Production of collagen-targeted recombinant human … · Fisiología de la Facultad de Ciencias de la Universidad de Málaga, ... que vas a ser mamá, compartir con vosotros la alegría

Dpto. Biología Celular, Genética y Fisiología Facultad de Ciencias

Production of collagen-targeted recombinant human growth factors for regenerative medicine purposes:

BMP-6 and bFGF

Memoria presentada por el Licenciado D. Rick Visser para optar al Grado de Doctor con la mención de Doctorado Europeo por la Universidad de Málaga.

Málaga, a 22 de junio de 2009.

Dr. D. Manuel Cifuentes Rueda, Profesor Titular del Departamento de Biología Celular,

Genética y Fisiología de la Facultad de Ciencias de la Universidad de Málaga, CERTIFICA

Que D. Rick Visser ha realizado bajo mi dirección el trabajo experimental que ha llevado a la redacción de la presente memoria de Tesis Doctoral, titulada “Production of collagen-targeted recombinant human growth factors for regenerative medicine purposes: BMP-6 and bFGF”. Considerando que constituye trabajo de Tesis Doctoral, autorizo su presentación para optar al Grado de Doctor con mención de Doctorado Europeo.

Y para que así conste y surta los efectos oportunos, firmo el presente documento en

Málaga, a 12 de mayo de 2009.

Fdo.: Manuel Cifuentes Rueda

Dr. D. José Becerra Ratia, Catedrático del Departamento de Biología Celular, Genética y

Fisiología de la Facultad de Ciencias de la Universidad de Málaga, CERTIFICA Que D. Rick Visser ha realizado bajo mi dirección el trabajo experimental que ha llevado a

la redacción de la presente memoria de Tesis Doctoral, titulada “Production of collagen-targeted recombinant human growth factors for regenerative medicine purposes: BMP-6 and bFGF”. Considerando que constituye trabajo de Tesis Doctoral, autorizo su presentación para optar al Grado de Doctor con mención de Doctorado Europeo.



Fdo.: José Becerra Ratia

Dr. D. José Becerra Ratia, director del Departamento de Biología Celular, Genética y Fisiología de la Facultad de Ciencias de la Universidad de Málaga,

CERTIFICA Que D. Rick Visser ha realizado el trabajo experimental, que ha llevado a la redacción de la

presente memoria de Tesis Doctoral, en los laboratorios del Área de Biología Celular y del Área de Fisiología, considerando que constituye trabajo de Tesis Doctoral.



Fdo.: José Becerra Ratia

Yo, Rick Visser, declaro que soy autor del presente trabajo de investigación y que lo he realizado en el Departamento de Biología Celular, Genética y Fisiología, bajo la dirección del Dr. Manuel Cifuentes Rueda y del Dr. José Becerra Ratia.

Y para que así conste, firmo el presente documento en Málaga, a 12 de mayo de 2009.

Fdo.: Rick Visser

A las tres grandes mujeres de mi vida: mi abuela, mi madre y mi hermana.

A Miguel.

“I – I hardly know who I am, sir, just at present – at least I know

who I was when I got up this morning, but I think I must have been changed

several times since then”

from Alice in Wonderland Lewis Carroll

Agradecimientos / Acknowledgements.

Éste ha sido un largo viaje, con una meta lejana, muchos caminos sin salida, atajos, penas

y alegrías, fracasos y pequeños éxitos. Como era de esperar en un peregrinaje de este calibre,

se hacen compañeros de viaje. Unos hacen todo el camino contigo, otros acaban eligiendo

caminos distintos y otros aparecen de repente, cuando uno no se lo espera, y se convierten en

compañías irreemplazables. No quiero dejar de agradecer a todas esas personas que, en algún

momento, han decidido calzarse las botas y caminar a mi lado.

A José (Pepe) Becerra por darme ese primer empujón para echar a andar y acogerme en

su grupo, por ser un ejemplo de impertérrita constancia y por mantenerse siempre accesible a

pesar de sus infinitas ocupaciones.

A Manuel (Manolo) Cifuentes, por hacerme descubrir el amor por la ciencia, por ser un

ejemplo de entusiasmo y por guiarme por este mundillo, pero dándome siempre la libertad de

cometer mis propias equivocaciones. A pesar de ser uno de los directores de este trabajo,

siempre ha estado cerca en el laboratorio y dispuesto a remangarse la bata para echar todas

las manos que hicieran falta. Pocos doctorandos tienen la suerte de poder decir que su director

de Tesis es, además, un buen amigo.

A Pilar Arrabal, por haber sido (y seguir siendo) la mejor compañera de viaje que uno

pueda desear. Emprendimos el viaje prácticamente juntos, pero ella siempre ha ido un paso por

delante, machete en mano, para abrir caminos en la maleza donde no los había. Juntos hemos

compartido todos los sinsabores de la ciencia, pero también momentos inolvidables de risas,

confidencias y pequeñas aventuras. Espero que sigas caminando infatigable hacia tus metas,

pero también poder seguir disfrutando de tu compañía y de tus sonrisas en el futuro. Y ahora

que vas a ser mamá, compartir con vosotros la alegría de este gran acontecimiento que tanto

se ha hecho esperar.

A mis amigos de Biología Celular, que han hecho el viaje mucho más agradable de lo que

hubiera podido ser. Eva, Leonor, Ana, Ángel, Mercedes, Iván, Irene, Lola, Juan Félix,

Jesús, Silvia, Pedro… y el resto del área en general.

Eva empezó colándose en Fisio Animal como si nada y acabó convirtiéndose en una amiga

que, con gran sinceridad, ha sabido apoyarme en los momentos malos y, con aún mayor gracia,

ha conseguido arrancarme carcajadas en los momentos buenos. ¡Mucha suerte en todo, rubia!

Leonor fue de las primeras que me dio prácticas en la carrera, allá por 1994 (!) y se ha

convertido en un gran referente para mi, tanto por su calidad humana como científica. Juntos

exploramos la Croacia profunda del “reis an biuti” y espero que aún podamos compartir muchas

más experiencias.

Ana apareció como “otra administrativa más” y acabó cautivándonos a “tous” con su

gracia, su sinceridad y su inocencia mezclada con picardía. No podría imaginarme el

departamento sin ella, ni el día a día sin su amistad.

Ángel, a pesar de quererle parecer serio y cortante a los que no le conocen, es una

fantástica persona a la que estimo mucho por su gran sentido del humor y por estar siempre

dispuesto a echar una mano.

Mercedes es un auténtico encanto y en más de una ocasión me ha prestado el apoyo que

necesitaba en ese momento. Pero mucho más importante es la alegría y el cariño que

transmite. A pesar de lo poco que le escribo, espero que sea consciente de que la echo de

menos y que siempre le desearé todo lo mejor.

Silvia Hernández enamora a cualquiera desde el primer momento. Su sentido del humor

y su enorme humanidad la convierten siempre en una compañía inestimable. Me alegro mucho

de tenerla cerca y espero que sea por mucho tiempo.

Iván es también de esos que se van ganando tu amistad sin darse uno cuenta (aparte de

ser el ganador indiscutible comiendo helados) por su gracia y compañerismo. Le deseo todo lo

mejor en su vida científica y personal.

Irene, Lola, Juan Félix, Jesús Santamaría, Pedro, Dani…, siempre han estado a

mano para hacerle a uno el día más agradable. Irene está en fase de reubicación y espero que

le vaya genial. Lola y Juan Félix son dos personas excepcionales y les deseo toda la felicidad

del mundo. Jesús es también una fantástica persona, siempre dispuesto a ayudar en cualquier

situación. ¡Eres un crack! Pedro y Dani han resultado ser una gran adquisición para el “club

del desayuno” y espero que les vaya todo muy bien.

Muchas gracias también a Inés y a Antonia por echarme una mano con los intrincados

formalismos de la burocracia.

A los demás compañeros de departamento que, en mayor o menor medida, han formado

alguna vez parte de mi viaje: José Manuel, Pepi, Antonio, J. Antonio, Diana, Mónica,

Adri, Lolín, Laura, Silvia, Alicia, Elena, Liz, Wilfredo…

Los integrantes del área de Fisiología Animal (Pedro, Juan, Jesús, Margarita, MariaDo,

Mamme…) han sido como una gran familia para mi, ya que me han acogido y aguantado

desde hace ya un montón de años.

Margarita es también de las primeras que conocí (¡hace ya unos 15 años!) y es una de las

personas que más admiro. No sólo ha sacado tiempo para escucharme siempre que me ha

hecho falta, sino que ha conseguido hacerme revolcarme de risa con su humor y alegría. Ojalá

hubiera más Margaritas en el mundo.

MariaDo también me ha brindado horas de buen humor, risas y afecto. Tu visión crítica del

mundo me cautiva y me ha hecho reflexionar en muchas ocasiones.

Pedro, Juan y Jesús han sido grandes puntos de apoyo, siempre dispuestos a ayudar en

todo. Vuestra compañía ha sido una alegría constante.

Mamme y Rafa, con los que he compartido muy buenos ratos, además de laboratorio y

despacho. Os deseo muchísima felicidad y espero poder ser partícipe de ella.

Carolina, Reme, las distintas Patricias… Gracias por vuestra compañía y los buenos

momentos.

Elena y Pablo llegaron sin previo aviso para traer sangre nueva al laboratorio. Ahora que

se han reubicado, espero que les vaya todo muy bien y poder ser testigo de sus futuras Tesis.

José Esteban siempre ha estado dispuesto a atender a todas las peticiones de ayuda. Le

agradezco mucho su dedicación durante ya muchos años.

For four and a half months, a great family adopted me in Braunschweig. Without knowing

me at all, Ursula (Uschi) Rinas opened me her house and her lab, not expecting anything in

exchange. I not only discovered in her a great researcher, from who I learned a lot, but also a

great woman, with who I hope to stay in contact forever.

I met Felipe in a quite tumultuous time of his life, but still he found some time to help me

with the protein folding. I am very thankful for that and wish him, his wife and their kid(s) all

the best.

With Heike and Xin I had a great feeling. They were always willing to help me with

whatever and I will never forget all the laughter we shared. I wish them all the best in their

personal and scientific life, and hope we will meet again someday (maybe another weekend in

Berlin?).

The beginning with Nadine was a little bit tense, but after just a few weeks I discovered in

her a fantastic person, very down to earth, with who I had some great moments. I wish her

also lots of success.

Ich danke euch allen für Ihre Hilfe und Unterstützung!

Gracias a mis compañeros del área de Genética (Eduardo, Cayo, Araceli, Cristina,

Gabriel, Fidela, Migue, Luis…) que tantas veces me han visto aparecer por ahí para pedirles

ayuda, ya fuera material o intelectual. Vuestra paciencia conmigo ha sido un alivio y vuestros

consejos muy tenidos en cuenta.

Mis vecinos de Fisiología Vegetal (Adolfo, Lourdes, Maca, Gema, Carmen, Elena, Sara,

Nieves, Lara, Sergio, Bea, Juan Antonio…) han sido siempre una compañía agradable y

han alegrado muchos almuerzos y “cigarritos en el porche”. Espero poder seguir disfrutando de

esa compañía mucho tiempo más.

Muchísimas gracias a mis compañeros de Zoología (Ramón Muñoz-Chápuli, José María,

Rita, Juan Antonio, Víctor…) por ofrecerme siempre su ayuda y consejos, y por los buenos

ratos que he pasado charlando con ellos. Es una tremenda alegría tener cerca a un grupo de

gente tan competente y tan dispuesta a echar una mano cuando uno lo pide, e incluso sin que

uno lo pida. Os deseo todo lo mejor.

Gracias también a mi pequeño grupo de acogida de Microbiología (Juanjo, Lola, Esther,

Irene, Benjamín…), que me dieron la oportunidad de vivir un pequeño cambio de aires y con

los que disfruté mucho trabajando.

Many thanks to Dr. A. Hari Reddi for his helpful comments and revision of the manuscript.

Mis amigos fuera de mi “mundo laboral” han sido una de las grandes bases en que me he

podido apoyar. A algunos de ellos les veo poco o casi nada, pero todos se han ganado a pulso

mi respeto y mi cariño, y no quisiera tener que prescindir nunca de ninguno de ellos. Los voy a

repasar por orden alfabético para que nadie se sienta menospreciado/a, aunque todos saben

que tienen su propia parcelita en mi corazón.

Si, en el pasado, alguien me hubiera dicho que alguna vez le iba a coger tanto cariño a

Cristina Draper le hubiera mirado con todo el escepticismo. Aunque poca gente sea capaz de

seguirle el ritmo, la alegría y la energía que transmite animan a cualquiera.

Cristina Muñoz y Rafa me llegaron casi por obligación pero, aún así, no me costó el más

mínimo trabajo empezar a cogerles un cariño que crece día a día. Ya he sido testigo (no literal)

de vuestra boda y de vuestra compra de casa y espero poder compartir muchas más alegrías

con vosotros en el futuro.

Eli ha estado muy presente en mi vida en los últimos años y se ha convertido en una gran

amiga, siempre dispuesta a tender una mano, a preocuparse por mis problemas y a compartir

muchísimas risas, comidas, partidas de cartas, ratos de playa, etc.

Con Esther y Manolo he podido pasar muy buenos momentos de norte a sur del país

(Oviedo, Madrid y Málaga). Ahora que ya parece que se han estabilizado, espero que nos

podamos hacer muchas visitas mutuas.

Isa ha sido una inseparable compañera de carrera y una gran amiga, con la que me he

reído hasta caer al suelo. Aunque ahora casi nunca la vea, espero que sepa el cariño que aún le

guardo y que le vaya todo muy bien.

Manolo se ha ganado el derecho a ser un amigo al que admiro mucho por su espectacular

humor, por su sinceridad y por estar siempre ahí. Espero que seas muy feliz y que coseches

muchos éxitos.

Mª Tere ha sido una especie de hermana durante un largo tiempo, con quien he

compartido muchas experiencias. Aunque nos hemos ido distanciando poco a poco, siempre

tendrá todo mi cariño y mi ayuda cuando la necesite.

En los principios de mi Tesis, Marisella me brindó muchos ratos de risas y conversaciones

surrealistas con su particular forma de ser. Ya hace varios años que perdimos el contacto, pero

espero que alcances todos tus sueños.

Raquel y Domingo son también de esos amigos poco vistos pero muy queridos. Siempre

recordaré los buenos momentos que hemos pasado juntos.

Teresa y Jose han sido de los últimos en aparecer en mi camino, pero rápidamente se han

hecho querer. Poco a poco se van acumulando aventuras vividas juntos a lo largo de toda la

geografía.

Gracias a mi “familia política”, que me ha acogido con tanto cariño y que me permiten

sentirme tan a gusto entre ellos. A Isabel y Antonio, por abrirme las puertas de su casa y

tenerme siempre un hueco reservado en su mesa para compartir esas paellas tan fantásticas. A

mis “cuñados” y “concuñadas”, Toni y Susana, Petri, David y Gladys, y Javi, por hacerme

partícipes de sus vidas y poder compartir unas cuantas risas con ellos.

Dank je wel, Willem, voor altijd klaar te staan om te helpen met wat dan ook. Dank zij jou

had ik zomaar een auto voor de deur en alles wat ik nodig had hoefde ik maar voor te vragen.

Je bent een kei!

Tante Rie, uw gevoel voor humor en doorzettingsvermogen is altijd een positief stimulance

geweest. Dank U wel!

Oma, dank U wel voor alles. U heeft altijd aan mij gedacht, en zelfs al had U niet veel, was

er altijd wat voor mij weggelegd.

Seda, jij bent niet alleen mijn enige zus, maar ook altijd een vriendin geweest. Ookal

hebben we soms ruzie gehad (en dat zal wel zo blijven), weet je dat ik zielsveel van je hou, dat

ik altijd aan je denk en dat ik hoop dat je heel gelukkig wordt.

Mama, wat kan ik zeggen? Je bent altijd, of je het gelooft of niet, mijn grootste steun

geweest. Alles wat ik heb bereikt en wat ik nog bereiken kan is dankzij jouw, jouw werk, jouw

moed en jouw liefde. Ookal hebben we soms onze oneinigheden, mijn liefde en dankbaarheid

voor jouw zit zo bij mij ingegroeid dat dat nooit kan verdwijnen.

Miguel, tú sabes bien lo que supones para mi. No habrá adversidad en el mundo capaz de

mermar lo mucho que te quiero ni la ilusión que tengo por compartirlo todo contigo. Gracias

por estar ahí a mi lado.

Gracias. De verdad. Rick

Durante la realización del presente trabajo de investigación, el doctorando ha disfrutado de una beca de Formación de Personal Docente e Investigador (FPDI) de la Junta de Andalucía, de una beca de Formación de Profesorado Universitario (FPU) del Ministerio de Educación y Ciencia, de un contrato de investigación del Centro de Investigación Biomédica en Red en Bioingeniería, Biomateriales y Nanomedicina (CIBER-BBN) y de un contrato de investigación de la Red de Terapia Celular (Red TerCel) del Instituto de Salud Carlos III.

El presente trabajo de investigación se ha financiado con fondos de los siguientes proyectos

e instituciones: - Ministerio de Educación y Ciencia (BIO2006-03599). - Junta de Andalucía, Consejería de Salud (TCRM 0012/2006). - Junta de Andalucía, Consejería de Innovación, Ciencia y Empresa (P07-CVI-2781). - Red de Terapia Celular. Instituto de Salud Carlos III (RD06/0010/0014).

_________________________________________________________________________Index

Text index.

Abbreviations used in this text ..................................................................................... 1

1. Introduction ............................................................................................................. 7

1.1. Bone and bone regeneration................................................................................. 9

1.1.1. The histology of bone.................................................................................. 9

1.1.2. Bone regeneration ..................................................................................... 10

1.1.3. Molecules involved in bone regeneration ...................................................... 11

1.1.4. Clinical and economic aspects of fracture healing.......................................... 15

1.2. Bone morphogenetic protein-6 ............................................................................. 17

1.2.1. The bone morphogenetic proteins ............................................................... 17

1.2.2. General structure of the BMPs..................................................................... 18

1.2.3. Structure of BMP-6..................................................................................... 20

1.2.4. BMP signalling ........................................................................................... 22

1.2.5. Biological activity of BMPs........................................................................... 25

1.2.6. Biological activity of BMP-6 in osteogenesis .................................................. 27

1.3. Basic fibroblast growth factor............................................................................... 29

1.3.1. The fibroblast growth factors ...................................................................... 29

1.3.2. Structure of bFGF ...................................................................................... 29

1.3.3. bFGF secretion........................................................................................... 30

1.3.4. bFGF receptors and bFGF-binding molecules ................................................ 31

1.3.5. bFGF signalling .......................................................................................... 33

1.3.6. bFGF biological activity ............................................................................... 33

1.4. Therapies for bone defect healing ........................................................................ 37

1.4.1. Growth factors for bone defect healing ........................................................ 37

1.4.2. Safety of the clinical use of growth factors ................................................... 38

1.4.3. Osteoconductive carriers............................................................................. 39

1.4.4. Modified growth factors for regenerative medicine ........................................ 41

1.5. Escherichia coli as an expression system............................................................... 43

1.5.1. Obtaining functional proteins from inclusion bodies....................................... 43

1.5.2. In vitro refolding of proteins ....................................................................... 44

1.6. Baculoviruses as expression systems .................................................................... 47

1.6.1. General information on baculoviruses. The baculovirus life-cycle .................... 47

1.6.2. Baculovirus-based expression systems ......................................................... 49

1.6.3. BacPak6™ and Sapphire™ .......................................................................... 51

i

Index_________________________________________________________________________

2. Hypothesis and objectives ....................................................................................... 53

2.1. Hypothesis.......................................................................................................... 55

2.2. Objectives .......................................................................................................... 57

3. Material and methods .............................................................................................. 59

3.1. Obtaining of the genes encoding h-bFGF and hBMP-6 ............................................ 61

3.1.1. Culture of U-2-OS cells ............................................................................... 61

3.2. Cloning into the pET17b expression vector and the pAcGP67B shuttle vector............ 61

3.2.1. Cloning of the hBMP-6 and the hBMP-6-CBD genes into the pET17b

and the pAcGP67B vectors .......................................................................... 62

3.2.2. Cloning of the h-bFGF and the h-bFGF genes into the pAcGP67B vector .......... 64

3.3. Protein production in Escherichia coli .................................................................... 66

3.3.1. Obtaining of bacterial clones for protein production....................................... 66

3.3.2. Protein expression...................................................................................... 68

3.3.3. Isolation of inclusion bodies ........................................................................ 68

3.3.4. Solubilization of inclusion bodies.................................................................. 69

3.3.5. In vitro refolding ........................................................................................ 69

3.4. Protein production in Sf9 insect cells ..................................................................... 73

3.4.1. Culture of Sf9 cells ..................................................................................... 75

3.4.2. Transfection of Sf9 cells.............................................................................. 75

3.4.3. Isolation of viral clones (plaque assay)......................................................... 76

3.4.4. PCR analysis of the viral clones.................................................................... 78

3.4.5. Expansion of the baculovirus clones............................................................. 78

3.4.6. Titering of viral suspensions ........................................................................ 79

3.4.7. Production assays ...................................................................................... 80

3.4.8. Large-scale protein production .................................................................... 80

3.4.9. Purification of rhBMP-6 produced in Sf9 cells ................................................ 81

3.4.10. Purification of rh-bFGF and rh-bFGF-CBD produced in Sf9 cells..................... 82

3.5. Biochemical analysis of the produced proteins ....................................................... 83

3.5.1. SDS-PAGE ................................................................................................. 83

3.5.2. Western blot.............................................................................................. 83

3.5.3. Dot blot..................................................................................................... 84

3.6. Collagen-binding affinity test ................................................................................ 84

3.7. In vitro biological activity tests ............................................................................. 85

3.7.1. Induction of ALP expression on C2C12 mouse myoblasts ............................... 85

3.7.2. Proliferation assay on MC3T3-E1 mouse preosteoblasts ................................. 86

3.7.3. Inhibition of differentiation assay on MC3T3-E1 mouse preosteoblasts ............ 86

3.8. In vivo heterotopic bone formation assay .............................................................. 87

ii

_________________________________________________________________________Index

3.9. Histological analysis of the implanted ACS............................................................. 88

3.9.1. Histochemical stains ................................................................................... 89

3.9.2. Immunohistochemistry ............................................................................... 89

3.10. Statistical analysis............................................................................................. 89

Appendix I. Protocols and recipes ............................................................................... 91

AI.1. Buffers of general use ....................................................................................... 93

AI.2. Recombinant DNA technology ............................................................................ 93

AI.2.1. Total RNA isolation ................................................................................... 93

AI.2.2. Reverse transcription - polymerase chain reaction....................................... 94

AI.2.3. Polymerase chain reaction......................................................................... 94

AI.2.4. Plasmid purification .................................................................................. 95

AI.2.5. DNA electrophoresis ................................................................................. 95

AI.2.6. RNA electrophoresis ................................................................................. 95

AI.2.7. DNA purification from agarose gels ............................................................ 96

AI.2.8. DNA digestion with endonucleases............................................................. 96

AI.2.9. DNA precipitation ..................................................................................... 97

AI.2.10. DNA ligation .......................................................................................... 97

AI.2.11. DNA sequencing..................................................................................... 98

AI.2.12. Plasmids................................................................................................ 98

AI.2.13. Oligonucleotides....................................................................................100

AI.3. Protein expression in E. coli...............................................................................102

AI.3.1. Bacterial cell culture ................................................................................102

AI.3.2. Bacterial cell culture media.......................................................................102

AI.3.3. Bacterial strains ......................................................................................103

AI.3.4. Storage of bacterial clones .......................................................................103

AI.3.5. Transformation of E. coli strains ...............................................................104

AI.3.5.1. Making of electrocompetents...........................................................104

AI.3.5.2. Electroporation ..............................................................................104

AI.3.6. Colony-PCR.............................................................................................105

AI.4. Eukaryotic cell culture.......................................................................................105

AI.4.1. Cell lines.................................................................................................106

AI.4.2. Cell culture media ...................................................................................106

AI.4.3. Cell counting and determination of cell viability..........................................107

AI.5. Protein analysis................................................................................................108

AI.5.1. Protein precipitation with trichloroacetic acid .............................................108

iii

Index_________________________________________________________________________

AI.5.2. SDS-PAGE.............................................................................................. 108

AI.5.2.1. Buffers and reagents ..................................................................... 108

AI.5.2.2. Gel preparation ............................................................................. 109

AI.5.3. Staining of gels with Coomassie blue........................................................ 110


AI.5.3.2. Staining protocol ........................................................................... 110

AI.5.4. Electrotransference of proteins to PVDF.................................................... 111


AI.5.4.2. Transference protocol .................................................................... 111

AI.5.5. Staining of proteins on PVDF with amido black.......................................... 111


AI.5.5.2. Staining protocol ........................................................................... 112

AI.5.6. Immunostaining of proteins on PVDF ....................................................... 112


AI.5.6.2. Immunostaining protocol................................................................ 112

AI.5.7. Development of immunostained proteins .................................................. 113

AI.6. Histological analyses ........................................................................................ 113

AI.6.1. Fixation, decalcification, dehydration and embedding in paraffin ................. 113

AI.6.2. Hematoxylin-eosin staining...................................................................... 114

AI.6.3. Masson’s trichrome staining..................................................................... 114

AI.6.4. Alcian blue staining................................................................................. 115

AI.6.5. Immunohistochemistry............................................................................ 115

Appendix II. Reagents and equipment...................................................................... 117

AI.1. Fungibles ........................................................................................................ 119

AI.2. Reagents ........................................................................................................ 119

AI.3. Equipment ...................................................................................................... 122

4. Results .................................................................................................................... 125

4.1. Obtaining of the gene encoding hBMP-6.............................................................. 127

4.2. Cloning of the genes into the expression vectors.................................................. 127

4.3. Production of rhBMP-6 in Escherichia coli ............................................................ 128

4.3.1. Obtaining of rhBMP-6-expressing clones of E. coli Rosetta™ (DE3) ............... 128

4.3.2. Expression of rhBMP-6 in Escherichia coli ................................................... 129

4.3.3. Refolding of rhBMP-6 produced in Escherichia coli....................................... 130

4.4. Production of rhBMP-6 and rhBMP-6-CBD in Sf9 cells ........................................... 134

4.4.1. Obtaining of rhBMP-6 and rhBMP-6-CBD expressing clones

of baculoviruses........................................................................................ 134

iv

_________________________________________________________________________Index

4.4.2. Production assays for rhBMP-6 and rhBMP-6-CBD ........................................137

4.4.2.1. Production assay for rhBMP-6 ..........................................................137

4.4.2.2. Production assay for rhBMP-6-CBD ...................................................139

4.4.3. Analysis of the influence of the PDI on rhBMP-6 production in Sf9 cells..........140

4.4.4. Expression and purification of rhBMP-6 and rhBMP-6-CBD ............................141

4.4.4.1. Purification of rhBMP-6 ....................................................................141

4.4.4.2. Purification of rhBMP-6-CBD..............................................................143

4.4.4.3. Obtaining of rhBMP-6 and rhBMP-6-CBD under native conditions..........145

4.4.5. In vitro analysis of the biological activity of rhBMP-6 and rhBMP-6-CBD .........146

4.5. Production of rh-bFGF and rh-bFGF-CBD in Sf9 cells..............................................147

4.5.1. Obtaining of rh-bFGF and rh-bFGF-CBD expressing clones

of baculoviruses ........................................................................................148

4.5.2. Production assays for rh-bFGF and rh-bFGF-CBD .........................................149

4.5.2.1. Production assay for rh-bFGF ............................................................149

4.5.2.2. Production assay for rh-bFGF-CBD.....................................................151

4.5.3. Expression and purification of rh-bFGF and rh-bFGF-CBD .............................151

4.5.3.1. Purification of rh-bFGF......................................................................152

4.5.3.2. Purification of rh-bFGF-CBD ..............................................................153

4.5.3.3. Obtaining of rh-bFGF and rh-bFGF-CBD under native conditions...........155

4.5.4. Collagen-binding affinity tests for rh-bFGF and rh-bFGF-CBD ........................157

4.5.5. In vitro analysis of the biological activity of rh-bFGF and rh-bFGF-CBD...........160

4.5.5.1. Mitogenic activity of rh-bFGF and rh-bFGF-CBD on MC3T3-E1 mouse

Preosteoblasts .................................................................................160

4.5.5.2. Inhibition of differentiation of MC3T3-E1 mouse preosteoblasts

by rh-bFGF and rh-bFGF-CBD ...........................................................163

4.6. In vivo heterotopic bone formation......................................................................164

4.6.1. Analysis of the implants with rhBMP-6 alone................................................169

4.6.2. Analysis of the implants with rhBMP-6 and commercial rh-bFGF ....................170

4.6.3. Analysis of the implants with rhBMP-6 and rh-bFGF produced in Sf9 cells.......171

4.6.4. Analysis of the implants with rhBMP-6 and rh-bFGF-CBD ..............................173

5. Discussion................................................................................................................175

5.1. Engineering of the growth factors........................................................................177

5.1.1. Engineering of the gene encoding the rhBMP-6-CBD ....................................177

5.1.2. Obtaining of the genes encoding the rh-bFGF and the rh-bFGF-CBD..............178

5.2. Production of rhBMP-6 in Escherichia coli .............................................................179

5.3. Production of rhBMP-6 and rhBMP-6-CBD in Sf9 cells ............................................181

5.4. Production of rh-bFGF and rh-bFGF-CBD in Sf9 cells..............................................186

v

Index_________________________________________________________________________

5.5. In vivo osteogenic activity of combinations of BMP-6 and bFGF ............................. 191

5.6. Perspectives for the future ................................................................................. 195

6. Conclusions............................................................................................................. 197

7. Bibliography............................................................................................................ 201

Appendix III. Abstract in Spanish / Resumen en español........................................ 221

Figures and tables index.

Fig. 1. Transversal section through the diaphysis of a long bone.......................................... 10

Table 1. Molecules involved in bone regeneration .............................................................. 14

Table 2. Fracture incidence and costs in the EU ................................................................. 15

Table 3. The BMP family .................................................................................................. 17

Table 4. Identity matrix for the BMP family ....................................................................... 18

Fig. 2. Schematic representation of the BMP-7 monomer .................................................... 19

Fig. 3. Schematic representation of the processing of a BMP pre-pro-protein

to obtain an active dimer....................................................................................... 20

Fig. 4. Three-dimensional representation of the BMP-6 dimer.............................................. 21

Table 5. The BMP receptors ............................................................................................. 22

Fig. 5. Representation of the “canonical” BMP-Smad signalling pathway............................... 23

Fig. 6. Representation of the “noncanonical” BMP-MAPK signalling pathway ......................... 24

Fig. 7. Hierarchic model of BMP-induced osteoblastic differentiation..................................... 26

Fig. 8. Schematic representation of the bFGF mRNA and the isoforms of bFGF

resulting from its alternative translation.................................................................. 30

Fig. 9. Three-dimensional representations of the bFGF molecule, based on

crystallization data at 2.2 Å resolution..................................................................... 31

Fig. 10. Schematic representation of the FGF Receptor....................................................... 32

Fig. 11. Representation of the two major pathways for bFGF signalling:

The MAPK and the PKC pathways ......................................................................... 34

Table 6. Recombinant fusion proteins with additional binding domains to cells

or extracellular matrix proteins ............................................................................ 42

Fig. 12. Schematic representation of the events that can happen during protein

folding............................................................................................................... 44

Fig. 13. Schematic representation of a typical baculovirus infection cycle ............................. 48

vi

_________________________________________________________________________Index

Fig. 14. Schematic representation of the homologous recombination event that

gives rise to an infective, recombinant baculovirus ................................................. 51

Fig. 15. The BacPak6™ viral DNA and the Sapphire™ viral DNA .......................................... 52

Fig. 16. Schematic representation of a recombinant engineered growth factor with a

decapeptidic collagen type I-binding domain fused to the N-terminal part

of the molecule................................................................................................... 56

Fig. 17. Schematic overview of the obtaining of the pET17b:BMP-6 and the

pET17b:BMP-6-CBD constructions ........................................................................ 63

Fig. 18. Schematic overview of the obtaining of the pAcGP67B:BMP-6 and the

pAcGP67B:BMP-6-CBD constructions..................................................................... 64

Fig. 19. Schematic overview of the obtaining of the pAcGP67B:bFGF and the

pAcGP67B:bFGF-CBD constructions ...................................................................... 65

Fig. 20. Schematic overview of the main steps needed for recombinant protein

production in E. coli.............................................................................................. 67

Table 7. Attempts on in vitro refolding of rhBMP6 monomers produced in

Escherichia coli .................................................................................................. 71

Fig. 21. Schematic overview of the steps needed to obtain a recombinant baculovirus .......... 73

Fig. 22. Formation of a lysis plaque .................................................................................. 77

Fig. 23. Real size photograph of the absorbable collagen sponge discs ................................ 85

Table 8. Combinations of growth factors tested by the heterotopic bone

formation assay in rats....................................................................................... 88

Fig. 24. Implantation of ACS loaded with growth factors into the dorsal muscles of rats ........ 88

Fig. 25. The pBlueScript® II SK(+) vector ......................................................................... 99

Fig. 26. The pET17b expression vector ............................................................................. 99

Fig. 27. The pAcGP67B shuttle vector ..............................................................................100

Fig. 28. RT-PCR with P5 vs. P6 on U-2 OS total RNA for the amplification of the sequence

encoding the mature domain of the hBMP-6.........................................................127

Fig. 29. PCR analysis of the obtained expression vectors ...................................................127

Fig. 30. rhBMP-6 production in Escherichia coli .................................................................130

Fig. 31. Effect of GSH:GSSG ratio on in vitro refolding of rhBMP-6 expressed in

Escherichia coli ...................................................................................................131

Fig. 32. Effect of antiaggregants, pH and GSH:GSSG ratio on in vitro refolding

of rhBMP-6 expressed in Escherichia coli ...............................................................132

Fig. 33. Effect of protein concentration and GSH:GSSG ratio on in vitro refolding

of rhBMP-6 expressed in Escherichia coli ...............................................................132

Fig. 34. Effect of redox pair, redox pair concentration and N2 supply on in vitro

refolding of rhBMP-6 expressed in Escherichia coli..................................................133

vii

Index_________________________________________________________________________

Fig. 35. Effect of the temperature on in vitro refolding of rhBMP-6 expressed in

Escherichia coli................................................................................................... 133

Fig. 36. Sf9 cells eight days after co-transfection with pAcGP67B:rhBMP-6 and

Sapphire™ linearized baculoviral DNA.................................................................. 135

Fig. 37. Isolation of baculoviral clones............................................................................. 136

Fig. 38. PCR analysis of the isolated baculoviral clones ..................................................... 136

Fig. 39. Production assay for rhBMP-6, analyzed by Western blot ...................................... 138

Fig. 40. Western blot analysis with reducing agents of the rhBMP-6 produced in Sf9 cells.... 138

Fig. 41. Production assay for rhBMP-6-CBD, analyzed by Western blot ............................... 139

Fig. 42. Production assay for rhBMP-6 expressed by BacPak6™ baculoviruses .................... 140

Fig. 43. Purification by heparin-sepharose chromatography of rhBMP-6 expressed

in Sf9 cells ......................................................................................................... 142

Fig. 44. Western blot analysis of the elution fractions obtained by heparin-sepharose

chromatography of rhBMP-6 expressed in Sf9 cells. ............................................... 142

Fig. 45. Proposed matrix elution model for rhBMP-6 forms expressed in Sf9 cells ................ 143

Fig. 46. Purification by heparin-sepharose chromatography of rhBMP6-CBD expressed

in Sf9 cells ......................................................................................................... 144

Fig. 47. Western blot analysis of the elution fractions obtained by heparin-sepharose

chromatography of rhBMP-6-CBD expressed in Sf9 cells......................................... 144

Fig. 48. Western blot analysis of the rhBMP-6 samples after removing the excess

of urea and NaCl ................................................................................................ 145

Fig. 49. ALP activity induced by rhBMP-6 produced in CHO cells on C2C12

mouse myoblasts................................................................................................ 147

Fig. 50. PCR analysis of the isolated baculoviral clones using primers that hybridize

with the baculoviral DNA flanking the insert .......................................................... 148

Fig. 51. PCR analysis of the isolated baculoviral clones ..................................................... 149

Fig. 52. Production assay for rh-bFGF, analyzed by Western blot....................................... 150

Fig. 53. Production assay for rh-bFGF-CBD, analyzed by Western blot................................ 151

Fig. 54. Purification by heparin-sepharose chromatography of rh-bFGF expressed

in Sf9 cells ......................................................................................................... 152

Fig. 55. Immuno-dot blot analysis of the collection fractions of rh-bFGF produced

in Sf9 cells and purified by heparin-sepharose chromatography .............................. 153

Fig. 56. Purification by heparin-sepharose chromatography of rh-bFGF-CBD expressed

in Sf9 cells ......................................................................................................... 154

Fig. 57. Immuno-dot blot analysis of the collection fractions of rh-bFGF-CBD produced

in Sf9 cells and purified by heparin-sepharose chromatography .............................. 154

Table 9. Samples of bFGF after purification by heparin-sepharose chromatography............. 155

Table 10. Samples of bFGF after buffer exchange and concentration................................. 156

viii

_________________________________________________________________________Index

Fig. 58. Analysis by immuno dot-blot of the rh-bFGF and the rh-bFGF-CBD

produced in Sf9 cells after purification and buffer exchange...................................156

Fig. 59. Collagen-binding test of rh-bFGF and rh-bFGF-CBD produced in Sf9 cells ................158

Fig. 60. Stability of the collagen-binding of rh-bFGF and rh-bFGF-CBD

produced in Sf9 cells ..........................................................................................159

Fig. 61. Phenotypical changes induced by rh-bFGF and rh-bFGF-CBD on MC3T3-E1

mouse preosteoblasts ..........................................................................................161

Fig. 62. Proliferation of MC3T3-E1 mouse preosteoblast induced by bFGF ...........................162

Fig. 63. Mitogenic activity curves of rh-bFGF and rh-bFGF-CBD ..........................................162

Fig. 64. ALP activity in cultures of MC3T3-E1 mouse preosteoblasts in the presence of

ascorbic acid and bFGF ........................................................................................163

Fig. 65. Staining of the implants without BMP-6 with H-E ..................................................165

Fig. 66. Staining of the implants with BMP-6 with H-E and Masson’s trichrome ....................167

Fig. 67. Staining of the implants with alcian blue ..............................................................168

Fig. 68. Immunostaining of the implants with an anti-osteopontin antibody ........................169

Fig. 69. Histological analysis of the implants with rhBMP-6 alone........................................170

Fig. 70. Histological analysis of the implants with rhBMP-6 + commercial rh-bFGF...............171

Fig. 71. Histological analysis of the implants with rhBMP-6 + rh-bFGF produced

in Sf9 cells.........................................................................................................172

Fig. 72. Histological analysis of the implants with rhBMP-6 + rh-bFGF-CBD

produced in Sf9 cells ..........................................................................................173

ix

__________________________________________________________________Abbreviations

Abbreviations used in this text.

A.

ACS: absorbable collagen sponge.

ActR: activin-like receptor.

aFGF (=FGF1): acidic fibroblast growth factor.

ALK: activin receptor-like kinase.

ALP: alkaline phosphatase.

AMSH: associated molecule with the SH3 domain of STAM.

APS: ammonium persulfate.

B.

β-1-LAP: latency-associated peptide of TGF-β.

BAMBI: BMP and activin membrane bound inhibitor.

bFGF (=FGF2): basic fibroblast growth factor.

BMP: bone morphogenetic protein.

BMPR: BMP receptor.

bp: base pair.

BSA: bovine serum albumin.

C.

CBD: collagen-binding domain.

CDMP: cartilage-derived morphogenetic protein.

cDNA: complementary DNA

CHES: 2-(N-Cyclohexylamino) ethanesulfonic acid.

CHO: Chinese hamster ovary.

CIZ: cas-interacting zinc finger protein.

CNS: central nervous system.

Co-Smad: common-partner Smad.

D.

DAB: 3,3’-diaminobenzidine.

DAG: diacyl glycerol.

DAN: differential screening-selected gene aberrative in neuroblastoma.

DBM: demineralized bone matrix.

DEPC: diethyl pyrocarbonate.

DMEM: Dulbecco's modified Eagle medium.

1

Abbreviations__________________________________________________________________

DMSO: dimethyl sulfoxide.

DNA: deoxyribonucleic acid.

dNTP: deoxyribonucleotide triphosphate.

dpp: decapentaplegic.

dsDNA: double stranded DNA.

DTT: dithiothreitol.

E.

ECM: extracellular matrix.

EDTA: ethylenediaminetetraacetic acid.

EGF: epidermal growth factor.

ELISA: enzyme-linked immunosorbent assay.

EMEA: European Medicines Agency.

F.

FBS: foetal bovine serum.

FDA: U.S. Food and Drug Administration.

FGF-BP: FGF-binding protein.

FGFR: FGF receptor.

FRS2: FGF receptor substrate 2.

G.

GDF: growth and differentiation factor.

Gnd-HCl: guanidine hydrochloride.

GOI: gene of interest.

GRB2: growth factor receptor-bound protein 2.

GSH: glutathione (reduced form).

GSSH: glutathione (oxidized form).

GV: granulovirus.

H.

HA: hydroxyapatite.

H-E: hematoxylin-eosin.

HGF: hepatocyte growth factor.

HRP: horseradish peroxidase.

HSPG: heparan sulphate proteglycan.

HVVS: high volume virus stock.

2

__________________________________________________________________Abbreviations

I.

Ibp: inclusion body protein.

IG: immunoglobulin.

IGF: insulin-like growth factor.

IGFBP: insulin-like growth factor binding protein.

IL: interleukin.

IPTG: isopropyl-β-D-thiogalactopyranoside.

IP3: inositol triphosphate.

I-Smad: inhibitory Smad.

K.

Kb: kilobase.

Kbp: kilobase pair.

KDa: kilodalton.

L.

LB: Luria-Bertani culture medium.

Lef-1: lymphoid enhancer-binding factor-1.

M.

MAPK: mitogen-activated protein kinase.

MCS: multiple cloning site.

MEM: minimum essential medium.

MES: 2-(N-morpholino)ethanesulfonic acid.

MNPV: multiple nucleopolyhedrovirus.

mRNA: messenger ribonucleic acid.

MSC: mesenchymal stem cell.

MSV: master stock of virus.

MTT: 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromid.

MW: molecular weight (molecular mass).

N.

NDSB256: non detergent sulfobetaine 256.

Nemo: nuclear factor kappa B essential Modulator.

NLK: Nemo-like kinase.

nls: nuclear localizing sequence.

NPV: nucleopolyhedrovirus.

3

Abbreviations__________________________________________________________________

O.

OC: osteocalcin.

ODx: optical density measured at “x” nm.

OP: osteogenic protein.

ORF: open reading frame.

P.

PBS: phosphate buffered saline.

PBST: phosphate buffered saline – tween20.

PCR: polymerase chain reaction.

PDI: protein disulfide isomerase.

PDGF: platelet-derived growth factor.

pfu: plaque forming units.

pfu polymerase: polymerase from Pyrococcus furiosus.

PG: proteoglycans.

p.i.: post-infection.

pI: isoelectric point.

PIP2: phosphatidyl-inositol-4, 5-bisphosphate.

PLCγ: phospholipase C gamma.

p-NPP: p-nitrophenyl phosphate.

Polh: polyhedrin.

PPolh: polyhedrin promoter.

PSA: ammonium persulphate.

PTHrP: parathyroid hormone-related protein.

PVDF: polyvinylidene fluoride.

R.

RNA: ribonucleic acid.

rpm: revolutions per minute.

R-Smad: receptor-activated Smad.

RT-PCR: reverse transcription – polymerase chain reaction.

S.

SD: standard deviation.

SDS: sodium dodecyl sulfate.

SDS-PAGE: sodium dodecyl sulfate polyacrylamide gel electrophoresis.

Sf: Spodoptera frugiperda.

SHC: Scr homologous and collagen protein.

4

__________________________________________________________________Abbreviations

Smad: small mothers against decapentaplegic.

SMURF: smad ubiquitin regulatory factor.

SNPV: single nucleopolyhedrovirus.

SOS: son of sevenless.

STAM: signal-transducing adaptor molecule.

STAT1: signal transduction and activator of transcription 1.

T.

Ta: annealing temperature.

TAB1: TAK1 binding protein.

TAK1: TGF-β activated kinase1.

TB: terrific broth culture medium.

TCA: trichloroacetic acid.

TCF-1: transcription factor-1.

TCID50: tissue culture infectious dose 50.

TEMED: N, N, N', N'-Tetramethyl-1-, 2-diaminomethane.

TGF-β: transforming growth factor-beta.

TK: tyrosine kinase.

TNF-α: tumor necrosis factor-alpha.

TNM-FH: Trichoplusia ni medium – formulation Hink.

Tris: tris (hydroxymethyl) aminomethane.

tRNA: transfer ribonucleic acid.

TS: transfection supernatant.

U.

UV: ultraviolet.

V.

Vgr: vegetal related.

VS: virus stock.

vWF: von Willebrand factor.

X.

XIAP: X-linked inhibitor of apoptosis.

2.

2xYT: 2 x yeast extract-tryptone culture medium.

5

Abbreviations__________________________________________________________________

4.

4-MPAA: 4-Mercaptophenylacetic acid.

6

1. Introduction.

7

8

Introduction

9

1.1. Bone and bone regeneration.

1.1.1. The histology of bone.

Osseous or bone tissue is a specialized type of connective tissue, of which the major

constituent is extracellular matrix. The organic part of this matrix is mainly composed of type I

collagen, though type V and, to a lesser extent, types III, XI and XIII are also found. Just 10%

of the proteins of the bone matrix are non-collagenic, these being proteoglycans, osteonectin,

osteopontin, sialoproteins I and II, osteocalcin (OC) and certain growth factors like insulin-like

growth factor (IGF), tumor necrosis factor-alpha (TNF-α), transforming growth factor-beta

(TGF-β), platelet-derived growth factor (PDGF), bone morphogenetic proteins (BMPs) and

interleukins -1 and -6 (IL-1 and IL-6). The inorganic part of the matrix is formed by calcium

phosphate in the form of hydroxyapatite (HA), which is deposited on the proteinaceous lattice.

In the protective exterior portion of all bones, the matrix is dense (compact bone tissue), while

in the inside it forms a porous network of trabeculae (cancellous or spongy bone tissue) filled

with bone marrow. Covering the outer surface of bones, for except the joints, is a thin layer of

dense, irregular connective tissue named periosteum, which contains fibroblasts and

osteoprogenitor cells. On the other side, lining the surface of the bony tissue that forms the

medullar cavity of long bones is another osteoprogenitor-containing, thin layer of connective

tissue, called endosteum.

Mature bone is mainly formed by cylindrical structural units called osteons or Haversian

systems (Fig. 1). These run parallel to the longitudinal axis of the bone, and possess a central

canal, called Haversian canal, which contains blood vessels and nerves. The Haversian canal is

surrounded by concentric layers of matrix called lamellae. In between the lamellae lay the

mature osseous cells, or osteocytes, which communicate with each other through long

cytoplasmic extensions that occupy tiny canals called canaliculi. The long, longitudinal

Haversian canals are communicated by shorter, perpendicular Volkmann´s canals.

The osteoprogenitor cells are derived from mesenchymal stem cells (MSCs), for which the

bone marrow seems to constitute the main reservoir. These osteoprogenitors can differentiate

into osteoblasts when they are triggered by the proper stimuli. Osteoblasts, which maintain the

capacity to divide their selves, are responsible for secreting the components of the organic part

of the extracellular matrix, also called osteoid. They also secrete matricial vesicles containing

high amounts of alkaline phosphatase (ALP), an enzyme that liberates phosphate groups (PO4-)

from different molecules of the extracellular matrix. These phosphates can react with Ca2+

cations fixed by osteocalcin and other sialoproteins, forming CaPO4 crystals, which trigger

mineralization of the osteoid surrounding the osteoblast by HA [Ca10(PO4)6(OH)2] deposition.

Once the bone matrix that surrounds an osteoblast is completely mineralized, the cell becomes

http://en.wikipedia.org/wiki/Medullary_cavity

http://en.wikipedia.org/wiki/Bone

http://en.wikipedia.org/wiki/Canaliculus_%28bone%29

Introduction _

10

less active and is now called an osteocyte. Osteocytes have just a limited capacity of both

forming and resorbing extracellular matrix, contributing to the homeostasis of calcium levels in

blood. They are also implicated in mechanotransduction, responding to mechanical stimuli

acting on the bone.

Figure 1. Transversal section through the diaphysis of a long bone. Modified from the U.S. National Cancer Institute's Surveillance, Epidemiology and End Results (SEER) Program.

The third type of cells present in bone is the osteoclast, which is formed by differentiation

of a mononuclear haematopoietic progenitor cell of the bone marrow. Osteoclasts are the cells

responsible for bone resorption by pumping protons to the extracellular matrix, causing a local

descent of the pH which leads to partial demineralization. This demineralized matrix is the

substrate for enzymes such as cathepsin K and matrix metalloproteinases, secreted by the

osteoclasts to digest collagen and other proteins of the bone matrix (Ross MH and Pawlina W,

2007).

1.1.2. Bone regeneration.

When a bone becomes broken, neutrophils and macrophages are the first cells to act by

cleaning up the site of fracture, as happens when any other type of tissue is damaged. Then,

new capillary vessels proliferate at the site of the fracture, and fibroblasts invade the damaged

tissue. This leads to the formation of a new loose connective tissue, called granulation tissue,

Osteocyte Osteon of compact bone

Spongy bone

Haversian canals

Volkmann´s canal

Periosteum

Osteon

Canaliculi

Lamellae

Introduction

11

which becomes gradually more compact and, at some places, gives rise to cartilage. The

formed dense connective tissue and cartilage proliferate to cover the bone at the site of the

fracture, forming a callus. During this process, the osteoprogenitor cells of the periosteum

proliferate and differentiate into osteoblasts, which start secreting new osseous tissue at the

external surface of the bone, at a certain distance from the site of fracture. This ossification

progresses towards the fracture until the new-formed bone constitutes a sheath that surrounds

the callus. This osseous sheath sends capillaries and osteoblasts into the callus to form new

bone tissue inside of it, converting it into an osseous callus. At the same time, cells from the

endosteum of the fractured bone also differentiate into osteoblasts, which synthesize new

spongy bone into the medullar cavity. As happens when normal ossification occurs, this spongy

bone will be gradually replaced by compact bone, at the same time the bony callus is being

eliminated by osteoclasts and remodelled to recover the original shape and function of the

bone. During the entire process, the new-forming blood vessels that grow inside the callus act

as a source of new MSCs. In fact, recent publications have given strong evidences of a

perivascular origin for the MSCs (Crisan M et al., 2008; da Silva Meirelles et al., 2008).

Usually, in healthy people, the entire healing process takes between six and twelve weeks,

depending on the seriousness of the fracture and the affected bone (Ross MH and Pawlina W,

2007).

1.1.3. Molecules involved in bone regeneration.

Fracture healing is a complex cascade of biological events that involves mechanical stress

and both intracellular and extracellular molecular signalling for osteoinduction and conduction.

These kind of multistage processes require regulation by many local and systemic regulation

factors, such as growth and differentiation factors, hormones and cytokines (Tsiridis E et al.,

2007).

The molecules that promote osteogenesis during a fracture healing can be divided into

three distinct groups: i) pro-inflammatory cytokines, ii) growth and differentiation factors, and

iii) metalloproteinases and angiogenic factors (Gerstenfeld LC et al., 2003).

I. Pro-inflammatory cytokines: Tumor necrosis factor-alpha (TNF-α) and interleukins-1 and

-6 (IL-1 and IL-6) show peak expression levels within the first 24 hours after fracture, initiating

the cascade of events that leads to healing (Einhorn TA et al., 1995; Gerstenfeld LC et al.,

2003). Secreted by macrophages and cells of mesenchymal origin located in the periosteum,

these cytokines induce downstream responses by exerting chemotaxis on inflammatory and

endogenous fibrogenic cells, enhancing extracellular matrix synthesis and stimulating

angiogenesis (Kon T et al., 2001).

Introduction _

12

II. Growth and differentiation factors: Transforming growth factor-beta (TGF-β) is released

by platelets at the initial inflammatory phase, and might be responsible for initiating callus

formation (Bostrom MP, 1998). This growth factor is also secreted by osteoblasts and

chondrocytes, and stored in the bone matrix, acting as a potent chemotactic stimulator of

MSCs, preosteoblasts, chondrocytes and osteoblasts (Lieberman JR et al., 2002). It also induces

the production of constituents of the extracellular matrix, such as collagen, proteoglycans,

osteopontin, osteonectin and alkaline phosphatase (Sandberg MM et al., 1993), and may initiate

signalling for bone morphogenetic protein synthesis by the osteoprogenitor cells (Lieberman JR

et al., 2002) as well as inhibiting osteoclast activation and promoting osteoclast apoptosis

(Mundy GR, 1996).

Bone morphogenetic proteins (BMPs) are members of the TGF-β superfamily and are

produced by osteoprogenitors, mesenchymal cells, osteoblasts and chondrocytes within the

extracellular matrix. They induce a sequential cascade of events to promote chondro-

osteogenesis, being responsible for chemotaxis, mesenchymal and osteoprogenitor cell

proliferation and differentiation, angiogenesis and controlled production of extracellular matrix

(Prisell PT et al., 1993). BMPs may also stimulate secretion of other bone and angiogenic

growth factors, such as insulin-like growth factor (IGF) and vascular-endothelial growth factor

(VEGF) (Deckers MM et al., 2002). Among the different subgroups into which these growth

factors are divided, each type of BMP has a unique role and distinct temporal expression

patterns during the fracture repair process.

Fibroblast growth factors (FGFs) are synthesized during bone healing by monocytes,

macrophages, mesenchymal cells, osteoblasts and chondrocytes, to promote growth and

differentiation of a variety of cells, such as fibroblasts, myocytes, osteoblasts and chondrocytes.

