Modificación génica no viral de células madre ...

Modificación génica no viral de

células madre mesenquimales en un

soporte tridimensional biocompatible

Ana Isabel Ramos Murillo

Universidad Nacional de Colombia sede Bogotá

Facultad de Ingeniería, Departamento de Ingeniería Química y Ambiental

Bogotá, Colombia

2020

Modificación génica no viral de

células madre mesenquimales en un

soporte tridimensional biocompatible


Tesis presentada como requisito parcial para optar al título de:

Doctora en Ingeniería – Ingeniería Química

Director:

Ph.D. Rubén Darío Godoy Silva

Codirector:

Ph.D. Gustavo Andrés Salguero López

Asesores nacionales:

Ph.D. Ingrid Silva Cote y M.Sc. Bernardo Camacho Rodríguez

Línea de Investigación:

Terapia génica e ingeniería de tejidos

Grupos de Investigación:

Grupo de Investigación en Procesos Químicos y Bioquímicos (GPQ&B)

Grupo de Investigación en Medicina Transfusional, Tisular y Celular (GIMTTyC)

Universidad Nacional de Colombia sede Bogotá

Facultad de Ingeniería, Departamento de Ingeniería Química y Ambiental

Bogotá D.C., Colombia

2020

Non-viral gene modification of

mesenchymal stem cells in a

tridimensional biocompatible scaffold


DISSERTATION - Presented in partial fulfillment of the requirements for the degree:

Doctor of Engineering – Chemical Engineering

Supervisor:

Ph.D. Rubén Darío Godoy Silva

Co-supervisor:

Ph.D. Gustavo Andrés Salguero López

External advisors:

Ph.D. Ingrid Silva Cote and M.Sc. Bernardo Camacho Rodríguez

Research area:

Gene therapy and tissue engineering

Research groups:

Research Group on Chemical and Biochemical Processes (GPQ&B)

Research Group in Transfusion, Tissue and Cellular Medicine (GIMTTyC)

Universidad Nacional de Colombia, Bogotá campus

Faculty of Engineering

Department of Chemical and Environmental Engineering

Bogotá D.C., Colombia

2020

A Leonor y Antonio, con amor infinito

Declaración de obra original

Yo declaro lo siguiente:

He leído el Acuerdo 035 de 2003 del Consejo Académico de la Universidad Nacional.

«Reglamento sobre propiedad intelectual» y la Normatividad Nacional relacionada al

respeto de los derechos de autor. Esta disertación representa mi trabajo original, excepto

donde he reconocido las ideas, las palabras, o materiales de otros autores.

Cuando se han presentado ideas o palabras de otros autores en esta disertación, he

realizado su respectivo reconocimiento aplicando correctamente los esquemas de citas y

referencias bibliográficas en el estilo requerido.

He obtenido el permiso del autor o editor para incluir cualquier material con derechos de

autor (por ejemplo, tablas, figuras, instrumentos de encuesta o grandes porciones de texto).

Por último, he sometido esta disertación a la herramienta de integridad académica, definida

por la universidad.

________________________________


Fecha 19/09/2020

Universidad Nacional de Colombia – Departamento de Ingeniería Química y Ambiental

Agradecimientos

Quiero expresar mi profundo agradecimiento a todas las personas y entidades que con

su apoyo incondicional permitieron la realización de esta tesis de doctorado.

Al Dr. Bernardo Camacho Rodríguez, director del Instituto Distrital de Ciencia,

Biotecnología e Innovación en Salud (IDCBIS), gracias infinitas. Gracias por su apoyo

desinteresado, por decir “Ana Isabel, te vamos a ayudar”, y por cumplirlo a rajatabla. Es

indudable que sin su apoyo y su firme decisión de sacar este proyecto adelante no lo

hubiera logrado. De todo corazón, gracias.

Al Dr. Rubén Darío Godoy Silva, director del Grupo de Investigación en Procesos

Químicos y Bioquímicos y director de mi tesis de doctorado. Millones de gracias.

Construimos un grupo de investigación maravilloso que llamamos nuestra gran familia.

Gracias infinitas por la confianza, por el apoyo desinteresado, por la generosidad. Gracias

por esforzarse en ser mejor y hacernos mejores a su paso.

Al Dr. Gustavo Salguero, director de la Unidad de Terapias Avanzadas del IDCBIS y

codirector de mi tesis de doctorado, mil y mil gracias. Gracias por el apoyo desinteresado,

por poner a disposición de este proyecto todo el conocimiento y experticia de su grupo de

investigación. Gracias por ser esa mirada crítica, cautelosa, gracias por darme ánimo en

esos momentos difíciles y estar ahí en los momentos determinantes.

A la Dra. Ingrid Silva Cote, directora del grupo de investigación en Ingeniería de

tejidos del IDCBIS, asesora “ad honorem” de mi tesis y apoyo inimaginable. Infinitas gracias

Agradecimientos XI


por el tiempo, la paciencia, por adoptarme en su grupo de investigación, por animarme

siempre, por alegrarse por mis resultados así fueran pequeños, y por hacer suyos mis

pequeños logros. En este último año su ayuda fue determinante; gracias de todo corazón.

A la Dra. Olga Villamizar del Centro de Terapia Génica del City of Hope National Medical

Center, mil gracias por su ayuda en la etapa final de la tesis, tanto en la preparación de las

publicaciones como en la defensa de la tesis. Olga, gracias infinitas por el tiempo, los

consejos y la asesoría que fueron definitivos en esta última etapa de todo mi proceso de

formación. Fue una enorme bendición toparme contigo en este camino.

A la Dra. Tatiana Segura de la Universidad de Duke, muchas gracias por su tiempo y

su asesoría que me permitieron finalizar la última parte de mi tesis.

A la Dra. Ana María Perdomo Arciniegas, directora del Banco de sangre de cordón

umbilical del IDCBIS por su apoyo constante. A la Dra. Carolina Aldana del Banco de

sangre del IDCBIS por su ayuda con las unidades de plasma humano y por enseñarme todo

lo relacionado con el procesamiento de las unidades de sangre. Mil gracias por la buena

disposición y por toda la ayuda brindada desde los tiempos de mi tesis de maestría.

Al Dr. Carlos Arturo Guerrero Fonseca, director del Laboratorio de Biología Molecular

del Virus de la Facultad de Medicina de la Universidad Nacional sede Bogotá, gracias por

ser mi primera casa, por enseñarme a pensar de forma analítica, a razonar, a tener una

estructura lógica. A Miguel Ospino, Rafael Antonio Guerrero, Carlos Maya y Orlando

Acosta, mil gracias por toda su ayuda.

Al Dr. Ravin Narain, director del Grupo de investigación en polímeros de la

Universidad de Alberta en Canadá, por abrirme las puertas de su laboratorio y ayudarme

en mi formación profesional. A la Dra. Bindu Tapha, Stephen Quan por su ayuda en el

XII Non-viral gene modification of mesenchymal stem cells in a tridimensional biocompatible scaffold


laboratorio. En memoria de Yinan Yang (Q.E.P.D.), el mejor profesional en el área de

polímeros que he conocido.

Al profesor Mario Enrique Velásquez Lozano, del Grupo de Investigación en Procesos

Químicos y Bioquímicos y a su estudiante de doctorado Margareth Patiño, por su ayuda

en la transformación de las bacterias y la obtención de los plásmidos.

Al Profesor Jaime Eduardo Castellanos, director de la maestría en Bioquímica de la

Universidad del Bosque y a la Dra. Eliana Calvo por su ayuda en la caracterización de los

plásmidos.

Al Profesor Camilo Ernesto López Carrascal y a la Dra. Edilene Ramírez del

Departamento de Biología de la Universidad Nacional de Colombia sede Bogotá por su

ayuda en la caracterización de los complejos.

Al profesor Jairo Ernesto Perilla, del Departamento de Ingeniería Química y Ambiental

por su ayuda con el préstamo del reómetro. Un agradecimiento similar a Diana Guzmán,

David Alejandro Daza y a Alejandro Serrano por su ayuda con las mediciones reológicas y

su asesoría con el uso del reómetro.

A la profesora Dolly Montoya Castaño, a los profesores Luis Francisco Boada Eslava,

Álvaro Orjuela Londoño, Paulo César Narváez Rincón, Néstor Ariel Algecira Enciso,

Rodrigo Jiménez Pizarro, Luis Ignacio Rodríguez Varela y Jorge Orlando Manrique

Perdomo, por su apoyo en la transición de las tesis de doctorado.

A mis jurados calificadores: la Dra. Luz Marina Restrepo Múnera, directora del Grupo

de Ingeniería de Tejidos y Terapias Celulares de la Universidad de Antioquia, la Dra.

Martha Raquel Fontanilla Duque, directora del Grupo de Ingeniería de Tejidos de la

Universidad Nacional de Colombia sede Bogotá y el Dr. Carlos Arturo Martínez Riascos,

Agradecimientos XIII


director del Instituto de Biotecnología (IBUN) de la Universidad Nacional de Colombia

sede Bogotá, mil gracias por sus aportes y contribuciones que mejoraron ostensiblemente

este trabajo.

Al Instituto Distrital de Ciencia, Biotecnología e Innovación en Salud (IDCBIS) por la

financiación recibida a través de los convenios especiales de cooperación entre el IDCBIS y

el Fondo Financiero Distrital de Salud y el Sistema General de Regalías.

A la Universidad Nacional de Colombia sede Bogotá, por su apoyo a través de la

Convocatoria para “Apoyo a proyectos de investigación y creación artística” del 2017.

A Intek Group SAS por su ayuda en la caracterización mediante microscopía

electrónica de barrido a través de la Convocatoria de “Apoyo a trabajos de investigación

en Educación Superior CLIG-001-2017”. A los ingenieros Ana Milena Piedra, Fabián

Espinel y Alejandra Escobar por su apoyo incondicional.

Al Ministerio de Ciencia, Tecnología e Innovación (MINCIENCIAS) por la beca

completa para participar de los cursos del Centro Argentino – Brasileño de Biotecnología

(CABBIO) desarrollado en la ciudad de Buenos Aires, Argentina en el año 2017. Así mismo,

por la financiación de mis estudios a través de la beca para Estudios de Doctorado en

Colombia - Convocatoria 567 del año 2012.

Al Servicio alemán de intercambio académico (DAAD) por la beca para “Viajes de

estudio para grupos de estudiantes extranjeros” del año 2017.

Al Gobierno Canadiense por la beca “Emerging Leaders in the Americas Program (ELAP)”

que me permitió realizar mi pasantía de investigación en el Laboratorio del Profesor Ravin

Narain en la Universidad de Alberta en el año 2014.

XIV Non-viral gene modification of mesenchymal stem cells in a tridimensional biocompatible scaffold


A Ana Rosa Salamanca Paternina, por su apoyo incondicional, por el café, las charlas,

los buenos momentos.

A Janeth Alejandra García Herrera de la Vicedecanatura de Investigación y Extensión

de la Facultad de Ingeniería por su ayuda con la corrección de estilo de esta tesis y de los

artículos publicados. Mil gracias por toda la paciencia y la ayuda.

A todo el personal del Laboratorio de Ingeniería Química (LIQ) de la Universidad

Nacional de Colombia, en especial a Ricardo Cortés Segura, Óscar Armando Camelo Roa,

Edgar Alfonso Martínez Ramírez y Martha Ramos. Mil gracias por su apoyo constante y

desinteresado. A Clarita y a don Julio de Servicios generales, muchas gracias. Al personal

de vigilancia del LIQ por su buena disposición: Sigifredo Ocampo, Horacio Ortiz, Hermes

Ariza, Oswaldo Zárate, Miriam Rojas y Gabriel Mesa.

A mis compañeros del Grupo de investigación de la Unidad de Terapias Avanzadas

del IDCBIS: Juli Buitrago, Carlos Medina, Leidi Yohana Méndez, Diana Mayorga, Valerie

Dorsant, Luz Stella Correa, Andrea Lizarazo, Darío Díaz, Mariana Cañas, Alejandro

Ricaurte, Karl Beltrán, Lorena González, William Cárdenas, Cristian Pinto, Adriana Lara,

Diana Londoño, Luisa Duarte y César Augusto Ramírez, mil gracias por toda su ayuda y

su buena disposición.

Al grupo del Banco de tejidos del IDCBIS, liderado por la Dra. Astrid Malagón e

integrado por Lina Marcela Rincón, Diana Carolina Nieto, Consuelo Forero, Luis Fernando

Huérfano, Diego Robayo y Cristian García. Gracias por acogerme, alimentarme y hacerme

sentir como en casa.

Al grupo del Banco de Sangre de cordón umbilical del IDCBIS: Andrea Moreno, Pavel

Medina, Laura Paipa y Ximena Bonilla. Mil gracias por todo el apoyo y la colaboración

brindada.

Agradecimientos XV


A los integrantes del área de Gestión de recursos operativos del IDCBIS, liderado por

la ingeniera Sandra Patricia Gómez y conformado en el área de Infraestructura por Edgar

Suárez, Javier Padilla del Castillo, Juan Gabriel García, Carlos Eduardo Torres, Omar

Augusto Villada, Feníbal González y en el área de Bioingeniería por Luis Fernando Mejía

y Jorge Alexander Pinzón. Así mismo, los integrantes del almacén Jennifer Bautista, José

David Zambrano, Richard Mesa y Sergio Eduardo García. Mil gracias por toda su ayuda

en la reparación y mantenimiento de los equipos, la adecuación de los espacios, la entrega

oportuna de los materiales y reactivos.

A la señora Verónica Ávila y Milena Mateus de Servicios generales del IDCBIS, mil y

mil gracias por sus cuidados constantes, por su preocupación, por tener siempre una

sonrisa y un saludo amable. A Adriana Castro Valbuena, asistente general del IDCBIS, mil

gracias por toda la ayuda desde los tiempos de la maestría, gracias por su disposición y por

estar siempre atenta a servir.

A todos los demás integrantes del IDCBIS que no alcanzo a nombrar y que enaltecen

el nombre de nuestro instituto y nos ayudan a crecer y mejorar cada día. Mil y mil gracias.

A mis amigos del grupo de investigación en Procesos Químicos y Bioquímicos: Andrés

Javier Bello, Leslie Vanessa Sánchez, Johan Andrés Pasos, Miguel Ángel Flórez, María

Alejandra García, Juan David Tibocha, Diego Alejandro Sánchez, Andrés Camilo Forero,

Juliana Vargas, Mateo Quintero, Natalia Cuéllar, Elizabeth Rodríguez, Fabián Rondón,

Mateo Rodríguez, Cindy Carolina Latorre, Valeria Colón, María Fernanda Candamil,

Andrés Galindo, Lorena Galván, Andrés Arias, Juan David Rangel y Astrid Nausa. Gracias

infinitas por creer, por hacerme una mejor persona, por permitirme crecer y aprender a su

lado. Gracias por su energía, por su deseo de ser mejores, por sus dudas, por sus

inquietudes, por sus temores, gracias por su confianza, por su apoyo infinito. Este camino

recorrido se hizo más divertido a su lado, más gratificante. Soy una mejor persona, un mejor

XVI Non-viral gene modification of mesenchymal stem cells in a tridimensional biocompatible scaffold


ser humano, una mejor profesional gracias a ustedes. Los llevo siempre en mi corazón;

fueron, son y serán una parte fundamental de mí. Gracias por montones.

A Duván, Isabella, Danna, Karen, Julián, Nathaly, Degly, Lina, Yara, Cheitania,

Michael, Lina, Mauricio, Julián, Juan José, Ana y demás integrantes del grupo de

investigación y participantes de los seminarios de los viernes, muchos de los cuales olvido

en estas cortas líneas; mil gracias por darle vida a este proyecto de grupo.

A Esteban Forero, Edgar Andrés Velázquez, Angie Rojas y Laura Ávila, muchas

gracias por su ayuda en la estandarización del proceso de cuantificación de las aminas

primarias.

A Elga Vargas, gracias por su apoyo y ayuda para entender mejor las células

estromales mesenquimales.

A mis amigos Víctor Manuel Trejos Montoya, Sandra Milena Castro Salazar, Juan

Gabriel Marín Gómez, Víctor Daniel Piedrahita y Danny Steven Gómez. Mil gracias por su

apoyo desde la distancia.

A mis amigos Luis Miguel Serrano Bermúdez, Camilo Antonio Monroy, Leidy Johana

Ortiz, Javier Fernando Méndez Monroy y Andrés Camilo, gracias por su apoyo, su cariño

y su soporte.

A Amalia Silva, Natalia Godoy, John Henry Pinzón, Sofía y Estefanía, gracias por todo

el cariño, los cuidados, el tiempo compartido. A Teresa Téllez y en memoria de Henry

Pinzón (Q.E.P.D.), gracias por los juegos de mesa y el cafecito. A Julieth Tijaro, Henry

Godoy, Paula y Laura, gracias por la compañía desde la distancia, las consultas médicas y

el apoyo constante.

Agradecimientos XVII


Finalmente, a mi familia, gracias por la paciencia, por entender mis ausencias y por el tiempo

juntos que les robé para hacer este sueño realidad. Gracias infinitas.

A mi mamá, Leonor, gracias por tanto. Gracias por estar ahí en el día a día, por las arepas del

desayuno, por el apoyo constante, por escucharme, por ayudarme a ser mejor persona cada día, por

inspirarme a ser lo que soy. Gracias por ser la mejor ingeniera de la casa, por enseñarnos a no

acostumbrarnos a las cosas como están, por renovar constantemente nuestra casa y de paso a nosotros

mismos. Gracias por enseñarnos la bondad, por hacer de nuestra casa ese lugar maravilloso llamado

hogar.

A mi papá, Antonio, gracias por tanto y tanto. Gracias por el apoyo, por las risas, por la

complicidad en los chistes, por siempre creer, por no desfallecer. Gracias por enseñarnos contabilidad

y reglas de tres y a hacer comida comunitaria. Gracias por siempre reír, por los chistes malos, por las

buenas historias. Gracias por ser esa mirada crítica, por enseñarnos a hacer lo correcto.

A mis hermanos, que sin su ayuda nada de esto hubiese sido posible. A Antonio, por decidir

estudiar por encima de todo, por su esfuerzo, por su tenacidad, por mostrarnos que era posible.

Gracias por llevarme a la escuela y cuidar de mí, por seguir haciéndolo hoy, por confiar y creer.

A Tobías, por su confianza ciega en mí, por forzarme a ser mejor, por llamar a preguntarme

cuándo vamos para la casa, por querer siempre a toda costa mantenernos unidos, por molestarme,

por ser la voz ecuánime de la casa, por llamar cada fin de semana a preguntar cómo voy, infinitas

gracias de todo corazón.

A Dubán, por su cariño, por su confianza infinita, porque hacernos mejores, por enseñarnos a

querer y a cuidar de alguien, porque lo hemos visto crecer y, hemos crecido a su lado y gracias a él.

(Peque, este esfuerzo es para ti). A Gabriela, gracias por el cariño, la paciencia y las risas. Gracias

por traer alegría a nuestra casa.

XVIII Non-viral gene modification of mesenchymal stem cells in a tridimensional biocompatible scaffold


A mi tía Alejandra, que siempre ha estado ahí para mí y para mi familia. Gracias infinitas por

abrirme las puertas de su casa y acogerme con tanto amor. Gracias por todo el esfuerzo, por

preguntarme día a día cómo me iba en la Universidad, por recoger mis historias con amor y por

contarme las suyas. Por entender y aceptar el momento cuando decidí abrir las alas y volar un poco

lejos. Sin ella y sin su ayuda no lo hubiera logrado.

A Martica, por ser nuestra segunda casa, por recibirnos después del Colegio, por su bondad

infinita, por su generosidad. Nunca encontraremos palabras y acciones suficientes para agradecer

todo lo que ha hecho por nosotros. A Daniel por su cariño y compañía, por estar pendiente de nosotros

y por cuidarnos. Mil gracias.

A mi abuelo Domingo, que cada fin de semana me llama a preguntar cómo estoy. Gracias por

esas preguntas difíciles que nos hacen reflexionar y por sus respuestas sencillas y profundas. Gracias

infinitas por cuidar de nosotros y llevarnos siempre en sus oraciones, gracias por la familia que formó

y por su amor a todo dar.

Resumen XIX


Resumen

Título: Modificación génica no viral de células madre mesenquimales en un soporte

tridimensional biocompatible

La necesidad creciente de tejidos y órganos para reemplazo y/o reparación impulsó el

desarrollo de nuevas disciplinas como la ingeniería de tejidos, que combina el uso de

células, andamios y moléculas biológicamente activas para reparar o restablecer la función

de los tejidos. Una aproximación para regular la liberación de las moléculas biológicamente

activas en los andamios de ingeniería de tejidos consiste en el uso de terapia génica, que

hace referencia a la introducción de material genómico exógeno (ARN o ADN) en las

células con el fin de generar un beneficio terapéutico.

