima, sino que es el resultado de la suma de muchos factores estructurales, tanto sobre la molécula 3  del sustrato como del sitio activo de la enzima. En el caso particular de los esteroides (que fueron los compuestos estudiados en mayor profundidad) estos factores incluyen el carácter polar/apolar de los grupos sustituyentes en las posiciones C-3/C-7 del anillo, la presencia/ausencia de dobles enlaces en el anillo esteroidal, y la longitud y características de la cadena lateral que afectan a la entrada del sustrato en el sitio activo de la enzima por C-25. En conclusión, las peroxigenasas estudiadas en esta Tesis (rCciUPO, AaeUPO y MroUPO) representan una alternativa sostenible a la síntesis química ya que catalizan la oxifuncionalización selectiva de diferentes compuestos alifáticos (tanto lineales como cíclicos) de interés en la industria química y farmacéutica, en presencia de H2O2 como único co-sustrato. 1. Introducción  10  Tabla 1 Clasificación de las diferentes oxidorreductasas según la reacción que catalizan y su dependencia de coenzimas (Feng Xu, 2005). Oxidasas Centro activo Oxidasas libres de cofactor Thr - Lys Tiol oxidasas Fe, Cu, o Cys + FAD Oxidasas que contienen Cu Cu Oxidasas que contienen flavina FMN o FAD Oxidasas con centros multi - redox Cu, Tyr, flavina, hemo De s hidrogenasas/reductasas (dependientes de NAD[P]) De s hidrogenasas que contienen flavina FMN o FAD De s hidrogenasas que contienen quinona PQQ De s hidrogenasas que contienen Zn Aminoácido + Zn De s hidrogenasas con centro multi - redox Flavina, PQQ, hemo, Fe - S, Mo Aldo - c eto reductasas Tyr - His - Asp - Lys Reductasas que contienen Cu Cu Reductasas que contienen flavina FMN o FAD Oxigenasas/hidroxilasas Oxigenasas libres de cofactor Aminoácidos Oxigenasas/hidroxilasas que contienen Fe Fe o hemo Oxigenasas/hidroxilasas que contienen Cu Cu Oxigenasas/ hidroxilasas que contienen flavina FMN o FAD Oxigenasas/hidroxilasas multicentro Fe, Mo, pterin, flavina, Fe - S Peroxidasas Hemo peroxidasas Hemo Catalasa s Hemo Haloperoxidasas Ser - His - Asp, hemo, o Val Otras peroxidasas Cys o Ser - Cys 1. Introducción  11  1.2.1.1 Oxidasas Las oxidasas, como la alcohol oxidasa, glucosa oxidasa y lacasa, entre otras, son enzimas que catalizan reacciones de oxidación o reducción utilizando simplemente como cofactor el oxígeno molecular (O2) que actúa como aceptor final de electrones. En estas reacciones el oxígeno molecular se reduce a agua (H2O) o a peróxido de hidrógeno (H2O2). Las lacasas, junto con la ascorbato oxidasa y bilirrubina oxidasa, son enzimas que contienen cuatro átomos de cobre en el sitio activo, y catalizan la oxidación (con transferencia de un electrón) de diferentes sustratos fenólicos y aminas aromáticas mientras que transfieren el electrón abstraído al O2, que es reducido a H2O (Hofrichter and Ullrich, 2010; Hollmann et al., 2011). 1.2.1.2 Deshidrogenasas Las deshidrogenasas son enzimas que catalizan la oxidación o reducción de un sustrato por sustracción de átomos de hidrógeno (deshidrogenación), los cuales son recibidos por coenzimas en su estado oxidado tales como NAD+, NADP+, FAD y FMN, que son reducidas a NADH, NADPH, FADH2 y FMNH2, respectivamente. Las deshidrogenasas catalizan la oxidación reversible de alcoholes y aminas. La alcohol deshidrogenasa, por ejemplo, es el catalizador más utilizado para la oxidación de alcoholes (Hollmann et al., 2011). 1.2.1.3 Oxigenasas/hidroxilasas Las oxigenasas e hidroxilasas son enzimas capaces de introducir un átomo de oxígeno en sustratos orgánicos con enlaces C-H y C=C desactivados y en heteroátomos, utilizando O2 y H2O2 como dadores de oxígeno. Las reacciones en que los compuestos orgánicos son oxigenados o hidroxilados son de gran valor para la síntesis orgánica. Sin embargo, como se ha mencionado anteriormente la oxifuncionalización selectiva de estos compuestos puede ser un problema en la síntesis química de compuestos orgánicos ya que a menudo se realizan con agentes oxidantes fuertes y ocurren con una baja quimio, enantio y regioselectividad (Torres Pazmiño et al., 2010; Holtmann et al., 2014). Por el contrario, estas enzimas son capaces de introducir un átomo de oxígeno en compuestos orgánicos como ácidos grasos, alcoholes, alcanos, esteroides y compuestos aromáticos, entre otros, incrementando la reactividad de la molécula 1. Introducción  12  inicial y dando lugar a la síntesis de metabolitos específicos como hormonas, antibióticos, fitoalexinas, etc., bajo condiciones medioambientalmente favorables (ej.: pH y temperaturas moderadas, solventes acuosos, etc.) (Ullrich and Hofrichter, 2007; Torres Pazmiño et al., 2010). Por otro lado, estos procesos de oxifuncionalización son de particular importancia para la iniciación de vías de degradación específicas de descontaminación. De este modo, contaminantes medioambientales como benceno, tolueno e hidrocarburos policíclicos aromáticos, por estar sujetos a la epoxidación e hidroxilación enzimática, son transformados en metabolitos asimilables (en el caso de bacterias y hongos) o en productos solubles en agua que pueden ser excretados (en el caso de animales) (Anh et al., 2007). Existen numerosas oxigenasas provenientes de diferentes organismos, que son capaces de catalizar la inserción de uno (monooxigenasas) o dos átomos de oxígeno (dioxigenasas) en moléculas orgánicas, como la tolueno monooxigenasa, el citocromo P450 monooxigenasa (P450) y la naftaleno dioxigenasa (Anh et al., 2007). La mayoría de las oxigenasas naturales utilizan como fuente el O2, pero hay también enzimas, presentes en plantas y hongos, que pueden actuar como peroxidasas/peroxigenasas transfiriendo oxígeno a partir del H2O2 o de peróxidos orgánicos (Fig. 2). La mayoría de estas reacciones, cuando son catalizadas por las mono- y dioxigenasas, suceden intracelularmente mientras que las reacciones catalizadas por las peroxidasas/peroxigenasas son extracelulares (Ullrich and Hofrichter, 2007). 1.2.1.4 Peroxidasas Las peroxidasas son enzimas secretadas, microsomales o citosólicas, que se encuentran en todos los dominios de la vida, y que catalizan reacciones de transferencia de oxígeno similares a las oxigenasas, utilizando H2O2 o hidroperóxidos orgánicos (ROOH) como co-sustratos aceptores de electrones (Hofrichter and Ullrich, 2010). Estas enzimas están involucradas en la degradación de polímeros aromáticos recalcitrantes (lignina y sustancias húmicas) y en la descontaminación oxidativa de componentes vegetales y contaminantes orgánicos. 1. Introducción  13  Fig. 2. Rutas básicas de hidroxilaciones enzimáticas de compuestos aromáticos. DO: dioxigenasa, MO: monooxigenasa, PO: peroxidasa/peroxigenasa, Tyr: Tirosinasa, DH: deshidrogenasa, Re: reordenamiento. (1) Sustrato aromático, (2) cis-dihidrodiol, (3) Producto catecólico, (4) Epóxido intermediario, (5) Producto fenólico, (6) cis,trans-dihidrodiol, (7) Sustrato fenólico, (8) Producto benzoquinona. (Adaptado de Ullrich and Hofrichter, 2007). Además, las peroxidasas son capaces de oxidar moléculas aromáticas recalcitrantes por la abstracción de un electrón, resultando en la formación de radicales inestables, que tienden a desintegrarse espontáneamente. Por tanto, debido a su versatilidad y estabilidad, las peroxidasas son de particular interés para las biooxidaciones industriales (Ullrich et al., 2004; Hofrichter et al., 2010). Según la reacción que catalizan, las peroxidasas están agrupadas bajo el número EC 1.11.1.x (peróxido de hidrógeno oxidorreductasas) en el sistema de clasificación NC-IUBMB. Actualmente se registraron 23 números EC diferentes para las peroxidasas, que comprenden desde el EC 1.11.1.1 hasta el EC 1.11.1.23, donde el número EC 1.11.1.4 se reclasificó como EC 1.13.11.11 en el grupo de las dioxigenasas (Passardi et al., 2007; Hofrichter et al., 2010) (Tabla 2). (1) (2) (3) (4) (5) (6) (7) (8) (3) (3) (4) (5) O 2 NAD + NADH DH NAD + NADH O 2 NAD + H 2 O + NADH Re H 2 O DH NAD + NADH R-OOH R-OH Re Tyr O 2 + 2H + H 2 O Tyr 2H + 1. Introducción  14  Tabla 2 Clasificación de las peroxidasas bajo los números EC 1.11.1.1 al EC 1.11.1.23 (peróxido de hidrógeno oxidorreductasas) en el sistema de clasificación y nomenclatura de enzimas del Comité de Nomenclatura de la Unión Internacional de Bioquímica y Biología Molecular (NC-IUBMB). (www.chem.qmul.ac.uk/iubmb/enzyme) Númer o EC Nombre recomendado Organismo EC 1.11.1.1 NADH p eroxidasa Clostridium acetobutylicum EC 1.11.1.2 NADPH peroxidasa Desulfovibrio desulfu ricans EC 1.11.1.3 Ácido graso peroxidasa Arachis hypogaea EC 1.11.1.4 EC 1.13.11.11, Triptófano 2,3 dioxigenasa Pseudomonas putida EC 1.11.1.5 Citocromo c peroxidasa Saccharomyces cerevisiae EC 1.11.1.6 Catalasa Mycobacterium tuberculosis EC 1.11.1.7 Peroxidasa Armoracia rusticana EC 1.11.1.8 Yoduro p eroxidasa Ascophyllum nodosum EC 1.11.1.9 Glutatión peroxidasa Mammalia EC 1.11.1.10 Cloro peroxidasa Caldariomyces fumago EC 1.11.1.11 L - A scorbato peroxidasa Pisum sativum EC 1.11.1.12 Fosfolípido hidroperóxido glutatión peroxidasa Sus scrofa EC 1.11.1.13 Manganeso peroxidasa Phanerochaete chrysosporium EC 1.11.1.14 Lignina p eroxidasa Phanerochaete chrysosporium EC 1.11.1.15 Peroxir r edoxina Haemophilus influenzae EC 1.11.1.16 Peroxidasa v ersátil Pleurotus eryngii EC 1.11.1.17 Glutatión p eroxidasa dependiente de amida Marichromatium gracile EC 1.11.1.18 Bromuro peroxidasa Corallina pilulifera EC 1.11.1.19 Peroxidasa decolorante Geotrichum candidum EC 1.11.1.20 Prostamida/prostaglandina F2 alfa sintasa Mus musculus EC 1.11.1.21 Catalasa - peroxidasa Mycobacterium tuberculosis EC 1.11.1.22 Hidroperóxido ácido graso reductasa Synechocystis PCC6803 EC 1.11.1.23 Ácido (S) - 2 - hidroxipropilfosfónico epoxidasa Streptomyces wedmorensis 1. Introducción  15  La mayoría de las peroxidasas son hemoproteínas (hemoperoxidasas) que llevan a cabo una variedad de funciones biosintéticas y degradativas actuando sobre un amplio espectro de sustratos que incluyen compuestos orgánicos e inorgánicos. El grupo prostético hemo que poseen las hemoperoxidasas, tiene un grupo protoporfirina IX donde en la parte central se encuentra el átomo de hierro (Fe) que puede formar hasta seis enlaces, de los cuales cuatro están unidos al anillo de porfirina (Fig. 3). El quinto enlace es un enlace coordinado con el nitrógeno de un residuo de histidina, la histidina distal, que sirve como catalizador ácido-base en la reacción entre el H2O2 y la enzima (Yarman et al., 2012; Bordeaux et al., 2012). El sexto enlace está ocupado por una molécula de agua fácilmente intercambiable que es desplazada por el sustrato cuando éste se une al sitio activo de la enzima (Hrycay and Bandiera, 2015). Fig. 3. Complejo Protoporfirina IX con Hierro (Fe). Aunque las hemoperoxidasas muestran diferentes especificaciones para los sustratos oxidables como indoles, fenoles, aminas aromáticas, lignina, iones Mn2+ e iones haluro, entre otros, todas ellas tienen el mismo ciclo catalítico que el P450, basado en tres etapas redox consecutivas (Fig. 4). En la primer etapa la enzima reacciona con una molécula de H2O2 en un proceso de N N O OH N O OH N Fe 1. Introducción  16  oxidación/reducción de dos electrones, donde el H2O2 se reduce a H2O y a su vez la enzima se oxida al compuesto I, que contiene un centro oxiferrilo [Fe(IV)=O] y un catión radical orgánico localizado en el grupo hemo (catión radical porfirina π). Posteriormente, el catión radical sufre una reducción de un electrón, oxidando una molécula de AH2 y dando lugar a la formación del compuesto II, el cual mantiene el centro oxiferrilo. Finalmente, el compuesto II es reducido volviendo la enzima al estado férrico inicial de reposo con la concominante oxidación de una segunda molécula de sustrato, en un proceso que es la etapa determinante de la velocidad (Ullrich and Hofrichter, 2007; Battistuzzi et al., 2010; Ortiz et al., 2010). Fig. 4. Ciclo catalítico de las hemoperoxidasas (adaptado de Battistuzzi et al., 2010). Fe III H2O H2O2 Fe IV O • + Compuesto I Fe IV O Compuesto II AH • AH2 AH2 AH • + H2O Enzima en reposo 1. Introducción  17  En base a sus propiedades estructurales, las hemoperoxidasas están clasificadas en dos superfamilias: a) la peroxidasa-ciclooxigenasa (anteriormente denominada animal o mamífero peroxidasa) y b) la peroxidasa no animal o peroxidasa vegetal. De esta manera, dependiendo de la secuencia, el organismo hospedador y la función fisiológica, la superfamilia de la peroxidasa no animal comprende tres clases principales: la Clase I, que incluye las peroxidasas intracelulares bacterianas y eucariotas localizadas en orgánulos (mitocondrias, cloroplastos, peroxisomas); la Clase II, donde se encuentran las hemoperoxidasas extracelulares fúngicas como la lignina peroxidasa (LiP), manganeso peroxidasa (MnP) y la peroxidasa versátil (VP); y finalmente la Clase III, donde están las hemoperoxidasas extracelulares vegetales que están involucradas en la síntesis de lignina. Además, existe un nuevo grupo de peroxidasas que contienen el grupo hemo y que no se encuentran incluidas en las dos superfamilias mencionadas anteriormente, las llamadas hemo-tiolato peroxidasas (HTP), que comprenden la peroxidasa clásica de Caldariomyces fumago cloroperoxidasa (CfuCPO) y las peroxigenasas de Agrocybe aegerita, Coprinopsis cinerea y Marasmius rotula, y las peroxidasas decolorantes del tinte, del hongo Auricularia auricula-judae (AauDyP) (Hofrichter et al., 2010). Las HTPs son enzimas pertenecientes a la superfamilia de las peroxidasas que, igual que éstas, poseen un grupo protoporfirina IX con la parte central unida a un átomo de Fe formando seis enlaces (Fig. 3), cuatro unidos al anillo porfirina y el sexto enlace unido a una molécula de H2O. A diferencia de las hemoperoxidasas, las HTPs tienen el quinto enlace unido al nitrógeno de un residuo de cisteína (Cys), como en los P450s (Ullrich and Hofrichter, 2007). Por otro lado, al contrario que el P450, las HTPs son enzimas extracelulares y por tanto más estables (Hofrichter and Ullrich, 2014). La CfuCPO (EC 1.11.1.10) fue la primera HTP aislada (1960) y fue la enzima de elección para las oxidaciones realizadas con peróxido y para la oxifuncionalización química (Hollmann et al., 2011; Hofrichter and Ullrich, 2014). El siguiente apartado se centra en las reacciones enzimáticas llevadas a cabo por las peroxigenasas fúngicas (objeto de esta Tesis), comparándolas con las reacciones enzimáticas realizadas por los P450s. 1. Introducción  18  1.2.2 Reacciones enzimáticas de Citocromo P450 y Peroxigenasas 1.2.2.1 Citocromo P450 monooxigenasas (EC 1.14.x.x) El citocromo P450 es una monooxigenasa que cataliza la inserción de un átomo de oxígeno en un sustrato orgánico (R-H) dando como producto final el correspondiente compuesto orgánico hidroxilado (R-OH): R-H + O2 + 2H+ + 2e– → R-OH + H2O El P450 representa una enorme y diversa superfamilia de hemoproteínas presentes en bacterias, arquaeas y eucariotas siendo la enzima hidroxilante más versátil. Su nombre viene dado porque al combinarse en su estado reducido con el monóxido de carbono (CO), forma un complejo ion Fe2+-CO que exhibe un máximo de absorción a 450 nm (Hrycay and Bandiera, 2015). Estas enzimas actúan sobre un amplio rango de compuestos exógenos como drogas, pesticidas, procarcinógenos, anestésicos y solventes orgánicos, y endógenos como esteroides, eicosanoides, ácidos grasos, hidroperóxidos lipídicos y retinoides, y además están involucradas en diferentes procesos bioquímicos intracelulares como el metabolismo de drogas, la detoxificación de pesticidas y xenobióticos, la modificación de ácidos grasos y la biosíntesis de esteroides y alcaloides (Orellana and Guajardo, 2004; Ullrich and Hofrichter, 2005; Ortiz et al., 2010; Brill et al., 2014; Shoji and Watanabe, 2014). En la actualidad, el interés industrial de los P450s se debe principalmente a la industria farmacéutica para la generación de metabolitos secundarios y la producción de precursores farmacéuticos activos, como los esteroides (Holtmann et al., 2014). Además, los P450s catalizan una amplia variedad de reacciones que incluyen epoxidación de insaturaciones C=C, N-alquilación, O-desalquilación, S- y N-oxidación, oxidación de aldehídos y alcoholes, deshalogenación, desnitración e hidroxilación de compuestos aromáticos y alifáticos y, dependiendo de la disponibilidad de oxígeno en el tejido, reacciones de reducción (Orellana and Guajardo, 2004; Ulracher and Eiben, 2006; Ullrich and Hofrichter, 2007). El ciclo catalítico de P450 (Fig. 5a) fue objeto de estudio durante décadas usando como modelo los citocromos P450CAM y P450BM3. Al iniciar el ciclo, el grupo hemo de la enzima está en su estado oxidado (Fe3+) de reposo (Fig. 5b). 1. Introducción   19   Fig. 5. (a) Ciclo catalítico del citocromo P450. Los pasos secuenciales mediante los cuales el P450 cataliza la oxidación de un sustrato (R-H) al producto monooxigenado (R-OH), son ilustrados numéricamente entre los pasos 1-7: 1) El paso 1 involucra la unión de R-H a la enzima (A), seguido de la pérdida de una molécula de H 2 O y luego, por la transferencia de un electrón (e - ), se genera la reducción del Fe 3+ presente en el grupo hemo a Fe 2+ formando un complejo P450-RH (B). 2) En el paso 2 el O 2 se une al grupo hemo-Fe 2+ para producir un complejo relativamente estable, complejo anión superóxido-ferroso [Cys-S-Fe 2+ -O 2- ] (C). 3) En el paso 3, mediante la transferencia de un e - , se genera un intermediario cargado dinegativamente, el peróxido-férrico [Cys-S-Fe 3+ -O 2- ] - (D). 4) En el paso 4, este último complejo se protona en el oxígeno distal para generar un complejo hidroperóxido-férrico [Cys-S-Fe 3+ -OOH] - (E). 5) En el paso 5, un segundo protón (H + ), produce la ruptura heterolítica del enlace O-O produciendo una molécula de H 2 O y el compuesto I (Cpd I) (F). 6) En el paso 6, el Cpd I transfiere su átomo de oxígeno ferrilo a R-H (G), generando R-OH y volviendo la enzima a su estado oxidado inicial (A) (paso 7). En el paso 2A, [Cys-S-Fe 2+ -O 2- ] puede disociarse liberando el ion Fe 2+ y O 2- y, O 2- , se disocia posteriormente en H 2 O 2 . En el paso 3A, [Cys-S-Fe 3+ -OO 2- ] - se disocia con una protonación generando H 2 O 2 . (b) El grupo hemo de la enzima en su estado oxidado (Fe 3+ ) de reposo. (Adaptado de Torres Pazmiño et al., 2010; Whitehouse et al., 2012; Krest et al., 2013; Hrycay and Bandiera, 2015). 