These factors play a critical role in angiogenesis and mesenchymal cell proliferation during the

early stages of fracture healing. Among the different types of FGFs, acidic fibroblast growth

factor (aFGF) mainly regulates chondrocyte proliferation and maturation, while basic fibroblast

growth factor (bFGF) is expressed by osteoblasts and seems to exert a more potent effect than

aFGF (Lieberman JR et al., 2002).

Platelet-derived growth factor (PDGF) is secreted by platelets at the early stages of fracture

healing, but is also released by monocytes, macrophages, endothelial cells and osteoblasts. It

acts as a potent chemotactic stimulator for inflammatory cells and exerts strong mitogenic

effects on MSCs and osteoblasts (Lieberman JR et al., 2002).

Insulin-like growth factors (IGFs) are found in the bone matrix, released by endothelial

cells, osteoblasts and chondrocytes. IGF-I induces bone matrix formation by fully differentiated

osteoblasts (Canalis E, 1980), while IGF-II seems to act at a later stage of fracture healing by

stimulating type I collagen production, cartilage matrix synthesis and cell proliferation (Prisell

PT et al., 1993).

Introduction

13

III. Metalloproteinases and angiogenic factors: At the final stages of endochondral

ossification, specific matrix metalloproteinases degrade cartilage and bone to allow infiltration of

blood vessels. The regulation of angiogenesis seems to be regulated by two separate pathways:

a VEGF-dependent pathway and an angiopoietin-dependent pathway (Gerstenfeld et al., 2003).

VEGF is a mediator of neoangiogenesis and a mitogen for endothelial cells (Ferrara N and

Davis-Smyth T, 1997), while angiopoietins are regulatory vascular morphogenetic molecules

related to the formation of larger vessels and development of ramifications from existing ones.

Obviously, to achieve restoration of the original shape and function of the damaged bone,

not only osteogenic molecules, but also inhibitory molecules are necessary. These inhibitors can

be divided into two groups: i) BMP inhibitors, and ii) other inhibitory molecules (Tsiridis E et al.,

2007).

I. BMP inhibitors: Many molecules that inhibit BMP signalling at the extracellular level have

been described. Some of these proteins, such as noggin, gremlin and chordin, antagonize BMP

signalling by binding to specific BMPs and blocking their coupling to their receptors (Groppe J et

al., 2002; Piccolo S et al., 1996; Hsu DR et al., 1998). Other antagonists of BMP activity, such

as sclerostin, directly bind to the BMP receptors (Sutherland MK et al., 2004), while follistatin

binds to BMP receptors through BMPs, forming a trimeric complex (Iemura S et al., 1998).

The BMP and activin membrane bound inhibitor (BAMBI) is structurally related to type I

BMP receptors in the extracellular domain, but it lacks the intracellular domain. Therefore, it

inhibits signalling within the cells by preventing the formation of active receptor complexes

(Onichtchouk D et al., 1999).

Since intracellular BMP signalling occurs via a smad (small mothers against

decapentaplegic) signalling cascade, it can also be modulated negatively by inhibitory smads

(Itoh F et al., 2001) or molecules that promote smad degradation, such as the smad ubiquitin

regulatory factor (SMURF)-1 and -2 (Zhu H et al., 1999). Other intracellular proteins that bind

to signalling smads to inhibit the BMP pathway are the oncoprotein ski (Wang W et al., 2000),

the anti-proliferative protein tob (Yoshida Y et al., 2000), smad-8B (Nishita M et al., 1999) and

the cas-interacting zinc finger protein (CIZ) (Shen ZJ et al., 2002).

II. Other inhibitory molecules: Certain cytokines, such as IL-1α, might inhibit osteogenesis

as it has been shown to decrease ALP activity and type I collagen production in osteoblasts in

vitro (Tanabe N et al., 2004). Also IGF-binding proteins (IGFBPs), such as IGFBP-2 and -4,

might inhibit osteogenesis, since it is thought that these molecules diminish the mitogenic

activities of IGF-I and IGF-II in human osteoblast-like cells (Mohan S et al., 1989; McCarthy TL

et al., 1994).

Introduction _

14

Surprisingly, many investigations point to the fact that the potent osteogenic factors TGF-β

and FGFs might also have inhibitory activities. For example, TGF-β was shown to block BMP-2-

mediated stimulation of terminal osteoblast differentiation in vitro (Spinella-Jaegle S et al.,

2001), while it has been suggested that FGF signalling might stimulate early differentiation of

osteogenic precursors, but inhibit late differentiation and mineralization (Fakhry A et al., 2005).

The most important molecules involved in bone regeneration can be found summarized in

Table 1.

OSTEOGENIC INDUCERS Factor Produced by Effects

• TNF-α • IL-1 • IL-6

• Macrophages • Mesenchymal cells

at the periosteum

• Chemotaxis on inflammatory and fibrogenic cells • ↑ extracellular matrix synthesis • ↑ angiogenesis

• TGF-β

• Platelets • Osteoblasts • Chondrocytes

• Initiates callus formation • Chemotaxis on MSCs, preosteoblasts, chondrocytes and osteoblasts • ↑ extracellular matrix synthesis • ↑ BMP synthesis • ↑ osteoclast apoptosis • ↓ osteoclast activation

• BMPs

• Osteoprogenitors • Mesenchymal cells • Osteoblasts • Chondrocytes

• Chemotaxis • Mesenchymal and osteoprogenitor cell proliferation and differentiation • ↑ angiogenesis • ↑ extracellular matrix synthesis • ↑ secretion of IGF and VEGF

• FGFs

• Monocytes • Macrophages • Mesenchymal cells • Osteoblasts • Chondrocytes

• Mesenchymal cell proliferation • Growth and differentiation of fibroblasts, myocytes, osteoblasts and

chondrocytes. • ↑ angiogenesis

• PDGF

• Platelets • Monocytes • Macrophages • Endothelial cells • Osteoblasts

• Chemotaxis on inflammatory cells • Growth of MSCs and osteoblasts

• IGFs

• Endothelial cells • Osteoblasts • Chondrocytes

• ↑ extracellular matrix synthesis • Cell proliferation

OSTEOGENIC INHIBITORS Factor Produced by Effects

• Noggin • Chordin • Gremlin • Follistatin

• Osteoblasts • Osteocytes

• ↓ BMP signalling

• Sclerostin • Osteoclasts • Osteocytes


• IL-1α • Monocytes • Macrophages

• ↓ ALP and type I collagen production by osteoblasts

• IGFBP • Osteoblasts • ↓ IGF activity • TGF-β

• Platelets • Osteoblasts • Chondrocytes


• FGFs

• Monocytes • Macrophages • Mesenchymal cells • Osteoblasts • Chondrocytes

• ↓ late osteoblastic differentiation

Table 1. Molecules involved in bone regeneration.

Introduction

15

1.1.4. Clinical and economic aspects of fracture healing.

Reparation of bone defects and fractures is a major clinical and economic concern. A study

carried out by Polinder, using data from ten European countries revealed that during the year

1999, nearly 2.5 million cases of injury were registered in the hospitals of these countries. From

all these cases, hip fracture was not only the most common, with an incidence of 2.3 per 1,000,

but also the most expensive injury to treat, with an average cost of € 5,530 (Polinder S et al.,

2005). The total cost of hip fractures in the EU was estimated at € 598 million per year (Finnern

HW and Sykes DP, 2003). The average costs of other fractures range from € 1,131 to € 3,504

(Table 2).

INJURY TYPE INCIDENCE RANK INCIDENCE MEAN COSTS (€)

Hip fracture 1 2.3 ‰ 5,530

Knee/lower leg fracture 5 0.9 ‰ 3,504

Wrist fracture 8 0.8 ‰ 1,374

Elbow/forearm fracture 9 0.6 ‰ 1,726

Ankle fracture 10 0.5 ‰ 2,632

Facial fractures 11 0.5 ‰ 1,379

Hand/finger fracture 16 0.4 ‰ 1,131

Upper arm fracture 17 0.3 ‰ 2,818

Rib/sternum fracture 20 0.2 ‰ 2,126

Foot/toe fracture 24 0.2 ‰ 2,514

Clavicle/scapula fracture 26 0.1 ‰ 2,152

Table 2. Fracture incidence and costs in the EU. Modified from Polinder S et al., 2005.

In the United States, over 7.9 million fractures are sustained each year, being trauma the

second most expensive medical problem, with a cost for the US health care system of $ 56,000

million per year. Nearly half of this amount is used for the treatment of broken bones (Bishop

GB and Einhorn TA, 2007).

The expected time for a fracture to heal naturally is between six and twelve weeks, but

there is a high rate of delayed unions, varying from 16-60% for less severe fractures to 43-

100% for more severe cases. A fracture that shows motion at the bony ends and is not

completely healed within 6 months is considered a non-union, which rate has been reported to

range from 4 to 10% (Garrison KR et al., 2007). Non-unions can not only lead to significant

pain, inhibition of function and decreases in personal and professional productivity, but also

enormously raise the economic implications for healthcare providers.

Introduction _

16

The rate of delayed or non-unions is especially high in elderly patients, in which the titer of

MSC within the bone marrow is diminished. While one of every 250,000 bone marrow cells is

estimated to be a MSC at the age of 30, the titer decreases to one of every 2,000,000 cells in

80 years old individuals (Caplan AI, 2007).

External fixation devices may help stabilizing fractures at risk from poor healing, but this

often result in the production of unstable bone with a high probability of refracture (Braddock M

et al., 2001). Extended bone defects following trauma or cancer resection, or non-union

fractures may require more sophisticated treatments than standard conservative or surgical

therapies. In these cases, segmental bone transport, distraction osteogenesis, bone grafting or

biomaterials must be applied for reconstruction (Kneser U et al., 2006).

Bone transport is based on the methods developed by Gavriil Ilizarov. For reparation of

bone defects by these approaches, a length of bone, above or below the defect, is fixed to an

external fixation device and separated from the remaining bone by an osteotomy. This chosen

piece of bone is then slowly (less than 1 mm/day) moved towards the defect, allowing the site

of the osteotomy to be filled with a new-formed callus, which will calcify and form new bone

once the distraction forces are removed (LaBianco et al., 1996). As the treatment with this

technique requires a long period of time, the surgeon may face many problems which may

negatively affect the final outcome (pin tract infection, early or delayed consolidation, axial

deviation, skin inversion, rupture of the bone by the wires, joint contractures, etc.).

Only in the United States, more than 1.5 million bone grafts are performed annually. By

bone grafting, the missing bone is replaced with material from the body of the own patient or

with a natural substitute. When autologous bone is used, it is typically harvested from the iliac

crest of the pelvis. Allografts from cadavers or living donors may also be used, and are usually

sourced from a bone bank. Although bone grafting is generally successful, the limited amount

of available donor tissue and the high associated morbidity, resulting in numbness or tingling at

the donor site, infection, or prolonged pain, make the need for development of alternative

therapies evident (Braddock M et al., 2001).

More recently, the medical field is focused on the use of natural or synthetic biomaterials

(i.e. materials which are compatible with living cells and tissues) for bone repair, being the aim

of these products to mimic the osteoconductive properties of bone grafts. To also confer

osteoinductive capacity to these grafts, their application in combination with osteogenic growth

factors is being widely studied. These alternatives will be further discussed in section 1.4.

Introduction

17

1.2. Bone morphogenetic protein 6.

1.2.1. The bone morphogenetic proteins.

At the end of the 19th century it was already demonstrated that decalcified bovine bone

could be used for the treatment of osteomyelitis (Senn N, 1889). Some decades later, in the

middle of the 20th century, Lacroix postulated that bone might contain an inductive substance,

which he named osteogenin. A few years later, Marshall R. Urist discovered that demineralized

lyophilized bone matrix was able to promote new endochondral bone formation when implanted

subcutaneously or in intramuscular pockets. This discovery led to the isolation of a low-

molecular mass glycoprotein from bone with the capacity of promoting bone formation when

ectopically located (Urist MR, 1965; Urist MR et al., 1976). In the early 1980s, Sampath and

Reddi made the first attempts on purifying the molecules responsible for bone formation

present in the bone matrix. They successfully isolated a pool of soluble proteins with a

molecular mass under 50 KDa which was shown to possess osteoinductive properties (Sampath

TK et al., 1982; Sampath TK and Reddi AH, 1983). But the real identity of the proteins

responsible for bone induction remained unknown until the purification and sequence of bovine

BMP-3 and the cloning of human BMP-2 and -4 in the late 1980s (Wozney JM et al., 1988;

Wang EA et al., 1988; Luyten FP et al., 1989; Celeste AJ et al., 1990; Wozney JM, 1992). To

date, around 20 BMP family members have been identified and characterized (Table 3).

BMP SUBFAMILY BMP OTHER NAMES RESIDUES (PRE-PRO-PROTEIN)

BMP-2 396 BMP-2/4

BMP-4 BMP-2B 408

BMP-3 Osteogenin 472 BMP-3

BMP-3B GDF-10 478

BMP-5 454

BMP-6 Vgr-1 513

BMP-7 OP-1 431

BMP-8A OP-2 402

OP-1/BMP-7

BMP-8B OP-3 402

BMP-12 CDMP-3 / GDF-7 450

BMP-13 CDMP-2 / GDF-6 455 CDMP

BMP-14 CDMP-1 / GDF-5 501

BMP-9 GDF-2 429

BMP-10 424

BMP-11 GDF-11 407

BMP-15 GDF-9B 392

Others

BMP-16 280

Table 3. The BMP family. Modified from Reddi AH, 2001.

Introduction _

18

Analysis of the amino acid sequences of these proteins revealed high identity with TGF-β,

leading to their inclusion as a family into the TGF-β superfamily. The BMP family genes are

highly conserved among evolution, and homologs can be found from Drosophila to Homo

sapiens. Due to their structural homology, these BMPs of distant species have shown to be

functionally interchangeable (Padgett RW et al., 1993; Sampath TK et al., 1993).

Today, BMPs are included in at least 4 subfamilies attending to the sequence homology of

the mature domain of the proteins (Table 4). Besides the BMPs, more than 40 other proteins

belonging to the same family have been identified in different tissues (Reddi AH, 1997).

Although sharing their name and location, BMP-1 is not a member of the TGF-β

superfamily, but a metalloprotease that cleaves the COOH-propeptides of procollagens I, II and

III (Kessler E et al., 1996).

BM

P-7

BM

P-5

BM

P-6

BM

P-8

A

BM

P-8

B

Vgr

-D

UN

IVIN

BM

P-2

BM

P-4

dpp

Vg-

1

BM

P-1

3

BM

P-1

2

BM

P-1

4

BM

P-9

DO

RSA

LIN

BM

P-1

0

BM

P-3

BMP-7 100

BMP-5 88 100

BMP-6 87 91 100

BMP-8A 74 74 75 100

BMP-8B 67 67 68 77 100

Vgr-D 69 74 71 63 55 100

UNIVIN 63 62 63 61 58 54 100

BMP-2 60 61 61 55 57 57 67 100

BMP-4 58 59 60 55 56 54 64 92 100

dpp 58 57 59 53 51 54 56 74 76 100

Vg-1 57 56 58 55 56 51 64 58 56 48 100

BMP-13 53 54 53 52 49 48 60 57 56 53 50 100

BMP-12 53 52 52 53 51 48 60 57 57 53 49 86 100

BMP-14 51 52 51 50 48 48 58 57 57 52 52 86 80 100

BMP-9 51 53 53 48 44 47 50 50 50 51 48 52 50 50 100

DORSALIN 49 51 53 47 44 48 47 53 54 53 46 55 56 53 79 100

BMP-10 47 48 48 46 45 48 49 55 52 49 50 57 55 49 63 66 100

BMP-3 42 43 44 41 41 41 48 48 47 43 49 46 46 47 38 38 39 100

Table 4. Identity matrix for the BMP family, constructed with the members that are more than 42% identical to BMP-7 in their mature domains. Highlighted in gray are groups of sequences having 75% or higher identity, which correspond to the subfamilies. Modified from Griffith DL et al., 1996.

1.2.2. General structure of the BMPs.

The genes encoding the BMP family members are mapped to different chromosomes,

indicating that these proteins have become widely dispersed during evolution (Dickinson ME et

al., 1990). The structure of the BMPs is highly conserved (Fig. 2). They are all produced as

large monomeric pre-pro-proteins formed by a 15-25 residue pre-peptide, a 50-375 residue pro-

domain, and a 110-139 residue mature domain at the C-terminus. The latter contains seven

conserved cystines which determine the formation of the characteristic structural motif of the

Introduction

19

members of the TGF-β superfamily: the cystine knot (McDonald and Hendrickson, 1993). The

cystine knot constitutes the core of the monomer and consists of three intracatenary disulfide

bonds. Two of these bonds form a ring through which the third passes, while the seventh

cystine remains free to form the single intercatenary disulfide bond that allows dimerization of

the molecule. Four strands of antiparallel β-sheets emanate from the knot, forming two finger-

like projections. On the opposite end of the knot, an α-helix lies perpendicular to the axis of the

two fingers, forming the heel of the hand (Griffith DL et al, 1996).

Figure 2. Schematic representation of the BMP-7 monomer, showing the four strands of antiparallel β-sheets forming the two finger-like projections, and the α-helix forming the heel of the hand. The core of the monomer is a cystine knot formed by three disulfide bonds, which are represented as orange lines. The N-linked sugar moiety attached to Asn-80 is represented as a green circle. The N-terminus is unresolved. Modified from Griffith DL et al., 1996.

All BMPs have one or more potential N-glycosylation sites but, in most cases (e.g. BMP-2,

BMP-6, BMP-7), only one of these have an N-linked sugar moiety attached. Glycosylation of the

molecule does not seem to be essential for the biological activity of BMP-2 (Ruppert et al.,

1996; Vallejo et al., 2002; Long et al., 2006), though very recent investigations point to a more

important role for glycosylation in BMP-6 (Saremba S et al., 2008). The pre-peptide is known to mediate translocation of the pre-pro-protein into the lumen of

the endoplasmic reticulum and, thus, secretion. In opposition, the function of the pro-domain is

still unknown. In the case of TGF-β, the pro-domain has been termed latency-associated

peptide (β-1-LAP), since it has been demonstrated to delay the function of the mature growth

factor (Gentry LE and Nash BW, 1990; Böttinger EP et al., 1996). A similar role has been

suggested for the BMP-2 pro-domain, despite its limited sequence homology with β-1-LAP. It

has also been suggested that the pro-domain may mediate oxidative structure formation of

β1 β2

β4 β3

β8 β6

β5 β7

α1

N36

S38

S104

C139

71S

138S

S67

S136

S103

Finger 1 Finger 2

Heel

Asn80

Introduction _

20

BMP-2, though it has been demonstrated that both BMP-2 and pro-BMP2 can be refolded in

vitro with comparable yields from the denatured state (Hillger F et al., 2005).

Dimerization of the molecule occurs after the mature domain of each monomeric precursor

is liberated by the action of a subtilisin-like proprotein convertase (SPC), which cleaves the pro-

domain at a conserved Arg-X-X-Arg maturation site (Akamatsu T et al., 1999; Constam DB and

Robertson EJ, 1999). Once cleaved, the mature domains dimerize and the active dimers are

secreted (Fig. 3) (Kingsley DM, 1994). It has been demonstrated that the dimeric form of the

molecule is the only one with biological activity and capable of triggering the BMP-signalling

pathways (Wang EA et al., 1990).

Figure 3. Schematic representation of the processing of a BMP pre-pro-protein to obtain an active dimer. Proteolytic digestion at the RXXR motif releases the mature domain. Two mature domains refold and dimerize to produce de active form of BMP. The disulfide bonds are drawn as orange lines.

1.2.3. Structure of BMP-6.

BMP-6 was first isolated from a murine embryonic cDNA library and named Vgr-1 due to its

homology with Xenopus Vg-1 (Lyons KM et al., 1989). The human and bovine homologues of

Vgr-1 were subsequently isolated from bone and named BMP-6 (Celeste AJ et al., 1990). The

human bmp-6 gene has been mapped to chromosome 6 (Hahn GV et al., 1992) and its

expression produces a 513-residue pre-pro-protein, formed by a 20-residue pre-peptide, a 354-

residue pro-domain, and a 139-residue mature domain, what makes the BMP-6 the largest

protein of the BMP family. The mature domain of the monomer is liberated after cleavage at an

NH2 RXXR C C C C C C C COOH

Pre-peptide Pro-domain Mature domain (15-25 residues) (50-375 residues) (110-139 residues)

NH2 C C C C C C C COOH

COOH C C C C C C C NH2

Processing, folding and dimerization.

Introduction

21

Arg-Thr-Thr-Arg motif localized in the pro-domain between residues 371 and 374. The seven

cystines implicated in constitution of the cystine knot are localized at sites 38, 67, 71, 103, 104,

136 and 138 of the mature domain. The disulfide bonds Cys67 – Cys136 and Cys71 – Cys138

form the ring structure through which the bond Cys38 – Cys104 passes, while Cys103 forms the

intercatenary bond implicated in dimerization.

BMP-6 is a basic protein, with an isoelectric point (pI) of 8.6 (Celeste AJ et al., 1990). Like

BMP-2 and BMP-7, BMP-6 is considered a strongly hydrophobic molecule, exhibiting large

nonpolar patches among its surface. This results in low solubility and tendency towards

aggregation in aqueous solutions. The N-terminal region of the molecule, preceding the first

cystine, contains 6 arginine residues and possesses a strongly positive net charge. This region,

as is the case for BMP-2 (Koenig BB et al., 1994; Ruppert R et al., 1996), may be responsible

for the affinity to heparin shown by most members of the subfamily.

The crystal structure of BMP-6 has been recently determined to a resolution of 2.1 Å

(Fig. 4) (Saremba S et al., 2008). The results of this study suggest that BMP-6 may adopt two

different conformations and that differences between both conformers are mainly localized in

the prehelix loops, which have been shown to contain the main binding and specificity

determinants for type I receptor recognition in other members of the BMP subfamily (Keller S

et al., 2004; Nickel J et al., 2005).

Each mature monomer of BMP-6 possesses three putative N-glycosylation sequences.

These three sites are also found in the BMP-7 molecule, where the two of them situated at the

N-terminus of the mature domain are not glycosylated (Sampath TK et al., 1992; Jones WK et

al., 1994). In contrast, the third N-glycosylation site, located in the cystine-knot motif, and

which is conserved among the BMP-2/4 and the OP-1/BMP-7 families, does carry carbohydrate

moieties (Groppe J et al., 2002; Greenwald J et al., 2003). Recent studies have revealed that

binding of BMP-6 to type I BMPRs may be strictly dependent on glycosylation at Asn73, since

BMP-6 from CHO cells deglycosylated by N-endoglycosidase F treatment completely lost its

capacity to bind ActR-I (Saremba S et al., 2008).

Figure 4. Three-dimensional representation of the BMP-6 dimer, based on crystallization data at 2.1 Å resolution obtained by Saremba S et al. The two monomers are colored blue and purple, respectively.

Prehelix loop

Prehelix loop

N1

N2

C1 C2

Introduction _

22

1.2.4. BMP signalling.

BMPs, like other members of the TGF-β superfamily, bind to two different types of

serine/threonine kinase receptors, being both type I and type II receptors required for

signalling. These receptors are structurally conserved, comprising an extracellular domain, a

single transmembrane domain, and a large intracellular kinase domain (Kawabata M et al.,

1998). Three type I and three type II receptors have been shown to bind BMPs (Table 5).

TYPE I RECEPTORS TYPE II RECEPTORS

NAME ALTERNATIVE NAME NAME ALTERNATIVE NAME

ActR-I ALK-2 ActR-II none

BMPR-IA ALK-3 ActR-IIB none

BMPR-IB ALK-6 BMPR-II none

Table 5. The BMP receptors (ten Dijke et al., 1994; Koenig et al., 1994; Yamashita H et al., 1996; Macias-Silva et al., 1998).

To initiate the signalling cascade, the ligand first binds two copies of its high-affinity

receptor, after which two copies of the lower-affinity receptor are able to bind, forming a six-

polypeptide chain complex (two monomers of the ligand and two pairs of each receptor type).

In this way, specificity in binding BMPR complex appears to be determined by the type I

receptor, being the type II receptor more important for the activation of signal transducing

mechanisms (Massague J, 1998). To promote osteoblast differentiation, BMP-6 binds ActR-I

with high affinity, using BMPR-II or ActR-II as lower-affinity receptors to form the signalling

complex (Ebisawa T et al., 1999).

Once the signalling complex is formed, the constitutively active type II receptors

phosphorylate a glycine/serine-rich domain of the type I receptors. This can lead to activation

of two different pathways: a “canonical” BMP-Smad pathway or a “non-canonical” BMP-MAPK

pathway (Botchkarev VA, 2003). Which pathway is activated seems to depend on the particular

mechanism of oligomerization of the BMPR complex (Nohe A et al., 2002).

When signal transduction occurs through the BMP-Smad pathway (Fig. 5), the activated

BMPR type I phosphorylates receptor-activated Smad proteins (Smad1, 5 and 8; also named

R-Smads) which can then form heteromeric complexes with a common-partner Smad (Smad4;

also named Co-Smad). This R-Smad/Co-Smad complex subsequently translocates to the

nucleus to regulate transcription of BMP responsive genes. Depending on which coactivator(s)

or corepressor(s) interact with the R-Smad/Co-Smad complex, this regulation will be negative or

positive (ten Dijke et al., 2002). In the particular case of BMP-6, the induction of osteoblastic

Introduction

23

differentiation of human MSCs seems to be mainly mediated by Smad5 and, to a lesser extent,

by Smad1 phosphorylation, with no apparent implication of Smad8 (Ebisawa T et al., 1999).

Inhibitory Smads (Smad6 and 7; also named I-Smads) antagonize the phosphorylation of

R-Smads by BMPR-I kinases (Imamura T et al., 1997). This antagonistic effect of Smad6 may

be blocked by AMSH, which directly binds to this I-Smad to prevent its interaction with the

R-Smads (Itoh F et al., 2001). Other inhibitors of this signalling pathway are Tob, which

interacts specifically with BMP activated Smads (Yoshida Y et al., 2000) and Smurf1, which

mediates the degradation of Smad 1 and 5 (Zhu H et al., 1999). Smurf1 also recognizes the

bone-specific transcription factor Runx2 to mediate its degradation (Zhao M et al., 2003) and

can form a complex with Smad6, which can be exported from the nucleus and targeted to the

type I BMPRs to promote their degradation (Murakami G et al., 2003).

Figure 5. Representation of the “canonical” BMP-Smad signalling pathway. Modified from Botchkarev VA, 2003 and from Chen D, 2004.

P P

PP

BMPR-II BMPR-I

BMP

Soluble antagonists (Noggin, Chordin,

Follistatin, DAN, Gremlin, Cerberus, Caronte).

R-SmadP

R-Smad

Smad6

Smurf1

I-Smad

Smurf1

TobR-Smad

P

R-Smad P

Co-Smad

AMSH

Co-activator / Co-repressor

R-SmadP

Co-Smad

Transcription factor

Smurf1

+/-

Cell membrane

nucleus

Smad6

Smurf1

BAMBI

Introduction _

24

The “non-canonical” BMP-MAPK pathway (Fig. 6) is triggered when the activated BMPR

complex interacts with the apoptosis inhibitor XIAP, which links BMP receptors with TAB1. The

latter, in turn, activates TAK1 (Yamaguchi K et al., 1999), which is a member of the MAPK-

kinase-kinase family. TAK1 subsequently activates NLK, which has been shown to inhibit

phosphorylation of TCF-1/Lef-1 transcription factors and downregulate Wnt/β-catenin-

dependent transcription (Ishitani T et al., 1999). It has also been shown that TAK1 can activate

p38 and JNK pathways, which are both involved in BMP-induced apoptosis (Kimura N et al.,

2000; Zhang D et al., 2000).

This signalling pathway might be linked at different levels with the BMP-Smad pathway,

since it has been shown that the I-Smad Smad6 is able to bind to TAK1 and inhibit its activity

(Kimura N et al., 2000).

Figure 6. Representation of the “noncanonical” BMP-MAPK signalling pathway. Modified from Botchkarev VA, 2003.

Different soluble antagonists, such as noggin, chordin, follistatin, or members of the

DAN/cerberus family, can negatively regulate BMP signalling at the extracellular level. These

P P

P P

BMP

Cell membrane

BMPR-II BMPR-I

nucleus

XIAP

TAB1

TAK1

NLK

β-Catenin

TCF

Smad6

p38 and JNK pathways

APOPTOSIS

Introduction

25

proteins bind to BMPs with different affinities, preventing them from binding to their receptors

(Zimmerman et al., 1996; Piccolo et al., 1996; Patel K, 1998; Massague J and Chen YG, 2000;

Gazzerro E and Canalis E, 2006). The insoluble type I BMPR-like protein BAMBI can also inhibit

BMP signalling by binding ligands without triggering an intracellular signal cascade (Onichtchouk

D et al., 1999).

As happens with other soluble growth factors, structural components of the extracellular

matrix may interact with the BMPs, preventing them from degradation and keeping a solid pool

of morphogens. This would augment the half-life of these molecules and ensure a slow and

controlled liberation of them into the extracellular milieu (Reddi AH, 2000).

1.2.5. Biological activity of BMPs.

BMPs are pleiotropic growth factors, which exert many different functions during both

development (organogenesis) and the adult life (tissue regeneration and renewal and wound

healing).

In development, the BMPs are implicated in the establishment of both the dorsoventral axis

and the left-right asymmetry during the early stages of ontogenesis (Piedra ME and Ros MA,

2002). They also participate in the organization of the embryonic ventral mesoderm and in the

development of almost all the tissues and organs, including the nervous system, heart, lungs,

kidneys, skin and gonads, controlling cell proliferation, differentiation, migration, apoptosis and

cell to cell adhesion (Lyons KM et al., 1991; Dale L et al., 1992; Hogan BL, 1996).

The lack of function of one or more BMPs, their receptors or proteins involved in their signal

transduction can cause premature death of the embryo due to incorrect development of meso-

ecto- and endodermic derivates (Hogan BL, 1996; Whitman M, 1998, Itoh S et al., 2000). All

knock-out mice for every BMP showed a mutant phenotype, indicating that each BMP play a

specific role during development and morphogenesis of one or more tissues, though some

functional redundancy can be observed among some members of the BMP family (Zhao GQ,

2003, Chen D et al., 2004).

The BMP activities during development are regulated by gradients of antagonists in the

extracellular space, although cellular responses to BMPs also depend on other issues, such as

the specific BMPR subtypes, the stage of differentiation of the target cell, other inhibitory or

stimulatory factors, the stage of development of the organism, etc (Yamamoto and

Oelgeschläger, 2004).

In the adult, the BMPs seem to be implicated in the regeneration of many tissues and in

protection and recovering after injuries. In this sense, it has been shown that BMP-7 expression

Introduction _

26

is decreased in various models of renal disease and that administration of BMP-7 can improve

the renal function after experimental kidney injury (Nguyen TQ and Goldschmeding R, 2008). It

is also well-known that BMPs act as neuroprotective agents in the central nervous system. For

example, BMP-6 expression increases in several regions of the central nervous system after a

mild ischemic damage, with this growth factor being apparently released from neurons into the

interstitial space at the cerebral cortex, hippocampus and cerebellum, suggesting a possible

regulation of neuronal resistance to insults (Martinez G et al., 2001). These results were

supported by studies in which BMP-6 attenuated the negative effect of H2O2 on primary cortical

cultures in vitro, and showed neuroprotective effects against ischemic injury in adult rats after

transient right middle cerebral artery ligation. This BMP-6-mediated neuroprotection seemed to

be through inhibition of apoptotic pathways and similar effects have been attributed to BMP-7

(Wang Y et al., 2001; Chang CF et al., 2002; Chou J et al., 2006). Other studies have

demonstrated that BMP-6 and/or BMP-7 act as neurotrophic factors for different cells of the

CNS (Gratacos E et al., 2002; Yabe T et al., 2002).

Nevertheless, the most studied and striking feature of BMPs is their role in the homeostasis

and regeneration of the skeletal system, being some BMPs the only growth factors known to

have the capacity to induce ectopic bone formation in adult vertebrates (Wang EA et al., 1990;

Volek-Smith H and Urist MR, 1996; Nakase T and Yoshikawa H, 2006).

It has been demonstrated that BMP signalling is necessary during all stages of osteoblastic

differentiation, which include proliferation, matrix formation, matrix maturation and

mineralization (Stein GS and Lian JB, 1993; van der Horst G et al., 2002). Furthermore, a study

carried out by Cheng H et al. using fourteen human BMPs showed that different BMPs act at

different stages during osteoblastic differentiation, with BMP-2, -6 and -9 exhibiting the greatest

ability to induce both early and late osteogenic markers, as well as matrix mineralization (Cheng

H et al., 2003). Their results led these authors to propose a model of osteogenic hierarchy of

the BMPs, in which BMP-2, 6 and -9 may be the most potent agents to induce lineage-specific

differentiation of mesenchymal progenitor cells, while most BMPs can promote terminal

differentiation of committed osteoblastic precursors and osteoblasts (Fig. 7).

Figure 7. Hierarchic model of BMP-induced osteoblastic differentiation. Modified from Cheng H et al., 2003.

Multipotential cell Osteoprogenitor Osteoblast Osteocyte

BMP-2, -6, -9 BMP-2, -4, -7, -9Most BMPs

(except BMP-3 and -12)

Introduction

27

1.2.6. Biological activity of BMP-6 in osteogenesis.

The BMP-6 protein is predominantly expressed in hypertrophic chondrocytes during

endochondral ossification (Lyons KM et al., 1989), and is known to stimulate expression of both

chondrogenic and osteogenic phenotypes in vitro (Gitelman SE et al., 1994; Yamaguchi A et al.,

1996) and to induce cartilage and bone formation in vivo (Gitelman SE et al., 1994).

The comparative analysis carried out by Cheng H et al. using adenovirus-mediated gene

transfer of BMPs to mesenchymal progenitor and osteoblastic cells revealed that BMP-6,

together with BMP-2 and -9, had the greatest ability to induce both early and late osteogenic

markers (ALP activity and osteocalcin, respectively), as well as matrix mineralization (Cheng H

et al., 2003). In a similar way, horse bone marrow derived mesenchymal stem cells transduced

with BMP-6-expressing adenoviruses achieved osteogenic differentiation attending to ALP

activity and mineralization levels (Zachos TA et al., 2006). When BMP-6 was administered to

human MSCs, these cells underwent drastic osteogenic differentiation, with bone-associated

gene and protein expression (Osterix, DLX-5, type I collagen and bone sialoprotein II),

extracellular matrix (ECM) mineralization, and hydroxyapatite formation at higher levels than

cells in presence of other BMPs (Friedman MS et al., 2006).

Despite this demonstrated osteogenic potential of BMP-6, surprisingly BMP-6 knock-out

mice are largely unremarkable, with exception of a delayed sternum ossification. Since the

expression of BMP-6 during embryogenesis is closely coupled with that of BMP-2, the lack of

other defects in BMP-6-deficient mice is thought to be due to functional compensation by BMP-2

(Solloway MJ et al., 1998).

It has been shown that addition of sulphated polysaccharides such as native heparan

sulphate and synthetic dextran sulphate to undifferentiated mesenchymal cells in vitro

enhances BMP-2-mediated osteoblastic differentiation. In contrast, the osteogenic activity of

BMP-6 seems to be inhibited by these molecules (Zhao B et al., 2006).

The capacity of this growth factor to promote osteogenesis has also been demonstrated in

vivo. Injection of BMP-6-expressing adenoviral constructions into the calf muscles of athymic

nude rats led to ectopic bone formation by way of mechanisms similar to both endochondral

and intramembranous ossification pathways (Jane JA Jr et al., 2002). The same approach has

been used to accelerate healing of rabbit bilateral ulnar osteotomies (Bertone AL et al., 2004).

In another study, BMP-6 was directly applied to periodontal fenestration defects in rats,

resulting in increased bone and cementum formation compared to control groups (Huang KK et

al., 2005).

Introduction _

28

The effectiveness of bone remodelling depends on a balance of bone formation

(osteoblastogenesis) and bone resorption (osteoclastogenesis). A recent study demonstrated

that BMP-6 has a biphasic effect on primary murine bone marrow cells, stimulating osteoclast

generation at low concentrations (1 ng/mL) and osteoblast generation at higher concentrations

(300 ng/mL) (Wutzl A et al., 2005). All these evidences point to the fact that BMP-6 might be

an important regulator of bone homeostasis.

Introduction

29

1.3. Basic fibroblast growth factor.

1.3.1. The fibroblast growth factors.

The fibroblast growth factors (FGFs) are a family of polypeptide growth factors present in

organisms ranging from nematodes to humans. In vertebrates, this family is composed of 22

members, which are highly conserved in both gene structure and amino-acid sequence among

species. FGFs are key players in the processes of proliferation, differentiation, migration and

survival of many cell types during embryonic development, while acting as homeostatic factors

in tissue repair and response to injury in the adult organism.

Among the FGFs, the most studied members are the two first discovered: acidic fibroblast

growth factor, aFGF or FGF1, and basic fibroblast growth factor, bFGF or FGF2, which have

special relevance on wound healing and blood vessel formation, being both more potent

angiogenic factors than VEGF (Ornitz DM and Itoh N, 2001).

1.3.2. Structure of bFGF.

Fibroblast growth factor 2 or basic fibroblast growth factor (FGF2 or bFGF) is one of the

members of the large family of the FGF family, being this cytokine initially purified from bovine

pituitary gland extracts in 1975 (Gospodarowicz D, 1975).

The human genome has one single copy of the fgf2 gene, localized in chromosome 4, and

extending over more than 36 Kbp. It is composed of three exons separated by two 16 Kbp

introns (Abraham JA et al., 1986). Translation using the conventional AUG codon gives rise to

the first-identified 18 KDa bFGF-isoform, while the existence of upstream in-frame CUG codons

allows alternative translation of higher molecular weight isoforms (Fig. 8). These 22, 22.5, 24

and 34 KDa isoforms possess a nuclear localizing sequence (nls) containing several Gly-Arg

repeats with methylated arginine residues, which directs the growth factor to the cell nucleus,

whereas the 18 KDa isoform is essentially cytosolic. The 34 KDa isoform is induced under cell

stress conditions and is poorly translated (Arnaud E et al., 1999).

Introduction _

30

Figure 8. Schematic representation of the bFGF mRNA and the isoforms of bFGF resulting from its alternative translation. Nuclear localizing sequences (nls) present in the high molecular weight isoforms are represented as orange boxes.

In the present work, we focused on the conventional 18 KDa isoform of bFGF, which is a

monomeric protein containing four cysteine residues with no intramolecular disulfide bonds, a

large number of basic aminoacids, and two sites (Ser64 and Thr112) that can be

phosphorylated by protein kinases A and C, respectively. Due to the predominance of basic

residues, bFGF has a pI of 9.6, being a pH range from 5 to 9 optimal for its stability in aqueous

solutions (Bikfalvi A et al., 1997). Crystallization of bFGF revealed that it consists of twelve anti-

parallel β-sheets organized into a trigonal pyramidal structure (Fig. 9). The region of the

molecule involved in receptor-binding is mainly localized between residues 40 and 100, whereas

the C-terminus is implicated in specific binding to heparan sulphate proteoglycans (HSPGs)

mediated by two lysine-rich surface loops (Asn28, Arg121, Lys126, Gln135, and Lys27, Asn102,

Lys136) (Eriksson AE et al., 1991).

1.3.3. bFGF secretion.

Soluble proteins destined for secretion use to possess N-terminal signal peptides to direct

their translocation into the lumen of the endoplasmic reticulum. From here they can be

packaged into transport vesicles and reach the Golgi apparatus, being finally sent to the cell

surface through vesicular transport. bFGF lacks such a signal peptide and its secretion to the

extracellular space seems to be unconventional, since it does not follow this classic secretory

pathway. It has been suggested that bFGF is released from cells as the result of cell damage,

death and non-lethal membrane disruptions (Conrad HE, 1998). Nevertheless, more recent

studies suggest that bFGF is secreted by direct translocation through the plasma membrane in

CUG 319 CUG 346 CUG 86 CUG 319 AUG 486 Stop 951

5’ 3’ mRNA

Translation

18 KDa

22 KDa

22.5 KDa

24 KDa

34 KDa

Introduction

31

an ATP- and membrane potential-independent manner (Backhaus R et al., 2004; Schäfer T et

al., 2004). It also seems that cell surface HSPGs act as a molecular trap which drives directional

transport of bFGF to the extracellular milieu, since C-terminus truncated bFGF forms are unable

to be secreted and cells lacking functional surface HSPGs do not secrete bFGF (Zehe C et al.,

2006).

1.3.4. bFGF receptors and bFGF-binding molecules.

Extracellular bFGF signals via an autocrine or a paracrine mechanism involving high affinity

transmembrane receptors derived from four separate genes (FGFR1-4). The existence of splice

variants for each receptor type results in differing ligand binding domains, which confer

specificity in signalling in response to the various FGF family members. Although all the

different splice variants of the four FGFRs can be activated by aFGF (acidic fibroblast growth

A B

C D

Figure 9. Three-dimensional representations of the bFGF molecule, based on crystallization data at 2.2 Å resolution obtained by Eriksson EA et al. A) Model of the bFGF molecule showing the twelve β-sheets. B) Wireframe model with the basic residues highlighted in blue and the acidic residues highlighted in red. C) Wireframe model highlighting the residues of the C-terminal surface loops implicated in binding to HSPGs. D) Wireframe model highlighting the central region of the molecule involved in receptor-binding.

Introduction _

32

factor), most of them have narrower specificity for the different FGF ligands (Kan M et al.,

1993). The FGFR2 consists of extracellular immunoglobulin (IG) domains, a transmembrane

domain, and intracellular tyrosine kinase (TK) domains (Fig. 10). The exon encoding the

C-terminus region of the IG loop3 undergoes alternative splicing to generate the IIIb and IIIc

isoforms of FGFR2, being bFGF only capable of binding to FGFR2-IIIc. In particular, bFGF binds

to both the extracellular IG loop2 and the interloop region of IG loop2 (McKeehan WL, 1992).

bFGF has specific affinity to heparin and heparan sulphate, which are linear sulphated

polysaccharides known as glycosaminoglycans. While heparin is only produced by mast cells,

heparan sulphate is present in all mammalian tissues attached to core proteins as heparan

sulphate proteoglycans (HSPGs), which are perlecan, syndecan, betaglycan, CD44 isoforms, and

glycosylphosphatidylinositol-linked forms, such as glypican and cerebroglycan. The binding to

these molecules protects bFGF against heat or acid denaturation, as well as protease cleavage,

maintaining an extracellular pool of active bFGF (Conrad HE, 1998). Binding between bFGF and

HSPGs also increases the binding affinity of bFGF to its receptors, being the basal lamina HSPG

perlican the major activator of bFGF. HSPG density within the extracellular matrix will dictate

bFGF transport and release rates, with cells competing for matrix released bFGF depending on

the density of cell surface binding sites (FGFR and membrane-associated HSPGs). The

magnitude and type of cellular response might depend on the ability to form ternary complexes

of bFGF, HSPGs and FGFR. These high-affinity complexes will have slower dissociation rates and

will thus show increased persistence on the cell surface and within the intracellular

compartments (Folkman J et al., 1988; Dowd CJ et al., 1999).

Figure 10. Schematic representation of the FGF Receptor.

bFGF

Signal peptide

IG Domain I loop 1 Acid box IG Domain II loop 2

IG Domain III loop 3

Transmembrane domain cell membrane

Tyrosine kinase domain I Tyrisone kinase domain II C-tail

Heparin-binding domain

Introduction

33

1.3.5. bFGF signalling.

bFGF signals through two major transduction pathways: the mitogen-activated protein

kinase (MAPK) pathway, and the protein kinase C (PKC) pathway (Fig. 11). Binding of bFGF to

its receptor leads to autophosphorylation of five tyrosine residues (Tyr586, Tyr656, Tyr657,

Tyr733 and Tyr769) of the TK domains, following dimerization of two FGFRs. Some of the

phosphotyrosines are binding sites for proteins containing phosphotyrosine-binding domains,

such as FGF receptor substrate 2 (FRS2) and Scr homologous and collagen protein (SHC).

These two proteins act as docking molecules that recruit a complex formed by growth factor

receptor-bound protein 2 (GRB2) and son of sevenless (SOS). The GRB2-SOS complex activates

the small G-protein RAS, which recruits and activates RAF, a serine/threonine kinase that

phosphorylates MEK. Activated MEK phosphorylates MAPK, which can translocate to the nucleus

to directly activate transcription factors by phosphorylation. On the other side, some

phosphotyrosines of the TK domain of FGFR can bind to the SH2 domain of phospholipase C

gamma (PLCγ) which, when activated, will cleave phosphatidyl-inositol-4, 5-bisphosphate (PIP2)

to diacylglycerol (DAG) and inositol triphosphate (IP3). The latter induces calcium liberation

from the endoplasmic reticulum, while DAG activates PKC in the presence of calcium.

The specific transduction pathways and molecules involved in bFGF signalling can depend

on the particular FGFR type activated, as well as on the type of cell surface HSPGs implicated in

receptor activation and direct activity of intracellular and intranuclear bFGF. Due to this

complexity, activation of various cell types by bFGF can lead to a variety of cellular responses,

such as proliferation, migration and/or stimulation/inhibition of the expression of a certain

phenotype (Nugent MA and Iozzo RV, 2000).

1.3.6. bFGF biological activity.

Though bFGF was first described as a mitogen for fibroblasts, it plays a key role in

development, remodelling and regeneration of almost every organ (Bikfalvi A et al., 1997). Most

of the reported activities are for the 18 KDa isoform, which is widely distributed in many

tissues. Anyway, as evidenced by knock-out experiments, no vital function is absolutely

dependent on bFGF, probably due to redundant functions of several FGFs triggering the same

FGFR variants. bFGF(-/-) mice lacking all the growth factor isoforms are viable, fertile and

phenotypically almost identical to wild-type mice, for except of some neuronal defects in the

cortex and reduced blood pressure due to an impaired neural reflex control of the vascular tone

(Dono R et al., 1998).

Introduction _

34

Nevertheless, bFGF is known to play important roles during the embryonic development of

vertebrates. For example, it has been demonstrated that bFGF acts synergically with BMPs to

induce controlled cell death during avian limb development (Montero JA et al., 2001)

Figure 11. Representation of the two major pathways for bFGF signalling: The MAPK and the PKC pathways.

One of the best characterized activities of this growth factor is the regulation of growth and

function of vascular cells, such as endothelial and smooth muscle cells. bFGF has been

implicated in angiogenesis and in the pathogenesis of vascular diseases, such as

atherosclerosis. Due to this feature, bFGF has been related with tumour progression and

metastasis (Basilico C and Moscatelli D, 1992), and a secreted FGF-binding protein (FGF-BP)

that mobilizes and activates locally stored FGF has been reported to act as an angiogenic switch

in human cancer (Czubayko F et al., 1997).

bFGF

PP

P

P

P

P

FRS2

GRB-2 SHC PLCγ

SOS

RAS GTP

RAF GDP+Pi

MEK

MAPK

MAPK

PIP2

IP3 + DAG

Ca2+

release

PKC

bFGF

bFGFFGFR

HSPG

Cell membrane

nucleus

Jun, Fos

Introduction

35

In bone regeneration, bFGF might play a double role. Its well known angiogenic activity can

stimulate neovascularization of the new formed bone (Lind M, 1998), while it might also be

important to induce proliferation and/or differentiation of mesenchymal osteoprogenitor cells.

The invasion of new blood vessels into the newly forming bone or bone grafts is considered

a source of potential osteoprogenitor cells. These blood vessels, growing between the neo-

formed woven-bone trabeculae, not only provide the nutrients for the new bone, but also form

the haematopoietic bone marrow, which is the origin of the osteoclasts responsible for bone

resorption (van de Wijngaert FP et al., 1987). In a later stage, this woven bone is remodelled

and replaced by mature lamellar bone. Stimulation of capillary invasion will lead to more bone

formation at the site of the defect.

On the other hand, there is a great controversy around the effect of bFGF on the

osteogenic cells themselves. Several in vitro studies revealed inhibition of cartilage and bone

differentiation by bFGF. In cultured chondrocytes, terminal differentiation was inhibited by

bFGF, since alkaline phosphatase activity, calcium deposition (Kato Y and Iwamoto M, 1990;

Iwamoto M et al., 1995) and proteoglycan synthesis (Demarquay D et al., 1990) was

diminished. This effect seems to be due to induction of parathyroid hormone-related protein

(PTHrP) expression (Terkeltaub RA et al., 1998). Similar inhibitory effects have been observed

in cultured osteoblastic cells, in which bFGF decreased the steady-state mRNA levels for

osteocalcin, type I collagen and alkaline phosphatase (Rodan SB et al., 1989; Hurley MM et al.,

1993; Delany AM and Canalis E, 1998). Also several in vivo studies revealed inhibitory effects of

bFGF on bone formation. It has been shown that exogenous bFGF did not accelerate fracture

healing in rabbits (Bland YS et al., 1995), while administration of this growth factor diminished

mechanical strength during repair of tibial segmental defects in rats (Andreshak JL et al., 1997).

Administration of bFGF mixed with bone matrix powder in hamstring muscles of mice showed a

dose-dependent inhibition of heterotopic endochondral bone formation (Sakano S et al., 2002).

In opposition to these evidences, many other studies have revealed important stimulatory

effects of bFGF on bone formation. In vitro, bFGF was shown to activate the transcription of the

human osteocalcin gene in ROS 17/2.8-transfected cells (Schedlich LJ et al., 1994), as well as it

enhanced the growth, expression of osteogenic markers (alkaline phosphatase and osteocalcin)

and formation of mineralized bone-like tissue in rat and human stromal bone marrow cells in

the presence of dexamethasone (Pitaru S et al., 1993; Pri-Chen S et al., 1998). In vivo, the

administration of low doses of this growth factor to growing rats stimulated endosteal and

endochondral bone formation, preceded by an initial increase in preosteoblastic cells from which

osteoblast were recruited (Nagai H et al., 1995). When administered together with

demineralized bone matrix powder, bFGF greatly enhanced reparation of mandibular critical-size

defects in rabbits (Lu M and Rabie AB, 2002). Direct infusion of 100 ng/day of bFGF into rat

femora after bone marrow ablation caused an increase in mRNA levels of osteopontin, but

Introduction _

36

decreased the expression of type I collagen, while higher doses inhibited gene expression of

osteogenic markers (Tanaka H et al., 2003). It has also been shown that intravenous

administration of bFGF to ovariectomized rats results in a significant increase in bone formation

and upregulation of transforming growth factor beta (TGF-β) and insulin-like growth factor-I

(IGF-I) expression (Power RA et al., 2004).

Probably, these contradictory observations just reflect the complexity of signalling by bFGF.

The effect of this growth factor is highly unpredictable depending on the concentration of

effective molecules, the stage of differentiation of the target cells and the presence of other

growth factors. Biphasic dose-dependent responses are common in growth factor biology, and

have already been reported for bFGF, as it enhanced bone formation in bone grafts (Wang JS

and Aspenberg P, 1996) and in a mandibular defect model (Zellin G and Linde A, 2000) at low

doses, while inducing fibrous tissue formation at higher doses. This kind of effect might be

explained by downregulation of FGF receptors in response to an excess of ligands (Moscatelli D,

1994), resulting in reduced bFGF action. It also seems that the bFGF effects are differentiation

stage-specific, as it stimulated cell growth and reduced the expression of osteoblast markers in

less mature human calvaria cells, whereas it induced OC production and matrix mineralization in

more mature osteoblasts (Debiais F et al., 1998). Other authors report that bFGF stimulates

proliferation of immature osteoblasts, but induces apoptosis in differentiating cells (Mansukhani

A et al., 2000). At last, bFGF activity in vivo may be modulated by a variety of other growth

factors with opposite activities or partially overlapping signalling pathways. Simultaneous

presence of bFGF and strong osteogenic molecules such as BMPs or IGF-I might cause a

conflict leading to inhibition of osteoblastic differentiation or even cell death, rather than

inducing osteogenesis. It has been reported that FGFs inhibit BMP receptor 1b (Merino R et al.,

1998) and FGFR3 seems to be a negative regulator of bone growth, since up-regulation of its

signalling represses hedgehog signalling and BMP-4 expression, leading to inhibition of

endochondral bone growth (Deng C et al., 1996; Naski MC et al., 1998). All these evidences

point to the fact that osteoblast responses are regulated by relative strengths of opposing

signalling pathways.

Introduction

37

1.4. Therapies for bone defect healing.

For the treatment of non-union fractures, autologous bone grafts are still considered the

gold standard nowadays, since they possess both important osteoconductive and osteoinductive

properties. The term osteoconduction refers to the process that supports the ingrowth of

capillaries, perivascular tissue and osteoprogenitors into the three-dimensional structure of the

graft, while osteoinduction is the process that supports the proliferation of undifferentiated

mesenchymal cells and the formation of osteoprogenitors with the capacity to form bone

(Bishop GB and Einhorn TA, 2007). Nevertheless, due to the limitations of autografts,

commented in section 1.1.4, the search for alternatives to autologous bone grafts has become

a need.

Human demineralized bone matrix (DBM) is commercially available but, when used alone,

has failed to demonstrate equivalent efficacy to autologous bone (Finkemeier CG, 2002).

Furthermore, the use of DBM mixed with CaSO4 in treatment of non-union fractures showed

high rates of wound drainage, infection and treatment failure (Ziran BH et al., 2007). On the

other hand, synthetic grafts, such as calcium phosphate, calcium sulphate, calcium

hydroxyapatite and collagen-calcium phosphate composites, can mimic the osteoconductive

properties of bone grafts, but fail in their osteoinductive properties.

Thanks to the recombinant DNA technology, many growth factors involved in osteogenesis,

angiogenesis and wound healing have become commercially available, and their potential use in

clinical bone repair is widely being studied. Combinations of an osteoconductive biomaterial and

osteoinductive growth factors have arisen as very promising alternatives to autografts.