La combinación de la terapia génica y la ingeniería de tejidos da lugar a la obtención

de andamios que se conocen bajo el nombre de matrices activadas génicamente o GAMs,

por sus siglas en inglés (Gene Activated Matrices). La presente tesis de doctorado consistió

en el desarrollo de una matriz activada génicamente, basada en andamios preparados a

partir de plasma humano crioconcentrado, combinado con células estromales

mesenquimales derivadas de la gelatina de Wharton (WJ-MSC) y complejos de

polietilenimina y ADN plasmídico.

Palabras clave: Transfección, matriz activada genéticamente, andamio, cordón umbilical,

terapia génica, plásmido

XX Non-viral gene modification of mesenchymal stem cells in a tridimensional biocompatible scaffold


Abstract XXI


Abstract

Title: Non-viral gene modification of mesenchymal stem cells in a tridimensional

biocompatible scaffold

The ever-increasing need for tissues and organs for replacement and / or repair

prompted the development of new disciplines such as tissue engineering, which combines

the use of biologically active cells, scaffolds, and molecules to repair or restore tissue

function. An approach to regulate the release of biologically active molecules in tissue

engineering scaffolds consists in the use of gene therapy, which refers to the introduction

of exogenous genomic material (RNA or DNA) into cells to generate a therapeutic benefit.

The combination of gene therapy and tissue engineering gives rise to scaffolds that are

known under the name of Gene Activated Matrices (GAMs). The present doctoral thesis

consisted in the development of a GAM, based on scaffolds prepared from

cryoconcentrated human plasma, combined with mesenchymal stromal cells derived from

Wharton's jelly (WJ-MSC) and complexes of polyethyleneimine and plasmid DNA.

Keywords: Transfection, gene activated matrix, biomaterial, umbilical cord, gene therapy,

plasmid

XXII Non-viral gene modification of mesenchymal stem cells in a tridimensional biocompatible scaffold


Content XXIII


Content

Pág.

Resumen ...................................................................................................................................... XIX

List of figures .......................................................................................................................... XXVI

Introduction..................................................................................................................................... 1

References ..................................................................................................................................... 5

1. Background ............................................................................................................................. 9

1.1 Mesenchymal stem cells (MSC): General information ............................................... 9

1.2 Different sources of MSC ............................................................................................. 10

1.2.1 Bone Marrow MSC .................................................................................................... 10

1.2.2 Placenta MSC ............................................................................................................. 10

1.2.3 Amniotic fluid MSC .................................................................................................. 11

1.2.4 Cord Blood MSC ........................................................................................................ 11

1.2.5 Adipose Tissue Mesenchymal Stem Cells ............................................................. 12

1.3 Human umbilical cord Wharton’s jelly-derived mesenchymal stromal cells (WJ-

MSC) 12

1.3.1 Umbilical cord structure ........................................................................................... 12

1.3.2 Differentiation of WJ-MSC ....................................................................................... 13

1.4 Non-viral gene delivery systems ................................................................................ 20

1.4.1 Cationic lipids ............................................................................................................ 20

1.4.2 Cationic polymers ..................................................................................................... 22

1.5 Complex formation ....................................................................................................... 23

1.5.1 Nucleic acids .............................................................................................................. 23

1.5.2 Entry of complexes into the cell .............................................................................. 25

1.6 References ....................................................................................................................... 28

2. Efficient non-viral gene modification of mesenchymal stromal cells from umbilical

cord Wharton’s jelly with polyethylenimine .......................................................................... 37

2.1 Introduction ................................................................................................................... 38

2.2 Materials and methods ................................................................................................. 41

2.2.1 Subsection ................................................................................................................... 41

2.2.2 Expansion of WJ-MSC .............................................................................................. 41

XXIV Non-viral gene modification of mesenchymal stem cells in a tridimensional biocompatible scaffold


2.2.3 Plasmid propagation ................................................................................................ 41

2.2.4 PEI and DNA nanoparticle formulations and physicochemical characterization

42

2.2.5 Growth kinetics of WJ-MSC .................................................................................... 44

2.2.6 Assessment of polyethylenimine (PEI) cytotoxicity on WJ-MSC ....................... 45

2.2.7 In vitro evaluation of transfection efficiency of PEI–DNA complexes in WJ-

MSC 45

2.2.8 Immunophenotype of WJ-MSC transfected with PEI .......................................... 46

2.2.9 Immunomodulation potency assay of PEI-transfected WJ-MSC ....................... 46

2.2.10 Evaluation of multilineage differentiation capacity of WJ-MSC ........................ 48

2.2.11 Statistical analysis ..................................................................................................... 48

2.3 Results and discussion ................................................................................................. 49

2.3.1 Mean hydrodynamic diameter and Zeta potential remain constant with the

increase in PEI concentration .............................................................................................. 49

2.3.2 Evaluation of cell toxicity induced by PEI complexes ......................................... 55

2.3.3 PEI Transfection Does Not Affect the Functional Properties of Wharton’s Jelly

MSC 61

2.4 Conclusions .................................................................................................................... 64

2.5 Supplementary materials ............................................................................................. 64

2.6 References....................................................................................................................... 67

3. Non-viral gene activated matrices based on human plasma for mesenchymal

stromal cells derived from human umbilical cord Wharton’s jelly ................................... 75

3.1 Introduction ................................................................................................................... 76

3.2 Materials and methods ................................................................................................. 79

3.2.1 Subsection .................................................................................................................. 79

3.2.2 Human plasma cryoconcentrated (HPCC) ............................................................ 79

3.2.3 Expansion of WJ-MSC and HEK cells .................................................................... 79

3.2.4 Culture of GFP-WJ-MSC over HPCC hydrogels at different fibrinogen

concentrations ........................................................................................................................ 80

3.2.5 Scanning Electron Microscopy (SEM) of HPCC hydrogels ................................ 82

3.2.6 GFP-WJ-MSC differentiation on HPCC hydrogels .............................................. 82

3.3 Gene activated matrices of HPCC .............................................................................. 85

3.3.1 Plasmid propagation ................................................................................................ 85

3.3.2 PEI and pGFP nanoparticle formulations and physicochemical

characterization ..................................................................................................................... 85

3.3.3 Transfection of WJ-MSC into HPCC scaffolds ...................................................... 86

3.3.4 Effect of fibrinogen concentration on the metabolic activity of WJ-MSC

embedded in fibrin scaffolds in the presence of PEI and PEI/DNA complexes .......... 87

3.3.5 Effect of WJ-MSC and PEI/DNA complexes on the rheological behavior of

fibrin gels ................................................................................................................................ 87

3.3.6 Effect of fibrinogen concentration and DNA content on the transfection

efficiency of HEK cells and WJ-MSC embedded in fibrin scaffolds .............................. 88

Content XXV


3.3.7 Statistical analysis...................................................................................................... 88

3.4 Results and discussion .................................................................................................. 89

3.4.1 Culture of WJ-MSC into HPCC gels ....................................................................... 89

3.4.2 Fibrinogen concentration and differentiation media in HPCC affects GFP-WJ-

MSC morphology .................................................................................................................. 91

3.4.3 Fibrinogen concentration in HPCC scaffolds does not affect the differentiation

of WJ-MSC .............................................................................................................................. 95

3.4.4 Fibrinogen concentration and culture media affect the inner morphology of

HPCC scaffolds. ................................................................................................................... 100

3.4.5 Fibrin scaffolds reduce PEI toxicity in WJ-MSC ................................................. 105

3.4.6 PEI/DNA nanoparticles characterization ............................................................. 113

3.4.7 PEI and DNA content regulate transfection efficiency in HPCC scaffolds ..... 113

3.5 Discussion..................................................................................................................... 119

3.6 Conclusions .................................................................................................................. 123

3.7 Supplementary material ............................................................................................. 124

3.8 References ..................................................................................................................... 125

4. Conclusions and recommendations ................................................................................ 135

4.1 Conclusions .................................................................................................................. 135

4.2 Recommendations ....................................................................................................... 135

Content XXVI


List of figures

Figure 1-1. Minimal criteria for defining multipotent mesenchymal stromal cells according

to the International Society for Cellular Therapy (ISCT) in 2006. ............................................ 9

Figure 1-2. Structure of the umbilical cord ................................................................................ 12

Figure 1-3. General structure of a cationic lipid. ...................................................................... 20

Figure 1-4. DOTMA. Chemical structure .................................................................................. 21

Figure 1-5. Liposome structure ................................................................................................... 21

Figure 1-6. Composition of linear PEI ........................................................................................ 22

Figure 1-7. Branched PEI .............................................................................................................. 23

Figure 1-8. RNA and DNA sugars .............................................................................................. 24

Figure 1-9. Nitrogenous bases (purines and pyrimidines) present in nucleic acids. .......... 24

Figure 1-10. Kopatz model for the entry of complexes into the cell. Reproduced with

authorization of Wiley Online Library ...................................................................................... 27

Figure 2-1. Detailed protocol used to transfect Wharton’s jelly-mesenchymal stem cells (WJ-

MSC) with PEI in 24-well plates by triplicate. .......................................................................... 47

Figure 2-2. Evaluation of PEI/DNA complexes formation. (a) Gel retardation assay of the

DNA plasmid to establish its size. (b) PEI/DNA complexes formation by gel retardation

assay. A value of 400 ng of DNA plasmid was mixed with PEI solutions in distilled water

(DW) at different concentrations to obtain complexes at several N/P ratios. N/P = 7

corresponds to 360 ng PEI. (c) Effect of N/P ratio on zeta potential. (d) Mean hydrodynamic

diameter and polydispersity of the sample (PDI). (a–c) denotes significance (n = 3 technical

replicates, p < 0.05) in comparison with all groups at the same time point. The error bars

represent 1 SD. ............................................................................................................................... 50

Figure 2-3. Transfection of WJ-MSC with PEI. (a) Effect of seeding density(C0) on the

growth kinetics of WJ-MSC. (b) Cell viability of 4.5 × 104 WJ-MSC/200 µL after 48 h of

treatment with PEI and PEI/DNA complexes at different N/P ratios. (c) Bright-field

photographs of non-treated WJ-MSC (without PEI) and treated with 720 and 1440 ng of

PEI/well after 24 h. Scale bar: 500 µm. (d) Percentage of WJ-MSC expressing green

fluorescent protein (GFP) by flow cytometry (y-axis, left) related with the percentage of

viable cells in the same assay (y-axis, right). (e,f) Fluorescent microscope images of

transfected WJ-MSC (48 h post-transfection). Scale bar: 1000 µm. (a–c) denotes significant

Content XXVII


differences (n = 3 biological replicates, p < 0.05) in comparison with all groups. The error

bars represent 1 SD. ...................................................................................................................... 56

Figure 2-4. Transfection with PEI did not affect the functional properties of WJ-MSC. (a)

Percentage of WJ-MSC expressing characteristic surface cell markers of mesenchymal

stromal cells. (b) Immunomodulatory potency of WJ-MSC over peripheral blood

mononuclear cells (PBMC). PBMC activated (Act) with CD2, CD3 and CD28 antibodies

were co-cultured with non-transfected (Act co-cul NT) and transfected (Act co-cul T) WJ-

MSC. Non-Activated (NAct) PBMC were used as controls. Data were expressed as absolute

CD3+ cell counts for each condition. Fold change (FC) was expressed as the ratio between

PBMC activated and non-activated. (n = 3 biological replicates, p < 0.05) in comparison with

all groups. The error bars represent 1 SD. (c) Adipogenic (scale bar: 100 μm) and osteogenic

(scale bar: 400 μm) differentiation of transfected (T) and non-transfected (NT) WJ-MSC

with PEI at an N/P ratio of 14 (720 ng PEI). ............................................................................... 63

Figure 3-1. General procedure to elaborate HPCC scaffolds with WJ-MSC ........................ 81

Figure 3-2. General procedure to differentiate WJ-MSC into HPCC scaffolds .................... 84

Figure 3-3. Methodology developed to prepare GAMs with WJ-MSC and HPCC. (A)

Preparation of the complexes by mixing equal volumes of PEI and DNA solutions in

distilled water. PEI is added to the DNA solution, the solution is vortexed for 10 s and

incubated for 30 min at 37°C. (B) WJ-MSC at 70% of confluence are recovered by

trypsinization and resuspended in DMEM 10% FBS. (C) A few seconds before preparing

the scaffolds, HPCC is mixed with 270 mM CaCl2 solution in a 10:1 proportion (HPCC-Ca).

(D) Complexes are mixed with WJ-MSC into the well plate, then HPCC-Ca solution is

added and mixed gently to avoid bubble formation. Fibrin gel is formed within 5-10

seconds. Then, DMEM 10% FBS and 0.5% tranexamic acid are added to each well (E)

Scaffolds are incubated at 37°C and 5% CO2, and transfection efficiency is evaluated 48 h

later. ................................................................................................................................................. 86

Figure 3-4. Effect of scaffold fibrinogen concentration and thickness on GFP-WJ-MSC

morphology and spreading. Fluorescence micrograph images of fibrin hydrogels at (A-C)

3, (D-F) 2, and (G-I) 1 µg of fibrinogen/µL containing 3.6x105 GFP-WJ-MSC/mL, in scaffolds

with different thicknesses: (A,D,G) 1, (B,E,H) 2, and (C,F,I) 3 mm. (J-L) GFP-WJ-MSC

cultured in monolayer were used as controls. Scale bar: 200 µm. Scaffolds were incubated

at 37°C and 5% CO2 for 2 days with DMEM supplemented with 10% FBS and tranexamic

acid (5% v/v). ................................................................................................................................. 90

Figure 3-5. Cell morphology evolution over time of GFP-WJ-MSC embedded in HPCC-

DMEM scaffolds. 2x105 GFP-WJ-MSC were seeded in scaffolds prepared at (A-C) 3, (D-F)

2, and (G-I) 1 µg of fibrinogen/µL. Scaffolds were incubated at 37°Cand 5% CO2 for 21 days

with DMEM supplemented with 10% FBS and tranexamic acid (5% v/v). (J-L) GFP-WJ-MSC

cultured in monolayer were used as controls. The media were changed every 3 days.

XXVIII Non-viral gene modification of mesenchymal stem cells in a tridimensional biocompatible scaffold


Fluorescence images were recorded at (A,D,G,J) day 1, (B,E,H,K) day 10, and (C,F,I,L) day

21. Scale bar: 200 µm. .................................................................................................................... 92

Figure 3-6. Schematic representation of GFP-WJ-MSC morphology embedded in HPCC-

DMEM scaffolds in comparison with monolayer cell culture, elaborated from fluorescence

images of cells from Figure 3-5. .................................................................................................. 93

Figure 3-7. Cell morphology evolution over time of GFP-WJ-MSC embedded in HPCC

scaffolds. Scaffolds prepared at 1 µg of fibrinogen/µLand 2x105 GFP-WJ-MSC per scaffold

were resuspended in (A-C) DMEM, (D-F) adipogenic, (G-I) chondrogenic, and (J-L)

osteogenic differentiation media. Scaffolds were incubated at 37°C, and 5% CO2 for 21 days.

Differentiation media supplemented with tranexamic acid (5% v/v) was changed every 3

days. Scale bar: 200 µm. ............................................................................................................... 95

Figure 3-8. Adipogenic differentiation of GFP-WJ-MSC embedded in HPCC-Adipogenic

media scaffolds. Adipogenic differentiation was evaluated at three cell seeding densities:

(A,E,I,M) 2, (B,F,J) 4, and (C,G,K) 6x105 GFP-WJ-MSC/scaffold. (D,H,L) 2x105 GFP-WJ-MSC

resuspended in DMEM media and cultured in HPCC-DMEM scaffolds were used as

control. Scaffolds prepared at (A-D) 3, (E-H) 2, and (I-L) 1 µg of fibrinogen/µL were stained

with Oil Red dye. (M) GFP-WJ-MSC cultured in monolayer and (N) HPCC scaffolds

without cells were used for differentiation control. Adipogenic differentiation media and

DMEM, both supplemented with tranexamic acid (5% v/v), were changed every 3 days.

Scale bar: 100 µm. .......................................................................................................................... 97

Figure 3-9. Chondrogenic differentiation of GFP-WJ-MSC embedded in HPCC-

chondrogenic media scaffolds. Chondrogenic differentiation was evaluated at three cell

seeding densities: (A,E,I,M) 2, (B,F,J) 4, and (C,G,K) 6x105 GFP-WJ-MSC/scaffold. (D,H,L)

4x105 GFP-WJ-MSC resuspended in DMEM media and cultured in HPCC-DMEM scaffolds

were used as control. Scaffolds prepared at (A-D) 3, (E-H) 2 and (I-L) 1 µg of fibrinogen/µL

were stained with Alcian Blue dye. (M) GFP-WJ-MSC cultured in monolayer and (N) HPCC

scaffolds without cells were used as differentiation controls. Chondrogenic differentiation

media and DMEM, both supplemented with tranexamic acid (5% v/v), were changed every

3 days. Scale bar: 100 µm. ............................................................................................................ 98

Figure 3-10. Osteogenic differentiation of GFP-WJ-MSC embedded in HPCC-Osteogenic

media scaffolds. Osteogenic differentiation was evaluated at three cell seeding densities:



control. Scaffolds prepared at (A-D) 3, (E-H) 2 and (I-L) 1 µg of fibrinogen/µL were stained

with Alizarin Red dye. (M) GFP-WJ-MSC cultured in monolayer and (N) HPCC scaffolds

without cells were used as differentiation controls. Osteogenic differentiation media and


Scale bar: 100 µm. .......................................................................................................................... 99

Content XXIX


Figure 3-11. Scanning electron microscopy (SEM) analysis of fibrin scaffolds. (A-D)

HPCC gels with variable fibrinogen concentrations (1.5, 1.7, 1.9, and 3.3 µg/µL) with 5.5

x105 GFP-WJ-MSC per scaffold after 7 days of culture in DMEM supplemented with 10%

FBS and tranexamic acid (0.5 % v/v) (DMEM-FBS-TA). HPCC gels (E) without cells after 7

days of incubation in DMEM-FBS-TA, and (F) without incubation (freshly prepared before

SEM analysis). Scale bar: 10 µm. (G) Average fiber diameter (bar height) of fibrin scaffolds

estimated by image processing using ImageJ software. The error bar corresponds to one

standard deviation (SD). (a,b,c,d) denotes significance (n = 20, p < 0.05) in comparison with all

groups. .......................................................................................................................................... 101

Figure 3-12. SEM photographs of increasingly smaller sections of the surface of a

scaffold made at a fibrinogen concentration of 6.6 µg/µL. The scaffold was made by

mixing HPCC with calcium, and WJ-MSC in a centrifuge tube of 50 mL to form a spherical

gel (diameter 8 mm) and cultured in DMEM+10%FBS+0.5% tranexamic acid (v/v) for 28

days. Note that the surface of the scaffold is characterized by the presence of a “skin”,

which is indicated in Figure 3-12B by the white arrows. Further magnification reveals the

presence of some sort of low-porosity material (white arrows in Figure 3-12E and F),

covering the most porous fibrin network (Figure 3-12F). ..................................................... 102

Figure 3-13. SEM photographs of increasingly smaller sections of two different adjacent

zones on the surface of a scaffold made at a fibrinogen concentration of 1.9 µg/µL. The

scaffold was made by mixing HPCC with calcium, DMEM+10%FBS and WJ-MSC, and

cultured for 7 days in DMEM+10%FBS+0.5% tranexamic acid (v/v). Orange-framed sections

(A-F) correspond to a fracture in the “skin” of the scaffold, which allows the visualization

of the fibrin network. Cyan-framed sections (G-J) focus on the structure of the skin. Greater

magnification reveals the radical difference in the structure of the skin vs. fibrin network,

visible at the same magnification in Figure 3-13Figure 3-13F, and J. .................................. 103


zones of the surface of a scaffold made at a fibrinogen concentration of 6.6 µg/µL without

cells. The scaffold was made by mixing 180 µL of HPCC with 10 µL solution of calcium

chloride 3% p/v. After gel formation, 400 µL of DMEM+10%FBS+ 0.5 % tranexamic acid

were added. The scaffold was incubated for 7 days at 37°C, and 5% CO2. ........................ 104

Figure 3-15. SEM photographs of increasingly smaller sections of the surface of a fresh

scaffold made by mixing HPCC with calcium at a final fibrinogen concentration of 6,6

µg/µL, without incubation. Note that the surface is homogeneous and characterized by

well-defined, clean fibers of fibrin. ........................................................................................... 104

Figure 3-16. Graphic representation of fibrin network .......................................................... 105

Figure 3-17. Effect of fibrinogen and PEI/DNA concentration on the metabolic activity

and rheological behavior of GFP-WJ-MSC embedded in HPCC scaffolds. 2x105 WJ-MSC

embedded in HPCC scaffolds at different fibrinogen concentrations were treated with 44

XXX Non-viral gene modification of mesenchymal stem cells in a tridimensional biocompatible scaffold


M resazurin in DMEM 10% FBS for 3 h. (A) Relative fluorescence of resazurin reduced to

resorufin as an indicator of metabolic activity was measured as a function of fibrinogen

concentration. The effect of the concentration of PEI (5, 10, and 15 µg/µL), both alone (P)

and complexed with DNA (C), was analyzed on the metabolic activity of WJ-MSC

embedded in HPCC scaffolds made at (B) 1, (C) 2 and (D) 3 µg/µL of fibrinogen. Cells

growing without PEI or DNA in the scaffolds at the proper fibrinogen concentration were

used as controls. In every case, the PEI/DNA mass ratio was maintained at 2. (a,b,c) denotes

significance (n = 3, p < 0.05) in comparison with all groups at the same fibrinogen

concentration. (E) Effect of GFP-WJ-MSC and PEI/DNA complexes on the elastic modulus

of HPCC scaffolds at 3 µg/µL of fibrinogen (strain: 0.1%). ................................................... 107

Figure 3-18. Agarose gel electrophoresis of PEI/DNA complexes at different PEI/DNA

mass ratios. Non-displacement in the band indicates the formation of complexes between

PEI and DNA. For this assay, 100 ng of DNA were mixed with PEI at different

concentrations to establish the PEI/DNA mass ratio at which complex formation occurs.