1. Introducción   26  Fig. 9. Ciclo catalítico hipotético de AaeUPO. El ciclo catalítico comienza cuando la enzima nativa AaeUPO (A) entra en contacto con el H 2 O 2 formando un complejo AaeUPO-Peróxido ([UPO-Cpd 0] - ) (B) que luego por la pérdida de una molécula de H 2 O forma el intermediario radical oxo-ferrilo (UPO-Cpd I) (C) (flechas en azul). A partir de este paso, el ciclo catalítico puede continuar por dos vías diferentes, una mediante acción peroxigenasa (flechas en rojo), en la cual el intermediario UPO-Cpd I abstrae un e - y un H + a partir del R-H dando un Cpd II protonado (C 1 ), el complejo ferril hidróxido y un sustrato radical (R  ) localizado cerca del sitio activo de AaeUPO, que rápidamente se convierte en un producto hidroxilado (R-OH). Posteriormente, el producto R-OH se disocia de la enzima y una molécula de agua se une al grupo hemo de la UPO dando lugar al comienzo de un nuevo ciclo catalítico. En la segunda vía, por acción peroxidasa (flechas en morado), se puede encontrar el típico ciclo hemoperoxidasa donde ambos, Cpd I (C) y Cpd II (C 2 ), abstraen un electrón cada uno a partir de dos moléculas de sustrato (A-OH), los cuales se liberan como radicales libres (A-O  ) y que pueden someterse a reacciones de acoplamiento. Finalmente, la oxidación del segundo A-OH da lugar a que UPO retorne a estado inicial para comenzar con un nuevo ciclo catalítico. (Adaptado de Hofrichter et al., 2015). 1. Introducción  27  En la vía peroxigenasa (Fig. 9a), UPO-Cpd I abstrae un e- y un H+ a partir del R-H dando un Cpd II protonado, un sustrato radical (R) localizado cerca del sitio activo de AaeUPO, que rápidamente se convierte en un producto hidroxilado (R-OH) dando lugar al comienzo de un nuevo ciclo catalítico. En la vía peroxidasa (Fig. 9b) ambos, Cpd I y Cpd II, abstraen un electrón cada uno a partir de dos moléculas de sustrato (A-OH), los cuales se liberan como radicales libres (A-O) que pueden someterse a reacciones de acoplamiento. Posteriormente la oxidación del segundo A-OH da lugar a que UPO retorne a su estado inicial para comenzar con un nuevo ciclo catalítico. De este modo se puede ver que la vía peroxidasa procede por la transferencia de electrones a través de la superficie del grupo hemo, mientras que en la vía peroxigenasa, se requiere del contacto directo del sustrato con el sitio activo del grupo hemo (Hofrichter and Ullrich, 2006; Ullrich and Hofrichter, 2007; Hofrichter and Ullrich, 2010; Yarman et al., 2011; Hofrichter et al, 2015). Por otro lado, AaeUPO presenta una gran ventaja con respecto a P450, catalizando las mismas reacciones, ya que el Cpd I en el ciclo catalítico de AaeUPO tan sólo necesita de peróxido para ser activado, y además cataliza la oxidación de dos electrones del sustrato durante la transferencia del átomo de oxígeno (Fig. 9). Por el contrario, en el ciclo catalítico de P450, el Cpd I necesita una segunda flavoproteína, y un agente reductor como el NAD[P]H, para ser activado por el oxígeno molecular (Martínez et al., 2014) (Fig. 5). AaeUPO cataliza un alto número de reacciones de oxidación interesantes, incluyendo entre otras, la epoxidación/hidroxilación regioselectiva del naftaleno, la sulfoxidación del dibenzotiofeno, la N-oxidación de derivados de piridina, la O-desalquilación de alquil-aril éteres, la oxidación bencílica de derivados del tolueno, la oxidación de aril-alcoholes y aldehídos, la bromación del fenol y la oxidación de cicloalcanos (Fig. 10) (Ullrich and Hofrichter, 2007; Aranda et al., 2009; Peter et al., 2011, Poraj-Kobielska et al., 2012; Peter et al., 2013; Karich et al., 2013; Peter et al., 2014; Martínez et al., 2014; Kluge et al., 2014). Además, tal como se describe a lo largo de esta Tesis, AaeUPO también cataliza la oxidación/oxigenación de ácidos grasos, alcoholes grasos, alcanos/alquenos, esteroides y secosteroides. Muchas de estas oxidaciones selectivas representan reacciones difíciles de catalizar por la química orgánica convencional. En este contexto, AaeUPO tiene un enorme potencial biotecnológico ya que las reacciones de oxifuncionalización 1. Introducción   28  selectiva que cataliza la convierten en una herramienta biocatalítica versátil para la producción de nuevos compuestos químicos de interés industrial (Hofrichter et al., 2010). Fig. 10. Resumen general de las diferentes reacciones catalizadas por AaeUPO (adaptado de Hofrichter et al., 2015). 1. Introducción  29  Después de la primera peroxigenasa descrita en el basidiomiceto A. aegerita (Ullrich et al., 2004), se encontraron otras peroxigenasas en hongos relacionados tales como Coprinellus radians y Marasmius rotula, siendo este hallazgo indicador de la presencia generalizada de estas enzimas en el reino fúngico (Anh et al., 2007; Hofrichter et al., 2010; Gröbe et al., 2011). La investigación realizada en bases de datos con el gen de AaeUPO como sonda, permitó detectar la presencia de genes de peroxigenasas en diversos hongos (Pecyna et al., 2009), como Coprinopsis cinerea (sinónimo: Coprinus cinereus). C. cinerea es un basidiomiceto multicelular que habita en el estiércol, y fue secuenciado por el Broad Institute (http://www.broadinstitute.org) en el marco del proyecto Fungal Genome Initiative (http://www.broadinstitute.org/science/projects/fungal-genome- initiative), cuyo genoma se encuentra disponible para la búsqueda de genes en el Joint Genome Institute (JGI; www.jgi.doe.gov). Muy recientemente se identificaron más de cien genes de tipo peroxigenasa, codificados por enzimas de la superfamilia de las HTP, durante el análisis de 24 genomas de basidiomicetos (Floudas et al., 2012). La peroxigenasa salvaje de C. cinerea aún no fue aislada y caracterizada, pero uno de los genes peroxigenasa de su genoma (JGI modelo CC1G_08427T0) se expresó de forma heteróloga por la empresa Novozymes A/S (Bagsvaerd, Dinamarca) lo que dio lugar a la publicación de una patente internacional (WO/2008/119780). Esta primera peroxigenasa recombinante (rCciUPO) puede ser una herramienta poderosa para aplicaciones sintéticas debido a los niveles altos de expresión que se pueden conseguir, y a la posibilidad de modificar a medida sus propiedades catalíticas y operacionales utilizando herramientas de ingeniería de proteínas. En al año 2011 se consiguió producir en gran cantidad una nueva HTP sintetizada por el hongo M. rotula (Gröbe et al., 2011). Este bacidiomiceto, es una especie saprófita y como tal obtiene sus nutrientes de la materia orgánica muerta en descomposición. Crece en grupos o racimos sobre la madera muerta, en restos de maderas como ramas o palos, y ocasionalmente sobre hojas caducas o podridas. La peroxigenasa de M. rotula (MroUPO) presenta una estructura más pequeña que AaeUPO y rCciUPO, sin embargo, MroUPO es capaz de oxidar sustratos más voluminosos debido a que la entrada al sitio activo de la enzima es más plana y más amplia que la de AaeUPO y rCciUPO (Poraj- Kobielska, 2013). 1. Introducción  30  Además de las tres peroxigenasas mensionadas anteriormente, existen otras especies de hongos que secretan UPOs pero la purificación o caracterización de estas enzimas no se realizó aún (Hofrichter et al., 2015). En la presente Tesis se describe por primera vez la actividad catalítica de las peroxigenasas AaeUPO, rCciUPO y MroUPO sobre compuestos alifáticos incluyendo ácidos grasos, alcanos, alcoholes grasos y ésteres de ácidos grasos, así como esteroides y secosteroides, entre otros.   2. Objetivos Coprinopsis cinerea  2. Objetivos  35  2. OBJETIVOS La presente Tesis tiene como objetivo principal el estudio de la oxifuncionalización enzimática de diferentes compuestos alifáticos, lineales (ácidos grasos, alcoholes grasos, alcanos/alquenos) y cíclicos (esteroides y secosteroides) por diferentes peroxigenasas fúngicas incluyendo las peroxigenasas salvajes de Agrocybe aegerita (AaeUPO) y Marasmius rotula (MroUPO) así como la peroxigenasa recombinante de Coprinopsis cinerea (rCciUPO). Los objetivos específicos de esta Tesis son los siguientes:  Estudiar la oxifuncionalización selectiva de diversos compuestos alifáticos lineales como ácidos grasos libres y esterificados, alcoholes grasos y alcanos, y cíclicos como esteroides (esteroles libres y esterificados, hidrocarburos y cetonas) y secosteroides, mediante la acción enzimática de diversas peroxigenasas fúngicas, incluyendo rCciUPO, AaeUPO y MroUPO, en presencia de H2O2 como único co-sustrato.  Estudiar los mecanismos de las reacciones de hidroxilación de dichos compuestos alifáticos por las peroxigenasas rCciUPO, AaeUPO y MroUPO (utilizando H218O2 como co-sustrato).  Investigar diferentes aspectos de las relaciones estructura-función de las peroxigenasas en las reacciones de hidroxilación de compuestos alifáticos mediante estudios computacionales. 3. Materiales y Métodos  42  Fig. 12. Estructuras químicas representativas de los diferentes compuestos alifáticos lineales utilizados en las reacciones enzimáticas: A) ácidos grasos saturados (C12 a C20), B) ácidos grasos insaturados (C12 a C20), C) alcoholes grasos (C14 a C16), D) alcanos (C12 a C18), E) cis-1- tetradeceno, F) trans-7-tetradeceno, G) miristato de metilo, H) octanoato de octilo, I) miristato de miristilo, J) monomiristina, K) dimiristina, L) trimiristina. Los experimentos control se llevaron a cabo bajo las mismas condiciones mencionadas anteriormente pero en ausencia de enzima. Las reacciones enzimáticas llevadas a cabo con H218O2 (90% de 18O, Sigma-Aldrich) también se realizaron en las mismas condiciones descritas anteriormente. O OH AO OH B OH C D E F O O GO O H O O I OH OH O O J O O OH O O K O O O O O O L 3. Materiales y Métodos  43  Fig. 13. Estructuras químicas de los diferentes compuestos alifáticos cíclicos utilizados en las reacciones enzimáticas: (A) colesterol, (B) campesterol, (C) ergosterol, (D) sitosterol, (E) estigmasterol, (F) colestan-3-ona, (G) 4-colesten-3-ona, (H) colesta-3,5-dien-7-ona, (I) testosterona, (J) colestano, (K) colesta-3,5-dieno, (L) pregnano, (M) colesteril acetato, (N) colesteril butirato, (O) colesteril caprilato, (P) colecalciferol y (Q) ergocalciferol. HO HO HO HO HO ABC DE O OOO OH F GHIJ KL O O O O O O M NO HO HO PQ 3. Materiales y Métodos   44  3.4 Recuperación de los productos de reacción Una vez finalizadas las reacciones enzimáticas, los productos se recuperaron mediante extracción líquido-líquido con metil tert-butil éter (MTBE) (Merck). Después de una agitación vigorosa, se separó la fase orgánica (fase superior) de la fase acuosa (fase inferior) en un matraz Erlenmeyer (Fig. 14). Con la fase acuosa resultante, se repitió dos veces más el mismo procedimiento de extracción. En la fase orgánica recuperada, que contiene el sustrato remanente y los productos formados, se agregó Na 2 SO 4 anhidro (Panreac) con el objeto de eliminar restos de humedad en la muestra. Finalmente, el material previamente evaporado con N 2 , se resuspendió en cloroformo para su posterior análisis mediante cromatografía de gases acoplada a espectrometría de masas (GC-MS). Para ello, las muestras se derivatizaron previamente utilizando Bis- (trimetilsilil)trifluoroacetamida (Supelco) en presencia de piridina (90 minutos, 80 ºC). Fig. 14. Representación esquemática de la extracción líquido-líquido. Metil tert-butil éter (MTBE). 3. Materiales y Métodos  45  3.5 Análisis mediante GC-MS Los análisis de los productos de la reacción mediante GC-MS se llevaron a cabo en un cromatógrafo Varian 3800 acoplado a un detector de trampa de iones (Varian 4000) utilizando una columna capilar de sílice fundida DB-5HT (12 m x 0.25 mm de diámetro interno, con un espesor de película de 0.1 μm) de J&W Scientific. También se utilizó un equipo de GC-MS Shimadzu QP2010 Ultra, con una columna capilar de sílice fundida DB-5HT (30 m x 0.25 mm de diámetro interno, con un espesor de película de 0.1 μm). Se utilizaron distintos programas de temperatura para los distintos compuestos estudiados, como se describe a continuación: 3.5.1 Alcanos y alquenos El programa de temperatura se inició de 50 ºC (2 min) a 80 ºC (6 min) a 30 ºC·min-1, posteriormente a 250 ºC (2 min) a 8 ºC·min-1. La temperatura del inyector se mantuvo a 250 ºC durante todo el análisis. La línea de transferencia se mantuvo a 300 ºC y el gas portador utilizado fue Helio (2 mL·min-1). 3.5.2 Ácidos grasos y ésteres (≤ C14) El programa de temperatura se inició desde 50 ºC a 90 ºC (2 min) a 30 ºC·min-1, posteriormente a 250 ºC (2 min) a 8 ºC·min-1. La temperatura del inyector se mantuvo a 250 ºC durante todo el análisis. La línea de transferencia se mantuvo a 300 ºC y el gas portador utilizado fue Helio (2 mL·min-1). 3.5.3 Ácidos grasos y ésteres (≥C14), esteroides y secosteroides El programa de temperatura se inició de 120 ºC (1 min) a 300 ºC (15 min) a 10 ºC.min-1 (5 ºC. min-1 en el Shimadzu). El inyector se programó de 60 ºC (0.1 min) a 300 ºC (28 min) a 200 ºC·min-1. La línea de transferencia se mantuvo a 300 ºC y el gas portador utilizado fue el Helio (2 mL·min-1) en ambos equipos. 3. Materiales y Métodos  46  3.5.4 Identificación y cuantificación de los productos de reacción Los compuestos se identificaron por espectrometría de masas comparando sus espectros de masas con los espectros existentes en las librerías Wiley y NIST y con estándares. La cuantificación se realizó a partir del área del pico total de iones, utilizando los factores de respuesta de los mismos compuestos o compuestos similares. Los perfiles cromatográficos de un solo ion se utilizaron para estimar la abundancia de un compuesto cuando dos picos se encontraban parcialmente solapados. 3.6 Análisis mediante Resonancia Magnética Nuclear (NMR) La estructura química de algunos productos de reacción, como el 25- hidroxicolecalciferol y 25-hidroxiergocalciferol (obtenidos en las reacciones realizadas con la Vitamina D), se confirmó por espectroscopía de Resonancia Magnética Nuclear (NMR) monodimensional (1H, 13C) y bidimensional (HSQC). Para ello se utilizó un equipo Bruker Biospin AVANCE de 500 MHz acoplado a una criosonda 5 mm TCI, utilizando secuencias estándar de pulsos. 3.7 Análisis computacional (PELE) Con el fin de comprender la interacción entre el centro activo de la enzima y los diferentes compuestos alifáticos se realizaron estudios computacionales utilizando el programa Protein Energy Landscape Exploration (PELE) en el Barcelona Supercomputing Center (grupo del Dr. Victor Guallar). En estos estudios se utilizaron como estructuras de partida para las simulaciones PELE la estructura cristalina de AaeUPO (PDB entry 2YOR) y un modelo homólogo para la estructura de rCciUPO obtenida a partir de 2YOR. Además, se utilizaron como compuestos alifáticos ocho esteroides representativos (colesterol, sitosterol, colestan-3-ona, colesta-3,5-dien-7-ona, colestano, colesta-3,5-dieno, colesteril acetato y colesteril butirato) y dos secosteroides (colecalciferol y ergocalciferol). El criterio que se utilizó para las simulaciones fue el de reducir la distancia entre el átomo de oxígeno del Cpd I de la HTP y el centro de masa del ligando. A partir de los resultados obtenidos se calcularon las fracciones del ligando que entraron al centro activo de la enzima por el extremo C-25 y las fracciones que entraron por el extremo C-3.   4. Resultados Generales y Discusión Agrocybe aegerita  4. Resultados Generales y Discusión  51  4. RESULTADOS GENERALES Y DISCUSIÓN 4.1 Oxifuncionalización selectiva de compuestos alifáticos con peroxigenasas fúngicas En la presente Tesis se ha estudiado la oxifuncionalización enzimática de diferentes compuestos alifáticos lineales (ácidos grasos, alcoholes grasos, y alcanos/alquenos), y cíclicos (esteroides y secosteroides), catalizadas por diferentes peroxigenasas fúngicas incluyendo las peroxigenasas salvajes de A. aegerita (AaeUPO) y M. rotula (MroUPO) y la peroxigenasa recombinante de C. cinerea (rCciUPO). Por otro lado, también se ha estudiado el mecanismo de las reacciones de oxifuncionalización enzimática (utilizando H218O2), y se han realizado estudios computacionales con el objeto de investigar diferentes aspectos de las relaciones estructura-función de las peroxigenasas. Los principales resultados obtenidos, así como la discusión de los mismos, se incluyen a continuación. 4.1.1 Reacciones de oxifuncionalización de compuestos alifáticos lineales 4.1.1.1 Ácidos grasos Se estudiaron diferentes ácidos grasos saturados (C12 a C18), como sustratos de AaeUPO y rCciUPO. Ambas enzimas transforman estos sustratos, obteniéndose los mejores resultados con rCciUPO (Tabla 4). Como productos de reacción se obtuvieron derivados monohidroxilados, preferentemente en las posiciones -1 y -2 (Fig. 15). Además de los derivados monohidroxilados, también se identificaron ceto-derivados en estas mismas posiciones. Por otro lado, los resultados mostraron que la distribución porcentual de los regioisómeros -1 y -2 obtenidos en las reacciones con rCciUPO depende de la longitud de la cadena del ácido graso, mientras que con AaeUPO se mantiene la misma regioselectividad independientemente de la longitud de la cadena del ácido graso. La hidroxilación de ácidos grasos por el citocromo P450 en las posiciones ω, ω-1 y ω-2, se ha estudiado extensamente (Miura and Fulco, 1975; Narhi and Fulco, 1986; Muerhoff et al, 1989; Boddupalli et al., 1992; Truan et al., 1999; Johnston et al., 2011), observándose similitudes y diferencias con las reacciones de las peroxigenasas fúngicas estudiadas en la presente Tesis. 4. Resultados Generales y Discusión  58  rCciUPO (81%) obteniéndose principalmente derivados monohidroxilados en las posiciones C-2 y C-3, y en menor grado derivados dihidroxilados. Por otro lado, con AaeUPO se observó tan sólo una conversión del 13% identificándose principalmente los derivados monohidroxilados en las posiciones C-2 y C-3. La falta de actividad de las peroxigenasas con alquenos terminales puede estar relacionada con la alquilación del grupo hemo por los 1-alquenos, que resulta en la inactivación de la enzima (Peter et al., 2013). En dicho estudio se midió la actividad residual de AaeUPO después de la conversión del 1-hexeno y n-hexano bajo las mismas condiciones. Mientras que en el último caso la actividad se mantuvo, la presencia de 1-hexeno provocó la pérdida completa de la actividad de la enzima. 4.1.2 Reacciones de oxifuncionalización de compuestos alifáticos cíclicos 4.1.2.1 Esteroides Los esteroides son compuestos orgánicos derivados del núcleo del ciclopentanoperhidrofenantreno o esterano formando cuatro anillos fusionados, tres con seis átomos y uno con cinco. En los esteroides esta estructura básica se modifica por adición de diversos grupos funcionales, como carbonilos e hidroxilos o cadenas hidrocarbonadas (Fig. 18a). Los esteroides representan una clase importante de compuestos naturales que tienen una multitud de propiedades farmacológicas. La actividad fisiológica de estos compuestos depende de su estructura, el estado de oxidación de los anillos y del tipo, número y posición de los grupos funcionales presentes en la molécula (Donova and Egorova, 2012). En este sentido, pequeños cambios generados en la estructura molecular de los esteroides pueden afectar su actividad biológica, por lo que se han realizado innumerables estudios basados en la modificación estructural de los esteroides de origen natural con el objeto de encontrar nuevos compuestos con mayor actividad biológica (Beneventi et al., 2009). La hidroxilación es una de las modificaciones estructurales más importantes en la oxifuncionalización de los esteroides. En la presente Tesis se estudiaron reacciones de oxifuncionalización enzimática de varios esteroides incluyendo: (i) esteroles libres, (ii) cetonas esteroidales, (iii) hidrocarburos esteroidales y (iv) ésteres de esteroles, utilizando las peroxigenasas fúngicas rCciUPO, AaeUPO y MroUPO. 