1.4.1. Growth factors for bone defect healing.

Among the wide variety of growth factors involved in bone homeostasis, the BMPs have

especially focused the attention of the researches, because of their strong osteogenic properties

and of being the only cytokines known to induce ectopic bone formation.

The use of recombinant human BMP-2 for the treatment of open tibial fractures was

investigated by the BESTT trial (the BMP-2 Evaluation in Surgery for Tibial Trauma) (Govender

S et al., 2002) and by a subgroup analysis (Swiontkowski MF et al., 2006). These studies

concluded that patients treated with 1.50 mg/Kg rhBMP-2 showed fewer hardware failures,

fewer infections and faster wound healing than patients in the control groups. These studies

finally led, in July 2002, to approval of the use of rhBMP-2 (InductOs®) by the European

Medicines Agency (EMEA) for treatment of severe tibial fractures in adults. A few months later,

in November 2002, the American Food and Drug Administration (FDA) approved the use of

Introduction _

38

rhBMP-2 in combination with absorbable bovine type I collagen sponges (INFUSE® Bone Graft

Device) for treatment of open fractures in long bones. rhBMP-2 has also been approved by the

FDA for use in spinal fusions, in the form of a cylindrical titanium fusion cage filled with

rhBMP-2/collagen sponge (InFuse® Bone Graft/LT-CAGE® Lumbar Tapered Fusion Device). This

approach has been proven to be effective to achieve anterior inter-body fusion in patients with

degenerative lumbar disc disease (Boden SD et al., 2000; Burkus JK et al., 2002, 2003, 2004).

Also rhBMP-7 for the treatment of tibial non-unions was investigated by a prospective,

randomised clinical trial, concluding that rhBMP-7 implanted with a type I collagen carrier is a

safe and effective alternative to autologous bone grafting for treatment of tibial non-unions

(Friedlaender GE et al., 2001). On this basis, the FDA issued a Humanitarian Device Exemption

for the application of BMP-7 implants (OP-1® Implant) in recalcitrant long bone non-unions

where autografts are unfeasible and alternative treatments had failed. Similarly, the EMEA

approved the use of Osigraft® for the same purposes. Since then, different clinical studies of

resistant tibial non-unions treated with rhBMP-7 have been published (Pecina M et al., 2001,

2003). In April 2004, the FDA also approved the use of a combination of rhBMP-7, bovine type I

collagen and carboxymethylcellulose (OP-1® Putty) for posterolateral spinal fusion after failure

of alternative treatments. This decision was made based on data obtained from previous

preclinical studies in dogs and clinical pilot studies (Cook SD, 1995; Vaccaro AR et al., 2002,

2004, 2005).

On the other hand, angiogenesis is known to play a critical role in both the systematic

growth and repair of bone, and the combination of angiogenic and osteogenic factors is thought

to enhance bone healing and regeneration (Kanczler JM and Oreffo RO, 2008). In this sense,

many in vivo studies have revealed that bone defects co-treated with bFGF and BMP-2 showed

improved healing when low concentrations of bFGF were used (Fujimura K et al., 2002;

Nakamura Y et al., 2005; Tanaka E et al., 2006; Kakudo N et al., 2006). In these studies, the

positive effect of bFGF might not only be due to enhancement of angiogenesis, but also to

stimulation of proliferation and/or differentiation of mesenchymal osteoprogenitors at the

implant site.

1.4.2. Safety of the clinical use of growth factors.

The clinical studies carried out with rhBMP-2 concluded that the effective osteoinductive

dose of this growth factor is 1.5 mg BMP-2 / mL ACS (Valentin-Opran A et al., 2002; Govender

S et al., 2002). Nevertheless, concentrations in the order of just hundreds of nanograms per

millilitre are sufficient to induce osteoblastic differentiation of mesenchymal cells in vitro while,

in the human body, normal concentrations of BMPs are estimated at 2 ng/g of bone

Introduction

39

(Rengachary SS, 2002). Thus, clinical application of BMPs implies raising their local

concentration more than 106-fold over the physiological levels.

It has been shown that, after administration of rhBMP-2, the amount of growth factor that

can be found in the systemic blood stream is about 0.1% of the used dose, and that these

molecules have a half-life of just a few minutes. Furthermore, neither ectopic ossification nor

calcification of soft tissues has been reported after clinical application of BMPs (Valentin-Opran

A et al., 2002). Although the use of BMP-2 and -7 is considered save, the long-term effects of

the application of such amounts of these potent, highly pleiotropic growth factors are not well

known. On the other hand, the immune mechanisms triggered upon BMP implantation are not

well defined due to controversy in the literature. It seems that single applications of allogenic

BMPs can promote recruitment of macrophages, lymphocytes and plasma cells, as well as

activate a moderate production of anti-BMP antibodies (Granjeiro JM et al., 2005). It has been

suggested that, because of these antibody responses, BMPs should not be used in pregnant

women and the repeated use of BMPs should be avoided (Carlisle E and Fischgrund JS, 2005).

Another disadvantage of the use of high doses of growth factors is the enormous economic

cost of the treatments. In the UK, it is estimated that the use of InductOs® for the treatment of

tibial fractures would cost £ 1,790 per fracture additional to the standard treatment and the

total incremental cost of adopting the use of BMPs in the treatment of open tibial fractures is

estimated to be approximately £ 3.5 million per year (Garrison KR et al., 2007).

1.4.3. Osteoconductive carriers.

It has been demonstrated that new bone formation can be achieved by direct application of

BMPs alone (Wozney JM et al., 1990; Einhorn TA et al., 2003). Nevertheless, these approaches

require the use of very high doses of growth factors, since they use to have a short half-life in

vivo and suffer a quick systemic dispersion after their injection. Application of the growth

factors in combination with specific carriers improves their osteogenic abilities (Peel SA et al.,

2003). The aim of the carrier is to retain the growth factors at the wound site, maintaining their

local concentrations, since it has been demonstrated that the bone healing efficiency is

correlated with the prolonged presence of BMPs at the wound site (Woo BH et al., 2001).

Furthermore, the carrier can act as an osteoconductive milieu, permitting its infiltration by

mesenchymal cells and the ingrowth of blood vessels (Peel SA et al., 2003).

It can be concluded that an ideal carrier for bone regeneration purposes should possess the

following qualities (Geiger M et al., 2003): i) biocompatibility, low immunogenicity and

antigenicity; ii) biodegradability with biocompatible components, in predictable manner in

concert with bone growth; iii) adequate porosity for cellular invasion and vascularization;

iv) adequate compressive and tensile strength; v) enhancement of cellular attachment, without

Introduction _

40

inducing soft tissue growth at the bone/implant interface; vi) amenability to sterilization without

loss of properties; vii) affinity to growth factors and host bone; viii) enhancement of osteogenic

activity of the growth factors with a restrictive release of them at an effective dose during a

period coincident with the accumulation of target cells; ix) adaptability to irregular wound site,

malleability; x) availability to surgeon on short notice.

Unfortunately, none of the currently available materials can be considered an ideal carrier.

Many investigations have been performed to test carriers based on inorganic materials

(tricalcium phosphate, HA, titanium, etc.), polymeric materials (polylactic acid, poly-lactic-co-

glycolic acid, poly-lactic acid/polyethylene glycol, etc.), and organic materials (collagen,

demineralized bone matrix, etc.) (Babensee JE et al., 2000; Kirker-Head CA, 2000; Li RH and

Wozney JM, 2001, Williams DF, 2008).

Among the organic carriers, DBM has shown many advantages and excellent BMP-

retention/liberation properties (Peel SA et al., 2003), but fails as a suitable carrier since it is

known to maintain endogenous growth factors, which are not eliminated during the

demineralization process (Blum B et al., 2004). In fact, DBM alone has some capacity of

inducing bone formation (Kale AA and Di Cesare PE, 1995; Hartman EH et al., 2004). Due to

the great amount and variety of growth factors that remains in DBM after demineralization, the

local effects of DBM implants are unpredictable, having the graft possibilities of being rejected

in long term, as frequently happens with allografts when used in bone surgery (Ziran BH et al.,

2007). Furthermore, other studies suggest that DBM contains proteins that can partially block

BMP activity (Behnam K et al., 2006). In contrast, despite its poor biomechanical properties,

collagen is the only carrier approved for clinical application of BMPs due to its high

biocompatibility and biodegradability and low immunogenicity (Hubbell JA, 1995; Friess W,

1998).

Collagen is the main protein of connective tissue in animals, and is considered the most

abundant protein in mammals. Among the 28 different types of collagen, type I is the most

represented in the human body and is found mainly in tendons, endomysium, fibrocartilage,

bone and in scar tissue. Clinical administration of BMPs for bone regeneration is done in

combination with bovine type I absorbable collagen sponges (ACS), which are soaked with the

growth factor before implantation (Valentin-Opran A et al., 2002). It has been shown that this

form of collagen allows proper cell infiltration during new bone formation (Friess W, 1998).

Unfortunately, most growth factors have little natural affinity to collagen. Pharmacokinetic

studies of rhBMP-2 retention/liberation from collagen sponges in vivo showed a rapid initial loss

followed by an exponential liberation of the growth factor (Hollinger JO et al., 1998).

Electrostatic attraction forces, due to the different pI of collagen and BMPs, are believed to be a

major factor controlling the protein-matrix interactions (Geiger M et al., 2003). When testing a

modified rhBMP-2, treated with plasmin to remove positive charges from the molecule, it was

demonstrated that its affinity to collagen diminished, showing a much faster liberation rate from

collagen sponges and, thus, a shorter local persistence. Although this modified growth factor

Introduction

41

exhibited an increased biological activity tested in vitro on cell cultures (Hollinger JO et al.,

1998), it osteogenic activity in vivo was partially lost in comparison to native rhBMP-2 (Israel DI

et al., 1992). According to these facts, it seems that a sustained liberation of BMPs in vivo may

be a critical pharmacokinetic parameter for osteoinduction, with the amount of new produced

bone increasing with the local concentration of the osteoinductive growth factor (Uludag H et

al., 2000, 2001).

1.4.4. Modified growth factors for regenerative medicine.

Most of the problems associated with clinical application of growth factors could be palliated

if these could be specifically retained at the wound site, with a slow and sustained liberation

from their carrier.

Several proteins have natural domains to confer them specific affinity to collagen. For

example, many pathogenic bacteria express virulence factors with collagen-binding properties,

which facilitate adhesion to the extracellular matrix of the host tissues (e.g. the Yersinia

enterolitica adhesin YadA) or its degradation (e.g. Clostridium histolyticum class I collagenase).

In higher organisms, most of the collagen-binding proteins are related with blood coagulation

and wound healing, such as fibronectin, thrombospondin and the von Willebrand Factor (vWF),

all of which are present in blood plasma (Takagi J et al., 1992).

Many different cytokines have already been produced as fusion proteins with some of these

additional domains to confer them specific affinity to cells or components of the ECM, without

loss of their natural biological activity (Table 6).

The collagen-binding domain (CBD) of the bovine vWF has been identified as a decapeptide

with the sequence Trp-Arg-Glu-Pro-Ser-Phe-Cys-Ala-Leu-Ser (Takagi J et al., 1992). This CBD

has already been used to successfully produce a fusion protein with bFGF in E. coli, and fusion

proteins with several members of the TGF-β superfamily, including BMPs (Table 6). All these

proteins showed increased collagen-binding properties without loss of their natural biological

activity. In both the cases of bFGF and BMP-2, the CBD was fused to the N-terminal part of the

growth factor, and the Cys-7 of the CBD was replaced by a methionine to avoid incorrect

disulphide bond formation during their production or posterior manipulation. In the case of

bFGF, the protein was also produced with a 6xHis purification tag and a thrombin cut site to

eliminate this tag after purification (Andrades JA et al., 2001), while this was already avoided

for the production of the collagen-targeted rhBMP-2, which was purified by its natural affinity to

heparin (Visser R et al., 2009).

Introduction _

42

PROTEIN MODIFICATION REFERENCE

HGF Collagen-binding domain of fibronectin Kitajima T et al., 2007

Cell-binding domain of fibronectin Kawase Y et al., 1992

Collagen-binding domain of C. hystolyticum collagenase Nishi N et al., 1998

EGF

Collagen-binding domain of fibronectin Ishikawa T et al., 2001

Collagen-binding domain of C. hystolyticum collagenase Nishi N et al., 1998

Collagen-binding domain of the vWF Andrades JA et al., 2001

bFGF

Fibrin-binding domain Zhao W et al., 2008

TGF-β1 Collagen-binding domain of the vWF Tuan TL et al., 1996

TGF-β2 Collagen-binding domain of the vWF Han B et al., 1997

BMP-3 Collagen-binding domain of the vWF Han B et al., 2002

Fibrin-binding domain Schmoekel HG et al., 2005

Collagen-binding domain of the vWF Chen B et al., 2007a

Collagen-binding domain of C. hystolyticum collagenase Chen B et al., 2007b

BMP-2

Collagen-binding domain of the vWF Visser R et al., 2009

Table 6. Recombinant fusion proteins with additional binding domains to cells or extracellular matrix proteins.

Since collagen is not just the only carrier approved by the EMEA and the FDA for bone

healing applications, but also a main natural constituent of bone, collagen-targeted growth

factors are of special clinical interest. After direct administration in soluble form, these

molecules could be used to augment their local concentrations by direct binding to collagen

fibres at the site of injection. On the other hand, when administered in combination with a

collagenic carrier, the latter would partially retain the growth factors, limiting their actions to

the wound site. These approaches could reduce the concentration of growth factors needed to

achieve tissue regeneration when compared to the use of native molecules, improving the

safety of the treatments and reducing their costs.

Besides their potential use by direct administration, collagen-targeted growth factors might

also be useful for the in vitro selection and amplification of osteogenic populations of bone

marrow derived cells cultured in collagen gels (Andrades JA et al., 1999; Becerra J et al., 2006).

These strategies may help solve the decrease in the osteogenetic capacity of the bone marrow

of aged patients.

Introduction

43

1.5. Escherichia coli as an expression system.

The enterobacterium Escherichia coli has been used for recombinant DNA technologies

since 1973, when Stanley N. Cohen constructed the first recombinant plasmid for heterologous

DNA transcription in this microorganism (Cohen SN et al., 1973). Since then, E. coli has become

probably the most studied prokaryote. The vast knowledge of its structure and metabolism has

led to the development of many different strains, a great diversity of cloning and gene

expression systems, and to optimisation of media and culture conditions.

The main advantages of the use of E. coli as expression system are:

- The costs of expression are much lower than those of other systems.

- Bacteria are easy to manipulate and to maintain in culture.

- Most of the E. coli strains are harmless and require no specific equipment for

manipulation.

- Many different cloning and expression systems are available. For most of them, all

cloning steps can be performed in vitro.

- Very strong expression of heterologous genes (up to 100 mg/L) can be achieved in

E. coli.

On the other hand, the main disadvantages of the use of bacteria are:

- E. coli is unable to carry out most of the posttranslational modifications that are often

required for eukaryotic protein production.

- Production attempts often result in insoluble, unfunctional proteins aggregated as

inclusion bodies.

- The simple, plasmid-based systems, only allow cloning of a limited amount of

heterologous DNA.

1.5.1. Obtaining functional proteins from inclusion bodies.

Inclusion bodies are insoluble protein aggregates formed by deposition of misfolded or

partially folded polypeptides, due to intermolecular interactions between their exposed

hydrophobic patches (Fig. 12). These structures are generated by the failure of chaperones and

proteases to either fold of degrade un- or misfolded polypeptides synthesized at high rates

(Villaverde A and Carrió MM, 2003).

Introduction _

44

Figure 12. Schematic representation of the events that can happen during protein folding. Correct folding is a multi-step pathway (1). Misfolding (2) can lead to protein aggregation (3). Aggregation can also affect correct folding intermediates with exposed hydrophobic patches. The green lines represent the hydrophilic parts of the protein, while the red lines are the hydrophobic patches. Modified from Vallejo LF and Rinas U, 2004.

During heterologous protein expression, the amount of recombinant protein represents

between 50 and 95% of the inclusion bodies, while the remaining percentage is formed by heat

shock proteins (inclusion body proteins A and B; IbpA and IpbB) (Allen SP et al., 1992),

chaperones like DnaK and GroEL (Carrió MM and Villaverde A, 2002), and some other

contaminants, all of which may be passively trapped in the inclusion bodies through their

natural interactions with the polypeptides during their formation (Hart RA et al., 1990; Rinas U

and Bailey JE, 1992).

The formation of inclusion bodies during recombinant protein expression can be seen as an

advantage when considering the high degree of purity of the target protein in the aggregate

fraction and the higher protection against proteolysis compared to the soluble counterpart.

Although inclusion bodies are easy to isolate (directly yielding a protein fraction highly

enriched in the goal protein), many further in vitro manipulation of the recombinant proteins is

needed to obtain them with their biological activity. In general, the strategy for protein recovery

includes four consecutive steps: i) isolation of the inclusion bodies; ii) solubilization of the

aggregated proteins; iii) in vitro refolding of the solubilized protein; iv) purification of the

refolded fraction (Villaverde A and Carrió MM, 2003).

1.5.2. In vitro refolding of proteins.

Inclusion bodies usually consist of inactive proteins, and a more or less complex refolding

process has to be carried out after solubilization to obtain active proteins in their native

conformation. Many different refolding techniques have been developed for successfully

refolding of proteins, with every one of these having specific requirements. The most commonly

applied techniques are (Vallejo LF and Rinas U, 2004):

1a 1b

3

2 3

Introduction

45

I. Direct dilution: This is the simplest refolding procedure, consisting in direct dilution of the

solubilized proteins into a proper refolding buffer. A method to improve the refolding yield of

this technique is the continuous or pulse addition of the denatured proteins.

II. Membrane controlled denaturant removal: These methods consist in the use of dialysis

and diafiltration systems to gradually change from denaturing to native buffer conditions. These

methods use to cause more aggregation during refolding compared to direct dilution, and

refolding yields may be reduced due to non-specific adsorption of the proteins to the

membrane.

III. Chromatographic methods: These techniques are based on denaturing buffer

substitution by refolding buffer inside a size exclusion or hydrophobic interaction

chromatography column.

IV. Matrix-assisted refolding: consisting in attaching the solubilized proteins to a solid

support prior to changing from denaturing to native buffer conditions by one of the above-

mentioned techniques. This method may avoid the intermolecular interactions between the

folding intermediates with tendency to aggregation.

Regardless of the chosen refolding technique, many physical and chemical variables have to

be taken into account to achieve successful refolding of proteins produced as inclusion bodies.

Among these, critical variables are:

I. Temperature: Each protein is thermodynamically stable in a limited temperature range.

Low temperatures can suppress hydrophobic aggregation during refolding, but also slow down

the refolding rate.

II. Pressure: High pressure can dissolve protein aggregates and inclusion bodies and

gradual depressurization may allow proteins to reach their native state.

III. Chemical additives: Several substances, such as L-arginine, 2-(N-Cyclohexylamino)

ethanesulfonic acid (CHES), sulfobetaines, etc., have shown to suppress aggregation and to

increase refolding yields. Although the way these agents interact with the folding intermediates

remains unclear, they are presumed to diminish aggregation by shielding hydrophobic regions

of the partially folded chains.

IV. Micelles and liposomes: Detergents and phospholipids have shown potential to aid

protein refolding since they can form micelles and liposomes, respectively, with which the

Introduction _

46

folding intermediates can establish transient non-polar interactions to avoid protein

aggregation.

V. Chaperones: Natural chaperones have been used for successful refolding of several

proteins in vitro. The main disadvantages of their application are their high cost and the need

for their removal once the refolding procedure has finished.

Besides the above mentioned general requirements, proteins containing disulfide bonds

have additional special requirements to achieve their native conformation. After their

solubilization in the presence of reducing agents, they have to be refolded under conditions that

allow the formation of their native disulfide bonds, having in mind that these proteins are often

very unstable and show a high tendency towards aggregation in their reduced states. Since air

oxidation of the free cysteine residues is slow and often yields mismatched disulfides, a mixture

of low molecular weight thiols in their reduced and oxidized state is usually added to a slightly

alkaline refolding buffer to permit rapid disulfide exchange reactions until the protein reaches

the thermodynamically most stable conformation.

The use of protein disulfide isomerase (PDI) in combination with a redox system has also

been shown to increase the refolding yields and/or rates of several disulfide-bonded proteins.

This protein is a folding catalyst known to participate in disulfide bond formation in vivo.

In any case, the best refolding conditions for each particular protein have to be empirically

determined, what is usually not a simple task.

Introduction

47

1.6. Baculoviruses as expression systems.

1.6.1. General information on baculoviruses. The baculovirus life-

cycle.

The baculoviruses are a family (Baculoviridae) of rod-shaped viruses that is divided into two

genera:

- The Granuloviruses (GVs) have only one nucleocapsid per envelope. They produce

granulin occlusion bodies, containing one single virion.

- The Nucleopolyhedroviruses (NPVs) contain one (SNPVs) or multiple (MNPVs)

nucleocapsids per envelope. They produce polyhedrin occlusion bodies (also called

polyhedra), which contain multiple embedded virions.

Baculoviruses infect many different invertebrates, being over 600 host species identified,

mainly larval stadiums of moth species. They are not known to replicate in vertebrate cells.

Nucleopolyhedrovirus infection is composed of two infection cycles (Fig. 13): The infection

begins when a healthy host-larva ingests the polyhedra released from an earlier host. These

polyhedra reach the alkaline environment of the midgut, where they are dissolved. The released

viruses will then fuse with the columnar epithelial cell membranes of the host intestine to

trigger primary infection. Viral transcription and replication occur in the cell nucleus, being two

types of progeny produced: a budded virus form and an occluded virus form.

The budded viruses collect cell membrane material as well as binding proteins when

budding through the basolateral cell membrane, being responsible for secondary infection

through an endocytosic process, spreading infection from the midgut cells to the rest of the

larva.

The occluded viruses accumulate as polyhedra in the animal during infection. Cell lysis and,

ultimately, disintegration of the larva will release the polyhedra to the environment to start a

new infection cycle. In their occluded form, baculoviruses are very resilient to environmental

factors and can survive for extended periods of time.

http://en.wikipedia.org/wiki/Cell_nucleus

Introduction _

48

Figure 13. Schematic representation of a typical baculovirus infection cycle.

Attending to protein expression, a typical baculovirus infection can be divided to three

distinct phases, though some genes can be expressed in more than one phase of the replication

cycle. Genes which promoter elements have strong similarity to insect promoters tend to be

expressed early in the cycle, whereas genes with specific viral promoter sequences tend to be

expressed during later phases. These phases are:

- Early phase (0-6 hours post-infection). During this phase, genes involved in the

regulation of the replication cascade and in preventing host responses are expressed,

as well as genes required for DNA synthesis, factors involved in late gene expression,

and a number of genes which modify aspects of the intra- and extracellular

environment.

Polyhedra are disolved in the midgut, liberating occlusion-derived viruses.

Virus fuses with midgut cell (primary infection)

Uncoating

Virogenic stroma Cell nucleus

Budding

Budded virus

Secondary infection

Envelopment and occlusion

Cell lysis and polihedra liberation

Polyhedra are ingested by host

Introduction

49

- Late phase (6-24 hours p.i.). The proteins expressed during this phase are involved in

the shutdown of host cell transcription and translation, in viral DNA packaging, as well

as GP64 (an envelope protein found on the surface of the budded viruses). In the late

phase, the viral DNA is replicated and the nucleocapsids are formed. These

nucleocapsids can either bud out through the cellular membrane and disseminate the

infection within the larva by infecting other cells by GP64-mediated envelope fusion and

endocytosis, or be occluded for horizontal transmission.

- Very late phase (24-72 hours p.i.). In this phase (also called occlusion phase), high

amounts of polyhedrin are produced to occlude the viruses in polyhedra. During the

final stage of this phase, the infected cells are lysed, resulting in death and liquefaction

of the host. P10, a 10 KDa microtubule-associated protein which seems to play a role in

host cell process formation during the infection, is also highly expressed in the very late

phase.

1.6.2. Baculovirus-based expression systems.

Since 1990, baculoviruses are used for expression of recombinant proteins. This is possible

thanks to the existence of high expression promoters in the viral genome, which are non-

essential for virus propagation in vitro. The main advantages of the use of baculoviruses as

expression system are:

- The costs of expression are lower than those when mammalian cells are used.

- Insect cells are easy to manipulate and to maintain in culture.

- Baculoviruses are harmless for vertebrates.

- Baculoviruses have the potential for very strong expression of heterologous genes.

Usual expression levels are in the range of 1-3 mg/L for intracellular proteins, and 3-15

mg/L for secreted proteins.

- Being a eukaryotic system, the baculovirus expression system is capable of proper

protein folding and performing posttranslational modifications, such as signal peptide

cleavage, phosphorylation, N- and O- glycosylation, disulfide bond formation, and

substitution of unusual analogs into proteins (e.g. selenomethionine, heme analogs,

etc.) (Luckow VA, 1991).

- Since the expression is performed at 27 ºC, this system is suitable for expression of

some temperature-sensitive proteins.

- The size of the baculovirus genome (ranging from 80 to 180 Kbp) permits cloning of

several heterologous genes for co-expression.

Introduction _

50

On the other hand, the disadvantages of the use of baculoviruses are:

- The costs of expression are higher than those for expression in prokaryotic systems.

- Expression levels are lower than those achieved with prokaryotic systems.

- The N-glycosylation pathway of baculovirus-infected cells differs from the pathway

found in higher eukaryotes. Glycoproteins produced in the baculovirus system typically

lack complex biantennary N-linked oligosaccharide side chains containing penultimate

galactose and terminal sialic acid residues. This could affect the biological properties of

the produced proteins.

- Extremely high level expression of proteins might overwhelm the ability of the cell to

modify the protein product, being secretion, phosphorylation and glycosylation affected

in particular.

Most of the today available baculovirus expression systems are based on substitution of a

viral non-essential gene by the heterologous gene of interest, maintaining the viral promoter to

control expression. The most commonly substituted genes are those encoding the very-late

expressed proteins P10 and Polh (polyhedrin). When cloned under the polh promoter, the

expression of a heterologous gene achieves maximum levels at 48-72 hours p.i., without

compromising the spreading of the infection among the cultured cells by viral budding.

The main strategy for obtaining a baculovirus expression system consists in cloning the

gene of interest into a shuttle vector, which possesses essential virus sequences flanking each

side of the MCS. On the other hand, the baculovirus DNA is digested with the restriction

enzyme Bsu36I, removing a fragment of an essential gene (ORF1629) for viral replication

(Possee RD et al., 1991) and producing a linear virus DNA that is unable to replicate within

insect cells. Co-transfection of insect cells with the linearized viral DNA and the shuttle vector

containing the gene of interest under the control of the polyhedrin promoter (or other

baculovirus or non-baculovirus promoters) flanked by baculovirus sequences homologous to

those removed by Bsu36I digestion, restores ORF1629 and re-circularises the virus by allelic

replacement (Fig. 14). The recombinant baculovirus DNA is then able to replicate in insect cells

and, in the late phase of infection, virions are assembled and recombinant baculoviruses are

produced. This mechanism ensures that all of the infective viruses are recombinant and that no

wild-type viruses will be produced.

Introduction

51

Figure 14. Schematic representation of the homologous recombination event that gives rise to an infective, recombinant baculovirus.

1.6.3. BacPak6™ and Sapphire™.

The two baculovirus-based expression systems used in the present work were BacPAK6™

(Clontech) and Sapphire™ (Orbigen). Both are designed for expression of recombinant proteins

under control of the polh promoter.

The BacPAK6™ system is a basic baculovirus expression system as explained in section

1.6.2. Recombinant baculoviruses are produced by homologous recombination after co-

transfection of the host cells with a shuttle vector in which the GOI is cloned, and the linearized

(Bsu36I digested) viral DNA, which is 137 Kbp in size (Fig. 15).

The Sapphire™ system is also a basic expression system, though it is improved for the

production of disulfide bond-containing proteins. Besides expression of the cloned GOI,

Sapphire™ baculoviruses co-express the protein disulfide isomerase (PDI) under control of the

p10 promoter (Fig. 15). This protein catalyzes oxidative protein folding in vivo by oxidizing pairs

of cysteines to form disulfide bonds, but can also shuffle incorrect disulfides into their correct

pairings (Gruber CW et al., 2006).

ORF603 ORF1629

GOI

Polh promoter MCS MCS

Shuttle vector with gene of interest (GOI)

Linearized (Bsu36I digested) baculovirus DNA

Infective, recombinant baculovirus DNA

Recombination

Introduction _

52

Figure 15. The BacPak6™ viral DNA and the Sapphire™ viral DNA.

Bsu36IBsu36I

Bsu36I

Bsu36IBsu36I

Bsu36I

PDI

2. Hypothesis and objectives.

53

54

________________________________________________________Hypothesis and Objectives

2.1. Hypothesis.

It has been demonstrated that collagen is a suitable osteoconductive carrier to be used in

combination with rhBMPs, leading to its approval for these purposes in clinical repair of osseous

defects. Nevertheless, the low natural affinity of BMPs to collagen implies that these approaches

require the use of very high concentrations of these growth factors to achieve the desired levels

of osteogenesis at the wound site, augmenting the risks of immune reactions and/or possible

undesired side effects due to diffusion of the molecules to the surrounding tissues or organs, or

into the blood stream. On the other side, bFGF is a potent mitogenic and angiogenic factor and

has been demonstrated to enhance osteogenesis. Nevertheless, the administration of bFGF for

bone repair purposes could also lead to side effects. Furthermore, growth factors are expensive,

and their clinical use significantly increases the costs of the treatments when compared to

standard intervention methods.

Previous studies have shown that the use of a recombinant collagen-targeted hBMP2 fusion

protein in combination with ACS increased osteogenesis at the site of implantation. This so-

called rhBMP2-CBD showed a higher affinity to ACS when compared to native rhBMP-2 and was

able to induce osteogenesis when used at lower concentrations than the threshold established

by other authors for rhBMP-2. This leads to the conclusion that the use of engineered collagen-

targeted growth factors in combination with collagenic carriers may be a better and safer

alternative for clinical repair of osseous defects than the at present available methods.

Among the BMP family members, BMP-6 has been demonstrated to be one of the most

potent inducers of both early and late osteogenic markers. In fact, BMP-6 can induce

osteogenesis at lower concentrations than BMP-2. On the other hand, bFGF is both an

important mitogenic factor for many different cell types, and a potent inducer of angiogenesis.

Several studies have shown that low concentrations of bFGF can enhance BMP-2 mediated

osteogenesis in vivo, but the osteogenic potential of combinations of BMP-6 and bFGF has not

been tested. The aim of the present work is to study this topic. Therefore, the goal is to

produce and purify both rhBMP-6 and rh-bFGF with and without an additional decapeptidic

collagen type I-binding domain derived from the vWF and to test the abilities of these growth

factors to induce osteogenesis in vivo. To avoid major changes to the molecules, the fusion

proteins will be produced without purification tags or any other additional domains. Although

the collagen-binding decapeptide (CBD) present in the vWF contains one cysteine residue, in

the fusion protein this residue will be replaced by a methionine to avoid disulfide scrambling or

unspecific disulfide bond formation during protein production (Fig. 16).

55

Hypothesis and Objectives________________________________________________________

NH2 CBD GROWTH FACTOR COOH

Trp-Arg-Glu-Pro-Ser-Phe-Met-Ala-Leu-Ser

Gly-Ala-Ser

Figure 16. Schematic representation of a recombinant engineered growth factor with a decapeptidic collagen type I-binding domain fused to the N-terminal part of the molecule. The original Cys-7 of the decapeptide has been replaced by a Met. An additional Gly-Ala-Ser tripeptide acts as a link.

We hypothesize that these collagen-targeted growth factors will be retained more efficiently

at the wound site when implanted together with absorbable collagen sponges, due to specific

binding to the carrier. This would allow the use of lower concentrations of these growth factors

to achieve significant levels of osteogenesis. In this sense, the combination of ACS with

collagen-targeted recombinant human bFGF and BMP-6 would show a greater osteogenic

activity than ACS combined with native rh-bFGF and BMP-6. Furthermore, we hypothesize that

the combination of rh-bFGF with rhBMP-6 will enhance osteogenesis when compared to

rhBMP-6 alone.

In conclusion, our hypothesis is that the use of ACS with collagen-targeted recombinant

human bFGF and BMP-6 could be a more effective and safer system for clinical repair of

osseous defects.

56

________________________________________________________Hypothesis and Objectives

2.2. Objectives.

The objectives established for the present work were:

1. To obtain the genes encoding the human bFGF and the human BMP-6, to add the

sequence encoding the collagen binding domain derived from the vWF to these genes

and to clone them into expression vectors.

2. To perform heterologous expression and to purify the native and collagen-targeted

rh-bFGF and rhBMP-6.

3. To determine the affinity to collagen type I of the collagen-targeted growth factors.

4. To characterize the osteogenic activity of the collagen-targeted growth factors in vitro

and in vivo and to compare it with the osteogenic activity of the native molecules.

5. To compare the osteogenic activity in vivo of combinations of bFGF and BMP-6 with

BMP-6 alone.

57

58

3. Material and methods.

59

60

____________________________________________________________Material and methods

61

3.1. Obtaining of the genes encoding h-bFGF and hBMP-6.

The sequence encoding the mature domain of the human bmp-6 gene was obtained by

RT-PCR on total RNA samples isolated from U-2 OS human osteosarcoma cells cultured in vitro.

The isolated RNA sample was previously analyzed by RNA electrophoresis (see Appendix I,

section AI.2.6) to check its quality. The RNA concentration in the sample was calculated from

the OD260 value, and the purity was determined by its OD260/OD280 ratio. Afterwards, the sample

was diluted to 1 µg/mL in highly pure, sterile, RNAse-free water. RNA isolation was performed

as described in Appendix I, section AI.2.1, while RT-PCR was performed with the specific

primers P5 vs. P6 (see Appendix I, section AI.2.13) as described in Appendix I, section AI.2.2.

The sequence encoding the human bfgf gene was obtained directly from the

pET28b:hbFGF-F1 and pET28b:hbFGF-F2 constructs described in Andrades JA et al., 2001.

3.1.1. Culture of U-2 OS cells.

U-2 OS human osteosarcoma cells are known to express rhBMP-6 and other members of

the BMP family. These cells were cultured as monolayers in 75 cm2 culture flasks with McCoy’s

5A medium supplemented with 10% foetal bovine serum (FBS) and 2 mM L-glutamine, at 37 ºC

in a humidified atmosphere with 5% CO2 (standard conditions). Cells were subcultured when

65-75% confluence was reached, by detaching them with 5 mL of a 0.25% tripsin, 0.03% EDTA

solution and diluting the cells 1:6 in fresh medium.

3.2. Cloning into the pET17b expression vector and the

pAcGP67B shuttle vector.

The gene encoding the hBMP-6 protein and the construction encoding the hBMP-6-CBD

were cloned into both the pET17b expression vector (for protein production in E. coli) and the

pAcGP67B shuttle vector (for protein expression in Sf9 cells). In contrast, the h-bFGF gene and

the construction encoding the h-bFGF-CBD were only cloned into the pAcGP67B shuttle vector.

For details on PCR, plasmid purification, DNA electrophoresis, DNA purification from agarose

gels, DNA digestion with endonucleases, DNA precipitation, DNA ligation, DNA sequencing,

transformation of bacteria or colony-PCR, see Appendix I, sections AI.2.3, AI.2.4, AI.2.5, AI.2.7,

AI.2.8, AI.2.9, AI.2.10, AI.2.11, AI.3.6 and AI.3.7, respectively.

Material and methods____________________________________________________________

62

3.2.1. Cloning of the hBMP-6 and the hBMP-6-CBD genes into the

pET17b and the pAcGP67B vectors.

The sequence encoding the mature domain of the hBMP-6, obtained by RT-PCR was ligated

into the pBIISK plasmid, previously digested with the EcoRV endonuclease, and the ligation

mixture was used to transform E. coli DH5α cells. In order to select a clone containing the

plasmid with the GOI correctly inserted, ten colonies were chosen and analyzed by colony-PCR

with the oligonucleotides P5 vs. P4.

The pBIISK:BMP-6 plasmid was isolated from the selected clone and used as template for

PCR reactions with a pfu polymerase and the oligonucleotides P10 vs. P11 (which yields the

mature bmp-6 sequence with an EcoRI restriction site upstream and a BamHI site downstream)

or P12 vs. P11 (which yields the mature bmp-6 sequence with an EcoRI restriction site and the

sequence for the CBD upstream and an BamHI site downstream). Both fragments were double-

digested with EcoRI and BamHI and ligated into the the pET17b vector previously digested with

the same combination of endonucleases. The ligation mixtures were used to transform E. coli

DH5α cells and the obtained clones were analyzed by colony-PCR using the oligonucleotide pair

P1 vs. P2. The selected, positive clones, were used for isolation of the pET17b:BMP-6 and

pET17b:BMP-6-CBD constructions (Fig. 17).

For cloning of the genes into the pAcGP67B shuttle vector, the obtained, isolated

pBSIIK:BMP-6 plasmid was used as a template for PCR reactions with a Pfu polymerase and the

oligonucleotides P7 vs. P8 (yielding the mature bmp-6 sequence with a BamHI restriction site

upstream and an EcoRI site downstream) or P9 vs. P8 (which yields the mature bmp-6

sequence with a BamHI restriction site and the sequence for the CBD upstream and an EcoRI

site downstream). Both fragments were double-digested with BamHI and EcoRI and ligated into

the pAcGP67B vector, previously digested with the same endonucleases. E. coli DH5α cells were

transformed with the ligation mixtures and the obtained clones were analyzed by colony-PCR

using the oligonucleotide pair P42 vs. P43. The selected, positive clones were used for isolation

of the pAcGP67B:BMP-6 and pAcGP67B:BMP-6-CBD constructions (Fig. 18)


63

Figure 17. Schematic overview of the obtaining of the pET17b:BMP-6 and the pET17b:BMP-6-CBD constructions.

P5 P6 vsRT-PCR

mature bmp-6 sequence

bmp-6 mRNA

Total RNA isolation

U-2 OS cells


Cloning

pBIISK (+)

EcoRV

pBIISK:BMP-6

Transformation of E. coli DH5α P5 P4vs

Plasmid isolation

PCR PCRP10 P11vs P12 P11 vs

mature bmp-6 sequence mature bmp-6 sequenceCBD

EcoRI/BamHI

EcoRI/BamHI

mature bmp-6 sequence mature bmp-6 sequenceCBD

pET17b


pET17b:BMP-6

mature bmp-6 sequence CBD

pET17b:BMP-6-CBD

EcoRI/BamHI

Cloning Cloning

Colony screening by PCR


64

Figure 18. Schematic overview of the obtaining of the pAcGP67B:BMP-6 and the pAcGP67B:BMP-6-CBD constructions.

3.2.2. Cloning of the h-bFGF and the h-bFGF genes into the

pAcGP67B vector.

For the obtaining of the pAcGP67B:bFGF and pAcGP67B:bFGF-CBD constructions, the

pET28b:hbFGF-F1 and pET28b:hbFGF-F2 constructs described in Andrades JA et al. (2001)

were used as templates for PCRs with a Pfu polymerase and the oligonucleotides P13 vs. P14

(yielding the bfgf sequence with a BglII restriction site upstream and an EcoRI site

downstream) or P15 vs. P14 (which yields the bfgf sequence with a BglII restriction site and the

sequence for the CBD upstream and an EcoRI site downstream). A BglII restriction site was

chosen instead of a BamHI restriction site since the sequence encoding the h-bFGF naturally

contains one BamHI site. Nevertheless, the digestion with BglII and BamHI generate


pBIISK:BMP-6

PCR PCRP7 P8 vs P9 P8vs

mature bmp-6 sequence mature bmp-6 sequence CBD

BamHI/EcoRI

BamHI/EcoRI

mature bmp-6 sequence mature bmp-6 sequence CBD

pAcGP67B


pAcGP67B:BMP-6

mature bmp-6 sequence CBD

pAcGP67B:BMP-6-CBD

BamHI/EcoRI

Cloning Cloning


65

compatible cohesive ends which allow the ligation of a BglII-digested fragment into a BamHI-

digested vector.

Both fragments were double-digested with BglII and EcoRI and ligated into the pAcGP67B

vector, previously digested with BamHI and EcoRI. E. coli DH5α cells were transformed with the

ligation mixtures and the obtained clones were analyzed by colony-PCR using the

oligonucleotide pair P42 vs. P43. The selected, positive clones were used for isolation of the

pAcGP67B:bFGF and pAcGP67B:bFGF-CBD constructions (Fig. 19)

Figure 19. Schematic overview of the obtaining of the pAcGP67B:bFGF and the pAcGP67B:bFGF-CBD constructions.

bfgf

pET28b:hbFGF-F1

PCR PCR P13 P14vs P15 P14 vs

bfgf bfgf CBD

BglII/EcoRI

BglII/EcoRI

bfgf bfgf CBD

pAcGP67B

bfgf

pAcGP67B:bFGF

bfgf CBD

pAcGP67B:bFGF-CBD

BamHI/EcoRI

Cloning Cloning

bfgf

pET28b:hbFGF-F2

CBD


66

3.3. Protein production in Escherichia coli.

To obtain purified recombinant proteins expressed in E. coli, the following steps must be

performed (Fig. 20):

- Transformation of E. coli with the expression vector (containing the gene of interest).

- Isolation of bacterial clones selected for antibiotic resistance.

- PCR analysis and/or sequencing of the selected clones to ensure that they contain the

expression vector and that it has suffered no mutations.

- Culture of the cells and protein expression.

- Cell disruption and isolation of inclusion bodies.

- Solubilization of the inclusion bodies.

- In vitro folding of the solubilized proteins, yielding a mixture of folded, misfolded and

unfolded proteins.

- Purification of the fraction containing the correctly folded proteins.

3.3.1. Obtaining of bacterial clones for protein production.

For information about culture media, culture conditions or bacterial strains, see Appendix I,

sections AI.3.1, AI.3.2 and AI.3.3, respectively.

E. coli Rosetta™ (DE3) cells were made competent and transformed by electroporation (see

Appendix I, section AI.3.5) with the pET17b:BMP-6 and the pET17b:BMP6-CBD constructions.

50 and 200 µL of each transformed aliquot were plated on LB agar dishes supplemented with

ampicillin and chloramphenicol, and incubated overnight to allow the bacteria that had

incorporated the expression vector to grow and form colonies. The next day, 10 isolated

colonies for each expression vector were selected and analyzed by colony-PCR (see Appendix I,

section AI.3.6) using specific primer pairs to detect the BMP-6 or the BMP6-CBD inserts. The

obtained PCR products were ran on an agarose gel (see Appendix I, section AI.2.5) to identify

which colonies were carrying the expression vector.

5 positive colonies for each expression vector were used to inoculate 5 mL 2xYT cultures,

which were incubated overnight. 1 mL of each culture was used to prepare a glycerol stock (see

Appendix I, section AI.3.4), while the other 4 mL were used for plasmid isolation (see Appendix

I, section AI.2.4). 2 plasmid samples for each expression vector were sequenced using specific

primers against the regions of the plasmidic DNA flanking the insert (T7 Promoter and

Terminator Primers, see Appendix I, section AI.2.13).


67

Figure 20. Schematic overview of the main steps needed for recombinant protein production in E. coli.

Expression vector Escherichia coli Transformation

Cell culture & protein expression

Inclusion body isolation

Inclusion body solubilization

In vitro refolding

Purification

Purified rhBMP-6 dimers

rhBMP-6 monomers

Inclusion bodies

Soluble rhBMP-6 monomers


68

3.3.2. Protein expression.

Isolated colonies from the clones containing the recombinant expression vectors, selected

by DNA sequencing, were obtained by resuspending a small amount of glycerol stock in 100 µL

of LB broth and seeding this suspension on an LB agar plate supplemented with ampicillin and

chloramphenicol. After incubating the plate for 24 hours at 37 ºC, one single, isolated colony

was picked and used to inoculate a 10 mL TB culture, which was grown overnight at 37 ºC with

200 rpm shaking. The following day, the OD600 of this inoculum was measured and a proper

volume of it was used to inoculate a new 100 mL TB culture to a starting OD600 value of 0.1.

This culture was grown at 37 ºC with shaking and its OD600 measured hourly. When the OD600

of the culture reached 0.8, IPTG was added to a final concentration of 1mM to induce

heterologous protein expression. The culture was incubated for an additional 4 hours, taking a

1 mL sample hourly for protein expression analysis. From each of these samples, cells were

harvested by centrifugation at 5,800 xg for 30 min at room temperature, resuspended in 50 mM

PB, pH 7.0 to a final OD600 value of 5.0, sonicated at 50 W for 2 minutes on ice and stored at

4 ºC until analysis by SDS-PAGE (see section 3.5.1). After 4 hours of induction, all the cells of

the culture were harvested by centrifugation and the pellet was stored at -80 ºC until inclusion

body isolation (see section 3.3.3).

3.3.3. Isolation of inclusion bodies.

The frozen cell pellet was resuspended in 50 mM PB to a final OD600 value of 5.0, pH 7.0

(one volume) and sonicated at 50 W for 2 minutes on ice. A small aliquot of this total protein

cell-content sample was taken for SDS-PAGE analysis, and the remaining sample was

centrifuged for 40 min at 38,000 xg, 4 ºC. The supernatant was removed and the insoluble

proteins contained in the pellet were resuspended with the same volume of 50 mM PB, pH 7.0.

A small aliquot of this insoluble protein sample was taken apart for SDS-PAGE analysis (see

section 3.5.1), and the remaining sample was used for isolation of inclusion bodies. For this

purpose, the sample was centrifuged for 40 min at 38,000 xg, 4 ºC, resuspended in ¼ the

volume of 50 mM PB, pH 7.0, centrifuged again and finally resuspended in twice the volume of

MR buffer (20 mM Tris-HCl, pH 8.5, 0.5 mM EDTA, 2% Tx-100), which removes the membrane-

associated and other lipophilic proteins. The proteins were centrifuged twice more at the same

conditions and resuspended in one volume and ½ the volume of MR buffer, respectively. A

small aliquot of the final suspension, containing the washed inclusion bodies, was taken apart

for SDS-PAGE analysis, and the remaining sample was used for inclusion body solubilization

(see section 3.3.4).


69

3.3.4. Solubilization of inclusion bodies.

In order to solubilize the recombinant proteins prior to in vitro refolding, the sample was

centrifuged for 40 min at 38,000 xg, 4 ºC, and the pellet was resuspended in 3 mL of

solubilization buffer (0.1 M Tris, pH 8.5, 6 M Gnd-HCl, 0.1 M DTT, 1 mM EDTA). Solubilization

was performed overnight, at room temperature, with constant stirring. The next day, the

sample was centrifuged for 45 min at 26,000 xg, 4 ºC to remove the remaining insoluble

particles, and the supernatant (containing the solubilized proteins) was transferred to another

tube. The pH of the sample was lowered below 6.0 by 16% HCl addition and the sample was

then dialyzed against MES-Gnd buffer (50 mM MES, pH 5.0, 6 M Gnd-HCl, 1 mM EDTA) for at

least 4 hours at 4 ºC in order to lower the DTT concentration below 1-2 mM, since the presence

of DTT would interfere with the protein refolding by reducing the cysteine residues involved in

disulfide bond formation.

After the dialysis, the final protein concentration of the sample was estimated by SDS-PAGE

analysis, since the concentration of the target protein during refolding is a critical parameter.

For this purpose, a small aliquot of the sample was diluted 1 :1,000 in 2x SDS-PAGE loading

buffer with DTT and heated at 95 ºC for 5 minutes. 1 :2, 1 :4, 1 :6, 1 :8 and 1 :10 dilutions in

2x SDS-PAGE loading buffer with DTT were made from the previous stock dilution and loaded

on a polyacrylamide gel for SDS-PAGE and Coomassie blue staining (see section 3.5.1 and

Appendix I, section AI.5.3, respectively). A sample of BMP-2 monomers of known concentration

was used as a standard for estimation of the rhBMP-6 concentration by digital image analysis

(Quantiscan v3.0, Biosoft®, Cambridge, UK).

3.3.5. In vitro refolding.

The attempts on refolding of rhBMP-6 produced in E. coli were based on the conditions

established for efficient rhBMP-2 refolding by Vallejo LF et al., 2002. These authors achieved

high refolding yields of this member of the BMP family by performing the renaturation for 72

hours at 10 ºC in a degassed refolding buffer containing 57.5 mM Tris, pH 8.5, 0.55 mM EDTA,

0.9 M NaCl, 0.75 M CHES, 0.5 M Gnd-HCl, 2 mM GSH and 1 mM GSSG. These were the starting

conditions used for rhBMP-6 refolding. The different variations on these conditions tested can

be found summarized in Table 7.

In general, 50 mL of redox base buffer A (55 mM Tris, pH 7.5 or pH 8.5 or pH 9.5, 1 M

NaCl, 0.5 mM EDTA, 0.82 M CHES or 0.55 M NDSB256 or 1.09 M NDSB256 or 0.55 mM

L-arginine or 1.64 M L-arginine) were mixed with 4.2 mL of redox base buffer B (100 mM Tris,

pH 7.5 or pH 8.5 or pH 9.5, 6 M Gnd-HCl, 1 mM EDTA). In some cases this mixture was

degassed for 20 minutes by N2 infusion while in others it was not. 0.55 mL of a 100x redox pair


70

stock solution (GSH :GSSG or 4-MPAA :GSSG) were added to the mixture and, afterwards,

0.39 mL of the rhBMP-6 solution, previously diluted with MES-Gnd buffer to obtain a 140-fold

concentration of the desired final concentration in the refolding mixture, were added. This

refolding mixture was placed in a cooling bath at 10 or 20 ºC and left for 72 hours. In some

cases, N2 was continuously supplied to the mixture during the entire refolding process to avoid

air-oxidation of the cysteine residues.

After 72 hours of incubation, the refolding yield for each condition tested was analyzed. For

this purpose, a 100 µL sample of the refolding mixture was taken apart and 11 µL of 1 M

iodoacetate was added to stop the refolding process by alkylation of the sulfhydryl groups. After

20 minutes of incubation at room temperature, the proteins in this aliquot were precipitated

with trichloroacetic acid (see Appendix I, section AI.5.1) and used for SDS-PAGE analysis (see

section 3.5.1).


71

Pro

tein

con

cen

trat

ion

Tem

p.

pH

Add

itiv

es

Red

ox p

air

Deg

assi

ng

CH

ES

ND

SB2

56

Arg

inin

e

GSH

:GSS

G

4-M

PA

A :

GS

SG

Yes

1 m

M

3 m

M

3 m

M Att

emp

t n

º

10

.7 µ

g/m

L

20

.0 µ

g/m

L

50

.0 µ

g/m

L

53

.4 µ

g/m

L

10

ºC

20

ºC

7.5

8.5

9.5

0.7

5 M

0.5

M

1.0

M

0.5

M

1.5

M

2 :

1

10

:1

2 :

1

10

:1

40

:1

2 :

1

10

:1

No

No

con

tin

uou

s N

2 s

upp

ly

Con

tin

uou

s N

2 s

upp

ly

1 ● ● ● ● ● ●

2 ● ● ● ● ● ●

3 ● ● ● ● ● ●

4 ● ● ● ● ● ●

5 ● ● ● ● ● ●

6 ● ● ● ● ● ●

7 ● ● ● ● ● ●

8 ● ● ● ● ● ●

9 ● ● ● ● ● ●

10 ● ● ● ● ● ●

11 ● ● ● ● ● ●

12 ● ● ● ● ● ●

13 ● ● ● ● ● ●

14 ● ● ● ● ● ●

15 ● ● ● ● ● ●

16 ● ● ● ● ● ●

17 ● ● ● ● ● ●

18 ● ● ● ● ● ●

19 ● ● ● ● ● ●

20 ● ● ● ● ● ●

21 ● ● ● ● ● ●

22 ● ● ● ● ● ●

23 ● ● ● ● ● ●

24 ● ● ● ● ● ●

25 ● ● ● ● ● ●


72

Pro

tein

con

cen

trat

ion

Tem

p.

pH

Add

itiv

es

Red

ox p

air

Deg

assi

ng

CH

ES

ND

SB2

56

Arg

inin

e

GSH

:GSS

G

4-M

PA

A :

GS

SG

Yes

1 m

M

3 m

M

3 m

M Att

emp

t n

º

10

.7 µ

g/m

L

20

.0 µ

g/m

L

50

.0 µ

g/m

L

53

.4 µ

g/m

L

10

ºC

20

ºC

7.5

8.5

9.5

0.7

5 M

0.5

M

1.0

M

0.5

M

1.5

M

2 :

1

10

:1

2 :

1

10

:1

40

:1

2 :

1

10

:1

No

No

con

tin

uou

s N

2 s

upp

ly

Con

tin

uou

s N

2 s

upp

ly

26 ● ● ● ● ● ●

27 ● ● ● ● ● ●

28 ● ● ● ● ● ●

29 ● ● ● ● ● ●

30 ● ● ● ● ● ●

31 ● ● ● ● ● ●

32 ● ● ● ● ● ●

33 ● ● ● ● ● ●

34 ● ● ● ● ● ●

35 ● ● ● ● ● ●

36 ● ● ● ● ● ●

37 ● ● ● ● ● ●

38 ● ● ● ● ● ●

39 ● ● ● ● ● ●

40 ● ● ● ● ● ●

41 ● ● ● ● ● ●

Table 7. Attempts on in vitro refolding of rhBMP6 monomers produced in Escherichia coli. Each row corresponds to one tested combination of parameters, which are highlighted at the upper part of the table.


73

3.4. Protein production in Sf9 insect cells.

To obtain a recombinant, infective baculovirus clone to be used for the production of

recombinant proteins in Sf9 cells, the following steps must be performed (Fig. 21):

- Co-transfection of Sf9 cells with the shuttle vector (containing the gene of interest) and

linearized baculovirus DNA. Homologous recombination events inside the cells will give

rise to complete viral DNA molecules.

- Separation of single viral clones by plaque assays.

- PCR analysis and/or sequencing of the selected clones to ensure that the gene of

interest is correctly inserted into de viral DNA and has suffered no mutations.