....................................................................................................................................................... 113

Figure 3-19. Effect of DNA content on the transfection efficiency of HEK cells in a

tridimensional scaffold of HPCC at 24 h. 4x105 HEK cells were seeded in scaffolds of

HPCC. Three different fibrinogen concentrations were evaluated: (A-E) 3 µg/µL, (F-J) 2

µg/µL and (K-O) 1 µg/µL. Complexes of PEI and DNA were prepared at five different

contents of DNA: (A, F and K) 1 µg, (B, G and L) 2 µg, (C, H and M) 3 µg, (D, I and N) 4 µg

and (E, J and O) 5 µg/scaffold. In every case, the PEI/DNA mass ratio was maintained at 2.

The final volume of each scaffold was 240 µL, scale bar: 500 µm ........................................ 115


tridimensional scaffold of HPCC at 60 h. Entire well reconstruction by image processing

using Gen5TM software from Biotek®. 4x105 HEK cells were seeded in scaffolds of HPCC.

Three different fibrinogen concentrations were evaluated (A-E) 3 µg/µL, (F-J) 2 µg/µL, and

(K-O) 1 µg/µL. Complexes of PEI, and DNA were prepared at five different contents of

DNA: (A,F, and K) 1 µg, (B,G, and L) 2 µg, (C,H, and M) 3 µg, (D,I, and N) 4 µg, and (E, J,

and O) 5 µg/scaffold. In every case, the PEI/DNA mass ratio was maintained at 2. Final

volume of each scaffold: 240 µL. Scale bar: 5 mm .................................................................. 116

Figure 3-21. Effect of DNA content on the transfection efficiency of WJ-MSC in a

tridimensional scaffold of HPCC at 48 h. 2x105 WJ-MSC were seeded in scaffolds of HPCC.

Three different fibrinogen concentrations were evaluated: (A-E) 3 µg/µL, (F-J) 2 µg/µL and


DNA: (A, F and K) 5 µg, (B, G and L) 6 µg, (C, H and M) 7 µg, (D, I and N) 8 µg, and (E, J


volume of each scaffold: 240 µL. Scale bar: 200 µm ............................................................... 117

Content XXXI




using Gen5TM software from Biotek®. 2x105 WJ-MSC were seeded in scaffolds of HPCC.


(K-O) 1 µg/µL. Complexes of PEI and DNA were prepared at five different contents of



volume of each scaffold: 240 µL. Scale bar: 5 mm .................................................................. 118

Figure 3-23. Effect of DNA content on the transfection efficiency of HEK cells, and WJ-

MSC in a tridimensional scaffold of HPCC. Quantitative analysis. 108 photos of each gel

at 4X were taken to reconstruct the conformation of the entire well. Images were processed

and analyzed using ImageJ software to establish the percentage of the fluorescent green

area into the well. Transfection efficiency was defined as the percentage of green area with

respect to the total area of the well. Effect of scaffold DNA content (PEI/DNA mass ratio of

2) was studied on transfection efficiency of (A) HEK cells and (B) WJ-MSC. Final volume

of each scaffold: 240 µL. ............................................................................................................. 119

XXXII Non-viral gene modification of mesenchymal stem cells in a tridimensional biocompatible scaffold



Introduction

Mesenchymal stem/stromal cells (MSC) are multipotent cells able to proliferate and

differentiate into specific lineages such as chondrogenic, adipogenic or osteogenic [1-3],

depending on the environmental conditions. MSC are obtained from different sources, such

as bone marrow [4-6], adipose tissue [7,8] dental pulp [9-11] amniotic fluid, [12,13]

placenta [14,15], umbilical cord blood [16-18], Wharton’s jelly umbilical cord [19-21],

synovial liquid [22-24], endometrium [25,26] and others.

Since MSC can be obtained from several sources and under different protocols (with

variations from laboratories), in 2006, the International Society for Cellular Therapy (ISCT)

established the “Minimal criteria for defining multipotent mesenchymal stromal cells”.

These criteria were plastic adherence (MSC must be plastic-adherent under standard

culture conditions), phenotype specificity (MSC must express CD105, CD73 and CD90, and

lack expression of hematopoietic markers such as CD45, CD34, CD14 or CD11b, CD79alpha

or CD19 and HLA-DR surface molecules) and in vitro differentiation (MSC must

differentiate into osteoblasts, adipocytes and chondroblasts) [27].

Recently, in 2019, the ISCT MSC committee made a new statement where they continue

to support the use of the acronym “MSC” but “recommend this be (i) supplemented by

tissue-source origin of the cells (which would highlight tissue-specific properties); (ii)

intended as MSC unless rigorous evidence for stemness exists that can be supported by

both in vitro and in vivo data; and (iii) associated with robust matrix of functional assays

to demonstrate MSC properties, which are not generically defined but informed by the

intended therapeutic mode of actions”[28].

One of the most remarkable sources of MSC are those obtained from Wharton's jelly

from the umbilical cord due to their high proliferation rates and low senescence in late

2 Non-viral gene modification of mesenchymal stem cells in a tridimensional biocompatible scaffold


passages, compared with stromal cells from other sources [19-21]. The umbilical cord is a

flexible conduct formed by two arteries and a vein surrounded by a gelatinous substance,

known by the name of Wharton's jelly, that joins the placenta with the navel of the embryo

and later of the fetus. This Wharton's jelly (WJ) is rich in MSC and its isolation from this

source does not generate any type of ethical conflict, as the umbilical cord is commonly

considered a biological waste [19-21].

To assess the impact of the use of WJ- MSC, 164 clinical trials were analyzed until the

end of 2019. Of these trials, 34 correspond to tissue engineering of skin, bone, cartilage,

muscle, and others. A percentage of 45% of these trials were in phase 1/phase 2 (studies

where the safety of the treatment, side effects and the best dose are tested) and 26% in phase

1 (determination of the highest dose of a new treatment that can be safely given without

causing serious side effects). These results suggest that the use of WJ-MSC is a technology

with a wide potential for development, which is susceptible to improvement if a series of

in vivo limitations are overcome after transplantation, such as low survival rates and

proliferation, hostile microenvironment of injured tissues, differentiation into undesirable

tissues, and impaired differentiation ability.

To overcome these difficulties, the use of gene therapy, which consists in the induction

of the expression of different factors, has been proposed as an alternative to increase its

survival and proliferation rate, and improve its pro-regenerative capacity, among others.

Gene therapy is an experimental technique used to introduce foreign genomic materials

(RNA or DNA) into host cells to generate a therapeutic benefit [29]. Gene supply vectors

are molecules that mediated the transfer of genetic material into the target cell, where it is

transcribed in parallel with genomic DNA [30]. Gene therapy systems can be classified as

viral, non-viral and hybrid systems. Viral methods consist of modified viruses with poor

replication but capable of delivering genetic material for expression. Examples of viral

system vectors include the use of adenovirus [31], associated adenovirus [32], retrovirus

[33], lentivirus [34], among others. Viral methods offer greater transduction efficiency and

gene expression over time, however, there are some limitations that restrict the use of these

Introduction 3


systems, as they may be associated with immunogenic problems, mutagenicity, inability to

transfer large genes, toxicity, and high costs [35-37].

Non-viral methods can be divided into physical and chemical. Physical methods use

mechanical and electrical forces to induce transient opening of the cell membrane for

transfection. Although these methods are relatively easy to develop, they need special

instruments, work best for adherent cells and can cause damage to nucleic acids [38]. The

most used physical methods include microinjection, electroporation and sonoporation,

among others. In contrast, the chemical approach involves the use of cationic lipids

(liposomes), cationic polymers or inorganic nanoparticles. The chemical vectors form

condensed complexes with negatively charged DNA through electrostatic interactions;

such complexes protect DNA from degradation and facilitate cell absorption and

intracellular supply. In addition, they are easy to scale and have a low stimulation of the

immune system. There is a wide variety of polymeric materials that can be modified to

modulate the efficacy and toxicity of transfection [35,36,39]. One of the chemical vectors

widely used in gene therapy is polyethylenimine, due to its high content of primary amines

that give it a positive net external charge that interacts with plasmid DNA, condense it and

facilitate its entry into the cell and subsequent release into the cytoplasm and entry into the

nucleus [40].

Furthermore, from the clinical point of view, they are more appealing, due to their ease

of scaling, low immune system stimulation and high versatility, and their use is preferred

when a transient expression is required. Among the non-viral systems, some are based on

reagents, among which the use of lipids, salts such as calcium phosphate, magnetic

particles and cationic polymers stand out. Polyethyleneimine is a cationic polymer

composed of ethyleneimine monomers CH3CH2NHx, where x is two for primary amines,

one for secondary amines and zero for tertiary amines. This distribution of amines gives

PEI an extraordinary cationic charge and a high buffering capacity, which benefits the

formation of complexes with nucleic acids (DNA/RNA), due to the differences in charge,

and favors endosomal/lysosomal release.



Gene therapy for the modulation of MSC characteristics has been widely studied in

two-dimensional models; however, despite the promising results obtained, these models

do not represent the three-dimensional conditions of the tissues to be repaired. To bring

these two-dimensional models to a three-dimensional one, the incorporation of a scaffold

is necessary. This scaffold must be biodegradable, biocompatible and must favor

interaction with cells.

Hydrogel scaffolds are widely used in tissue engineering due to their high

compatibility, mechanical properties like those of soft tissues, efficient transport of

nutrients and waste, uniform encapsulation of cells, and their capability to be injected in

situ. Fibrin is one of the hydrogels most used in tissue engineering, which meets the

previously mentioned characteristics and is also easy to handle. Fibrin is the polymeric

form of fibrinogen, which is present in mammals blood plasma and is involved in the

process of blood clotting. Human plasma cryoconcentrated (HPCC) is a by-product rich in

fibrinogen, obtained by freezing and thawing cycles of human plasma, resulting in the

precipitation of the protein (fibrinogen), which is further concentrated by centrifugation.

The gels obtained from HPCC have better rheological properties than gels prepared only

with human plasma.

Based on the above, the research question addressed in this work was: Is it possible to

genetically modify WJ-MSC using a three-dimensional reverse transfection model,

incorporating non-viral nanoparticles (NPs) and cells into a biocompatible hydrogel? The

hypothesis is that CEM-GW can adhere and grow in a hydrogel of HPCC and as they

colonize the support, they can be transfected with polyethyleneimine nanoparticles and

plasmid.

Accordingly, this work was aimed to analyze the effect of a cationic polymer in the

transfection of human MSC in a three-dimensional scaffold of human plasma

cryoprecipitate.

In order to achieve this general objective, the present work was divided into three

stages, which sought first the optimization of the transfection conditions in monolayer

culture before transferring the experiments to a three-dimensional model. In the first stage,

Introduction 5


the ratio between WJ-MSC, polyethyleneimine and a plasmid DNA, containing the genetic

information for the expression of the Green Fluorescent Protein (GFP) was explored, and

optimized in monolayer cultures, looking for the highest transfection efficiencies with the

least cellular toxicity (Chapter 3). The second stage aimed to standardize the conditions for

the preparation of a scaffold made from human plasma cryoprecipitate favoring both the

growth of mesenchymal stromal cells and their differentiation. For this stage, stromal cells

were used that expressed the green fluorescent protein in a constitutive way and that had

been previously developed at IDCBIS. Finally, in the third and final stage, the experiences

and results from the two previous stages were brought together, combining them for the

elaboration of a scaffold of human plasma cryoprecipitate and serving as a model for in

situ three-dimensional transfection of mesenchymal stromal cells (Chapter 4). This

document concludes with a section corresponding to the main conclusions reached, as well

as recommendations and suggestions for future work.

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https://doi.org/10.1016/B978-0-7234-3691-1.00099-4pp

1. Background

1.1 Mesenchymal stem cells (MSC): General information

Mesenchymal stem cells (MSC) are multipotent cells able to proliferate and

differentiate into several lineages depending on the environmental conditions [1,2]. In 2006,

the International Society for Cellular Therapy (ISCT) established a position statement about

the minimal criteria for defining multipotent mesenchymal stromal cells.

Figure 1-1. Minimal criteria for defining multipotent mesenchymal stromal cells

according to the International Society for Cellular Therapy (ISCT) in 2006.

z

α



Those criteria are: adherence to plastic in standard culture conditions; specific surface

antigen (Ag) expression; positive (≥95%) for CD105, CD73 and CD90, and negative (≤2%)

for CD45, CD34, CD14 or CD11B, CD79α or CD19 and HLA-DR measured by flow

cytometry and multipotent differentiation potential (osteoblasts, adipocytes and

chondroblasts) demonstrated by staining of in vitro cell culture [3].

1.2 Different sources of MSC

1.2.1 Bone Marrow MSC

Bone marrow was one of the first sources of stem cells studied. First works isolating

MSC from rat bone marrow were reported by Friedenstein et al.(1966) [4]. Arnold Caplan

of Case Western University called mesenchymal stem cells (MSC) a type of adult cells that

can be found in different tissues throughout life and that have to be affected and

propagated at the laboratory level [5]. In 1995, Lazarus et al. evaluated the therapeutic

potential of MSC and their safety in humans through a phase 1 trial to determine the

viability of harvesting, expansion of ex vivo culture and intravenous infusion of progenitor

stromal cells derived from human bone marrow (mesenchymal progenitor cells – MPC).

From these experiments, it was concluded that no adverse reactions were observed with

the infusion of the MPC [6]. Later, in 1999, Pittenger et al. demonstrated the ability to

multiline differentiation of clonal populations of MSC [7].

1.2.2 Placenta MSC

Fukuchi et al. (2004) isolated adherent cells from human placenta by digestion with

trypsin. They established two clones by successive (limiting) dilutions and examined cell

morphology, surface markers, gene expression patterns and potential of differentiation,

and proved that those cells are a useful source of mesenchymal stem cells [8]. Miao et al.

(2006) compared placenta MSC with bone marrow MSC (BM-MSC) in terms of growth

Chapter 1. Background 11


kinetics, immunophenotype and differentiation capability. Placenta MSC present no

significant difference in cell numbers and growth kinetics in comparison with BM-MSC,

and were positive for CD105, CD29 and CD44 and negative for the hematopoietic surface

markers of CD34, CD45, HLA-DR. Additionally, both sources of MSC were able to

differentiate into neurogenic and endothelial cells [9].

1.2.3 Amniotic fluid MSC

in´t Anker et al. (2003) found that second-trimester amniotic fluid (AF) is an abundant

source of MSC, and those cells exhibit a phenotype and multilineage differentiation

potential like that of postnatal BM-MSC [10]. In 2007, Roubelakis et al. evaluated the

immunophenotype of cultured MSC from second-trimester amniotic fluid and compared

the results with BM-MSC. They demonstrated that cultured MSC from both sources exhibit

similar expression patterns. They also carried out a protein map of both sources of cells,

evaluated the functional pattern and found that proteins from both sources were similar.

However, cultured AF-MSC displayed unique proteins related to proliferation and

primitive phenotype [11].

1.2.4 Cord Blood MSC

Lee et al. (2004) isolated blood from umbilical cords of 11 donors. Umbilical cord blood

(UCB) mononuclear cells were obtained by negative immunodepleting of CD3+, CD14+,

CD19+, CD38+, CD66b+, and glycophorin A+. Then, the cells obtained were separated by

Ficoll-Paque and cultured, while adherent cells were isolated. Finally, cells were serially

diluted to obtain single cells and colonies that grew were evaluated by morphology,

phenotype, osteogenic, chondrogenic, and adipogenic differentiation and finally by

ectodermal and endodermal differentiation. The results of those assays allowed concluding

that UCB does contain MSC [12].



1.2.5 Adipose Tissue Mesenchymal Stem Cells

In 2001, Zuk et al. analyzed if a population of MSC could be isolated from human

adipose tissue and found that adipose tissue may contain a significant fraction of cells with

multilineage capacity [13]. Puissant el al. (2005) discovered that adipose tissue-derived MSC

had a similar mechanism for immunomodulation to MSC from bone marrow [14].

1.3 Human umbilical cord Wharton’s jelly-derived

mesenchymal stromal cells (WJ-MSC)

1.3.1 Umbilical cord structure

The umbilical cord (UC) is a conduit between the placenta and fetus and normally

contains one vein and two arteries. It has a length of approximately 60 cm and an average

diameter of 1.5 cm. It is surrounded by a gelatinous substance composed of

mucopolysaccharides (chondroitin sulfate and hyaluronic acid), named Wharton's jelly

(WJ) in honor of Thomas Wharton who was the first to describe it in 1656.

The umbilical cord vein supplies oxygenated and nutrient-rich blood to the fetus from

the placenta and the fetal heart pumps low oxygen and nutrient-depleted blood back to the

placenta through the umbilical cord arteries [15] . Figure 1-2 shows a diagram of a cross

section of the umbilical cord where the distribution of its parts can be observed.

Figure 1-2. Structure of the umbilical cord

UC vein UC arteries

Wharton’s jelly

UC blood



McElreavey et al. were the first to isolate MSC from a portion of WJ from an umbilical

cord in 1991 [16]. In 2004, Wang et al. evaluated 30 fresh human umbilical cords obtained

after birth and demonstrated that mesenchymal cells from the human umbilical cord (UC-

MSC), expanded in culture, express adhesion molecules (CD44, CD105), integrin markers

(CD29, CD51), and mesenchymal stem cell (MSC) markers (SH2, SH3), but no markers of

hematopoietic differentiation (CD34, CD45). Additionally, the authors successfully

differentiated those cells into the three linages (osteoblasts, adipocytes and chondroblasts)

under suitable culture conditions and suggest that stroma cells from Wharton’s jelly are

similar to mesenchymal stem cells (MSC) [17].

Weiss et al. (2006) made a preliminary characterization of cells obtained from 15

different umbilical cords. They found that those cells have several important properties for

therapeutic use, since they can be isolated in large numbers, are negative hematopoietic

markers (CD34 and CD45), grow robustly after frozen/thawed process, and can be clonally

expanded and engineered to express exogenous proteins [18]. Weiss et al. (2008)

characterized the immune properties of UC-MSC and found that those cells have

immunosuppressive properties in vitro and low immunogenicity. Those results suggest

that UC-MSC would be permissive to allogeneic transplantation [19].

Since its first isolation in 1991, WJ-MSC has been used in different applications, mainly

they have been differentiated in several lineages using differentiation medium with or

without scaffolds.

1.3.2 Differentiation of WJ-MSC

Auditory Hair Cells Neuronal Progenitor Cells

Kil et al. (2016) isolated WJ-MSC and evaluated their differentiation into auditory hair

cells and neuronal progenitor cells. First, WJ-MSC were differentiated into neuronal

progenitor cells employing epidermal growth factor (EGF) and basic fibroblast growth

factor (bFGF) for 14 days. Each 3 days fresh bFGF was added to the culture medium. Then,

cells were differentiated into hair cells and neurons using neurobasal medium containing



glial cell-derived neurotrophic factor (GDNF), brain-derived neurotrophic factor (BDNF)

and neurotrophic factor 3 (NT-3) [20].

Cardiac Differentiation

In order to differentiate MSC from cardiac cells, inhibitors of DNA methyltransferases

are used, such as 5-azacytidine via ERK signal pathway [21-24] or DC301. Histone

deacetylase (HDAC) inhibitors are also used in combination with DNA methyl transferase

inhibitors and differentiation takes place with an active participation of Wnt mediators [25].

Other authors have reported that the scaffold affects cells differentiation. Lin et al.

(2016) found that WJ-MSC were able to differentiate into cardiac progenitor cells and

express cardiac specific markers in a scaffold of collagen coated with 0.1% of gelatin, in the

presence or absence of TGF- β2 [26]. Rabbani et al. (2017) elaborated a scaffold composed

of 70% of hyaluronic acid 50 kDa, 28% of PEG 6000 kDa and 2% chitosan. WJ-MSC were

combined with this scaffold and implanted in infarcted rabbit myocardium. The authors

found that cardiac function, angiogenesis and cardiogenesis was improved in comparison

with cells and scaffold alone.