4. Resultados Generales y Discusión  59  Fig. 18. Estructura química de un esteroide (colesterol) a) y un secosteroide (colecalciferol) b). (i) Esteroles libres Se llevaron a cabo reacciones enzimáticas de cinco esteroles libres (colesterol, campesterol, ergosterol, sitosterol y estigmasterol), siendo el colesterol el sustrato con el que se obtuvo un mayor grado de conversión con respecto a los demás esteroles libres (Tabla 5). En este sentido, entre las enzimas estudiadas, rCciUPO fue la única capaz de transformar por completo el colesterol (Fig. 19). En cuanto a los productos de reacción, se obtuvieron derivados monohidroxilados principalmente en la posición C-25 y, en algunos casos, aunque en menor proporción, en las posiciones C-24, C-26 (o C-27) y C-28. En el caso de rCciUPO, se obtuvo el 25-hidroxicolesterol como único derivado monohidroxilado (Tabla 5). Por otro lado, AaeUPO presentó una mayor actividad con el campesterol (47%), ergosterol (10%) y sitosterol (13%) frente a las otras dos enzimas. Finalmente, el estigmasterol fue transformado en mayor medida por MroUPO (5%) (Tabla 5). a HO AB CD 1 2 3 456 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 HO A CD b 1 2 3 4 5 67 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 4. Resultados Generales y Discusión  60  Tabla 5 Conversión de diferentes compuestos alifáticos cíclicos por las peroxigenasas de C. cinerea, A. aegerita y M. rotula (% de sustrato transformado). Conversión de sustra to (%) Sustrato r Cc i UPO Aae UPO Mro UPO Esteroides Esteroles libres Colesterol 100 64 23 Campesterol 30 47 10 Ergosterol 6 10 7 Sitosterol 6 13 4 Es tigmasterol 2 2 5 Cetonas esteroidales 5 - Colestan - 3 - ona 54 42 12 4 - Colesten - 3 - ona 100 67 14 C olesta - 3,5 - dien - 7 - on a 97 57 39 Testosterona 0 0 0 Hidrocarburos esteroidales 5 - Colestano 14 3 1 C olesta - 3,5 - dien o 27 18 23 Pregnano 0 0 37 Ésteres de esteroles C olester i l acetat o 34 16 5 C olester il butirato 7 1 1 Colester il caprilato 0 0 0 Secosteroides Colecalciferol (Vit. D 3 ) 100 90 --- Ergocalciferol (Vit. D 2 ) 90 81 --- La presencia de un grupo metilo o etilo en el C-24, no presente en el colesterol, parece dificultar la hidroxilación en el campesterol y sitosterol, 4. Resultados Generales y Discusión  61  respectivamente. De igual forma, el doble enlace en el C-22, además del grupo metilo o etilo en el C-24 en el ergosterol y estigmasterol, respectivamente, parece ser el responsable de la baja conversión de estos compuestos por las tres peroxigenasas, en comparación con el colesterol. Fig. 19. GC-MS de la reacción de oxifuncionalización del colesterol con rCciUPO. (ii) Cetonas esteroidales En las reacciones enzimáticas realizadas con las cetonas esteroidales (colestan-3-ona, 4-colesten-3-ona, colesta-3,5-dien-7-ona y testosterona), se observó que rCciUPO fue la enzima más eficiente en transformar estos compuestos, a excepción de la testosterona que no fue modificada por ninguna 12 14 18 10 16 20 Tiempo de retención (min) Colesterol (C27) HO 25-OH-colesterol HO OH 4. Resultados Generales y Discusión  62  de las tres peroxigenasas estudiadas (Tabla 5). En cuanto a los productos obtenidos, se observó en todos los casos la hidroxilación preferente en la posición C-25 de la cadena lateral. Por otro lado, en las reacciones de la colesta-3,5-dien- 7-ona como sustrato, se observó además que AaeUPO fue la única enzima capaz de monohidroxilar esta molécula también en la posición C-26 (o C-27) y que la hidroxilación en el anillo esteroidal fue catalizada por MroUPO, y en menor medida por rCciUPO y AaeUPO. (iii) Hidrocarburos esteroidales En las reacciones enzimáticas realizadas con los hidrocarburos esteroidales (colestano, colesta-3,5-dieno y pregnano), se observó que rCciUPO fue la enzima más eficiente en transformar estos compuestos a excepción del pregnano que tan sólo fue modificado por MroUPO (Tabla 5). En cuanto a los productos obtenidos con el colesta-3,5-dieno como sustrato se observó que, además de los derivados monohidroxilados en la posición C-25 de la cadena lateral, rCciUPO, AaeUPO y MroUPO fueron capaces de hidroxilar este compuesto en el anillo esteroidal dando como productos derivados dihidroxilados (3,4-dihidroxicolest-5-eno; 3,6- y 3,6-dihidroxicolest-4- eno y x,25-dihidroxicolest-3,5-dieno) y trihidroxilados (3,6,25-trihidroxicolest- 4-eno). Finalmente, el pregnano fue modificado tan sólo por MroUPO dando lugar a la formación de varios isómeros monohidroxilados en el anillo esteroidal y un derivado oxidado, el 3-cetopregnano. La posición de los grupos hidroxilo no pudo ser determinada. (iv) Ésteres de esteroles Se estudiaron tres ésteres del colesterol (colesteril acetato, colesteril butirato y colesteril caprilato) como sustratos de las reacciones enzimáticas. Los mejores resultados se obtuvieron con rCciUPO y el colesteril acetato (Tabla 5), siendo el 25-hidroxiderivado el único producto identificado. No se observó hidroxilación en el grupo acilo. Los resultados revelaron que la presencia de un grupo esterificado dificulta la conversión de sustrato, y que este efecto está directamente relacionado con el aumento de la longitud de la cadena de los ácidos grasos. 4. Resultados Generales y Discusión  63  4.1.2.2 Secosteroides Los secosteroides son terpenoides que pertenecen a una subclase de los esteroides, similares en estructura pero que se diferencian en que el anillo B está abierto (Fig. 18b). La vitamina D pertenece a un grupo de secosteroides liposolubes que son esenciales en la homeostasis del calcio y fosfato en mamíferos. La deficiencia de la vitamina D puede causar raquitismo en los niños, osteomalacia en adultos, aumentar el riesgo de fracturas óseas en personas mayores y aumentar el riesgo de cánceres, enfermedades cardiovasculares, inmunodeficiencia, y diabetes (Holick, 2007, Bikle, 2014). La vitamina D3 (colecalciferol) se produce en la piel humana por irradiación de rayos UV provenientes de la luz del sol. Posteriormente, ésta es monohidroxilada en el hígado y dihidroxilada en los riñones dando lugar a la 25-hidroxivitamina D3 (25-hidroxicolecalciferol) y a la 1α,25-dihidroxivitamina D3 (1α,25- dihidroxicolecalciferol), respectivamente. La vitamina D2 (ergocalciferol), sigue unos pasos de activación análogos a los de la vitamina D3 para dar la 25- hidroxivitamina D2 (25-hidroxiergocalciferol) y 1α,25-dihidroxivitamina D2 (1α,25-dihidroxiergocalciferol). Se ha demostrado que los metabolitos 25- hidroxilados de la vitamina D, incluyendo 25-hidroxicolecalciferol (25- hidroxivitamina D3) y 25-hidroxiergocalciferol (25-hidroxivitamina D2), son considerablemente mejores agentes terapéuticos para diferentes enfermedades que la propia vitamina D, debido a su actividad biológica directa y su mejor absorción intestinal en algunos casos (Sitrin and Bengoa, 1987; Leichtmann et al., 1991; Jean et al., 2008). El reconocimiento de que la administración de ambas formas de la 25- hidroxivitamina D exhibe varias ventajas frente a la vitamina D, ha generado un gran interés en la síntesis química de estos compuestos (Blunt and Deluca, 1969; Batcho et al., 1982; Kyler and Watt, 1983; Adrews et al., 1986; Manchand et al., 1995; Fritts and Waldroup, 2003). Sin embargo, estas reacciones requieren a menudo agentes oxidantes fuertes que incluyen varios pasos, son costosas, y tienen lugar con una baja regioselectividad en los pasos de hidroxilación. Además del interés fisiológico de la 25-hidroxivitamina D en la salud humana, este compuesto también es de interés para la alimentación de aves de corral y otros animales de granja (Fritts and Waldroup, 2003; Simoes-Nunes and Weber, 2004; Michalczuk et al., 2010; Käppeli et al., 2011; Hernández, 2013). 4. Resultados Generales y Discusión  64  Con el objeto de sintetizar los hidroxiderivados de la vitamina D (vitamina D3 y vitamina D2), se estudiaron las reacciones enzimáticas de las peroxigenasas fúngicas AaeUPO y rCciUPO con vitamina D y se observó que son capaces de catalizar la oxifuncionalización selectiva de estos secosteroides en los derivados monohidroxilados 25-hidroxivitamina D3 y 25-hidroxivitamina D2. En este sentido, se observó que el colecalciferol fue completamente transformado por rCciUPO (100%) y en un 90% por AaeUPO. El grado de conversión del ergocalciferol fue similar para las peroxigenasas AaeUPO (81%) y rCciUPO (85%) (Tabla 5). Por otro lado, la peroxigenasa rCciUPO resultó ser altamente regioselectiva con el colecalciferol y el ergocalciferol observándose exclusivamente la formación del 25-hidroxicolecalciferol (Fig. 20) y el 25- hidroxiergocalciferol, respectivamente. En el caso de la peroxigenasa AaeUPO, se observó una alta regioselectividad para el ergocalciferol dando como único producto el 25-hidroxiergocalciferol, mientras que en el caso del colecalciferol, además del 25-hidroxiderivado, se observó en menor proporción el 24- y el 26/27-hidroxicolecalciferol. Fig. 20. GC-MS de la reacción de oxifuncionalización del colecalciferol con rCciUPO. 29 31 33 35 37 Tiempo de retención (min) 25-OH-D 3 25-OH-D 3 HO OH HO Colecalciferol 4. Resultados Generales y Discusión  65  4.2 Mecanismo de oxigenación/oxidación de los compuestos alifáticos con peroxigenasas Con el fin de estudiar el origen del oxígeno incorporado durante la oxigenación de los diferentes compuestos alifáticos y demostrar la actividad peroxigenasa de estas enzimas, se estudiaron las reacciones enzimáticas de las peroxigenasas AaeUPO, MroUPO y rCciUPO en presencia de H218O2 como cosustrato. En estos estudios se utilizaron como compuestos representativos el ácido mirístico y el tetradecanol. Las reacciones enzimáticas realizadas con H218O2 demostraron que el oxígeno introducido en la hidroxilación de los ácidos grasos en las posiciones ω-1 y ω-2 proviene del H2O2 y no del O2, a diferencia de lo que ocurre en reacciones similares catalizadas por P450BM3 (Munro et al., 2002). Por otro lado, se observó que el oxígeno incorporado en el grupo carboxílico durante la oxidación del 1-tetradecanol a ácido mirístico, y sus derivados hidroxilados ω-1 y ω-2, también proviene del H2O2 (Fig. 21), demostrando así la actividad peroxigenasa de estas enzimas en estas reacciones. En este sentido, el primer producto que se formaría en la oxidación del alcohol por las peroxigenasas sería un gem-diol debido a la hidroxilación del C-1 (Fig. 21A), el cual posteriormente podría ser por un lado directamente hidroxilado produciendo un gem-triol que se deshidrataría irreversiblemente dando el grupo ácido (Fig. 21B), o bien primero deshidratado en el aldehído correspondiente y posteriormente hidroxilado para dar el ácido (Fig. 21C). 4.3 Estudios computacionales Con el objeto de investigar diferentes aspectos de las relaciones estructurafunción de las peroxigenasas se realizaron estudios computacionales utilizando el programa Protein Energy Landscape Exploration (PELE). En estos estudios la estructura de partida para las simulaciones PELE fue la estructura cristalográfica de AaeUPO (PDB entry 2YOR). Se utilizaron como compuestos alifáticos ocho esteroides y dos secosteroides. El criterio que se tuvo para las simulaciones fue el de reducir la distancia entre el átomo de oxígeno del Cpd I de la HTP y el centro de masa del ligando. A partir de los resultados se calcularon las fracciones del ligando que entraron al centro activo de la enzima por el extremo C-25 y las fracciones que entraron por el extremo C-3. 4. Resultados Generales y Discusión  66  R─ CH 2 ─ (CH 2 ) n ─ CH 16 OH 18 OH C) + >1/2 R─ CH 2 ─ (CH 2 ) n ─ C 16 O H <1/2 R─ CH 2 ─ (CH 2 ) n ─ C 18 O H >1/2 H 218 O <1/2 H 216 O HTP H 218 O 2 HTP H 218 O 2 HTP H 218 O 2 R─ CH 2 ─ (CH 2 ) n ─ CH 216 OH R─ CH 2 ─ (CH 2 ) n ─ CH 16 OH 18 OH A) 12 R─ CH 2 ─ (CH 2 ) n ─ CH 16 OH 18 OH B) HTP H 218 O 2 R─ CH 2 ─ (CH 2 ) n ─ C ─ 18 OH 16 OH 18 OH 2/3 H 218 O 1/3 H 216 O 1/3 R─ CH 2 ─ (CH 2 ) n ─ C + 2/3 R─ CH 2 ─ (CH 2 ) n ─ C 18 O 18 O 16 O 18 O - - 1/3 R─ CH ─ (CH 2 ) n ─ C HTP H 218 O 2 HTP H 218 O 2 + 2/3 R─ CH ─ (CH 2 ) n ─ C 18 O 18 O 16 O 18 O - - 18 OH 18 OH 23 45 67 289 4 and 65 and 7 Fig. 21. Posible mecanismo de oxidación de alcoholes grasos, por la hemo-tiolato peroxidasa de A. aegerita, mediante el cual se producen los ácidos grasos correspondientes. Los estudios realizados con H218O2 incluyen: A) La hidroxilación del alcohol (1) a un gem-diol con 18O marcado (2); B) La oxidación directa del gem-diol a un gem-triol intermedio (3) que por deshidratación irreversible produce ácidos grasos (4 y 5) con uno o dos 18O marcados, que posteriormente son enzimáticamente hidroxilados (en la posición ω-1 en el esquema) produciendo los correspondientes hidroxiácidos (6 y 7); y C) La deshidratación del gem-diol produce el aldehido, con 18O marcado (8) y 16O no marcado (9), que puede experimentar un intercambio variable de hidroxilo con el solvente resultando en la reducción del 18O marcado, antes de ser hidroxilado sucesivamente por la enzima produciendo los mismos ácidos grasos (4 y 5) e hidroxiácidos (6 y 7) que en el paso B. 4. Resultados Generales y Discusión   67  Los resultados permitieron establecer una correlación directa entre el porcentaje de conversión del sustrato (Tabla 5) y el porcentaje de entrada del sustrato por C-25 (Fig. 22). Fig. 22. Gráfica de correlación entre el porcentaje de conversión del sustrato y la entrada del sustrato por C-25 (cadena lateral) en el sitio activo de la enzima. A: colesterol, D: sitosterol, F: colestan-3-ona, H: colesta-3,5-dien-7-ona, J: colestano, K: colesta-3,5-dieno, M: colesteril acetato, O: colesteril caprilato. 4.3.1 Esteroides Teniendo en cuenta los criterios mencionados anteriormente, se utilizaron como esteroides representativos el colesterol, sitosterol, colestan-3-ona, colesta- 3,5-dien-7-ona, colestano, colesta-3,5-dieno, colesteril acetato y colesteril caprilato. De estos esteroides, el colesterol fue el compuesto de mejor trayectoria de entrada al sitio activo de la enzima por el C-25 (82 %) (Fig. 22), debido a que el canal de acceso al hemo hidrófobo de la peroxigenasa de AaeUPO favorece la entrada de este compuesto por el lado apolar de la molécula. Además, una vez que la molécula se encuentra posicionada en el sitio activo de la enzima, ésta se  5. Referencias Coprinopsis cinerea  5. Referencias  77  5. REFERENCIAS Andrews DR, Barton DHR, Hesse RH, Pechet MM. 1986. Synthesis of 25- hydroxy- and 1-alpha-25-dihydroxy vitamin D3 from vitamin D2 (calciferol). Journal of Organic Chemistry 51: 4819-4828. Anh DH, Ullrich R, Benndorf D, Svatos A, Muck A, Hofrichter M. 2007. The coprophilous mushroom Coprinus radians secretes a haloperoxidase that catalyzes aromatic peroxygenation. Applied and Environmental Microbiology 73: 5477-5485. Aranda E, Kinne M, Kluge M, Ullrich R, Hofrichter M. 2009. 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Hofrichter M, Ullrich R. 2014. Oxidations catalyzed by fungal peroxygenases. Current Opinion in Chemical Biology 19: 116-125. Hofrichter M, Ullrich R. 2010. New trends in fungal Biooxidation. The Mycota. Industrial applications 10: 425-449. Hofrichter M, Ullrich R, Pecyna MJ, Liers C, Lundell T. 2010. New and classic families of secreted fungal heme peroxidase. Applied Microbiology and Biotechnology 87: 871-897. Hofrichter M, Ullric R. 2006. Heme-thiolate haloperoxidases: versatile biocatalysts with biotechnological and environmental significance. Applied Microbiology and Biotechnology 71: 276-288. Holtmann D, Fraaije MW, Arends IWCE, Opperman DJ, Hollmann F. 2014. The taming of oxygen: biocatalytic oxyfunctionalisations. Chemical Communications 50: 13180-13200. Hollmann F, Arends I, Buehler K, Schallmey A, Büuhler B. 2011. Enzymemediated oxidations for the chemist. Green Chemistry 13: 226-265. Hrycay EG, Bandiera SM. 2015. The monooxygenase, peroxidase, and peroxygenase properties and reaction mechanisms of cytochrome P450 enzymes. Advances in Experimental Medicine and Biology 851: 1-61. Hrycay EG, Bandiera SM. 2012. The monooxygenase, peroxidase, and peroxygenase properties of cytochrome P450. Archives of Biochemistry and Biophysics 522: 71-89. 5. Referencias  81  Hudlick T, Reed JW. 2009. Applications of biotransformations and biocatalysis to complexity generation in organic synthesis. Chemical Society Reviews 38: 3117-3132. Ishimaru A. 1980. On the structure-function relationship of peroxidases and peroxygenase. Bioorganic Chemistry 9: 472-481. Jean G, Terrat JC, Vanel T, Hurot JM, Lorriaux C, Mayor B, Chazot C. 2008. Daily oral 25-hydroxycholecalciferol supplementation for vitamin D deficiency in haemodialysis patients: effects on mineral metabolism and bone markers. Nephrol Dial Transplant 23: 3670-3676. Johnston JB, Ouellet H, Podust LM, Ortiz de Montellano PR. 2011. Structural control of cytochrome P450-catalyzed ω-hydroxylation. Archives of Biochemistry and Biophysics 507: 86-94. Karich A, Kluge M, Ullrich R, Hofrichter M. 2013. Benzene oxygenation and oxidation by the peroxygenase of Agrocybe aegerita. Applied and Industrial Microbiology and Biotechnology Express 3: 5. Kellner H, Luis P, Pecyna MJ, Barbi F, Kapturska D, Krüger D, Zak DR, Marmeisse R, Vandenbol M, Hofrichter M. 2014. Widespread occurrence of expressed fungal secretory peroxidases in forest soils. PLoS ONE 9(4): e95557. Kinne M, Poraj-Kobielska M, Ullrich R, Nousiainen P, Sipilä J, Scheibner K, Hammel KE, Hofrichter M. 2011. Oxidative cleavage of non-phenolic β- O-4 lignin model dimers by an extracellular aromatic peroxygenase. Holzforschung 65: 673-679. Kinne M, Zeisig C, Ullrich R, Kayser G, Hammel KE, Hofrichter M. 2010. Stepwise oxygenations of toluene and 4-nitrotoluene by a fungal peroxygenase. Biochemical and Biophysical Research Communications 397: 18-21. 