- Expansion of the selected clones and titering of the obtained viral suspensions.

- Production assays, to determine the optimal conditions of MOI and days post-infection

to obtain the highest yield of correctly expressed proteins.

Co-transfection into Sf9 cells

Shuttle vector Linearized baculovirus DNA

Transfection supernatant

Plaque assay

Single plaque picking Recombinant virus clone (Virus stock)


74

Figure 21. Schematic overview of the steps needed to obtain a recombinant baculovirus.

Master stock of virus (MSV)Titering

Clone expansion

Titering High volume virus stock (HVVS)

Production assay

Western blot analysis

PCR analysis and/or sequenciation

Clone expansion


75

3.4.1. Culture of Sf9 cells.

The Sf9 cell line was obtained from pupal ovarian tissue of the fall armyworm Spodoptera

frugiperda, by isolation of a clone of the parental IPLB-SF21-AE cell line (Vaughn JL et al.,

1977) by G. Smith and C. Cherry in 1983 (O’Reilly DR et al., 1994). This cell line is highly

susceptible to infection with Autographa californica nuclear polyhedrosis virus, and is widely

used for the production of recombinant proteins. Sf9 cells are spherical, and can be grown both

as an adherent monolayer in culture flasks or dishes, or as a suspension culture in spinner

vessels.

Culture of Sf9 cells was always performed at 28 ºC in an incubator without CO2 supply, in

TNM-FH Medium (Trichoplusia ni Medium – Formulation Hink), composed of Grace Medium for

insect cells (Grace TD, 1962) supplemented with trace metals, lactalbumin hydrolysate,

yeastolate and 10% (v/v) heat inactivated foetal bovine serum (FBS). To ensure sterility of the

cultures, amphotericin B (2.5 mg/L, final concentration) and a penicillin/streptomycin solution

(105 U/L and 100 mg/L, final concentrations, respectively) were also added to the medium. Only

during the production of the recombinant proteins, the cells were maintained in TNM-FH

Medium without FBS, but supplemented with 2 mM L-glutamine, since some authors have

published that high levels of L-glutamine can enhance recombinant protein production in these

cells (Sanders MM and Kon C, 1992; Nguyen B et al., 1993; Benslimane C et al., 2005).

Monolayer cultures were performed in Petri dishes (90 or 150 mm diameter) or in culture

flasks (T-75 cm2 or T-175 cm2) at a starting density of 3x105 cells/mL. When the monolayer

became confluent, cells were harvested by gently scraping with a sterile Digralsky spreader

covered by teflon® and subcultured at a 1:5 dilution.

Suspention cultures were started seeding cells harvested from adherent cultures at 5x105

cells/mL in a spinner vessel. Cells were grown with gentle agitation (50-100 rpm) until a density

of 2 – 3x106 cells/mL was reached before subculturing.

To determine the cell density of any suspension, cells were diluted in a trypan blue solution

and counted using a Neubauer haemocytometer (see Appendix I, section AI.4.3). All

experiments were performed seeding cells with a viability of 95% or higher.

3.4.2. Transfection of Sf9 cells.

Once the GOI was correctly cloned into the pAcGP67B shuttle vector, Sf9 cells had to be co-

transfected with this construction and linear baculovirus DNA molecules (BacPak6™ or

Sapphire™) in order to generate complete, infective baculoviruses by homologous

recombination within the cells.


76

For this purpose, cells from a suspension culture growing in log phase were seeded in 6-

well culture plates at a density of 1.5x106 cells/well and incubated for 1 h at 28 ºC to allow cells

to adhere to the bottom of the wells. Afterwards, the medium was removed from the wells,

cells were washed twice with serum-free TNM-FH and incubated another 30 minutes at 28 ºC

with 1.5 mL of serum-free TNM-FH/well. Meanwhile, the transfection mixtures were prepared in

sterile polyestirene tubes, gently mixed and left for 15 minutes at room temperature to allow

formation of complexes between the DNA molecules and the liposomes provided by the

transfection reagent. A control transfection mixture (reagent C-) was also prepared, lacking the

baculoviral DNA. A second control (cell C-) consisted of just 100 µL sterile water.

Transfection Reagent C- Cell C-

Sterile water 86 µL 91 µL 100 µL

pAcGP67B:GOI (100 ng/µL) 5 µL 5 µL -

Baculovirus DNA (20 ng/µL) 5 µL - -

Transfection reagent 4 µL 4 µL -

Each transfection mixture was added, drop by drop, to a different well of the culture plate,

and the plate was left for 4 h at room temperature with gentle shaking. Afterwards, 1.5 mL of

TNM-FH + 10% FBS was added to each well and the plate was left in a 28 ºC incubator for 6-9

days.

Finally, once the transfected cultures started showing signs of infection (increased cell

diameter, granular appearance of the cytoplasm and decreased growth rate), the medium

containing the recombinant baculoviruses was harvested, centrifuged at 1,000 xg for 5 minutes

to eliminate cells and cell debris, and the supernatant stored at 4 ºC. This supernatant was

denominated transfection supernatant (TS).

3.4.3. Isolation of viral clones (plaque assay).

The transfection supernatant obtained in the previous step is a heterogeneous mixture of

recombinant baculoviruses. To ensure the production of a homogeneous pool of recombinant

proteins, the cells have to be infected with one single viral clone.

The method used for isolation of clones from the TS was the plaque assay, by which cell

monolayers were infected with serial dilutions of the TS and covered with agarose to limit the

mobility of the viruses. When a cell is infected by one virus, the budded viruses liberated at the

end of the infection cycle are prevented by the agarose from freely disseminating through the


77

culture and will, thus, infect only the neighbour cells. This leads to the formation of lysis

plaques, which can be visualized by staining the living cells surrounding them (Fig. 22).

Figure 22. Formation of a lysis plaque. 1. One single virus infects a cell. 2. First infected cell lyses and newly formed viruses infect the surrounding cells. 3. Staining of the living cells with a vital stain reveals the lysis plaque.

To perform a plaque assay, cells from a suspension culture growing in log phase were

seeded in 6-well culture plates at a density of 1.2x106 cells/well and incubated for 1 h at 28 ºC

to allow cells to adhere to the bottom of the wells. Afterwards, the medium was retired from

the wells, cells were washed twice with serum-free TNM-FH and incubated another 30 minutes

at 28 ºC with 1.5 mL of serum-free TNM-FH/well. In the meantime, serial dilutions of the TS

were prepared in serum-free TNM-FH, from 10-3 to 10-8. Afterwards, the medium was removed

from the wells and 300 µL of each TS dilution were added, drop by drop, on top of the cell

monolayers of the different wells. Viruses were allowed to infect the cells during 2 h at room

temperature, gently inclining the plate every 5-10 minutes to avoid drying of the cells.

Meanwhile, a 3% low melting agarose solution was mixed 1:1 with TNM-FH + 10% FBS,

and kept in a heating bath at 37 ºC. The virus solutions were retired from the wells and 2 mL

agarose solution were carefully added to each well, covering the cell monolayers. Agarose was

allowed to solidify at room temperature for 30 minutes before adding 1 mL of TNM-FH + 10%

FBS to each well. The plate was then incubated for 5 days at 28 ºC.

Once the incubation time was finished, the medium was carefully removed from the wells

and 1 mL of a 0.33% neutral red solution diluted 1/16 in sterile PBS was added to each well.

After 2 h of incubation at room temperature, the staining solution was retired and the plate was

left upside down overnight to drain the remaining liquid.

The following day, the lysis plaques were observed through an inverted microscope and the

agarose covering each plaque was picked out using a sterile Pasteur pipette. Each piece of

agarose, containing a single viral clone, was placed in a microcentrifuge tube with 500 µL of

TNM-FH + 10% FBS and stored at 4 ºC. These samples were denominated virus stocks (VS)

1 2 3


78

3.4.4. PCR analysis of the viral clones.

Although the use of linearized (Bsu36I digested) viral DNA guarantees that no wild-type,

infective baculoviruses can rise in the TS, the clones selected by plaque assay were submitted

to PCR analysis to verify correct cloning of the GOI into the viral DNA (Malitschek B and Schartl

M, 1991; Webb AC et al., 1991). The list of oligonucleotides used for this purpose can be found

in Appendix I, section AI.2.13.

To obtain the viral DNA suitable for PCR analysis, 10 µL of each VS were mixed with 89 µL

of virus lysis buffer (10 mM Tris-HCl, pH 8.3, 100 µg/mL gelatine, 0.45% (v/v) Triton X-100,

0.45% (v/v) Tween-20, 50 mM KCl) and 1 µL of proteinase K solution (6 mg/mL). These

mixtures were first incubated for 1 h at 60 ºC in a dry block heated thermostat and, afterwards,

for 10 minutes at 90 ºC. This procedure destroys the viral coating and liberates the DNA

molecules.

PCR reactions were prepared using the 5Prime® MasterMix system. For each reaction

(25 µL total volume), 11.5 µL sterile water, 10 µL 5Prime® MasterMix, 2.5 µL virus sample and

0.5 µL of each 25 pM oligonucleotide solution were mixed in a 0.2 mL PCR tube. PCR was

performed using the Ta specified for each oligonucleotide pair.

Finally, the obtained PCR products were analyzed by DNA electrophoresis in agarose gels

(see Appendix I, section AI.2.5).

3.4.5. Expansion of the baculovirus clones.

For the production of the recombinant proteins, a large, homogeneous and titered batch of

baculoviruses is fundamental. This ensures that every production can be performed under the

same conditions and with the same MOI (multiplicity of infection; i.e. the number of pfu per cell

used to infect a culture).

94 ºC 94 ºC 1 min

72 ºC 72 ºC

x35 cycles

1 min 6 min 2 min 1 min Ta


79

The first amplification of the VS gives rise to a master stock of virus (MSV), which is titered

by a serial dilution assay (see section 3.4.6). Once the titer of the MSV is known, a second

amplification is performed to obtain a high volume virus stock (HVVS). This HVVS is produced

infecting a culture with a low MOI (0.1), so that the viral population can grow exponentially

before the majority of the cells die, yielding a high-titer suspension of virus. Once the HVVS is

harvested, it also has to be titered, so that every future protein production can be performed

using the same MOI.

Every MSV was obtained by seeding 5x106 cells on a 90 mm culture dish with, 10 mL of

TNM-FH + 10% FBS. Once the cells were attached to the bottom of the dish, the medium was

removed before adding 100 µL of the VS, drop by drop, on top of the monolayer of cells.

Afterwards, 10 mL of fresh TNM-FH + 10% FBS were added and the culture was incubated at

28 ºC for 7 days. The viral suspension was harvested, centrifuged for 5 minutes at 1,000 xg in

a swing bucket centrifuge to remove cell debris, and the supernatant stored at 4 ºC before

titering (Jarvis DL and Garcia A Jr, 1994).

The HVVS were produced by seeding 3x107 cells on a 160 mm culture dish, with 30 mL of

TNM-FH + 10% FBS. The cells were allowed to adhere to the bottom of the dish for 1 hour

before removing the medium and infecting the culture with the appropriate volume of MSV,

corresponding to a MOI of 0.1. Afterwards, 30 mL of fresh TNM-FH + 10% FBS were added to

the dish, and the culture was left in a 28 ºC incubator for 7 days. Finally, the viral suspension

was harvested, centrifuged for 5 minutes at 1,000 xg in a swing bucket centrifuge, and the

supernatant stored at 4 ºC before titering.

3.4.6. Titering of viral suspensions.

The concentration of infective viruses in a suspension can be determined using a variant of

the limit dilution method described by Reed and Muench (1938). This method yields a value for

the 50% tissue-culture infectious dose (TCID50), which is the dilution at which 50% of the

inoculated cultures become infected. The TCID50 of a viral suspension can be converted into a

titer expressed as pfu/mL applying a Poisson distribution.

Titering was performed seeding cells from a suspension culture growing in log phase in 96-

well culture plates at a density of 8x103 cells/well and incubating the plates for 1 h at 28 ºC to

allow cells to adhere to the bottom of the wells. Meanwhile, serial ten-fold dilutions of the viral

suspension were prepared, from 10-5 until 10-16. Afterwards, 10 µL of each dilution were added

to 12 wells and the plate was left for 6 days at 28 ºC.

At the end of the process, each well was observed through an inverted microscope and the

number of infected wells for each dilution was determined. With these numbers, the TCID50

value and the titer in pfu/mL was calculated.


80

3.4.7. Production assays.

Before starting large-scale protein productions, the best conditions of MOI and production

time had to be determined for each recombinant protein. Usually, protein productions are

initiated infecting a culture with a high MOI (1-10) to ensure a high protein synthesis rate and a

rapid increase of the recombinant protein levels in the culture medium. The main disadvantage

of this strategy is the also high rate of cell lysis of the culture, which implies a significant

release of contaminants and proteases.

Combinations of MOI values of 2.5 and 10, and production times of 72, 96 and 120 hours

were tested for each protein to produce. Note that the production time was started counting

once the culture medium was changed by serum-free TNM-FH, what was done 24 hours p.i.

To perform a production assay, cells from a suspension culture growing in log phase were

seeded on 24-well culture plates at a density of 5x105 cells/well. Plates were then left for 1 hour

at 28 ºC to allow the cells to adhere to the bottom of the wells. The medium was removed and

the proper volume of HVVS was added to 16 of the wells before adding 500 µL of fresh TNM-FH

+ 10% FBS. Eight wells were infected with a MOI of 2.5, and the other eight wells with a MOI

of 10. The remaining wells were left uninfected to serve as a negative control. After 24 hours of

incubation at 28 ºC, the cells were washed twice with serum-free TNM-FH which was finally

replaced by 500 µL of serum-free TNM + 10% FBS + 2 mM L-glutamine. At this point the

production time started counting.

72 hours later, the medium of two wells infected with MOI 2.5 and two wells infected with

MOI 10 was harvested, centrifuged for 5 minutes at 1,000 xg, and the supernatants were

stored at -20 ºC for its further analysis. Every next 24 hours, the medium of two wells for each

MOI was harvested in the same way. The negative controls were harvested together with the

samples corresponding to a 120 hour production time.

400 µL of each sample were precipitated with trichloroacetic acid (see Appendix I, section

AI.5.1) and analyzed by SDS-PAGE and Western blot (see sections 3.5.1 and 3.5.2).

3.4.8. Large-scale protein production.

Once the optimal conditions of MOI and production time have been established, the goal

proteins can be produced at large scale. For this purpose, cells from a suspension culture

growing in log phase were seeded in T-175 cm2 culture flasks at a density of 4 x 107 cells/flask.

Once the cells had adhered to the bottom of the flasks, the medium was removed and the cells

were infected with the proper volume of HVVS to achieve the MOI selected by the previous

production assay. 40 mL of fresh TNM-FH + 10% FBS were added to each flask.


81

After 24 hours of incubation at 28 ºC, the cells were washed twice with serum-free TNM-FH

and, at the end, 40 mL of serum-free TNM-FH + 2 mM L-glutamine were added to each flask.

The cultures were then left in a 28 ºC incubator until the optimal production time was reached.

At this moment, the medium was harvested and centrifuged for 10 min at 1,000 xg in a swing

bucket centrifuge. In case the supernatant was not directly used for protein purification, it was

stored at -80 ºC.

3.4.9. Purification of rhBMP-6 produced in Sf9 cells.

The rhBMP-6 produced in Sf9 cells was purified by its natural affinity to heparin, using 1 mL

HiTrap™ Heparin HP columns (Amersham Biosciences / GE Healthcare). Because of the strong

tendency towards aggregation of this protein, the entire process was carried out in presence of

6 M urea to prevent it from precipitating inside the column or in the elution fractions. Therefore,

urea was dissolved directly into the conditioned medium to reach a 6 M concentration. Before

passing this sample through the column, it was filtered through a 2 mL Sephadex® G25 column

to eliminate any possible precipitates and/or particles.

The entire purification was performed using a BioLogic Duo Flow chromatographer (Biorad),

at room temperature, and at a flow rate of 1 mL/min. First, the column was equilibrated with 5-

10 mL of serum-free TNM-FH + 6 M urea before loading the sample (100-500 mL). Once the

sample had entirely passed through the column, the latter was washed with 10 mL BMP-

washing buffer (50 mM MES, pH 5.5, 6 M urea, 0.15 M NaCl) prior to eluting the proteins.

Elution of the retained proteins was induced by increasing the conductivity in the column.

In some cases, this was done using a linear gradient from BMP-washing buffer + 0.15 M NaCl

to 1 M NaCl, and in other cases a two-step elution was performed (a first step elution with BMP-

washing buffer + 0.43 M NaCl and a second step elution with BMP-washing buffer + 1 M NaCl).

The elution fractions were analyzed by Western dot-blot (see section 3.5.3) to identify those

containing the protein of interest.

To obtain the proteins in their native, active forms, the urea and the excess of NaCl had to

be removed from the samples. To achieve this, different strategies were tried:

1. Samples were directly dialyzed for at least 4 hours against DMEM, pH 7.0.

2. Samples were directly dialyzed for at least 4 hours against DMEM, pH 4.9.

3. Samples were directly dialyzed for at least 4 hours against 4 mM HCl.

4. Samples were dialyzed for at least 4 hours against 10 mM ammonium acetate, pH 4.0,

subsequently lyophilized and resuspended in sterile, highly pure water.


82


subsequently lyophilized and resuspended in sterile, highly pure water with 0.1% BSA.


subsequently lyophilized and resuspended in 4 mM HCl.


subsequently lyophilized and resuspended in 4 mM HCl, 0.1% BSA.

8. Samples (2 mL) were loaded on Vivaspin 2 columns (GE Healthcare) and centrifuged at

10,000 xg at room temperature to reduce their volume. Then, 4 mM HCl was added to

the column to dilute the urea and the NaCl of the sample. These steps were repeated

until the urea and NaCl concentration was below 5 mM and the volume of the sample

was reduced to, approximately, 0.5 mL.

The biological activity of all the samples was tested in vitro on C2C12 mouse myoblasts

(section 3.7.1), using serial dilutions of the samples from 1/2 to 1/1024 in DMEM + 2% FBS.

3.4.10. Purification of rh-bFGF and rh-bFGF-CBD produced in Sf9 cells.

The rh-bFGF and rh-bFGF-CBD produced in Sf9 cells were purified by its natural affinity to

heparin, using 1 mL HiTrap™ Heparin HP columns (Amersham Biosciences / GE Healthcare).

Before passing the samples through the column, they were pre-filtered through a 2 mL

Sephadex® G25 column to eliminate any possible precipitates and/or particles.

The entire purification was performed using a BioLogic Duo Flow chromatographer (Biorad),

at room temperature, and at a flow rate of 1 mL/min. The column was first equilibrated with 5-

10 mL of serum-free TNM-FH before loading the sample (250 mL). Once the sample had

entirely passed through the column, the latter was washed with 10 mL FGF-washing buffer

(20 mM Tris, pH 7.1, 1 mM EDTA, 1 mM DTT, 0.15 M NaCl) prior to eluting the proteins.

Elution of the retained proteins was induced by increasing the conductivity in the column

using a 60 mL linear gradient from FGF-washing buffer + 0.15 M NaCl to 2 M NaCl, harvesting

1 mL elution fractions.

The elution fractions were analyzed by Western dot-blot (see section 3.5.3) to identify

those containing the protein of interest. Similar elution fractions were then reunited and treated

for removal of excess of NaCl. For this purpose, the samples were loaded on Vivaspin 2 columns

(GE Healthcare) and centrifuged at 10,000 xg, 20 ºC until the volume was reduced to

approximately 0.3 mL. Sterile PBS, pH 7.3, 1 mM EDTA, 1 mM DTT was added to the column to

dilute the salt concentration in the sample, being these steps repeated until the NaCl

concentration in the samples was below 20 mM and the total volume was reduced to

approximately 0.3 mL.


83

3.5. Biochemical analysis of the produced proteins.

3.5.1. SDS-PAGE.

Analysis of the production, refolding and/or renaturation of the proteins was performed by

sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE), following the method

described by Laemmli (1970).

Samples were diluted 1:1 in 2x SDS-PAGE Sample Buffer and heated at 95 ºC for 5 minutes

in a dry block heating thermostat prior to loading them into a 90x60x0.75 or 90x60x1.50

(mm WxHxT) gel. Polyacrylamide concentration of the gels was usually 12.5% for BMP-6

analysis, and 15.0% for bFGF analysis. In case the presence of disulfide bonds in the proteins

needed to be determined, DTT was added to the Sample Buffer to a final concentration of

135 mM. Besides the samples, one well of each gel was used for loading 10 µg of a commercial

mixture of molecular mass standard proteins: myosin (200.00 KDa), β-galactosidase

(116.25 KDa), phosphorylase b (97.40 KDa), BSA (66.20 KDa), ovoalbumin (45.00 KDa),

carbonic anhydrase (31.00 KDa), soybean trypsin inhibitor (21.50 KDa), lysozyme (14.40 KDa)

and aprotinin (6.50 KDa) (BioRad).

Electrophoresis was performed at 200 Volt, at room temperature, for approximately 1 hour,

in Laemmli Running Buffer. Once the electrophoretic separation of the samples was finished,

proteins in the gel were stained with Coomassie Blue (see Appendix I, section AI.5.3) or

transferred to a PVDF membrane for Western Blot analysis (see section 3.5.2).

3.5.2. Western blot.

After their electrophoretic separation, proteins in the gel were transferred to a PVDF

membrane following the method described by Towbin H et al. (1979). For the detailed protocol

see Appendix I, section AI.5.4.

Transference was performed at 100 mA, at room temperature, for approximately 3 hours,

in transference buffer. Once the transference was completed, the lane corresponding to the

molecular mass standard proteins was stained with amido black (see Appendix I, section

AI.5.5), while the membrane with the transferred samples was immunostained (see Appendix I,

section AI.5.6).


84

3.5.3. Dot blot.

In some cases, protein samples were not separated electrophoretically before

immunostaining, but loaded directly onto a PVDF membrane. This permits identification of the

goal protein within a sample, without determining its molecular mass.

For dot blotting, 10 µL of each sample was directly pipetted as a spot onto a PVDF sheet,

previously activated with methanol and washed with PBST. Once the drop of sample was

absorbed by the membrane, the samples were treated for immunostaining as described in

Appendix I, section AI.5.6.

3.6. Collagen-binding affinity test.

In order to test the affinity of the growth factors to collagen in the form of absorbable

collagen sponges (ACS), a method described by T. Kitajima was used (Kitajima T et al., 2007).

An ACS sheet (obtained from highly pure, bovine skin-derived, native collagen; kindly provided

by Dr. M. E. Nimni; US patent 5374539) was cut into discs (5 mm diameter, 1 mm thickness;

Fig. 23) and washed with PBST (PBS + 0.1% Tween-20). Each disc was impregnated with

10 µL of a solution containing 1.25 pmol of each growth factor, and incubated for 2 hours at

37 ºC. Afterwards, the ACS discs were washed with PBST for 1 hour to remove the unbound

molecules and immunostained with an anti-bFGF antibody as described for immunostaining of

proteins on PVDF in Appendix I, section AI.5.6.

To test the stability of the binding to collagen, the same protocol was used, but the

washing step after binding of the proteins to the ACS was prolonged for 6 days before

immunostaining, with the PBST being renewed twice every day. This washing was performed at

4 ºC to minimize protein loss by degradation.

Quantification of the amount of proteins bound to the collagen sponges was done using

digital image analysis with the free software program ImageJ (Rasband, WS., ImageJ, U.S.

National Institute of Health, Bethesda, Maryland, USA; http://rsb.info.nih.gov.ij), version 1.38x.

A circular area of approximately 5 mm diameter was placed on the centre of each spot and the

pixel density within that area was measured and used for comparison between the different

sponges. The background of the image, corresponding to the measurement of pixel density on

the negative control sponge, was subtracted from all the other obtained values for graphic

representation of the data.


85

Figure 23. Real size photograph of the absorbable collagen sponge discs used for the collagen-binding affinity tests and for the in vivo heterotopic bone formation assay.

3.7. In vitro biological activity tests.

3.7.1. Induction of ALP expression on C2C12 mouse myoblasts.

C2C12 mouse myoblasts were cultured as monolayers in 175 cm2 culture flasks with

Dulbecco's Modified Eagle's Medium (DMEM) supplemented with 10% foetal bovine serum

(FBS) and 2 mM L-glutamine, at 37 ºC in a humidified atmosphere with 5% CO2 (standard

conditions). Cells were subcultured when 65-75% confluence was reached, by detaching them

with 5 mL of a 0.25% trispin, 0.03% EDTA solution and diluting the cells 1:6 in fresh medium.

C2C12 cells are known to transdifferentiate from the myoblastic to the osteoblastic lineage

in presence of BMPs, which induce the expression of osteogenic markers, such as alkaline

phosphatase, in these cells. A method described by Katagiri T et al. (1994) was used to test the

biological activity of rhBMP-6. For this purpose, 30,000 cells/well were seeded on 96-well

culture plates with DMEM + 10% FBS, and incubated for 3 hours under standard conditions.

Afterwards, the medium was removed from the wells, replaced by DMEM + 2% FBS and the

cells incubated with this medium for another hour. The medium was again removed, replaced

by 100 µL of DMEM + 2% FBS containing the rhBMP-6 to test, and the cells were incubated for

72 hours under standard conditions.

After this time, the cells were washed with PBS and 100 µL of p-nitrophenyl phosphate

(p-NPP, Sigma Fast™) was added to each well. After 5-15 minutes of incubation under standard

conditions, the reaction was stopped by addition of 100 µL 0.1 M NaOH and the OD405 in each

well was measured using an ELISA plate reader. Finally, the OD405 values obtained were

transformed into U/L.

As a positive control, commercial rhBMP-6 produced in CHO cells (R&D Systems) was used.


86

3.7.2. Proliferation assay on MC3T3-E1 mouse preosteoblasts.

MC3T3-E1 mouse preosteoblasts were cultured as monolayers in 175 cm2 culture flasks

with alpha-Minimum Essential Medium (alpha-MEM) supplemented with 10% fetal bovine serum

(FBS) and 2 mM L-glutamine, at 37 ºC in a humidified atmosphere with 5% CO2 (standard

conditions). Cells were subcultured when 65-75% confluence was reached, by detaching them

with 5 mL of a 0.25% trispin, 0.03% EDTA solution and diluting the cells 1:4 in fresh medium.

To test the ability of rh-bFGF and rh-bFGF-CBD to induce proliferation of these cells, a

method based on the cleavage of the tetrazolium salt MTT to formazan crystals by metabolic

active cells was used (Vistica DT et al., 1991). When cells are incubated with the yellow MTT

solution, this tetrazolium salt is cleaved inside the mitochondria, yielding purple, insoluble

formazan salt crystals. These crystals can be solubilized and spectrophotometrically quantified

using an ELISA reader. An increase in number of living cells results in an increase of the total

metabolic activity in the sample, and this increase directly correlates to the amount of purple

formazan crystals formed.

For this purpose, 10,000 cells/well were seeded on 96-well culture plates with alpha-MEM +

2 mM L-glutamine + 10% FBS, and incubated for 3 hours under standard conditions.

Afterwards, the medium was removed from the wells, replaced by alpha-MEM +

2mM L-glutamine + 2% FBS and the cells incubated with this medium for another hour. The

medium was again removed, replaced by 100 µL of alpha-MEM + 2 mM L-glutamine + 2% FBS

containing the rh-bFGF or the rh-bFGF-CBD to test, and the cells were incubated for 72 hours

under standard conditions.

After this time, 10 µL of MTT labelling reagent (final concentration of 0.5 mg/mL) was

added to each well and the cells were incubated for 4 hours under standard conditions to allow

them to form the formazan crystals. Finally, 100 µL of solubilization solution was added to each

well and the plates were left overnight at 37 ºC to ensure complete lysis of the cells and

solubilization of the formazan crystals. The OD570 of the wells was measured using an ELISA

plate reader and the values transformed into number of cells by interpolation into a standard

curve. Commercially available rh-bFGF (R&D Systems) was used as a positive control.

3.7.3. Inhibition of differentiation assay on MC3T3-E1 mouse

preosteoblasts.

This assay was performed to test the ability of rh-bFGF and rh-bFGF-CBD to inhibit the

differentiation of MC3T3-E1 cells into the osteogenic lineage induced by ascorbic acid. Under

limitant serum conditions and in the presence of ascorbic acid, these cells are known to stop


87

proliferating and initiate their osteogenic differentiation, while the presence of bFGF should

partially block this differentiation.

For this purpose, 10,000 cells/well were seeded on two 96-well culture plates with alpha-

MEM + 2 mM L-glutamine + 10% FBS + 0.2 mM L-ascorbic acid, and incubated for 3 hours

under standard conditions. Afterwards, the medium was removed from the wells, replaced by

alpha-MEM + 2 mM L-glutamine + 2% FBS + 0.2 mM L-ascorbic acid and the cells were

incubated with this medium for another hour. The medium was again removed, replaced by

100 µL of alpha-MEM + 2 mM L-glutamine + 2% FBS + 0.2 mM L-ascorbic acid containing the

rh-bFGF or the rh-bFGF-CBD to test, and the cells were incubated for 120 hours under standard

conditions. One of the plates was used for determination of the number of cells per well by MTT

labelling, as described in section 3.7.2, while the other plate was used for measurement of ALP

activity, as described in section 3.7.1. Finally, the ALP activity per cell was calculated.

3.8. In vivo heterotopic bone formation assay.

To evaluate the biological activity of the rh-bFGF and rh-bFGF-CBD produced in Sf9 insect

cells and their capacity to enhance bone formation in combination with BMP-6, a heterotopic

bone formation assay in rats was used.

For this purpose, an ACS sheet (obtained from highly pure, bovine skin-derived, native

collagen; kindly provided by Dr. M. E. Nimni; US patent 5374539) was cut into discs (5 mm

diameter, 1 mm thickness). Under sterile conditions, 10 µL of growth factor solution, containing

13.89 pmol (0.5 µg) dimeric rh-BMP-6, 1.25 pmol rh-bFGF or rh-bFGF-CBD, or a combination of

both growth factors were loaded on each disc. A set of discs loaded with 10 µL of sterile PBS

was used as a negative control (n=4, for each assayed condition). The assayed conditions are

summarized in Table 8.

Six, four months old, male Wistar rats with a weight of 250-280 g were used for this study.

Animals were anesthetized by an intraperitoneal injection of 2,2,2-tribromo-ethanol (1% in

0.9% NaCl), using 1 mL of anaesthetic solution per 100 g animal weight. Once anesthetized,

the dorsal skin of the animals was disinfected with 70% ethanol and shaved. An incision was

made along the dorsal midline of the skin to expose the underlying dorsal muscles and small

cuts were made in the epimysium and the dorsal muscles to form a small muscular pocket. The

collagen sponges were randomly introduced in these pockets (Fig. 24), the epimysium was

sutured, the skin closed with surgical clamps and the wound disinfected once more with iodine.

Animals were housed in individual cages, with full access to standard food and water, a

controlled temperature of 20±2 ºC and a 12 hour-photoperiod. 21 days later, the animals were

sacrificed by CO2 inhalation and the implants dissected for histological analysis.


88

Condition rhBMP-6 rh-bFGF

(R&D Systems) rh-bFGF rh-bFGF-CBD

1 ●

2 ●

3 ●

4 ●

5 ● ●

6 ● ●

7 ● ●

8 (C-)

Table 8. Combinations of growth factors tested by the heterotopic bone formation assay in rats.

Animal housing and experimentation were carried out according to international

(86/609/EU, 2003/65/CE, 2007/526/CE) and national (RD 1201/2005, LEY 32/2007) laws

concerning animal welfare and scientific experimentation, and were approved by the Committee

for Ethics in Investigation of the University of Málaga.

Figure 24. Implantation of ACS loaded with growth factors into the dorsal muscles of rats for the heterotopic bone formation assay. Asterisk: ACS implanted into a muscular pocket.

3.9. Histological analysis of the implanted ACS.

The dissected implants were fixed with 3.7-4% buffered formaldehyde for 24 hours. After

fixation, the implants that contained rhBMP-6 were decalcified for 4 hours prior to dehydration,

while the rest of the implants were directly dehydrated and embedded in paraffin (see

Appendix I, section AI.6.1).

The samples were transversally cut into 10 µm-thick sections with help of a microtome and

these were placed on poly-L-lysine coated glass slides and incubated at 37 ºC for drying.

*


89

3.9.1. Histochemical stains.

The following histochemical stains were used for analysis of the samples:

Hematoxylin-Eosin. For simple observation of the tissue samples, these were stained

with hematoxylin and eosin (H-E), as detailed in Appendix I, section AI.6.2. By this staining

method, the osteoid appears light pink, while mineralized bone gains a dark pink colour.

Masson’s trichrome. The Masson’s trichrome staining uses three stains to differentiate

muscle, collagen fibers, reticulin and erythrocytes. By use of this technique, the nuclei appear

black, the cytoplasm, muscle and erythrocytes appear red and the collagen and reticulin fibers

appear green. The staining protocol is detailed in Appendix I, section AI.6.3.

Alcian blue. This stain binds to acid and sulphated residues of the glycosaminoglycans

present in the cartilaginous matrix, giving cartilage a bluish colour, while the rest of the tissues

remain unstained. See Appendix I, section 6.4.

3.9.2. Immunohistochemistry.

For determination of the expression of the osteogenic marker osteopontin in the tissue

samples, immunohistochemistry with a specific anti-osteopontin antibody was performed. The

detailed protocol for tissue immunostaining can be found in Appendix I, section AI.6.5.

3.10. Statistical analysis.

All statistic analyses were performed using SigmaStat version 3.1 (Systat Software Inc., San

Jose, CA, USA). Student’s t-test was applied for paired samples comparisons. For multiple

comparison tests, a one-way ANOVA (Holm-Sidak test) was performed.

90

Appendix I. Protocols and recipes.

91

92

__________________________________________________Appendix I. Protocols and recipes

93

AI.1. Buffers of general use.

PBS, 0.1 M:

Na2HPO4 • 2H2O 1.9 g/L

KH2PO4 0.43 g/L

NaCl 7.2 g/L

pH 7.3.

Tris-PBS, 0.1 M:

Na2HPO4 • 2H2O 1.48 g/L

KH2PO4 0.48 g/L

NaCl 7.0 g/L

Tris 5.0 g/L

NaN3 0.2 g/L

pH 7.8.

AI.2. Recombinant DNA technology.

AI.2.1. Total RNA isolation.

Isolation of total RNA content from eukaryotic cells was performed using a commercial kit

(Nucleospin® RNA II, Clontech), following the manufacturers instructions.

The ARN yield of each purification was measured according to the OD260 of the sample,

applying the formula:

[ARN](µg/mL) = OD260 x 40 x dilution factor

The purity and quality of the sample was estimated both by spectrophotometry and RNA

electrophoresis (see Appendix I, section AI.2.6). For spectrophotometric analysis, the

OD260/OD280 relation was calculated. A sample was considered free of contaminant proteins

when this coefficient was higher than 1.8.

Appendix I. Protocols and recipes__________________________________________________

94

AI.2.2. Reverse transcription – polymerase chain reaction.

RT-PCR on the isolated total RNA samples was performed using a commercial kit (Titan™

One Tube RT-PCR System, Roche), which allows sequential retrotranscription and PCR in the

same reaction tube. The list of oligonucleotides used can be found in section AI.2.13.

A first 30 minutes step at 55 ºC allowed synthesis of the cDNA (RT), while a second, 35-

cycle, step is used for amplification of dsDNA (PCR). In the last 25 cycles, the elongation time

had a 5 second/cycle increase.

1) X sec = 60 seconds + ∆ 5 sec/cycle

AI.2.3. Polymerase chain reaction.

PCR reactions were prepared using the 5Prime MasterMix system. For each reaction (25 µL

total volume), 11.5 µL sterile water, 10 µL 5Prime MasterMix, 2.5 µL DNA sample and 0.5 µL of

each 25 pM oligonucleotide solution were mixed in a 0.2 mL PCR tube. PCR was performed

using the Ta specified for each oligonucleotide pair. The oligonucleotides used are listed in

section AI.2.13.

94 ºC 94 ºC 94 ºC 30 sec 30 sec

68 ºC 68 ºC 68 ºC

x10 cycles x25 cycles

1 min 2 min 30 sec Ta 30 sec Ta

55 ºC

30 min

X sec1 7 min

94 ºC 94 ºC 1 min

72 ºC 72 ºC

x30 cycles



95

AI.2.4. Plasmid purification.

For isolation of plasmidic DNA from bacterial cultures a commercial kit (GFX™ Micro Plasmid

Prep Kit, Amersham Biosciences) was used, following the manufacturers instructions. The DNA

yield of each purification was roughly measured by semi-quantitative DNA electrophoresis (see

Appendix I, section A.2.5), comparing the intensity of the plasmid bands with the bands of the

molecular mass standard, each of which contains a known amount of DNA.

AI.2.5. DNA electrophoresis.

DNA electrophoresis was used to visualize any DNA sample (estimation of plasmid yield

after plasmid isolation, evaluation of restriction analyses, analysis of PCR products, etc.).

Gels were prepared by dissolving agarose (1.5% for plasmid visualization or 0.8% for PCR

products visualization) in TAE buffer (40 mM Tris, 20 mM sodium acetate, 1 mM EDTA) with

help of a microwave oven. Ethidium bromide was added to a final concentration of 0.5 µg/mL

and the solution was poured into a gel caster. Once the gel had solidified, it was transferred to

a horizontal electrophoresis chamber filled with TAE buffer.

The samples were prepared by addition of a proper volume of 6x DNA loading buffer

(0.25% (w/v) bromophenol blue, 0.25% (w/v) xylene cyanole, 30% (v/v) glycerol) and loaded

into the wells of the agarose gel. Lambda phage DNA cut with HindIII endonuclease (Sigma-

Aldrich) was used as molecular mass standard for plasmid visualization, while a mixture of low

molecular mass DNA fragments (precision molecular mass standard, BioRad) was used when

running PCR products.

Electrophoretical separation of the samples was performed at 70-100 Volt during 30-90

minutes, and bands were observed by placing the gel on an ultraviolet-light transilluminator.

For detailed analysis, the gel was photographed with help of a gel documentation system

(UVItec).

AI.2.6. RNA electrophoresis.

For RNA visualization, the samples were ran on a denaturing agarose gel. The presence of

formaldehyde and formamide in the gel denatures the secondary structures that most RNA form

via intramolecular base pairing, allowing RNA molecules to migrate strictly according to their

size.


96

Gels were prepared by dissolving 0.3 g agarose in a mixture of 25 mL DEPC-treated, sterile

water, 2 mL 37% formaldehyde and 3 mL sterile 10x MESA buffer (200 mM MOPS, 10 mM

EDTA, 50 mM sodium acetate, pH 7.0), by heating in a microwave oven. Ethidium bromide was

added to a final concentration of 0.5 µg/mL and the solution was poured into a gel caster. Once

the gel had solidified, it was transferred to a horizontal electrophoresis chamber filled with 1x

MESA buffer (20 mM MOPS, 1 mM EDTA, 5 mM sodium acetate, pH 7.0).

The samples were prepared by addition of 3.45 volumes of RNA loading buffer (0.25%

(w/v) bromophenol blue, 50% (v/v) formamide, 6.5% (v/v) formaldehyde, 6% (v/v) glycerol,

10% 10x MESA buffer), denatured for 5 minutes at 70 ºC in a dry block heating thermostat and

loaded into the wells of the agarose gel.

Electrophoretical separation was performed at 70 Volt during 45 minutes, and bands were

observed by placing the gel on an UV light transilluminator. For detailed analysis, the gel was

photographed with help of a gel documentation system (UVItec).

AI.2.7. DNA purification from agarose gels.

For purification of a DNA sample previously separated electrophoretically, the agarose gel

was placed on a 312 nm UV transilluminator to allow visualization of the bands. The band of

interest was cut out with help of a surgical blade and the obtained piece of agarose was

weighed. To remove agarose and ethidium bromide from the sample, a commercial purification

kit (GFX™ PCR DNA and Gel Band Purification Kit, Amersham Biosciences) was used, following

the manufacturers instructions.

AI.2.8. DNA digestion with endonucleases.

Endonuclease enzymes for digestion of DNA molecules were from Amersham Biosciences.

1/10 of the final reaction volume (generally 50 µL) of the proper 10x restriction buffer was

added to the DNA solution. In case the enzyme needed the presence of 0.01% BSA and/or

0.01% Triton X-100, these supplements were also added to the reaction. The volume of the

reaction was adjusted with sterile MilliQ water. A suitable amount of enzyme (10-15 U/µg DNA)

was added, never exceeding the volume of enzyme stock used 1/20 of the reaction volume.

The tubes containing the endonuclease reaction were incubated for at least 2 hours in a 37 ºC

(30 ºC for BamHI restriction) water bath.

Simultaneous digestions with two different endonucleases were only performed when both

enzymes required the same restriction buffer. Else, the DNA was first cut with one enzyme,


97

purified using a commercial kit (GFX™ PCR DNA and Gel Band Purification Kit, Amersham

Biosciences) or precipitated, and finally cut with the second enzyme.

The composition of the different 10x restriction buffers can be found below.

10x L restriction buffer: 100 mM Tris-HCl, pH 7.5, 100 mM MgCl, 10 mM Dithiothreitol.

10x M restriction buffer: 100 mM Tris-HCl, pH 7.5, 100 mM MgCl, 10 mM Dithiothreitol,

500 mM NaCl.

10x H restriction buffer: 500 mM Tris-HCl, pH 7.5, 100 mM MgCl, 10 mM Dithiothreitol,

1 M NaCl.

10x K restriction buffer: 200 mM Tris-HCl, pH 8.5, 100 mM MgCl, 10 mM Dithiothreitol,

500 mM NaCl.

10x T restriction buffer: 330 mM Tris-acetate, pH 7.9, 100 mM Mg-acetate, 5 mM

Dithiothreitol, 660 mM K-acetate.

AI.2.9. DNA precipitation.

For precipitation of DNA in an aqueous solution, 1/10 volume of 3 M sodium acetate, pH 5.2

and 2.5 volumes of pre-chilled absolute ethanol were added to the sample. The sample was

then incubated at -80 ºC for at least 30 minutes before centrifuging at 15,000 xg for 15

minutes at 4 ºC to precipitate de nucleic acids. The supernatant was removed and the DNA

pellet was washed with 500 µL of cold 70% ethanol. The sample was again centrifuged under

the same conditions, ethanol was removed and the pellet was left air-drying before

resuspending it in a suitable volume of MilliQ water.

AI.2.10. DNA ligation.

Ligation of cohesive-ended fragments of DNA into digested plasmids was performed using

the T4 DNA ligase (Usb Corporation). The ligation reactions were prepared by mixing 50 ng of

plasmid with a proper volume of the fragment solution to reach a 1:3 or 1:6 molar proportion.

1 µL (1 U) of T4 DNA ligase and sterile MilliQ water were added to the mixture to complete

10 µL of reaction volume. Different negative controls were prepared for testing both the

plasmid digestion-efficiency and the DNA ligase activity. One control lacked the fragment

(C- VL) and another control lacked both the enzyme and the fragment (C- V).

All reactions were placed in a 16 ºC water bath for 16 hours. After this time, 1 µL of each

sample and each control was used for transformation of E. coli DH5α (see Appendix I, section

AI.3.4). Another aliquot was transformed with 5 ng of non-digested plasmid (C- P). 200 µL of


98

each transformed bacterial aliquot were plated on a LB agar dish and incubated overnight at

37 ºC. The next day, the number of colonies grown on each dish was counted.

The colonies grown on the dish corresponding to the C- V are due to plasmid copies that

had remained circular (i.e. that have not been cut with any endonuclease). Comparing this

number with the number of colonies grown on the C- P dish, a percentage of non-digestion can

be calculated:

% Non-digestion = (C- V) x 100 / (C- P)

On the other hand, the number of colonies grown on the C- VL dish can be used to

calculate the percentage of plasmid copies that had been double-digested:

% Double digestion = 100 – [ (C- VL) x 100 / (C- P) ]

AI.2.11. DNA sequencing.

Sequencing of DNA fragments was performed by the Nucleic Acid Sequencing Service at the

University of Málaga.

AI.2.12. Plasmids.

The plasmids used in this work were:

pBlueScript® II SK(+): Fig. 25. Abbreviated pBIISK, this phagemid is a vector used for

routine cloning and sequencing procedures. The MCS is located inside a lacZ gene fragment,

allowing blue-white screening for plasmids with an insert when using strains containing

lacZ∆M15 on an F’ episome. Bacteria containing wild-type pBSIIK copies will produce blue

colonies when grown in presence of IPTG and X-gal, while bacteria containing the plasmid with

a cloned insert will produce white colonies under the same conditions, due to disruption of the

lacZ gene by the insert.


99

Figure 25. The pBlueScript® II SK(+) vector.

pET17b: Fig. 26. This plasmid is specifically designed for protein expression in E. coli. The

target gene is cloned under control of bacteriophage T7 transcription and translation signals.

When an expression host, carrying a chromosomal copy of the T7 RNA polymerase gene under

lacUV5 control, is transformed with this plasmid, target protein expression can be induced by

IPTG addition.

Figure 26. The pET17b expression vector.

pBIISK (+)

3.0 Kbp

ampR

(1976-2833)

pUC ori (1158-1825)

Plac

Kpn I (653) Xho I (668) Sal I (674) Hind III (689) EcoR V (695) EcoR I (701) Pst I (707) Sma I (713) BamH I (719) Spe I (725) Xba I (731) Not I (737) Eag I (738) BstX I (744) Sac II (747) Sac I (755)

MCS

f1 ori (135-441)

lacZ’

PT7

Xho I (141) Not I (147) BstX I (160) EcoR V (166) EcoR I (174) BstX I (186) Spe I (199) BamH I (205) Ban II (215) Sac I (215) Kpn I (221) Hind III (223) Nhe I (261) Nde I (268) Xba I (306)

MCS

pET17b

3.3 Kbp

ampR

(2241-3098)

ori (1480)


100

pACGP67B: Fig. 27. This plasmid is a shuttle vector for transferring heterologous genes

into baculoviral DNA. It contains the gp67 signal sequence in front of the MCS, and the target

protein will be expressed as a gp67 signal peptide fusion protein under the control of the

baculovirus polyhedrin promoter. Baculovirus DNA flanking the MCS allow homologous

recombination with the baculovirus genome when co-transfected into insect cells.

Figure 27. The pAcGP67B shuttle vector.

AI.2.13. Oligonucleotides.

The oligonucleotides used in this work were:

NUMBER FULL NAME SEQUENCE DESCRIPTION

P1 T7 Promoter Primer 5'-TAATACGACTCACTATAGGG-3' Hybridizes with the pET17b plasmid, upstream of the MCS.

P2 T7 Terminator Primer 5'-GCTAGTTATTGCTCAGCGG-3' Hybridizes with the pET17b plasmid, downstream of the MCS.

P42 Bac1 5’-ACCATCTCGCAAATAAATAAG-3’ Hybridizes with the pAcGP67B plasmid, upstream of the MCS.

P43 Bac3 5’-TCCCAGGAAAGGATCAG-3’ Hybridizes with the pAcGP67B plasmid, downstream of the MCS.

P3 M13 Forward Primer 5’-TGTAAAACGACGGCCAGT-3’ Hybridizes with the pBIISK plasmid, upstream of the MCS.

P4 M13 Reverse Primer 5’-CAGGAAACAGCTATGACC-3’ Hybridizes with the pBIISK plasmid, downstream of the MCS.

P5 RT-BMP6-UP 5’-CCTGGTGGGCAGAGACG-3’ Antisense primer for amplification of the bmp-6 mRNA.

P6 RT-BMP6-DOWN 5’-GAACCAGCTGATCCTTTAGCC-3’ Sense primer for amplification of the bmp-6 mRNA.

P7 BamHI-BMP6-UP 5’-AAGGATCCAGGTCAGCCTCCAG-3’ Antisense primer for amplification of the

bmp-6 gene, introducing the BamHI restriction site.

pAcGP67B

9.7 Kbp

ampR

Ppolyhedrin

Col E ori

BamH I (4258) Xma I (4262) Sma I (4262) Nco I (4268) EcoR I (4274) Not I (4274) Eag I (4285) Pst I (4291) Bgl II (4295)

MCS

gp67 secretion signal


101

NUMBER FULL NAME SEQUENCE DESCRIPTION

P8 EcoRI-BMP6-DOWN 5’-AAGAATTCAGTTAGTGGCATCCAC AAGCTC-3’

Sense primer for amplification of the bmp-6 gene, introducing the EcoRI

restriction site.

P9 BamHI-CBDBMP6-UP

5’-AAGGATCCTGGCGCGAACCGAGCT TCATGGCTCTGAGCGGTGCTAGCAGG

TCAGC-3’

Antisense primer for amplification of the bmp-6 gene, introducing the BamHI

restriction site and the CBD sequence.

P10 EcoRI-BMP6-UP 5’-AAGAATTCAGGTCAGCCTCCAG-3’ Antisense primer for amplification of the

bmp-6 gene, introducing the EcoRI restriction site.

P11 BamHI-BMP6-DOWN

5’-AAGGATCCAGTTAGTGGCATCCAC AAGCTC-3’

Sense primer for amplification of the bmp-6 gene, introducing the BamHI

restriction site.

P12 EcoRI-CBDBMP6-UP 5’-AAGAATTCTGGCGCGAACCGAGCT TCATGGCTCTGAGCGGTGCTAGCAGG

TCAGC-3’

Antisense primer for amplification of the bmp-6 gene, introducing the EcoRI

restriction site and the CBD sequence.

P13 BglII-FGF-UP 5’- TTAGATCTGCAGCCGGGAGCATCACC-3’ Antisense primer for amplification of the

bfgf gene, introducing the BglII restriction site.

P14 EcoRI-FGF-DOWN 5’- AAGAATTCTCAGCTCTTAG CAGACATTGG -3’

Sense primer for amplification of the bfgf gene, introducing the EcoRI

restriction site.

P15 BglII-CBDFGF-UP 5’-TTAGATCTTGGCGCGAACCGAGC

TTCATGGCTCTGAGC-3’

Antisense primer for amplification of the bfgf-CBD gene, introducing the BglII

restriction site.

The restriction sites are highlighted in colours: blue = BamHI; orange = EcoRI;

green = BglII. The sequence of the CBD is highlighted in red.

The Ta used for each specific oligonucleotide pair was:

OLIGONUCLEOTIDE PAIR Ta (ºC)

P1 vs. P2 52.6

P5 vs. P6 56.0

P5 vs. P4 56.2

P10 vs. P11 58.2

P12 vs. P11 59.1

P7 vs. P8 58.2

P9 vs. P8 59.1

P42 vs. P43 54.2

P13 vs. P14 53.5

P15 vs. P14 57.2


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AI.3. Protein expression in E. coli.

AI.3.1. Bacterial cell culture.

All bacterial cultures were incubated at 37 ºC. Liquid cultures were performed in shake

flasks with vigorous shaking (200-300 rpm). The volume of the culture did never exceeded 1/5

of the capacity of the flask to ensure proper oxygenation of the medium. Preparation and

manipulation of the cultures was always performed under semi-sterile conditions, using

sterilized equipment and material and working nearby a Bunsen burner.

AI.3.2. Bacterial cell culture media.

Culture medium for standard liquid culture of E. coli strains (I). Luria-Bertani (LB)

broth, composed of 5 g/L NaCl, 5 g/L yeast extract and 10 g/L tryptone, pH 7.0. This medium

was autoclaved for sterilisation. If necessary, antibiotics and/or supplements were added before

use.

Culture medium for standard liquid culture of E. coli strains (II). 2xYT broth was

used when a higher cell density or faster growth of the cultures was needed. This medium is

composed of 5 g/L NaCl, 10 g/L yeast extract and 20 g/L tryptone, pH 7.0, and was autoclaved

for sterilisation. If necessary, antibiotics and/or supplements were added before use.

Culture medium for protein production in E. coli. Terrific broth (TB), which was

prepared by mixing in a 9:1 proportion the following solutions, previously autoclaved

separately:

YTG Base:

Yeast extract 26.67 g/L

Tryptone 13.33 g/L

Glycerol 8 mL/L

B solution:

KH2PO4 23.1 g/L

K2HPO4 125.4 g/L

Medium was supplemented with antibiotics before use.


103

Culture medium for standard solid-phase culture of E. coli strains. Luria-Bertani

(LB) agar, composed of LB broth and 20 g/L of bacteriological European type agar. After

autoclaving, medium was cooled to a temperature below 60 ºC before adding the proper

antiobiotics and/or supplements and finally poured into sterile, 90 mm diameter, Petri dishes

(approximately 20 mL/dish). Once the medium had solidified, the dishes were stored at 4 ºC

before use.

Ampicillin was prepared as a 25 mg/mL filtered, aqueous stock solution and added to the

media to a 100 µg/mL final concentration. Chloramphenicol was prepared as a 34 mg/mL

filtered stock solution in absolute ethanol and added to the media to a 25 µg/mL final

concentration. IPTG was prepared as a 100 mg/mL filtered, aqueous stock solution and added

to the media to a 240 µg/mL (1 mM) final concentration.

AI.3.3. Bacterial strains.

All the cloning procedures and protein production were performed in Escherichia coli. The

specific strains used in this work were:

DH5α. This strain is a non-expression host, used for general purpose cloning and plasmid

propagation. It has no inherent resistance to any antibiotics.

Rosetta™ (DE3). This strain is a lactose permease (lacY) mutant, deficient in lon and

ompT proteases. It contains a plasmid encoding argU, argW, glyT, ileX, leuW, metT, proL, thrT,

thrU, and tyrU, allowing expression of genes encoding the tRNAs for rare argenine codons AGA,

AGG, and CGA, glycine codon GGA, isoleucine codon AUA, leucine codon CUA, and proline

codon CCC. This accessory plasmid also includes the chloramphenicol resistance gene. These

features make this strain suitable for heterologous protein expression.

AI.3.4. Storage of bacterial clones.