In 2012, Quian et al. demonstrated that 5-azacytidine induces cardiac differentiation of

UC-MSC by activating extracellular regulated kinase (ERK) [23]. 5-Azacytidine acts as an

epigenetic modifier by incorporating into DNA, where it irreversibly binds to DNA

methyltransferases, thus inhibiting their activity. Several groups have reported that 5-

azacytidine induced the differentiation of mesenchymal stem cells into cardiomyocytes in

vitro [21-23]. Lian et al. (2016) also used 5-azacytidine and evaluated the effects of serial

passage on the characteristics and cardiac and neural differentiation of WJ-MSC. The

authors found that in the early and middle phases WJ-MSC were relatively stable and

cardiac differentiation capability decreased, whereas the propensity for neural

differentiation increased significantly in the middle phase compared with the early phase

[24]. Bhuvanalakshmi et al. (2017) also used inhibitors of DNA methyl transferase (DNMT)

histone deacetylase (HDAC) to reach cardiac differentiation of WJ-MSC [25].



Scaffolds are also used to induce differentiation into cardiac cells. Lin et al. (2016) used

a collagen coated with gelation scaffold to differentiate WJ-MSC into cardiac progenitor

cells. The medium was supplemented with TGF- β2, however, authors found that WJ-MSC

were able to differentiate into cardiac progenitor cells and express cardiac specific markers

in the presence or absence of TGF- β2. This demonstrates that the characteristics of the

scaffold significantly influence cell differentiation [26].

In 2017, Rabbani et al. evaluated the differentiation capability of WJ-MSC into cardiac

cells in vivo. They seeded WJ-MSC in a scaffold composed of hyaluronic acid, polyethylene

glycol (PEG) and chitosan and then implanted the scaffold with cell into cardiac defects

(infarcted myocardium) in rabbits. They found a reduction in the size of the defect and an

increase in neoangiogenesis and cardio myogenesis [27].

Chondrogenesis

In 1998, Johnstone et al. demonstrated that MSC can be differentiated into

chondrocytes using dexamethasone and TGF-β1. [28]. Dexamethasone (DEX) is a synthetic

glucocorticoid that combined with TGF- β1 increases aggrecan biosynthesis and gene

expression in BMSC [29], as well as the size of the pellets [30], in comparison with cells

treated with TGF-β1 alone [31]. It has been found that DEX upregulated protein levels and

gene expression of Col XI, a cartilage matrix marker, and enhanced the expression of

proteins mediated by TGF-β, such as aggrecan, COMP and Col II [32]. It was also evidenced

that TGF-β induces the expression of Sox9 (a transcriptional factor), an early gene of

chondrogenesis, enhanced mRNA expression of collagen type II, an important marker of

hyaline cartilage, enhanced aggrecan expression and reduced the expression of collagen I.

It is important to maintain hyaline cartilage phenotype of MSC-derived

chondrocytes[33,34]. TGF-β1 is mostly related to fibrosis [35,36] and wound healing [37],

while TGF- β3 is mostly used for chondrogenic induction [34,38].

Chondrogenic medium is also supplemented with sodium ascorbate (a salt of ascorbic

acid or vitamin C), which increases type II procollagen mRNA and type I procollagen



mRNA [39]. Ascorbic acid also promotes 1,25‐dihydroxy vitamin D3 synthesis and cartilage

matrix production [40].

ITS is a mixture of recombinant human insulin, human transferrin, and sodium

selenite. Insulin is a peptide hormone that promotes the uptake of amino acids and glucose,

lipogenesis, synthesis of proteins and nucleic acids, and intracellular transport. Transferrin

is an iron-transport glycoprotein. Iron in its free form can be toxic, so transferrin is used as

an iron-carrier to nourish cells in culture. Selenium is an essential trace element, as sodium

selenite is a co-factor for proteins such as glutathione peroxidase and it is used as

antioxidant. So, ITS is used as a basal medium supplement in order to reduce the amount

of fetal bovine serum (FBS) used to culture cells. Some formulations contain Ethanolamine

(ITS-X), which is a precursor of phospho-glycerides, essential to the structure of the plasma

membrane and cellular organelles. Sodium pyruvate is commonly added to cell culture

media as an additional source of energy.

Tanthaisong et al. (2017) evaluated chondrogenic differentiation of WJ-MSC through

GSK-3 inhibitors. It has been demonstrated that Lithium chloride (LiCl) has the potential

to be a GSK-3 inhibitor, which deactivates phosphorylation of β-catenin to initiate the Wnt

signaling pathway involved in chondrogenesis and cartilage development [41, 42].

Tanthaisong et al. (2017) used a chondrogenic standard medium composed of TGF-β3, ITS-

X, dexamethasone, ascorbate-2-posphate, L-proline and sodium pyruvate and

supplemented with LiCl. They found that the presence of LiCl upregulated the expression

of several genes, including ACAN, Col2a1, and Sox9. Also, LiCl in the presence of TGF-β3

can induce the Wnt signaling pathway and promote chondrogenic differentiation of WJ-

MSC, without inducing chondrocyte hypertrophy [41].

Widowati et al. (2018) evaluated the effects of insulin-like growth factor (IGF-I) in

chondrogenesis of WJ-MSC in an osteoarthritis model. IGF-I is a hormone similar in

molecular structure to insulin which acts as anabolic for articular chondrocytes and

stimulates the production of two key extracellular matrix proteins: proteoglycans and

type-II collagen, both in vitro [42] and in vivo [43]. The IGF1-induced WJ-MSC increased

expression of SOX9 and COL2 and decreased expression of matrix metalloproteinase, such



as ADAMTS1, ADAMTS5, MMP3, MMP1, and RANKL, which possible play a pathogenic

role in osteoarthritis [44]. This indicates that IGF1-induced WJ-MSC were capable of

enhancing chondrogenesis [45]

In 2017, Liu et al. employed WJ-MSC to repair osteochondral defects in a rabbit model.

They used a scaffold derived from the extracellular matrix (ECM) of swine cartilage and

prepare the chondrogenic induction medium with TGF-β1, ITS, dexamethasone and FGF

without FBS. They observed a spatial temporal remodeling orderly of WJ-MSC into

cartilage tissues for 16 months, with hyaline-like neocartilage formation and a complete

integration with adjacent host cartilage and regenerated subchondral bone. Besides, no

immune rejection was detected where xenografts were implanted into rabbit cartilage

defects [46].

Differentiation using commercial scaffolds has also been studied. Marmotti et al. (2017)

evaluated two scaffolds. The first one was based on hyaluronic acid and the second one

was composed of a double layer matrix of pig collagen type I and type III [47].

MSC from umbilical cord might have a potential for tissue engineering of bone and

cartilage, according to the results of Marmotti et al. (2017), who evaluated the chondrogenic

and osteogenic potential of UC-MSC in commercial tridimensional scaffolds. For

osteogenic differentiation they evaluated the expression of osteocalcin and RunX-2 and for

chondrogenic differentiation. They measured the expression of Sox-9 and type II collagen

[47].

Sadlik et al. (2018) developed a technique of cartilage repair in the knee using

Wharton's jelly derived MSC embedded onto a type I/III collagen scaffold and implanted

in a minimally invasive fashion using dry arthroscopy. The scaffold was moistened with

saline solution and then immersed in the WJ-MSC suspension for a period of 5 minutes to

create the final WJ-MSC embedded scaffold graft. They carried out magnetic resonance

imaging every 6 weeks, 12 weeks, and 6 months postoperatively to monitor the repaired

lesion and rehabilitation progression. After 9 months of implantation, they observed the

integration of regenerative tissue with surrounding cartilage and subchondral lamina [48].



Widowati et al. (2018) evaluated the effect of insulin-like growth factor (IGF) on

chondrogenesis of WJ-MSC in an osteoarthritis model and found that IGF1-induced WJ-

MSC increased expression of COL2 and SOX9, which indicates IGF1-WJMSC are capable

of enhancing chondrogenesis [45].

Glycogen synthase kinase 3 (GSK-3) inhibitors can induce the Wnt signaling pathway,

which is involved in cartilage development and chondrogenesis processes. Tanthaisong et

al. (2017) used GSK-3 inhibitors and TGF-B3 to enhance chondrogenic differentiation of

WJ-MSC without inducing chondrocyte hypertrophy [41].

The BCOR gene encodes a protein known as the BCL6 co-repressor. Wang et al. (2017)

found that the BCOR negatively regulates insulin-like growth factor binding protein 2

(IGFBP2). They also discovered that an overexpression of IGFBP2 enhances adipogenic

differentiation of WJ-MSC through the activation of c-Jun N terminal kinase (JNK) and

protein kinase B (Akt) signaling pathways. Those results suggest that IGFBP2 may be a

potential target to promote the adipose tissue regeneration [49].

Famian et al. (2017) evaluated the effect of a conditioned medium of WJ-MSC on

cartilage specific genes expression by using chondrocytes in mass culture systems. Their

results suggest that WJ-MSC can increase the expression of cartilage-specific genes and can

be introduced as a promoting factor for cartilage regeneration [50].

Pereira et al. (2017) co-cultured WJ-MSC with articular chondrocytes (hAC) and a

micromass differentiation model. WJ- hMSC exposed to soluble factors secreted by hACs,

were able to express higher levels of chondrogenic genes with deposition of extracellular

matrix components of cartilage. Those results suggest their use as an alternative cell source

for treating degenerated cartilage [51].

Alves da Silva et al. (2017) compared WJ-MSC and hAC cultured in polycaprolactone

(PCL) nanofibrous meshes. They observed higher glycosaminoglycans production and

over-expression of cartilage-related genes from WJ-MSC cultured with basal medium in

comparison with hAC isolated from osteoarthritic joints. The authors observed the

presence of sulfated proteoglycans and collagen type II on both types of cell cultures. Their



results suggest that this effect is due to the electrospun nanofibers topography and the

intrinsic chondrogenic differentiation potential of WJ-MSC [52].

Liu et al. (2014) cultured WJ-MSC in pellets combined with rotary cell-culture system

(RCCS) to fabricate an engineering cartilage tissue without scaffold. They found that in

assays in vitro, WJ-MSC can differentiate to cartilage and produce a substantial cartilage

matrix. They also evidenced that RCCS culture with a chondrogenic medium can facilitate

the formation of larger and condensed cartilage like tissue [53].

Reppel et al. (2015) evaluated the chondrogenic potential of WJ-MSC embedded in

alginate/hyaluronic acid hydrogel over 28 days without adding growth factors at transcript

and protein levels. Those cells were able to adapt to their environment and express specific

cartilage-related genes and matrix proteins [54].

Aleksander-Konert (2016) et al. evaluated two commercially hyaluronic acid-based

hydrogels, HyStem and HyStem-C, for the cultivation of WJ-MSC and their differentiation

towards chondrocytes. Their results indicate that WJ-MSC have some degree of

chondrogenic potential in HyStem and HyStem-C hydrogels, showing promising

properties for the engineering of damaged articular cartilage [55]. During in vitro

chondrogenic differentiation of WJ-MSC on PLGA scaffolds, Paduszyńsk et al. (2016)

evaluated changes in the expression of cartilaginous genes. Those changes suggest that the

PLGA scaffolds may be applied to WJ-MSC differentiation [56].

Osteogenesis

Like the chondrogenic medium, the osteogenic medium is composed of

dexamethasone and ascorbic acid. The main difference between both of them is the

presence of β-glycerophosphate in osteogenic media. β-glycerophosphate (Glycerol 2-

phosphate) is a phosphoric ester of glycerol [57-62]. Langenbach and Handschel (2013)

analyzed the effects of dexamethasone, ascorbic acid and β-glycerophosphate on the

osteogenic differentiation of stem cells in vitro [63]. In 2007, Baksh et al. compared

proliferative and multilineage differentiation potential of MSC derived from umbilical cord

and bone marrow [64]. In 2009, Hou et al. discovered that BMP2 successfully mediates the



osteogenic differentiation of UC-MSC using a signal transduction pathway like that for BM-

MSC [57].

1.4 Non-viral gene delivery systems

Nucleic acids (DNA or RNA) are negatively charged as is the cell membrane; therefore,

delivery of genetic information through the cell membrane is challenging. Non-viral gene

delivery systems involve the conjunction of nucleic acids carrying specific genetic

information, and certain chemical moieties to form small particles able to interact with the

cell surface and being internalized by endocytosis, thus effectively delivering the genetic

information into the cell. The two main approaches to non-viral gene delivery involve the

combination of nucleic acids with cationic lipids to form lipoplexes and/or cationic

polymers to form polyplexes.

1.4.1 Cationic lipids

Cationic lipids are formed by four components (Figure 1-3): a cationic headgroup

(hydrophilic), a hydrophobic tail, a backbone domain and a linker group [65] . Because of

its positively charged nature, the cationic headgroup interacts with nucleic acids to form a

lipid-DNA (or ARN) complex (commonly named lipoplex) via electrostatic interactions

[66].

Figure 1-3. General structure of a cationic lipid.

In 1987, Felgner et al.[67] developed DOTMA, the first synthetic cationic lipid used in

gene therapy which was able to interact with DNA (Figure 1-4). DOTMA or Trimethyl[2,3-

(dioleyloxy) propyl] ammonium Chloride N-(1-(2,3-dioleyloxy)propyl)-N,N,N-

Linker group

Hydrophobic

tail

Hydrophilic headgroup

Backbone domain

It is between two

Positively charged and interacts with

nucleic acids

Steroid or alkyl chains (saturated or unsaturated)

Separates the hydrophilic headgroup from the

hydrophobic tail. Acts as a platform on which the cationic

lipid is built (the most common is glycerol)



trimethylammonium), has a molecular weight of 670.6 g/mol (molecular formula

C42H84ClNO2). Since then, a large amount of diverse cationic lipids has been synthetized.

Figure 1-4. DOTMA. Chemical structure

Figure 1-5. Liposome structure

Bhattacharya et al.(2009) classified cationic lipids into the next 11 categories according

to their headgroups, as they are a strong determinant in transfection efficiency: cationic

lipids with ammonium headgroups (either with hydrocarbon chains or cholesterol-based),

cationic lipids with polyammonium and multivalent head groups, nucleolipids, cationic

lipids with amino acid head group, sugar-based cationic lipids, chemically triggered

cationic lipids for effective DNA release, dimeric surfactants, cationic phospholipids, lipids

with targeting ligands, polymerizable cationic lipids and a series of miscellaneous cationic

lipids based on calixarenes, steroids and other hydrophobic moieties[66].



The main advantages of cationic lipids are their biocompatibility and their ability to

form liposomes (Figure 1-5), a structure that surrounds the molecule of interest (DNA,

drug, enzyme, others) and can protect it from degradation [68] while effectively

transporting it through the cell membrane into the cytoplasm. Because their

biocompatibility, cationic lipids can be used in in-vivo assays. The main disadvantage of

cationic polymers is that the transfection efficiency depends on its chemical identity and on

the cell line; therefore, the transfection conditions for each cell line and cationic lipid must

be established. Cationic lipids have been used to transfect a wide variety of cell lines such

as COS-7 [69], CHO [70], HepG2 [71], HeLa [72], and HEK [71], among others.

1.4.2 Cationic polymers

Polyethylenimine

Polyethyleneimine (PEI) is a polymer composed of repetitions of ethyleneimine (—

CH2CH2NH—) subunits. Each subunit has a molecular weight of 43 Dalton. There are two

types of PEI: linear and branched. In linear PEI, nitrogen atoms are always bonded to two

carbon atoms and one hydrogen; therefore, linear PEI will only have secondary amine

groups, except for one primary amine group at the end and another at the beginning of the

chain (Figure 1-6).

Figure 1-6. Composition of linear PEI

In the case of branched PEI, as shown in Figure 1-7, the molecule will have primary

amines (in green), secondary amines (red) and tertiary amines (blue). Akinc et al.(2005) and



Dai et al.(2012) point out that the ratio of primary, secondary, and tertiary amine groups in

branched PEI is approximately 1: 2: 1. [73,74].

Figure 1-7. Branched PEI

At physiological conditions, whenever there is a secondary or tertiary amine, the

molecule is positively charged. These positive charges allow PEI to interact by electrostatic

attraction with molecules of opposite charge, such as nucleic acids.

1.5 Complex formation

1.5.1 Nucleic acids

Nucleic acids, such as the deoxyribonucleic acid (DNA) and the ribonucleic acid

(RNA), are polymers composed of repeated units of nucleotides, in the case of DNA, or

ribonucleotides, in the case of RNA. Both nucleotides and ribonucleotides are organic

molecules composed of a phosphate group, a sugar (a pentose) and a nitrogenous base,

which are joined together by phosphodiester bonds (covalent bond between the phosphate

group of one nucleotide and the hydroxyl group of the pentose of the next nucleotide),

forming single strands of DNA or RNA (Figure 1-9). The differences between nucleotides

and ribonucleotides lie in the type of nitrogenous base and sugar that make them up.



Sugars can be β-D-ribofuranose (usually known as ribose) or 2-deoxyribose (a ribose

without the oxygen atom from hydroxyl 2’) (Figure 1-8). Nitrogenous bases are derived

from purines and pyrimidines (heterocyclic aromatic compounds). Adenine and Guanine

are derived from purines, while Cytosine, Thymine and Uracil are derived from

pyrimidines (Figure 1-9).

Figure 1-8. RNA and DNA sugars

On one side, a ribonucleotide contains the sugar ribose (β-D-ribofuranose) while the

nitrogenous base can be uracil, adenine, cytosine or guanine (never thymine). On the other

side, a nucleotide contains deoxyribose —(2-deoxyribose)—, and a nitrogenous base that

can be adenine, thymine, cytosine, or guanine (never uracil).

Figure 1-9. Nitrogenous bases (purines and pyrimidines) present in nucleic acids.

Double strands of RNA or DNA are formed by double hydrogen bonds between the

nitrogenous bases of the single chains, always retaining the complementarity of the base

junction, where adenine always binds with thiamine in the case of DNA or with uracil in

the case of RNA, while the cytosine in both nucleic acids always bind with guanine.

When the double helix structure of the nucleic acids is formed, the nitrogenous bases

remain inside the structure, while the sugars and phosphate groups remain outside. These

phosphate groups are negatively charged ([—PO4—]-), thus giving nucleic acids a negative



overall charge. Some authors affirm that since PEI is a molecule with a high positive charge,

it can interact by electrostatic attraction with nucleic acids, forming "complexes" that

protect said nucleic acids from degradation by, for example, endonucleases (enzymes

catalyzing the breakdown of phosphodiester bonds); however Dai et al.(2012) controvert

such claim and affirm that “complexation is driven by the gain of translational entropy,

namely, the releasing of counterions, instead of the gain of enthalpy via electrostatic

interaction” [73,75].

1.5.2 Entry of complexes into the cell

Regarding the entrance of the complexes to the cell, it has been postulated that the

entry occurs through the interaction of the complexes with syndecans, ubiquitous

transmembrane proteoglycans (glycosaminoglycan + a protein skeleton) which are

expressed by all adherent cells and can interact with a variety of ligands through their

protein skeleton and chondroitin or heparan sulfate chains [76].

Kopatz et al.(2004) proposed a model for the entry of the complexes into the cell (Figure

1-10). First, the complex bind to the Syndecans, then occurs a lateral diffusion of more

syndecan molecules near to the complex, grouping in cholesterol rafts. This triggers

activation of protein kinase C, phosphorylation of the tails of the syndecans and binding of

actin (protein-mediated) to the cytoplasm tail of syndecans. This actin forms a network and

the formed tension fibers attract the particles into the cell via phagocytosis [77].

Once inside the cell, the complexes must escape from the lysosomes to be able to exert

their function. Benjaminsen et al.(2013) postulate three possible escape mechanisms of the

complexes [78]:

1. Not all the complexes reach the lysosomes and that small part that does not arrive is

the one that mediates the transfection in the cell, while the others that follow the endocytic

pathway are degraded and exoculated.

2. The complexes escape from lysosomes because of the effect known as "proton

sponge", without this effect generating an increase in pH in the lysosomal space.



This is where the controversy begins, because, although Benjaminsen et al.(2013) state

that the proton sponge can occur, it might not necessarily be a “buffer” effect. It is known

that PEI can bind a large number of protons and this would "buffer" the lysosomes; in

response to this capacity of the PEI, the ATP-driven V-ATPase pump would inject more

protons into the lysosome in order to compensate for the effect of PEI. In that sense, the

proton sponge hypothesis may be correct, without necessarily changing the lysosomal-

space pH.

Another aspect that has been controversial in the “proton sponge” hypothesis is the

relationship between the concentration of PEI in the lysosomal space and release of the

complex. Kopatz et al.(2004) measured the concentration of PEI in the lysosome space and

found[77]:

- Despite the nitrogen concentration of PEI be greater than 300 mmol/liter, less than 1%

of lysosomes reach that concentration

- Approximately 50% of liposomes have a concentration lower than 40 mmol/L of PEI-

nitrogen.