5. Referencias  82  Kinne M, Poraj-Kobielska M, Ralph SA, Ullrich R, Hofrichter M, Hammel KE. 2009. Oxidative cleavage of diverse ethers by an extracellular fungal peroxygenase. The Journal of Biological Chemistry 284(43): 29343-29349. Kinne M, Poraj-Kobielska M, Aranda E, Ullrich R, Hammel KE, Scheibner K, Hofrichter M. 2009. Regioselective preparation of 5-hydroxypropranolol and 4-hydroxydiclofenac with a fungal peroxygenase. Bioorganic & Medicinal Chemistry Letters 19: 3085-3087. Kinne M, Ullrich R, Hammel KE, Scheibner K, Hofrichter M. 2008. Regioselective preparation of (R)-2-(4-hydroxyphenoxy) propionic acid with a fungal peroxygenase. Tetrahedron Letters 49: 5950-5953. Kluge M, Ullrich R, Scheibner K, Hofrichter M. 2014. Formation of naphthalene hydrates in the enzymatic conversion of 1,2- dihydronaphthalene by two fungal peroxydases and subsequent naphthalene formation. Journal of Molecular Catalysis B: Enzymatic 103: 56-60. Kluge M, Ullrich R, Scheibner K, Hofrichter M. 2012. Stereoselective benzylic hydroxylation of alkylbenzenes and epoxidation of styrene derivatives catalyzed by the peroxygenase of Agrocybe aegerita. Green Chemistry 14: 440-446. Kluge M, Ullrich R, Dolge C, Scheibner K, Hofrichter M. 2009. Hydroxylation of naphthalene by aromatic peroxygenase from Agrocybe aegerita proceeds via oxygen transfer from H2O2 and intermediary epoxidation. Applied Microbiology and Biotechnology 81: 1071-1076. Kluge MG, Ullrich R, Scheibner K, Hofrichter M. 2007. Spectrophotometric assay for detection of aromatic hydroxylation catalyzed by fungal haloperoxidase-peroxygenase. Applied Microbiology and Biotechnology 75: 1473-1478. 5. Referencias  83  Krest CM, Onderko EL, Yosca TH, Calixto JC, Karp RF, Livada J, Rittle J, Green MT. 2013. Reactive intermediates in cytochrome P450 catalysis. The Journal of Biological Chemistry 288 (24): 17074-17081. Kyler KS, Watt DS. 1983. Partial synthesis of (20R)-25-hydroxycholesterol involving a nickel (II)-promoted dienol rearrangement. Journal of the American Chemical Society, 105: 619-621. Leichtmann GA, Bengoa JM, Bolt MJG, Sitrin MD. 1991. Intestinal absorption of cholecalciferol and 25-hydroxycholecalciferol in pathients with both Crohn's disease and intestinal resection. The American Journal of Clinical Nutrition, 54: 548-552. Li H. 2001. Cytochrome P450, in: Messerschmidt A, Huber R, Poulos TL, Wieghardt K. (Eds.), Handbook of Metalloproteins. Wiley, Baffins Lane, UK, pp. 267-282 Li HY, Poulos TL. 1997. The structure of the cytochrome P450BM-3 haem domain complexed with the fatty acid substrate, palmitoleic acid. Nature Structural and Molecular Biology 4: 140-146. Manchand PS, Yiannikouros GP, Belica PS, Madan P. 1995. Nickel-mediated conjugate addition. Elaboration of calcitriol from ergocalciferol. Journal of Organic Chemistry 60: 6574-6581. Martínez AT, Ruiz-Dueñas FJ, Gutiérrez A, del Río JC, Alcalde M, Liers C, Ullrich R, Hofrichter M, Scheibner K, Kalum L, Vind J, Lund H. 2014. Search, engineering, and applications of new oxidative biocatalysts. Biofuels, Bioproducts and Biorefining 8: 819-835. May SW. 1999. Applications of oxidoreductases. Current Opinion in Biotechnology 10: 370-375. Miura Y, Fulco AJ. 1975. ω-1, ω-2 and ω-3 hydroxylation of long-chain fattyacids, amides and alcohols by a soluble enzyme-system from Bacillus 6. Publicaciones Marasmius rotula  6. Publicaciones  93  6. PUBLICACIONES Los principales resultados obtenidos durante la realización de esta Tesis han dado lugar a las siguientes publicaciones y patente: 6.1 Artículos científicos I. Gutiérrez A., Babot E.D., Ullrich R., Hofrichter M., Martínez A.T., del Río J.C. (2011). Regioselective oxygenation of fatty acids, fatty alcohols and other aliphatic compounds by a basidiomycete hemo-thiolate peroxidase. Archives of Biochemistry and Biophysics 514: 33-43. II. Babot E.D., del Río J.C., Kalum L., Martínez A.T. and Gutiérrez A. (2012). Oxyfunctionalization of aliphatic compounds by a recombinant peroxygenase from Coprinopsis cinerea. Biotechnology and Bioengineering 110(9): 2323- 2332. III. Babot E.D., del Río J.C., Cañellas M., Sancho F., Lucas F., Guallar V., Kalum L., Lund H., Gröbe G., Scheibner K., Ullrich R., Hofrichter M., Martínez A.T. and Gutiérrez A. (2015) Steroid hydroxylation by basidiomycete peroxygenases: A combined experimental and computational study. Applied and Environmental Microbiology 81:4130-4142. IV. Babot E.D., del Río J.C., Kalum L., Martínez A.T. and Gutiérrez A. (2014). Regioselective hydroxylation in the production of 25-hydroxyvitamin D by Coprinopsis cinerea peroxygenase. ChemCatChem 7: 283-290. V. Lucas F., Babot E.D., Cañellas M., del Río J.C., Kalum L., Ullrich R., Hofrichter M., Guallar V., Martínez A.T. and Gutiérrez A. (2016). Molecular determinants for selective C25-hydroxylation of vitamins D2 and D3 by fungal peroxygenases. Catalysis Science & Technology. 6: 288-295. 6. Publicaciones  94  6.2 Patente I. Lund H., Brask J., Kalum L., Gutiérrez Suárez A., Babot E. D., Ullrich R., Hofrichter M., Martínez Ferrer A. T., del Río Andrade J. C. (2013). Enzymatic preparations of diols. WO 2013/004639. PCT/EP2012/062763.  6. Publicaciones  97  6.1. Publicación I: Gutiérrez A., Babot E.D., Ullrich R., Hofrichter M., Martínez A.T., del Río J.C. (2011). Regioselective oxygenation of fatty acids, fatty alcohols and other aliphatic compounds by a basidiomycete hemo-thiolate peroxidase. Archives of Biochemistry and Biophysics 514: 33-43. 6. Publicaciones  99  Regioselective oxygenation of fatty acids, fatty alcohols and other aliphatic compounds by a basidiomycete heme-thiolate peroxidase Ana Gutiérreza*, Esteban D. Babota, René Ullrichb, Martin Hofrichterb, Angel T. Martínezc, José C. del Ríoa aInstituto de Recursos Naturales y Agrobiología de Sevilla, CSIC, PO Box 1052, E-41080 Seville, Spain bUnit of Environmental Biotechnology, International Graduate School of Zittau, Markt 23, 02763 Zittau, Germany cCentro de Investigaciones Biológicas, CSIC, Ramiro de Maeztu 9, E-28040 Madrid, Spain * Corresponding author Abstract: Reaction of fatty acids, fatty alcohols, alkanes, sterols, sterol esters and triglycerides with the so-called aromatic peroxygenase from Agrocybe aegerita was investigated using GC-MS. Regioselective hydroxylation of C12-C20 saturated/unsaturated fatty acids was observed at the -1 and -2 positions (except myristoleic acid only forming the -2 derivative). Minor hydroxylation at  and -3 to -5 positions was also observed. Further oxidized products were detected, including keto, dihydroxylated, keto-hydroxy and dicarboxylic fatty acids. Fatty alcohols also yielded hydroxy or keto derivatives of the corresponding fatty acid. Finally, alkanes gave, in addition to alcohols at positions 2 or 3, dihydroxylated derivatives at both sides of the molecule; and sterols showed side-chain hydroxylation. No derivatives were found for fatty acids esterified with sterols or forming triglycerides, but methyl esters were -1 or -2 hydroxylated. Reactions using H218O2 established that peroxide is the source of the oxygen introduced in aliphatic hydroxylations. These studies also indicated that oxidation of alcohols to carbonyl and carboxyl groups is produced by successive hydroxylations combined with one dehydration step. We conclude that the A. aegerita peroxygenase not only oxidizes aromatic compounds but also catalyzes the stepwise oxidation of aliphatic compounds by hydrogen peroxide, with different hydroxylated intermediates. A. Gutiérrez et al. / Archives of Biochemistry and Biophysics (2011) 514: 33-43. 106  Fig. 1. GC-MS analysis of the peroxygenase reactions with saturated C12 to C20 (left) and mono-unsaturated C12:1 to C20:1 (right) fatty acids showing the remaining substrate, and the different -1 to -2 keto,  to -5 hydroxyl, dicarboxylic, hydroxy-keto and dihydroxy derivatives (see SD, Fig.S1 for chemical structures, and Table 1 for quantitative values). The most adequate chromatographic program was used in each case. C 12 C 14 C 16 C 18 C 20 (-2)-OH (-1)-OH (-1)-keto (-2)-keto -OH di-COOH (-3)-OH OH-keto di-OH (-4)-OH (-5)-OH (-2)-OH (-1)-OH (-3)-OH (-2)-keto (-1)-keto -OH (-2)-OH (-1)-OH (-2)-keto (-1)-keto -OH (-2)-OH (-1)-OH (-2)-keto (-1)-keto Retention time (min) (-2)-OH (-1)-OH C 12:1 C 14:1 C 16:1 C 18:1 C 20:1 (-2)-OH (-1)-OH -OH (-1)-keto (-2)-keto -OH (-2)-OH (-2)-keto (-2)-OH (-1)-OH -OH (-1)-keto (-2)-keto -OH (-1)-keto (-2)-keto (-2)-OH (-1)-OH (-3)-OH (-2)-OH (-1)-OH 510 10 15 510 10 10 15 10 15 10 15 20 510 10 10 15 Retention time (min) 6. Publicaciones  107  Fig. 2. Mass spectra of -1 (A) and -2 (B) monohydroxylated, and -1 (C) and -2 (D) keto derivatives of myristic acid (C14) from peroxygenase reactions with myristic acid, as trimethylsilyl (TMS) derivatives. 50 100 150 200 250 300 350 400 m/z 100% 73 75 129 131 149 359 373 204 283 217 [M + ] 388 OTMS OTMS m/z 131 m/z 359 B 50 100 150 200 250 300 350 400 m/z 100% 73 75 117 129149 204 357373 283 217 [M + ] 388 OTMS OTMS m/z 117 m/z 373 A 50 100 150 200 250 300 350 400 100% 73 75 117 129 153 243 299 281 93 OTMS [M + ] 314 [M - CH 3 -CH 3 CH 2 CO] + [M - CH 3 –H 2 O] + [M - CH 3 ] + m/z D 50 100 150 200 250 300 350 400 100% 73 75 117 129 149 257 299 281 167 93 OTMS [M + ] 314 [M - CH 3 -CH 3 CO] + [M - CH 3 ] + [M - CH 3 –H 2 O] + m/z O C O O O O O A. Gutiérrez et al. / Archives of Biochemistry and Biophysics (2011) 514: 33-43. 108  Table 2 Abundance (relative percentage) of different monohydroxylated, keto, dihydroxylated, ketohydroxy and dicarboxylic derivatives (at ω to ω-5) identified by GC-MS in the reactions of saturated and unsaturated fatty acids (from 12 to 20 carbons) with the peroxygenase (see Fig. 1 for chromatographic profiles and SD, Fig.S1 for chemical structures), and -1/-2 ratio of the main (monooxygenated) products formed. ω- OH ω-1 OH ω-2 OH ω-3 OH ω-4 OH ω-5 OH ω-1 keto ω-2 keto di- OH OH- keto di- COOH ω-1/ ω-2 C12 1.3 39.7 32.0 0.2 <0.1 <0.1 5.8 1.0 4.4 15.5 0.3 1.4 C12:1 3.3 37.4 59.2 0 0 0 <0.1 <0.1 0 0 0 0.6 C14 3.5 34.4 30.5 0.3 0 0 20.8 3.3 0.5 6.2 0.6 1.6 C14:1 1.8 0 94.6 0 0 0 0 3.6 0 0 0 0 C16 1.4 23.6 23.6 0.3 0 0 34.5 16.3 0 0 0.3 1.5 C16:1 2.5 35.7 47.0 0.1 0 0 10.4 4.4 0 0 0 0.9 C18 <0.1 22.7 27.0 0.1 0 0 32.8 17.0 0 0 0.5 1.3 C18:1 1.6 40.8 39.0 0.2 0 0 13.0 5.3 0 0 0 1.2 C18:2 1.0 50.2 33.5 2.5 0 0 10.0 2.9 0 0 0 1.7 C20 <0.1 16.0 28.1 0 0 0 38.7 17.3 0 0 0 1.2 C20:1 1.2 35.0 38.7 0.4 0 0 18.8 6.0 0 0 0 1.2 An 18O-labeling study, using myristic acid as substrate, and either H218O2 or H216O2 as enzyme cosubstrate, was performed to investigate the origin of the oxygen incorporated during the oxygenation of fatty acids. In the reaction using H218O2, mass spectral analysis of the resulting monohydroxylated fatty acids showed that characteristic fragments for the (-1)-hydroxyfatty acid had ~90% shifted from the natural abundance m/z 117 and m/z 373 found in the unlabeled peroxide reaction (Fig. 2A) to m/z 119 and m/z 375 (Fig. 3A), respectively. Likewise, the characteristic fragments for the (-2)-hydroxyfatty acid shifted from the natural abundance m/z 131 and m/z 359 found in the unlabeled peroxide reaction (Fig. 2B) to m/z 133 and m/z 361 (Fig. 3B), respectively. In both hydroxyfatty acids, ~10% of the original fragments remained in the H218O2 reactions due to the 90% 18O isotopic purity of the labeled peroxide used. 6. Publicaciones  109  Fig. 3. Mass spectra of -1 (A) and -2 (B) 18O-labeled monohydroxylated myristic acid (C14) from peroxygenase reactions with myristic acid in the presence of H218O2 (90% isotopic purity) as trimethylsilyl (TMS) derivatives. In addition to hydroxylated derivatives, keto derivatives at positions -1 and -2 were also identified (Fig. 1). The position of the carbonyl group was determined by the mass spectra of the trimethylsilyl derivatives, as shown in Figs. 2C and 2D for (-1) and (-2)-ketomyristic acid, respectively. No molecular ions were observed in the mass spectra, although they could be determined from the [M - CH3]+ fragment (m/z 299 in both ketomyristic acids). Additional losses of the methylketo [M - CH3 - CH3CO]+ or ethylketo [M - CH3 - CH3CH2CO]+ groups from α-cleavage of the carbonyl group in (-1) and (-2)-ketofatty acids, respectively, produced other diagnostic fragments (at 73 100% 50 100 150 200 250 300 350 400 75 95 133 204 361 375 217 117 m/z 283 [M + ] 390 73 75 95117 119 204 217 359375 50 100 150 200 250 300 350 400 100% m/z 283 [M + ] 390 OTMS 18 OTMS m/z 119 m/z 375 B A m/z 133 OTMS 18 OTMS m/z 361 90% 90% O O A. Gutiérrez et al. / Archives of Biochemistry and Biophysics (2011) 514: 33-43. 110  m/z 257 for 13-ketomyristic acid, and m/z 243 for 12-ketomyristic acid). Another characteristic fragments resulted from the loss of a water molecule [M - CH3 -H2O]+ (at m/z 281 in both ketomyristic acid isomers). The ratio between the two main isomers (-1/-2) among the total monooxygenated derivatives (hydroxy and ketofatty acids) for the different compounds assayed is shown in Table 2. This ratio was very similar for all the saturated fatty acids (from 1.2 to 1.6). Unsaturated fatty acids often showed lower ratios (ranging from 0.6 to 1.2) with the exception of linoleic acid (1.7 ratio), and myristoleic acid that only formed the -2 derivative. In the enzyme-catalyzed reaction of myristic acid with H218O2, mass spectral analysis of the (-1) and (-2)-ketofatty acids formed (together with the same hydroxyfatty acids) showed that the above mentioned [M - CH3]+ fragment (shared by both keto isomers) had ~10% shifted from the natural abundance m/z 299 found in the unlabeled peroxide reaction (Fig. 2) to m/z 301 (spectrum not shown). This 18O-labeling is much lower than the 90% observed during formation of the corresponding hydroxyfatty acids, and revealed 18O loss from the carbonyl formed, due to hydroxyl exchange with water. Further oxygenation/oxidation of the above mono-substituted hydroxy and keto derivatives were observed in the enzymatic reactions with fatty acids, including dihydroxylated compounds and several combinations of keto and hydroxy derivatives. The chemical structures of all the fatty-acid derivatives identified in the different enzymatic reactions performed are shown in Supplementary data. SD. Fig. S1. With the aim of investigating whether it was necessary that the carboxyl group was in free form for enzyme activity, methyl laurate was assayed as substrate. The results showed that the enzyme is able to transform the methyl esters (Table 1). Like in the reaction with free lauric acid, the reaction gave two main monohydroxylated derivatives at -1 and -2 positions, with a -1/-2 ratio of 1.3 (similar to that obtained for the free fatty acids). Fatty-alcohol oxidation studies Two primary fatty alcohols (1-tetradecanol and 1-hexadecanol) were assayed as substrates for the A. aegerita peroxygenase (Table 1). Among the reaction products identified (Fig. 4 and Fig. S2 of SD), the presence of the 6. Publicaciones  111  corresponding fatty acids (myristic and palmitic acids, respectively) is noteworthy. The majority of the additional products were hydroxy or keto derivatives at -1 and -2 positions of the fatty acids (-hydroxymyristic and -hydroxypalmitic acids were also identified). The -1/-2 ratio of these monooxygenated derivatives was 1.4 (similar to that obtained directly from myristic and palmitic acids). Some minor peaks were tentatively assigned to monohydroxylated (and keto) derivatives of the fatty alcohols at -1 and -2 positions, which were more evident in the reaction with tetradecanol. Only traces of the corresponding aldehydes were observed in the reactions. Fig. 4. GC-MS analysis of the peroxygenase reaction with tetradecanol, showing the remaining fatty alcohol substrate (Alc), the corresponding fatty acid (Acd) and its different - 1 and -2 keto,  to -5 hydroxyl, dicarboxylic, hydroxy-keto and dihydroxy derivatives formed (see SD, Fig. S2 for chemical structures, and Table 1 for quantitative values). Other minor compounds such as -1 and -2 keto and hydroxy derivatives of the tetradecanol and traces of the aldehyde (Ald) are also shown. 10 15 Retention time (min) Alc Acd -1 OH Acd -2 OH Acd -1+ -2 keto Acd + -1+ -2 OH Alc Ald OH Acd -1+-2 keto Alc A. Gutiérrez et al. / Archives of Biochemistry and Biophysics (2011) 514: 33-43. 112  18O-labeling studies were also performed in the reactions of 1-tetradecanol with the enzyme (and similar results were obtained in the 1-hexadecanol reactions). Incorporation of 18O from H218O2 to the carboxyl group was observed during the oxidation of 1-tetradecanol to myristic acid (Fig. 5). Mass spectral analysis of the myristic acid formed, showed that the characteristic fragment at [M - CH3]+ had shifted from the natural abundance m/z 285 found in the unlabeled peroxide reaction (Fig. 5A) to m/z 289 (incorporation of two 18O atoms at the carboxyl group) and m/z 287 (incorporation of one 18O atom at the carboxyl group) (Fig. 5B). The relative abundances of the 18O mono (74%) and bi-labeled (16%) myristic acid, estimated from the corresponding fragment peak areas, are included in Fig. 5B, together with their formulae (10% unlabeled acid was also formed due to the partial isotopic purity of the peroxide used). Fig. 5. Mass spectra of myristic acid (C14), from peroxygenase reactions with 1-tetradecanol in the presence of H216O2 (A) and H218O2 (90% isotopic purity) (B), as trimethylsilyl (TMS) derivatives, and formulae for the unlabeled compound found in the A, and the mono-, and bilabeled (top to bottom) compounds found in B with indication of their characteristic ions and relative abundances (10% unlabeled acid was also found in the second reaction). 285 280 290 300 310 286 300287 100% m/z A 280 290 300 310 285 287 289 302 100% m/z B m/z 285 16 OTMS 16 O m/z 287 18 OTMS 16 O 74% m/z 289 18 OTMS 18 O 16% 6. Publicaciones  113  The incorporation of 18O from H218O2 was also evidenced in the mass spectra of the -1 or -2 hydroxylated derivatives of myristic acid, formed in the same 1-tetradecanol reaction. The mass spectral analysis showed that the characteristic fragments for the (-1) and (-2)-hydroxylated derivatives of fatty acids had ~90% shifted from the natural abundance m/z 117 and m/z 131 to m/z 119 and m/z 133, respectively (spectra not shown) due to peroxide 18O incorporation. Moreover, the characteristic fragments for the -1 at [M - CH3]+ had shifted from the natural abundance m/z 373 found in the unlabeled peroxide reaction (Fig. 6A) to m/z 379 (incorporation of three 18O atoms), m/z 377 (incorporation of two 18O atoms) and m/z 375 (incorporation of one 18O atom) (Fig. 6C). Likewise, the characteristic fragments for the -2 at [M - CH3CH2]+ (including both the hydroxyl and carboxyl groups) had shifted from the natural abundance m/z 359 found in the unlabeled peroxide reaction (Fig. 6B) to m/z 365, m/z 363 and m/z 361 (Fig. 6D). The relative abundances of the 18O mono, bi- and tri-labeled hydroxymyristic acid, estimated from the corresponding fragment peak areas, are also included in Fig. 6C (-1 isomer) and Fig. 6D (-2 isomer), together with their formulae and fragment generation (0-2% unlabeled hydroxyacid was also formed). Alkane oxidation studies Four saturated alkanes (dodecane, tetradecane, hexadecane and octadecane) were tested as A. aegerita peroxygenase substrates (Table 1). No reactivity with octadecane was observed under the experimental conditions used. It should be noted that hydroxylation of fatty acids with the same chain length (C18) was observed. In contrast, the reactions with dodecane, gave monohydroxylated derivatives at positions 2 and 3. In addition to the monohydroxylated derivatives, products dihydroxylated at the positions 2 and 3 from both ends of the molecule (i.e. +1 and -1/-2, or +2 and -1/-2) were identified as the predominant compounds. Several combinations of keto and hydroxy derivatives were also formed although in minor amounts (see SD, Fig. S3). Unlike that observed in the reactions with fatty acids and fatty alcohols, terminal hydroxylation was not general in the reactions with alkanes. Similar derivatives to those found for dodecane were obtained from tetradecane and hexadecane although in lower amounts. A. Gutiérrez et al. / Archives of Biochemistry and Biophysics (2011) 514: 33-43. 