For long-term storage of a bacterial clone, a fresh-grown colony was picked from an LB

agar dish and used to inoculate a 5 mL 2xYT culture, which was incubated overnight at 37 ºC

with vigorous shaking. The next day, 1 mL of the culture was transferred to a microcentrifuge

tube and the cells were pelleted at 10,000 xg for 2 minutes. The supernatant was removed and

the cells were resuspended in 500 µl sterile filtered 50% glycerol and stored at -80 ºC.


104

AI.3.5. Transformation of E. coli strains.

The bacterial strains were transformed with the plasmid of interest by electroporation, by

which internalization of the plasmids by the cells, previously made competent, is triggered by

an electric impulse.

AI.3.5.1. Making of electrocompetents.

To obtain electrocompetent cells, bacteria from a glycerol stock were spread on a Petri dish

with LB agar and incubated overnight at 37 ºC to obtain single colonies. One of these colonies

was picked and used to inoculate a 10 mL LB broth culture, which was also incubated overnight

at 37 ºC. The following day, a proper volume of this culture was used to inoculate 200 mL of

fresh LB broth to obtain an initial OD600 of 0.1. This new culture was incubated at 37 ºC and its

OD600 was measured hourly until a value of 0.5 was reached. At this point, cells were harvested

by centrifugation at 15,000 xg, 15 min, 4 ºC and the pellet was resuspended in 200 mL sterile,

ice-cold milliQ water. Centrifugation was repeated three more times under the same conditions,

resuspending the cells in 100, 20 and 2 mL sterile, ice-cold milliQ water, respectively. Finally,

cells were centrifuged one more time and resuspended in 600 µL pre-chilled, sterile 10%

glycerol. This suspension was aliquoted in 40 µL fractions in microcentrifuge tubes, frozen

immediately in liquid N2 and stored at -80 ºC.

AI.3.5.2. Electroporation.

In general, 100 ng of purified plasmid was used for each electroporation. The aliquots of

competent cells were slowly thawed on ice before adding 1 µL of the 0.1 mg/mL plasmid

solution and mixing by gently pippeting. As a negative control, one aliquot of cells with 1 µL of

sterile milliQ water was used. Cells were left on ice for 1 minute before transferring to an

electroporation cuvette. Electroporation was performed at 2.5 Volt, 200 Ohm of resistance and

25 µF of capacitance. The length of the electric impuls was between 4.5 and 4.8 milliseconds.

After electroporation, each aliquot of cells was resuspended with 960 µL 2xYT (without any

antibiotics or supplemented with 25 µg/mL chloramphenicol when transforming the Rosetta™

(DE3) strain) and transferred to a sterile microcentrifuge tube. Cells were incubated for 45

minutes at 37 ºC with stirring to allow them to start synthesizing the β-lactamase enzyme,

which confers resistance to ampicillin. Finally, 50 and 200 µL of each sample were seeded on an

LB agar dish. The C- cells were pelleted for 3 minutes at 10,000 xg in a microcentrifuge,


105

resuspended with 100 µL 2xYT and seeded on another LB agar dish. Dishes were placed upside

down in a 37 ºC incubator and colonies were allowed to grow for at least 12 hours.

AI.3.6. Colony-PCR.

For rapid selection of colonies of transformed cells, PCR was performed directly on bacteria,

without previous plasmid isolation. PCR reactions were prepared using the 5Prime® MasterMix

system. For each reaction (50 µL total volume), a small portion of a colony was picked using

the tip of a micropipette and resuspended in 29 µL sterile water in a 0.2 mL PCR tube. 20 µL

5Prime® MasterMix and 0.5 µL of each 25 pM oligonucleotide solution were added to each tube

and gently mixed. PCR was performed using the Ta specified for each oligonucleotide pair. The

list of oligonucleotides used can be found in section AI.2.13.

The obtained PCR products were analysed by DNA electrophoresis in agarose gels (see

Appendix I, section AI.2.5).

AI.4. Eukaryotic cell culture.

Manipulation of eukaryotic cells was always done under sterile conditions, working with

sterilized material, equipment and reagents. All cultures and assays involving eukaryotic cells

were performed on a clean bench (Telstar AH100).

94 ºC 94 ºC 1 min

72 ºC 72 ºC

x25 cycles



106

AI.4.1. Cell lines.

U-2 OS cells. ECACC, catalogue number 92022711. This human osteosarcoma cell line was

derived from a moderately differentiated sarcoma of the tibia of a 15 year old girl. U-2 OS cells

are epithelial-like and grow as an adherent monolayer in culture flasks or dishes. They are

known to express high levels of BMP-2, -4, -5, -6 and -7 (Raval P et al., 1996).

Sf9 cells. ECACC, catalogue number 89070101. This cell line was obtained from pupal

ovarian tissue of the fall armyworm Spodoptera frugiperda, by isolation of a clone of the

parental IPLB-SF21-AE cell line (Vaughn JL et al., 1977) by G. Smith and C. Cherry in 1983

(O’Reilly DR et al., 1994). Sf9 cells are highly susceptible to infection with Autographa

californica nuclear polyhedrosis virus (AcNPV), and they are widely used for the production of

recombinant proteins. Sf9 cells are spherical, and can be grown both as an adherent monolayer

in culture flasks or dishes, or as a suspension culture in spinner vessels.

C2C12 cells. ECACC, catalogue number 91031101. This mouse myoblastic cell line was

established by Yaffe D and Saxel O in 1977. C2C12 cells are fibroblast-shaped and grow as an

adherent monolayer in culture flasks or dishes. They differentiate rapidly, forming contractile

myotubes and producing characteristic muscle proteins. Treatment with BMP-2 is known to

cause a shift in their differentiation pathway from myoblastic to osteoblastic (Katagiri T et al.,

1994).

MC3T3-E1 cells. ECACC, catalogue number 99072810. This mouse preosteoblastic cell line

was established from a C57BL/6 mouse calvaria and selected on the basis of high alkaline

phosphatase activity in the resting state. MC3T3-E1 cells are fibroblast-shaped and grow as

adherent monolayers in culture flasks or dishes. They have the capacity to differentiate into

osteoblasts and osteocytes and can form calcified bone tissue in vitro.

AI.4.2. Cell culture media.

Culture medium for U-2 OS osteosarcoma cells. MacCoy’s 5A, which is the medium of

election for most osteosarcoma cell lines. This medium is bought liquid, sterile and with a pH of

6.9. Before use, 1.25 mg/L amphotericin B, 105 U/L penicillin and 100 mg/L streptomycin were

added, and the medium was supplemented with 10% (v/v) heat inactivated FBS and 2 mM

L-glutamine.


107

Culture medium for Sf9 insect cells. Trichoplusia ni Medium – Formulation Hink (TNM-

FH), composed of Grace Medium for insect cells supplemented with trace metals, 3.3 g/L

lactalbumin hydrolysate and 3.3 g/L yeastolate, pH 6.0. This medium is bought in powder form

and reconstituted following the manufacturers indications. Once the medium is sterilized by

filtering through a 0.22 µm filter, 2.5 mg/L amphotericin B, 105 U/L penicillin and 100 mg/L

streptomycin were added. When necessary, the medium was also supplemented with 10% (v/v)

heat inactivated FBS and/or 2 mM L-glutamine.

Culture medium for C2C12 mouse myoblasts. Dulbecco's Modified Eagle's Medium

(DMEM), which is a modification of Basal Medium Eagle (BME), containing four-fold

concentrations of the amino acids and vitamins to improve growth of primary cultures of mouse

and chicken cells. It also contains 4.5 g/L glucose and 110 mg/L sodium pyruvate. This medium

is bought liquid, sterile and with a pH value of 6.9. Before use, 1.25 mg/L amphotericin B,

105 U/L penicillin, 100 mg/L streptomycin and 2 mM L-glutamine were added. When necessary,

the medium was also supplemented with 2 or 10% (v/v) heat inactivated FBS.

Culture medium for MC3T3-E1 mouse preosteoblasts. Alpha-Minimum Essential

Medium (alpha-MEM), which is a modification of standard MEM, supplemented with vitamin B12,

non-essential amino acids, sodium pyruvate, lipoic acid and D-biotin. This medium is bought

liquid, sterile and with a pH of 6.9. Before use, 1.25 mg/L amphotericin B, 105 U/L penicillin,

100 mg/L streptomycin and 2 mM L-glutamine were added. When necessary, the medium was

also supplemented with 2 or 10% (v/v) heat inactivated FBS. For the inhibition of differentiation

assay, the medium was supplemented with 0.2 mM L-ascorbic acid.

AI.4.3. Cell counting and determination of cell viability.

In order to seed the amount of cells required for each experiment, the cell density and the

percentage of living cells in the starting suspension had to be determined. For this purpose, a

5-fold dilution of the suspension in trypan blue was prepared and used to fill both chambers of

a Neubauer haemocytometer. With help of a microscope, the cells on the four corner squares

were counted and the final number was divided by four to obtain the mean number of cells per

square. Since the volume of the column of liquid on top of one square is 1x1x0.1 = 0.1 mm3 =

10-4 mL and the cell suspension was 5-fold diluted, the cell density of the starting suspension

was calculated by multiplying the mean of cells/square by 5 and by 104.

Cells/mL = (total cells / 4) x 5 x 104


108

To estimate the viability of the cells, the living cells (not stained with trypan blue) and the

dead cells (stained blue) were counted separately. The percentage of viability was calculated

applying the formula:

% Viability = (living cells / total cells) x 100

AI.5. Protein analysis.

AI.5.1. Protein precipitation with trichloroacetic acid.

Precipitation of proteins in aqueous solutions with trichloroacetic acid (TCA) can be used for

analysis of low concentration protein samples by SDS-PAGE or immunoblotting (Hames BD and

Rickwood D, 1981). Previous addition of deoxycholate improves the concentration for samples

with protein contents below 1 µg/mL (Peterson GL, 1983).

For every 100 µL of sample, 1 µL of a 20 mg/mL deoxycholate solution was added and the

sample was left on ice for 10 minutes. Afterwards, 43 µL of a pre-chilled 20% TCA solution was

added and the sample was left on ice for another 30 minutes. Proteins were then pelleted at

38,000 xg for 10 minutes at 4 ºC and the pellet was washed with 200 µL cold acetone. The

sample was again centrifuged as before and the acetone was carefully removed. Tubes were

left open at room temperature to eliminate remaining traces of acetone by evaporation. Finally,

the proteinaceous pellet was resuspended in 2x SDS-PAGE Sample Buffer.

AI.5.2. SDS-PAGE.

AI.5.2.1. Buffers and reagents.

4x Laemmli running buffer:

2 M Tris-HCl 50 mL/L

Glycine 58 g/L

10% SDS 40 mL/L


109

2x SDS-PAGE loading buffer:

2 M Tris-HCl, pH 6.8 60 µL/mL

10% SDS 0.4 mL/mL

Glycerol 0.2 mL/mL

Bromophenol blue 1 mg/mL

Stacking gel buffer:

Tris 181.5 g/L

pH 8.8

Resolving gel buffer:

Tris 60 g/L

pH 6.8

AI.5.2.2. Gel preparation.

SDS-PAGE was performed using 12.5 and 15% polyacrylamide gels, with 90 x 60 mm

(WxH) dimensions and 0.75 or 1.5 mm thickness. The equipment used was from BioRad (Mini-

Protean III).

An outer glass and a short plate were assembled and placed in a gel caster. The liquid

resolving gel mixture was prepared on ice to avoid premature polymerization, poured between

the two plates and covered with 50% isopropanol until complete polymerization (approx.

1 hour). Afterwards, isopropanol was removed and the liquid stacking gel mixture was poured

on top of the resolving gel. Immediately after, a 10-well comb was introduced into the stacking

gel and the latter was left at room temperature for polymerization for at least 45 minutes.

Resolving gel 12.5% 15%

30% acrylamide:bis-acrylamide (37.5:1) 4125 µL 4950 µL

MilliQ water 3220 µL 2395 µL

Resolving gel buffer 2500 µL 2500 µL

10% SDS 100 µL 100 µL

10% PSA 50 µL 50 µL

TEMED 5 µL 5 µL


110

Stacking gel 5%

30% acrylamide:bis-acrylamide (37.5:1) 1020 µL

MilliQ water 3420 µL

Stacking gel buffer 1500 µL

10% SDS 60 µL

10% PSA 30 µL

TEMED 6 µL

AI.5.3 Staining of gels with Coomassie blue.


Fixing solution:

Isopropanol 250 mL/L

Acetic acid 100 mL/L

Coomassie blue staining solution:

Coomassie blue G-250 1 g/L

Methanol 500 mL/L


Coomassie blue destaining solution:

Methanol 100 mL/L


AI.5.3.2. Staining protocol.

After electrophoretic separation of the proteins, the gel was incubated for 30 minutes in

fixing solution to avoid protein diffusion through the gel. Afterwards, the proteins were stained

with Coomassie blue staining solution for 45 minutes and the gel was destained overnight in

destaining solution. All incubations were done with gentle agitation on a horizontal shaker.


111

AI.5.4. Electrotransference of proteins to PVDF.


5x Transference buffer:

2 M Tris-HCl 62.5 mL/L

Glycine 72 g/L

1x Transference buffer:

5x Transference buffer 200 mL/L

Methanol 200 mL/L

10% SDS 10 mL/L

AI.5.4.2. Transference protocol.

PVDF sheets were cut into the same dimensions as the polyacrylamide gels, activated by

immersion in methanol for 10 seconds, and washed with transference buffer.

The gel was also equilibrated in transference buffer for 10 minutes, after which the PVDF

membrane was placed on the gel in a transference cassette. Transference was performed at

100 mA, at room temperature, for approximately 3 hours, in transference buffer.

AI.5.5. Staining of proteins on PVDF with amido black.


Amido black staining solution:

Amido black 10B 1 g/L

Methanol 450 mL/L


Amido black destaining solution:

Methanol 500 mL/L



112

AI.5.5.2. Staining protocol.

The sheet of PVDF containing the molecular mass standard proteins was directly incubated

for 5-10 minutes in amido black staining solution after transference from the SDS-PAGE gel.

Afterwards, the sheet was placed in amido black destaining solution for 20-30 minutes with

several renewals of the solution until the excess of stain was removed. The PVDF sheet was

then rinsed with distilled water and placed between sheets of laboratory paper for drying.

AI.5.6. Immunostaining of proteins on PVDF.


PBST:

PBS (see Appendix I, section AI.1) with 0.1% Tween-20.

Blocking solution:

PBST with 2% BSA and 5% non-fat dehydrated milk.

AI.5.6.2. Immunostaining protocol.

Proteins transferred or loaded on PVDF sheets were immunostained using specific

antibodies. All washing and incubating steps were performed at room temperature with gentle

agitation.

The PVDF sheet was washed with PBST (2 x 5 min, 2 x 10 min, 2 x 5 min) before blocking

the sheet with blocking solution (2 x 30 min) to avoid non-specific antibody binding. The sheet

was then washed again with PBST (2 x 5 min) to remove the excess of blocking solution. The

specific primary antibody was diluted at the proper concentration (1:2,500 for anti-BMP-6 and

1:1,000 for anti-bFGF) in PBS + 2% BSA + 0.02% NaN3 and the sheet was incubated overnight

in this solution. The next day, the sample was again washed with PBST (2 x 5 min, 2 x 10 min,

2 x 5 min) and incubated for 1 hour in darkness in the secondary antibody solution (anti-mouse

or anti-rabbit IgG conjugated with HRP, diluted 1:5,000 in blocking solution). From this step on,

the sample was preserved from light to avoid photoinactivation of the peroxidase. The sheet

was washed once more with PBST (2 x 5 min, 2 x 10 min, 2 x 5 min) to remove traces of

unbound secondary antibody, and finally washed with PBS (2 x 5 min).


113

AI.5.7. Development of immunostained proteins.

Detection of the immunostained proteins was performed in a dark room. The PVDF sheet

was placed in a plastic envelope and covered with ECL™ detection reagent (Amersham) for 2-3

minutes. The detection reagent was removed and a Kodak Biomax® chemiluminescence film

was placed on top of the envelope. The exposure time depended on the sample type and

concentration.

After exposure, the film was developed for 5 minutes by submersion in Kodak GBX

developer, washed for 1 minute with water and, fixed for 10 minutes in Kodak GBX fixer.

Finally, the film was extensively washed with water and left drying.

AI.6. Histological analyses.

AI.6.1. Fixation, decalcification, dehydration and embedding in

paraffin.

The fixation of tissue samples to maintain their histological integrity was done by

submersion in buffered 3.7-4.0% formaldehyde for 24 hours, followed by washing with distilled

water for 30 minutes. The samples that were presumed to contain calcified tissue were

decalcified for 4 hours in Decalcifier II®, which is a commercial hydrochloric acid – EDTA

solution, and extensively washed with distilled water for 1 hour, prior to dehydration. The rest

of the samples were directly dehydrated and embedded in paraffin as described below:

- 80º ethanol 30 min at room temperature.





- 1:1 100º ethanol:butanol 60 min at room temperature.

- Butanol 2 x 15 min at room temperature.

- 1:1 butanol:paraffin 120 min at 60 ºC.

- paraffin overnight at 60 ºC.

- paraffin 2 x 120 min at 60 ºC.

Afterwards, the samples were placed into paraffin casts filled with liquid paraffin and left at

room temperature until its complete solidification. Finally, the samples were transversally cut


114

into 10 µm-thick slices whit help of a microtome and these were placed on poly-L-lysine coated

glass slides and incubated at 37 ºC for drying.

AI.6.2. Hematoxylin-eosin staining.

Before staining, the samples were deparaffined with xylene and hydrated. Afterwards,

staining was performed by the following steps:

- Harris hematoxylin 40 sec.

- Ethanol 96º + 8 drops of acetic acid 5 min.

- Running water 5 min.

- Eosin yellowish, hydroalcoholic solution 8 sec.

- Distilled water 1 min.

Afterwards, the samples were dehydrated and mounted with Eukitt.

AI.6.3. Masson’s trichrome staining.



- Bouin liquor 60 min.

- Extensive washing with distilled water

- Ferric hematoxylin 10 min.

- Running water 10 min.


- Scarlet - acid fuchsin 3 min.


- Phosphotungstic acid 5% 15 min.

- Light green 2% 8 min.


- Acetic acid 1% 4 min.



115

Preparation of the different staining solutions was done as detailed below:

Ferric hematoxylin. Solutions A and B were mixed well before use according to the

manufacturer instructions.

Light green.

Light green 2 g

Acetic acid glacial 1 mL

Distilled water 99 mL

AI.6.4. Alcian blue staining.



- Acetic acid 3% 3 min.

- Alcian blue 1 hour

- Running water


Alcian blue.

Alcian blue 1 g

3% acetic acid 100 mL

pH 2.5

AI.6.5. Immunohistochemistry.

Before staining, the samples were deparaffined with xylene and hydrated, and the

endogenous peroxidase activity was inactivated by incubating the samples for 1 hour in

10% H2O2 + 10% methanol in PBS, pH 7.3. The samples were then washed for 3 x 10 minutes

with PBS and saturated with 10% normal sheep serum in PBS for 1 hour.


116

Afterwards, immunostaining of the samples was performed by the following steps:

- PBS 3 x 5 min.

- Primary antibody 18 hours.

- PBS 3 x 5 min.

- Biotinylated secondary antibody 1 hour.

- PBS 3 x 5 min.

- ABC reagent (avidin-biotin complex) 30 min.

The peroxidase activity was developed for 10 minutes with a solution containing 0.2% 3,3’-

diaminobenzidine (DAB) and 0.03% H2O2 in PBS. Then, the nuclei were counterstained with

Harris Hematoxylin for 15 seconds followed by a 7 minutes incubation in ethanol 96º + 8 drops

of acetic acid. Finally, the sections were dehydrated and mounted with Eukitt.

The primary antibodiy used was:

- Anti-osteopontin mouse monoclonal antibody 1:500 in 0.1 M PB + 0.3% triton x-100 +

0.3% BSA

The secondary antibody used was:

- Biotinylated anti-mouse IgG rabbit antibody 1:1,000 in 0.1 M PB + 0.3% triton x-100 +

0.3% BSA

As a negative control, a set of samples was incubated for 18 hours in 0.1 M PB + 0.3%

triton x-100 + 0.3% BSA without primary antibody.

Appendix II. Reagents and equipment.

117

118

_______________________________________________Appendix II. Reagents and equipment

AII.1. Fungibles.

Conical bottom tubes 15 mL and 50 mL (Sterilim) Cryogenic vials 1 mL (377224, Nunc) Culture flask T-75 cm (Nunc) 2

Culture flask T-175 cm (Nunc) 2

Culture plates 6, 12, 24, 48 and 96 wells (Nunc)Dyalisis membranes (D-9652, Sigma) Electroporation cuvettes, Gene Pulser (165-2086, BioRad)®

Filters 0.22 µm, Millex-GP50 (SLGPB5010, Millipore) Haemocytometer (717815, Brand)HiTrap™ Heparin HP columns, 1 mL (17-0406, GE Healthcare) Microcentrifuge tubes 1.5 mL Micropipette filter tips 0.5-10 µL, 5-30 µL, 5-200 µL and 100-1000 µL (Bioscience, Inc) Microscope coverslips (Menzel-Gläser) Microscope glass slides (Thermo Scientific)Pasteur pipettes (Normax) PCR tubes, 0.2 mL (PCR-02D-C, Axygen)Petri dishes for bacterial culture 90 mm (Soria Gelab, S.A.) Petri dishes for cell culture 60, 90 and 150 mm (Nunc)Serological pipettes 2, 5, 10 and 25 mLSuspension culture vessels 1 L (2605-0001, Nalgene) Suspension culture vessels 250 mL (129500, Pobel) Suspension culture vessels 500 mL (129510, Pobel) Syringe filters 0.2 µm (190-2520, Nalgene)Vivaspin 2, 5 kDa MWCO (28-9322-45, GE Healthcare) Ultraviolet spectrophotometry cuvettes (7591 50, Brand)

AII.2. Reagents.

ABC peroxidase staining kit (32020, Pierce) Acetic acid glacial (CH3COOH) (141008, Panreac) Acetone (CH3COCH3) (141007, Panreac) Acrilamide-bisacrilamide solution 30% 37.5:1 (161-0158, BioRad) Agarose (11404, Serva) Agarose, low melting point (A-4018) Alcian blue (A3157, Sigma) Amido black 10B (252036, Panreac) Ammonium acetate (CH3COONH4) (131114, Panreac) Ammonium persulphate (PSA) (161-0700, BioRad) Amphotericin B (A-2942, Sigma) Ampicillin, sodium salt (A-9518, Sigma) Anti-BMP-6 mouse monoclonal antibody (MAB507, R&D Systems) Anti-bFGF rabbit polyclonal antibody (858-450-5558, Calbiochem) Anti-mouse IgG, biotinylated, developed in goat (Ab7067, Abcam) Anti-mouse IgG, HRP-conjugated, developed in rabbit (A-5906, Sigma) Anti-osteopontin mouse monoclonal antibody (MPIIIB10(1), Hybridoma bank) Anti-rabbit IgG, HRP-conjugated, developed in goat (A-6145, Sigma) BacPAK6™ DNA (631401, Clontech)Bacteriological agar (402302, Cultimed)

119

Appendix II. Reagents and equipment_______________________________________________

BamHI endonuclease (E1010V, Amersham) BglII endonuclease (E1021Y, Amersham) Biebrich scarlet-acid fuchsin solution (HT151, Sigma) Bouin liquor (254102, Panreac) Bovine serum albumin (BSA) (12018, Merck) Bromophenol blue (B-8502, Sigma) Butanol (141082, Panreac) Chloramphenicol (C-0378, Sigma) Coomassie brilliant blue G-250 (B-1131, Sigma) Decalcifier II® (00460, Surgipath) Diethyl pyrocarbonate (DEPC) (D-5758, Sigma) Dimethyl sulfoxide (DMSO) (C2H6OS) (D-8779, Sigma) di-Potassium hydrogen phosphate anhydrous (K2HPO4) (121512, Panreac) di-Sodium hydrogen phosphate 2-hydrate (Na2HPO4•2H2O) (122507, Panreac) Dithiothreitol (DTT) (C4H10O2S2) (D-9779, Sigma) DMEM (D-6546, Sigma) Dodecyl sodium sulphate (SDS) (C12H25O4SNa) (142363, Panreac) ECL chemiluminiscence detection reagents (RPN2209, Amersham) EcoRI endonuclease (E1040Y, Amersham) EcoRV endonuclease (10667145001, Roche) Eosin yellowish hydroalcoholic solution 1% (251301, Panreac) Escort transfection reagent (E-9770, Sigma) Ethanol absolute (C2H5OH) (212801, Panreac) Ethanol 96º (212800, Panreac) Ethidium bromide (E-1510, Sigma) Ethylenediaminetetraacetic acid (EDTA) (C10H16N2O8) (131669, Panreac) Ethylenediaminetetraacetic acid (EDTA) (disodium salt) 0.5 M (E-7889, Sigma) Eukitt (O. Kindler GmbH) EZMix™ Terrific broth (T-9179, Sigma) Foetal bovine serum gold (FBS) (A15-649, PAA Laboratories) Formaldehyde 3.7-4.0 % (252931, Panreac) Formaldehyde 36.5-38% (F-8775, Sigma) Formamide (F-7508, Sigma) Gelatin (G-9391, Sigma) GFX™ Micro Plasmid Prep Kit (27-9601-01, Amersham) GFX™ PCR DNA and Gel Band Purification Kit (27-9602-01, Amersham) Glutathione, oxidized form (GSSG) (G-6654, Sigma) Glutathione, reduced form (GSH) (G-4251, Sigma) Glycerol (G-5516, Sigma) Glycine (C2H5NO2) (131340, Panreac) Guanidine hydrochloride (G-3272, Sigma) Harris hematoxylin (253949, Panreac) Hydrochloric acid 37% (HCl) (131019, Panreac) Isopropyl alcohol (C3H8O) (I-0398, Sigma) Isopropyl-β-D-thiogalactopyranoside (IPTG) (I-5502, Sigma) Kodak BioMAx films (Z370398, Sigma) Kodak GBF developing solution (P-7042, Sigma) Kodak GBF fixing solution (P-7167, Sigma) Lambda DNA HindIII digest (D-9780, Sigma) L-arginine (H2NC(=NH)NH(CH2)3CH(NH2)CO2H) (A-5006, Sigma) LB broth (L-3022, Sigma) L-glutamine (G-7513, Sigma) Light green SF yellowish (62110, Sigma) Low fat dehydrated milk (Central Lechera Asturiana) McCoy’s 5A medium (M-8403, Sigma) MEM-alpha (M-8042, Sigma)

120


MEM-alpha (A1049001, Gibco) Methanol (CH3OH) (141091, Panreac) MOPS (M-1254, Sigma) NDSB256 (480010, Calbiochem) Neutral red (N-2889, Sigma) N, N, N', N'-Tetramethyl-1-, 2-diaminomethane (TEMED) (161-0800, BioRad) Normal sheep serum (S-2263, Sigma) Nucleospin® RNA II (635990, Clontech) pAcGP67B shuttle vector (554757, Pharmingen) Paraffin pellets, Histosec® (11609, Merck) pBIISK(+) cloning vector (212205, Stratagene) PCR Mastermix (2200100, 5Prime) Penicilin-streptomicin solution (P-0781, Sigma) pET17b expression vector (69663, Stratagene) PfuTurbo® DNA polymerase (600250, Stratagene) Phosphotungstic acid (121033, Panreac) Poly-L-lysine solution (P-8920, Sigma) Potassium chloride (KCl) (131494, Panreac) Potassium di-hydrogen phosphate (KH2PO4) (131509, Panreac) Precision molecular mass standard (170-8207, BioRad) Proteinase K (E76230Y, Amersham) PVDF sheets, Immobilon-P (IPVH-20200, Millipore) rh-bFGF (233-FB, R&D Systems) rhBMP-6 (507-BP, R&D Systems) Sapphire™ linearized baculovirus DNA (BVD-10001, Orbigen) SDS-PAGE Broad Range molecular weight markers (161-0317, BioRad) Sephadex® G-25, fine (G-2580, Sigma) Sf9 insect cells (71104-3, Novagen) Sigma Fast™ p-nitrophenyl phosphate tablets (N-2770, Sigma) Sodium acetate (C2H3O2Na) (S-2889, Sigma) Sodium azide (NaN3) (122712, Panreac) Sodium bicarbonate (NaHCO3) (S-5761, Sigma) Sodium carbonate (Na2CO3) (6392, Merck) Sodium chloride (NaCl) (141659, Panreac) Sodium deoxycholate (D-6750, Sigma) Sodium di-hydrogen phosphate monohydrate (NaH2PO4•H2O) (6346, Merck) Sodium hydroxide (NaOH) (141687, Panreac) Sodium iodoacetate (I-9148, Sigma) Titan™ One Tube RT-PCR System (11888382001, Roche) TNM-FH (T-1032, Sigma) Tribromoethanol (90710, Fluka) Trichloroacetic acid (TCA) (T-9159, Sigma) Tris (hydroxymethyl)-aminomethane (TRIS) (C4H11NO3) (TR0424, Scharlau) Triton X-100 (t-octylphenoxypoly-ethoxyethanol) (T-8787, Sigma) Trypan blue (T-8154, Sigma) Trypsin-EDTA (T-3924, Sigma) Tryptone (T-9410, Sigma) Tween®20 (Polyoxyethylene sorbitan monolaurate) (P-1379, Sigma) T4 DNA ligase (7005Y, Usb) Urea (131754, Panreac) Weigert’s iron hematoxylin solution (HT1079, Sigma) Xylene (141769, Panreac) Xylene cyanol (X-4126, Sigma) Yeast extract (Y-1625, Sigma) 2-mercaptoethanol (C2H6OS) (M-3148, Sigma) 2-(N-Cyclohexylamino) ethanesulfonic acid (CHES) (C-2885, Sigma) 2-(N-morpholino)ethanesulfonic acid (MES) (M-3671, Sigma)

121

Appendix II. Reagents and equipment_______________________________________________

2-propanol ((CH3)2 CHOH) (141090, Panreac) 2X YT microbial medium (Y-2377, Sigma) 3,3’-diaminobenzidine (DAB) (D-5637, Sigma)

AII.3. Equipment.

Autoclave, Presoclave 75, Selecta. Balance, EB 300, Salter. Chromatographer, Biologic Duo Flow, BioRad. Cell freezing container, Cryo 1ºC, 5100-0001, Nalgene. Centrifuge, MC-15, Cobos. Centrifuge, 320 R, Hettich. Conductimeter, 524, Crison. Container for cell cryopreservation in liquid N2, XC47/11-6, MVE. Electronic propipette, Accu-jet, Brand. Electronic propipette, Pipetboy acu, Integra Biosciences. Electroporator, GenePulser™, BioRad. ELISA plate reader, ELx800, Bio-Tek Instruments. Fixed rotor centrifuge, 3K30, Sigma. Gel documentation system, Uvidoc GAS9000, UVItec. Heating plate, Plan-Tronic, Selecta. Horizontal shaker, OVAN. Horizontal shaker, ProMax 1020, Heidolph. Incubator, Digitronic, Selecta. Incubator (28 ºC, for insect-cell culture), HotCold-S, Selecta. Incubator (37 ºC, for bacterial cultures), 205, Selecta. Incubator/Shaker (37 ºC for bacterial cultures), G25, New Brunswick Scientific. Incubator with CO2 supply (37 ºC, for mammalian cells and tissues), HeraCell, Heraus. Inverted optic microscope, Nikon. Laminar flow cabinet, AV-30/70, Telstar. Lyophilizer, Cryodos-45, Telstar Magnetic shaker, Asincro, Selecta. Magnetic shaker, IKA. Microcentrifuge, 110, Sigma. Micropipettes Eppendorf, Biohit Proline and Boeco. Microtome, HM 340, Microm Heidelberg. Microwave oven, LG. Multiple magnetic shaker, A-04, SBS. Multiple magnetic shaker, A-013, SBS. Multiple micropipette, Biohit Proline. Nucleic acid electrophoresis chamber, Mini-Sub® Cell DT, BioRad. Nucleic acid electrophoresis power supply, EPS300, Pharmacia. Optic microscope, Labophot2, Nikon. Orbital shaker, Heidolph. Pasteur sterilization oven, Conterm 2000 210, Selecta. pHmeter, micropH 2000, Crison. pHmeter, pH211 Microprocessor pH Meter, Hanna Instruments. Precision balance, ER120A, AND. Precision balance, Mettler AM100, German Weber. Protein electrophoresis and electrotransference power supply, PowerPac 300, BioRad. Protein electrophoresis and electrotransference system, MiniProtean III, BioRad. Quantiscan, Biosoft, Ferguson. Spectrophotometer, UV-1603, Shimadzu.

122


Swinging bucket centrifuge, Centronic, Selecta. Swinging bucket centrifuge, Rotofix 32, Hettich. Thermocycler, GeneAmp PCR system 2400, Perkin Elmer. Thermostatic block, BioTDB-100, Boeco. Thermostatic water bath, Digiterm 100, Selecta. Thermostatic water bath, Memmert. Thermostatic water bath, Precisterm, Selecta. Tube shaker, Movil-Rod, Selecta. Ultraviolet light lamp, 312 nm, UVIlite, LF 206 LM, UVItec. Water deionizer, ATAPA 25, ATAPA.

123

124

4. Results.

125

126

________________________________________________________________________Results

127

4.1. Obtaining of the gene encoding hBMP-6.

Total RNA was isolated from cultured U-2 OS human osteosarcoma cells and used as a

template for the amplification of the sequence encoding the mature domain of the hBMP-6 by

RT-PCR. As expected, the RT-PCR yielded a 460 bp band (Fig. 28), which was purified from the

agarose gel and cloned into the pBIISK maintenance vector.

Figure 28. RT-PCR with P5 vs. P6 on U-2 OS total RNA for the amplification of the sequence encoding the mature domain of the hBMP-6.

4.2. Cloning of the genes into the expression vectors.

By PCR, using specific oligonucleotides and the pBIISK:BMP-6 construction as template,

both the sequences of the BMP-6 and the BMP-6-CBD were obtained with an EcoRI restriction

site upstream and a BamHI restriction site downstream (for cloning into the pET17b expression

vector) or with a BamHI restriction site upstream and an EcoRI restriction site downstream (for

cloning into the pAcGP67B shuttle vector). After digestion of these fragments with the proper

endonucleases, they were ligated into the two above mentioned plasmids. Cloning of the

fragments was confirmed by PCR analysis using oligonucleotides against the sequences flanking

the MCSs of the plasmids (Fig. 29 A and B).

Figure 29. PCR analysis of the obtained expression vectors. A) PCR with P1 vs. P2 on the pET17b:BMP-6 (lane 1) and pET17b:BMP-6-CBD (lane 2) constructions. B) PCR with P42 vs. P43 on the pAcGP67B:BMP-6 (lane 1) and pAcGP67B:BMP-6-CBD (lane 2) constructions. C) PCR with P42 vs. P43 on the pAcGP67B:bFGF (lane 1) and pAcGP67B:bFGF-CBD (lane 2) constructions.

500 bp

1 Kbp

700 bp

1 Kbp

500 bp

700 bp

700 bp

1 Kbp

A B C 1 2 1 2 1 2

Results________________________________________________________________________

128

For the cloning of the sequences encoding the rh-bFGF and the rh-bFGF-CBD into the

pAcGP67B shuttle vector, a PCR with specific oligonucleotides was performed on the

pET28b:hbFGF-F1 and pET28b:hbFGF-F2 constructs to obtain the fragments with a BglII

restriction site upstream and an EcoRI restriction site downstream. After digestion of the

fragments with the proper endonucleases, they were ligated into the pAcGP67B shuttle vector

and the cloning was confirmed by PCR analysis using oligonucleotides against the sequences

flanking the MCS of the plasmid (Fig. 29 C).

4.3. Production of rhBMP-6 in Escherichia coli.

Since Escherichia coli is known to be a potent system for the production of heterologous

proteins, and other members of the BMP family have been successfully produced using this tool

(Vallejo LF et al., 2002), even with an additional collagen binding domain (Chen B et al., 2007a;

Chen B et al., 2007b; Visser R et al., 2009), the first attempts on rhBMP-6 and rhBMP-6-CBD

production were made using this system.

4.3.1. Obtaining of rhBMP-6-expressing clones of E. coli

Rosetta™ (DE3).

E. coli Rosetta™ (DE3) cells were made competent and transformed with the pET17b:

BMP-6 and the pET17b:BMP6-CBD constructions. The transformed cells were plated on LB agar

dishes supplemented with antibiotics and ten of the obtained colonies for each expression

vector were analyzed by colony-PCR using specific primers against the rhBMP-6 or rhBMP-6-

CBD inserts. All of the analyzed colonies resulted to be positive for their respective

constructions.

Five positive colonies for each expression vector were grown in liquid cultures and their

plasmids were isolated. Two plasmid samples for each expression vector were sequenced using

specific primers against the regions of the plasmidic DNA flanking the insert. Since the

sequence of both genes was correct, with no mutations being found, and they were all cloned

in frame, one clone for each expression vector was selected for protein expression.

________________________________________________________________________Results

129

4.3.2. Expression of rhBMP-6 in Escherichia coli.

For rhBMP-6 production, the selected E. coli Rosetta™ (DE3) clone, containing the

recombinant pET17b expression vector, was grown in TB with a starting OD600 of 0.1, until an

OD600 of 0.8 was reached. At this point, IPTG was added to the culture to induce the expression

of the heterologous proteins, and the production was continued for four hours, taking samples

and measuring the OD600 of the culture hourly. During this period, a clear variation of the

growth rate could be observed after addition of IPTG when compared to a control culture of

non-transformed bacteria (Fig. 30 A). While the control culture continued its exponential growth

during the entire analyzed period, the rhBMP-6-expressing culture slowed its growth after the

addition of IPTG, as showed by the lowering of the slope of the growth curve. This was

presumed to be a direct consequence of a high rate of heterologous protein expression, which

should interfere with the normal physiological behaviour of the bacteria in a highly rich culture

medium such as TB.

Four hours after the induction of heterologous protein expression, the cells of the culture

were harvested, obtaining approximately 612 mg (wet weight) of bacterial biomass. A 1 mL

aliquot of the culture was used for SDS-PAGE analysis of the protein production procedure. For

this purpose, the cells were lysed by sonication, the soluble protein fraction was separated from

the insoluble protein fraction, and the produced inclusion bodies were subsequently isolated

from the latter by washing in presence of triton x-100. As shown in figure 30 B, before the

addition of IPTG to the culture, the total protein content of the bacteria included one most

represented protein that migrated as a ±16.5 KDa band in the polyacrylamide gel. The facts

that this band did not appear when analyzing the total protein content of non-transformed

control cultures, and that its molecular mass corresponded to that of non-glycosilated rhBMP-6

monomers, made us assume that this was our goal protein. The presence of rhBMP-6 in the

bacteria before the induction of heterologous protein expression showed that the chosen

expression system has a low stringency, and that the T7 RNA polymerase responsible for

heterologous protein expression is present in the host cells at basal levels in the absence of

lactose or its analogue IPTG.

After IPTG addition to the cultures, a significant increase of the band corresponding to the

rhBMP-6 and of the rhBMP-6/total protein ratio could be observed. This increase was especially

reached during the first hour after induction, being the protein expression rate lower during the

following three hours. Four hours after IPTG addition, the rhBMP-6 monomers were estimated

to represent 40-50% of the total protein content of the bacteria.

The high amounts of rhBMP-6 monomers produced and the fact that they could be

recovered in the insoluble protein fraction made us assume that they were accumulated in the

form of inclusion bodies. Washing of these inclusion bodies in the presence of triton x-100 to

Results________________________________________________________________________

130

remove contaminant, insoluble, membrane-associated proteins yielded a highly pure sample of

inclusion bodies containing the rhBMP-6 monomers.

Figure 30. rhBMP-6 production in Escherichia coli. A) Growth of a not transformed control culture (blue line) versus a culture expressing rhBMP-6 (red line). The arrow indicates addition of IPTG to the culture to induce recombinant protein expression. B) Cell protein content analyzed by SDS-PAGE and Coomassie blue staining. Lane 1, total protein content before induction of expression. Lane 2, total protein content 1 hour after induction of expression. Lane 3, total protein content 2 hours after induction of expression. Lane 4, total protein content 3 hours after induction of expression. Lane 5, total protein content 4 hours after induction of expression. Lane 6, insoluble protein content 4 hours after induction of expression. Lane 7, washed inclusion bodies.

The obtained inclusion bodies were solubilized with 6 M Gnd-HCl and finally dialyzed against

MES-Gnd buffer, yielding a 4 mL solution. Quantification of the rhBMP-6 in this sample by SDS-

PAGE and digital image analysis revealed that the concentration of rhBMP-6 was approximately

2.7 mg/mL and that the yield of monomeric rhBMP-6 production with the used expression

system and host cell strain was, thus, approximately 108 mg/L.

4.3.3. Refolding of rhBMP-6 produced in Escherichia coli.

Since the active form of rhBMP-6 is a homodimer with one intercatenary disulfide bond and

three intracatenary disulfide bonds per monomer, in vitro refolding of the produced monomers

became a need. BMPs do not spontaneously acquire their native conformation when the

chaotropic agent is removed from the solubilization buffer, so a complex refolding procedure

has to be performed, in which many different variables have to be taken into account (i.e.

protein concentration, refolding time, temperature, pH, redox pairs, antiaggregants, chaotropic

0

2

4

6

8

10

12

14

16

0 2 4 6 8 10 12

OD600

Time (hours)

1 2 3 4 5 6 7

14 KDa

21 KDa

31 KDa

45 KDa

66 KDa

rhBMP-6

A B

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131

agent concentration, presence of molecular oxygen in the refolding mixture, etc). The starting

conditions assayed for rhBMP-6 refolding were based on those described for the successful

refolding of rhBMP-2 by Vallejo LF et al., 2002, which also resulted useful for refolding of a

collagen-targeted rhBMP-2 (Visser R et al., 2009). These conditions were refolding for 72 hours

at 10 ºC and a protein concentration of 20 µg/mL in a refolding buffer containing 57.5 mM Tris,

pH 8.5, 0.55 mM EDTA, 0.9 M NaCl, 0.75 M CHES, 0.5 M Gnd-HCl, 2 mM GSH and 1 mM GSSG.

The first attempt on rhBMP-6 refolding was done using these conditions and testing the

possible effect of three different GSH :GSSG ratios.

SDS-PAGE analysis of the refolding procedure showed that, in the three cases, most of the

rhBMP-6 remained in the monomeric form (Fig. 31). Besides the ±16.5 KDa band, a

characteristic ladder appeared, formed of a ±32 KDa, a ±47 KDa, a ±63 KDa and higher bands.

Since the molecular mass of these bands could correspond to dimeric, trimeric, tetrameric and

polymeric forms of rhBMP-6, respectively, they were assumed to be due to unspecific

intermolecular disulfide bond formation. This was confirmed by analyzing these samples by

SDS-PAGE in the presence of DTT, in which the higher bands disappeared and only the

16.5 KDa band could be seen.

Figure 31. Effect of GSH:GSSG ratio on in vitro refolding of rhBMP-6 expressed in Escherichia coli. Monomers (x1) give rise to dimers (x2), trimers (x3), tetramers (x4) and higher polymers. Lane 1, GSH:GSSG ratio 2:1. Lane 2, GSH:GSSG ratio 10:1. Lane 3, GSH:GSSG ratio 40:1.

Since our aim was to obtain a better refolding yield and a more specific disulfide bond

establishment, different variations on the previous standard conditions were assayed. In total,

41 combinations of parameters were tested and analyzed by SDS-PAGE (Figs. 32, 33, 34, 35).

1 2 3

14 KDa

21 KDa

31 KDa

45 KDa

66 KDa

x1

x2

x3

x4

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Figure 32. Effect of antiaggregants, pH and GSH:GSSG ratio on in vitro refolding of rhBMP-6 expressed in Escherichia coli. A) Refolding in presence of NDSB256. Lane 1, 0.5 M NDSB256, pH 7.5, GSH:GSSG ratio 2:1. Lane 2, 0.5 M NDSB256, pH 7.5, GSH:GSSG ratio 40:1. Lane 3, 0.5 M NDSB256, pH 8.5, GSH:GSSG ratio 2:1. Lane 4, 0.5 M NDSB256, pH 8.5, GSH:GSSG ratio 10:1. Lane 5, 0.5 M NDSB256, pH 8.5, GSH:GSSG ratio 40:1. Lane 6, 0.5 M NDSB256, pH 9.5, GSH:GSSG ratio 2:1. Lane 7, 0.5 M NDSB256, pH 9.5, GSH:GSSG ratio 40:1. Lane 8, 1 M NDSB256, pH 7.5, GSH:GSSG ratio 2:1. Lane 9, 1 M NDSB256, pH 7.5, GSH:GSSG ratio 40:1. Lane 10, 1 M NDSB256, pH 8.5, GSH:GSSG ratio 2:1. Lane 11, 1 M NDSB256, pH 8.5, GSH:GSSG ratio 10:1. Lane 12, 1 M NDSB256, pH 8.5, GSH:GSSG ratio 40:1. Lane 13, 1 M NDSB256, pH 9.5, GSH:GSSG ratio 2:1. Lane 14, 1 M NDSB256, pH 9.5, GSH:GSSG ratio 40:1. B) Refolding in presence of arginine. Lane 1, 0.5 M arginine, pH 7.5, GSH:GSSG ratio 40:1. Lane 2, 0.5 M arginine, pH 8.5, GSH:GSSG ratio 2:1. Lane 3, 0.5 M arginine, pH 8.5, GSH:GSSG ratio 10:1. Lane 4, 0.5 M arginine, pH 8.5, GSH:GSSG ratio 40:1. Lane 5, 0.5 M arginine, pH 9.5, GSH:GSSG ratio 40:1. Lane 6, 1.5 M arginine, pH 7.5, GSH:GSSG ratio 2:1. Lane 7, 1.5 M arginine, pH 7.5, GSH:GSSG ratio 40:1. Lane 8, 1.5 M arginine, pH 8.5, GSH:GSSG ratio 2:1. Lane 9, 1.5 M arginine, pH 8.5, GSH:GSSG ratio 10:1. Lane 10, 1.5 M arginine, pH 8.5, GSH:GSSG ratio 40:1. Lane 11, 1.5 M arginine, pH 9.5, GSH:GSSG ratio 40:1.

Figure 33. Effect of protein concentration and GSH:GSSG ratio on in vitro refolding of rhBMP-6 expressed in Escherichia coli. A) Refolding of rhBMP-6 at low concentration (10.7 µm/mL). Lane 1, GSH:GSSG ratio 40:1. Lane 2, GSH:GSSG ratio 10:1. Lane 3, GSH:GSSG ratio 2:1. B) Refolding of rhBMP-6 at high concentration (53.4 µm/mL). Lane 1, GSH:GSSG ratio 40:1. Lane 2, GSH:GSSG ratio 10:1. Lane 3, GSH:GSSG ratio 2:1.

1 2 3 4 5 6 7 8 9 10 11 12 13 14 1 2 3 4 5 6 7 8 9 10 11

A B

14 KDa

21 KDa

31 KDa

45 KDa

66 KDa

97 KDa

1 2 3 1 2 3

A B

14 KDa

21 KDa

31 KDa

45 KDa

66 KDa

97 KDa

x1

x2

x3

x4

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Figure 34. Effect of redox pair, redox pair concentration and N2 supply on in vitro refolding of rhBMP-6 expressed in Escherichia coli. A) Refolding of rhBMP-6 with 1 mM of GSH:GSSG, without N2 supply. Lane 1, GSH:GSSG ratio 2:1. Lane 2, GSH:GSSG ratio 10:1. B) Refolding of rhBMP-6 with 3 mM of GSH:GSSG, with continuous N2 supply. Lane 1, GSH:GSSG ratio 2:1. Lane 2, GSH:GSSG ratio 10:1. C) Refolding of rhBMP-6 with 3 mM of 4-MPAA:GSSG with continuous N2 supply. Lane 1, 4-MPAA:GSSG ratio 2:1. Lane 2, 4-MPAA:GSSG ratio 10:1.

Figure 35. Effect of the temperature on in vitro refolding of rhBMP-6 expressed in Escherichia coli. Lane 1, refolding performed at 10 ºC. Lane 2, refolding performed at 20 ºC.

None of the assayed combinations was able to produce a satisfactory yield of rhBMP-6

dimers. In all cases, the previously described ladder of bands appeared, though the degree of

polymerization versus dimerization varied among the different conditions used. For example,

the highest levels of polymerization (unspecific disufide bond formation) were obtained when

1 2 1 2 1 2

A B C

14 KDa

21 KDa

31 KDa

45 KDa

66 KDa

97 KDa

x1

x2

x4

x3

1 2

x1

x2

X3

14 KDa

21 KDa

31 KDa

45 KDa

66 KDa

Results________________________________________________________________________

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using 1 M NDSB 256 or 0.5 M L-arginine as antiaggregants (Fig. 32) or when the protein

concentration was increased (Fig. 33 B).

4.4. Production of rhBMP-6 and rhBMP-6-CBD in Sf9 cells.

Since the refolding of the rhBMP-6 produced in Escherichia coli was unsuccessful, we

decided to try the production of these proteins in a eukaryotic expression system, in which the

folding of the monomers to obtain the native quaternary structure of the protein occurs within

the cell. The observation that rhBMP-6 monomers seem to be highly susceptible to form

polymers through unspecific disulfide bonds led us to use a baculoviral expression system in

which our goal proteins were co-expressed with the PDI. We hypothesized that this enzyme

could be helpful to shuffle incorrect disulfides into their correct pairings, what would improve

the yield of correctly folded dimers. Furthermore, in order to facilitate the recovery and

posterior purification of the produced proteins, we used a system by which the goal proteins are

produced with a fused signal peptide which directs them through the endoplasmic reticulum

and the Golgi apparatus for secretion into the culture medium. This signal peptide is excised

from the molecule by specific endopeptidases of the host cell before secretion.

4.4.1. Obtaining of rhBMP-6 and rhBMP-6-CBD expressing clones of

baculoviruses.

Once the genes encoding the rhBMP-6 and rhBMP-6-CBD were correctly cloned into the

pAcGP67B plasmid, Sf9 cells were co-transfected with the recombinant donor plasmid and

linearized Sapphire™ baculoviral DNA in order to obtain infective, heterologous protein-

expressing baculoviruses by homologous recombination. Eight days after transfection, the

cultures showed clear signs of infection (i.e., presence of both enlarged and lysed cells)

(Fig. 36), and the transfection supernatants (TS) were harvested. These TS are heterogeneous

mixtures of recombinant baculoviruses, cell debris and DNA molecules derived from the

degradation of donor plasmid and baculoviral DNA molecules.

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135

Figure 36. Sf9 cells eight days after co-transfection with pAcGP67B:rhBMP-6 and Sapphire™ linearized baculoviral DNA. A) Control culture transfected only with the donor plasmid. B) Culture co-transfected with the donor plasmid and the baculoviral DNA. Small arrows point to lysed cells. Arrowheads point to cells with increased diameter, which are in the late phase of the infection cycle.

To ensure the reproducible production of the proteins, these must be expressed by a single

baculoviral clone. In order to obtain pure clones of the recombinant baculoviruses, the TS were

used to perform a plaque assay, infecting cultures with 10-3 to 10-8 dilutions and covering the

cells with agarose. Five days p.i., the cultures were stained with neutral red to identify the lysis

plaques (Fig. 37 A), which were found in the cultures infected with 10-3 to 10-6 dilutions of the

TS. Ten well-isolated lysis plaques were picked and the viruses they contained were analyzed

by PCR using specific primers against the viral DNA flanking the inserts (Fig. 37 B). Since the

lysis plaques were picked from cultures infected with high dilutions of the TS and the selected

plaques were small, round-shaped and well isolated from others, it was assumed that each of

them corresponded to one single viral clone.

One clone for each protein was selected and used for amplification to obtain a master stock

(MSV), which was then titered by a limit dilution assay. The calculated titer of the MSV of

rhBMP-6 was 4.35 x 109 pfu/mL, while the titer of the MSV of rhBMP-6-CBD was 1.16 x 1010

pfu/mL. Although the viruses isolated by the plaque assay were roughly analyzed by PCR to

confirm that they had the heterologous genes inserted into the viral DNA, a second PCR

analysis was performed on the master stocks. In this case, PCR reactions on both clones were

performed using a specific primer against the 3’-end of the bmp6 gene and a primer against the

5’-end of the gene or against the sequence of the CBD (Fig. 38). Furthermore, both clones were

sequenced with the primers against the viral DNA flanking the inserts to ensure that no

mutations or frame-shifting had occurred during the generation of the recombinant

baculoviruses.

A B

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Figure 37. Isolation of baculoviral clones. A) Plaque assay for isolation of clones expressing rhBMP-6-CBD. The arrows point to the lysis plaques. B) PCR analysis of the selected clones using primers that hybridize with the baculoviral DNA flanking the insert. Lane 1, analysis of a clone expressing rhBMP-6. Lane 2, analysis of a clone expressing rhBMP-6-CBD.

Figure 38. PCR analysis of the isolated baculoviral clones. Lanes 1 and 2, Analysis of the rh-BMP6-CBD expressing clone with specific primers against the CBD and the BMP-6 sequence (P9 vs. P8, and P7 vs. P8), respectively. Lanes 3 and 4, Analysis of the rh-BMP6 expressing clone with specific primers against the CBD and the BMP-6 sequence (P9 vs. P8, and P7 vs. P8), respectively.

To obtain a high volume virus stock (HVVS) with which all the future protein productions

can be made, the MS were amplified by infecting cultures with a MOI of 0.1. This low MOI

allows the viral population to grow exponentially before the majority of the cells die, yielding a

high-titer suspension of virus. Finally, the obtained HVVS were titered to allow us to perform

the protein productions with a known and previously established MOI. The calculated titer of

the HVVS of rhBMP-6 was 3.48 x 108 pfu/mL, while the titer of the HVVS of rhBMP-6-CBD was

estimated to be 4.92 x 1014 pfu/mL.