- As the concentration of PEI increases, there is more damping, that is, more chlorine

ions enter, which leads to differences in osmotic pressure, due to the entry of water and

therefore a greater tension in the membrane.



Figure 1-10. Kopatz model for the entry of complexes into the cell. Reproduced with

authorization of Wiley Online Library

Kopatz et al.(2004) made an approximation of the osmotic pressure in the lysosome as

a function of the concentration of PEI and found that lysosomes with 100 and 50 mmol/L

of PEI-nitrogen can reach sizes around 775 and 1500 nm, respectively, before exploding.

Through simulations, they found that concentrations of PEI higher than those values are

needed for the lysosome to burst and those higher concentrations are the least likely;

therefore, an escape of the complexes through this pathway is unlikely[77].

3. The complexes escape from lysosomes through pores (holes) in the membrane, due

to an interaction of PEI with the endosomal membrane and an increase in membrane

tension due to the “proton sponge”.



On the other hand, Merdan et al.(2002) demonstrated that the complexes accumulate

in acidic vesicles, probably lysosomes and the release occur suddenly, probably due to the

bursting of these organelles. When the ATPase pump is inhibited with bafilomycin A1

(vacuolar proton pump inhibitor) no lysosomal escape was observed [79]

Once the complex is released from the endosome and is free in the cytosol, the

characteristics of the nucleic acid fraction play an important role in the fate of transfection.

RNA must get rid of the polymer (i.e. PEI), fold properly and reach ribosomes to be

expressed, while DNA must reach the nucleus for transfection to take place. In the cytosol,

nucleases can degrade nucleic acids that are not protected by the polymer; therefore, if the

nucleic acid is complexed with PEI, it will be protected. The most likely way for DNA to

access the nucleus is during mitosis (Pollard et al., 1998). It has been shown, however, that

mitosis is not the only route, since Horbinski et al.(2001) efficiently transfected postmitotic

neurons using PEI [77,80,81].

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2. Efficient non-viral gene modification of

mesenchymal stromal cells from umbilical cord

Wharton’s jelly with polyethylenimine

The content of this chapter was published in Pharmaceutics 2020, 12(9), 896.

https://doi.org/10.3390/pharmaceutics12090896

Ana Isabel Ramos-Murillo 1,2 , Elizabeth Rodríguez1, Karl Beltrán2, Cristian Ricaurte2,

Bernardo Camacho2, Gustavo Salguero2 and Rubén Darío Godoy-Silva1,*

1 Chemical and Biochemical Processes Research Group, Department of Chemical and

Environmental Engineering, Faculty of Engineering, Universidad Nacional de Colombia,

111321, Bogotá D.C., Colombia.

2 Advanced Therapies Unit, Instituto Distrital de Ciencia, Biotecnología e Innovación en Salud

(IDCBIS), 111611, Bogotá D.C.

* Correspondence: [email protected]; Tel.: +57-1316-5000 (ext. 14307)

Abstract: Mesenchymal stromal cells (MSC) derived from human umbilical cord

Wharton’s jelly (WJ) have a wide therapeutic potential in cell therapy and tissue

engineering because of their multipotential capacity, which can be reinforced through

gene therapy in order to modulate specific responses. However, reported methodologies

to transfect WJ-MSC using cationic polymers are scarce. Here, WJ-MSC were transfected

using 25 kDa branched- polyethylenimine (PEI) and a DNA plasmid encoding GFP.

PEI/plasmid complexes were characterized to establish the best transfection efficiencies

with lowest toxicity. Expression of MSC-related cell surface markers was evaluated.

Likewise, immunomodulatory activity and multipotential capacity of transfected WJ-

https://doi.org/10.3390/pharmaceutics12090896

https://orcid.org/0000-0001-9578-7480

https://orcid.org/0000-0002-5018-9927

https://orcid.org/0000-0001-6064-3743



MSC were assessed by CD2/CD3/CD28-activated peripheral blood mononuclear cells

(PBMC) cocultures and osteogenic and adipogenic differentiation assays, respectively. An

association between cell number, PEI and DNA content, and transfection efficiency was

observed. The highest transfection efficiency (15.3 ± 8.6%) at the lowest toxicity was

achieved using 2 ng/L DNA and 3.6 ng/L PEI with 45,000 WJ-MSC in a 24-well plate

format (200 L). Under these conditions, there was no significant difference between the

expression of MSC-identity markers, inhibitory effect on CD3+ T lymphocytes

proliferation and osteogenic/adipogenic differentiation ability of transfected WJ-MSC, as

compared with non-transfected cells. These results suggest that the functional properties

of WJ-MSC were not altered after optimized transfection with PEI.

Keywords: gene therapy; differentiation; cationic polymer; immunophenotype;

immunomodulation; cell therapy; standardization; polyplexes

2.1 Introduction

Mesenchymal stem cells (MSC) was the name proposed in 1991 by Arnold Caplan for

a class of cells isolated from human and mammalian periosteum and bone marrow, able to

be expanded in culture while preserving their in vitro capacity to form several mesodermal

phenotypes and tissues. Additionally, they were proposed as multipotent cells able to

proliferate and differentiate into several lineages depending on the environmental

conditions [1]. Due to the heterogeneity of isolation and cultivation procedures among

laboratories, in 2006, the International Society for Cellular Therapy (ISCT) proposed that

the abbreviation MSC should be used to designate multipotent mesenchymal stromal cells

and established a position statement about the minimal criteria to call them this way. These

criteria were adherence to plastic in standard culture conditions, specific surface antigen

(Ag) expression, positive (≥95%) for CD73, CD90 and CD105 and negative (≤2%) for CD12B

or CD14, CD34, CD45, CD79α or CD19 and HLA-DR, as measured by flow cytometry and

Chapter 2. Transfection of WJ-MSC with PEI in monolayer 39


multipotent differentiation potential (osteoblasts, adipocytes and chondroblasts)

demonstrated by staining of in vitro cell culture [2].

Bone marrow (BM) has been the most common source of MSC [3-5]; however, their

clinical application is limited because their collection is invasive, painful [6], and has low

cell yield [7]. As a result, new sources of MSC have been explored. The umbilical cord (UC)

is a conduit between the placenta and fetus, and it is surrounded by a gelatinous substance

made up of mucopolysaccharides (chondroitin sulfate and hyaluronic acid) named

Wharton’s jelly (WJ) [8,9]. WJ has been characterized as a promising source of MSC since

derived cells have some advantages over other sources of multipotent stromal cells [9,10].

WJ-MSC display higher proliferation, lower senescence rates, and relatively higher

expression of pluripotency markers than stromal cells obtained from other sources [11-14].

Furthermore, since WJ-MSC are obtained from the neonatal umbilical cord, which is

considered a biological waste [15], the ethical concerns are strongly reduced [10,11,14].

Several researchers have found that regenerative capacity in vivo of MSC-based

therapies is sub-optimal due their low survival, poor biodistribution and reduced

differentiation rates [16-18]. To overcome these difficulties, MSC can be modified via genetic

engineering to improve their survival rate, increase the secretion of differentiation factors

needed to induce a specific lineage [19], improve the immunomodulatory capacity and

increase the secretion of cytokines [20], among others [21,22]. Additionally, the discovery of

several genes related to the repair of damaged tissues brings the opportunity to genetically

modified cells, such as MSC, with huge potential in the treatment of diseases that benefit

from transitory or durable expression of therapeutic genes [21].

The success of gene delivery is highly dependent on the carrier employed to transfer

DNA or RNA. Non-Viral vectors such as liposomes and cationic polymers are preferred

when transient modifications are expected because they are safe and easily scalable, albeit

their low transfection efficiency and short transgene half-life, thereby limit their clinical

application [23]. In this regard, the use of polyethylenimine (PEI) as a transfection agent has

emerged as a potential candidate for engineering MSC from different sources [24-26]



displaying satisfactory results. PEI is a cationic polymer widely used for non-viral

transfection because it combines strong DNA compaction with a potential for endosomal

escape [27].

While there is a vast amount of literature regarding methodologies for PEI-driven

transfection of MSC from diverse sources, to date, quite few reports exist on the use of

cationic polymers in the transfection of umbilical cord MSC. Bahadur et al. (2015),

conjugated 1.2 kDa PEI with linoleic acid and hyaluronic acid and evaluated transfection

in MSC of bone marrow (BM) and umbilical cord (UC) and used 25 kDa branched PEI as

transfection control [28]. Although they reached transfection efficiencies of up to 40% in

UC-MSC, the viability was reduced to 60% or lower, making it a methodology that can be

improved [29]. With the goal of transfecting UC-MSC with functionalized particles, Wang

et al. (2017) included a modified HIV-1 trans-activator of transcription protein (TAT) into

the transfecting particles; they reached a peak transfection efficiency of 12% with 25 kDa

branched PEI vs. 14% with the functionalized particles, although they found significant

differences in cell viability with both approaches, going from 70 to 90%, respectively [29].

Unfortunately, both mentioned papers evaluated the expression of surface markers to

confirm the identity of MSC before transfection but there is no indication in their work

related to the effect of transfection on the expression of those markers, which is essential

for further use of transfected MSC in clinical applications.

The main aim of this work was to evaluate the non-viral gene modification of WJ-MSC

by using PEI. We evaluated a methodology that combined optimal PEI ratios using smaller

amounts of DNA to increase transfection efficiencies. Additionally, we were able to confirm

that major biological properties of WJ-MSC including cell viability, immunophenotype,

immunomodulatory capacity and expression of MSC-markers were preserved after PEI

transfection, favoring the use of PEI as the optimal methodology for transient gene of WJ-

MSC for future clinical translation.



2.2 Materials and methods

2.2.1 Subsection

Cell culture: Growth media Dulbecco's Modified Eagle's medium (DMEM)11885-084,

Gibco, Life Technologies Corp., Carlsbad, CA, USA) and fetal bovine serum FBS (12103C,

Sigma Aldrich, St. Louis, MO, USA) were employed for WJ-MSC culture. Roswell Park

Memorial Institute (RPMI) 1640 medium (61870-036, Gibco, Life Technologies Corp.,

Carlsbad, CA, USA) was used to culture peripheral blood mononuclear cells (PBMC).

Transfection: 25 kDa branched polyethylenimine (PEI) (408727, Sigma Aldrich, St. Louis,

MO, USA) was employed in all transfection assays. Nucleus staining DAPI (4′,6-diamidino-

2-phenylindole) was used at 300nM (D9542, Sigma Aldrich, St. Louis, MO, USA) in

phosphate-buffered solution 1X (PBS) at pH 7.4. Cell viability: Resazurin sodium salt

(121519, PanReac AppliChem, Darmstadt, Germany) in PBS at pH 7.4 was used to evaluate

the metabolic activity of cells as a measurement of cell viability. Escherichia coli DH5-α

culture: Luria Bertani (LB) broth, Miller (Ref. L3152), and ampicillin sodium salt (Ref.

A0166) were acquired from Sigma Aldrich (St. Louis, MO, USA) and agar-agar (A0949) was

supplied by PanReac AppliChem (Darmstadt, Germany).

2.2.2 Expansion of WJ-MSC

WJ-MSC were obtained from umbilical cord donors (donors 40 and 148) from the

Advanced Therapies Unit at the IDCBIS, following informed consent from mothers. WJ-

MSC at passages 1 to 6 were cultured in low-glucose DMEM supplemented with 10% FBS

at 37 °C and 5% CO2.

2.2.3 Plasmid propagation

Competent E. coli DH5-α cells (kindly donated by Dr. Velásquez laboratory at

Universidad Nacional de Colombia) were transformed with a DNA plasmid

(RRL.SIN.cPPT-hCMV-eGFP) coding for GFP and resistance to ampicillin. Transformed E.



coli cells were cultured in Luria Bertani (LB) medium and LB with agar-agar and selected

with ampicillin. The GFP plasmid (DNA) was purified using a ZymoPURE™ plasmid

maxiprep kit (catalog #D4203, Zymo Research, Irvine, CA, USA) following the

recommended manufacturer procedure. The DNA concentration and purity were

measured using a NanoDrop™ 2000/2000c Spectrophotometer (ThermoScientific®,

Wilmington, DE, USA).

2.2.4 PEI and DNA nanoparticle formulations and physicochemical

characterization

Branched PEI (25 kDa) was employed as the transfection agent. A PEI stock solution at

10 µg/µL was prepared in distilled water (DW), and the pH was adjusted to 3.15 with

hydrochloric acid (HCl) (H1758, Sigma Aldrich, St. Louis, MO, USA). The PEI stock

solution was filtered using a 0.2 µm filter. PEI working solutions at 1 µg/µL were prepared

by dilution of the PEI stock solution in sterile DW and were aliquoted at 500 µL before

freezing at −20 °C. In all experiments, fresh working PEI solutions were used to avoid

repeating cycles of freezing and thawing. Transfection efficiency was evaluated at different

N/P ratios (molar ratio of nitrogen in PEI/molar ratio of phosphorus in DNA). It was

considered that every basic amino group of PEI was potentially responsible for DNA

binding. One nitrogen (N) per repeat unit of PEI is CH3CH2NHx, where x is the average

number of protons attached to N, (x = 2 × %NH2 (percentage of primary amines)+ 1 × %NH

(percentage of secondary amines)). Thus, Mw: 43.1 g/mol [30,31]. At DNA, the repeating

unit is a nucleotide, which has one phosphorus atom and an average molecular weight of

330 g/mol, approximately. As a rule, there are 3 nmol of P every µg of DNA, approximately

[24,30,32,33]. In this work, the overall nitrogen content was quantified using a Total Organic

Carbon/Total Nitrogen (TOC/TN) analyzer (Analytik Jena, Jena, Germany). Our analyses

showed that a 50 mg/L solution of PEI in DW contained 14.6 mg/L of TN. According to the

elemental composition of the repeating PEI unit, the theoretical TN should be 16.3 mg/L;

thus, we adjusted our PEI content to 90% purity, considering that PEI is highly hygroscopic,

and this difference corresponds to hydrated PEI. One of the main limitations of using PEI



is the difficulty in obtaining reproducible results and comparing them with the results

reported by other authors. To avoid these difficulties, Table 1 displays the detailed N/P

ratio calculations used in this work.

Table 1. N/P ratio calculations to prepare polyethylenimine (PEI)/DNA complexes.

PEI (ng)

Without

Correction

PEI (ng) Corrected

by TN 1 Content

DNA

(ng)

PEI/DNA

Mass Ratio

Moles of N

From PEI

Moles of P

From DNA

N/P

Ratio

100 90 400 0.2 2.1 1.2 1.7

200 180 400 0.5 4.2 1.2 3.5

400 360 400 0.9 8.4 1.2 7.0

800 720 400 1.8 16.7 1.2 14.0

1200 1080 400 2.7 25.1 1.2 20.9

1600 1440 400 3.6 33.5 1.2 27.9

2000 1800 400 4.5 41.9 1.2 34.9

1 TN: Total Nitrogen.

According to Table 1, different N/P ratios were evaluated using 400 ng of DNA. DNA

plasmid aliquots and PEI concentrations were adjusted to maintain the same volume of

both solutions in every mixture. Transfection assays were carried out in 24-well plates. Each

well had a superficial area of 2 cm2; thus, a minimum volume of 0.2 cm3 (200 µL) was

required to cover all the surface. Several authors recommend avoiding exceeding 10% of

the volume with complexes [30,34]. Therefore, considering that all the experiments were

made in triplicate, the complexes were prepared in a final volume of 70 µL. This means that

35 µL of PEI solutions in DW at different concentrations were added to 35 µL DNA plasmid

solution at 40 ng/µL, and the solution was vortexed immediately. After 30 min of

incubation at 37 °C, the complexes were resuspended in 625 µL of prewarmed DMEM to

obtain a final volume of 700 µL. Next, 200 µL of the complexes in DMEM were added to

each well.



To evaluate the effect of the N/P ratio on the mean hydrodynamic diameter and zeta

potential, several measurements were carried out using dynamic light scattering (DLS) and

laser doppler micro-electrophoresis (LDME) (Zetasizer Nano ZS, Malvern Panalytical,

Worcestershire, UK), respectively. This technique requires the dilution of the sample in 1

mL of distilled water doubly filtered (DWDF) (0.22 µm). In this case, 120 µL of complexes

prepared in 4 batches of 30 µL (15 µL of DNA and 15 µL of PEI solution) were mixed with

1 mL of DWDF. The first measurements carried out were the mean hydrodynamic

diameters. Then, the same sample was used to measure the Zeta potential. All

measurements were carried out in triplicates. Complexes formed were also evaluated by

electrophoresis in agarose gel, which contained 0.79 g of agarose and 100 mL of TAE buffer

0.5X stained with Ethidium Bromide. Bio-Rad power supply was employed to apply 100V

through the agarose gel for 30 min. 1 Kb Plus DNA Ladder (10787-018, Invitrogen, Vilnius,

Lithuania) was used for sizing and quantification of double stranded DNA on agarose gels.

BlueJuiceTM (10816-015, Invitrogen, Vilnius, Lithuania) was used as loading buffer.

2.2.5 Growth kinetics of WJ-MSC

To evaluate the effect of the initial cell concentration on the growth kinetics of WJ-MSC,

two different seeding densities were evaluated: 2000 and 9000 WJ-MSC/cm2. WJ-MSC were

cultured in a 48-well plate for 24 h, and then the medium was removed from the first 6

wells of the 48-well plate and was replaced with 100 µL of resazurin solution in

supplemented DMEM at 44 µM. The whole plate was incubated at 37 °C and 5% CO2 in a

Cytation® 3 microplate reader (Biotek®, Winooski, VT, USA), and metabolic activity was

monitored for 24 h by fluorescence. At this time, the same procedure was carried out for

the next six different wells until day 6. In this way, the error due to the possible cytotoxic

effect of resazurin on the cells is reduced.



2.2.6 Assessment of polyethylenimine (PEI) cytotoxicity on WJ-MSC

Metabolic activity was established as a measure of the variation of cell viability,

through the reduction of resazurin to resorufin. PEI cytotoxicity was evaluated using the

resazurin assay technique. For this purpose, 4.5 × 104 WJ-MSC per well were cultured in a

24 well plate for 24 h. Next, culture medium was removed, and cells were treated with PEI

and PEI/DNA complexes in DMEM without FBS at different concentrations for 4 h at 37 °C

and 5% CO2. After that time, PEI or PEI/DNA solutions were removed, and 100 µL of

resazurin solution 44 µM in DMEM + 10% FBS were added to each well and fluorescence

intensity was quantified every hour using a Cytation 3 microplate reader with wavelengths

of 530 and 590 nm for excitation and emission, respectively. All assays were conducted in

triplicates. Cells treated with DMEM without FBS were used as controls.

2.2.7 In vitro evaluation of transfection efficiency of PEI–DNA

complexes in WJ-MSC

Samples of 1 × 104 WJ-MSC/cm2 in DMEM + 10% FBS were cultured in 24-well plates

for 48 h before transfection at 37 °C, 5% CO2. At this time, media were removed from wells

and 20 L of complexes of PEI and DNA (prepared as previously described) were

resuspended in 180 L of DMEM without FBS and the mixture was added to each well

carefully. The plate was centrifuged at 280 g for 5 min and the cells were incubated for 4 h

with the complexes. Then complexes were removed, replaced with DMEM + 10% FBS and

cells were incubated again. After 48 h of transfection, WJ-MSC were trypsinized, recovered

by centrifugation and resuspended in 100 μL of PBS 1X; GFP expression was evaluated by

flow cytometry using a FACSCanto II (Becton Dickinson, San Jose, CA, USA). Data were

analyzed using FlowJo™ Software V10.7.1 (Becton, Dickinson and Company, Ashland, OR,

USA). Untreated cells and cells treated with non-complexed plasmid were used as controls.

In every treatment, data are represented by the mean and standard deviation (SD) of three

replicates.



2.2.8 Immunophenotype of WJ-MSC transfected with PEI

To evaluate the presence of characteristic cell surface markers of MSC and the absence

of hematopoietic cell markers, WJ-MSC were harvested by trypsinization, 48 h after the

transfection protocol with PEI at an N/P ratio of 14 (720 ng PEI/90000 cells). The expression

of MSC-related cell surface antigens was assessed by flow cytometry using the membrane

markers CD90 (APC), CD73 (PECy7), CD105 (PE), CD45 (APC/Cy7), CD34 (PerCP-Cy5.5),

HLA-DR (Pacific Blue), and HLA-ABC (Clone W6/32, FITC). Cells were incubated for 30

min at 4 °C, then, centrifuged at 300 g for 6 min and resuspended in 0.2 mL PBS. Cell

acquisition and MSC phenotyping was performed using a FACSCanto II flow cytometer

and data were analyzed with FlowJo™ Software V10.7.1 (Becton, Dickinson and Company,

Ashland, OR, USA). Non-Transfected WJ-MSC were used as the negative control.