114  Fig. 6. Mass spectra of -1 (A and C) and -2 (B and D) monohydroxylated myristic acid (C14), from peroxygenase reactions with 1-tetradecanol in the presence of H216O2 (A and B) and H218O2 (90% isotopic purity) (C and D), as trimethylsilyl (TMS) derivatives, and formulae for the mono-, bi- and tri-labeled (top to bottom) compounds with indication of their characteristic ions and relative abundances in the -1 (C) and -2 (D) isomers of hydroxymyristic acid (0-2% unlabeled product was also found). CD AB 370 375 380 385 390 395 400 375 377 379 100% m/z 350 355 360 365 370 375 380 361 363 365 100% m/z 370 375 380 385 390 395 400 373 374 100% m/z 350 355 360 365 370 375 380 359 361 100% m/z 357 16 OTMS 16 OTMS 16 O m/z 359 16 OTMS 16 OTMS m/z 373 16 O 16 OTMS 18 OTMS 16 O m/z 361 7% 18 OTMS 18 OTMS 16 O m/z 363 76% 18 OTMS 18 OTMS 18 O m/z 365 15% 16 OTMS 18 OTMS m/z 375 16 O 9% 18 OTMS 18 OTMS m/z 377 16 O 76% 18 OTMS 18 OTMS m/z 379 18 O 15% 6. Publicaciones  115  Steroid oxidation studies Reactions of both free (cholesterol and sitosterol) and esterified (cholesteryl butyrate, caprylate and linoleate) sterols with the A. aegerita peroxygenase were studied (Table 1). The enzyme showed low reactivity with free sterols, and only a very low conversion rates were attained under the experimental conditions used. The reaction products of cholesterol were monohydroxylated derivatives at positions C-24, C-25, and C-26 (or C-27) (see SD, Fig. S4A) with monohydroxylated C-25 predominating (see SD, Fig. S4B). In the reaction with sitosterol, only the monohydroxylated derivatives at positions C-24 and C- 25 were observed. On the other hand, in the reactions with cholesterol esters no derivatives were observed. Triglyceride oxidation studies In addition of testing the reactivity of free lauric acid and lauric acid methyl ester, the peroxygenase reaction with a glycerol ester of lauric acid (trilaurin) was also studied. Unlike the reaction of free and methylated fatty acid, the triglyceride reaction did not yield any product. Discussion A. aegerita peroxygenase and other heme-thiolate enzymes Heme-containing enzymes using H2O2 as electron acceptor (peroxidases) are a fascinating group of biocatalysts with a variety of ecological and biotechnological implications [9]. The current work deals with one of the most novel hemeperoxidase types, the so-called aromatic peroxygenase, that catalyze remarkable reactions such as peroxide-driven oxygen transfer, together with typical peroxidase reactions, being part of a separate peroxidase superfamily for which the name heme-thiolate peroxidases has been suggested [4,9,16]. A similar heme pocket is shared by P450s and results in similar ability to catalyze monooxygenation reactions, although P450s use O2 as electron acceptor. However, the so-called "peroxide-shunt" exceptionally operates in some of them [17], and an unusual P450 being preferentially activated by H2O2 has been described [18]. The main difference between the P450s and peroxygenase reactions concerns formation of the cofactor reactive Compound I (a porphyrin cation-radical Fe4+=O complex) that in peroxidases results from H2O2 generated A. Gutiérrez et al. / Archives of Biochemistry and Biophysics (2011) 514: 33-43. 122  The first product of fatty alcohol oxidation by the peroxygenase will be a gem-diol from C1 hydroxylation (Fig. 7A), which will be either directly hydroxylated (even at the nascent stage) yielding a gem-triol intermediate, irreversibly dehydrating to release the acid group (Fig. 7B); or first dehydrated pto the corresponding aldehyde and then hydroxylated to the acid group (Fig. 7C). The predominant simple-labeling of the carboxyl group in fatty and hydroxyfatty acids (in the latter case accompanied by a second 18O atom at the hydroxyl group) indicates the existence of such dehydration step, resulting in partial loss of the previously introduced 18O-labeling and formation of both 16O and 18O carboxylic group. The same has been shown in stepwise oxidations of ethanol by P450s [29,44]. The 18O-labeling patterns obtained, which are compatible with both oxidation pathways mentioned above, reveal no extensive hydroxyl exchange with the solvent at the aldehyde/gem-diol stage, which was hardly detectable in the chromatographic analysis (only aldehyde traces found). In the former aspect, the reaction differs from P450 cascade oxidation of hexadecanol, where complete 18O-hydroxyl exchange with the solvent is produced before the final oxidation to palmitic acid, resulting in ~2 ratio between unlabeled and 18O-monolabeled palmitic acid [30]. Hydroxylation of the aldehyde form was the mechanism suggested in the A. aegerita peroxygenase oxidation of benzyl alcohol to benzoic acid, where, in contrast with that observed here, the aldehyde accumulates in substantial amount [45]. However, gem-triol intermediates are suggested in several P450-catalyzed reactions, including oxygenation of some of the aliphatic compounds investigated in the present study [30]. Conclusions The new A. aegerita peroxidase combines P450 and typical peroxidase properties, resulting in activation by peroxide and catalysis of monooxygenasetype reactions (peroxygenase activity). Detailed GC-MS analyses revealed that this enzyme is able to oxidize/oxygenate a variety of aliphatic compounds. Therefore, the name "aromatic peroxygenase" should be substituted by “unspecific peroxygenase” and the corresponding entry (EC 1.11.2.1) has been accepted in the IUBMB Enzyme Nomenclature (www.chem.qmul.ac.uk/iubmb/enzyme). In spite of its classical peroxidative (i.e. non-oxygenase) activity, H218O2 reactions indicated that the enzyme oxidizes 6. Publicaciones  123  aliphatic alcohols to the corresponding acids by enzymatic hydroxylation and dehydration reactions, taking advantage from its monooxygenase activity (also shown in fatty acid 18O hydroxylation). Interestingly, most of the aliphatic oxygenations catalyzed by the A. aegerita peroxygenase (resulting in new hydroxy and keto derivatives) are regioselective taking place at the -1 and -2 positions, irrespective of the aliphatic chain length (myristoleic acid hydroxylation being only produced at the -2 position). This regioselectivity, together with the self-sufficient oxygenase activity, make these new hemethiolate peroxidases interesting biocatalysts for oxidative modification of different aliphatic compounds. Acknowledgements This study was funded by the PEROXICATS (KBBE-2010-4-265397) and BIORENEW (NMP2-CT-2006-026456) EU-projects, and the ELLE (AGL2008-00709) and RAPERO (BIO2008-01533) Spanish MICINN projects co-financed by FEDER funds. J. Rencoret (IRNAS, Seville) is acknowledged for preliminary enzyme assays. E.D. Babot thanks the Spanish CSIC for a JAE fellowship. References [1] R. Ullrich, M. Hofrichter, FEBS Lett. 579 (2005) 6247-6250. [2] R. Ullrich, J. Nuske, K. Scheibner, J. Spantzel, M. Hofrichter, Appl. Environ. Microbiol. 70 (2004) 4575-4581. [3] M.J. Nuell, G.H. Fang, M.J. Axley, P. Kenigsberg, L.P. Hager, J. Bacteriol. 170 (1988) 1007-1011. [4] M. Hofrichter, R. Ullrich, M.J. Pecyna, C. Liers, T. Lundell, Appl. Microbiol. Biotechnol. 87 (2010) 871-897. [5] R. Bernhardt, J. Biotechnol. 124 (2006) 128-145. [6] H. Li, in: A. Messerschmidt, R. Huber, T.L. Poulos, K. Wieghardt (Eds.), Handbook of metalloproteins, Wiley, Baffins Lane, UK, 2001, pp. 267-282. [7] R. Ullrich, M. 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Biology. 4 (1997) 140-146. [41] M.A. Noble, C.S. Miles, S.K. Chapman, D.A. Lysek, A.C. Mackay, G.A. Reid, R.P. Hanzlik, A.W. Munro, Biochem. J. 339 (1999) 371-379. [42] M. Sundaramoorthy, J. Terner, T.L. Poulos, Structure. 3 (1995) 1367-1377. [43] K. Piontek, R. Ullrich, C. Liers, K. Diederichs, D.A. Plattner, M. Hofrichter, Acta Crystallogr. F. 66 (2010) 693-698. [44] F.P. Guengerich, C.D. Sohl, G. Chowdhury, Arch. Biochem. Biophys. 507 (2011) 126-134. [45] M. Kinne, C. Zeisig, R. Ullrich, G. Kayser, K.E. Hammel, M. Hofrichter, Biochem. Biophys. Res. Commun. 397 (2010) 18-21. 6. Publicaciones  127  Fig. S1. Chemical structures of the main oxidation products identified in the peroxygenase reactions with fatty acids, including  to -5 hydroxylated derivatives, -1 and -2 keto derivatives, and dicarboxylic fatty acids. OH O ()n HTP, H2O2 OH O ()n OH OH O ()n OH OH ()n HO O -OH (-1)-OH (-2)-OH OH O ()n OH OH O ()n OH OH O ()n OH (-3)-OH (-4)-OH (-5)-OH O OH O ()nOH O ()n O (-1)-keto (-2)-keto OH O ()n O HO di-COOH A. Gutiérrez et al. / Archives of Biochemistry and Biophysics (2011) 514: 33-43. 128  Fig. S2. Chemical structures of the main oxidation products identified in the peroxygenase reactions with fatty alcohols, including fatty acids (Acd) and their  to -2 hydroxylated, and -1 and -2 keto derivatives. Minor compounds such as -1 and -2 hydroxy and keto derivatives of the fatty alcohols (Alc) are also shown in brackets. nOH () HTP, H 2 O 2 OH O ()n OH OH O ()n OH OH ()n HO O -OH-Acd (-1)-OH-Acd (-2)-OH-Acd O OH O ()nOH O ()n O (-1)-keto-Acd (-2)-keto-Acd OH O ()n Acd nOH ()nOH () nOH ()nOH () OH OH O O (-1)-OH-Alc (-2)-OH-Alc (-1)-keto-Alc (-2)-keto-Alc Alc 6. Publicaciones  129  Fig. S3. Chemical structures of the main oxidation products identified in the peroxygenase reactions with alkanes, including monohydroxylated (positions 2 and 3), and dihydroxylated (positions 2 and -1, 2 and -2, and 3 and -2) fatty alcohols, and ketofatty alcohols ((-1) and (-2)-keto-2-hydroxyfatty alcohol; and (-1) and (-2)-keto-3-hydroxyfatty alcohol). n () HTP, H 2 O 2 OH n ()n () OH (-1)-OH (-2)-OH OH OH n () OH n () OH n () OH OH 2,(-1)-diOH 2,(-2)-diOH 3,(-2)-diOH O OH n () O n () OH n () OH O (-1)-keto-2-OH (-2)-keto-2-OH (-2)-keto-3-OH n () OH (-1)-keto-3-OH O A. Gutiérrez et al. / Archives of Biochemistry and Biophysics (2011) 514: 33-43. 130  Fig. S4. Chemical structures of the main oxidation products identified in the peroxygenase reactions with cholesterol including 24-hydroxy, 25-hydroxy and 26-hydroxycholesterol (A), and GC-MS analysis of these products (B). B 12 13 14 15 16 17 25-OH 26-OH 24-OH HTP, H 2 O 2 HO 2 3 1 4 567 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 HO OH HO OH HO OH 24-OH 25-OH 26-OH A 6. Publicaciones  235  Molecular determinants for selective C25-hydroxylation of vitamins D2 and D3 by fungal peroxygenases Fátima Lucas,‡*a Esteban D. Babot,‡b Marina Cañellas,‡a,c José C. del Río,b Lisbeth Kalum,d René Ullrich,e Martin Hofrichter,e Victor Guallar,a,f Angel T. Martínezg and Ana Gutiérrez*b. a Joint BSC-CRG-IRB Research Program in Computational Biology, Barcelona Supercomputing Center, Jordi Girona 29, E-08034 Barcelona, Spain. E-mail: [email protected] b Instituto de Recursos Naturales y Agrobiología de Sevilla, CSIC, Reina Mercedes 10, E- 41012 Seville, Spain. E-mail: [email protected]; Fax: +32 954624002; Tel: +32 954624711 c Anaxomics Biotech, Balmes 89, E-08008 Barcelona, Spain d Novozymes A/S, Krogshoejvej 36, 2880 Bagsvaerd, Denmark e TU Dresden, Department of Bio- and Environmental Sciences, Markt 23, 02763 Zittau, Germany f ICREA, Passeig Lluís Companys 23, E-08010 Barcelona, Spain g Centro de Investigaciones Biológicas, CSIC, Ramiro de Maeztu 9, E- 28040 Madrid, Spain ‡ These three authors equally contributed to this work * Corresponding author Abstract: Hydroxylation of vitamin D by Agrocybe aegerita and Coprinopsis cinerea peroxygenases was investigated in a combined experimental and computational study. 25-Monohydroxylated vitamin D 3 (cholecalciferol) and D2 (ergocalciferol), compounds of high interest in human health and animal feeding, can be obtained through reaction with both fungal enzymes. Differences in conversion rates and, especially, in site selectivity were nevertheless observed. To rationalize the results, diffusion of D2 and D3 on the molecular structure of the two enzymes was performed with PELE software. In good agreement with experimental conversion yields, simulations indicate more favorable energy profiles for the substrates’ entrance in than for A. aegerita enzyme. On the other hand, GC-MS Lucas F. et al. / Catalysis Science & Technology (2016) 6: 288-295. 236  analyses show that while a full regioselective conversion into the active C25 form is catalyzed by C. cinerea peroxygenase for D2 and D3, A. aegerita yielded a mixture of the hydroxylated D3 products. From the molecular simulations, relative distance distributions between the haem compound I oxygen and H24/H25 atoms (hydrogens on C24 and C25 respectively) were plotted. Results show large populations for O-H25 distances below 3 Å for D2 and D3 in C. cinerea in accordance with the high reactivity observed for this enzyme. In A. aegerita, however, cholecalciferol has similar populations (below 3 Å) for O-H25 and O-H 24 which can justify the hydroxylation observed in C24. In the case of ergocalciferol, due to the bulky methyl group in position C24, very few structures are found with O-H24 distances below 3 Å and thus, as expected, reaction was only observed at C25 position. Introduction Selective oxygenations of aliphatic compounds are among the most challenging reactions in organic chemistry for the regio and/or stereo specific synthesis of pharmaceuticals and fine chemicals. Monooxygenases catalyzing such hydroxylation reactions include cytochromes P450, a family of haem proteins playing a variety of physiological roles but often requiring an auxiliary flavoenzyme (or flavin-containing module) and a source of reducing power to be activated by O2, two facts that limit their biotechnological applicability. Recently, a new peroxidase type, which shares the active-site architecture and reaction mechanism of cytochromes P450, but has the advantage of being activated directly by H2O2, was isolated from Agrocybe aegerita,46 and later identified in a variety of sequenced basidiomycete genomes including that of Coprinopsis cinerea.14 Due to the above characteristics, these unspecific peroxygenases (EC 1.11.2.1) have a huge biotechnological potential as selfsufficient monooxygenases,10,20 for hydroxylation of both aromatic2,3,25-28,43-45 and aliphatic compounds.4,16,34 The A. aegerita enzyme has been the most widely investigated basidiomycete peroxygenase, but recent studies have shown that the C. cinerea enzyme has comparative advantages related to its high conversion yield/selectivity for some 6. Publicaciones  237  hydroxylation reactions, and its production as a recombinant protein in an industrial expression host (by Novozymes, Bagsvaerd, Denmark).5,7 Hydroxylation of vitamin D for the selective production of its active C25- hydroxylated derivatives is one of the reactions where the C. cinerea peroxygenase can be of biotechnological interest (Fig. 1).6 Supplementation with 25-hydroxyvitamin D has a positive effect in different human diseases,12,21,22,29,42 and also raises considerable interest for feeding broiler chickens15,18,24,33 and other farm animals,41 to reduce skeleton problems caused by rapid growth and reduced mobility. Therefore, the use of a peroxygenase in vitamin D hydroxylation represents an attractive alternative to the chemical synthesis. Fig. 1. Enzymatic conversion of cholecalciferol (vitamin D3; 1) and ergocalciferol (vitamin D2; 3) into their bioactive 25-hydroxylated derivatives (2 and 4) by basidiomycete peroxygenases. Lucas F. et al. / Catalysis Science & Technology (2016) 6: 288-295. 238  The goal of the present work was to rationalize the differences observed in cholecalciferol and ergocalciferol (vitamins D3 and D2, respectively) conversion rates and site selectivity by the A. aegerita and C. cinerea peroxygenases. The work presented here consists first of the enzymatic conversion of these compounds by the two peroxygenases, under the same reaction conditions, and gas chromatography-mass spectrometry (GC-MS) analyses to identify all reaction products. Then, energy profiles and binding modes of vitamins D2 and D3 were determined by structure-based computational simulations, using the PELE software.11,13 Finally, differences in site selectivity were investigated through the analysis of the most favorable binding orientations in the active site. Results show that molecular simulations can effectively discriminate experimentally observed differences in conversion rates and site selectivity. Results and discussion Experimental hydroxylation reactions Conversion of cholecalciferol and ergocalciferol was experimentally determined for both the A. aegerita and C. cinerea peroxygenases. In the case of cholecalciferol (0.1 mM), GC-MS analyses of the reaction mixture revealed that this compound was completely (100%) converted by the C. cinerea enzyme within 60 min reaction (Fig. 2C) as compared with the control reaction without peroxygenase (Fig. 2A). In the A. aegerita peroxygenase reaction up to 90% conversion was observed (Fig. 2B). In the case of ergocalciferol (Figs. 2D-F), a conversion of 85% was produced by C. cinerea while in A. aegerita reaction 81% product was observed. When higher substrate concentrations were tested (e.g. 0.5 mM) higher differences in the conversion rates by the two enzymes were observed (40-50% and 20% conversions by the C. cinerea and A. aegerita peroxygenases, respectively). Moreover, cholecalciferol and ergocalciferol conversion by the C. cinerea peroxygenase showed a strict site selectivity since it gave exclusively 25- hydroxycalciferol. Likewise, conversion by the A. aegerita enzyme of ergocalciferol yielded exclusively 25-hydroxyergocalciferol but for cholecalciferol a mixture of products was observed. 