1 Kbp

700 pb

1 2 A B

500 bp

200 bp

1 2 3 4

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137

4.4.2. Production assays for rhBMP-6 and rhBMP-6-CBD.

To determine the best conditions for large scale protein production, a production assay was

performed with the two HVVS obtained. This was done infecting Sf9 cultures with a MOI of 2.5

or 10, and harvesting the medium of the cultures 72, 96, 120 and 144 hours p.i. (i.e.,

production times of 48, 72, 96 and 120 hours). Since previous works carried out by our group

on the production of other BMPs in Sf9 cells already demonstrated that the yield of this

expression system is not high enough to detect the heterologous proteins secreted into the

culture medium by SDS-PAGE and Coomassie blue staining, 400 µL of the harvested

conditioned media were precipitated with TCA and the samples analyzed by SDS-PAGE and

Western blot.

4.4.2.1. Production assay for rhBMP-6.

In the case of rhBMP-6, immunostaining of the proteins with a specific anti-BMP-6 antibody

revealed the presence of at least three proteins in the culture media which were recognized by

the antibody (Fig. 39). The rhBMP-6 produced in CHO cells, used as a C+, appeared as a triplet

of bands, with the central band having a MW of ±37 KDa. In the analyzed samples, the lowest

of the bands also had an estimated MW of ±37 KDa and could correspond to the dimeric,

glycosilated form of rhBMP-6. The intensity of this band increased from 48 to 72 hours and

decreased afterwards as the production time progressed. A second band, with an estimated

MW of ±56 KDa, was also found in all of the analyzed samples. This band, which intensity

slightly increased from 48 to 72 hours and then remained stable, could correspond to rhBMP-6

trimers. A third band, with an estimated MW of ±73 KDa, appeared slightly in the samples

corresponding to cells infected with a MOI of 2.5 but was more intense in the samples of cells

infected with a MOI of 10. This band could correspond to a tetrameric organization of rhBMP-6

molecules.

To determine whether the proteins observed in the Western blot analysis of the production

assay were disulfide bonded di-, tri-, tetra-, or polymers, or other non BMP-related proteins, the

samples of the cultures infected with a MOI of 10 were analyzed by western blot under

reducing conditions (Fig. 40). In this analysis it became clear that the three previously

described bands are more or less unique with 48 and 72 hours of production time, but that they

are not with longer production times. With 120 hours of production time, instead of a single

±37 KDa band, a triplet of bands (±34, ±37 and ±40 KDa) appeared.

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Figure 39. Production assay for rhBMP-6, analyzed by Western blot. Lane 1, rhBMP-6 produced in CHO cells (R&D Systems). Lanes 2-5, rhBMP-6 produced with a MOI of 2.5 and 48, 72, 96 and 120 hours production time, respectively. Lanes 6-9, rhBMP-6 produced with a MOI of 10 and 48, 72, 96 and 120 hours production time, respectively.

In the presence of DTT, the three unique bands previously described for the 48 and 72

hours production time conditions disappeared, and were substituted by a doublet of bands (±18

and ±23 KDa), which molecular mass could correspond to that of the rhBMP-6 monomer. This

fact demonstrated that the higher molecular mass proteins observed under non-reducing

conditions are disulfide bonded proteins. This doublet also appeared after 96 and 120 hours,

although a great variety of higher molecular mass bands could also be observed in these

samples.

Figure 40. Western blot analysis with reducing agents of the rhBMP-6 produced in Sf9 cells. Lane 1, rhBMP-6 produced in CHO cells (R&D Systems). Lanes 2-5, rhBMP-6 produced with a MOI of 10 and 48, 72, 96 and 120 hours production time, respectively. Lanes 6-9, rhBMP-6 produced with a MOI of 10 and 48, 72, 96 and 120 hours production time, respectively, in the presence of DTT.

1 2 3 4 5 6 7 8 9

31 KDa

45 KDa

66 KDa

97 KDa

21 KDa

1 2 3 4 5 6 7 8 9

31 KDa

45 KDa

66 KDa

97 KDa

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139

According to these results, the best conditions for rhBMP-6 production were determined to

be infecting with a MOI of 10 and harvesting the conditioned medium after 48-72 hours of

production time since, under these conditions, the highest yields of the ±37 KDa protein

(rhBMP-6 dimers) were found, with less contamination of tri-, tetra-, or polymers, or other non

BMP-related proteins.

4.4.2.2. Production assay for rhBMP-6-CBD.

In the case of rhBMP-6-CBD, immunostaining with the anti-BMP-6 antibody also revealed

three bands (Fig. 41). The lowest of these had an estimated MW of ±39 KDa and could

correspond to the dimeric, glycosilated form of rhBMP-6-CBD, which predicted MW is 38.56

KDa. The intensity of this band increased from 48 to 72 hours and decreased afterwards as the

production time progressed, has happened with the production assay analysis for rhBMP-6. The

second band had an estimated MW of ±58 KDa and the third band had an estimated MW of

±77 KDa. These bands could correspond to a trimeric and tetrameric organization of rhBMP-6-

CBD, respectively, and the evolution of their intensity with production time mimicked that of the

±39 KDa band. Under all circumstances, the intensity of the ±58 KDa band was higher than

that of the lower, ±39 KDa band

Figure 41. Production assay for rhBMP-6-CBD, analyzed by Western blot. Lane 1, rhBMP-6 produced in CHO cells (R&D Systems). Lanes 2-5, rhBMP-6-CBD produced with a MOI of 2.5 and 48, 72, 96 and 120 hours production time, respectively. Lanes 6-9, rhBMP-6-CBD produced with a MOI of 10 and 48, 72, 96 and 120 hours production time, respectively.

According to these results, the best conditions for rhBMP-6-CBD production were

determined to be infecting with a MOI of 10 and harvesting the conditioned medium after 72

hours of production time since, under these conditions, the highest yields of the ±39 KDa

1 2 3 4 5 6 7 8 9

31 KDa

45 KDa

66 KDa

97 KDa

Results________________________________________________________________________

140

protein (rhBMP-6-CBD dimers) were found, although the amounts of the other detected

proteins was also very high.

4.4.3. Analysis of the influence of the PDI on rhBMP-6 production in

Sf9 cells.

To determine if the excessive production of disulfide bonded polymers of rhBMP-6 was due

to the co-expression of the PDI by the Sapphire™ baculovirus, Sf9 cells were co-transfected

with the pAcGP67B:BMP-6 donor plasmid and BacPak6™ linearized baculoviral DNA in order to

obtain recombinant baculoviruses to express rhBMP-6 without co-expression of the PDI

chaperone. Eight days after co-transfection, the cells showed signs of infection, so the culture

medium was harvested and 1 mL was used to infect 6 x 106 Sf9 cells seeded on a 90 mm Petri

dish with the aim of amplifying the generated recombinant baculoviruses. Four days p.i., the

culture medium was harvested and used for a production assay for rhBMP-6. For this purpose,

Sf9 cells were seeded on a 24 well culture dish at a density of 5 x 105 cells/well and infected

with 25 µL of serial dilutions of the obtained baculovirus suspension, from 0 to 10-6. 96 hours

p.i. (72 hours production time), the conditioned media were harvested and analyzed by

Western blot (Fig. 42).

Figure 42. Production assay for rhBMP-6 expressed by BacPak6™ baculoviruses. Lane 1, uninfected control. Lane 2, rhBMP-6 produced in CHO cells (R&D Systems). Lanes 3-9, cultures infected with 0 to 10-6 dilutions of the viral suspension, respectively.

1 2 3 4 5 6 7 8 9

31 KDa

45 KDa

66 KDa

97 KDa

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141

The cultures infected with the non-diluted viral suspension or with 10-1 or 10-2 dilutions

yielded the same dimer:polymer ratio as the rhBMP-6-expressing Sapphire™ baculoviruses, with

a great proportion of trimers. In contrast, no bands were recognized by the anti-BMP-6

antibody in the uninfected cultures.

These results indicate that the PDI is not responsible for polymer formation during rhBMP-6

expression in Sf9 cells. Nevertheless, since the PDI might be helpful for disulfide scrambling

among the dimeric rhBMP-6 fraction, what would increase the number of correctly folded

molecules in this population, the rest of the experiments were done using the rhBMP-6 and

rhBMP-6-CBD expressing Sapphire™ baculoviruses.

4.4.4. Expression and purification of rhBMP-6 and rhBMP-6-CBD.

Once the best conditions for rhBMP-6 and rhBMP-6-CBD had been established, larger scale

productions of both proteins were performed, culturing the cells at a density of 4 x 107

cells/flask in T-175 cm2 flasks and infecting them with a MOI of 10. 96 hours p.i. (72 hours

production time), the conditioned media were harvested and 6 M urea was added to prevent

protein precipitation within the purification column.

Purification of the protein was done by heparin-affinity chromatography, since the BMPs are

known to possess an amino-terminal heparin-binding site (Ruppert R et al., 1996). After

washing of the column and loading of the sample, a two-step elution of the proteins was

performed, using elution buffers with conductivity values of 31 mS/cm (0.43 M NaCl) and

63 mS/cm (1 M NaCl).

4.4.4.1. Purification of rhBMP-6.

The purification profile for rhBMP-6 showed one clear peak of the OD280 for each

conductivity value, indicating that at least two protein populations with different affinities to

heparin were separated (Fig. 43).

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142

Figure 43. Purification by heparin-sepharose chromatography of rhBMP-6 expressed in Sf9 cells. The analyzed collection fractions are highlighted in a grey box.

The elution fractions corresponding to both OD280 peaks were analyzed by Western blot, as

well as the non-purified conditioned medium supernatant and the flow-through fraction

(Fig 44). Immunostaining with a specific anti-BMP-6 antibody showed the presence of the three

previously described bands in the conditioned medium supernatant, which are assumed to

correspond to rhBMP-6 dimers, trimers and tetramers, respectively. As happened in the

production assays, trimers were the most represented form of rhBMP-6 in this sample. No

proteins were recognized by the anti-BMP-6 antibody in the flow-through fraction.

Figure 44. Western blot analysis of the elution fractions obtained by heparin-sepharose chromatography of rhBMP-6 expressed in Sf9 cells. Lane 1, 20 ng of rhBMP-6 produced in CHO cells (R&D Systems). Lane 2, 25 µL of conditioned culture medium supernatant. Lane 3, 25 µL of the flow-through fraction. Lanes 4-18, 5 µL of the elution fractions e12-e26, respectively.

-0.06

-0.04

-0.02

0

0.02

0.04

0.06

0.08

0

10

20

30

40

50

60

70

OD280 Conductivity

e12-e26

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18

31 KDa

45 KDa

66 KDa

97 KDa

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143

The ±39 KDa protein (rhBMP-6 dimer) appeared in fractions e12-e16, with a maximum of

intensity in fractions e13 and e14 (conductivity: 27.6-30.4 mS/cm). The ±56 KDa protein

(rhBMP-6 trimer) appeared in fractions e13-e26, with a maximum of intensity in fractions e14

and e15. The ±73 KDa protein (rhBMP-6 tetramer) appeared in fractions e14-e26, with a

maximum of intensity in fractions e21 and e22. Thus, the elution profiles of the different forms

of rhBMP-6 partially overlapped as proposed by the model shown in figure 45, what implies that

it is impossible to completely purify one of the forms using a standard FPLC system.

Figure 45. Proposed matrix elution model for rhBMP-6 forms expressed in Sf9 cells.

This behaviour could be considered a logic consequence of the fact that rhBMP-6 dimers

only possess two heparin-binding domains, while the trimeric and tetrameric forms possess

three and four of them, respectively. This also supports the idea that the heparin-binding

properties of BMP-6 are not due to structural aspects of the heparin-binding site, but only

depends on the net charge of the amino acid sequence that forms this domain.

The elution fractions e13, e14 and e15 were selected for further analysis, since they

contained the highest amounts of dimeric rhBMP-6. These fractions were reunited in one single

sample (rhBMP6 e13-e15).

4.4.4.2. Purification of rhBMP-6-CBD.

The purification profile for rhBMP-6-CBD could be considered identical to that of rhBMP-6,

showing one clear peak of the OD280 for each conductivity value, being the peaks comparable in

height and width to those of the rhBMP-6 purification profile (Fig. 46).

The elution fractions corresponding to both OD280 peaks were analyzed by Western blot, as

well as the non-purified conditioned medium supernatant and the flow-through fraction

n n+1 n+2 n+3 n+4 n+5 n+6

Elution fraction

rhBMP-6 tetramers

rhBMP-6 trimers

rhBMP-6 dimers

Results________________________________________________________________________

144

(Fig 47). Immunostaining with the specific anti-BMP-6 antibody showed the presence of the

three previously described bands in the conditioned medium supernatant, which are assumed to

correspond to rhBMP-6-CBD dimers, trimers and tetramers, respectively. In this case, rhBMP-6-

CBD dimers were found in more elution fractions (e12-e19), with a maximum of intensity in

fractions e13 to e18 (conductivity: 27.4-30.9 mS/cm). The ±58 KDa protein (rhBMP-6-CBD

trimer) appeared in fractions e13-e37, with a maximum of intensity in fractions e16 to e18. The

±77 KDa protein (rhBMP-6-CBD tetramer) appeared in fractions e35-e37, with a maximum of

intensity in fractions e35 and e36.

Figure 46. Purification by heparin-sepharose chromatography of rhBMP6-CBD expressed in Sf9 cells. The analyzed collection fractions are highlighted in grey boxes.

Figure 47. Western blot analysis of the elution fractions obtained by heparin-sepharose chromatography of rhBMP-6-CBD expressed in Sf9 cells. Lane 1, 20 ng of rhBMP-6 produced in CHO cells (R&D Systems). Lane 2, 25 µL of conditioned culture medium supernatant. Lane 3, 25 µL of the flow-through fraction. Lanes 4-14, 5 µL of the elution fractions e9-e19, respectively. Lanes 15-18, 5 µL of the elution fractions e35-e38, respectively.

-0.08

-0.06

-0.04

-0.02

0

0.02

0.04

0.06

0.08

10

20

30

40

50

60

70

530 535 540 545 550 555 560 565

Vol (mL)

0.08 70 0.06 0.04

0.02 0 -0.02

-0-04 -0.06 -0.08

60

50

40

30

20

10 OD280 Conductivity

e35-e38 e9-e19

31 KDa

45 KDa

66 KDa

97 KDa

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18

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145

1 2 3 4 5 6

31 KDa

45 KDa

66 KDa 97 KDa

The elution fractions e12, e13 and e14 were selected for further analysis, since they

contained the highest amounts of dimeric rhBMP-6 with less contamination of trimers. These

fractions were reunited in one single sample (rhBMP6-CBD e12-e14).

4.4.4.3. Obtaining of rhBMP-6 and rhBMP-6-CBD under native conditions.

Since the purification of rhBMP-6 and rhBMP-6-CBD was performed in presence of 6 M urea,

and the elution of the proteins was induced by increased NaCl concentrations, the excess of

urea and NaCl had to be removed from the selected samples to obtain samples suitable for in

vitro testing of their biological activity. For this purpose, different strategies were tried:

- Dialysis of the samples against 4 mM HCl, against DMEM, pH 7.0, or against DMEM,

pH 4.9. This resulted in a slight increase of volume of the samples.

- Dialysis of the samples against 10 mM ammonium acetate, pH 4.0, lyophilization and

resuspension with water alone, water with 0.1% BSA, 4 mM HCl alone or 4 mM HCl

with 0.1 % BSA.

- Buffer exchange to 4 mM HCl in Vivaspin 2 columns (GE Healthcare).

After removing of the urea and the NaCl, the obtained samples were analyzed by Western

blot (Fig. 48).

Figure 48. Western blot analysis of the rhBMP-6 samples after removing the excess of urea and NaCl. Lane 1, 20 ng of rhBMP-6 produced in CHO cells (R&D Systems). Lane 2, rhBMP-6 e13-e15 before removing the excess of urea and NaCl. Lanes 3 and 4, rhBMP-6 e13-e15 dialyzed against DMEM, pH 7.0 or against DMEM, pH 4.9, respectively. Lane 5, rhBMP-6 e13-e15 dialyzed against 10 mM ammonium acetate, pH 4.0, lyophilized and resuspended with 4 mM HCl + 0.1% BSA. Lane 6, rhBMP-6 e13-e15 recovered from a Vivaspin 2 column after buffer exchange to 4 mM HCl.

Immunostaining of the samples with a specific anti-BMP-6 antibody showed that little

protein was lost in the samples that were dialyzed against DMEM or 4 mM HCl, since the loss of

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intensity corresponded with the increase of the sample volumes. In the case of the lyophilized

samples, the proteins became more concentrated without detecting signs of protein loss due to

precipitation. In the case of the samples loaded on a Vivaspin column, most of the protein was

lost during the process. This could be attributed to protein precipitation within the column, or to

the proteins becoming trapped by the filter membrane.

4.4.5. In vitro analysis of the biological activity of rhBMP-6 and

rhBMP-6-CBD.

The biological activity of the obtained samples was assayed by testing their ability to induce

ALP expression on C2C12 mouse myoblasts cultured in vitro, since BMPs are known to have the

capacity to transdifferentiate these cells into the osteoblastic lineage.

For this purpose, C2C12 cells were seeded on 96-well culture plates with DMEM +

10% FBS. Once attached to the bottom of the plate, the cells were washed with DMEM +

2% FBS and finally incubated with DMEM + 2% FBS containing the different samples of

rhBMP-6 or rhBMP-6-CBD for 72 hours before measuring the ALP activity in the cultures.

The following samples were used for this assay:

- Serial 1/2 dilutions of the rhBMP-6 and rhBMP-6-CBD samples dialyzed against DMEM,

pH 4.9 or 7.0, in DMEM + 2% FBS, from 1/2 to 1/1,024.

- Serial 1/2 dilutions of the rhBMP-6 and rhBMP-6-CBD samples dialyzed against 10 mM

ammonium acetate and lyophilized, in DMEM + 2% FBS, from 1/10 to 1/5,120.

- Serial 1/2 dilutions of the rhBMP-6 and rhBMP-6-CBD samples recovered from the

Vivaspin 2 columns, in DMEM + 2% FBS, from 1/2 to 1/1,024.

- Serial 1/2 dilutions of the rhBMP-6 and rhBMP-6-CBD conditioned medium

supernatants dialyzed against DMEM, in DMEM + 2% FBS, from 1/2 to 1/1,024.

- Serial 1/2 dilutions of the rhBMP-6 and rhBMP-6-CBD conditioned medium

supernatants, in DMEM + 2% FBS, from 1/1 to 1/512.

- Serial 1/2 dilutions of rhBMP-6 produced in CHO cells (R&D Systems), in DMEM + 2%

FBS, from 1 µg/mL to 15.625 ng/mL.

After 72 hours of incubation, ALP activity was only detected in the C+ cultures incubated

with rhBMP-6 produced in CHO cells (Fig. 49). None of the samples containing rhBMP-6 or

rhBMP-6-CBD produced in Sf9 cells was able to induce ALP expression at any tested dilution.

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Figure 49. ALP activity induced by rhBMP-6 produced in CHO cells on C2C12 mouse myoblasts. Data are represented as the mean ± SD. n=3.

4.5. Production of rh-bFGF and rh-bFGF-CBD in Sf9 cells.

Since an rh-bFGF-CBD has already been produced in Escherichia coli and both its biological

activity and collagen-binding ability tested (Andrades JA et al., 2001), we wanted to produce

rh-bFGF and rh-bFGF-CBD in a eukaryotic expression system, which resembles more to human

bFGF-expressing cells than E. coli.

The native bFGF is a monomeric protein with no intracatenary disulfide bonds, though

several cysteine residues, susceptible of forming unspecific disulfide bonds under permissive

conditions, are present in its primary structure. This fact led us to use a baculovirus system that

does not co-express the PDI, since the presence of this chaperone could lead to incorrect

folding of the recombinant proteins by establishing unspecific disulfide bonds.

Although natural bFGF seems to be secreted by an unconventional secretion pathway, by

which the protein is directly translocated through the plasma membrane without entering the

endoplasmic reticulum or the Golgi apparatus, we decided to produce the recombinant bFGF

proteins with a fused signal peptide to direct them through the conventional secretion pathway.

Since Sf9 cells are not natural producers of bFGF, these cells may not possess the elements

involved in the bFGF secretion mechanism, so production of bFGF without a signal peptide in

these cells could lead to accumulation of the recombinant proteins in the form of inclusion

bodies within them.

05

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rhBMP-6 (ng/mL)

rhBMP-6 (R&D Systems)

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4.5.1. Obtaining of rh-bFGF and rh-bFGF-CBD expressing clones of

baculoviruses.

Once the genes encoding the rh-bFGF and rh-bFGF-CBD were correctly cloned into the

pAcGP67B plasmid, Sf9 cells were co-transfected with the recombinant donor plasmid and

linearized BacPak6™ baculoviral DNA in order to obtain infective, heterologous protein-

expressing baculoviruses by homologous recombination. Six or nine days after transfection (for

rh-bFGF and rh-bFGF-CBD, respectively), the cultures showed signs of infection, so the

transfection supernatants (TS) were harvested.

The obtained TS were used to perform a plaque assay, infecting Sf9 cultures with 10-3 to

10-8 dilutions and covering the cells with agarose. Five days p.i., the cultures were stained with

neutral red to identify the lysis plaques, which were found in the cultures infected with 10-5 to

10-7 dilutions of the TS. Four well-isolated lysis plaques for rh-bFGF and six well-isolated lysis

plaques for rh-bFGF-CBD were picked and the viruses they contained were analyzed by PCR

using specific primers against the viral DNA flanking the inserts (Fig. 50).

Figure 50. PCR analysis of the isolated baculoviral clones using primers that hybridize with the baculoviral DNA flanking the insert (P42 vs. P43). A) Analysis of the rh-bFGF-CBD-expressing clones. Lane 1, negative control reaction on the pAcGP67B plasmid. Lanes 2-7, analysis of clones 1-6, respectively. B) Analysis of the rh-bFGF-expressing clones. Lane 1, negative control reaction on the pAcGP67B plasmid. Lanes 2-5, analysis of clones 1-4, respectively.

One clone for each protein was selected and used for amplification to obtain a master stock

(MSV), which was titered by a limit dilution assay. The estimated titer of the MSV of rh-bFGF

was 7.31 x 108 pfu/mL, while the titer of the MSV of rh-bFGF-CBD was 1.62 x 1011 pfu/mL.

Although the viruses isolated by the plaque assay were roughly analyzed by PCR to confirm that

they had the heterologous genes inserted into the viral DNA, a second PCR analysis was

performed on the master stocks. In this case, PCR reactions on both clones were performed

using a specific primer against the 3’-end of the bfgf gene and a primer against the 5’-end of

1 Kbp

700 bp 500 bp

200 bp

1 2 3 4 5 6 7 1 2 3 4 5 A B

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the gene or against the sequence of the CBD (Fig. 51). Furthermore, both clones were

sequenced with the primers against the viral DNA flanking the inserts to ensure that no

mutations or frame-shifting had occurred during the generation of the recombinant

baculoviruses.

Figure 51. PCR analysis of the isolated baculoviral clones. Lanes 1 and 2, Analysis of the rh-bFGF expressing clone with specific primers against the bFGF and the CBD sequence (P13 vs P14, and P15 vs P14), respectively. Lanes 3 and 4, Analysis of the rh-bFGF-CBD expressing clone with specific primers against the CBD and the bFGF sequence (P15 vs P14, and P13 vs P14), respectively.

To obtain a high volume virus stock (HVVS) with which all the future protein productions

can be made, the MS were amplified by infecting cells cultured on 160 mm Petri dishes with a

MOI of 0.1. The obtained HVVS were titered to allow us to perform the protein productions with

a known and previously established MOI. The estimated titer of the HVVS of rh-bFGF was 2.76

x 1010 pfu/mL, while the estimated titer of the HVVS of rh-bFGF-CBD was 9.59 x 109 pfu/mL.

4.5.2. Production assays for rh-bFGF and rh-bFGF-CBD.

To determine the best conditions for large scale protein production, a production assay was

performed with the two HVVS obtained. For this purpose, Sf9 cultures were infected with a MOI

of 2.5 or 10, and the medium of the cultures was harvested 96, 120 and 144 hours p.i. (i.e.,

production times of 72, 96 and 120 hours). 400 µL of the harvested conditioned media were

precipitated with TCA and the samples analyzed by SDS-PAGE and Western blot.

4.5.2.1. Production assay for rh-bFGF.

Immunostaining of the proteins with a specific anti-bFGF antibody revealed one single band

in all the lanes corresponding to the conditioned culture media (Fig. 52). This band had an

1 2 3 4

500 pb

700 pb

200 pb

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estimated molecular mass of ±16 KDa, which is the value described for the commercial bFGF

produced in Escherichia coli.

The highest intensity and integrity of this band was found in the medium of cultures

infected with a MOI of 10 and after a production time of 72 hours. With longer production

times, the intensity of the band diminished and a lower MW smear (possibly due to protein

degradation or incorrectly produced, truncated bFGF accumulation) became more patent.

Infection of the cultures with a MOI of 2.5 yielded less of this ±16 KDa protein and a similar

smear was found under the band with progression of the production time.

In the lane loaded with the commercial rh-bFGF (R&D Systems), the antibody recognized at

least three bands, with an estimated MW of ±16 KDa, ±33 KDa and ±50 KDa. According to

their molecular masses, the two highest bands could correspond to aggregated bFGF molecules

in the form of dimers and trimers, though these aggregations could not be due to unspecific

disulfide bonds since the buffer in which this commercial protein is diluted contains DTT to

avoid cysteine oxidation.

Figure 52. Production assay for rh-bFGF, analyzed by Western blot. Lane 1, 25 ng of commercial rh-bFGF (R&D Systems). Lanes 2-4, rh-bFGF produced with a MOI of 10 after 72, 96 and 120 hours of production time, respectively. Lanes 5-7, rh-bFGF produced with a MOI of 2.5 after 72, 96 and 120 hours of production time, respectively.

According to these results, the best conditions for rh-bFGF production were found to be

infecting with a MOI of 10 and harvesting the conditioned medium after 72 hours of production

time since, under these conditions, the highest yields of the ±16 KDa protein (rh-bFGF) were

found, with apparently less protein degradation.

1 2 3 4 5 6 7

14 KDa

21 KDa

31 KDa

45 KDa

66 KDa

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4.5.2.2. Production assay for rh-bFGF-CBD.

In this case, immunostaining of the proteins with a specific anti-bFGF antibody also

revealed the presence of one single band in the conditioned culture media, with their behaviour

strongly resembling that of the bands found in the production assay for rh-bFGF (Fig. 53).

The estimated MW of this band was ±18 KDa, which could correspond to the predicted

value for rh-bFGF-CBD (± 17.28 KDa). Little differences were found between the cultures

infected with a MOI of 10 or 2.5. In both cases, the intensity and integrity of the band was

higher with shorter production times (72 hours), and diminished with longer production times.

After 120 hours of production time, a lower MW smear appeared under the ±18 KDa band.

Figure 53. Production assay for rh-bFGF-CBD, analyzed by Western blot. Lanes 1-3, rh-bFGF-CBD produced with a MOI of 10 after 72, 96 and 120 hours of production time, respectively. Lanes 4-6, rh-bFGF-CBD produced with a MOI of 2.5 after 72, 96 and 120 hours of production time, respectively. Lane 7, 50 ng of commercial rh-bFGF (R&D Systems).

According to these results, we decided that the best conditions for rh-bFGF production were

infecting indistinctly with a MOI of 10 or 2.5 and harvesting the conditioned medium after 72

hours of production time since, under these conditions, the highest yields of the ±18 KDa

protein (rh-bFGF-CBD) were detected, with apparently less protein degradation.

4.5.3. Expression and purification of rh-bFGF and rh-bFGF-CBD.

Once the best conditions for rh-bFGF and rh-bFGF-CBD had been established, larger scale

productions of both proteins were performed, culturing Sf9 cells at a density of 4 x 107

1 2 3 4 5 6 7

14 KDa

21 KDa

31 KDa

45 KDa

66 KDa

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cells/flask in T-175 cm2 flasks and infecting them with a MOI of 10. 96 hours p.i. (72 hours of

production time), the conditioned media were harvested and directly loaded on heparin-

sepharose chromatography columns for purification. Elution of the proteins was performed

using a linear gradient from 0.15 M to 2 M NaCl (18.9 to 153.0 mS/cm).

4.5.3.1. Purification of rh-bFGF.

280 mL of conditioned medium were used for purification of rh-bFGF. The purification

profile for rh-bFGF showed two overlapping peaks of the OD280 between 22 and 80 mS/cm,

corresponding to at least two different protein populations with different, but relatively low,

affinities to heparin. Two smaller peaks were detected at the end of the profile, which

correspond to proteins that elute between 112 and 150 mS/cm (Fig. 54).

Figure 54. Purification by heparin-sepharose chromatography of rh-bFGF expressed in Sf9 cells. The analyzed collection fractions are highlighted in grey boxes.

Since the analysis of the production assays demonstrated that the rh-bFGF appears as a

single band, 10 µL of the elution fractions corresponding to the four OD280 peaks, the non-

purified conditioned medium supernatant and the flow-through fraction were analyzed by dot

blot instead of by Western blot, because this technique allows a faster analysis of more samples

at once (Fig. 55).

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OD280 Conductivity

e7-e26 e42-e60

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A

B

C

D

E

F

G

1 2 3 4 5 6 7

Figure 55. Immuno-dot blot analysis of the collection fractions of rh-bFGF produced in Sf9 cells and purified by heparin-sepharose chromatography. Spot 1A, 10 ng of commercial rh-bFGF. Spot 2A, culture medium supernatant. Spot 3A, flow-through fraction. Spot 4A, column washing with 20 mM Tris, pH 7.1, 1 mM EDTA, 1 mM DTT, 150 mM NaCl. Spots 5A-3D, collection fractions e7-e26. Spots 1E-5G, collection fractions e42-e60.

Immunostaining of the proteins with a specific anti-bFGF antibody showed the presence of

the rh-bFGF in the conditioned culture medium, while no rh-bFGF was detected in the flow-

through fraction, indicating that all of the recombinant protein was retained within the column

after loading of the sample. The rh-bFGF was also found in all of the analyzed elution fractions,

although a drastic decrease of intensity was detected after fraction e50 (138 mS/cm).

Since the elution fractions between e27 and e41 were not analyzed, it was not possible to

determine if the elution of rh-bFGF was continuous along the entire NaCl gradient, or if the

decrease of the OD280 between these fractions correlated with a decrease of concentration of

recombinant proteins within these fractions. Nevertheless, the strong negative slope of the

purification curve in this area made us assume that the rh-bFGF found between e7 and e26

(23-77 mS/cm), and the one found between e42 and e50 (112-129 mS/cm) correspond to at

least two different bFGF populations with different affinities to heparin. According to this, both

groups of fractions were reunited and further analyzed separately (rh-bFGF e7-e26 and rh-bFGF

e42-e50; or rh-bFGF “low heparin-affinity” and “high heparin-affinity” samples, respectively).

4.5.3.2. Purification of rh-bFGF-CBD.

210 mL of conditioned medium were used for purification of rh-bFGF-CBD. The purification

profile for rh-bFGF-CBD strongly resembled the one observed for rh-bFGF. Again, two

overlapping peaks of the OD280 were found when eluting with relatively low conductivity values

(26-79 mS/cm) and a group of small overlapping peaks were detected at the end of the

gradient, between 116 and 150 mS/cm. In this case, another relevant peak of the OD280 was

found between 90 and 101 mS/cm (Fig. 56).

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100

120

140

160

180

OD280 Conductivity

e7-e15 e16-e31 e32-e39 e45-e71e40-e44

Figure 56. Purification by heparin-sepharose chromatography of rh-bFGF-CBD expressed in Sf9 cells. The analyzed collection fractions are highlighted in grey boxes.

10 µL of all of the elution fractions between e7 and e71 (26-150 mS/cm) were analyzed by

dot blot to determine which of the observed peaks of the OD280 correspond to rh-bFGF-CBD

elution. The non-purified conditioned culture medium supernatant and the flow-through fraction

were also analyzed (Fig. 57).

Figure 57. Immuno-dot blot analysis of the collection fractions of rh-bFGF-CBD produced in Sf9 cells and purified by heparin-sepharose chromatography. Spot 1A, culture medium supernatant. Spot 2A, flow-through fraction. Spots 3A-3I, collection fractions e7-e71. Spots 4I and 5I, 10 ng of commercial rh-bFGF.

1 2 3 4 5 6 7 8

A B C D E F G

I

H

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Immunostaining of the proteins with the specific anti-bFGF antibody showed the presence

of the rh-bFGF-CBD in the conditioned culture medium, while no rh-bFGF was detected in the

flow-through fraction, indicating that all of the recombinant protein was retained within the

column. The rh-bFGF-CBD was found in all of the analyzed elution fractions, until e46 (121

mS/cm), although the intensity of the spots showed clear variations along the elution gradient.

Most of the rh-bFGF-CBD was found between e16 and e31 (49-87 mS/cm), corresponding with

the highest observed peak of the OD280. Nevertheless, a second, unexpected, intensity peak

was found between e41 and e 43 (106-113 mS/cm), not corresponding to any clear peak of the

OD280.

Since the elution gradient was linear, it could be concluded that both intensity peaks

correspond to different rh-bFGF-CBD populations with distinct affinities to heparin. According to

this, both groups of fractions were reunited and further analyzed separately (rh-bFGF-CBD

e16-e31 and rh-bFGF-CBD e40-e44; or rh-bFGF-CBD “low heparin-affinity” and “high heparin-

affinity” samples, respectively).

4.5.3.3. Obtaining of rh-bFGF and rh-bFGF-CBD under native conditions.

In summary, the samples obtained after purification of the recombinant proteins were:

Sample Volume (mL) Conductivity range (mS/cm) Average NaCl concentration (M)

rh-bFGF e7-e26 20 23-77 0.594

rh-bFGF e42-e50 9 112-129 1.530

rh-bFGF-CBD e16-e31 16 49-87 0.800

rh-bFGF-CBD e40-e44 5 104-116 1.395

Table 9. Samples of bFGF after purification by heparin-sepharose chromatography.

Although the purification of rh-bFGF and rh-bFGF-CBD was preformed in the absence of

urea or other denaturing or chaotropic agents, the high NaCl molarity of the samples did not

allow their use on cells or living tissues. To remove the excess of salt, 4 mL of each sample

were loaded on a Vivaspin 2 column with a 5 KDa MW cut-off and the elution buffer was

exchanged to PBS, pH 7.3, 1 mM DTT, 1 mM EDTA by repeated centrifugation and dilution

steps. At the end of the process, the NaCl concentration in the samples was reduced to values

below 20 mM and the final volumes of the samples were approximately 250 µL (Table 10).

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Sample Volume (µL) Estimated NaCl concentration (mM)

rh-bFGF e7-e26 ±220 ±11

rh-bFGF e42-e50 ±250 ±18

rh-bFGF-CBD e16-e31 ±240 ±14

rh-bFGF-CBD e40-e44 ±250 ±15

Table 10. Samples of bFGF after buffer exchange and concentration.

To determine if the recombinant proteins were lost or could be recovered after the buffer

exchange, the obtained samples before and after the buffer exchange procedure were analyzed

by dot blot for comparison (Fig. 58).

Figure 58. Analysis by immuno dot-blot of the rh-bFGF and the rh-bFGF-CBD produced in Sf9 cells after purification and buffer exchange. Row 1, 1 µL of rh-bFGF e7-e26, rh-bFGF e42-e50, rh-bFGF-CBD e16-e31 and rh-bFGF-CBD e40-e44, respectively, before buffer exchange and concentration. Row 2, 1 µL of rh-bFGF e7-e26, rh-bFGF e42-e50, rh-bFGF-CBD e16-e31 and rh-bFGF-CBD e40-e44, respectively, after buffer exchange and concentration.

Immunostaining of the proteins with the specific anti-bFGF antibody revealed that both the

rh-bFGF e7-e26 and the rh-bFGF-CBD e16-e31 samples had suffered a significant loss of

proteins during buffer exchange and/or concentration. Although the spots corresponding to

these samples increased their intensity after this procedure, this increase was not high enough

to correlate with the 16-fold decrease of the sample volume achieved. In contrast, the proteins

with high affinity to heparin (rh-bFGF e42-e50 and rh-bFGF-CBD e40-e44) greatly increased

their concentration after the buffer exchange procedure, so it was assumed that no significant

protein loss had occurred in these samples.

An active bFGF molecule has to possess a high affinity to heparin, since binding of bFGF to

glycosaminoglycans is necessary for the formation of the bFGF-FGFR complex. The facts that

the samples rh-bFGF e7-e26 and rh-bFGF-CBD e16-e31 seemed to contain mostly bFGF forms

with a diminished affinity to heparin, and that most of the proteins of these samples were lost

by precipitation or by becoming trapped in the column membrane during buffer exchange,

made us decide to reject them. From now on, we will refer to the samples rh-bFGF e42-e50 and

rh-bFGF-CBD e40-e44 as rh-bFGF and rh-bFGF-CBD, respectively.

1 2

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To determine the collagen-binding affinity and the biological activity of the proteins in the

samples, the concentration of the recombinant growth factors had to be estimated. For this

purpose, serial dilutions of both samples and of a commercial standard rh-bFGF solution of

known concentration were analyzed by Western blot. Densitometric analysis of the bands were

performed using the program ImageJ (Rasband, WS., ImageJ, U.S. National Institute of Health,

Bethesda, Maryland, USA). The estimated recombinant protein concentrations in the samples

were:

rh-bFGF: 16.67 µg/mL, i.e., 926.11 nM

rh-bFGF-CBD: 9.27 µg/mL, i.e., 481.81 nM

According to these values, the final yield of the production of these proteins in Sf9 cells

with the used expression system could be calculated:

rh-bFGF: ±33.50 µg/L, i.e., 1.86 nmol/L

rh-bFGF-CBD: ±13.80 µg/L, i.e., 0.72 nmol/L

4.5.4. Collagen-binding affinity tests for rh-bFGF and rh-bFGF-CBD.

To determine if the addition of the CBD in the engineered rh-bFGF-CBD conferred an

increased affinity to collagen to the molecule, a collagen-binding affinity test was performed

following a method described by T. Kitajima (Kitajima T et al., 2007). Absorbable collagen

type I sponge discs (1 mm thick, 5 mm diameter) were incubated with 10 µL of a solution

containing 1.25 pmol of either commercial rh-bFGF (R&D Systems), rh-bFGF or rh-bFGF-CBD for

2 hours at 37 ºC. Afterwards, the collagen sponges were washed for 1 hour with PBST and

immunostained with a specific anti-bFGF antibody. Finally, the amount of proteins bound to the

collagen sponges was quantified by digital image analysis using the program ImageJ (Rasband,

WS., ImageJ, U.S. National Institute of Health, Bethesda, Maryland, USA).

Immunostaining of the sponges revealed that almost all of the commercial rh-bFGF,

produced in Escherichia coli, was washed from the ACS (Fig. 59). Surprisingly, a slight staining

of the ACS incubated with rh-bFGF was observed, indicating that the rh-bFGF produced in Sf9

cells with the used expression system had an increased affinity to collagen when compared to

the growth factor produced in E. coli. These differences were found to be statistically significant

(p<0.05).

As hypothesized, the sponges incubated with rh-bFGF-CBD showed an increased staining

when compared to both the commercial rh-bFGF and the rh-bFGF produced in Sf9 cells, what

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indicated that the addition of the CBD conferred a specific affinity to collagen type I to the

molecule. This increase of the collagen-binding properties of rh-bFGF-CBD was statistically

significant when compared to those of rh-bFGF, reaching a value close to 50%.

Figure 59. Collagen-binding test of rh-bFGF and rh-bFGF-CBD produced in Sf9 cells. A) Immunostaining with an anti-bFGF antibody of absorbable collagen sponges incubated with 1.25 pmol of rh-bFGF or rh-bFGF-CBD and washed for 1 hour with PBST. Black circles indicate the location of the sponges incubated with commercial rh-bFGF; the red circle indicates the location of the negative control sponge. B) Graphic showing the densitometric analysis of the spots. Data are represented as the blanked means ± SD, using a relative scale (0-255). n = 5; *p < 0.05.

Since the aim of the production of rh-bFGF-CBD was to obtain an active form of bFGF with

the ability of being retained for a longer period of time at the wound site when implanted in

vivo for bone repair purposes, with a consequent enhanced effect on tissue healing and safety

of the clinical approach, we wanted to determine if the binding of rh-bFGF-CBD to the

absorbable collagen sponges was stable in time. For this purpose, a new set of ACS discs were

incubated with 1.25 pM of either rh-bFGF or rh-bFGF-CBD for 2 hours at 37 ºC. Afterwards, the

sponges were washed for 6 days with PBST at 4 ºC before immunostaining of the proteins.

The immunostaining of the collagen sponges revealed that both the rh-bFGF and the

rh-bFGF-CBD remained bound after 6 days of extensive washing (Fig. 60). As observed when

the sponges were washed during only one hour, the collagen-targeted rh-bFGF-CBD showed an

increased affinity to collagen when compared to native rh-bFGF, being the difference

statistically significant (p<0.05).

0

5

10

15

20

25

30

35

rh-bFGF (R&D Systems)

rh-bFGF

rh-bFGF-CBD

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rh-bFGF


C-

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*rh-bFGF (R&D Systems)

rh-bFGF

rh-bFGF-CBD

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Figure 60. Stability of the collagen-binding of rh-bFGF and rh-bFGF-CBD produced in Sf9 cells. A) Immunostaining with an anti-bFGF antibody of absorbable collagen sponges incubated with 1.25 pmol of rh-bFGF or rh-bFGF-CBD and washed for 6 days with PBST. The red circle indicates the location of the negative control sponge. B) Graphic showing the densitometric analysis of the spots. Data are represented as the blanked means ± SD, using a relative scale (0-255). n = 5; *p < 0.05.

According to these results, it could be concluded that the rh-bFGF and rh-bFGF-CBD

produced in Sf9 cells showed an enhanced affinity to collagen when compared to commercial

rh-bFGF produced in E. coli, and that binding of the growth factors to collagen type I sponges

was stable during long periods of time. Furthermore, the CBD of the rh-bFGF-CBD conferred to

the molecule an additional affinity to collagen type I.

0

10

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50

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rh-bFGF-CBD

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rh-bFGF-CBD rh-bFGF

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4.5.5. In vitro analysis of the biological activity of rh-bFGF and

rh-bFGF-CBD.

4.5.5.1. Mitogenic activity of rh-bFGF and rh-bFGF-CBD on MC3T3-E1 mouse

preosteoblasts.

Since bFGF is especially known for being a potent mitogen for cells of mesodermic and

neuroectodermic origins, the biological activity of the produced recombinant growth factors was

tested in vitro by a proliferation assay on mouse preosteoblastic cells.

MC3T3-E1 mouse preosteoblast were seeded on 96-well culture plates at a density of

10,000 cells/well and incubated with 1.25 nM, 625, 312.50 or 156.25 pM of either rh-bFGF

produced in E. coli (R&D Systems), rh-bFGF or rh-bFGF-CBD. 72 hours later, the number of

cells/well was measured using an MTT-based colorimetric method.

After 72 hours of incubation with the highest concentration of any of the recombinant

growth factors, the cells showed evident morphological changes when compared to the control

cultures without bFGF (Fig. 61). The first had experienced a remarkable decrease of size,

showing a small, slightly fibroblastic or round-shaped cell body and long, thin cytoplasmic

prolongations which connected them with their neighbour cells or to the plate. These cells also

had a small, condensed nucleus and a well-defined cell border. In contrast, the cells of the

control cultures were much bigger and had a polygonal shape, with smaller cytoplasmic

prolongations if any. They showed a big nucleus and the borders of the cells were difficult to

identify.

These morphological changes were found to be dose-dependent, since the cells incubated

with the lowest concentrations of the recombinant growth factors showed an intermediate

phenotype between that of the cells incubated with 1.25 mM of bFGF and that of the cells in

the control cultures.

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161

Figure 61. Phenotypical changes induced by rh-bFGF and rh-bFGF-CBD on MC3T3-E1 mouse preosteoblasts. Photomicrographs were taken after 72 hours of incubation with the growth factors and 2 hours of incubation with MTT labelling reagent. A, B and C) Cells incubated with 1.25 nM rh-bFGF (R&D Systems), rh-bFGF and rh-bFGF-CBD, respectively. D, E and F) Cells incubated with 156.25 pM of rh-bFGF (R&D Systems), rh-bFGF and rh-bFGF-CBD, respectively. G) Negative control culture of cells incubated without bFGF. Scale bars = 100 µm.

The conversion of the obtained OD570 values into number of cells per well showed that,

after 72 hours of incubation, the cells in the C- cultures had grown to a density of 30,121 ±

3,573 cells/well (Fig. 62). After the same period of incubation with any form of bFGF, the

cultures all reached higher densities (ranging from 39,790 ± 2390 to 54,960 ± 7861 cells/well),

being these differences statistically significant (p<0.05). Furthermore, no significant differences

were found between the cell proliferation achieved by the commercial rh-bFGF or by the

rh-bFGF or rh-bFGF-CBD produced in Sf9 cells when used at low concentrations (156.25 or

312.50 pM). Only in the cultures incubated with higher concentrations of growth factors

(625.00 pM or 1.25 nM), the commercial rh-bFGF was able to induce a significant higher level of

proliferation when compared to the growth factors produced in Sf9 cells (p<0.05).

A B C

D E F

G

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162

Figure 62. Proliferation of MC3T3-E1 mouse preosteoblast induced by bFGF. At any concentration tested, the proliferation achieved in the cultures treated with any bFGF was significantly higher than the one observed in the control cultures. Only at the highest concentrations (1.25 nM and 625.00 pM), the commercial rh-bFGF was significantly more active than the rh-bFGFs produced in Sf9 cells. Data are represented as the means ± SD. n = 7; *p < 0.05.

Graphic representation of the data as activity curves showed more clearly how the initial

slopes of the curves of the three growth factors were identical (Fig. 63). Furthermore, no

differences between the curves corresponding to both growth factors produced in Sf9 cells

could be observed at any of the tested concentrations. Only at the highest concentrations

assayed, the commercial rh-bFGF showed slightly significant more mitogenic activity than the

recombinant growth factors produced with the baculoviral expression system.

Figure 63. Mitogenic activity curves of rh-bFGF and rh-bFGF-CBD. Data are represented as the means ± SD. n = 7; *p < 0.05.

25000

30000

35000

40000

45000

50000

55000

60000

65000

bFGF concentration

Num

ber

of c

e


rh-bFGF

rh-bFGF-CBD

0 156.25 625.00 1250 pM 312.50

*

*


rh-bFGF

rh-bFGF-CBD

C-

1.25 nM 625.00 pM 312.50 pM 156.25 pM bFGF concentration

0

10000

20000

30000

40000

50000

60000

70000Nu

mbe

r of c

ells

* *

* *

*

*

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163

4.5.5.2. Inhibition of differentiation of MC3T3-E1 mouse preosteoblasts by rh-bFGF

and rh-bFGF-CBD.

To determine if the produced recombinant growth factors had the ability to inhibit the

osteoblasic differentiation of MC3T3-E1 preostoblastic cells induced by ascorbic acid, cells were

seeded on 96-well culture plates at a density of 10,000 cells/well in medium containing 0.2 mM

L-ascorbic acid and incubated with 1.25 nM, 625, 312.50 or 156.25 pM of either rh-bFGF

produced in E. coli (R&D Systems), rh-bFGF or rh-bFGF-CBD. Negative control cultures were

incubated under the same conditions without bFGF. 120 hours later, the number of cells/well

and the ALP activity in the cultures were measured using an MTT-based and a p-NPP-based

colorimetric method, respectively.

After 120 hours of incubation, no ALP activity could be detected in any of the cultures

treated with bFGF. Only in the negative control cultures, very low levels (0.338 ± 0.109 U/L) of

ALP activity were found. Nevertheless, when calculating the ALP activity per cell, these

differences were found to be statistically significant (Fig. 64).

Fig. 64. ALP activity in cultures of MC3T3-E1 mouse preosteoblasts in the presence of ascorbic acid and bFGF. Data are represented as the means ± SD. n = 7; **p < 0.01.

According to these results, it could be concluded that the recombinant growth factors

produced in Sf9 cells were able to inhibit the osteoblastic differentiation of preostoblasts in the

presence of ascorbic acid.

1.25 nM 625.00 pM 312.50 pM 156.25 pM -5E-10

0

5E-10

0.000000001

1.5E-09

0.000000002

2.5E-09

0.000000003

bFGF concentration

ALP

act

ivit

y (U

/cel

l)

rh-bFGF (R&D Systems)rh-bFGFrh-bFGF-CBDC-

** ** ** **3 x 10-9

2.5 x 10-9

2 x 10-9

1.5 x 10-9

1 x 10-9

5 x 10-10

-5 x 10-10

0

Results________________________________________________________________________

164

4.6. In vivo heterotopic bone formation.

To evaluate the biological activity of the rh-bFGF and rh-bFGF-CBD produced in Sf9 insect

cells and their capacity to enhance bone formation in combination with BMP-6, ACS discs

carrying 13.89 pmol rhBMP-6 alone, 1.25 pmol of rh-bFGF or rh-bFGF-CBD alone, or

combinations of these factors, were implanted into the dorsal muscles of rats. Commercial

rh-bFGF produced in E. coli was used as a positive control, while ACS discs loaded with vehicle

only served as a negative control.

21 days after surgery, all of the implants loaded with growth factors could be recovered

and no macroscopic signs of immune rejection or fibrotic encapsulation were observed. In

contrast, none of the negative control ACS, loaded with vehicle only, could be recovered due to

reabsorption.

All the implants that were loaded with rhBMP-6 (alone or in combination with bFGF) showed

a well-defined border and hard consistency, though varied in size. In opposition, most of the

implants that were loaded with commercial rh-bFGF, rh-bFGF produced in Sf9 cells or

rh-bFGF-CBD alone, were difficult to localize in the muscle and mainly appeared as more loose,

sometimes disperse, particles.

The histological analysis of the implants loaded with commercial rh-bFGF, rh-bFGF

produced in Sf9 cells or rh-bFGF-CBD alone showed that these all had given raise to a more or

less dense, fibrotic tissue in which no signs of ossification and almost no signs of angiogenesis

could be detected (Fig. 65). Spread throughout the implants, dense accumulations of an

apparently disorganized, unidentified extracellular matrix were observed (asterisks in Fig. 65),

being these accumulations especially frequent and compact in the implants loaded with rh-bFGF

and rh-bFGF-CBD produced in insect cells. Staining of these implants with Masson’s trichrome

revealed the presence of collagen molecules within the extracellular matrix accumulations,

especially in those found in the implants loaded with commercial rh-bFGF.

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Figure 65. Staining of the implants without BMP-6 with H-E (A, B and C) and Masson’s trichrome (D, E and F). A, D) Implant with commercial rh-bFGF. B, E) Implant with rh-bFGF produced in Sf9 cells. C, F) Implant with rh-bFGF-CBD. asterisks: extracellular matrix accumulations; m: muscle. Scale bars = 100 µm.

A

B

D

E

C Fm

m

m

m

*

* *

*

*

*

*

*

* *

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166

All the implants loaded with rhBMP-6 (alone or in combination with bFGF) showed clear

signs of osteogenesis, leading to the formation of a bony tissue with a trabecular appearance

(Fig. 66). Staining of the implants with hematoxylin-eosin revealed the existence of trabeculae,

especially at the periphery and, to a greater or lesser extent, also at the inner part of the

samples, with numerous osteocytes embedded inside of them. By staining of the samples with

Masson’s trichrome these trabeculae appeared green, indicating that they were mainly

composed of collagen.

The external surface of the implants was, in most cases, covered by a dense, fibrous cluster

of mesenchymal cells, organized as more or less concentric layers around the external

trabeculae. In contrast, the inner part of the implants was mainly filled with a bone marrow-like

tissue, constituted of large quantities of undifferentiated mesenchymal cells. In between these

cells it was possible to observe blood lacunae, which had an irregular shape and poorly defined

borders, corresponding to blood vessels in a very immature state. Other implants showed more

mature blood vessels with clearly differentiated endothelial cells and, in some cases, cells

forming a partial second layer around the endothelium, resembling perivascular cells. These

blood vessels varied in diameter but exceeded 200 µm in some implants. Adipocytes were

found spread throughout the inner part of the implants, more or less isolated or forming small

groups.

In some implants, it was possible to find zones of hypertrophic cartilage, indicating that

bone formation occurred, at least at some areas, by the endochondral ossification mechanism.

Staining of the implants with alcian blue, which binds to the acid and sulphated residues of the

glycosaminoglycans present in the cartilaginous matrix, revealed the presence of more

immature patches within the bony trabeculae, corresponding to areas where the cartilaginous

matrix was not yet completely substituted by osteoid and mineralized (Fig. 67). As expected,

alcian blue staining was especially intense in the areas where the hypertrophic chondrocytes

were localized.

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167

Figure 66. Staining of the implants with BMP-6 with H-E (A, B, C and D) and Masson’s trichrome (E, F, G and H). A, E) Implant with rhBMP-6 alone. B, F) Implant with rhBMP-6 + commercial rh-bFGF. C, G) Implant with rhBMP-6 + rh-bFGF produced in Sf9 cells. D, H) Implant with rhBMP-6 + rh-bFGF-CBD. bl: blood lacunae. bm: bone marrow-like tissue; f: fibrous layers. m: muscle; t: trabeculae; v: blood vessels. Scale bars = 100 µm.

A E

B F

C G

m m

m m

mm

m

t

t

t

t

t

t

t

t

t

t

t

t

t t

f

f

f

f f

f

f

f

bl

bm

bm

bmv

v v

v v

v bm

bl bl

bl

bl bl

bl

bm bm

D H

m

bm

t t

t

t m

bm

Results________________________________________________________________________

168

Figure 67. Staining of the implants with alcian blue. A) Implant with rhBMP-6 alone. B) Implant with rhBMP-6 + commercial rh-bFGF. C) Implant with rhBMP-6 + rh-bFGF produced in Sf9 cells. D) Implant with rhBMP-6 + rh-bFGF-CBD. bl: blood lacunae. bm: bone marrow-like tissue; m: muscle; t: trabeculae; v: blood vessels. Scale bars = 100 µm.