2.2.9 Immunomodulation potency assay of PEI-transfected WJ-MSC

MSC have been reported to inhibit the activation and proliferation of cells of the

immune system. To assess the ability of WJ-MSC to maintain this ability after PEI

transfection, an immunomodulation assay was performed, measuring the ability of

transfected WJ-MSC to inhibit the proliferation of CD3+ T lymphocytes. Peripheral blood

mononuclear cells (PBMC) were cultured in RPMI medium supplemented with 10% v/v

FBS and used as a source of T lymphocytes. PBMC were previously activated with CD2,

CD3 and CD28 monoclonal antibodies (130-091-441, T cell Activation/Expansion Kit,

Miltenyi Biotec GmbH, Bergisch Gladbach, Germany). Initially, WJ-MSC were transfected

following the procedure described in Figure 2-1, using an N/P ratio of 14. After 48 h of

transfection, transfected cells were trypsinized, recovered by centrifugation, seeded into

24-well plates at 5 × 104 cells per well, and cultured for 24 h. Resulting cells were then co-

cultured with 5 × 105 of activated PBMC (Act co-cul T) at 37 °C with 5% humidified CO2 for

5 days. Activated PBMC co-cultured with WJ-MSC untransfected (Act co-cul NT), activated

PBMC (Act) and non-activated PBMC (NAct) were used as controls. After 5 days of culture,

PBMC were recovered from wells, incubated at 4 °C for 30 min with antiCD3 antibody



(300329, Pacific Blue™ anti-human CD3 Antibody, Clone HIT3a, Biolegend, San Diego, CA,

USA), and then examined by flow cytometry.

Figure 2-1. Detailed protocol used to transfect Wharton’s jelly-mesenchymal stem cells

(WJ-MSC) with PEI in 24-well plates by triplicate.

Prepare a PEI stock in distilled deionized water (DDW)

10 mg/mL

Adjust pH at 3.6 with HCl and filter (0.22 m)

Prepare a PEI working solution (1 mg/mL), diluting 1 mL of

PEI stock solution (10 mg/mL) in 9 mL of sterile DDW

Prepare aliquots (200 L each one) of PEI working solution (1

mg/mL) and store at -20°C (avoid frozen-thaw cycles)

Seed a 24-well plate with 2 x104 WJ-MSC/well (1x104

WJ-MSC/cm2 ) in DMEM+10 %FBS and incubate at 37°C and 5% CO2.

*Start transfection when cells reach 2x104 WJ-MSC/cm2 (double of seeding

density)

COMPLEXES PREPARATION

TIPS- Use same volume of DNA

and PEI solution- The volume of the

complexes must not exceed 10% of the total volume of transfection

Prepare 300 L of plasmid DNA solution 40 ng/L in sterile DDW

Prepare 50 L of PEI solutions at several concentrations (0, 10, 20, 40, 80, 120, 160 and 200

ng/L) in DDW

Add 35 L of DNA plasmid solution to each microcentrifuge tube (8 tubes in total)

To one tube containing DNA plasmid add 35 L of

one of the PEI solution. Repeat for the other seven

PEI solutions

Vortex for 5 seconds

Incubate complexes for 30 minutes at 37°C

Add 630 L of DMEM to each tube and mix gently

Add 200 L/well of diluted PEI/DNA complexes to cells. Calculations are based in assays carried out by

triplicate

Centrifuge the plate at 230g for 5 minutes

Incubate cells with the diluted complexes for 4 hours at 37°C and 5% CO2

Remove complexes and add 400 L pre-warmed DMEM+10% FBS to each well

Incubate cells for 24-48 hours at 37°C and 5% CO2

before visualizing/analyzing transfected cells

PRELIMINAR

Remove the culture media just before adding the complexes

Day

0

CELL CULTURE

Day

1 –

2*

Day 3-4

Day

0

TRANSFECTIONFormat

Superficial area

(cm2/well)

Transfection volume

(L/well)

24-well plate

2 200



2.2.10 Evaluation of multilineage differentiation capacity of WJ-MSC

The multipotential capacity of the WJ-MSC after transfection with PEI at an N/P ratio

of 14 (720 ng PEI/9 × 104 cells) was examined by culturing transfected cells in secondary

cultures using osteogenic and adipogenic differentiation media. To induce adipogenic

differentiation, transfected cells were seeded into 24-well plates at 5 × 104 cells per well and

cultured until 60% confluency was achieved. At this point, medium was replaced with

adipogenic induction medium (Stromal Pro Adipogenesis Differentiation Kit, Life

Technologies, USA). The medium was changed every 3 days and after 14 days, cells were

fixed in 4% paraformaldehyde (Sigma Aldrich, St Louis, MO, USA) before staining lipid

vacuoles with Oil Red O (Sigma Aldrich, St Louis, MO, USA). For osteogenic

differentiation, transfected cells were seeded similarly and exposed to osteogenic

differentiation medium (Stromal Pro Osteogenesis Differentiation Kit, Life Technologies).

Calcium deposition was analyzed using Alizarin red-S (Sigma Aldrich, St Louis, MO, USA)

staining. For all controls (non-differentiation conditions), cells were kept in DMEM + 10%

FBS under standard culture conditions. The procedure was repeated using non-transfected

(NT) WJ-MSC. In all cases, observations were made using microscopic and photographic

records.

2.2.11 Statistical analysis

Statistical analyses were performed using Statgraphics Centurion 18 software

(Statgraphics Technologies, Inc., The Plains, VA). One-way ANOVA was used for analysis

of variance with Tukey’s post hoc test for comparison between groups. Numerical and

graphical results are displayed as mean ± standard deviation. Significance was accepted at

a level of p < 0.05.



2.3 Results and discussion

2.3.1 Mean hydrodynamic diameter and Zeta potential remain

constant with the increase in PEI concentration

DNA/PEI complexes used for transient transfection of WJ-MSC were characterized by

electrophoresis and dynamic light scattering (DLS). First, in order to evaluate the

performance of a gene delivery vector, it is necessary to establish the minimum N/P ratio

for the effective complexation of a given amount of DNA. Furthermore, evaluation of

transfection conditions to obtain an optimum N/P ratio at which transfection efficiency is

maximized, is then required to reduce cell toxicity. For this purpose, the size of the plasmid

used in the formation of complexes was evaluated by electrophoresis. Two bands were

distinguished when plasmid was not digested in the second lane of a; the same two bands

were visible also in the second lane (DNA) of the gel shown in Figure 2-2a and very likely

corresponded to the open-circular (oc) and covalently closed circular (ccc) DNA, which

suffer from friction against the agarose matrix. The size of the linearized plasmid was

resolved after digestion with EcoRI and corresponded to approximately 8 kb, as shown in

Figure 2-2a. Once the plasmid size was established, complexes prepared at different PEI

concentrations using the same amount (400 ng) of plasmid DNA at 40 ng/L were

evaluated by electrophoresis in agarose gel, where a delay shift of any given band

suggested DNA complexation with PEI. At N/P ratios equal or lower than 1.7, we observed

bands migrating in a similar pattern to the one observed for the plasmid alone, indicating

that higher N/P ratios are required to allow the formation of PEI-DNA complexes (Figure

2-2b). At N/P ratios equal or greater than 3.5, DNA bands are not seen in the electrophoresis

gel once there is enough PEI to complex with DNA, because the condensation of DNA with

PEI either excludes or displaces ethidium bromide from intercalating with DNA;

additionally, the cationic character of both ethidium bromide and PEI causes a repulsion

between the two molecules, reducing or preventing the electrostatic interaction between

DNA and ethidium bromide. As a result, the fluorescence will be reduced to the basal level



of ethidium bromide in the aqueous solution of the gel and will no longer be detectable as

a band in the corresponding lanes of the electrophoresis.

Figure 2-2. Evaluation of PEI/DNA complexes formation. (a) Gel retardation assay

of the DNA plasmid to establish its size. (b) PEI/DNA complexes formation by gel

retardation assay. A value of 400 ng of DNA plasmid was mixed with PEI

solutions in distilled water (DW) at different concentrations to obtain complexes

at several N/P ratios. N/P = 7 corresponds to 360 ng PEI. (c) Effect of N/P ratio on

zeta potential. (d) Mean hydrodynamic diameter and polydispersity of the sample

(PDI). (a–c) denotes significance (n = 3 technical replicates, p < 0.05) in comparison

with all groups at the same time point. The error bars represent 1 SD.

After establishing the conditions to form complexes, zeta potential and particle size

were evaluated. As expected, due to the presence of phosphate groups on its surface, a

PEI content (ng) (N/P ratio)

DN

A

PE

I

36

0 (

7)

72

0 (

14

)

10

80

(2

1)

14

40

(2

8)

18

00

(3

5)

Zet

a p

ote

nti

al (

mV

)

-40

-20

0

20

40

60

a

b

c

PEI content (ng) (N/P ratio)

360

(7)

720

(14)

1080

(21

)

1440

(28

)

1800

(35

)

Mea

n h

yd

rod

yn

amic

dia

met

er (

nm

)

0

50

100

150

200

PD

I

0,0

0,2

0,4

0,6

0,8

1,0

PDI

a

b b

N/P ratio in the complexa b

Marker DNA PEI 0.9 1.7 3.5 14 17.5 21 2810.57.0

Marker DNA P/EcoRI

10.08.0

3.0

2.0

1.5

kb

c d

Marker

10.06.0

1.5

kb

0.1



negative zeta potential of approximately −27.0 ± 0.9 mV was obtained for a DNA stock

solution at 400 ng/mL in DW; this value is slightly more negative than obtained by Mady

et al. (2011) [35]. In contrast, the zeta potential of a PEI solution (33.3 µg/mL) in DW was

positive, but oscillated between 6 and 26 mV, meaning the highest variability (coefficient

of variation of 51%); such value is also slightly smaller than the reported by Mady et al.

(2011) [35] and using the same analytic technique. The lowest N/P ratios evaluated were 1.7

and 3.5, and their zeta potential values were 17 ± 1 and 23 ± 2 mV, respectively. Under these

conditions, it was not possible to measure the hydrodynamic sizes by DLS because samples

were too polydisperse for the cumulative analysis used in the DLS technique. This behavior

can be explained by comparing our results with those obtained by Mady et al. (2011) [35]

who evaluated complex formation by measuring the relative fluorescence intensity emitted

upon the addition of PEI to ethidium bromide (EB)-DNA complex (fluorescent light could

be quenched by the addition of PEI that competes with EB to bind DNA). They reported

that supernatants of complexes prepared at N/P values lower than 3.5, showed fluorescence

suggest incomplete DNA condensation by PEI [35].

Figure 2-2c shows zeta potential measurements varied between 33 ± 3 and 45 ± 1 mV

for N/P ratios between seven and 35, respectively. A Tukey multiple comparison test was

performed to determine which zeta potential averages were significantly different from the

others. Statistically significant differences were only found at a N/P ratio of 14 (confidence

level of 95%). For the other N/P ratios, no differences were found (p < 0.05). Finally, the

mean hydrodynamic diameter of the complexes was measured as a function of the N/P

ratio. Figure 2-2d shows a slight but significant reduction in the mean hydrodynamic

diameter when the N/P ratio was 14 (123 ± 3 nm) in comparison with the other four N/P

ratios evaluated, which on average had a diameter of 152 ± 7 nm. It must be mentioned that

the polydispersity index of those measurements ranged from 0.25 to 0.45, which means that

in reality there was a wide distribution of particle sizes for the evaluated conditions; such

distribution will be important when discussing the possible route of entrance of PEI/DNA

complexes into the cell.



Han et al. (2009), indicated that, in the phase of formation of the PEI / DNA complexes,

the outcome of the transported transgenes is largely defined [36]. Therefore, complex

preparation is one of the critical steps to maximize the results of PEI-mediated protein

expression processes [36]. Van Gaal et al. (2011) analyzed the different parameters varied

in transfection assays with non-viral vectors, which are: a buffer used for complexation

(water, NaCl, HEPES buffer), DNA dose, cell confluency, incubation period of complexes

with cells and incubation medium (with or without serum) [37]. In addition to these

parameters, Bono et al. (2020) also suggest an effect of the method of complex formation

(by dripping, mixing with a pipette, homogenizing with vortex) and the relationship

between the volume of the complex and the volume of culture medium. These operational

aspects influence complex size, zeta potential, toxicity, and transfection efficiency [30].

Bono et al. (2020) compared the transfection efficiency of the complexes prepared in

water with those prepared in 150 mM NaCl, using murine fibroblasts (L929 cell line) and

25 kDa linear PEI and found that the transfection efficiency of the complexes prepared in

NaCl was increased 10 times compared to the complexes prepared in water [30]. They also

observed that the particle size increased from 4.5 to nine times, going from particles with

sizes between 100–200 nm when complexes were prepared in water, to others of 900 nm

when prepared in NaCl. According to the authors, transfection efficiency is favored by

larger particles, which settle faster than small ones, according to Stokes’ law that describes

the movement of a sphere of diameter (d) in a gravitational field. According to Stokes’

equation (Equation (2-1)), the velocity of sedimentation (v) is given by the relation of the

diameter of the sphere, the particle (p) and the medium (L) density, the gravitational force

(g) and the viscosity of medium (n):

v = d2(p − L)g/18n (2-1)

In presence of a gravitational force, particles of higher density or larger size typically

travel at a faster rate than particles less dense or smaller.

d2 = 900 nm, v2 = 810,000 k2, where k2 = (p2 − L)g/18n



d1 = 100 nm, v1 = 10,000 k1, where k1 = (p1 − L)g/18n

If we assume that the particle density remains constant, then k2 = k1, and the ratio (r)

between the velocity of sedimentation will be r = v2/v1 = 81, thus, particle of 900 nm

diameter, sediment 81 times faster than 100 nm diameter particle.

Meanwhile, Han et al. (2009) evaluated the effect of incubation time on the size of the

complexes prepared in NaCl and found that the size of the complexes increases with the

incubation time going from 750 nm (30 min) to 1750 nm (120 min). Han et al. (2009)

postulated that the complexes must be stable enough not to dissociate in the cytoplasm but

to dissociate in the nucleus for transfection and they also found that when the stability of

the complexes is increased, the efficiency of endocytosis/phagocytosis is decreased [36].

Complex formation time was also evaluated. An increase from 10 to 120 min, reduced the

transfection efficiency from 40 to 13%. They also found that the total apparent activity of

both gene transcription and protein synthesis was negatively affected by complex

formation time. That is, the longer the complex formation time, the greater the particle size,

the less transfection efficiency [36].

Ogris et al. (1998) found that smaller particles (60–120 nm) which compacted too

strongly to start transcription were formed in the absence of NaCl [38]. They found that

intracellular complexes were dispersed mainly within 48 h after transfection and

anticipated that high stability endocytosed complexes are likely to cause a delay in gene

expression [38]. In our work, we prepared the PEI/DNA complexes in distilled water and

found that the particle size (approximately 150 nm) and the zeta potential (greater than 30

mV) remained constant with the increase in PEI. Regarding the size of the particles, they

are involved in the selection of the absorption pathways of non-viral genetic complexes, as

well as the surface charge of the particle, the shape of the particle, the type of cell, and even

the condition of the culture. PEI/DNA complexes with sizes less than 500 nm are mainly

taken by CME (Clathrin-mediated endocytosis) and CvME (caveolae-mediated

endocytosis), while PEI/DNA complexes with sizes greater than 500 nm are mainly



internalized by the macropinocytosis pathway [39,40]. In COS-7 cells, von Gersdorff et al.

(2006) found that transfection with PEI was mediated by the clathrin pathway. For HUH-7

transfection with linear PEI occurs mainly by the clathrin-dependent pathway, whereas for

branched PEI transfection was mediated by CME and CvME. The same occurred for HeLa

cells, where both pathways mediated transfection, with the caveolae pathway being the

most efficient [41]. In CHO-1 and Hela, Hufnagel et al. (2009) found that for particles of the

order of 500 nm, macropinocytosis played an important role in the absorption of the

complexes [40].

We obtained particles of approximately 150 nm, however, the polydispersity values

(PDI) varied between 0.3 and 0.5, which indicates that the size of the particles is highly

heterogeneous and therefore, several pathways may be intervening in the entrance of the

complexes to the cell. In summary, the size of the complexes can define the cell entry

mechanism, as well as its disassembly and subsequent entry into the nucleus. Additionally,

size is also associated with the stability of the complexes. Smaller sizes lead to the formation

of more stable complexes that take time to disassemble, which could favor the transfection

of cells such as WJ-MSC that have a lower proliferation rate than cell lines. Finally, in the

case of WJ-MSC, it is necessary to assess whether the transfection is dependent on the cell

cycle, since this dependence is a function of the cell type evaluated. Further studies should

evaluate the dependence of WJ-MSC transfection on the cell cycle, since several authors

have reported that the relation between transfection efficiency and cell cycle is cell

dependent, and there are reports were authors there were found a correlation [42-44], while

others have found that there is no dependency [36,43,45,46]. Additionally, it is necessary to

evaluate the growth rate of WJ-MSC, as well as the effect of concentration on transfection

efficiency and select conditions that optimize transfection efficiency—since PEI has been

reported to be toxic [47-55]—and it is necessary to find the conditions that maximize

transfection efficiency and at the same time have the least effect on cell viability.



2.3.2 Evaluation of cell toxicity induced by PEI complexes

Cell type and source are some of the most critical factors impacting on transfection

efficiency. The growth kinetics of WJ-MSC and the effect of seeding density (C0) were first

assessed in order to standardize the transfection conditions of WJ-MSC. Results evidence

that increasing 4.5 times the seeding density (from 2000 to 9000 cells/cm2) produces a

decrease in the duplication rate at the first 19 h (Figure 2-3Figure 3-1a). When the seeding

concentration was 2000 cells/cm2 (approximately 5% confluence), exponential growth was

observed with a doubling time of 40.5 h. However, at an initial cell concentration of 9000

cells/cm2, cell doublings were observed at 19 h, 33 and 50 h. Thus, increasing of cell seeding

density reduce duplication rate, therefore, 9000 cells/cm2 was used for further assays.

Likewise, the addition of PEI complexes into culture was fixed after 24 h of seeding, while

the evaluation of transfection was conducted 24–48 h later to ensure that cells had divided

at least once. In this way, at least one WJ-MSC generation is expected to be formed in the

presence of PEI complexes, facilitating its entry into the cell, assuming that transfection of

the MSC was mitosis-dependent.

Next, the effect of PEI and PEI/DNA complexes on the metabolic activity of WJ-MSC

was evaluated using the resazurin assay technique. There were no statistically significant

differences (p < 0.05) between the cell viability of WJ-MSC treated with PEI alone and

PEI/DNA complexes (Figure 2-3b). This result suggests that the toxicity is caused both by

the PEI that forms complexes with DNA, and by the PEI that remains free in the culture

medium.

Additionally, the metabolic activity of WJ-MSC was reduced with the increase in PEI

concentration. WJ-MSC treated with 1440 and 1800 ng of PEI in DW and PEI complexed

with DNA (at 28 and 35 N/P ratios, respectively) reduced its viability to 50 and 70%,

respectively, compared to untreated WJ-MSC. Figure 3c corresponds to bright field

photographs of WJ-MSC used to evaluate metabolic activity of cells after treatment with

PEI and PEI/DNA complexes (Figure 2-3b). Metabolic activity was related with cell

viability. According to Figure 2-3b, cell viability of WJ-MSC treated with 720 and 1440



ng/well of PEI was 92.6 ± 4.6 and 53.3 ± 2.3%, respectively. Some morphological changes in

the cells were observed with the increase in PEI content, accompanied by the appearance

of small black dots, which probably correspond to cellular debris. In PEI-treated wells there

are fewer cells attached to the plate, some cells begin to leave their elongated shapes and

become circular.

Figure 2-3. Transfection of WJ-MSC with PEI. (a) Effect of seeding density(C0) on

the growth kinetics of WJ-MSC. (b) Cell viability of 4.5 × 104 WJ-MSC/200 µL after

48 h of treatment with PEI and PEI/DNA complexes at different N/P ratios. (c)

Bright-field photographs of non-treated WJ-MSC (without PEI) and treated with

720 and 1440 ng of PEI/well after 24 h. Scale bar: 500 µm. (d) Percentage of WJ-

MSC expressing green fluorescent protein (GFP) by flow cytometry (y-axis, left)

related with the percentage of viable cells in the same assay (y-axis, right). (e,f)

Fluorescent microscope images of transfected WJ-MSC (48 h post-transfection).

Scale bar: 1000 µm. (a–c) denotes significant differences (n = 3 biological replicates,

p < 0.05) in comparison with all groups. The error bars represent 1 SD.

Culture time (h)

0 24 48 72 96 120 144 168

Nu

mb

er o

f W

J-M

SC

/cm

2

0

2x104

4x104

6x104

8x104

C0=2000 WJ-MSC/cm2

C0=9000 WJ-MSC/cm2

PEI content in ng

(N/P ratio)

90 (

1.7)

180

(3.5

)

360

(7)

540

(10.

5)

720

(14)

900

(17.