6. Publicaciones  239  Fig. 2. GC-MS analyses of cholecalciferol (left) and ergocalciferol (right) hydroxylation by the A. aegerita (B, E) and C. cinerea peroxygenases (C, F), compared with a control without enzyme showing the substrate peaks (A, D), as TMS derivatives from 60-min reactions. In all cases the isopyro (left) and pyro (right) isomers from secosteroid thermal rearrangement are obtained (two small peaks in A and D correspond to minor additional isomers). D D 2 29 31 33 35 37 Retention time (min) D2 100% Relative response F A D 3 29 31 33 35 37 Retention time (min) D3 100% Relative response C 29 31 33 35 37 Retention time (min) 25-OH-D3 25-OH-D3 100% Relative response B 29 31 33 35 37 Retention time (min) 25-OH-D3 25-OH-D3 100% Relative response D3D3 24-OH-D 3 24-OH-D 3 26-OH-D 3 26-OH-D 3 E 29 31 33 35 37 Retention time (min) 25-OH-D2 25-OH-D2 100% Relative response D2D2 29 31 33 35 37 Retention time (min) 25-OH-D2 25-OH-D2 100% Relative response D2D2 Lucas F. et al. / Catalysis Science & Technology (2016) 6: 288-295. 240  The products include 25-hydroxycholecalciferol (64% of the initial substrate) together with 24-hydroxycholecalciferol (21%) and 26/27- hydroxycholecalciferol (7%), as confirmed by comparison with true standards showing identical retention times and mass spectra. The double peaks observed in the chromatograms for both substrate and product (Fig. 2) correspond to the isopyro (19, 9) and pyro (19α, 9α) isomers formed by thermal rearrangement involving ring-B closure, that vitamin D and its hydroxylated derivatives undergo due to the temperature at which GC-MS EI(+) is carried out. Indeed, the presence of the two isomers during GC separation is a useful indication that a secosteroid of the vitamin D type was injected into the system.32 The position of the hydroxyl group at the target C25 position was established by MS of the TMS derivative. The spectrum shows a prominent ion from C24-C25 bond cleavage, with characteristic fragment at m/z 131 and molecular ion at m/z 544. Additionally, characteristic fragments at m/z 529 ([M-15]+), m/z 454 ([M- 90]+), m/z 439 ([M-90-15]+), m/z 349 ([M-90-90-15]+) and m/z 413, were also present. Therefore, both the chromatographic profiles and the mass spectra correspond to the isomerized secosteroids. Peroxygenase structure A superimposition of the general structure and the haem pocket residues in the A. aegerita and C. cinerea peroxygenases is shown in Fig. 3. The main differences are two longer loops (Fig. 3A, arrows) and the substitution of Phe69 by Met69 (Fig. 3B, red label) in the C. cinerea enzyme. All other haem pocket residues, including the proximal cysteine acting as the fifth ligand of the haem iron (Cys36) and the distal glutamic acid and arginine involved in haem activation by H2O2 (Glu169 and Arg189),35 are conserved in the two enzymes. Moreover, differences in the channel providing access to the haem cofactor and the neighbor residues at the channel entrance are shown in Fig. 4. 6. Publicaciones  241  Fig. 3. Comparison of peroxygenase molecular structures. A) General superimposition with the A. aegerita and C. cinerea proteins as pink and yellow ribbons, respectively. The haem group (CPK-colored spheres) and several haem pocket residues (pink- and CPK-colored sticks, respectively) are shown, and two larger loops in the second enzyme are indicated by arrows. B) Haem pocket residues in the A. aegerita (pink sticks) and C. cinerea (CPK-colored sticks) peroxygenases (red label indicates non-conserved methionine in position 69 of the latter enzyme). A hypothetical product molecule identified as 4-hydroxymethylimidazole is shown in A (brown-colored vdW spheres) and B (MZ0, brown-colored sticks).35 From A. aegerita PDB 2YOR, and C. cinerea homology model (provided as Supplemental file 1). Lucas F. et al. / Catalysis Science & Technology (2016) 6: 288-295. 242  Fig. 4. Solvent access surfaces showing differences in the size of the haem (CPK-colored sticks) access channel in the peroxygenases of A. aegerita (A) and C. cinerea (B). Several neighbor residues are shown as CPK-colored vdW spheres, including Gly323 and Ser324 contributing to occlude the haem channel in the C. cinerea enzyme. Computational modeling Ligand diffusion energy profiles For the simulations of cholecalciferol and ergocalciferol access to the H 2 O 2 - activated haem in the A. aegerita and C. cinerea peroxygenases, the haem cofactor was modeled as compound I (a FeIV=O, porphyrin cation radical). The substrate was placed close to the entrance of the haem-access channel of the proteins prepared at the optimal pH for peroxygenase activity (pH 7). This initial location was identified using SiteMap, 40 and from there, the ligand was spawned inside the protein by PELE, 11 following the distance between the reactive O atom in the haem compound I and the cholecalciferol/ergocalciferol C 25 atom (Fig. 1). PELE simulations were done in two stages: first the substrate is perturbed to reduce the C 25 -O distance and, when this distance is below 5 Å, the substrate is free to explore the active site cavity (at the haem distal side) with a 15 Å restrain. In the first step, the ligand is perturbed with a combination of large and small translations and rotations, ranging from 0.5 to 1.5 Å for translations, and 6. Publicaciones   243   0.05 to 0.25 radians for rotations. However, during the second stage, translation range is reduced (0.75-0.25 Å) to perform a finer active site exploration. The plots shown in Fig. 5 correspond to three 48 h simulations each with 80 processors, and show the substrate-C 25 to haem-O distance vs. the interaction energy between the protein and the substrate at each of the different poses explored. Fig. 5. Interaction energies vs. ligand distances from PELE simulations for cholecalciferol (left) and ergocalciferol (right) entrance by the C 25 end in the peroxygenases from A. aegerita (A, B) and C. cinerea (C, D). The distances shown (Å) are between the reactive O atom in the haem compound I and the calciferol C 25 (for substrate numbering see Fig. 1). Simulations show that for the A. aegerita peroxygenase the entrance of the substrates from the surface of the protein is quite favorable but then, the access to the activated haem is obtained against an uphill potential (Figs. 5A,B). For the C. cinerea enzyme the entrance is less open than in the A. aegerita Lucas F. et al. / Catalysis Science & Technology (2016) 6: 288-295. 250  was performed at 300 ºC, the transfer line was kept at 300 ºC, and helium was used as carrier gas. Compounds were identified by mass fragmentography, and by comparing their mass spectra with standards, and quantitation was obtained from total-ion peak area, using molar response factors obtained from cholecalciferol, ergocalciferol, 25-hydroxycholecalciferol and 25- hydroxyergocalciferol standards. The two latter compounds were also used as external standards for calculation of product yields. PELE and other computational analyses The starting structures for PELE simulations were the A. aegerita peroxygenase crystal (2YOR)35 and a homology model for the C. cinerea peroxygenase structure obtained using 2YOR as template.9 As the optimum pH for peroxygenase activity is 7, the structures were prepared accordingly using Schrodinger’s Protein Preparation Wizard36 and H++ web server.1 Histidines were -protonated, with the exception of His82 (-protonated) and His118 and His251 (double protonated). All acidic residues were deprotonated, except Asp85 that was kept in its protonated state. The ergocalciferol and cholecalciferol molecules were optimized with Jaguar38 at the DFT/M06 level with the 6-31G** basis and a PBF implicit solvent in order to obtain their electrostatic potential atomic charges. Finally, the haem site was modeled as thiolate-ligated compound I after being fully optimized in the protein environment with quantum mechanics/molecular mechanics (QM/MM) using QSite.39 The electronic calculations show three unpaired electrons: two located on the oxoiron group and a third on the heme and less than 1% spin contamination. Once the initial protein structure was prepared and ligands optimized, these were placed manually in identical positions at the entrance of the haemaccess channel and PELE simulations were performed.11 PELE is a Monte Carlo based algorithm that produces new configurations through a sequential ligand and protein perturbation, side chain prediction and minimization steps, freely available at https://pele.bsc.es. New configurations are then filtered with a Metropolis acceptance test, where the energy is described with an all-atom OPLS force field23 and a surface generalized Born solvent 8. In this way it is possible to locate and characterize local and global minima structures for the 6. Publicaciones  251  most favorable protein-ligand interactions. PELE has been successfully used in a number of ligand migration studies with both small and large substrates 19,30,31. Distance and angle distributions were computed after screening all structures from PELE simulations by interaction energy and distance to the compound I oxygen. Structures that showed interaction energies below -20 kcal/mol for ergocalciferol and -30 kcal/mol for cholecalciferol were selected. Reactive hydrogen atoms where considered below 5 Å to the compound I oxygen atom for the distance analysis and 3Å for the angle study (angles are much more sensitive and thus only near attack conformations were selected). All the selected structures where then used to compute the relative frequencies of the ligand’s hydrogen atoms at reactive distances to the haem oxygen and Fe=O- H and O-H-C angles. Acknowledgements This work was supported by the INDOX (KBBE-2013-7-613549) and PELE (ERC-2009-Adg 25027) EU projects, and by the BIO2011-26694 and CTQ2013-48287 projects of the Spanish Ministry of Economy and Competitiveness. Electronic Supplementary Information (ESI) available: Molecular model for C. cinerea peroxygenase obtained using the A. aegerita crystal structure as template. See DOI: 10.1039/b000000x/. 1. R. Anandakrishnan, B. Aguilar and A. V. Onufriev, Nucleic Acids Res., 2012, 40, W537- W541. 2. E. Aranda, M. Kinne, M. Kluge, R. Ullrich and M. Hofrichter, Appl. Microbiol. Biotechnol., 2009, 82, 1057-1066. 3. E. Aranda, R. Ullrich and M. Hofrichter, Biodegradation, 2010, 21, 267-281. 4. E. D. Babot, J. C. del Río, M. Cañellas, F. Sancho, F. Lucas, V. Guallar, L. Kalum, H. Lund, G. Gröbe, K. Scheibner, R. Ullrich, M. Hofrichter, A. T. Martínez and A. Gutiérrez, Appl. Environ. Microbiol., 2015, 81, 4130-4142. 5. E. D. Babot, J. C. del Río, L. Kalum, A. T. Martínez and A. Gutiérrez, Biotechnol. Bioeng., 2013, 110, 2332. Lucas F. et al. / Catalysis Science & Technology (2016) 6: 288-295. 252  6. E. D. Babot, J. C. del Río, L. Kalum, A. T. Martínez and A. Gutiérrez, Chemcatchem, 2014,(on line). 7. E. D. Babot, J. C. del Río, L. Kalum, A. T. Martínez and A. Gutiérrez, Chemcatchem, 2014,(on line). 8. D. Bashford and D. A. Case, Annual Review of Physical Chemistry, 2000, 51, 129-152. 9. L. Bordoli, F. Kiefer, K. Arnold, P. Benkert, J. Battey and T. Schwede, Nat. Protoc., 2009, 4, 1-13. 10. S. Bormann, A. G. Baraibar, Y. Ni, D. Holtmann and F. Hollmann, Catal. Sci. Technol., 2015, 5, 2038-2052. 11. K. W. Borrelli, A. Vitalis, R. Alcantara and V. Guallar, J. Chem. Theory Comput., 2005, 1, 1304-1311. 12. N. R. Buck, W. Claerhout, B. H. Leuenberger, E. Stoecklin, K. Urban and S. Wolfram, Patent (USA), 2013,US 20130210782 A1. 13. B. P. Cossins, A. Hosseini and V. Guallar, J. Chem. Theory Comput., 2012, 8, 959-965. 14. D. Floudas, M. Binder, R. Riley, K. Barry, R. A. Blanchette, B. Henrissat, A. T. Martínez, R. Otillar, J. W. Spatafora, J. S. Yadav, A. Aerts, I. Benoit, A. Boyd, A. Carlson, A. Copeland, P. M. Coutinho, R. P. de Vries, P. Ferreira, K. Findley, B. Foster, J. Gaskell, D. Glotzer, P. Górecki, J. Heitman, C. Hesse, C. Hori, K. Igarashi, J. A. Jurgens, N. Kallen, P. Kersten, A. Kohler, U. Kües, T. K. A. Kumar, A. Kuo, K. LaButti, L. F. Larrondo, E. Lindquist, A. Ling, V. Lombard, S. Lucas, T. Lundell, R. Martin, D. J. McLaughlin, I. Morgenstern, E. Morin, C. Murat, M. Nolan, R. A. Ohm, A. Patyshakuliyeva, A. Rokas, F. J. Ruiz-Dueñas, G. Sabat, A. Salamov, M. Samejima, J. Schmutz, J. C. Slot, F. St.John, J. Stenlid, H. Sun, S. Sun, K. Syed, A. Tsang, A. Wiebenga, D. Young, A. Pisabarro, D. C. Eastwood, F. Martin, D. Cullen, I. V. Grigoriev and D. S. Hibbett, Science, 2012, 336, 1715-1719. 15. C. A. Fritts and P. W. Waldroup, Journal of Applied Poultry Research, 2003, 12, 45-52. 16. A. Gutiérrez, E. D. Babot, R. Ullrich, M. Hofrichter, A. T. Martínez and J. C. del Río, Arch. Biochem. Biophys., 2011, 514, 33-43. 17. A. Gutiérrez, J. C. del Río, F. J. González-Vila and F. Martín, J. Chromatogr., 1998, 823, 449-455. 18. J. M. Hernández, Patent (USA), 2013,US 20130137662 A1. 19. A. Hernández-Ortega, F. Lucas, P. Ferreira, M. Medina, V. Guallar and A. T. Martínez, J. Biol. Chem., 2011, 286, 41105-41114. 20. M. Hofrichter and R. Ullrich, Curr. Opin. Chem. Biol., 2014, 19, 116-125. 21. G. Jean, J. C. Terrat, T. Vanel, J. M. Hurot, C. Lorriaux, B. Mayor and C. Chazot, Nephrol. Dial. Transplant., 2008, 23, 3670-3676. 6. Publicaciones  253  22. G. Jones, Annu. Rev. Nutr., 2013, 33, 23-44. 23. G. A. Kaminski, R. A. Friesner, J. Tirado-Rives and W. L. Jorgensen, J. Phys. Chem. B, 2001, 105, 6474-6487. 24. S. Käppeli, E. Frohlich, S. G. Gebhardt-Henrich, A. Pfulg, H. Schäublin, R. Zweifel, H. Wiedmer and H. H. Stoffel, Arch. Geflügelk., 2011, 75, 179-184. 25. A. Karich, M. Kluge, R. Ullrich and M. Hofrichter, AMB Express, 2013, 3:5, 26. M. Kinne, C. Zeisig, R. Ullrich, G. Kayser, K. E. Hammel and M. Hofrichter, Biochem. Biophys. Res. Commun., 2010, 397, 18-21. 27. M. Kluge, R. Ullrich, C. Dolge, K. Scheibner and M. Hofrichter, Appl. Microbiol. Biotechnol., 2009, 81, 1071-1076. 28. M. Kluge, R. Ullrich, K. Scheibner and M. Hofrichter, Green Chem., 2012, 14, 440-446. 29. G. A. Leichtmann, J. M. Bengoa, M. J. G. Bolt and M. D. Sitrin, Amer. J. Clin. Nutr., 1991, 54, 548-552. 30. D. Linde, R. Pogni, M. Cañellas, F. Lucas, V. Guallar, M. C. Baratto, A. Sinicropi, V. Sáez-Jiménez, C. Coscolín, A. Romero, F. J. Medrano, F. J. Ruiz-Dueñas and A. T. Martínez, Biochem. J., 2014, on-line doi:10.1042/BJ20141211, 31. M. F. Lucas and V. Guallar, Biophys. J., 2012, 102, 887-896. 32. Makin, H. L. J. and D. B. Gower. Steroid analysis. Springer, NY, 2010. 33. M. Michalczuk, D. Pietrzak, J. Niemiec and J. Mroczk, Pol. J. Food Nutr. Sci., 2010, 60, 121-126. 34. S. Peter, M. Kinne, X. Wang, R. Ulrich, G. Kayser, J. T. Groves and M. Hofrichter, FEBS J., 2011, 278, 3667-3675. 35. K. Piontek, E. Strittmatter, R. Ullrich, G. Grobe, M. J. Pecyna, M. Kluge, K. Scheibner, M. Hofrichter and D. A. Plattner, J. Biol. Chem., 2013, 288, 34767-34776. 36. G. M. Sastry, M. Adzhigirey, T. Day, R. Annabhimoju and W. Sherman, Journal of Computer-Aided Molecular Design, 2013, 27, 221-234. 37. J. C. Schöneboom, S. Cohen, H. Lin, S. Shaik and W. Thiel, J. Am. Chem. Soc., 2004, 126, 4017-4034. 38. Schrödinger. Jaguar 7.8. LCC, New York, 2011. 39. Schrödinger. QSite 5.7. LCC, New York, 2011. 40. Schrödinger. SiteMap 2.5. LCC, New York, 2011. 41. C. Simoes-Nunes and G. M. Weber, Patent (European), 2004,EP 1516540 B1. 42. M. D. Sitrin and J. M. Bengoa, Amer. J. Clin. Nutr., 1987, 46, 1011-1015. 43. R. Ullrich, C. Dolge, M. Kluge and M. Hofrichter, FEBS Lett., 2008, 582, 4100-4106. Lucas F. et al. / Catalysis Science & Technology (2016) 6: 288-295. 254  44. R. Ullrich and M. Hofrichter, FEBS Lett., 2005, 579, 6247-6250. 45. R. Ullrich and M. Hofrichter, Cell. Mol. Life Sci., 2007, 64, 271-293. 46. R. Ullrich, J. Nuske, K. Scheibner, J. Spantzel and M. Hofrichter, Appl. Environ. Microbiol., 2004, 70, 4575-4581. Supporting Information The Supporting Information shows the main interactions of cholecalciferol and ergocalciferol at the peroxygenase heme access channel (Fig. S1 and S2, respectively), the effect of side-chain structure on the conversion rates of five sterols by the A. aegerita and C. cinerea peroxygenases (Table S1), and distributions of estimated Fe=O-H and O-H-C angles in the peroxygenase reactions with cholecalciferol and ergocalciferol (Fig. S3). Fig. S1. Main interactions (below 3 Å) for cholecalciferol with the peroxygenases of A. aegerita (A) and C. cinerea (B). 6. Publicaciones   255   Fig. S2. Main interactions (below 3 Å) for ergocalciferol with the peroxygenases of A. aegerita (A) and C. cinerea (B). Lucas F. et al. / Catalysis Science & Technology (2016) 6: 288-295. 256  Table S1. Conversion degree of steroids with different side-chains (A-E: cholesterol, campesterol, ergosterol, sitosterol and stigmasterol, respectively) by A. aegerita and C. cinerea peroxygenasesa. Conversion (%) A. aegerita peroxygenase C. cinerea peroxygenase A 64 100 B 47 30 C 10 6 D 13 6 E 2 2 aFrom E. D. Babot, J. C. del Río, M. Cañellas, F. Sancho, F. Lucas, V. Guallar, L. Kalum, H. Lund, G. Gröbe, K. Scheibner, R. Ullrich, M. Hofrichter, A. T. Martínez, and A. Gutiérrez. 2015. Steroid hydroxylation by basidiomycete peroxygenases: A combined experimental and computational study. Appl. Environ. Microbiol. 81:4130-4142 C 19 H 30 O 22 23 24 21 20 25 26 27 C 19 H 30 O 22 23 24 21 20 25 26 27 28 C 19 H 28 O 22 23 24 21 20 25 26 27 28 C19H30O 22 23 24 21 20 25 26 27 28 29 C 19 H 30 O 22 23 24 21 20 25 26 27 28 29 6. Publicaciones   257   Fig. S3. Relative distributions of computed Fe=O-H and O-H-C angles for the reactions of A. aegerita (A, C) and C. cinerea (B, D) peroxygenases at the C 24 , C 25 , C 26 and C 27 , positions of cholecalciferol (blue) and ergocalciferol (green) and the C 28 position of ergocalciferol. Structures were filtered by energy and distance between the pertinent hydrogen atom and the haem compound I oxygen (below 3 Å). QM/MM studies for P450 camb show that in a prearranged reactive position camphor adopts angles of 130º for Fe=O-H and 170º for O-H-C. While for that latter no correlation is found to the theoretical predictions, in the case of Fe=O-H 24 angles between 120 and 140º are favoured in C 24 for cholecalciferol reacting with Lucas F. et al. / Catalysis Science & Technology (2016) 6: 288-295. 258  A. aegerita (A) in agreement with the experimentally observed formation of 21% product in this position. In the case of the remaining 3 reactions, few structures are found with angles below 145º. In the case of Fe=O-H25 (as expected) all compounds present favourable angles. Analogous to the case of the distance distribution analysis, no conclusions can be made for the reactivity of C26, C27 and C28. bJ. C. Schöneboom, S. Cohen, H. Lin, S. Shaik and W. Thiel, J. Am. Chem. Soc., 2004, 126, 4017-4034.  Lund H. et al. / WO 2013/004639. PCT/EP2012/062763 266 Fig 1. Chromatographic profile of tetradecane incubated with C. cinerea peroxigenase (0.5 mM H2O2). Fig 2. Chromatographic profile of tetradecane incubated with A. aegerita peroxigenase (2.5 mM H2O2). 