Immunolocalization of osteopontin with a specific anti-osteopontin antibody revealed the

expression of this early osteogenic marker in all the implants loaded with BMP-6, alone or in

combination with bFGF (Fig. 68). Osteopontin was found in the bone trabeculae, where it was

heterogeneously localized but mainly concentrated at their external surfaces, defining the areas

of active synthesis of new osteoid. Some of the osteocytes embedded in the trabecular osteoid

also showed certain immunoreactivity with the antibody used for staining.

Among the bone marrow, osteopontin could be found as more or less small accumulations

in the extracellular matrix. It was also possible to observe numerous osteoblasts expressing this

protein between the undifferentiated mesenchymal cells.

A

m t

bm

B

vv

v

t

t bm

D

m

t

v

m

t

m

bm

C

v

v

v v

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bm

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169

Figure 68. Immunostaining of the implants with an anti-osteopontin antibody. A) Implant with rhBMP-6 alone. B) Implant with rhBMP-6 + commercial rh-bFGF. C) Implant with rhBMP-6 + rh-bFGF produced in Sf9 cells. D) Implant with rhBMP-6 + rh-bFGF-CBD. bm: bone marrow-like tissue; m: muscle; t: trabeculae; v: blood vessels; arrowheads: osteopontin accumulations; Scale bars = 100 µm.

4.6.1. Analysis of the implants with rhBMP-6 alone.

The implants that were loaded with rhBMP-6 alone were, in general, smaller in size than the

rest of the rhBMP-6-containing implants (Fig. 69). They showed bone trabeculae containing

osteocytes mainly at the outer surface but almost not at the inner part of the implant. These

trabeculae showed little staining with alcian blue, indicating that almost no cartilaginous matrix

was left within them. The inner part of the implants was constituted of a very dense

accumulation of undifferentiated mesenchymal cells in between which only few, disperse

adipocytes could be observed. Immunostaining with a specific anti-osteopontin anitbody

revealed that this protein was accumulated mainly within the osteoid of the trabeculae and that

only small groups of osteoblast were expressing this osteogenic marker in the rest of the

implant, indicating that there was little osteogenic activity in the sample. Only some immature

blood lacunae and small blood vessels could be observed throughout these implants.

A m

t

bm

C

v

v v

t

bm

B

v

v

bm t

D

m

t

bm

Results________________________________________________________________________

170

Figure 69. Histological analysis of the implants with rhBMP-6 alone. A) H-E staining. B, C and D) Immunolocalization of osteopontin. D shows a magnification of the area marked in C. bm: bone marrow-like tissue; f: fibrous layers; t: trabeculae; arrowheads: osteoblasts expressing osteopontin. Scale bars = 100 µm in A, B and C; 20 µm in D.

4.6.2. Analysis of the implants with rhBMP-6 and commercial rh-bFGF.

The implants that were loaded with rhBMP-6 and commercial, E. coli-derived rh-bFGF

showed a clear border formed by bone trabeculae containing numerous osteocytes, as observed

in the implants that were loaded with rhBMP-6 alone (Fig. 70). Nevertheless, the inner part of

the implants was also occupied by a broad network of trabeculae. In between the trabeculae, a

bone marrow-like tissue was observed, containing undifferentiated mesenchymal cells but also

many adipocytes. Osteopontin was localized not only in the osteoid of the trabeculae, but some

of the implants also showed areas of osteopontin-containing material with a fibrous,

disorganized appearance (arrowheads in Fig. 70 B). Many blood lacunae and some relatively big

blood vessels could be found along the implant, with some of these blood vessels showing a

well-defined endothelium. In some cases, additional layers of cells could be observed associated

with the endothelium, resembling perivascular organizations (asterisk in Fig. 70 E) and

indicating a greater maturity of these vessels.

BA

C

t

bm

t

bm

t

bm

D

f

f

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171

Figure 70. Histological analysis of the implants with rhBMP-6 + commercial rh-bFGF. A, E and F) H-E staining. B, C and D) Immunolocalization of osteopontin. F shows a magnification of the area marked in E. a: adipocytes; bl: blood lacunae; bm: bone marrow-like tissue; t: trabeculae; v: blood vessels; arrowheads: osteopontin accumulations; arrows: osteocytes; double arrows: endothelial cells; asterisks: perivascular organizations. Scale bars = 100 µm in A, B, C and D; 50 µm in E; 20 µm in F.

4.6.3. Analysis of the implants with rhBMP-6 and rh-bFGF produced in Sf9 cells.

The implants that were loaded with rhBMP-6 and rh-bFGF produced in insect cells were

very similar to the ones that were loaded with rhBMP-6 and commercial rh-bFGF, showing bone

trabeculae with numerous osteocytes at the surface and at the inner part, and a bone marrow-

like tissue with areas of adipose tissue and disperse adipocytes in between the undifferentiated

mesenchymal cells (Fig. 71). Nevertheless, these implants contained small osteopontin

A B

C

a

v v

v

t

t

bm

v

a

a

D

bl

t

t

E

bm

t

v

*

F

*

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Results________________________________________________________________________

172

accumulations within the bone marrow, but no disorganized, fibrous, osteopontin-containing

material could be observed. Furthermore, all of the implants contained numerous blood lacunae

and very big blood vessels. The latter reached 200 µm of diameter in some cases and showed a

well defined endothelial border. As happened in the implants loaded with rhBMP-6 and

commercial rh-bFGF, groups of perivascular cells could be found associated to some of the

blood vessels (asterisk in Fig. 71 E). These implants showed the most mature appearance of all

the analyzed samples.

Figure 71. Histological analysis of the implants with rhBMP-6 + rh-bFGF produced in Sf9 cells. A and F) H-E staining. B, C, D and E) Immunolocalization of osteopontin. a: adipocytes; bm: bone marrow-like tissue; t: trabeculae; v: blood vessels; arrowheads: osteopontin accumulations; arrows: osteocytes; double arrows: endothelial cells; asterisks: perivascular organizations. Scale bars = 100 µm in A, B and C; 50 µm in D and F; 20 µm in E.

A B

C D

v v

a

bmt

a

v

v

tbm

t

bm

bm

F

bm

t

v

*

E

t

________________________________________________________________________Results

173

4.6.4. Analysis of the implants with rhBMP-6 and rh-bFGF-CBD.

The implants that were loaded with rhBMP-6 and rh-bFGF-CBD were comparable in size to

those loaded with rhBMP-6 and commercial or Sf9-derived rh-bFGF. Nevertheless, the general

aspect of these implants was clearly different than the latter, having the appearance of bony

tissue in an earlier stage of maturation (Fig. 72). Bone trabeculae containing osteocytes were

heterogeneously distributed along the implants, defining areas of more mature and areas of

less mature bone. In these less mature areas it was possible to observe many hypertrophic

chondrocytes embedded in a strong alcian blue-positive matrix (Figs. 67 D and 72 C and D).

Although some hypertrophic chondrocytes could be found in the other implants, these were

especially abundant in the ones loaded with rhBMP-6 and rh-bFGF-CBD. In between these

patches of hypertrophic cartilage, abundant fibrous-like accumulations of osteopontin-

containing material could be observed (arrowheads in Fig. 72 C), similar to those found in some

of the implants loaded with rhBMP-6 and commercial rh-bFGF.

The bone marrow-like tissue of these implants contained only few adipocytes and almost no

signs of angiogenesis.

Figure 72. Histological analysis of the implants with rhBMP-6 + rh-bFGF-CBD produced in Sf9 cells. A) H-E staining. B and C) Immunolocalization of osteopontin. D) Alcian blue staining. a: adipocytes; bm: bone marrow-like tissue; hc: hypertrophic chondrocytes; m: muscle; t: trabeculae; v: blood vessels; arrowheads: osteopontin accumulations; arrows: osteocytes. Scale bars = 100 µm.

C

A

D

B

hc

hc

hc

hc

m

t

bm

v

t

t

174

5. Discussion.

175

176

Discussion

5.1. Engineering of the growth factors.

5.1.1. Engineering of the gene encoding the rhBMP-6-CBD.

Since U-2 OS osteosarcoma cells are known to express high levels of BMP-6, they were

used as the primary source of the bmp-6 mRNA. By RT-PCR, the sequence that encodes the

mature domain of the hBMP-6 was retro-transcribed to cDNA, amplified and cloned into the

routine maintenance vector pBIISK. We decided to use only the sequence of the mature

domain, removing the sequences for the preceding pre-peptide and pro-domain, to simplify the

expression in the host cells and to allow us to add the sequence of the CBD derived from the

vWF to the N-terminus of the protein. The pre-peptide and pro-domain are excised from the

BMP during its processing inside the cell and are not known to be necessary for the protein to

acquire its native, folded and active structure (Hillger F et al., 2005). Furthermore, the pre-

peptide seems to mediate translocation of the pre-pro-protein into the lumen of the

endoplasmic reticulum, what becomes unnecessary when the heterologous proteins are

expressed in a prokaryotic system. In the case of the eukaryotic expression systems used, the

shuttle vector in which the gene is cloned provides the GP64 signal peptide sequence to direct

the produced heterologous proteins to the lumen of the endoplasmic reticulum. This signal

peptide, located at the N-terminus of the protein, is eliminated by the endopeptidases of the

host cell before secretion of the protein to the culture medium.

On the other hand, the function of the pro-domain is still unclear, although it has been

suggested that it could delay the maturation of the growth factor until its excision, controlling

the BMP activity in vivo (as happens with the latency-associated peptide of TGF-β) or mediate

oxidative structure formation of the BMPs.

These facts, together with the demonstration that both BMP-2 and pro-BMP2 can be

refolded in vitro from the denatured state (Hillger F et al., 2005) made us decide to use only

the sequence of the mature domain.

For the production of the rhBMP-6-CBD, the sequence encoding the decapeptidic collagen

binding domain derived from the von Willebrand factor was added to the bmp-6 sequence to

obtain this CBD fused to the N-terminus of the molecule. This location was chosen since the

cysteine-rich region involved in the constitution of the cysteine knot and the regions involved in

binding to the BMP receptors are mainly located at the C-terminus (McDonald and Hendrickson,

1993). Thus, addition of the CBD to the C-terminus might cause incorrect folding of these

structures or, if not, partially hide the correctly folded BMPR-recognition sites.

In opposition, the N-terminus of the BMPs contains 6 arginine residues and possesses a

strongly positive net charge. This region seems to be responsible for the affinity to heparin

177

Discussion_____________________________________________________________________

shown by the BMPs, which may help to modulate their biological effects in vivo, but is not

directly involved in signal transduction (Ruppert R et al., 1996).

The original sequence of the CBD contains one cysteine residue (Takagi J et al., 1992),

which could react with any of the other 7 cysteines present in the mature domain to form an

unspecific disulfide bond, hindering the correct establishment of the cysteine knot and, thus,

giving rise to an inactive, aberrant BMP. In our construction, the cysteine of the CBD was

replaced by a methionine (Tuan TL et al., 1996) in order to limit the probability of the CBD to

interfere with the correct folding of the protein.

Other recombinant growth factors produced with this same CBD by other groups also

contained a hexa-histidine purification tag and a thrombin recognition site to excise this tag

after recovery and purification of the protein (Tuan TL et al., 1996; Han B et al, 1997; Andrades

JA et al, 2001). Since the presence of six histidine residues (which have a positive net charge)

could interfere with the folding of the protein, and our aim was to obtain the growth factors

with the fewer modifications and to perform as little in vitro manipulations as possible, we only

added the decapeptidic CBD and a tripeptidic (Gly-Ala-Ser) linker sequence to the molecule. The

presence of the glycine in the linker sequence should allow free rotation of the CBD and

minimize possible conformational impediments between the CBD and the N-terminal domain of

the growth factor.

5.1.2. Obtaining of the genes encoding the rh-bFGF and the

rh-bFGF-CBD.

The genes encoding the rh-bFGF and rh-bFGF-CBD were directly obtained from the two

constructions (pET28b:hbFGF-F1 and pET28b:hbFGF-F2) that were used for their expression in

Escherichia coli as described in Andrades JA et al., 2001. Although these constructions

contained the above mentioned sequences for a His6 purification tag and a thrombin recognition

site, these elements were not amplified together with the growth factors during the PCRs

performed to transfer them to the pAcGP67B shuttle vector since we wanted to avoid the extra

manipulation steps necessary to remove them after the purification of the molecules.

178

Discussion

5.2. Production of rhBMP-6 in Escherichia coli.

The first attempt on rhBMP-6 production was performed in E. coli, since this expression

system was demonstrated to be suitable for the production of active rhBMP-2 and rhBMP-2-CBD

with acceptable yields (Visser R et al., 2009). As in this case, the Rosetta™ (DE3) strain was

used for expression of the growth factor, since this strain possesses the tRNA molecules for

codons that are rare in prokaryotic organisms. The sequence encoding the mature domain of

the hBMP-6 includes 12 of these codons (1 AGA, 1 CGA, 1 ATA, 3 AGG, 3 GGA and 3 CCC),

which could cause frame-shifting during the translation of the sequence (with the consequent

formation of truncated proteins) if the used strain does not express sufficient of these rare

tRNAs (Kane JF, 1995).

The addition of IPTG to the culture of bacteria transformed with the pET17b:rhBMP-6

expression vector resulted in a clear reduction of the growth rate, what could be due to high-

level expression of heterologous proteins. Indeed, SDS-PAGE analysis of the total protein

content of the bacteria revealed the presence of one highly-represented, ±16.5 KDa protein

which was assumed to correspond to the rhBMP-6 molecule. Although this protein constituted

more than 40% of the total protein content and most of it could be isolated in the insoluble

protein fraction of the cells, it is well known that the following in vitro manipulations needed to

obtain a purified sample of correctly refolded and biologically active growth factors imply a

great loss of proteins. This is especially important in the case of BMPs, which need a complex

refolding procedure to obtain correctly folded dimers and have a great tendency towards

aggregation and precipitation in their unfolded states.

Most of the methods used to obtain active BMPs expressed in Escherichia coli consist in

complex purification steps to separate the monomers from the rest of the protein content of the

cells, followed by dialysis, in vitro refolding and one or two more purification steps to isolate the

obtained dimers from the remaining monomers (Ruppert R et al., 1996; Long S et al., 2006). All

this makes the obtaining of active BMPs a long and complex process with many in vitro

manipulations and important losses of recombinant proteins during the entire procedure. To

partially avoid these problems we followed the method described by Vallejo LF et al., 2002 for

the renaturation and purification of BMP-2, which also resulted useful for the successful

refolding and purification of an rhBMP-2-CBD (Visser R et al., 2009). This method simplifies the

needed procedure as it consists in isolation of the produced inclusion bodies by simple washing

steps, followed by solubilization, direct refolding of the non-purified monomers, and one single

purification step to separate the dimeric molecules from the remaining monomers.

One of the main issues during the in vitro refolding of BMPs is the loss of proteins due to

aggregation of the folding intermediates by their exposed hydrophobic patches and consequent

179

Discussion_____________________________________________________________________

precipitation. Different strategies can be used to minimize this aggregation, such as performing

the renaturation at very low protein concentrations, at low temperature and/or including

antiaggregants into the refolding mixture. The method described by Vallejo LF et al. assessed

renaturation of rhBMP-2 and rhBMP-2-CBD at high concentrations (up to 50 µg/mL) by the

addition of the antiaggregant CHES, which seems to stabilize the folding intermediates,

preventing them from establishing hydrophobic interactions with other molecules.

Unfortunately, the refolding conditions established by these authors resulted not to be suitable

for the in vitro refolding of rhBMP-6 and, in consequence, forty other conditions were designed

and assayed.

All the 41 combinations of parameters tested for the refolding of the rhBMP-6 monomers

gave rise to a mixture of monomeric, dimeric, trimeric, tetrameric and polymeric associations,

although the ratio of dimeric BMP-6 versus other forms was always low. The fact that this effect

could be reverted in the presence of a reducing agent such as DTT demonstrated that these

associations of monomers were due to the establishment of disulfide bonds between them and

not to hydrophobic or other interactions. In most of the cases in which CHES was used as

antiaggregant, the degree of association was relatively low, and most of the rhBMP-6 remained

monomeric. In contrast, when the antiaggregant was changed to a non-detergent sulfobetaine

or to L-arginine, the majority of the peptides formed high molecular mass polymers. This effect

was only mimicked by a CHES-containing refolding mixture when the protein concentration was

raised to 53.4 µm/mL. Thus, it could be thought that NDSB265 and L-arginine are less suitable

antiaggregants than CHES for BMP-6 refolding, since these molecules are supposed to interact

with the peptides, hiding their hydrophobic patches and exposing their reactive cysteines to

allow the establishment of disulfide bonds. NDSB265 and L-arginine seem to interact less with

the BMP-6 molecules, allowing the formation of many unspecific disulfides in a similar way as

when the protein concentration is increased and the probability of encounter between two

peptides is raised. In contrast, the interaction between CHES and the BMP-6 peptides seems to

result in less exposure of the cysteines and, thus, fewer disulfide bond establishments.

When the same conditions are used for the refolding of rhBMP-2, only monomeric and

dimeric molecules can be observed after 72 hours (Vallejo LF et al., 2002; Chen B et al.,

2007a ; Visser R et al., 2009) . This indicates that, in contrast to BMP-2, BMP-6 has multiple

thermodynamically stable conformations, although only one of these is supposed to correspond

to the biologically active molecule.

In any case, since the rate of unspecific disulfide bond formation was very high during the

refolding process, it might be assumed that even among the dimeric fraction observed under

some conditions, only a small percentage (if any) would correspond to correctly folded BMP-6.

According to this, attempts on purifying these putative active dimers were considered unviable

and it was concluded that in vitro refolding of rhBMP-6 expressed in Escherichia coli is not

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Discussion

suitable. This conclusion was later supported by other groups, which also made unsuccessful

approaches to E. coli-derived BMP-6 refolding in vitro (Saremba S et al., 2008; personal

communications).

5.3. Production of rhBMP-6 and rhBMP-6-CBD in Sf9 cells.

Since we were unsuccessful in obtaining biologically active rhBMP-6 by the Escherichia coli

expression system, we decided to make a second attempt using a eukaryotic expression

system. We chose a baculovirus-Sf9 system, since these cells provide a suitable cellular milieu

for the production of heterologous proteins and can carry out the main posttranslational

modifications, such as signal peptide cleavage, phosphorylation, N- and O- glycosylation,

disulfide bond formation, and substitution of unusual analogs into proteins, in a similar way as

mammalian cells (Luckow VA, 1991). These expression systems are also known to be cheaper

and to produce higher yields than typical mammalian cell expression systems. Furthermore,

besides many structurally simpler proteins, some proteins containing a cysteine knot have been

produced with success using these kinds of expression systems, including murine and Xenopus

PDGF (Wang C et al., 1992), a human BMP-2 (Maruoka Y et al., 1995) and several Xenopus

BMPs (Hazama M et al., 1995; Aono A et al., 1995).

The high degree of unspecific disulfide bond formation observed after the attempts on in

vitro refolding of rhBMP-6 produced in E. coli made us decide to use a baculoviral system that

co-expresses the protein disulfide isomerase (PDI), since this foldase is known to facilitate both

disulfide bond formation and isomerization (Gruber CW et al., 2006) and it has been shown that

its co-expression can enhance the secretion of heterologous proteins expressed in insect cells

(Hsu TA et al., 1996).

We also decided to use an expression system in which recombinant protein expression is

under control of the polyhedrin promoter (PPolh), which is strongly activated during the late

stages of the infection cycle. This promoter is the most used for heterologous protein

expression by baculoviral systems, since it ensures the highest yields of recombinant proteins.

The analysis of the production assays performed for rhBMP-6 and rhBMP-6-CBD showed

that both proteins were mainly found in the culture media in the form of trimers, although a

significant percentage of dimers and small amounts of tetramers and higher polymers were also

observed, somewhat resembling the results obtained for the in vitro refolding of the rhBMP-6

produced in E. coli. Nevertheless, no signs of unfolded monomers were found.

The concentration of the recombinant proteins in soluble form was usually highest when

production times of 72 hours were used, diminishing when the infection was continued for

longer periods. This effect could be due to protein aggregation followed by precipitation when

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Discussion_____________________________________________________________________

the concentration in the culture medium reaches a certain threshold. Nevertheless, since the

protein concentration is far below the detection limit for Coomassie staining and the intensity of

the bands is even lower than the one corresponding to the commercial rhBMP-6 used as a

positive control, protein loss due to precipitation seems improbable. More likely, the loss of

proteins could be due to proteolytic degradation by proteases secreted by the Sf9 cells or

liberated to the medium by them after cell lysis.

In order to determine if the BMP-6 dimers and trimers were disulfide linked, the samples

were analyzed by Western blot under reducing conditions. In presence of DTT, which reduces

the cysteine residues of the peptides, the bands corresponding to the dimeric, trimeric and

tetrameric forms disappeared and were substituted by a doublet of low molecular mass bands,

what indicates that the oligomers were stabilized by disulfide bonds. Although the predicted

molecular masses of the rhBMP-6 and rhBMP-6-CBD monomers are ±18 KDa and ±19.3 KDa,

respectively (attending to their amino acid sequences), the appearance of a slightly higher

molecular mass band was not surprising, since the commercial rhBMP-6 is described to run as a

triplet of bands. These additional bands could be due to non-homogeneous glycosylation of the

BMP-6 population or to the production of truncated or elongated peptides by frame shifting

during translation.

Since the only biologically active form of the BMPs is a dimer, we wanted to determine if

the excessive establishment of intercatenary disulfides, which give rise to the trimeric and

tetrameric BMP-6, was due to the co-expression of the PDI and if the yield of BMP-6 dimers

could be improved by expressing the proteins in the absence of heterologous PDI. The analysis

of the rhBMP-6 production assay without PDI co-expression showed that the formation of these

higher oligomers was PDI-independent, since the pattern of bands that could be observed was

identical to the one obtained when using the Sapphire™ expression system. According to this, it

could be assumed that the presence of this foldase does not negatively affect the BMP-6

production, since it did not increase the formation of intercatenary disulfides. Nevertheless, the

PDI could still be beneficial for the formation of correctly folded BMP-6 dimers among this

fraction, since this chaperone not only promotes the formation of disulfides, but also catalyzes

their isomerization. Thus, if the correctly folded structure of the BMP-6 dimer is

thermodynamically more stable than the other possible conformations, the presence of the PDI

might help to increase the rate of correctly versus incorrectly folded dimers.

Another possible explanation for the high yield of incorrectly folded BMP-6 molecules

obtained is the election of the polyhedrin promoter to drive the heterologous protein

expression. This promoter is strongly activated during the very late phase of the viral infection

cycle, a stage at which the cellular protein-production machinery, re-programmed by the viral

DNA, is fully committed to the production of large quantities of new virions. In this

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Discussion

compromised stage, the activation of the PPolh forces the synthesis of huge amounts of

rhBMP-6(-CBD) monomers, which are translocated into the endoplasmic reticulum by the GP67

signal peptide. It does not seem improbable that the accumulation of heterologous BMP-6

inside the endoplasmic reticulum of a cell in an altered state due to infection by a lytic virus

could lead to an alteration of the endoplasmic luminal microenvironment. This, together with

the high concentration of BMP-6 monomers inside the endoplasmic reticulum, could cause

protein aggregation and/or establishment of unspecific disulfides. Furthermore, possibly many

of these incorrectly folded proteins do not even reach the Golgi apparatus for their vesicular

secretion, but are directly liberated to the culture medium when the host cell becomes lysed. If

this were the case, these problems might have been overcome by the election of an earlier

promoter to drive the heterologous protein expression. If we would have used a milder

promoter we probably would have obtained lower yields of total rhBMP-6(-CBD), but possibly

the ratio of correctly versus incorrectly folded proteins would have been higher.

The purification of both the rhBMP-6 and rhBMP-6-CBD by heparin-sepharose

chromatography yielded almost identical elution profiles, with the maximum of dimeric

molecules obtained when eluting with conductivity values around 28 mS/cm. This indicates that,

in contrast to rhBMP-2 (Visser R et al., 2009), the addition of the CBD to the N-terminus of the

BMP-6 molecule does not alter its heparin-binding properties.

Unfortunately, the fact that the conditioned culture media contained not only dimeric but

also high quantities of trimeric and tetrameric BMP-6 molecules, made the purification of the

dimeric fraction by this technique impossible, since the trimeric BMP-6 only showed a slightly

increased affinity to heparin when compared with the dimers. This behaviour could be

considered a logic consequence of the fact that rhBMP-6 dimers only possess two heparin-

binding domains, while the trimeric and tetrameric forms possess three and four of them,

respectively. This also supports the idea that the heparin-binding properties of BMP-6 are not

due to structural aspects of the heparin-binding site, but are only dependent on the net charge

of the amino acid sequence that forms this domain, since it might be assumed that in the

incorrectly folded, unnatural trimers and tetramers, the folding of the heparin-binding sites is

also compromised.

The selected elution fractions were those containing the highest dimer:trimer ratio. Before

we could test the biological activity or the collagen-binding properties of these samples, the

excess of salts and urea had to be removed. Nevertheless, this has to be done gradually, since

the BMPs are highly hydrophobic and show a great tendency towards aggregation and

precipitation in aqueous solutions. To achieve this, different strategies were tried: direct dialysis

against culture medium, dialysis against ammonium acetate followed by lyophilization and

buffer exchange to 4 mM HCl in a buffer exchange/concentration column. Only in the latter

case an important loss of proteins was observed, though no visible signs of protein precipitation

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Discussion_____________________________________________________________________

were detected during the process. It seems improbable that the protein was lost by passing the

column filter, since a column with a molecular mass cut-off of 5 KDa was used, so the only

possible explanation is that the BMP-6 molecules were partially lost by being retained by the

filter itself. In fact, when treating E. coli derived, in vitro refolded, purified rhBMP-2(-CBD) in

similar columns, more than 70% of the proteins are lost (unpublished data).

Serial dilutions of all the obtained samples were tested on their ability to induce the

transdifferentiation of cultured C2C12 mouse myoblastic cells into the osteogenic lineage. These

cells are not able to differentiate into osteoblasts by themselves but, in presence of BMPs, they

start expressing osteogenic markers (including ALP) and acquire an osteoblastic phenotype

(Katagiri T et al., 1994).

While the commercial rhBMP-6 produced in CHO cells induced the expression of ALP in a

typical dose-dependent manner, with an estimated ED50 close to 300 ng/mL, none of the tested

samples was able to induce the expression of this osteogenic marker, even at the lowest

dilutions/highest concentrations tested.

There are different possible causes which, alone or in combination, could explain the

inactivity of the recombinant BMP-6 molecules produced in Sf9 cells with the chosen expression

system:

1. The sequencing of the genes cloned in the pAcGP67B shuttle vector showed that they

were correctly inserted into the plasmidic DNA and that no mutations were introduced during

the amplification and/or cloning procedures. Furthermore, the Western blot analyses of the

conditioned culture media with a specific monoclonal anti-BMP-6 antibody revealed a pattern of

bands that corresponds to the expected pattern for a mixture of dimeric, trimeric and tetrameric

BMP-6 molecules. When the Western blot was performed under reducing conditions, almost all

of the proteins became reduced, giving rise to a doublet of bands which molecular mass

corresponds to the expected molecular mass for rhBMP-6 monomers. All these facts made us

assume that the bmp6 and bmp6-cbd constructs were correctly expressed and the peptides

translocated into the endoplasmic reticulum for posttranslational processing. Nevertheless, a

high percentage (or all) of the obtained dimeric rhBMP-6(-CBD) might be incorrectly folded due

to saturation and/or malfunctioning of the cellular protein production and processing machinery

in the late stages of the baculoviral infection cycle, during which the PPolh becomes activated.

Since the quaternary structure of the native BMPs is relatively complex, an incorrectly folded

BMP-6 dimer would probably not have a functional BMPR-binding site and would consequently

not be able to trigger the BMP signal transduction within the target cells.

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Discussion

2. Even if a certain percentage of the purified dimers would have the correct quaternary

structure, the presence of high concentrations of incorrectly folded dimers or higher oligomers

could be inhibiting the activity of the native molecules by acting as competitive inhibitors. This

possibility is supported by the fact that Weber FE et al. (2001) reported that dimeric BMP-2

molecules with incorrect intracatenary disulfides were able to inhibit both the in vivo heterotopic

ossification induced by BMP-2 and the maturation of the preosteoblastic MC3T3-E1 cell line to

osteoblasts in vitro. Although these aberrant dimers did not have the native quaternary

structure, they were still able to bind the BMP receptors but not to trigger the BMP signal

transduction within the cell, acting as real agonists for the native BMP-2 molecules.

3. Insect cells are able to carry out most of the posttranslational modifications described in

mammalian cells, including signal peptide cleavage, phosphorylation, N- and O- glycosylation,

disulfide bond formation, and substitution of unusual analogs into proteins. Nevertheless,

although the processing of N-glycans in insect and mammalian cells appears to follow a similar

initial pathway, the final processing seems to be different, being the N-glycans from insect cell

lines usually modified to paucimannosidic or oligomannose structures instead of terminally

sialylated complex-type structures (Marchal I et al., 2001; Tomiya N et al., 2004). Since

terminal sialic acid residues are known to play diverse roles in many glycoconjugates and the

glycosylation of BMP-6 seems to be strictly necessary for the binding of the growth factor to

type I BMPRs (Saremba S et al., 2008), it is possible that the glycosylation provided by the Sf9

cells is not sufficient to obtain active rhBMP-6 molecules. If this were the case, a possible

solution would be the expression of rhBMP-6 in a genetically engineered insect cell line that

carries out the terminal sialylation of its N-glycans. These type of cell lines are nowadays being

improved by introducing the genes for the expression of missing glycosyltransferases and of the

enzymes responsible for generating the essential donor sugar nucleotide required for sialylation,

as well as inhibiting the N-acetylglucosaminidase responsible for removing a terminal

N-acetylglucosamine from the N-glycan. In this sense, the aim of these genetically engineered

insect cell lines is to mimic the N-glycosylation pathway that occurs in mammalian cells to

improve the expression of heterologous glycoproteins (Tomiya N et al., 2004).

In conclusion, the chosen baculoviral/insect cell expression system seems not to be suitable

for the production of complex, cysteine knot proteins such as rhBMP-6. In fact, similar attempts

on rhBMP-2 production with this system performed in our group also resulted unsuccessful,

since the obtained proteins were also inactive (unpublished data). Perhaps the election of a

milder and earlier promoter to drive the heterologous protein expression in combination with a

so-called “humanized” insect cell line as a host would have yielded better results. These options

might be taken in consideration if future attempts on rhBMP-6 production in insect cells were

done.

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Discussion_____________________________________________________________________

5.4. Production of rh-bFGF and rh-bFGF-CBD in Sf9 cells.

In order to produce these growth factors with a non-prokaryotic expression system, we

decided to make an attempt on the production of rh-bFGF and rh-bFGF-CBD in insect cells.

Since the structure of bFGF is much simpler than the one of BMP-6 (monomeric, without

disulfides, not glycosylated), some of the problems that arose during the production of rhBMP-6

might not be an issue for the production of bFGF.

The lack of disulfide bonds in the tertiary structure of the bFGF molecule made us decide to

use a simple baculoviral expression system, with no co-expression of any heterologous

chaperone. Nevertheless, we still used a system in which the proteins carry the GP67 signal

peptide for their translocation into the endoplasmic reticulum. Although in mammalian cells the

secretion of bFGF seems to be by direct translocation through the plasma membrane mediated

by cell surface HSPGs (Backhaus R et al., 2004; Schäfer T et al., 2004; Zehe C et al., 2006), we

were not able to predict if the Sf9 insect cells would have the capacity to secrete the

heterologous bFGF(-CBD) to the culture medium in a vesicle-independent manner. Although

three homologs to FGFs (Pyramus (Pyr), Thisbe (Ths) and Branchless (Bnl)) have been

identified in Drosophila (Kadam S et al., 2009), the secretion mechanisms that affect these

growth factors in insect cells are not known. Thus, the expression of a heterologous bFGF in

insect cells without addition of any signal peptide might lead to accumulation of the proteins in

the form of inclusion bodies within the cells, being this something we wanted to avoid, since

this would have forced us to carry out additional manipulations of the proteins in order to

isolate and solubilize them.

The analysis by Western blot of the production assays performed for rh-bFGF and

rh-bFGF-CBD showed that the specific anti-bFGF antibody recognized one single band in each

sample. The molecular masses of these bands corresponded to the predicted molecular masses

for the rh-bFGF and the rh-bFGF-CBD, according to their amino acid sequences. Thus, it seems

that the passing of these proteins through the endoplasmic reticulum does not provoke

oligomerization by intercatenary disulfide establishment, or apparent unspecific glycosylation.

Nevertheless, we were not able to ensure that the produced proteins exhibited their native

conformation since, for example, the formation of unspecific intracatenary disulfides could have

lead to their incorrect folding.

For both rh-bFGF and rh-bFGF-CBD, the highest concentrations of heterologous proteins in

the culture media were achieved with production times of 72 hours. When the proteins were

harvested after longer production times, the intensity of the bands decreased and a lower

molecular mass smear appeared. The appearance of this smear could be due to partial

proteolytic degradation of the rh-bFGF(-CBD) by proteases secreted by the insect cells or

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Discussion

187

liberated to the medium by them after their lysis. Another possible explanation is that, while the

infection of the culture progresses, the number of cells that becomes lysed increases. By their

lysis, these cells liberate their total cell content to the culture medium, including truncated or

incompletely synthesized rh-bFGF(-CBD) molecules.

The purification of both rh-bFGF and rh-bFGF-CBD were performed by heparin-sepharose

affinity chromatography, eluting the proteins with a linear gradient of NaCl, from 0.15 to

2.00 M. Since bFGF is known to be much more stable than BMPs in aqueous solutions, its

purification was performed in absence of urea but in presence of DTT to avoid air oxidation of

the cysteine residues. Apparently, the elution profiles for both proteins were more or less

similar, with bFGF found along a great portion of the profile. Since the achieved gradient of

NaCl was almost completely linear, the fact that bFGF molecules were eluted from the column

along almost the entire gradient made us conclude that the conditioned culture media used for

purification contained a heterogeneous mixture of rh-bFGF(-CBD) molecules with different

affinities to heparin. This was further supported by the fact that the concentration of the

heterologous proteins suffered variations along the elution profile, defining at least two distinct

populations for each of the produced bFGFs: a “low heparin-affinity” population (constituted of

the majority of the produced growth factors, which eluted with low to moderate concentrations

of NaCl) and a “high heparin-affinity” population (represented by a smaller percentage of the

produced growth factors, which eluted with high NaCl concentrations). Although the behaviours

of rh-bFGF and rh-bFGF-CBD were slightly different during the purification, these differences

were not great enough to allow us to ensure that the addition of the CBD sequence is

significantly altering the heparin-binding properties of the bFGF molecule. In the case of the

rh-bFGF, the protein was detected in the elution fractions until 138 mS/cm, while in the case of

the rh-bFGF-CBD, all of the proteins bound to the column could be eluted with 121 mS/cm.

Nevertheless, more purifications of both proteins should be performed to determine if these

differences are reproducible.

There are different possible causes which, alone or in combination, could explain the

existence of these populations with different affinities to heparin in the culture media:

1. The fact that the expressed rh-bFGF(-CBD) molecules are driven into the endoplasmic

reticulum might be causing changes to the molecules that could not be detected by Western

blot analysis. It is possible that the special microenvironment inside the endoplasmic reticulum

could be inducing the formation of intracatenary disulfides or partial glycosylation of the

molecules, affecting the tertiary structure of the protein and/or the conformation of the amino

acid residues involved in binding to heparin.

Discussion_____________________________________________________________________

2. As hypothesized for the production of BMP-6 in Sf9 cells, the election of a promoter that

becomes activated during the very late phase of the infection cycle to drive the heterologous

protein expression might be responsible for the synthesis of truncated or misfolded proteins

since, at this stage, the host cell is close to its lysis and the protein production machinery of the

cell might be starting to malfunction.

After purification, we obtained one “low heparin-affinity” and one “high heparin-affinity”

sample for each of the heterologous growth factors. Although the purification of the proteins

was performed in the absence of urea, the high salt concentrations (0.6 - 0.8 M in the “low

heparin-affinity” samples and 1.4 – 1.5 M in the “high heparin-affinity” samples) had to be

reduced to more physiological levels before testing the biological activity or the collagen-binding

properties of the purified proteins. For this purpose, the samples were loaded on buffer

exchange columns and the elution buffer was changed to PBS, pH 7.3, 1 mM DTT, 1 mM EDTA.

By repeated dilution and centrifugation steps, the salt molarity in all the samples was reduced

below 20 mM. Western dot blot analysis of the samples after the buffer exchange showed that,

surprisingly, both “low heparin-affinity” samples suffered an important loss of proteins during

the process, while the proteins in the “high heparin affinity” samples were successfully

concentrated. This indicates that the majority of the heterologous rh-bFGF(-CBD) produced in

Sf9 cells not only has a diminished affinity to heparin, but also a reduced solubility in aqueous

solutions, pointing to more important disorders in the molecular structure than initially thought

which, among others, could have varied the pI of the proteins. In fact, these observations made

us assume that the probabilities of these molecules to have a diminished or lost biological

activity were quite high.

It is known that binding of bFGF to HSPGs present in the extracellular matrix or on the

target cell surface is important for bFGF signal transduction (Folkman J et al., 1988; Dowd CJ et

al., 1999). Furthermore, since the protein concentration in the “low heparin-affinity” samples

after the buffer exchange was too low to allow proper testing of their biological activity and

collagen-binding properties, and so was their probability of being active, we decided to discard

them and focus only on the “high heparin-affinity” samples.

The concentration of the samples was estimated by Western blot and digital image analysis.

With the obtained values, the yields of heterologous proteins per litre of conditioned culture

medium were calculated to be ±33.50 µg/L for rh-bFGF and ±13.80 µg/L for rh-bFGF-CBD.

These values are 3 orders of magnitude below the ones described for typical heterologous

protein productions with baculoviral/insect cell expression systems, which are between 10 and

100 mg/L. Nevertheless, the facts that a very high percentage of the rh-bFGF(-CBD) was

incorrectly synthesized and discarded after purification and buffer exchange and that probably

another percentage of the “high heparin affinity” samples was lost during these processes

could, in part, explain these low yields.

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Discussion

Since the addition of the sequence of the CBD implies that the rh-bFGF-CBD has an 8%

increase in its molecular mass over the rh-bFGF, the in vitro and in vivo experiments to

determine their biological activity and collagen-binding properties were designed using the

molar concentrations of the samples, in order to compare the effects of equal numbers of

effective molecules.

To determine if the addition of the CBD in the engineered rh-bFGF-CBD conferred to this

molecule an increased affinity to collagen, a collagen-binding affinity test was performed

following a method described by T. Kitajima in 2007. Therefore, we used a 1 mm thick collagen

type I sheet obtained from highly pure, native, bovine skin collagen, from which 5 mm diameter

discs were cut. This collagenic carrier has been recently approved by the FDA for its use in

combination with BMP-2 and is similar to the other ones used in clinical applications.

After loading the samples and washing the ACSs for one hour with buffer, almost no

immunostaining was detected on the sponges incubated with commercial rh-bFGF, indicating

that this growth factor has no natural affinity to collagen type I in this presentation. Very

surprisingly, the sponges that were incubated with 1.25 pmol of the rh-bFGF produced in Sf9

cells showed immunostaining with the anti-bFGF antibody after the washing with the buffer.

This indicates that, although the sequence of the bfgf gene was cloned into the used shuttle

vector without any modifications on the native sequence and the performed PCR analyses and

sequencing revealed no mutations or alterations of the gene, the rh-bFGF molecule produced in

Sf9 insect cells has to have some slight chemical or structural differences when compared with

the commercial rh-bFGF. Nevertheless, since the commercial growth factor used for comparison

was produced in Escherichia coli, these observations might be a simple consequence of the

natural differences between prokaryotic and eukaryotic expression systems.

The immunostaining of the sponges that were incubated with 1.25 pmol of rh-bFGF-CBD

showed that these proteins had a significant higher retention than the non-modified rh-bFGF. It

was not possible to determine if the total affinity to collagen type I of the rh-bFGF-CBD when

compared to the commercial rh-bFGF was due only to the CBD, or if it was the sum of the

effect of the CBD and the effect of the used expression system. In any case, it could be clearly

concluded that the addition of the decapaptidic collagen binding domain to the bFGF molecules

increases its affinity to collagen type I in the form of an absorbable collagen sponge.

Since our goal was the production of an active form of bFGF with the ability of being

retained for longer periods of time at the wound site when implanted in vivo for bone repair

purposes, with a consequent enhanced effect on tissue healing and safety of the clinical

approach, we needed to demonstrate that the binding of the rh-bFGF-CBD to ACSs was stable

in time. Therefore, the same collagen-binding affinity test was repeated, though

immunostaining of the sponges was performed after 6 days of extensive washing with buffer.

According to the results, a high percentage of both rh-bFGF and rh-bFGF-CBD remained bound

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Discussion_____________________________________________________________________

to the collagen sponges after this period and, also under these conditions, the amount of

rh-bFGF-CBD found in the sponges was higher than the amount of rh-bFGF. These results

indicate that the rh-bFGF and rh-bFGF-CBD not only posses a higher affinity to ACSs compared

to E. coli-derived rh-bFGF, but also that this binding is stable for long periods of time. In

addition, the CBD confers an additional affinity to collagen to the molecule, resulting in over

50% more binding. In consequence, the use of these growth factors for clinical bone repair in

combination with absorbable collagen sponges could possibly allow the implantation of lower

concentrations of them, reducing the system dispersion and, thus, enhancing the effectiveness

and safety of the treatment. Furthermore, the specific binding of the proteins to the carrier

might avoid the great initial loss of proteins that occurs due to manipulation of the sponge by

the surgeon prior to its implantation.

Since bFGF is known to be a mitogen for most cells of mesodermic origin, the biological

activity of the produced growth factors was determined in vitro by a proliferation assay on

MC3T3-E1 murine preosteoblasts, calculating the number of cells in the cultures after 72 hours

of incubation with the different bFGFs or without any growth factor. For this purpose we used

an MTT-based method, which is generally considered reliable and highly reproducible. Since the

use of MTT implies the counting of only living, metabolically active cells, possible cell death

events in the cultures are not taken into account and, in consequence, this was not a

proliferation assay sensu stricto. Nevertheless, by direct observation of the cultures during

incubation with the MTT-labelling reagent, no signs of death cells or debris could be observed in

any of the cultures, so the rate of cell death was considered negligible.

The results showed that, at low concentrations (156.25 or 312.50 pM), both rhbFGFs

produced in Sf9 cells and the commercial rh-bFGF were able to significantly increase the

number of cells in the cultures, achieving similar yields in all the three cases. Although these

concentrations are denoted as “low”, they are actually around or even above the ED50 described

for the activity of bFGF in many processes. In contrast, at higher concentrations (625.00 pM or

1.25 nM), the growth factors produced in Sf9 cells were not able to further increase the cell

density in the cultures, whereas the commercial rh-bFGF induced a significantly higher

proliferation. Nevertheless, it should be noticed that the variability was much higher among the

cultures treated with the commercial rh-bFGF than the ones found among the cultures treated

with the rh-bFGFs produced in insect cells.

To further analyze the biological activity of the growth factors in vitro, we decided to test

their ability to inhibit the osteoblastic differentiation of MC3T3-E1 mouse preosteoblasts induced

by ascorbic acid. Since the expression of alkaline phosphatase is considered an early marker of

osteoblastic differentiation, we measured the inhibition by bFGF as the inhibition of the activity

of this enzyme within the culture media and the cells. After 120 hours of incubation with the

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Discussion

growth factors, both the number of cells and the ALP activity in the cultures were determined

and the average ALP activity per cell calculated.

The results showed that, after 120 hours of incubation with any of the bFGFs, no traces of

ALP activity could be detected in the cultures. In contrast, the control cultures grown in the

absence of bFGF showed very low, though significant levels of ALP activity after this period of

time. The differences between the cultures incubated with bFGF and the control cultures

became higher when comparing the average ALP activity per cell, since the number of cells in

the latter was lower than in the cultures incubated with commercial or Sf9-derived

rh-bFGF(-CBD). Nevertheless the period of time established for this assay was not long enough

to affirm that the ability of the three bFGFs to inhibit the osteoblastic differentiation of the cells

was the same since, at this time, the cells were just starting to express the studied osteogenic

marker. Thus, longer incubations with the growth factors should be performed in future assays

in order to detect possible differences between their activities. The obtained results after 120

hours only allowed us to conclude that both the commercial rh-bFGF and the rh-bFGF(-CBD)

produced in Sf9 cells were able to, at least, delay the osteoblastic differentiation of mouse

preosteoblasts.

5.5. In vivo osteogenic activity of combinations of BMP-6 and bFGF.

In order to evaluate the biological activity of the rh-bFGF and rh-bFGF-CBD produced in Sf9

cells and their capacity to enhance bone formation when administered in combination with

BMP-6, absorbable collagen sponge discs carrying 13.89 pmol (0.5 µg) rhBMP-6 alone, 1.25

pmol of rh-bFGF or rh-bFGF-CBD alone, or combinations of these factors, were implanted into

the dorsal muscles of rats. Commercial rh-bFGF produced in E. coli was used as a positive

control, while ACS discs loaded with vehicle only served as a negative control. Previous works of

our group showed that, when inducing heterotopic osteogenesis with 0.5 µg of rhBMP-2, no

cartilage rests could be found when analyzing the samples 28 days after surgery (unpublished

data). Since we wanted to compare the quality and maturity of the bone induced by the

different combinations of growth factors tested in this work, we decided to analyse the samples

in an earlier stage (21 days after surgery).

As expected, when implanted alone with ACSs, none of the bFGFs were able to induce

heterotopic osteogenesis by themselves. It has been described that low concentrations of bFGF

can act synergically with BMP-2 to enhance osteogenesis in vivo (Fujimura K et al., 2002;

Nakamura Y et al., 2005; Tanaka E et al., 2006; Kakudo N et al., 2006) but, as stated earlier,

BMPs are the only growth factors known to have the capacity to induce ectopic bone formation

191

Discussion_____________________________________________________________________

in adult vertebrates (Wang EA et al., 1990; Volek-Smith H and Urist MR, 1996; Nakase T and

Yoshikawa H, 2006).

Nevertheless, the fact that all of the implanted control sponges, loaded with vehicle only,

were reabsorbed and could not be recovered 21 days after surgery, while most of the implants

loaded with bFGF had formed a fibrotic nodule within the rats muscle, indicated that both the

commercial rh-bFGF and the rh-bFGFs produced in insect cells possess biological activity in vivo.

All the three growth factors induced the formation of a dense, fibrotic tissue in which irregular

accumulations of basic, eosinophyllic extracellular material were observed. These were

especially abundant and apparent in the implants that were loaded with Sf9 cell-derived

rh-bFGF or rh-bFGF-CBD. Since these two growth factors demonstrated to possess specific

affinity to the carrier in vitro, probably the accumulation of this unidentified extracellular

material was due to the retention of these bioactive proteins at the implant site. If this were the

case, it would be indicating that the binding to ACS of both rh-bFGF and rh-bFGF-CBD produced

in insect cells remains stable when implanted within living tissue.

In contrast to the implants with bFGF alone, the implants loaded with 13.89 pmol (0.5 µg)

rhBMP-6 (alone or in combination with bFGF) all gave rise to osteogenic events, resulting in the

formation of a trabecular bony tissue. Nevertheless, clear differences could be observed among

the different combinations assayed.

The implants that were loaded with rhBMP-6 alone were, in general, smaller in size than the

ones that were loaded with BMP-6 and bFGF. In these implants, osteogenesis was mainly

limited to the periphery, where clear bone trabeculae were formed. The fact that only little

staining with alcian blue (which stains the components of the cartilaginous matrix) could be

detected in these trabeculae made us assume that they had reached a certain degree of

maturity. This assumption was based on the work of other authors, which have described that

heterotopic bone formation induced by BMP-2 occurs through endochondral ossification (Wang

EA et al., 1990; Nakagawa T et al., 2003). Furthermore, since traces of hypertrophic cartilage

could be detected in some of the analyzed implants, it seems that BMP-6-mediated ossification

also follows the endochondral pathway.

The inner part of these implants was constituted of a dense accumulation of mesenchymal

cells, not resembling a real bone marrow tissue. Only few disperse adipocytes could be

observed and almost no vascularization. Immunostaining of these implants with a specific anti-

osteopontin antibody revealed the existence of this early marker of osteogenesis mainly in the

edges of the trabeculae. Osteopontin is a cell-to-matrix adhesion molecule which is secreted by

osteoblasts after they start synthesizing ALP. Its expression is maintained during the entire

period of bone matrix synthesis so that the protein diffuses through the new-formed osteoid

and becomes trapped within this osteoid after its mineralization (Roach HI, 1994). According to

this, the presence of osteopontin mainly in the external parts of the trabeculae indicated that,

192

Discussion

although new osteoid was still being synthesized on their surface, the inner areas of the

trabeculae were formed by more mature, mineralized osteoid. In contrast, almost no

osteopontin was detected in the inner part of these implants and only few osteoblasts

expressing this marker could be found, indicating that osteogenesis was very limited in this

area. This is consistent with the observation that almost no angiogenesis occurred within these

implants, since the ingrowth of new blood vessels is known to sustain osteogenesis by providing

osteoprogenitor cells to the new-forming bone.

In contrast, the implants that were loaded with rhBMP-6 in combination with commercial or

insect cell-derived bFGF were not only bigger in size in general, but also showed osteogenic

activity throughout their entire volume, with bony trabeculae being found at the periphery and

at the inner part of the implants. In both cases, these trabeculae were intensely stained with

the light green present in the Masson’s trichrome, but showed only small, heterogenously

distributed alcian blue-positive patches, indicating that they were mainly composed of collagen

but that little glycosaminoglycans were present within them. These observations, together with

the fact that almost no hypertrophic chondrocytes could be detected, were indicating that most

of the trabeculae found in these implants were in an advanced stage of the chondro-osseous

transition. In between the trabeculae, a tissue that resembled bone marrow was found,

presenting many adipocytes and small blood vessels. Immunostaining of these implants with a

specific anti-osteopontin antibody showed that this bone marrow-like tissue contained abundant

osteoblasts expressing this protein and small osteopontin accumulations in the extracellular

matrix, indicating that these implants were exhibiting a greater osteogenic activity than the

ones loaded with rhBMP-6 alone.

One of the differences found between these two implant types was that some of the ones

loaded with rhBMP-6 + commercial rh-bFGF showed areas of osteopontin-containing material

with a fibrous, disorganized appearance, while this was not observed in the implants loaded

with rhBMP-6 + Sf9 cell-derived rh-bFGF.

The second main difference affects the blood vessels found in the samples. Both implant

types showed a high degree of vascularization, presenting immature blood lacunae (in which no

clear defined endothelium could be observed), many small and some big blood vessels.

Nevertheless, vascularization seemed to be slightly lower in the implants loaded with rhBMP-6 +

commercial rh-bFGF than in the ones loaded with rhBMP-6 + Sf9 cell-derived rh-bFGF, with the

latter showing both more and bigger vessels. In any case, some of the blood vessels found in

both implant types showed, on their abluminal side, small groups of periendothelial cells

immediately opposed to the endothelium. This could be interpreted as a sign of maturation of

these blood vessels, what could be important for the osteogenic potential within these samples

since recent studies have given strong evidences of a perivascular origin for mesenchymal stem

cells. According to these studies, pericytes (or adventitial reticular cells in bone marrow) present

in perivascular locations would actually be the MSCs (Crisan M et al., 2008; da Silva Meirelles L

193

Discussion_____________________________________________________________________

et al., 2008). Thus, the presence of more and more mature blood vessels in these samples

could be responsible for the higher osteogenic activity observed and, since almost no

angiogenesis could be detected in the implants loaded with rhBMP-6 alone, it could be

concluded that the formation of these blood vessels was induced by the presence of the

exogenous bFGF. Furthermore, this bFGF might also be contributing to the osteogenic activity

within the samples by inducing the proliferation and/or differentiation of the mesenchymal

osteoprogenitor cells.

From all these facts it could be concluded that, when 1.25 pmol of growth factor are used,

the commercial E. coli-derived rh-bFGF is able to enhance BMP-6-induced osteogenesis in vivo

and that the Sf9 cell-derived rh-bFGF is at least as efficient as the commercial protein.

Surprisingly, although the implants that were loaded with rhBMP-6 + rh-bFGF-CBD were

comparable in size to those loaded with rhBMP-6 + commercial or Sf9 cell-derived rh-bFGF, the

bony tissue they contained showed signs of being less mature. Well defined trabeculae were

found at the periphery and, to a greater or lesser extent, also at the inner part, but not always

throughout the entire implant. Furthermore, staining of the samples with alcian blue revealed

the existence of some extensive positive areas which contained many hypertrophic

chondrocytes, indicating that the chondro-osseous transition in these areas was in a relatively

early stage. Anti-osteopontin immunostaining of these implants showed that this osteogenic

marker was heterogenously localized among the trabeculae. Especially surrounding the areas

containing cartilage, osteopontin was detected as abundant fibrous-like accumulations.

Other signs of the immaturity of these samples were the little angiogenic activity observed

and the lack of adipocytes in the bone marrow-like tissue found in between the trabeculae.

These observations are in opposition to the in vitro properties exhibited by the rh-bFGF-

CBD. The ability of this molecule to induce the proliferation of MC3T3-E1 mouse preosteoblasts

in vitro was equal to that of the Sf9 cell-derived rh-bFGF, so the effectiveness of both molecules

to enhance heterotopic osteogenesis in vivo was expected to be, at least, the same.