5)

1440

(28

)

1800

(35

)

WJ-

MS

C v

iab

ilit

y (

%)

0

20

40

60

80

100

120

140

PEI

PEI/DNA

PEI content (ng)/well

(N/P ratio)

Co

ntr

ol

18

0 (

3.5

)

36

0 (

7)

72

0 (

14

)

10

80

(2

1)

14

40

(2

8)

18

00

(3

5)

WJ-

MS

C G

FP

po

siti

ve

(%)

0

20

40

60

80

100

Via

ble

cel

ls (

%)

0

20

40

60

80

100

Viable cells (%)

1440

ng

PE

I72

0 n

g P

EI

Wit

ho

ut

PE

I

d

a c

f

500 µm

500 µm

500 µm

b

e



Free PEI contributes to cell toxicity and—considering that these free molecules drive a

faster and more efficient cellular internalization of polyplexes and contribute to the

subsequent intracellular trafficking [56]—it is necessary to establish a balance between cell

toxicity and transfection efficiency. In this regard, the percentage of WJ-MSC treated with

PEI/DNA complexes expressing green fluorescent protein was quantified at day two post-

transfection by flow cytometry (Figure 2-3d).

Figure 2-3d reported separately, the percentage of cells GFP positive (y axis, left) and

the number of viable cells (y axis, right) as a function of PEI content. In this figure, it is

possible to observe that maximum transfection obtained was 63%; however, at this

condition cell viability was only 20%. Additionally, in Figure 2-3d data on transfection

efficiency (y) and PEI concentration (x) show a trend, closely described by a second order

polynomic equation y = 1.3897E − 05x2 + 8.5976E − 03x + 8.3111E − 01, R² = 0.996. (Figure 2-

S1, Supplementary materials). Additionally, when the cell viability data, measured by both

flow cytometry and resazurin assay technique are analyzed; in both cases, the percentage

of viable cells (y axis) declined following a lineal pattern starting from N/P ratio (x) of 3.5

(180 ng of PEI) up to N/P ratio of 35 (1800 ng of PEI), though there are slight variations

depending on the used technique (Figures 2-S2 and 2-S3, Supplementary materials).

In mathematical terms, it is possible to find an equation to maximize transfection by

multiplying both equations (Table 2-S1, Supplementary materials), in order to obtain the

number of viable GFP positive cells as a function of PEI content, then differentiating such

function and equating to zero to find the maximum. Accordingly, the maximum number

of transfected viable cells would be reached using between 1581 and 1606 ng of PEI;

however, at this point cell viability would be near to 33–40%, as calculated from equations

for flow cytometry and resazurin assay, respectively.

Despite its remarkable DNA condensation ability and transfecting efficiency, PEI also

induces cytotoxicity in a concentration-dependent manner. Recent studies show the

cytotoxic effects of PEI are caused by a series of different mechanisms, mainly disruption

of the different membranes of the cell [57], including the outer cell membrane as well as



internal membranes (i.e., endosome, mitochondria, endoplasmic reticulum (ER), Golgi

apparatus and nuclear membrane) [47-55]. Other mechanisms include affected gene

expression [58]. As a result, cytotoxicity of PEI remains a challenging issue, further

complicated by the lack of clarity related to what PEI’s ultimate fate is, whether it is

exocytosed or degraded (at least partially). This gap in knowledge is of clinical concern.

However, at low enough concentrations, PEI has been cleared by the FDA for applications

as an impregnant in the food-contact surface of regenerated cellulose sheets

(21CFR177.1200); the FDA also approved, in march of 2015, the use of Adherus® AutoSpray

Dural Sealant, manufactured by Hyperbranch Medical Technology, Inc. (Durham, NC,

USA), to prevent cerebrospinal fluid leakage in brain surgery; it contains a mixture of

polyethylene glycol (PEG) ester solution and a polyethylenimine (PEI) solution which the

body absorbs after 90 days [59]. Therefore, finding an appropriate concentration range to

work with this polycationic molecule is essential.

Cautiously, we selected a PEI content of 720 ng (N/P ratio of 14) and used it as the

“optimal” condition for further experiments; this “optimal” is relatively far from the

calculated maximum, reducing the number of transfected viable cells by about 45%, but

this lower content of PEI guarantees a much lower cytotoxicity (cell viability is larger than

80% as measured by resazurin).

Finally, in order to analyze cell morphology of transfected cells, fluorescent microscope

images of WJ-MSC, stained with DAPI and transfected with complexes at N/P ratios of 14

and 21, were analyzed and are shown in Figure 2-3e and Figure 2-3f, respectively. It is

observed that the green fluorescent protein comprises the entire cell and that the

morphology of the transfected cells is not affected by the transfection.

Our results are comparable with those obtained by Wang et al. (2011) and Tierney et

al. (2013) who evaluated the transfection efficiency of PEI (branched, 25 kDa) in MSC from

human bone marrow. Both authors obtained the same highest transfection efficiency of 25%

under different conditions. On the one hand, Wang et al. (2011) used an N/P ratio of eight

and a dose of 6 g DNA/cm2, and calculated the N/P ratio based on the premise that only

primary amines interact with DNA to form complexes and assuming that primary amines



content in PEI is 25%. Therefore, in order to compare their results with ours, the N/P ratio

reported by Wang et al. (2011) was corrected by multiplying it by four. Tierney et al. (2013),

on the other hand, considered all the amines presented in the PEI and used a N/P ratio of

seven and a dose of 1 g DNA/cm2. Wang et al. (2011) and Tierney et al. (2013) used similar

seeding densities (4180 to 5000 cells/cm2) and reported variations in cell viability after PEI

transfection from 60 to 80%, respectively. In our assays, we use the double of the seeding

density (9000 cells/cm2) and we reached transfection efficiencies near to 15% at a N/P of 14

and 0.4 g of DNA with a cell viability of 80%. In comparison, we used 30 times less DNA

than Wang et al. (2011), and five times less DNA than Tierney et al. (2013) to transfect the

same number of cells [25,60].

Considering the work of Mady et al. (2011), where PEI added at a N/P value greater

than 3.5 formed no complexes and remained free in solution, perhaps increasing the

amount of DNA will also likely increase the number of particles and, therefore, the

transfection efficiency. Taking into account that mean hydrodynamic diameter and zeta

potential remained constant at N/P ratios from seven to 35 (Figure 2-2c and Figure 2-2d),

the variation in transfection efficiency is not likely associated with size and charge of the

particles, but rather linked to free PEI that does not complex with DNA. According to Mady

et al. (2011), when an N/P ratio of 3.5 is reached, the additional PEI added is not used in the

formation of complexes, that is, it remains free in solution. This explains why transfection

begins to be observed after a N/P ratio of 3.5. Transfection increased when N/P reached

seven, peaked between 14 and 28, and then decreased when the N/P went up to 35, mainly

due to toxicity [35]. Yue et al. (2011) showed that an excess of PEI in the solution improved

the internalization of the complexes and contributed to the subsequent intracellular traffic

[56]. Likewise, Hanzlíková et al. (2011) found that these free molecules are the ones that

contribute the most to the toxicity during transfection [61]. These results suggest that there

must be enough free PEI to promote transfection, but not that much to affect cell viability.

Thus, optimal transfection of WJ-MSC is reached at a N/P ratio of 14.



In addition to PEI, various non-viral transfection systems have been employed for the

genetic modification of MSC from different sources. The most studied MSC are those from

bone marrow (hBM-MSC). Hoare et al. (2010) transfected hBM-MSC with commercially

available cationic lipid, Lipofectamine (LF) 2000, and achieved transfection efficiencies

between 20 and 40%, with the plasmid: lipid ratio increasing from five to 20, however the

viability decreased from 80 to 50% [62].

To improve the viability and scalability of the agents used for transfection, the use of

molecules such as glucocorticoids (GC) that are steroid hormones that regulate metabolic

activity by binding the GC receptor and translocating it to the nucleus, where the receptor

acts as a transcription factor to modulate gene expression, has been evaluated. Kelly et al.

(2016) combined Lipofectamine-LTX (LF-LTX) with cortisol and dexamethasone and

significantly improved hBM-MSC transfection, increasing transfection efficiency by up to

three times and raising transgene expression by four to 15 times compared to non-

glucocorticoid transfection [63].

Likewise, PEI has been used to improve transfection efficiency in MSC and reduce the

toxicity of complexes. Saraf et al. (2008) modified branched polyethyleneimine (bPEI) with

hyaluronic acid (HA), which is a natural ligand for CD 44, CD 54, and CD 168 receptors in

hMSC, accomplishing an increased transfection efficiency of up to 4.6-fold relative to PEI

alone and a concurrent reduction in toxicity by up to 80% [64]. Similarly, Delyagina et al.

(2014) combined PEI/DNA complexes with biotin-streptavidin-adorned magnetic

nanoparticles (MNP) to transfect hBM-MSC reaching efficiencies of 9.8 ± 7.5%, which are

up to five times greater than those obtained with PEI alone [65].

These methodologies have been tested in MSC from different sources such as bone

marrow and adipose tissue, among others. However, to date, few reports exist of the use

of cationic polymers in the transfection of umbilical cord MSC. Bahadur et al. (2015),

conjugated 1.2 kDa PEI with linoleic acid and hyaluronic acid and evaluated transfection

in MSC of bone marrow (BM) and umbilical cord (UC) and used 25 kDa PEI as a

transfection control. Although they reach transfection efficiencies of up to 40% in UC-MSC,

the viability was reduced up to 60%, making it a methodology that can be improved [29].



Similar results were obtained with the functionalized particles used to transfect UC-MSC,

where Wang et al. (2017) reached a transfection efficiency of 12% with PEI 25 kDa and 14%

with the functionalized particles with a TAT peptide, even though with significant

differences in cell viability, going from 70 to 90%, respectively [29].

Therefore, starting from the methodology presented here, it is possible to further

improve transfection efficiencies using the variations proposed by other authors. While

optimizing transfection, it is necessary to analyze the characteristic properties of MSC such

as immunophenotype, immunomodulatory properties, and differentiation ability in vivo,

considering that one of the goals of gene therapy is to improve or modulate these

characteristics.

2.3.3 PEI Transfection Does Not Affect the Functional Properties of

Wharton’s Jelly MSC

Following the international criteria established by the Society for Cell and Gene

Therapy (ISCT) to define multipotent mesenchymal stromal cells, we evaluated the effect

of PEI transfection on WJ-MSC according to three identity factors: adherence to plastic in

standard culture conditions, specific surface antigen (Ag) expression, multipotent

differentiation potential (osteoblasts and adipocytes) demonstrated by in vitro cell culture

staining [2] and one parameter of biological activity: the immunomodulatory properties.

WJ-MSC transfected with PEI/DNA at an N/P ratio of 14 were evaluated. First, the presence

of characteristic surface cell markers was analyzed by flow cytometry. WJ-MSC expressed

no hematopoietic markers (CD34, CD45, and HLA-DR) in both transfected (T) and non-

transfected (NT) cells (≤2%, Figure 2-4a). The presence of identity markers for MSC (CD73,

CD90, and CD105) was positive (≥95%) in both transfected and non-transfected WJ-MSC.

The immunomodulatory capacity of WJ-MSC was evaluated as the ability to inhibit

the proliferation of activated peripheral blood mononuclear cells (PBMC), measured as the

number of CD3+ cells (specific marker for T lymphocytes). Figure 2-4b shows the number



CD3+ cells for each evaluated condition. PBMC activated and co-cultured with transfected

WJ-MSC (Act co-cul T) or non-transfected WJ-MSC (Act co-cul NT) were compared with

PBMC non-activated (NAct). The ratio between them was expressed such as the fold

change (FC) and was compared with the FC of PBMC activated (Act) and non-activated

(NAct), used as the control. Figure 2-4b shows proliferation of activated PBMC in co-

culture is inhibited by the presence of WJ-MSC. This inhibition in the growth of PBMCs is

associated with the immunomodulatory capacity of WJ-MSC. Given that the inhibition of

activated PBMC proliferation occurred, both in the co-cultures of the transfected and non-

transfected WJ-MSC, these results suggest that the immunomodulatory capacity of the WJ-

MSC was not affected by the transfection with PEI. Finally, we evaluated the impact of PEI-

mediated transfection in the multipotent differentiation towards adipogenic and

osteogenic lineages of WJ-MSC. The evaluation was performed in both transformed (T) and

non-transformed (NT) cells. For adipogenic differentiation, we observed the formation of

lipid vacuoles by staining with Oil Red in both experimental groups (Figure 2-4c),

indicating the preservation of adipogenic potential after PEI transfection. Likewise,

osteogenic lineage differentiation was evaluated using alizarin red-S. We observed the

formation of calcium deposits in both T and NT WJ-MSC, as shown in Figure 2-4c. Then,

we confirmed that MSC subjected to PEI transfection did not lost differentiation potential

and immune-modulatory effects, suggesting PEI-based gene transfer as a safe method for

MSC gene modification.



Figure 2-4. Transfection with PEI did not affect the functional properties of WJ-MSC. (a)

Percentage of WJ-MSC expressing characteristic surface cell markers of mesenchymal

stromal cells. (b) Immunomodulatory potency of WJ-MSC over peripheral blood

mononuclear cells (PBMC). PBMC activated (Act) with CD2, CD3 and CD28 antibodies

were co-cultured with non-transfected (Act co-cul NT) and transfected (Act co-cul T) WJ-

MSC. Non-Activated (NAct) PBMC were used as controls. Data were expressed as

absolute CD3+ cell counts for each condition. Fold change (FC) was expressed as the ratio

between PBMC activated and non-activated. (n = 3 biological replicates, p < 0.05) in

comparison with all groups. The error bars represent 1 SD. (c) Adipogenic (scale bar: 100

μm) and osteogenic (scale bar: 400 μm) differentiation of transfected (T) and non-

transfected (NT) WJ-MSC with PEI at an N/P ratio of 14 (720 ng PEI).

PBMC

Act

Act

co

-cu

l N

T

NA

ct

Act

Act

co

-cu

l T

NA

ct

CD

3+ cel

ls

0

200x103

400x103

600x103

800x103

1x106 FC:2.7FC:2.9

FC:0.5

FC:0.6

NT

T

Control Adipogenic Osteogenicc

a b

Cell surface markers

CD

34

CD

45

HL

A-D

R

CD

73

CD

90

CD

105

HL

A-A

BC

Per

cen

tag

e o

f W

J-M

SC

0

20

40

60

80

100

120 NT

T

400 µm100 µm100 µm

400 µm100 µm100 µm



2.4 Conclusions

The overall aim of this study was to evaluate the efficacy of polyethylenimine (PEI) as

a non-viral gene delivery system that can be optimized for gene therapy in human

mesenchymal stromal cells isolated from umbilical cord Wharton’s jelly. Transfection

efficiency achieved with our methodology gave a similar performance, using a lower

quantity of DNA, compared to the results reported by other authors. Additionally, our

results showed that WJ-MSC transfected with PEI retained their morphology, plastic

adherence, immunophenotype, immunomodulatory function, and multi-lineage

differentiation potential. Thus, PEI can be used as a transfection system in applications for

therapeutic purposes. This improved methodology for transfection of WJ-MSC, based on

PEI, has great potential for tissue engineering applications given the attractive

differentiation capacity of MSC. Since engineered tissue constructs are made of three-

dimensional structures, the next stage of the present work is to incorporate PEI-based

transfection strategies to a three-dimensional scaffold that emulates the characteristics of

the extracellular matrix. In this scenario, gene-activated three-dimensional scaffolds would

serve as support for WJ-MSC, thus enabling gene transfer of inductive factors to favor

osteogenic, adipogenic or chondrogenic differentiation to improve tissue regeneration.

2.5 Supplementary materials

Transfection efficiency (y) and PEI concentration (x) followed a trend, closely described by a

second order polynomic equation y = 1.3897E-05x2 + 8.5976E-03x + 8.3111E-01, R² =0. 996. (Figure 2-

S1)



Figure 2-S1. Second order polynomic patter of transfection by flow cytometry.

Analyzing cell viability data, measured by both flow cytometry and resazurin assay technique;

in both cases, the percentage of viable cells (y axis) declined following a lineal pattern starting from

N/P ratio (x) of 3.5 (180 ng of PEI) up to N/P ratio of 35 (1800 ng of PEI), though there are slight

variations depending on the used technique, as shown in figures 2-S2 and 2-S3:

Figure 2-S2. Linear patter of cell viability by flow cytometry.

y = 1,3897E-05x2 + 8,5976E-03x + 8,3111E-01

R² = 9,9641E-01

0

10

20

30

40

50

60

70

80

90

100

0 360 720 1080 1440 1800

WJ-

MS

C G

PF

po

siti

ve

(%)

PEI content (ng)

y = -0,035x + 88,069

R² = 0,9936

0

10

20

30

40

50

60

70

80

90

100

0 360 720 1080 1440 1800

Cel

l v

iab

ilit

y (

%)

PEI content (ng)

Flow cytometry



Figure 2-S3. Linear patter of cell viability by Resazurin assay technique.

In mathematical terms, it is possible to find an equation to maximize transfection by

multiplying both equations, in order to obtain the number of viable GFP positive cells as a function

of PEI content, then differentiating such function and equating to zero to find the maximum, as

shown in the table 2-S1.

Table 2-S1. Maximization of transfection and cell viability.

Transfection efficiency by

Flow cytometry

y = 1.3897E-05x2 + 8.5976E-

03x + 8.3111E-01, R² =0. 996

Value of x (PEI

content) when

equation is equal to

zero (xmax)

Viability by flow

cytometry

(y = -0.035x + 88.069,

R² = 0.9936)

Function to maximize:

-4.86E-07x3+9.23E-

04x2+7.28E-01x+7.32E+01

Derivative of the

function to maximize:

-1.46E-06x2+1.85E-

03x+0.73

xmax = 1581 ng of PEI

(the other value of x is

negative, having no

physical meaning)

Cell viability calculated

at xmax = 33%

Transfection efficiency

calculated at xmax = 49%

Viability by

resazurin

(y = -0.0418x +

106.82, R² = 0.9866)

Function to maximize:

-5.81E-07x3+1.12E-

03x2+8.84E-01x+8.88E+01

Derivative of the

function to maximize:

-1.74E-06x2+2.25E-

03x+0.88

xmax = 1606 ng of PEI

(the other value of x is

negative, having no

physical meaning)

Cell viability calculated

at xmax = 40%

Transfection efficiency

calculated at xmax = 50%

y = -0,0418x + 106,82

R² = 0,9866

0

20

40

60

80

100

120

0 360 720 1080 1440 1800

Cel

l v

iab

ilit

y (

%)

PEI content (ng)

Resazurin assay technique



Accordingly, table 2-S1, the maximum number of transfected viable cells would be reached

using between 1581 and 1606 ng of PEI; however, at this point cell viability would be near to 33 –

40%, as calculated from equations for flow cytometry and resazurin assay, respectively.

Author Contributions: Conceptualization, methodology and validation, A.I.R.-M., G.S. and R.D.G.-

S.; formal analysis, A.I.R.-M.; investigation, A.I.R.-M., E.R., K.B., C.R.; resources, G.A.S., B.C. and

R.D.G.-S.; writing—original draft preparation, A.R.; writing—review and editing, A.I.R.-M., K.B.,

G.A.S., R.D.G.-S.; visualization, A.I.R.-M.; supervision, project administration and funding

acquisition G.A.S., R.D.G.-S. and B.C. All authors have read and agreed to the published version of

the manuscript.

Funding: This research was funded by Universidad Nacional de Colombia under Grant

203010026990, the Ministry of Science, Technology, and Innovation (MINCIENCIAS) through the

Doctoral Scholarship Program 567-2012 and grant BPIN2012000100186 from Fondo de Ciencia,

Tecnología e Innovación, Sistema General de Regalías, Colombia.

Acknowledgments: A.I.R.-M. thanks Margareth Patiño and Leslie V. Sanchez-Castillo for their help

in E. coli DH5-α transformation and plasmid extraction, respectively, and thanks to Alejandra García

Herrera who provided language help and writing assistance of the article.

Conflicts of Interest: The authors declare no conflict of interest.

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mesenchymal stem cells. Biomacromolecules 2008, 9, 818-827, doi:10.1021/bm701146f.

65. Delyagina, E.; Schade, A.; Scharfenberg, D.; Skorska, A.; Lux, C.; Li, W.; Steinhoff,

G. Improved transfection in human mesenchymal stem cells: Effective intracellular

https://www.accessdata.fda.gov/cdrh_docs/pdf13/P130014b.pdf



release of pDNA by magnetic polyplexes. Nanomedicine 2014, 9, 999-1017,

doi:10.2217/nnm.13.71.

3. Non-viral gene activated matrices based on

human plasma for mesenchymal stromal cells

derived from human umbilical cord Wharton’s

jelly

Ana Isabel Ramos-Murillo 1,2 , Ingrid Silva Cote2, Leslie V. Sánchez2, Bernardo

Camacho2, Gustavo Salguero2 and Rubén Darío Godoy-Silva1,*

1 Chemical and Biochemical Processes Research Group, Department of Chemical and

Environmental Engineering, Faculty of Engineering, Universidad Nacional de Colombia,

111321, Bogotá D.C., Colombia.