5101520 minutes C14 -1 OH  - 2 OH (-1, -1) di OH (-1, -2) di OH (-2, -2) di OH 5101520 minutes -1 OH -2 OH C14 (-1, -1) di OH (-2, -2) di OH (-1, -2) di OH 6. Publicaciones  267 Fig 3. Chromatographic profile of tetradecanol incubated with C. cinerea peroxigenase (0.5 mM H 2 O 2 ). Fig 4. Chromatographic profile of tetradecanol incubated with A. aegerita peroxigenase (2.5 mM H 2 O 2 ). Lund H. et al. / WO 2013/004639. PCT/EP2012/062763 268 DEFINITIONS Peroxygenase activity: The term "peroxygenase activity" is defined herein as "unspecific peroxygenase" according to EC 1.11.2.1. This is a heme-thiolate protein. Enzymes of this type include glycoproteins secreted by agaric basidiomycetes. They catalyse the insertion of an oxygen atom from H2O2 into a wide variety of substrates, such as naphthalene, 4-nitrobenzodioxole; and alkanes such as propane, hexane and cyclohexane. They have little or no activity toward chloride. For purposes of the present invention, peroxygenase activity is determined according to the spectrophotometric procedure described by Kluge et al. (2007, Appl Microbiol Biotechnol 75:1473-1478). Isolated polypeptide: The term "isolated polypeptide" as used herein refers to a polypeptide that is isolated from a source. In a preferred aspect, the polypeptide is at least 1% pure, preferably at least 5% pure, more preferably at least 10% pure, more preferably at least 20% pure, more preferably at least 40% pure, more preferably at least 60% pure, even more preferably at least 80% pure, and most preferably at least 90% pure, as determined by SDS PAGE. Substantially pure polypeptide: The term "substantially pure polypeptide" denotes herein a polypeptide preparation that contains at most 10%, preferably at most 8%, more preferably at most 6%, more preferably at most 5%, more preferably at most 4%, more preferably at most 3%, even more preferably at most 2%, most preferably at most 1%, and even most preferably at most 0.5% by weight of other polypeptide material with which it is natively or recombinantly associated. It is, therefore, preferred that the substantially pure polypeptide is at least 92% pure, preferably at least 94% pure, more preferably at least 95% pure, more preferably at least 96% pure, more preferably at least 96% pure, more preferably at least 97% pure, more preferably at least 98% pure, even more preferably at least 99%, most preferably at least 99.5% pure, and even most preferably 100% pure by weight of the total polypeptide material present in the preparation. The polypeptides of the present invention are preferably in a substantially pure form, i.e., that the polypeptide preparation is essentially free of other polypeptide material with which it is natively or recombinantly associated. 6. Publicaciones 269 This can be accomplished, for example, by preparing the polypeptide by wellknown recombinant methods or by classical purification methods. Mature polypeptide: The term "mature polypeptide" is defined herein as a polypeptide having peroxygenase activity that is in its final form following translation and any posttranslational modifications, such as N-terminal processing, C-terminal truncation, glycosylation, phosphorylation, etc. In a preferred aspect, the mature polypeptide has the amino acid sequence shown in positions 1 to 330 of SEQ ID NO: 1 based on the N-terminal peptide sequencing data (Ullrich et al., 2004, Appl. Env. Microbiol. 70(8): 4575-4581), elucidating the start of the mature protein of AaP peroxygenase enzyme. In another preferred aspect, the mature polypeptide has the amino acid sequence shown in positions 1 to 328 of SEQ ID NO: 2. Identity: The relatedness between two amino acid sequences or between two nucleotide sequences is described by the parameter "identity". For purposes of the present invention, the degree of identity between two amino acid sequences is determined using the Needleman-Wunsch algorithm (Needleman and Wunsch, 1970, J. Mol. Biol. 48: 443-453) as implemented in the Needle program of the EMBOSS package (EMBOSS: The European Molecular Biology Open Software Suite, Rice et al., 2000, Trends in Genetics 16: 276-277; http://emboss.org), preferably version 3.0.0 or later. The optional parameters used are gap open penalty of 10, gap extension penalty of 0.5, and the EBLOSUM62 (EMBOSS version of BLOSUM62) substitution matrix. The output of Needle labeled "longest identity" (obtained using the -nobrief option) is used as the percent identity and is calculated as follows: (Identical Residues x 100)/(Length of Alignment - Total Number of Gaps in Alignment). For purposes of the present invention, the degree of identity between two deoxyribonucleotide sequences is determined using the Needleman-Wunsch algorithm (Needleman and Wunsch, 1970, supra) as implemented in the Needle Lund H. et al. / WO 2013/004639. PCT/EP2012/062763 270 program of the EMBOSS package (EMBOSS: The European Molecular Biology Open Software Suite, Rice et al., 2000, supra; http://emboss.org), preferably version 3.0.0 or later. The optional parameters used are gap open penalty of 10, gap extension penalty of 0.5, and the EDNAFULL (EMBOSS version of NCBI NUC4.4) substitution matrix. The output of Needle labeled "longest identity" (obtained using the -nobrief option) is used as the percent identity and is calculated as follows: (Identical Deoxyribonucleotides x 100)/(Length of Alignment - Total Number of Gaps in Alignment). Modification: The term "modification" means herein any chemical modification of the polypeptide consisting of the mature polypeptide of SEQ ID NO: 1, 2, 3, 4, 5, 6, 7, 8, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28 or 29; or a homologous sequence thereof; as well as genetic manipulation of the DNA encoding such a polypeptide. The modification can be a substitution, a deletion and/or an insertion of one or more (several) amino acids as well as replacements of one or more (several) amino acid side chains. DETAILED DESCRIPTION OF THE INVENTION Polypeptides having peroxygenase activity (peroxygenases) The present invention relates to uses of an isolated polypeptide, which is preferably recombinantly produced, having peroxygenase activity, which comprises an amino acid sequence having at least 30% identity, preferably at least 35%, 40%, 45%, 50%, 55%, 60%, 65%, 70%, 75%, 80%, 85%, 90%, 95%, 97%, or 98% identity to the polypeptide of SEQ ID NO: 1, 2, 3, 4, 5, 6, 7, 8, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28 or 29; preferably SEQ ID NO: 2 or SEQ ID NO: 4. In a preferred embodiment, the polypeptide comprises an amino acid sequence represented by one or more of the following motifs, preferably comprising two or more, three or more, four or more, five or six of the following motifs: 6. Publicaciones 271 In a more preferred embodiment, the peroxygenase comprises an amino acid sequence represented by the motif: E[HG]DXSX[ST]RXD. In another embodiment, the polypeptide comprises an amino acid sequence having a substitution, deletion, and/or insertion of one or several amino acids of the mature polypeptide of SEQ ID NO: 1, 2, 3, 4, 5, 6, 7, 8, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28 or 29; preferably SEQ ID NO: 2 or SEQ ID NO: 4. In yet another embodiment, the polypeptide of the first aspect comprises or consists of the amino acid sequence of SEQ ID NO: 1, 2, 3, 4, 5, 6, 7, 8, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28 or 29; preferably SEQ ID NO: 2 or SEQ ID NO: 4; or a fragment thereof having peroxygenase activity; preferably the polypeptide comprises or consists of the mature polypeptide of SEQ ID NO: 1, 2, 3, 4, 5, 6, 7, 8, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28 or 29; preferably SEQ ID NO: 2 or SEQ ID NO: 4. Preferably, amino acid changes are of a minor nature, that is conservative amino acid substitutions or insertions that do not significantly affect the folding and/or activity of the protein; small deletions, typically of one to about 30 amino acids; small amino- or carboxyl-terminal extensions, such as an amino-terminal methionine residue; a small linker peptide of up to about 20-25 residues; or a small extension that facilitates purification by changing net charge or another function, such as a poly-histidine tract, an antigenic epitope or a binding domain. Motif I: [FL]XX[YF]S[AN]X[FHY]G[GN]GX[YF]N (SEQ ID NO: 9); Motif II: G[GN]GX[YF]NXX[VA]AX[EH][LF]R (SEQ ID NO: 10); Motif III: RXXRI[QE][DEQ]S[IM]ATN (SEQ ID NO: 11); Motif IV: S[IM]ATN[PG][EQN][FM][SDN][FL] (SEQ ID NO: 12); Motif V: P[PDK][DG]F[HFW]R[AP] (SEQ ID NO: 13); Motif VI: [TI]XXXLYPNP[TK][GV] (SEQ ID NO: 14); Motif VII: E[HG]DXSX[ST]RXD (SEQ ID NO: 15). Lund H. et al. / WO 2013/004639. PCT/EP2012/062763 272 Examples of conservative substitutions are within the group of basic amino acids (arginine, lysine and histidine), acidic amino acids (glutamic acid and aspartic acid), polar amino acids (glutamine and asparagine), hydrophobic amino acids (leucine, isoleucine and valine), aromatic amino acids (phenylalanine, tryptophan and tyrosine), and small amino acids (glycine, alanine, serine, threonine and methionine). Amino acid substitutions that do not generally alter specific activity are known in the art and are described, for example, by H. Neurath and R.L. Hill, 1979, In, The Proteins, Academic Press, New York. The most commonly occurring exchanges are Ala/Ser, Val/Ile, Asp/Glu, Thr/Ser, Ala/Gly, Ala/Thr, Ser/Asn, Ala/Val, Ser/Gly, Tyr/Phe, Ala/Pro, Lys/Arg, Asp/Asn, Leu/Ile, Leu/Val, Ala/Glu, and Asp/Gly. In addition to the 20 standard amino acids, non-standard amino acids (such as 4-hydroxyproline, 6-N-methyl lysine, 2-aminoisobutyric acid, isovaline, and alpha-methyl serine) may be substituted for amino acid residues of a wild-type polypeptide. A limited number of non-conservative amino acids, amino acids that are not encoded by the genetic code, and unnatural amino acids may be substituted for amino acid residues. "Unnatural amino acids" have been modified after protein synthesis, and/or have a chemical structure in their side chain(s) different from that of the standard amino acids. Unnatural amino acids can be chemically synthesized, and preferably, are commercially available, and include pipecolic acid, thiazolidine carboxylic acid, dehydroproline, 3- and 4- methylproline, and 3,3-dimethylproline. Alternatively, the amino acid changes are of such a nature that the physicochemical properties of the polypeptides are altered. For example, amino acid changes may improve the thermal stability of the polypeptide, alter the substrate specificity, change the pH optimum, and the like. Essential amino acids in the parent polypeptide can be identified according to procedures known in the art, such as site-directed mutagenesis or alaninescanning mutagenesis (Cunningham and Wells, 1989, Science 244: 1081-1085). In the latter technique, single alanine mutations are introduced at every residue in the molecule, and the resultant mutant molecules are tested for biological activity (i.e., peroxygenase activity) to identify amino acid residues that are critical to the activity of the molecule. See also, Hilton et al., 1996, J. Biol. Chem. 271:4699-4708. The active site of the enzyme or other biological interaction can also be determined by physical analysis of structure, as determined by such 6. Publicaciones 273 techniques as nuclear magnetic resonance, crystallography, electron diffraction, or photoaffinity labeling, in conjunction with mutation of putative contact site amino acids. See, for example, de Vos et al., 1992, Science 255: 306-312; Smith et al., 1992, J. Mol. Biol. 224: 899-904; Wlodaver et al., 1992, FEBS Lett. 309: 59-64. The identities of essential amino acids can also be inferred from analysis of identities with polypeptides that are related to a polypeptide according to the invention. Single or multiple amino acid substitutions, deletions, and/or insertions can be made and tested using known methods of mutagenesis, recombination, and/or shuffling, followed by a relevant screening procedure, such as those disclosed by Reidhaar-Oison and Sauer, 1988, Science 241: 53-57; Bowie and Sauer, 1989, Proc. Natl. Acad. Sci. USA 86: 2152-2156; WO 95/17413; or WO 95/22625. Other methods that can be used include error-prone PCR, phage display (e.g., Lowman et al., 1991, Biochem. 30: 10832-10837; U.S. Patent No. 5,223,409; WO 92/06204), and region-directed mutagenesis (Derbyshire et al., 1986, Gene 46: 145; Ner et al., 1988, DNA 7: 127). Mutagenesis/shuffling methods can be combined with high-throughput, automated screening methods to detect activity of cloned, mutagenized polypeptides expressed by host cells (Ness et al., 1999, Nature Biotechnology 17: 893-896). Mutagenized DNA molecules that encode active polypeptides can be recovered from the host cells and rapidly sequenced using standard methods in the art. These methods allow the rapid determination of the importance of individual amino acid residues in a polypeptide of interest, and can be applied to polypeptides of unknown structure. The total number of amino acid substitutions, deletions and/or insertions of the mature polypeptide of SEQ ID NO: 1, 2, 3, 4, 5, 6, 7, 8, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28 or 29; preferably SEQ ID NO: 2 or SEQ ID NO: 4; is 10, preferably 9, more preferably 8, more preferably 7, more preferably at most 6, more preferably 5, more preferably 4, even more preferably 3, most preferably 2, and even most preferably 1. Another preferred embodiment relates to the polypeptide having peroxygenase activity of the first aspect of the invention, wherein the mature polypeptide is amino acids 1 to 330 of SEQ ID NO: 1. Lund H. et al. / WO 2013/004639. PCT/EP2012/062763 274 Yet another preferred embodiment relates to the polypeptide having peroxygenase activity of the first aspect of the invention, wherein the mature polypeptide is amino acids 1 to 328 of SEQ ID NO: 2. Yet another preferred embodiment relates to the polypeptide having peroxygenase activity of the first aspect of the invention, wherein the mature polypeptide is amino acids 1 to 344 of SEQ ID NO: 4. Yet another preferred embodiment relates to the polypeptide having peroxygenase activity of the first aspect of the invention, wherein the mature polypeptide is amino acids 1 to 261 of SEQ ID NO: 23. Hydrogen peroxide The hydrogen peroxide required by the peroxygenase may be provided as an aqueous solution of hydrogen peroxide or a hydrogen peroxide precursor for in situ production of hydrogen peroxide. Any solid entity which liberates upon dissolution a peroxide which is useable by peroxygenase can serve as a source of hydrogen peroxide. Compounds which yield hydrogen peroxide upon dissolution in water or an appropriate aqueous based medium include but are not limited to metal peroxides, percarbonates, persulphates, perphosphates, peroxyacids, alkyperoxides, acylperoxides, peroxyesters, urea peroxide, perborates and peroxycarboxylic acids or salts thereof. Another source of hydrogen peroxide is a hydrogen peroxide generating enzyme system, such as an oxidase together with a substrate for the oxidase. Examples of combinations of oxidase and substrate comprise, but are not limited to, amino acid oxidase (see e.g., US 6,248,575) and a suitable amino acid, glucose oxidase (see e.g., WO 95/29996) and glucose, lactate oxidase and lactate, galactose oxidase (see e.g., WO 00/50606) and galactose, and aldose oxidase (see e.g., WO 99/31990) and a suitable aldose. By studying EC 1.1.3._, EC 1.2.3._, EC 1.4.3._, and EC 1.5.3._ or similar classes (under the International Union of Biochemistry), other examples of such combinations of oxidases and substrates are easily recognized by one skilled in the art. Hydrogen peroxide or a source of hydrogen peroxide may be added at the beginning of or during the method of the invention, e.g., as one or more separate additions of hydrogen peroxide; or continously as fed-batch addition. Typical 6. Publicaciones 275 amounts of hydrogen peroxide correspond to levels of from 0.001 mM to 25 mM, preferably to levels of from 0.005 mM to 5 mM, and particularly to levels of from 0.01 to 1 mM hydrogen peroxide. Hydrogen peroxide may also be used in an amount corresponding to levels of from 0.1 mM to 25 mM, preferably to levels of from 0.5 mM to 15 mM, more preferably to levels of from 1 mM to 10 mM, and most preferably to levels of from 2 mM to 8 mM hydrogen peroxide. Surfactants The method of the invention may include application of a surfactant (for example, as part of a detergent formulation or as a wetting agent). Surfactants suitable for being applied may be non-ionic (including semi-polar), anionic, cationic and/or zwitterionic; preferably the surfactant is anionic (such as linear alkylbenzenesulfonate, alpha-olefinsulfonate, alkyl sulfate (fatty alcohol sulfate), alcohol ethoxysulfate, secondary alkanesulfonate, alpha-sulfo fatty acid methyl ester, alkyl- or alkenylsuccinic acid or soap or non-ionic such as alcohol ethoxylate, nonylphenol ethoxylate, alkylpolyglycoside, alkyldimethylamineoxide, ethoxylated fatty acid monoethanolamide, fatty acid monoethanolamide, polyhydroxy alkyl fatty acid amide, or N -acyl N-alkyl derivatives of glucosamine ("glucamides"), or a mixture thereof. When included in the method of the invention, the concentration of the surfactant will usually be from about 0.01% to about 10%, preferably about 0.05% to about 5%, and more preferably about 0.1% to about 1% by weight. Aliphatic Hydrocarbons The hydrocarbons, which are oxidized in the method of the invention, are aliphatic hydrocarbons having a chain of at least five carbons. Preferably, the aliphatic hydrocarbon is an alkane or an alkene; more preferably, the aliphatic hydrocarbon is an alkane, such as pentane, hexane, heptane, octane, nonane, decane, undecane, dodecane, tridecane, tetradecane, pentadecane or hexadecane, or isomers thereof. Even more preferably, the aliphatic hydrocarbon is undecane, dodecane, tridecane, tetradecane, pentadecane or hexadecane, or isomers thereof. In an embodiment, the aliphatic hydrocarbon is not n-hexane or n - decane. Lund H. et al. / WO 2013/004639. PCT/EP2012/062763 282 hydrocarbon having at least five carbons to the corresponding acid, comprising contacting the alcohol of an aliphatic hydrocarbon with hydrogen peroxide and a polypeptide having peroxygenase activity; wherein the polypeptide comprises: i) an amino acid sequence which has at least 30% identity to SEQ ID NO: 1, 2, 3, 4, 5, 6, 7, 8, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28 or 29; preferably SEQ ID NO: 2 or SEQ ID NO: 4; and j) an amino acid sequence represented by one or more of the following motifs: For example, pentanol may be changed (oxidized) to pentanoic acid (valeric acid) , hexanol may be changed