Nevertheless the augmented affinity to collagen type I of the rh-bFGF-CBD could explain the

obtained results. Different authors have reported that low concentrations of bFGF enhanced

bone formation in vivo when co-administered with BMP-2, but also that higher concentrations of

bFGF had the opposite effect (Fujimura K et al., 2002; Nakamura Y et al., 2005; Tanaka E et

al., 2006; Kakudo N et al., 2006). In all these studies, above a certain threshold concentration,

bFGF inhibited heterotopic bone formation in a dose-dependent manner. In this line,

Kakudo N et al. showed that co-administration of 2 µg rhBMP-2 and 16 ng (1.00 pmol) rh-bFGF

with a collagenic carrier resulted in more bone formation than when rhBMP-2 alone was used,

whereas bone formation was suppressed when the amount of rh-bFGF was raised to 80 ng

(5 pmol).

194

Discussion

In the present work, 500 ng rhBMP-6 were co-administered with 21.6 ng (1.25 pmol)

rh-bFGF-CBD. This amount of rh-bFGF-CBD should have been low enough to enhance

osteogenesis as observed when the same amount of rh-bFGF was used. Nevertheless, since the

CBD had demonstrate to confer an increased affinity to collagen to the molecule, the

persistency of the rh-bFGF-CBD at the implant site could have caused the same effect as

administering bFGF above the osteogenesis-supressing threshold. If this were the case, the

rh-bFGF-CBD dose should be lowered to obtain the desired enhancement of bone formation

5.6. Perspectives for the future.

Among the BMPs, BMP-6 is one of the most potent osteogenic growth factors known. In

fact, only 300 ng of rhBMP-6 were able to induce the formation of a well-defined bony tissue in

a rat heterotopic osteogenesis model (unpublished data), whereas at least 460 ng of rhBMP-2

are needed to observe some signs of osteogenesis (Wang EA et al., 1990). Nevertheless, the

medical and scientific communities are mainly focused on BMP-2, rather than on other BMPs.

The search in PubMed Central (http://www.ncbi.nlm.nih.gov/pubmed/) for the keywords

“BMP2” or “BMP-2” or “bone morphogenetic protein-2” yielded 4,115 articles until may 2009,

while the search for the keywords “BMP6” or “BMP-6” or “bone morphogenetic protein-6”

yielded only 411 articles.

On the other hand, we and other authors have demonstrated that low doses of bFGF can

enhance bone induction when administered together with BMPs. We have also demonstrated

that the addition of a decapeptidic collagen-binding domain derived from the von Willebrand

Factor to rh-bFGF produced in insect cells increases its affinity to absorbable collagen type I

sponges without affecting its biological activity in vitro.

The results of the heterotopic bone formation assay in rats, implanting ACSs with rhBMP-6

and the different bFGFs used in this work, were stimulating enough to induce us to believe that

the combination of absorbable collagen, BMP-6 and collagen-targeted bFGF could be a more

effective and safer alternative to the today available biomaterials for clinical repair of bone

defects.

Nevertheless, additional studies should be performed in the future to prove this hypothesis.

The aim of these studies would be the optimization of the production and purification of the

heterologous proteins produced in insect cells, as well as further characterization of the

structural and biochemical features of these molecules. Also, additional in vivo heterotopic bone

formation assays should be planned for determination of the most effective dose for each

growth factor and for their combination. Furthermore, both the osteogenic and angiogenic

events occurring in the implants should be studied in detail, analyzing samples recovered earlier

195

Discussion_____________________________________________________________________

and later after surgery, determining the expression of osteogenic markers and the maturity of

the blood vessels and their perivascular structures. At last the combination of ACS, BMP-6 and

collagen-targeted bFGF should be used in a bone defect model.

196

6. Conclusions.

197

198

Conclusions

1. All the combinations of variables tested for the in vitro refolding of the rhBMP-6 produced in

Escherichia coli resulted in low levels of disulfide formation between the monomeric rhBMP-6

molecules, with most of the established disulfides being unspecific. Thus, in vitro refolding of

monomeric rhBMP-6 must require other conditions than those assayed in the present work.

2. The rhBMP-6 and rhBMP-6-CBD dimers produced with a baculoviral/insect cell expression

system were inactive due to incorrect folding or insufficient glycosylation. Thus, this

expression system seems not to be appropriate for the production of these growth factors.

3. The rh-bFGF and rh-bFGF-CBD molecules produced with a baculoviral/insect cell expression

system resulted to be biologically active, although the majority of them were produced with a

low affinity to heparin.

4. The commercially available rh-bFGF produced in Escherichia coli did not show any specific

affinity to absorbable collagen type I sponges, whereas the rh-bFGF molecules produced in

insect cells specifically bound to this material. Furthermore, the addition of the CBD to the

rh-bFGF did not alter its native structure or its affinity to heparin, but enhanced the affinity of

the molecule to absorbable collagen type I sponges.

5. The in vitro biological activity of the bFGFs produced in insect cells was comparable to that

of the commercially available rh-bFGF produced in Escherichia coli when used at low

concentrations. At higher concentrations, the biological activity of the bFGFs produced in

insect cells was slightly lower than that of the commercially available rh-bFGF produced in

Escherichia coli.

6. The combination of rhBMP-6 and rh-bFGF enhanced heterotopic osteogenesis in vivo when

compared to rhBMP-6 alone. The rh-bFGF produced in insect cells was at least as effective as

the commercially available rh-bFGF produced in Escherichia coli, while the use of rh-bFGF-

CBD at the same concentration resulted in the formation of an apparently more immature

bone. This was probably because of an increased retention of the growth factor at the

implant site due to the enhanced affinity to the carrier conferred by the CBD.

199

200

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201

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Appendix III. Abstract in Spanish.

Apéndice III.

Resumen en español.

221

222

Appendix III. Abstract in Spanish / Resumen en español

1. Introducción.

1.1. El tejido óseo y su regeneración tras una fractura.

El tejido óseo es un tipo especializado de tejido conectivo cuyo constituyente mayoritario es

matriz extracelular. La parte orgánica de esta matriz, denominada osteoide, está formada

principalmente por colágeno tipo I, aunque también es posible encontrar otros tipos de

colágeno y proteínas no colagénicas como proteoglicanos, osteonectina, osteopontina,

sialoproteínas, osteocalcina y determinados factores de crecimiento. La parte inorgánica está

constituida por fosfato cálcico en forma de hidroxiapatita, la cual se encuentra depositada sobre

la fase proteica.

Las células osteoprogenitoras derivan de células tronco mesenquimales que se encuentran

en la médula ósea. Estos osteoprogenitores pueden dar lugar a osteoblastos en respuesta a los

estímulos adecuados. Los osteoblastos, que mantienen la capacidad de dividirse, son las células

que secretan los componentes de la matriz osteoide, al igual que vesículas matriciales con altos

contenidos en fosfatasa alcalina (ALP), el enzima responsable de la mineralización de la matriz

por depósito de hidroxiapatita. Una vez mineralizado el osteoide que rodea a un osteoblasto,

éste se convierte en un osteocito, el cual posee una limitada capacidad de formación y

reabsorción de matriz extracelular, y está implicado en fenómenos de mecanotransducción,

respondiendo a estímulos mecánicos ejercidos sobre el hueso. El tercer tipo celular presente en

el hueso es el osteoclasto, que deriva de progenitores hematopoyéticos mononucleados de la

médula ósea. Estas células son las principales encargadas de reabsorber la matriz ósea,

contribuyendo así a la homeostasis del sistema esquelético.

Cuando un hueso resulta fracturado se desencadena una compleja cascada de eventos

biológicos que, en la mayoría de los casos, culmina con la completa recuperación del hueso,

tanto a nivel estructural como funcional, tras un periodo de seis a doce semanas. En este

proceso intervienen tanto factores mecánicos como de señalización intra- y extracelular para

promover la osteoinducción y la osteoconducción. La regulación de esta compleja cadena de

eventos es llevada a cabo por una gran cantidad de factores locales y sistémicos, tales como

factores de crecimiento y de diferenciación, hormonas y citoquinas.

Las moléculas que promueven la osteogénesis durante la reparación de una fractura ósea

pueden dividirse en tres grupos principales:

- Citoquinas pro-inflamatorias, tales como el factor de necrosis tumoral alfa (TNF-α) o

las interleucinas 1 y 6 (IL-1 e IL-6), que ejercen quimiotaxis sobre células

fibrogénicas endógenas y células de la respuesta inflamatoria, promueven la síntesis

de matriz extracelular y estimulan la angiogénesis.

223

Appendix III. Abstract in Spanish / Resumen en español____________________________________________________

- Factores de crecimiento y diferenciación, tales como el factor de crecimiento

transformante beta (TGF-β), proteínas morfogenéticas de hueso (BMPs), factores de

crecimiento fibroblástico (FGFs), el factor de crecimiento derivado de plaquetas

(PDGF) o los factores de crecimiento tipo insulina (IGFs).

- Metaloproteinasas y factores angiogénicos. De éstos, las metaloproteinasas son las

encargadas de degradar el cartílago y el hueso para permitir la entrada de nuevos

vasos sanguíneos, mientras que los factores angiogénicos (factor de crecimiento de

endotelio vascular o VEGF y las angiopoyetinas) promueven la formación, crecimiento

y ramificación de dichos vasos.

Obviamente, para conseguir la restauración de la forma y función original del hueso dañado

no sólo se requiere la acción de moléculas osteogénicas, sino también la modulación por parte

de moléculas inhibidoras, las cuales pueden ser clasificadas en dos grupos:

- Inhibidores de BMPs, tales como nogina, gremlin, cordina, esclerostina, folistatina y el

inhibidor de BMP y activina unido a membrana (BAMBI), que antagonizan a las BMPs

de una u otra manera a nivel extracelular. Otros inhibidores de BMPs actúan a nivel

intracelular, interfiriendo con la cascada de señalización que se desencadena cuando

las BMPs se unen a sus receptores en las células diana.

- Otras moléculas inhibidoras, tales como la interleucina 1 alfa (IL-1α) o las proteínas

de unión a IGF (IGFBPs).

Ciertos estudios apuntan a que varios de estos factores pueden tener un comportamiento

dual durante la reparación de fracturas. Así, se ha descrito que el TGF-β puede bloquear la

diferenciación osteoblástica inducida por BMP-2, mientras que los FGFs podrían estimular la

diferenciación temprana de precursores osteogénicos, pero inhibir los procesos de

diferenciación tardía y la mineralización.

1.2. Aspectos clínicos y económicos de la reparación de fracturas óseas.

La reparación de defectos óseos tiene enormes implicaciones a nivel socio-económico. Se

estima que, sólo en la Unión Europea, el coste derivado del tratamiento de fracturas de cadera

(el tipo de fractura de mayor incidencia) supera los 598 millones de euros anuales.

Si bien el tiempo esperado de reparación de forma natural de una fractura oscila entre seis

y doce semanas, hay una alta incidencia de reparaciones retardadas o de fracturas que no

llegan a soldar. Estos casos no sólo suponen una considerable disminución de la calidad de vida

de los pacientes, sino que también incrementan enormemente el coste económico a los

sistemas sanitarios.

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Mientras que ciertos casos menos severos pueden ser tratados mediante el uso de

estabilizadores mecánicos, los casos más graves suelen requerir tratamientos más sofisticados,

tales como distractores óseos, injertos óseos o la aplicación de biomateriales.

Actualmente, el injerto óseo, mediante el cual el hueso que falta es reemplazado por

material extraído del propio paciente o de donantes, es considerado el tratamiento más eficaz

para la reparación de fracturas que no sueldan. Sin embargo, la escasa cantidad de tejido que

puede ser extraído del donante y la alta morbilidad asociada a la extracción convierten en

patente la necesidad de desarrollar nuevas estrategias alternativas.

Recientemente, la comunidad médica ha centrado su atención en el uso de biomateriales

(materiales compatibles con células y tejidos vivos) naturales o sintéticos para la reparación del

tejido óseo, pretendiendo que estos materiales simulen la acción osteoconductora de los

injertos óseos. Para, además, conferirle a estos materiales propiedades osteoinductoras, se está

estudiando extensamente el uso de éstos en combinación con factores de crecimiento

osteogénicos.

1.3. Las proteínas morfogenéticas de hueso (BMPs).

Las proteínas morfogenéticas de hueso (BMPs) constituyen una familia de más de 20

factores de crecimiento, perteneciente a la superfamilia del TGF- β. Estas proteínas presentan

una estructura altamente conservada, siendo todas ellas producidas como grandes pre-pro-

proteínas monoméricas. El dominio maduro de estas proteínas contiene siete residuos

conservados de cisteína, los cuales determinan la formación de un motivo estructural

característico de estos factores: el nudo cisteínico. Este nudo constituye el núcleo del

monómero y se debe a la formación de tres puentes disulfuro intracatenarios que involucran a

seis de las cisteínas. El séptimo residuo de cisteína interviene en la formación de un único

puente disulfuro intercatenario, lo cual permite la dimerización de la molécula una vez

escindidos los dominios maduros de las pre-pro-proteínas precursoras. Se ha demostrado que la

forma dimérica de la molécula es la única que posee actividad biológica.

Si bien las BMPs poseen varios sitios susceptibles de ser N-glucosilados, en la mayoría de

los casos (BMP-2, BMP-7, BMP-7, etc), sólo uno de estos sitios se encuentra glucosilado.

Aunque esta glucosilación no parece ser esencial para la actividad biológica de BMP-2,

investigaciones muy recientes le otorgan mayor importancia a la glucosilación de la BMP-6.

La señalización por BMPs se debe a la unión de estos factores a dos receptores

serín/treonín quinasas distintas (tipo I y tipo II). El ligando se une primero a dos copias de su

receptor de alta afinidad, tras lo cual se asocian al complejo dos receptores de baja afinidad,

constituyéndose así un complejo de seis cadenas polipeptídicas. En este punto, los receptores

tipo II, que son constitutivamente activos, fosforilan a los receptores tipo I, lo cual puede

conllevar a la activación de dos vías de señalización intracelular distintas: una vía “canónica”

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mediada por proteínas Smads, o una vía “no canónica” mediada por BMP-MAPK. Cuál de estas

vías resulta activada parece depender de los mecanismos particulares de oligomerización del

complejo BMPR.

Las BMPs son factores de crecimiento pleiotrópicos que ejercen multitud de funciones

distintas durante el desarrollo embrionario y en la vida adulta.

En el desarrollo, las BMPs están implicadas en el establecimiento del eje dorsoventral

durante las primeras fases de la ontogénesis. Igualmente, participan en la organización del

mesodermo ventral y en el desarrollo de casi todos los tejidos y órganos, controlando procesos

de proliferación, diferenciación, migración, apoptosis y adhesión celular.

En el organismo adulto, las BMPs parecen estar implicadas en la regeneración de una gran

variedad de tejidos y en la protección y recuperación tras un daño tisular. De todas estas

funciones, la más estudiada es su participación en la regeneración del sistema esquelético,

siendo algunas de estas proteínas las únicas descritas con capacidad de inducir la formación de

hueso ectópico en vertebrados adultos. Se ha demostrado que la señalización por BMPs es

necesaria durante todas las fases de la diferenciación osteoblástica, incluyendo la proliferación y

la formación, maduración y mineralización de la matriz. Sin embargo, se ha visto que las

distintas BMPs actúan en distintas etapas de este proceso, siendo BMP-2, -6 y -9 las que

poseen una mayor capacidad de inducir la expresión de marcadores osteogénicos, tanto

tempranos como tardíos, y la mineralización de la matriz.

La BMP-6 es principalmente expresada en condrocitos hipertróficos durante el proceso de

osificación endocondral y se ha demostrado su capacidad de estimular la expresión de fenotipos

condrogénicos y osteogénicos in vitro, y de inducir la formación de cartílago y hueso in vivo.

1.4. Los factores de crecimiento fibroblástico (FGFs).

Los factores de crecimiento fibroblástico (FGFs) constituyen una familia de 22 miembros de

proteínas de gran importancia en procesos de proliferación, diferenciación, migración y

supervivencia de una gran variedad de tipos celulares durante el desarrollo embrionario,

mientras que actúan como factores homeostáticos durante la reparación tisular y las respuestas

ante daños en los organismos adultos. De entre todos estos factores, los más estudiados son el

FGF básico (bFGF) y el FGF ácido (aFGF), que tienen especial relevancia en los procesos de

cicatrización y de formación de vasos sanguíneos, teniendo un potencial angiogénico mayor que

el del VEGF.

El bFGF presenta 5 isoformas debido a procesos de traducción alternativa del ARN

mensajero, aunque la isoforma de menor masa molecular (18 KDa) es la única que no es

dirigida al núcleo celular tras su síntesis y es la más estudiada. Esta proteína es un monómero

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que presenta 12 láminas beta en su estructura terciaria, sin puentes disulfuro ni glucosilaciones.

En el extremo carboxilo presenta una región rica en lisinas, responsable de la afinidad de esta

molécula por proteoglucanos heparán sulfatados (HSPGs).

Al contrario que la mayoría de proteínas cuyo destino sea ser secretadas al espacio

extracelular, el bFGF no posee ninguna secuencia señal que la dirija al retículo endoplásmico.

Por lo tanto, en vez de ser secretado por una vía vesicular, se cree que el bFGF es exportado al

exterior de la célula por translocación directa a través de la membrana plasmática, en un

proceso independiente de ATP. En este fenómeno parecen estar involucrados los HSPGs de la

superficie celular, los cuales actuarían a modo de “trampas moleculares”, dirigiendo el

transporte unidireccional del bFGF hacia el ambiente extracelular.

El bFGF se une a su receptor FGFR2-IIIc en las células diana. La unión del bFGF a las

moléculas de HSPGs de la membrana celular parece aumentar la afinidad del ligando por su

receptor, de forma que la magnitud y tipo de respuesta celular podría depender de la formación

de complejos ternarios entre el bFGF, los HSPGs y el FGFR. El bFGF también tiene afinidad

específica por heparina y heparán sulfato. La unión del bFGF a estas moléculas protege al factor

frente a la desnaturalización por calor o ácidos, al igual que frente a la acción de proteasas,

manteniendo un reservorio extracelular de bFGF activo.

La señalización intracelular por bFGF presenta dos vías principales: una vía dependiente de

MAPK y una vía dependiente de PKC. El tipo específico de transducción de señal que se

desencadena puede depender del tipo de FGFR que sea activado, al igual que del tipo de

HSPGs de la superficie celular involucrado y de la acción directa de bFGFs intracelulares e

intranucleares. Debido a esta complejidad, la activación de varios tipos celular por bFGF puede

desencadenar una gran variedad de respuestas celulares distintas, tales como proliferación,

migración y/o el estímulo/inhibición de la expresión de un determinado fenotipo.

Una de las actividades mejor caracterizadas de este factor de crecimiento es la de regular el

crecimiento y la función de células vasculares, tales como células endoteliales y musculares

lisas. Durante la regeneración del tejido óseo, el bFGF podría desempeñar un doble papel, ya

que su actividad angiogénica puede estimular la neovascularización del hueso neoformado,

mientras que también podría ser importante para inducir la proliferación y/o diferenciación de

células osteoprogenitoras mesenquimales. Además, la invasión del hueso en formación por

nuevos vasos sanguíneos se considera una fuente de potenciales células osteoprogenitoras.

En lo que respecta a la acción directa del bFGF sobre las propias células osteogénicas hay

una gran controversia. Muchos estudios in vitro han descrito efectos inhibitorios del bFGF sobre

osteoblastos, al igual que varios estudios in vivo han mostrado una disminución de la cantidad

de hueso formado al aplicar altas concentraciones de bFGF. Sin embargo, otros estudios han

encontrado efectos positivos del bFGF sobre la diferenciación de células osteoprogenitoras in

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vitro, y se han descrito tasas mayores de formación de hueso in vivo al implantar dosis bajas de

este factor. Probablemente, estos efectos contradictorios simplemente reflejen la complejidad

de la señalización por bFGF, siendo el efecto de este factor altamente impredecible y

dependiente de la concentración empleada, del estadio de diferenciación de las células diana y

de la influencia de otros factores de crecimiento. Sin embargo, in vivo, se ha demostrado en

numerosas ocasiones el efecto bifásico del bFGF, estimulando la formación neta de hueso a

bajas concentraciones e inhibiéndola a altas concentraciones.

1.5. Terapias para la reparación de fracturas óseas.

Para el tratamiento de fracturas que no consolidan por sí mismas, los injertos óseos son

considerados la mejor opción, aunque las limitaciones que presentan, comentadas

anteriormente, han obligado a la búsqueda de terapias alternativas. La matriz ósea

desmineralizada (DBM) es una sustancia comercializada aunque, cuando se emplea sola, no

posee la misma eficacia que los injertos de hueso autólogos. Por otra parte, los injertos

sintéticos, como los de fosfato cálcico, sulfato cálcico, hidroxiapatita o composites de colágeno

y calcio, pueden simular las propiedades osteoconductoras de los injertos óseos, pero no

poseen capacidad osteoinductora.

Gracias a la tecnología del ADN recombinante, muchos factores de crecimiento relacionados

con la osteogénesis, angiogénesis y cicatrización han sido comercializados y su uso para

aplicaciones clínicas está siendo extensamente estudiado. Combinaciones de estos factores con

biomateriales están siendo considerados alternativas muy prometedoras a los autoinjertos.

Actualmente, las Agencias del Medicamento Europea y Americana (EMEA y FDA,

respectivamente) aprueban el uso de BMP-2 en combinación con esponjas absorbibles de

colágeno (ACS) para el tratamiento de fracturas severas de tibia en humanos adultos.

Igualmente, la FDA aprueba el empleo de BMP-2 en combinación con ACS para las fusiones

espinales. Por su parte, en los casos de fracturas recalcitrantes de huesos largos, se ha

aprobado el uso compasivo de BMP-7.

Por otro lado, es bien conocida la importancia de la angiogénesis durante la reparación del

hueso y numerosos estudios han demostrado que la co-administración de BMP-2 con bajas

concentraciones de bFGF mejora este proceso.

En la clínica, la BMP-2 es usada a concentraciones de 1.5 mg por mililitro de ACS, lo que

supone un incremento a nivel local de más de seis órdenes de magnitud sobre los niveles

fisiológicos. Si bien el tratamiento con BMP-2 o BMP-7 se considera seguro, los efectos a largo

plazo de la aplicación de estas cantidades de unos factores tan altamente pleiotrópicos aún se

desconocen. Además, parece que estos tratamientos estimulan al sistema inmune, pudiéndose

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detectar anticuerpos anti-BMP en un cierto porcentaje de los pacientes. Otra desventaja del uso

de estas grandes cantidades de BMPs es el astronómico coste económico de los tratamientos,

habiéndose estimado que, en el Reino Unido, el uso de BMPs para el tratamiento de fracturas

abiertas de tibia supone un gasto anual adicional de 3.5 millones de libras esterlinas sobre el

gasto de los tratamientos estándar.

Una forma de disminuir la cantidad de factor necesaria para un tratamiento y de

incrementar su eficacia es la aplicación de estas proteínas en combinación con un transportador

osteoconductor que las retenga en el lugar del implante, manteniendo sus concentraciones

locales. Desafortunadamente, ningún transportador disponible puede ser considerado ideal.

De entre los posibles transportadores orgánicos, la DBM ha demostrado tener excelentes

propiedades de retención/liberación de BMPs, pero falla como transportador útil al contener

numerosos factores de crecimiento que no son eliminados durante el proceso de

desmineralización. Esto hace que los efectos de la implantación de DBM sean altamente

impredecibles. Por el contrario, a pesar de sus pobres propiedades biomecánicas, el colágeno es

el único transportador aprobado para la aplicación clínica de BMPs debido a su alta

biocompatibilidad y biodegradabilidad y a su baja inmunogenicidad.

Por desgracia, la mayoría de los factores de crecimiento tienen una baja afinidad natural

por el colágeno. Para remediar esto, muchas citoquinas han sido producidas en el laboratorio

con dominios adicionales que les confieren una mayor afinidad por el colágeno. Uno de estos

dominios es el decapéptido de unión al colágeno/gelatina (CBD) del factor de von Willebrand

(vWF), que ya ha sido empleado para producir una proteína de fusión bFGF-CBD en un sistema

de expresión procariótico, al igual que proteínas de fusión con distintos miembros de la

superfamilia del TGF- β, incluyendo BMPs. Tanto en el caso del bFGF como de la BMP-2, el CBD

fue añadido al extremo N-terminal del factor y la cisteína presente en la secuencia del CBD fue

reemplazada por una metionina para impedir la formación de puentes disulfuro inespecíficos

durante los procesos de producción de las proteínas. En el caso del bFGF, la proteína fue

además producida con un domino de purificación consistente en una secuencia hexahistidínica y

un sitio de corte con trombina. Esto ya fue evitado para la producción de la BMP-2-CBD, la cual

fue purificada gracias a su afinidad natural por heparina.

Al ser el colágeno no sólo el único transportador aprobado por la FDA y la EMEA para

aplicaciones clínicas para la reparación ósea, sino también uno de los componentes principales

del hueso, los factores recombinantes con afinidad añadida por colágeno son considerados de

gran interés. Tras su administración en forma soluble, estas moléculas podrían ser usadas para

aumentar su concentración local al unirse directamente al colágeno propio del tejido en el lugar

del implante. Por otro lado, al ser administradas en combinación con un transportador

colagénico, este último podría retener parcialmente a los factores, limitando su acción al lugar

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del implante. Por tanto, estas estrategias podrían reducir la cantidad de factores necesarios

para conseguir la regeneración tisular, aumentando la seguridad de estos tratamientos y

disminuyendo su coste.

2. Objetivos.

Los objetivos concretos que se plantearon para el presente trabajo fueron:

1. Obtener los genes que codifican para los factores bFGF y BMP-6 humanos, añadir a estas

secuencias la que codifica para el dominio de afinidad por el colágeno derivado del factor

de von Willbrand y clonarlas en vectores de expresión.

2. Expresar de manera heteróloga y purificar la rhBMP-6 y el rh-bFGF tanto en forma nativa

como con el dominio de afinidad por el colágeno.

3. Determinar la afinidad por el colágeno de los factores producidos con el domino adicional

de afinidad por el colágeno.

4. Caracterizar la actividad osteogénica de los factores dirigidos a colágeno in vitro e in vivo y

compararla con la de los factores nativos.

5. Comparar la actividad osteogénica in vivo de combinaciones de bFGF y BMP-6 con BMP-6

solo.

3. Material y métodos.

3.1. Producción de rhBMP-6 y rh-BMP-6-CBD en un sistema de expresión en

Escherichia coli.

En primer lugar, se trataron de producir las proteínas rhBMP-6 y rhBMP-6-CBD en un

sistema de expresión en Escherichia coli, el cual rinde las proteínas heterólogas en forma de

cuerpos de inclusión insolubles que, posteriormente, han de ser solubilizados para obtener las

moléculas monoméricas solubles. Finalmente, estos monómeros han de ser replegados in vitro

para conseguir los dímeros activos, que deberán ser purificados para separarlos de los

monómeros remanentes y otros posibles contaminantes.

El gen codificante para la BMP-6 humana se obtuvo mediante RT-PCR sobre muestras de

ARN total de células de osteosarcoma humano U-2 OS y la secuencia obtenida fue clonada en

un vector de mantenimiento (pBIISK). Mediante PCR se generaron las secuencias necesarias

para obtener los genes de la BMP-6 y la BMP-6-CBD con los sitios de restricción apropiados

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para poder ser clonados en el plásmido pET17b. Las construcciones obtenidas se usaron para

transformar la cepa de E. coli Rosetta™ (DE3).

Se seleccionó un clon para cada una de las proteínas a expresar y se analizó el plásmido

que contenían para comprobar la correcta inserción de la secuencia heteróloga y descartar

posibles mutaciones.

El clon seleccionado para la expresión de rhBMP-6 fue crecido en cultivo líquido e inducido

a expresar la proteína por adición de IPTG. Cuatro horas tras la adición del IPTG se analizó la

producción mediante SDS-PAGE y tinción con azul de Coomassie, y se aislaron los cuerpos de

inclusión generados, los cuales fueron solubilizados con 6 M guanidina hidrocloruro. Los

monómeros de rhBMP-6 fueron finalemente cuantificados por SDS-PAGE, tinción con azul de

Coomassie y análisis de imagen digital y densitometría.

Con los monómeros así obtenidos, se probaron 41 condiciones de replegamiento in vitro,

empleando combinaciones de distintas variables: concentración de proteínas, temperatura, pH,

tipo y concentración de antiagregantes, tipo y concentración de par redox, y degaseado o no

del tampón de replegamiento. El resultado de cada uno de estos intentos de replegamiento fue

analizado mediante SDS-PAGE y tinción con azul de Coomassie.

3.2. Producción de rhBMP-6 y rh-BMP-6-CBD en un sistema de expresión en células

de insecto/baculovirus.

Para la producción de rhBMP-6 y rhBMP-6-CBD en un sistema de expresión en células de

insecto, las secuencias de ambos genes fueron clonadas en un vector donador de un sistema

basado en baculovirus (pACGP67B). A continuación se cotransfectaron células de insecto Sf9

con las construcciones obtenidas y ADN baculoviral Sapphire™ linealizado. En el interior de la

células, la existencia de secuencias virales en el plásmido donador, flanqueando el sitio de

restricción múltiple, permite su recombinación homóloga con el ADN vírico, generándose una

partícula vírica infectiva que porta el gen heterólogo bajo el control del promotor de la

polihedrina, el cual se activa intensamente durante las últimas fases del ciclo infectivo de los

baculovirus. Los baculovirus Sapphire™ coexpresan, junto con la proteína heteróloga, la

proteína isomerasa de puentes disulfuro (PDI), que es una chaperona que contribuye a la

correcta formación de puentes disulfuro en las proteínas. Además el uso del pACGP67B implica

que las proteínas producidas portan el péptido señal GP67, que dirige a las proteínas hacia el

retículo endoplásmico y aparato de Golgi para su secreción al medio de cultivo por vía vesicular.

Antes de la secreción, el péptido señal es escindido de la proteína por endopeptidasas propias

del virus.

Los sobrenadantes de transfección se emplearon para aislar clones individuales de

baculovirus recombinantes mediante un ensayo de placas de lisis en cultivos de Sf9 cubiertos

con agarosa y los clones seleccionados fueron analizados mediante PCR. Un clon para cada

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proteína a expresar fue amplificado y la suspensión vírica obtenida fue titulada mediante un

ensayo de infección por dilución limitante. Una vez conocido el título de ambas suspensiones,

los virus fueron nuevamente amplificados usando una multiplicidad de infección (MOI; es decir,

el número de partículas víricas por célula empleado para realizar la infección) de 0.1. Se

obtuvieron así suspensiones de virus de gran volumen, que fueron igualmente titulados. Estas

suspensiones permitieron realizar todas las producciones posteriores de forma reproducible y

con un MOI conocido.

Para determinar las condiciones óptimas de tiempo de producción y MOI para cada

proteína, se realizaron ensayos de producción, infectando cultivos de Sf9 con valores de MOI de

2.5 y 10 y recogiendo los medios de cultivo condicionados tras 48, 72, 96 y 120 horas de

tiempo de producción. Estas muestras fueron analizadas mediante Western blot con un

anticuerpo específico anti-BMP-6.

Una vez establecidas las condiciones óptimas para cada proteína, se realizaron

producciones de rhBMP-6 y rhBMP-6-CBD. Para su purificación, se añadió 6 M urea a los medios

condicionados y éstos se cargaron en columnas de heparín-sefarosa. Las proteínas retenidas en

la columna se eluyeron en dos pasos, con tampones con concentraciones incrementadas de

NaCl y las fracciones de elución se analizaron mediante Western blot.

Las fracciones de elución con mayor contenido en dímeros de rhBMP-6 o rhBMP-6-CBD

fueron tratadas para eliminar el exceso de urea y NaCl. Para ello se probaron los siguientes

métodos:

- Diálisis frente a medio DMEM, pH 7.0 ó pH 4.9.

- Diálisis frente a 4 mM HCl.

- Diálisis frente a 10 mM acetato amónico, pH 4.0, liofilización y resuspensión en agua,

agua con 0.1 % BSA, 4 mM HCl ó 4 mM HCl con 0.1% BSA.

- Cambio de tampón a 4 mM HCl en columnas Vivaspin 2.

La actividad biológica de todas las muestras fue probada mediante un ensayo de inducción

de expresión de fosfatasa alcalina (ALP) en mioblastos de ratón C2C12.

3.3. Producción de rh-bFGF y rh-bFGF-CBD en un sistema de expresión en células de

insecto/baculovirus.

Los genes codificantes para el bFGF y el bFGF-CBD fueron clonados en el plásmido donador

del sistema de expresión baculoviral pACGP67B y las construcciones obtenidas se usaron para

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cotransfectar células Sf9 junto con ADN de baculovirus BacPak6™. Los virus BacPak6™ no

coexpresan la PDI, ya que el bFGF no presenta puentes disulfuro en su estructura nativa.

Al igual que para el caso de la rhBMP-6 y la rhBMP-6-CBD, se aislaron clones productores

de rh-bFGF y rh-bFGF-CBD mediante ensayos de placas de lisis, se analizaron los clones

mediante PCR, y se amplificaron y titularon los virus seleccionados. Igualmente, se

determinaron las condiciones óptimas de producción mediante ensayos de producción y análisis

por Western blot con un anticuerpo específico anti-bFGF.

Una vez producidas las proteínas, se purificaron en columnas de heparín-sefarosa en

ausencia de urea y se eluyeron las proteínas retenidas en la columna mediante un gradiente de

NaCl. Las fracciones de elución fueron analizadas mediante dot blot y las muestras de rh-bFGF

y rh-bFGF-CBD obtenidas se cargaron en columnas de filtración Vivaspin 2 para sustituir el

tampón de elución por PBS, pH 7.2 con 1 mM DTT y 1 mM EDTA.

Para determinar la afinidad de las proteínas producidas por el colágeno tipo I se cargaron

1.25 pmoles de cada una de ellas y de bFGF comercial en esponjas de colágeno tipo I de 1 mm

de grosor y 5 mm de diámetro. Las esponjas fueron lavadas con PBS + tween-20 durante una

hora o 6 días y la cantidad de proteínas retenidas cuantificadas mediante inmunotinción con un

anticuerpo anti-bFGF y análisis densitométrico digital.

La actividad biológica de las proteínas obtenidas se probó in vitro mediante un ensayo de

proliferación sobre preosteoblastos de ratón MC3T3-E1, comparando el número de células de

los cultivos tras 72 horas de incubación con los factores con cultivos incubados con bFGF

comercial o sin factor. Para ello se empleó un método colorimétrico basado en MTT.

Igualmente, se determinó la capacidad de los factores producidos de inhibir la

diferenciación osteoblástica de las células MC3T3-E1 en presencia de ácido ascórbico. Para ello

se determinó el número de células en los cultivos incubados durante 120 horas con los factores

recombinantes, con bFGF comercial o sin factor. Igualmente, se cuantificó la expresión de ALP

en estos cultivos mediante un método basado en pNPP y se calculó la actividad ALP/célula.

3.4. Determinación de la capacidad osteogénica de combinaciones de rh-bFGF o

rh-bFGF-CBD con rhBMP-6 in vivo.

Para determinar el efecto de las proteínas bFGF producidas en combinación con rhBMP-6 se

realizó un ensayo de osificación ectópica en ratas. Para ello, seis ratas Wistar macho de 4

meses de edad y 250-280 gramos de peso fueron anestesiadas e implantadas con discos de

ACS de 5 mm de diámetro y 1 mm de grosor, a las que se habían añadido los factores a probar.

Los discos fueron implantados en pequeñas incisiones realizadas en los músculos dorsales de

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los animales, cerrándose posteriormente las heridas con puntos de sutura y la piel dorsal con

grapas quirúrgicas. Las combinaciones de factores probados fueron:

- C- (discos incubados con PBS)

- 13.89 pmol (0.5 µg) de rhBMP-6 comercial (R&D Systems)

- 1.25 pmol de rh-bFGF comercial (R&D Systems)

- 1.25 pmol de rh-bFGF producido en células Sf9

- 1.25 pmol de rh-bFGF-CBD producido en células Sf9

- 13.89 pmol de rhBMP-6 comercial + 1.25 pmol de rh-bFGF comercial

- 13.89 pmol de rhBMP-6 comercial + 1.25 pmol de rh-bFGF producido en células Sf9

- 13.89 pmol de rhBMP-6 comercial + 1.25 pmol de rh-bFGF-CBD producido en células

Sf9

Al cabo de 21 días, los animales fueron sacrificados y los implantes diseccionados y fijados

en formaldehído tamponado al 4%. Los implantes que portaban rhBMP-6 fueron descalcificados

antes de continuar el proceso de inclusión, mientras que los demás implantes fueron

directamente deshidratados con alcohol de gradación creciente e incluídos en parafina. Se

realizaron secciones de 10 µm de grosor, que fueron montados en portaobjetos para su

posterior análisis histológico mediante las siguientes técnicas:

Tinciones histoquímicas:

- Hematoxilina-eosina. Tinción general para la observación de los tejidos.

- Tricrómico de Masson. Tinción triple que tiñe al osteoide de rojo, y al hueso

mineralizado de verde.

- Azul alciano. Tiñe los mucopolisacáridos ácidos y los glucosaminoglucanos de la

matriz cartilaginosa de azul.

-

Tinciones inmunohistoquímicas:

- Inmunotinción del marcador de osteogénesis osteopontina.

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4. Resultados y Discusión.

4.1. Producción de rhBMP-6 en Escherichia coli.

Tras la obtención y clonación de los genes para la rhBMP-6 y la rhBMP-6-CBD en el vector

de expresión pET17b, las construcciones fueron usadas para transformar la cepa de E. coli

Rosetta™ (DE3). Se seleccionó un clon que contuviera la construcción pET17b:BMP-6 y se

procedió a la producción de la proteína heteróloga en cultivos líquidos. El análisis por SDS-PAGE

reveló que, cuatro horas tras la adición de IPTG al cultivo, las bacterias contenían una gran

cantidad (40-50% del contenido de proteína total) de una proteína cuya masa molecular podría

corresponder a la predicha para los monómeros de BMP-6 (±16.5 KDa). Esta proteína se

encontraba en la fracción de proteínas insolubles en forma de cuerpos de inclusión y pudo ser

aislada con alta pureza tras lavar esta fracción con un tampón con triton x-100.

Los cuerpos de inclusión fueron solubilizados con 6 M guanidina hidrocloruro y la rhBMP-6

monomérica soluble cuantificada por análisis digital de imagen sobre geles de poliacrilamida

teñidas con azul de Coomassie, estimándose un rendimiento de aproximadamente 108 mg/L.

Se probaron 41 condiciones de replegamiento in vitro de la rhBMP-6, obteniéndose en casi

todos los casos resultados similares: la mayoría de la rhBMP-6 permanecía en su forma

monomérica; sólo una pequeña fracción de las proteínas pasaba a formar parte de

organizaciones oligoméricas, constituyendo dímeros, trímeros, tetrámeros o polímeros

superiores. Estas agregaciones se debían a la formación de puentes disulfuro intercatenarios

inespecíficos ya que el análisis por SDS-PAGE de estas muestras en presencia de un agente

reductor mostraba la reversión total del proceso de oligomerización.

El hecho de que se formaran gran cantidad de puentes disulfuro inespecíficos nos hizo

suponer que la fracción dimérica probablemente estaría constituida de una mezcla heterogénea

de proteínas correcta e incorrectamente plegadas. Esto, junto con el hecho de que en todos los

casos esta fracción dimérica era muy minoritaria, nos hizo considerar inviable su purificación ya

que nuestra experiencia previa con proteínas de la misma familia expresadas con este mismo

sistema nos había indicado que un alto porcentaje de proteínas se pierden durante la

purificación y posterior manipulación.

4.2. Producción de rhBMP-6 y rhBMP-6-CBD en células Sf9.

Al haber resultado el replegamiento de rhBMP-6 producido en Escherichia coli infructuoso,

decidimos abordar la producción de estos factores de crecimiento en un sistema de expresión

eucariótico, en el que las proteínas son plegadas en el interior de la célula para obtener su

estructura cuaternaria nativa.

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Una vez que los genes codificantes para el rhBMP-6 y el rhBMP-6-CBD fueron

correctamente clonados en el plásmido pAcGP67B se cotransfectaron células Sf9 con el

plásmido donador recombinante y ADN baculoviral Sapphire™ linealizado para obtener

baculovirus recombinantes capaces de expresar las proteínas heterólogas. Los sobrenadantes

de transfección fueron empleados para realizar un ensayo de placas de lisis mediante el cual se

aislaron clones de baculovirus recombinantes. Estos clones fueron amplificados y titulados como

descrito anteriormente.

Los ensayos de producción mostraron que, en ambos casos, las proteínas heterólogas eran

secretadas al medio de cultivo en forma de dímeros, trímeros y tetrámeros, no detectándose

moléculas monoméricas. Estos oligómeros se encontraban estabilizados por puentes disulfuro

ya que el análisis de las muestras por Western blot en condiciones reductoras mostró una

reversión total de la oligomerización.

Para determinar si la excesiva formación de puentes disulfuro se debía a la coexpresión de

la PDI, generamos un baculovirus BacPak6™ recombinante con el fin de expresar la rhBMP-6

sin esta chaperona. El análisis por Western blot de la producción con este baculovirus mostró

que la PDI no era responsable de la formación de puentes disulfuro inespecíficos, ya que el

patrón de oligómeros obtenido con este virus era igual que con el anterio.

Los dímeros de rhBMP-6(-CBD) fueron purificados mediante cromatografía de afinidad a

heparina. Sin embargo, no nos fue posible separar completamente la fracción dimérica de la

trimérica mediante FPLC. Las fracciones enriquecidas en dímeros fueron tratados como se

describe en “material y métodos” para eliminar el exceso de urea y NaCl. Ninguna de las

muestras así obtenidas resultó tener actividad biológica en un ensayo de inducción de actividad

ALP sobre mioblastos de ratón C2C12.

Hay distintas posibles causas que, solas o en combinación, podrían explicar la ausencia de

actividad biológica de las proteínas heterólogas expresadas en células Sf9:

- La elección del promotor de la polihedrina para controlar la expresión de las proteínas

heterólogas. Un alto porcentaje (o todo) de la rhBMP-6(-CBD) dimérica podría estar

incorrectamente plegada debido a saturación y/o desregulación de la maquinaria

celular de producción y procesamiento de proteínas durante las fases tardías del ciclo

infectivo del virus, durante las cuales se activa el promotor de la polihedrina.

- Incluso aunque un cierto porcentaje de los dímeros purificados tuvieran la estructura

cuaternaria correcta, la presencia de altas concentraciones de dímeros incorrectos u

oligómeros superiores podría estar inhibiendo la actividad de las moléculas nativas al

actuar como inhibidores competitivos.

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- Dado que los residuos de ácido siálico terminales juegan un importante papel en

muchos glucoconjugados y la glucosilación de la BMP-6 parece ser fundamental para

su unión a los receptores de BMPs, es posible que la glucosilación llevada a cabo por

las células Sf9 no sea suficiente para obtener moléculas de rhBMP-6 activas.

4.3. Producción de rh-bFGF y rh-bFGF-CBD en células Sf9.

Como la producción de un rh-bFGF-CBD en Escherichia coli ya ha sido publicada y se probó

que esta proteína tenía actividad biológica y afinidad por el colágeno, decidimos producir el rh-

bFGF y rh-bFGF-CBD en un sistema de expresión eucariótico, que recuerda más a las células

humanas productoras de bFGF que E. coli.

Las secuencias codificantes para rh-bFGF y rh-bFGF-CBD fueron obtenidas mediante PCR

sobre las construcciones usadas para su producción en E. coli y clonadas en el plásmido

donador del sistema de baculovirus pAcGP67B. El bFGF nativo es una proteína monomérica que

no presenta ningún puente disulfuro, pero sí varios residuos de cisteína susceptibles de formar

puentes disulfuro inespecíficos bajo condiciones permisivas. Este hecho nos condujo a emplear

un sistema de expresión con baculovirus que no coexpresara la PDI, ya que la presencia de

esta chaperona podría provocar el incorrecto plegamiento de las proteínas.

Una vez obtenidos los baculovirus recombinantes para la expresión de las proteínas

heterólogas mediante cotransfección de células Sf9 con las construcciones de pAcGP67B y ADN

viral linealizado, se aislaron clones infectivos de virus mediante ensayos de placas de lisis. Estos

clones fueron después amplificados y titulados.

Los análisis por Western blot de los ensayos de producción para ambas proteínas revelaron

que el anticuerpo anti-bFGF reconocía una única proteína en los medios de cultivo

condicionados, cuya masa molecular correspondía con la predicha para el rh-bFGF o el rh-bFGF-

CBD, respectivamente. Una vez que se habían determinado las mejores condiciones de MOI y

tiempo de producción se realizaron producciones de gran volumen de las dos proteínas y éstas

fueron purificadas mediante cromatografía de afinidad por heparina. Para eluir las proteínas de

la columna se empleó un gradiente lineal de NaCl.

En ambos casos, esto resultó en la elución de, al menos, dos poblaciones distintas de bFGF:

una mayoritaria, con “baja afinidad por heparina” y una minoritaria, con “alta afinidad por

heparina”. Ambas fracciones con “baja afinidad por heparina” se perdieron durante el proceso

de cambio de tampón en columnas Vivaspin2, mientras que las fracciones con “alta afinidad por

heparina” pudieron ser satisfactoriamente concentradas y recuperadas. En cualquier caso, como

la unión a heparina parece ser importante para la actividad biológica de estos factores,

quisimos centrarnos especialmente en las muestras con mayor afinidad por la heparina.

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Las muestras recuperadas fueron cuantificadas mediante Western blot y análisis de imagen

digital antes de realizar con ellas un ensayo de pegada a colágeno para determinar su afinidad

por el colágeno tipo I. Para ello se cargaron discos de ACS con rh-bFGF o rh-bFGF-CBD y se

lavaron con PBS + tween-20 durante 1 hora ó 6 días. Discos cargados con rh-bFGF comercial,

producido en E. coli, o únicamente con tampón se emplearon como controles. Mientras que

prácticamente todo el rh-bFGF comercial era lavado de las esponjas tras una hora, cantidades

significativas de los rh-bFGF y rh-bFGF-CBD producidos en células Sf9 permanecían retenidas

en ellas. Sin embargo, la cantidad de rh-bFGF-CBD que quedaba retenida era significativamente

mayor que la de rh-bFGF. Además, esta unión al colágeno era estable en el tiempo ya que, tras

seis días de lavado, aún se podían detectar ambas proteínas en las esponjas. Estos resultados

nos indicaron, por tanto, que los factores rh-bFGF y rh-bFGF-CBD producidos en células Sf9 no

sólo poseen una mayor afinidad por ACS comparadas con el rh-bFGF producido en E. coli, sino

que esta unión es perdurable en el tiempo. Además, el CBD le confiere a la molécula una

afinidad por el colágeno tipo I adicional que resulta en un, aproximadamente, un 50% más de

unión. En consecuencia, el empleo de estos factores en combinación con esponjas absorbibles

de colágeno para la reparación clínica de fracturas óseas podría permitir la implantación de

menores concentraciones, reduciendo la dispersión sistémica y, por tanto, aumentando la

eficiencia y seguridad del tratamiento. Por último, la pegada específica de las proteínas al

biomaterial transportador podría evitar la gran pérdida de proteína que ocurre debido a la

manipulación de las esponjas por parte del cirujano antes de su implantación.

Para determinar la actividad biológica de los factores de crecimiento obtenidos se realizó un

ensayo de proliferación sobre la línea celular de preosteoblastos de ratón MC3T3-E1. No se

observaron diferencias entre el efecto de las dos proteínas, ni entre ellas y el factor comercial,

cuando eran usadas a bajas concentraciones (156.25 o 312.50 pM). Sin embargo, a mayores

concentraciones (625.00 pM o 1.25 nM), el rh-bFGF comercial presentaba una actividad ligera,

aunque significativamente, mayor que los factores producidos en células Sf9.

4.4. Formación de hueso heterotópico in vivo con rh-bFGF and rhBMP-6.

Para evaluar la actividad biológica del rh-bFGF y rh-bFGF-CBD producidos en células Sf9 y

su capacidad para aumentar la formación de hueso en combinación con rhBMP-6, se prepararon

discos de ACS con 13.89 pmoles de rhBMP-6, 1.25 pmoles de rh-bFGF o rh-bFGF-CBD o

combinaciones de estos factores, que fueron implantados en los músculos dorsales de ratas

Wistar jóvenes. Como control positivo se empleó el rh-bFGF comercial producido en E. coli y,

como control negativo, esponjas cargadas únicamente con tampón.

21 días tras la cirugía, todos los implantes que habían sido cargados con factores de

crecimiento pudieron ser recuperados sin que se percibieran signos de rechazo inmune ni

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encapsulación fibrótica. Por el contrario, ninguno de los implantes cargados con sólo tampón

pudo ser recuperado debido a la reabsorción del colágeno. Los implantes cargados con rhBMP-6

(sola o en combinación con cualquier bFGF) mostraban unos bordes bien definidos y

consistencia dura, aunque variaban en tamaño. En contraposición, la mayoría de los implantes

que fueron cargados con rh-bFGF comercial o rh-bFGF o rh-bFGF-CBD producidos en células de

insecto fueron difíciles de localizar en el músculo y solían encontrarse como partículas de menos

consistencia, en ocasiones dispersas.

El análisis histológico de los implantes mostró que, aquellos que sólo habían sido cargados

con bFGF, habían formado un tejido fibroso más o menos denso, sin mostrar indicios de

osteogenésis ni angiogénesis. Por el contrario, todas las esponjas cargadas con BMP-6 sola o

con bFGF habían formado tejido óseo.

Los implantes cargados con sólo BMP-6 mostraban trabéculas óseas fundamentalmente en

la superficie, estando la zona central llena de densas acumulaciones de células mesenquimales

indiferenciadas. Aunque era posible encontrar algunos vasos sanguíneos en estos implantes,

éstos eran relativamente pequeños y se encontraban de forma dispersa. El marcador temprano

de osteogénesis osteopontina se localizaba fundamentalmente en las trabéculas y únicamente

en algunas células aisladas en el interior del implante.

Todos los implantes con BMP-6 y bFGF mostraban trabéculas óseas no sólo en la superficie

exterior, sino también por todo el interior. En la mayoría de los casos era posible distinguir un

tejido con apariencia de médula ósea entre las trabéculas, con gran cantidad de lagunas

vasculares o vasos sanguíneos grandes, adipocitos y osteoblastos expresando osteopontina.

En el caso de los implantes cargados con rhBMP-6 y rh-bFGF-CBD se encontraban algunas

zonas de cartílago hipertrófico, al igual que acumulaciones de osteopontina de aspecto fibroso.

Se pudo concluir que el uso de 1.25 pmoles de bFGF puede aumentar la formación de

hueso en combinación con BMP-6 y que el rh-bFGF producido en células Sf9 era, al menos,

igual de efectivo que el rh-bFGF comercial, producido en E. coli. Los implantes con rhBMP-6 y

rh-bFGF-CBD mostraban una apariencia intermedia entre los implantes con rhBMP-6 + rh-bFGF

y los que habían sido cargados con sólo bFGF. Como se ha demostrado que altas

concentraciones de bFGF pueden inhibir la osteogénesis inducida por BMP-2, podría pensarse

que la mayor afinidad por el colágeno del rh-bFGF-CBD supone un aumento de la concentración

efectiva local del factor por encima del límite osteogénico. Si este fuera el caso, podrían usarse

cantidades menores de este factor para alcanzar los mismos resultados que con cantidades

mayores del factor sin modificar. En cualquier caso, el hecho de que el rh-bFGF producido en

células Sf9 tenga una cierta afinidad por el colágeno tipo I ya podría incrementar la seguridad

de los tratamientos con este factor en comparación con el rh-bFGF comercial a las mismas

concentraciones.

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5. Conclusiones.

1. Todas las combinaciones de variables probadas para el plegamiento in vitro de la rhBMP-6

producida en Escherichia coli resultaron en bajos niveles de formación de puentes disulfuro,

siendo la mayoría de las formadas inespecíficas. Por tanto, el plegamiento in vitro de la

rhBMP-6 monomérica debe requerir otras condiciones distintas a las empleadas en el

presente trabajo.

2. Los dímeros de rhBMP-6 y rhBMP-6-CBD producidas con un sistema de expresión en

baculovirus/células de insecto resultaron ser inactivos debido a un plegamiento incorrecto o

a una glucosilación insuficiente. Por tanto, este sistema de expresión no parece ser

apropiado para la expresión heteróloga de estos factores de crecimiento.

3. Las moléculas de rh-bFGF y rh-bFGF-CBD producidas con un sistema de expresión en

baculovirus/células de insecto resultaron poseer actividad biológica, aunque la mayoría de

ellas presentaban una baja afinidad por la heparina.

4. El rh-bFGF comercial producido en Escherichia coli no mostró tener ninguna afinidad

específica por esponjas absorbibles de colágeno tipo I, mientras que las moléculas de

rh-bFGF producidas en células de insecto se pegaban de manera específica a este material.

Además, la adición del CBD al rh-bFGF no alteró su estructura nativa ni su afinidad por la

heparina, pero sí aumentó aún más la afinidad de la molécula por las esponjas absorbibles

de colágeno tipo I.

5. La actividad biológica in vitro de los bFGFs producidos en células de insecto fue comparable

a la de la proteína comercial, producida en Escherichia coli, a bajas concentraciones. A

mayores concentraciones, la actividad biológica de los bFGFs producidos en células de

insecto fue ligeramente inferior a la del factor comercial.

6. La combinación de rhBMP-6 y rh-bFGF incrementó la osteogénesis heterotópica in vivo frente

a la rhBMP-6 en solitario. El rh-bFGF producido en células de insecto fue, al menos, igual de

efectivo que el factor comercial producido en Escherichia coli, mientras que el uso del

rh-bFGF-CBD a la misma concentración resultó en la formación de un hueso de apariencia

más inmadura. Esto probablemente se debió a una mayor retención del factor de crecimiento

en el sitio del implante debido al incremento de su afinidad por el material transportador

que le transfiere el CBD.

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