to hexanoic acid (caproic acid), heptanol may be changed to heptanoic acid (enanthic acid), octanol may be changed to octanoic acid (caprylic acid) , nonanol may be changed to nonanoic acid (pelargonic acid), decanol may be changed to decanoic acid (capric acid), dodecanol may be changed to dodecanoic acid (lauric acid) , tetradecanol may be changed to tetradecanoic acid (myristic acid), hexadecanol may be changed to hexadecanoic acid (palmitic acid), octadecanol may be changed to octadecanoic acid (stearic acid), and eicosanol may be changed to eicosanoic acid (arachidic acid) . Motif I: [FL]XX[YF]S[AN]X[FHY]G[GN]GX[YF]N (SEQ ID NO: 9); Motif II: G[GN]GX[YF]NXX[VA]AX[EH][LF]R (SEQ ID NO: 10); Motif III: RXXRI[QE][DEQ]S[IM]ATN (SEQ ID NO: 11); Motif IV: S[IM]ATN[PG][EQN][FM][SDN][FL] (SEQ ID NO: 12); Motif V: P[PDK][DG]F[HFW]R[AP] (SEQ ID NO: 13); Motif VI: [TI]XXXLYPNP[TK][GV] (SEQ ID NO: 14); Motif VII: E[HG]DXSX[ST]RXD (SEQ ID NO: 15); preferably, Motif VII: E[HG]DXSX[ST]RXD. 6. Publicaciones 283 The present invention is further described by the following examples that should not be construed as limiting the scope of the invention. EXAMPLES The amino acid sequence of the peroxygenase from Agrocybe aegerita is shown as SEQ ID NO: 2 ; and the amino acid sequence of the peroxygenase from Coprinopsis cinerea is shown as SEQ ID NO: 4. EXAMPLE 1 Enzymatic oxidation of dodecane, tetradecane and hexadecane The extracellular peroxygenase of A. aegerita (isoform II , 44 kDa, SEQU ID NO: 2) was used. The enzyme preparation was homogeneous by sodium dodecylsulfate-polyacrylamide gel electrophoresis, an exhibited and A418/A280 ratio of 1.75. Its specific activity was 117 units·mg-1 , where 1 unit represents the oxidation of 1 μmol of veratryl alcohol to veratraldehyde (310 9300 M-1· cm-1) in 1 minute at 23 ºC and pH 7, in the presence of 2.5 mM H2O2. Three alkanes: dodecane (C12) , tetradecane (C14) and hexadecane (C16) were obtained from Sigma-Aldrich. Five mL reaction s of the above model substrates (1 mM) with the A. aegerita peroxygenase (1 U) were performed in 50 mM sodium phosphate buffer (pH 7) at 25 ºC for 2 h, in the presence of 2.5 mM H2O2 . The substrates were previously dissolved in acetone and added to the buffer (the acetone concentration in the reaction was 15%). In control experiments, substrates were treated under the same conditions but without enzyme. After the enzymatic reactions, water was immediately removed in a rotary evaporator, and the products recovered with chloroform, dried under nitrogen, and redissolved in chloroform for GC-MS analyses. Bis(trimethylsilyl)-trifluoroacetamide (Supelco) in the presence of pyridine was used to prepare trimethylsilyl derivatives. The GC-MS analyses were performed with a Varian 3800 chromatograph coupled to an ion-trap detector (Varian 4000) using a medium-length fusedsilica DB-5HT capillary column (12 m x 0.25 mm internal diameter, 0.1 μm film thickness) from J&W Scientific, enabling simultaneous elution of the different compound classes. The oven was heated from 120 ºC (1 minute) to Lund H. et al. / WO 2013/004639. PCT/EP2012/062763 284 380 ºC at 10 ºC per minute, and held for 5 minutes. Other temperature program, from 50 ºC to 110 ºC (at 30 ºC per minute) and then to 320 ºC (at 6 ºC per minute), was used when necessary. In all GC-MS analyses, the transfer line was kept at 300 ºC, the injector was. Compounds were identified by mass fragmentography, and by comparing their mass spectra with those of the Wiley and NIST libraries and standards, and quantification was obtained from total-ion peak area, using response factors of the same or similar compounds. Single-ion chromatographic profiles (of base or other specific ions) were used to estimate compound abundances when two peaks partially overlapped. Results Three saturated alkanes (dodecane, tetradecane and hexadecane) were tested as A. aegerita peroxygenase substrates. The reactions with dodecane, gave monohydroxylated derivatives at positions 2 and 3. In addition to the monohydroxylated derivatives, dihydroxylations at the positions 2 and 3 from both ends of the molecule (i.e., α+1 and ω -1/ ω -2, or α+2 and ω -1/ ω -2) were identified as the predominant compounds. Table 1 GC-MS peak areas for the peroxygenase reactions. Substrate (ω-1) OH (ω-2) OH 2,(ω-1) di OH 2,(ω-2) di OH 3,(ω-2) di OH ω -1-OH (2+3 keto ) ω -2-OH (2+3 keto ) dodecane 90,000 80,000 2.4x106 4.3x106 2.1x106 1.3x106 1.1x106 tetradecane 120,000 140,000 260,000 420,000 350,000 520,000 740,000 hexadecane 90,000 70,000 60,000 150,000 100,000 100,000 120,000 6. Publicaciones 285 EXAMPLE 2 Enzymatic oxidation of saturated and unsaturated fatty acids The extracellular peroxygenase of A. aegerita (isoform II , 44 kDa, SEQ ID NO: 2) was used. The enzyme preparation was homogeneous by sodium dodecylsulfate-polyacrylamide gel electrophoresis, an exhibited and A418/A280 ratio of 1.75. Its specific activity was 117 units·mg-1 , where 1 unit represents the oxidation of 1 μmol of veratryl alcohol to veratraldehyde (310 9300 M -1 . cm -1 ) in 1 minute at 23 ºC and pH 7, in the presence of 2.5 mM H2O2. Saturated and unsaturated acids were obtained from Sigma-Aldrich: Lauric ( dodecanoic, C 12 ), myristic (tetradecanoic, C14 ), palmitic (hexadecanoic, C 16 ), stearic (octadecanoic, C18), arachidic (eicosanoic, C20 ), lauroleic ( cis -9- dodecenoic, C 12:1 ), myristoleic ( cis -9-tetradecenoic, C 14:1 ), palmitoleic ( cis -9- hexadecenoic, C 16:1 ), oleic ( cis -9-octadecenoic, C 18:1 ), linoleic ( cis,cis- 9,12- octadecadienoic, C 18:2 ) and eicosenoic (C20:1) acids. Five mL reaction s of the above model substrates (1 mM) with the A. aegerita peroxygenase (1 U) were performed in 50 mM sodium phosphate buffer (pH 7) at 25 ºC for 2 hours, in the presence of 2.5 mM H2O2. The substrates were previously dissolved in acetone and added to the buffer (the acetone concentration in the reaction was 15%). In control experiments, substrates were treated under the same conditions but without enzyme. After the enzymatic reactions, water was immediately removed in a rotary evaporator, and the products recovered with chloroform, dried under nitrogen, and redissolved in chloroform for GC-MS analyses. Bis(trimethylsilyl)trifluoroacetamide (Supelco) in the presence of pyridine was used to prepare trimethylsilyl derivatives. The GC-MS analyses were performed with a Varian 3800 chromatograph coupled to an ion-trap detector (Varian 4000) using a medium-length fusedsilica DB-5HT capillary column (12 m x 0.25 mm internal diameter, 0.1 μm film thickness) from J&W Scientific, enabling simultaneous elution of the different compound classes. The oven was heated from 120 ºC (1 minute) to 380 ºC at 10 ºC per minute, and held for 5 minutes. Other temperature program, from 50 ºC to 110 ºC (at 30 ºC per minute) and then to 320 ºC (at 6 ºC per minute), was used when necessary. In all GC-MS analyses, the transfer line was kept at 300 ºC, the injector was programmed from 120 ºC (0.1 minute) to 380 ºC at 200 ºC·per minute and held until the end of the analysis, and helium was used as carrier gas at a rate of 2 ml per minute. Lund H. et al. / WO 2013/004639. PCT/EP2012/062763 286 Compounds were identified by mass fragmentography, and by comparing their mass spectra with those of the Wiley and NIST libraries and standards, and quantification was obtained from total-ion peak area, using response factors of the same or similar compounds. Single-ion chromatographic profiles (of base or other specific ions) were used to estimate compound abundances when two peaks partially overlapped. Results Eleven saturated and unsaturated fatty acids were tested as substrates of the A. aegerita peroxygenase and all fatty acids showed reactivity towards the enzyme. The abundance (relative percentage) of different monohydroxylated , keto, dihydroxylated, keto-hydroxy and dicarboxylic derivatives identified by GC-MS in the reactions of saturated and unsaturated fatty acids are listed in Table 2 . Oxidation of the terminal methyl group (ω-OH) was observed for all tested free fatty acids, in some cases this was further oxidized leading to formation of dicarboxylic acids ( di-COOH ) . EXAMPLE 3 Enzymatic oxidation of tetradecane in 40% acetone The extracellular peroxygenase of A. aegerita (isoform II , 44 kDa, SEQ ID NO: 2) and the recombinant peroxygenase of Coprinopsis cinerea (WT392, SEQ ID NO: 4) were used. The activity of the preparations was determined by oxidation of veratryl alcohol. 1 unit represents the oxidation of 1 μmol of veratryl alcohol to veratraldehyde ( 310 9300 M-1.cm-1) in 1 minute at 23ºC and pH 7, in the presence of 2.5 mM H2O2. Tetradecane (C14 ) was obtained from Sigma-Aldrich. Five mL reaction s of the above model substrate (0.3 mM) with 1 U of peroxygenase were performed in 50 mM sodium phosphat e buffer (pH 7) at 40 ºC for 2 h, in the presence of H 2 O 2 . The concentration of H 2 O 2 was 2.5 mM when A. aegerita peroxygenase was applied and 0.5 mM when using C. cinerea peroxygenase. The substrate was previously dissolved in acetone and added to the buffer ( the acetone concentration in the reaction was 40%). In control experiments, 6. Publicaciones 287 substrates were treated under the same conditions but without enzyme. After the enzymatic reactions, water was immediately removed in a rotary evaporator, and the products recovered with chloroform, dried under nitrogen, and redissolved in chloroform for GC-MS analyses. Bis(trimethylsilyl)-trifluoroacetamide (Supelco) in the presence of pyridine was used to prepare trimethylsilyl derivatives. Table 2 Relative abundance of reaction products. Fatty acid ω-OH ω - 1 OH ω - 2 OH ω - 3 OH ω - 1 Keto ω - 2 Keto di - OH OH - Keto di - COOH C 12 1.3 39.7 32.0 0.2 5.8 1.0 4.4 15.5 0.3 C 12:1 3.3 37.4 59.2 0 <0.1 <0.1 0 0 0 C 14 3.5 34.4 30.5 0.3 20.8 3.3 0.5 6.2 0.6 C 14:1 1.8 0 94.6 0 0 3.6 0 0 0 C 16 1.4 23.6 23.6 0.3 34.5 16.3 0 0 0.3 C 16:1 2.5 35.7 47.0 0.1 10.4 4.4 0 0 0 C 18 <0.1 22.7 27.0 0.1 32.8 17.0 0 0 0.5 C 18:1 1.6 40.8 39.0 0.2 13.0 5.3 0 0 0 C 18:2 1.0 50.2 33.5 2.5 10.0 2.9 0 0 0 C 20 <0.1 16.0 28.1 0 38.7 17.3 0 0 0 C 20:1 1.2 35.0 38.7 0.4 18.8 6.0 0 0 0 The GC-MS analyses were performed with a Varian 3800 chromatograph coupled to an ion-trap detector (Varian 4000) using a medium-length fusedsilica DB-5HT capillary column (12 m x 0.25 mm internal diameter, 0.1μm film thickness) from J&W Scientific, enabling simultaneous elution of the different compound classes. The oven was heated from 120ºC (1 minute) to 380 ºC at 10 ºC per minute, and held for 5 minutes. Other temperature program, from 50 ºC to 110 ºC (at 30 ºC per minute) and then to 320 ºC (at Lund H. et al. / WO 2013/004639. PCT/EP2012/062763 288 6 ºC per minute), was used when necessary. In all GC-MS analyses, the transfer line was kept at 300 ºC, the injector was programmed from 120 ºC (0.1 minute) to 380 ºC at 200 ºC per minute and held until the end of the analysis, and helium was used as carrier gas at a rate of 2 ml per minute. Compounds were identified by mass fragmentography, and by comparing their mass spectra with those of the Wiley and NIST libraries and standards, and quantification was obtained from total-ion peak area, using response factors of the same or similar compounds. Single-ion chromatographic profiles (of base or other specific ions) were used to estimate compound abundances when two peaks partially overlapped. Results Chromatographic profiles resulting from hydroxylation of the saturated alkane tetradecane (C14) are shown in Figure 1 for C. cinerea peroxygenase and Figure 2 for A. aegerita peroxygenase. The reactions with tetradecane, resulted in monohydroxylated derivatives at positions 2 and 3 ( ω -1 and ω -2 OH) and dihydroxylations at the positions 2 and 3 from both ends of the molecule (i.e., ω -1/ ω -1, ω -2/ ω -2 or ω -1/ ω -2 di OH ) . EXAMPLE 4 Enzymatic oxidation of 1-tetradecanol in 20% acetone The extracellular peroxygenase of A. aegerita (isoform II , 44 kDa, SEQ ID NO: 2) and recombinant peroxygenase of Coprinopsis cinerea (WT392, SEQ ID NO: 4) were used. The activity of the preparations was determinated by oxidations of veratryl alcohol. 1 unit represents the oxidation of 1 μmol of veratryl alcohol to veratraldehyde (  310 9300 M-1.cm-1 in 1 minute at 23 ºC and pH 7, in the presence of 2.5 mM H 2 O 2 . 1-Tetradecanol (C14 ) was obtained from Sigma-Aldrich. Five mL reaction s of the above model substrate (0.1 mM) with 1 U of peroxygenase were performed in 50 mM sodium phosphate buffer (pH 7) at 30 ºC for 1 minute, in the presence of H 2 O 2 . The concentration of H 2 O 2 was 2.5 mM when A. aegerita peroxygenase was applied and 0.5 mM when using C. cinerea 6. Publicaciones 289 peroxygenase. The substrate was previously dissolved in acetone and added to the buffer (the acetone concentration in the reaction was 20%). In control experiments, substrates were treated under the same conditions but without enzyme. After the enzymatic reactions, water was immediately removed in a rotary evaporator, and the products recovered with chloroform, dried under nitrogen, and redissolved in chloroform for GC-MS analyses. Bis(trimethylsilyl)-trifluoroacetamide (Supelco) in the presence of pyridine was used to prepare trimethylsilyl derivatives. The GC-MS analyses were performed with a Varian 3800 chromatograph coupled to an ion-trap detector (Varian 4000) using a medium-length fusedsilica DB-5HT capillary column (12 m x 0.25 mm internal diameter, 0.1 μm film thickness) from J&W Scientific, enabling simultaneous elution of the different compound classes. The oven was heated from 120 ºC (1 minute) to 380 ºC at 10 ºC per minute, and held for 5 minutes. Other temperature program, from 50 ºC to 110 ºC (at 30 ºC per minute) and then to 320 ºC (at 6 ºC per minute), was used when necessary. In all GC-MS analyses, the transfer line was kept at 300 ºC, the injector was programmed from 120 ºC (0.1 minute) to 380 ºC at 200 ºC per minute and held until the end of the analysis, and helium was used as carrier gas at a rate of 2 ml per minute. Compounds were identified by mass fragmentography, and by comparing their mass spectra with those of the Wiley and NIST libraries and standards, and quantification was o btained from total-ion peak area, using response factors of the same or similar c ompounds. Single-ion chromatographic profiles (of base or other specific ions) were used to estimate compound abundances when two peaks partially overlapped. Results Chromatographic profiles resulting from oxidation of 1-tetradecanol (Alc) are shown in Figure 3 for C. cinerea peroxygenase and Figure 4 for A. aegerita peroxygenase. The reactions with 1-tetradecanol, resulted in formation of 1- tetradecanoic acid (Acd) and hydroxylated decanoic acid ( ω -1 OH Acd and ω - 2 OH Acd) and two dihydroxylated products (ω-1 OH Alc and ω -2 OH Alc ) . Lund H. et al. / WO 2013/004639. PCT/EP2012/062763 290 CLAIMS 1. A method for introducing a hydroxy or a keto group at the second or third carbon of at least two ends of a substituted or unsubstituted, linear or branched, aliphatic hydrocarbon having at least five carbons and having at least one hydrogen attached to said second or third carbon, comprising contacting the aliphatic hydrocarbon with hydrogen peroxide and a polypeptide having peroxygenase activity; wherein the polypeptide comprises: k) an amino acid sequence which has at least 30% identity to SEQ ID NO: 1, 2, 3, 4, 5, 6, 7, 8, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, or 29; and l) an amino acid sequence represented by one or more of the following motifs: 2. The method of claim 1, wherein the second or third carbon is unsubstituted until contacted with the peroxygenase. 3. The method of claims 1 or 2, wherein the polypeptide comprises or consists of an amino acid sequence having at least 35% identity, preferably at least 40% identity, more preferably at least 45% identity, more preferably at least 50% identity, more preferably at least 55% identity, more preferably at least 60% identity, more preferably at least 65% identity, more preferably at least Motif I: [FL]XX[YF]S[AN]X[FHY]G[GN]GX[YF]N (SEQ ID NO: 9 ) ; Motif II: G[GN]GX[YF]NXX[VA]AX[EH][LF]R (SEQ ID NO: 10 ) ; Motif III: RXXRI[QE][DEQ]S[IM]ATN (SEQ ID NO: 1 1 ) ; Motif IV: S[IM]ATN[PG][EQN][FM][SDN][FL] (SEQ ID NO: 12 ) ; Motif V: P[PDK][DG]F[HFW]R[AP] (SEQ ID NO: 13 ) ; Motif VI: [TI]XXXLYPNP[TK][GV] (SEQ ID NO: 14 ) ; Motif VII: E[HG]DXSX[ST]RXD (SEQ ID NO: 15 ) . 6. Publicaciones 291 70% identity, more preferably at least 75% identity, more preferably at least 80% identity, more preferably at least 85% identity, most preferably at least 90% identity, and in particular at least 95% identity to the mature polypeptide of SEQ ID NO: 1, 2, 3, 4, 5, 6, 7, 8, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28 or 29; preferably SEQ ID NO: 2 or 4. 4. The method of any of claims 1-3, wherein the polypeptide comprises or consists of the amino acid sequence of SEQ ID NO: 1, 2, 3, 4, 5, 6, 7, 8, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28 or 29; preferably SEQ ID NO: 2 or 4; or a fragment thereof having peroxygenase activity. 5. The method of any of claims 1-4, wherein the substituents of the aliphatic hydrocarbon are selected from the group consisting of halogen, hydroxyl, carboxyl, amino, nitro, cyano, thiol, sulphonyl, formyl, acetyl, methoxy, ethoxy, phenyl, benzyl, xylyl, carbamoyl and sulfamoyl. 6. The method of any of claims 1-5, wherein the substituents are selected from the group consisting of chloro, hydroxyl, carboxyl and sulphonyl; in particular chloro and carboxyl. 7. The method of any of claims 1-6, wherein the aliphatic hydrocarbon is an alkane. 8. The method of claim 7, wherein the alkane is pentane, hexane, heptane, octane, nonane, decane, undecane, dodecane, tridecane, tetradecane, pentadecane or hexadecane, or isomers thereof. 9. The method of claim 7, wherein the alkane is undecane, dodecane, tridecane, tetradecane, pentadecane or hexadecane, or isomers thereof. 10. The method of any of claims 1-9, wherein the aliphatic hydrocarbon is unsubstituted. 11. The method of any of claims 1-10, wherein the aliphatic hydrocarbon is linear. 12. The method of any of claims 1-11, wherein the aliphatic hydrocarbon is converted to a diol, by introduction of two hydroxy groups. 13. An enzymatic method for producing polyurethane, comprising converting an aliphatic hydrocarbon to a diol, according to the method of claim 12, and using the diol for producing polyurethane. 14. Use of a diol produced according to claim 12, for producing polyurethane. 15. Use of a peroxygenase for introducing a hydroxyl or a keto group at the second or third carbon of two or more ends of an aliphatic hydrocarbon. 