2 Advanced Therapies Unit, Instituto Distrital de Ciencia, Biotecnología e Innovación en Salud

(IDCBIS), 111611, Bogotá D.C., Colombia.

* Correspondence: [email protected]; Tel.: +57-1316-5000 (ext. 14307)

Abstract: Gene therapy using non-viral vectors in a tridimensional scaffold is an effective

approach to overcome the shortcomings of protein delivery of differentiation factors in

human mesenchymal stromal cells (hMSC). In this work, a gene activated matrix based on

human plasma cryoconcentrated (HPCC) was prepared using polyethyleneimine (PEI) and

plasmid DNA (pDNA) encoding for green fluorescent protein (GFP). For this purpose,

complexes were formed by mixing PEI with pDNA at the same mass ratio, varying its final

content into the scaffold, and cytotoxicity and transfection efficiency were evaluated.

HPCC scaffolds were prepared at different fibrinogen concentrations. The effect of

fibrinogen content in the inner structure of the scaffolds was evaluated by scanning electron

https://orcid.org/0000-0001-9578-7480

https://orcid.org/0000-0002-5018-9927

https://orcid.org/0000-0001-6064-3743



microscopy (SEM), and its effect on transfection efficiency was also evaluated. Transfection

efficiency increases with the content of DNA and PEI in the scaffold but decreases with

increasing concentration of fibrinogen in the scaffold for WJ-MSC. Together, these findings

suggest that HPCC gene-activated scaffolds are an attractive gene delivery system with

significant potential for clinical applications.

Keywords: Gene activated matrix, non-viral transfection, mesenchymal stromal cells

(MSC), Green-Fluorescent Protein (GFP), polyethylenimine (PEI), Wharton’s Jelly, fibrin,

scaffold.

3.1 Introduction

Every day in the world, millions of surgical procedures are carried out, with the aim

of replacing or repairing tissues that have been damaged as a result of diseases or accidents

[1]. Currently, the standard treatment for this type of condition consists in the

transplantation of healthy tissue from one site to another in the same patient (autograft) or

from another individual of the same species (transplant or allograft). This treatment,

however, has different limitations. On the one hand, autografts require a double surgical

intervention (one to remove healthy tissue and another to graft it at the site of the lesion),

which, apart from being expensive, is painful for the patient and can often lead to infections

or bruises [2].

On the other hand, the use of allografts carries the risk of rejection of the tissue or organ

by the patient, who usually requires the permanent use of immunosuppressive

medications. Another problem associated with allografts is the low availability of organs

and tissues to meet their demand. According to figures from the Ministry of Health and

Social Protection of Colombia, in 2018, 410 (effective) donors were presented, which

allowed 1013 transplants to be performed. However, by November 30th of that same year,

2,500 Colombians were still on the waiting list for a transplant [3].

Chapter 3. Tridimensional transfection of WJ-MSC into a HPCC scaffold 77


The growing need for tissues and organs for replacement and/or repair gave rise to

what is known today as Tissue Engineering (TE), which comprises a number of areas of

knowledge that allow the combination of cells with biologically active molecules in a

porous biomaterial (scaffold) that acts as a scaffold for the regeneration and orientation of

the formation of new tissue [1,4-6].

Different cells are used in TE, according to the tissue to restore. Mesenchymal stromal

cells (MSC) are a type of cells widely used in TE because of their ability to differentiate into

specific lineages such as chondrogenic, adipogenic, or osteogenic [7-9]. MSC are obtained

from different sources such as bone marrow [10-12], adipose tissue [13,14], dental pulp [15-

17], amniotic fluid [18,19], placenta [20,21] , umbilical cord blood [22-25] , umbilical cord

Wharton´s jelly [26-28], synovium and synovial fluid [29-31], endometrium [32,33], among

others [34-36]. MSC from the Wharton’s Jelly of umbilical cord (WJ-MSC), which is

considered a biological waste, have attracted the interest of researchers not only because

they are easy to obtain, but also because they display a higher proliferation rate and low

senescence at later passages compared to stromal cells obtained from other sources [26,28].

Scaffolds for TE must be biodegradable [37], biocompatible, and must favor interaction

with cells [1]. Different biomaterials are used in TE according to the tissue to repair. Fibrin

is a natural biomaterial that provides a similar structure to that of the native extracellular

matrix. Scaffolds based on fibrin are used in several TE applications, which include skin

[38,39], cardiac/vascular [40-42], musculoskeletal and nervous [43,44], among others. An

important source of fibrin is that obtained from autologous blood, whose cost is reduced,

and the risk of viral transmission is absent [45].

Finally, the third component of TE are biologically active molecules that can be

supplied by gene therapy. In 1996, Fang et al. conceived the idea of a biodegradable scaffold

containing cells and a gene delivery vector [46], and called this structure gene activated

matrix (GAM).

Gene therapy is an experimental technique used to introduce foreign genomic

materials (RNA or DNA) into host cells to prompt a therapeutic benefit. [47]. Gene delivery



vectors are molecules that mediate the transfer of genetic material into the nucleus of the

target cell, where the transfected gene is transcribed in parallel with genomic DNA[47,48].

Gene delivery systems can be classified into viral, non-viral, and hybrid systems. Viral

methods consist in modified viruses with deficient replication but able to deliver genetic

material for expression. Note that natural viruses act as cellular parasites, taking advantage

of the existing cellular machinery of the host cell to propagate their own genetic material.

If the viral genome is manipulated to replace pathogenic sequences with specific genes,

modified viruses can be used to introduce ARN o DNA into a target cell.

Some examples of viral systems vectors are adenoviruses[49-51], adeno-associated

viruses[52,53], retroviruses, [54-56] and lentiviruses [57-60]. Viral methods offer higher

transduction efficiency and long-term gene expression. However, there are some

limitations that restrict the use of these systems because they may be associated with

immunogenicity, mutagenicity, poor target cell specificity, inability to transfer large size

genes, toxicity, and high costs [61-63].

Non-viral methods can be divided into physical and chemical. Physical methods utilize

mechanical and electrical forces to induce the transient opening of the cell membrane for

transfection and are easy to develop. However, these methods need special instruments,

and nucleic acids are vulnerable and work better for adherent cells [64]. Physical methods

include microinjection, needle injection, jet injection, gene gun, electroporation,

sonoporation, hydrodynamic gene transfer and mechanical massage, among others. In

contrast, the chemical approach comprises the use of cationic lipids (liposomes), cationic

polymers, and inorganic nanoparticles. Chemical vectors form condensed complexes with

negatively charged DNA through electrostatic interactions. These complexes protect DNA

from degradation and facilitate cellular absorption and intracellular delivery. Furthermore,

they are easy to scale and have low stimulation of the immune system. Likewise, there is a

wide variety of polymeric materials that can be modified to modulate transfection

efficiency and toxicity [61-63]. A widely known chemical vector is branched

polyethylenimine, which has been used to mediate gene delivery into mesenchymal stem

cells of human bone marrow with successful results [65].



This chapter comprises the development of a gene activated matrix for TE, composed

of a scaffold of human plasma cryoconcentrated, combined with mesenchymal stromal

cells from Wharton´s Jelly and a non-viral vector formed by polyethylenimine and plasmid

DNA. The HEK-293 cell line was used as a transfection control.

3.2 Materials and methods

3.2.1 Subsection

Growth media Dulbecco's Modified Eagle Medium (DMEM) (Gibco, Life Technologies

Corp, USA) and Fetal Bovine Serum (FBS) (Biowest, USA and Sigma Aldrich, USA) were

employed for HEK-293 cells and WJ-MSC culture. Polyethylenimine (PEI) branched (25

kDa) from Sigma (USA) was employed in all transfection assays.

3.2.2 Human plasma cryoconcentrated (HPCC)

Three units (~27 mL/unit) of frozen Human Plasma Cryo-Concentrated (HPCC) (Rh

O+) were provided by the local blood bank of the Instituto Distrital de Ciencia,

Biotecnología e Innovación en Salud - IDCBIS, Bogotá D.C., Colombia. The units were

completely thawed in a thermostat bath at 37°C and mixed under sterile conditions to

create a single lot, therefore, the variability in outcomes attributed to differences in plasma

among donors was reduced. The HPCC mixture was aliquoted (7 mL per tube) and stored

at -20°C to avoid repeated freeze-thaw cycles. Levels of fibrinogen in HPCC were

quantified by an external laboratory using the Clauss method [66,67] (Supplementary

Figure 3-S1).

3.2.3 Expansion of WJ-MSC and HEK cells

WJ-MSC were obtained from umbilical cord donors (number 40 and 148) of the

Advanced Therapies Unit at the IDCBIS, following informed consent from the mothers.

WJ-MSC at passage 1 to 6 were cultured in low-glucose DMEM supplemented with 10%



FBS at 37 °C and 5% CO2. Human embryonic kidney cells 293 (HEK-293) were cultured in

T-75 flasks with high glucose DMEM supplemented with 10% FBS.

WJ-MSC were transduced in our laboratory with a GFP plasmid to obtain a stable cell

line that expresses endogenous green fluorescent protein. In this study, these cells were

named GFP-WJ-MSC.

3.2.4 Culture of GFP-WJ-MSC over HPCC hydrogels at different

fibrinogen concentrations

The cell growth of GFP-WJ-MSC into HPCC scaffolds at different fibrinogen

concentrations (0 to 3.4 µg/µL) was evaluated. Three different scaffold thicknesses (1, 2,

and 3 mm) were also analyzed to establish the best culture conditions. The final

concentration of GFP-WJ-MSC into the HPCC gel was 3.6x106 cells/mL. The detailed

procedure to prepare HPCC gels with embedded GFP-WJ-MSC is presented in Figure 3-1.

First, GFP-WJ-MSC were added into the well in a proportion of 50% v/v, then, variable

volumes of DMEM (10% v/v FBS) and HPCC supplemented with 270 mM CaCl2 were

added into the well (variable rate between 0-50% v/v) to complete the final volume. For

example, the highest fibrinogen concentration evaluated was 3.4 µg/µL. In the 24-well plate

format, the total volume was 200 µL. This gel was formed by mixing 100 µL of HPCC-Ca

with 100 µL of cell stock (7x105 GFP-WJ-MSC per mL). HPCC-Ca is the solution formed by

HPCC concentrated (~7.5 µg/µL) with CaCl2 270 mM (in DMEM) in a 10:1 proportion to

reach a final concentration of ~6.8 µg of fibrinogen per µL of solution. Therefore, the final

composition of the scaffold was 70.000 GFP-WJ-MSC and 680 µg of fibrinogen in 200 µL,

leading to a final concentration of 3.5x105 cells/mL and 3.4 µg of fibrinogen/µL. Detailed

calculations for each fibrinogen concentration and well-plate format are presented in the

Supplementary Figure 3-S2

Gel formation occurred at room temperature within 5 to 10 s. Nevertheless, HPCC gels

with cells were further incubated for 30 min at 37 °C and subsequently scaffolds were

covered with 400 µL of DMEM (10% v/v FBS) supplemented with 0.5% v/v tranexamic acid

(Tranexam® injectable solution 100 mg/mL, Ropsohn Therapeutics Ltda., Colombia). Cell



growth was carried out at 37°C and 5%CO2, and scaffolds with embedded cells were

monitored in days 0, 1 and 2 by fluorescence microscopy.

Figure 3-1. General procedure to elaborate HPCC scaffolds with WJ-MSC

0.64

0.1

0.32

0.31

HPCC + CaCl2 3%w/v (10:1) DMEM + FBS (9:1)

GFP WJ-MSCs (7.2x106 cells/ml)

24-well plate 48-well plate 96-well plate

The proportion of the cell stock (50%) remained

constant and the volume of HPCC and DMEM was varied

to modify the concentration of fibrinogen in the gel.

Cells were incubated for 30 minutes at 37 ° C and

subsequently 400 ul of DMEM 10% v/v FBS

supplemented with tranexamic acid (0.5mg/ml) was

added to each well.

The growth of the cells on the gel was monitored for 3

days and the monitoring was performed by fluorescence

microscopy every day.

1GFP WJ-MSCs were added

to the well (50%v/v)2

HPCC-CaCl2 with variable fibrinogen

concentration was added to the well

(variable rate between 0-50% v/v)

3Gel formation occurred at room

temperature almost instantaneously.

4The same procedure was repeated

in each well to obtain 12 different

growing conditions.

HPCC

scaf

fold

Diameter (cm)

Volume (ml or cm3)

area (cm2)

h (cm)

1.1

0.2

0.95

0.21

1.56

0.2

1.91

0.10



3.2.5 Scanning Electron Microscopy (SEM) of HPCC hydrogels

The effect of fibrinogen concentration in HPCC and cell growth of GFP-WJ-MSC on

the distribution of the fibrin fibers in the scaffold was evaluated by scanning electron

microscopy (SEM). Four different concentrations of fibrinogen (1.5 to 3.3 µg/µL per

scaffold) were analyzed. Scaffolds with GFP-WJ-MSC were cultured for seven days at 37°C

and 5%CO2. For analysis using SEM, scaffolds were cut into 4 pieces and fixed with 4%

w/v paraformaldehyde solution for 20 min, followed by one wash with PBS 1X,

dehydration through seven ethanol series (30%, 50%, 70%, 90%, 96%, and 100%) and critical

point drying with hexamethyldisilane. Samples were then mounted onto aluminum

specimen stubs using adhesive carbon tape and coated by ion sputtering with conductive

gold set at 5mA for 120 secs (SPT-20, Coxem, Korea). The inner morphology was examined

using a SEM EM30AX Plus (Coxem, Korea) operated at 16kV accelerating voltage.

3.2.6 GFP-WJ-MSC differentiation on HPCC hydrogels

The effect of fibrinogen concentration was evaluated on the differentiation of GFP-WJ-

MSC embedded in scaffolds of HPCC. Additionally, three seeding densities of GFP-WJ-

MSC were analyzed in order to determine the effect of cell concentration on the

differentiation ability of WJ-MSC into the HPCC scaffolds. HPCC gels were prepared into

a 24 well-plate to obtain scaffolds of 1 mm thickness and 15.6 mm diameter. The detailed

procedure is presented in Figure 3-2. Differentiation of GFP-WJ-MSC to adipogenic,

chondrogenic, and osteogenic lineages was evaluated. GFP-WJ-MSC cultured in DMEM

into the HPCC scaffolds were used as a control. Briefly, GFP-WJ-MSC were recovered from

T-flasks by trypsinization and centrifuged at 200g for 6 min. Then, the supernatant was

removed and the cells were resuspended to a concentration of 1.74x106 cells/mL in the

appropriate amount of differentiation medium, i.e. adipogenic induction medium

(StemProTM Adipogenesis Differentiation Kit, GibcoTM by Life Technologies, USA),

osteogenic induction medium (StemProTM Osteogenesis Differentiation Kit, GibcoTM by

Life Technologies, USA) and chondrogenic induction medium (StemProTM Chondrogenesis



Differentiation Kit, GibcoTM by Life Technologies, USA). The cell suspension was mixed

with HPCC-Ca to form gels at three different concentrations of fibrinogen into the scaffold:

1, 2, and 3 µg/µL. Then, 400 µL of the corresponding differentiation media (0.5% v/v

tranexamic acid) were added to each well and gels were incubated at 37°C and 5 %CO2.

The differentiation medium was changed every 3 days for 21 days. Then, scaffolds

were fixed in 4% paraformaldehyde (Sigma Aldrich, St Louis, MO, USA) for 30 min. To

analyze adipogenic differentiation, fixed HPCC gels were washed two times with PBS 1X

and incubated at room temperature with 400 µL of isopropanol 66% v/v for 20 min. Then,

isopropanol was removed and 400 µL of Oil Red O (Sigma Aldrich, St Louis, MO, USA)

0.3% w/v in isopropanol 99% (Sigma Aldrich, St Louis, MO, USA) were added to each well.

After 5 min, Oil Red was removed, and gels were washed 5 times with deionized water to

remove excess dye. The staining of lipid vacuoles was analyzed by bright-field microscopy.

Chondrogenic differentiation was analyzed using Alcian Blue (Sigma Aldrich, St Louis,

MO, USA). After fixation with paraformaldehyde, gels were treated with 400 µL of HCl

0.1N for 20 min at room temperature, then, HCl was removed and gels were stained with

400 µL of Alcian Blue 0.1% w/v (in HCl 0.1 N). After 15 min, Alcian Blue was removed and

gels were washed several times with deionized water until the removed water was

colorless (~10 times). Bright field microscopy was used to analyze the staining of aggrecan,

which is a proteoglycan used as an indicator of cartilage formation.

Gels formed with osteogenic differentiation medium were treated with Alizarin red-S

(Sigma Aldrich, St Louis, MO, USA) to analyze the formation of calcium deposits. Fixed

gels were treated with 400 µL of Alizarin red solution (2% w/v in deionized water). After 5

min at room temperature, the dye was removed, and gels were washed several times (more

than 10) to remove excess dye. GFP-WJ-MSC cultured in DMEM into HPCC scaffolds and

HPCC gels without cells were used as controls.



Figure 3-2. General procedure to differentiate WJ-MSC into HPCC scaffolds

24-well plate

1

GFP WJ-MSCs resuspended

in each differentiation

medium were added to the

well (50%v/v)

4 After 30 minutes, 400 µl of

differentiation medium were added to

the well

2HPCC-CaCl2 was added to

the well (variable rate

between 0-50% v/v)

3Gel formation occurred at room

temperature almost instantaneously.

5The same procedure was repeated in each well to obtain 4 different growing

conditions. Differentiation media evaluated were adipogenic, chondrogenic and

osteogenic. DMEM supplemented with FBS was used as control.

DMEM 10% FBS ADIPOGENIC OSTEOGENICCHONDROGENIC



3.3 Gene activated matrices of HPCC

3.3.1 Plasmid propagation

One ShotTM TOP10 Chemically Competent E. coli cells (Invitrogen, USA) were

transformed with a Human TGFB1 ORF (C-GFP-Spark tag) mammalian expression

plasmid (Sino Biological, USA). Transformed E. coli were cultured in Luria Bertani (LB)

medium (Ref. L3152, Sigma Aldrich, USA) and LB with agar-agar (Ref. A0949, PanReac

AppliChem, Germany), and selected with kanamycin sulfate (Ref. 11815024,

ThermoScientific, USA). GFP plasmid (pGFP) was purified using a ZymoPURETM Plasmid

Maxiprep Kit (catalog #D4203, Zymo Research, Irvine, USA). The DNA concentration and

purity were measured using a NanoDropTM 2000/2000c spectrophotometer

(ThermoScientific®, USA).

3.3.2 PEI and pGFP nanoparticle formulations and physicochemical

characterization

Gene transfection of HEK and WJ-MSC in human plasma cryoconcentrated (HPCC)

scaffolds was carried out at a PEI/DNA mass ratio of 2. The DNA content was varied

between 1 and 5 µg per scaffold for HEK transfection and between 5 and 9 µg per scaffold

for WJ-MSC. The final volume of the complexes was 20 µL per scaffold.

Briefly, pGFP and PEI concentrations were adjusted to always mix the same volume of

each solution. PEI was added to the DNA and the solution was vortexed immediately.

Then, the solution was incubated at 37°C for 30 min to favor complex formation, which was

evaluated by electrophoresis in agarose gel. Afterward, 0.79 g of UltrapureTM agarose

(ThermoFisher Scientific, USA) were dissolved in 100 mL of Tris-acetate-EDTA (TAE)

buffer. A Bio-Rad® power supply was employed to apply 100V through the agarose gel for

30 min. A 100bp marker was used as molecular weight marker (GelPilot 100 bp Plus Ladder

(100), Qiagen, USA) and SYBRTM safe dye (ThermoFisher Scientific, USA) was used as DNA

gel stain.



3.3.3 Transfection of WJ-MSC into HPCC scaffolds

The detailed procedure of transfection of WJ-MSC into the fibrin (from human plasma)

scaffolds is presented in Figure 3-3. Complexes were prepared using equal volumes of PEI

and plasmid as described in 5.5.2. Subsequently, complexes were incorporated into the

HPCC gels as follows. First, 20 µL of complexes were added into the well and mixed with

115 µL of WJ-MSC resuspended in DMEM 10% FBS. Afterward, 165 µL of HPCC-Ca were

added to the respective concentration. HPCC-Ca solution was obtained by mixing HPCC

at an appropriate fibrinogen concentration with 270 mM CaCl2 solution in 10:1 proportion.

The fibrin gel was formed 5-10 seconds after mixing all the previously described

components. Gels were kept at 37°C for 30 min before adding 400 µL of DMEM 10% FBS

(0.5% v/v tranexamic acid), then, the scaffolds were incubated at 37°C and 5% CO2.

Figure 3-3. Methodology developed to prepare GAMs with WJ-MSC and HPCC. (A)

Preparation of the complexes by mixing equal volumes of PEI and DNA solutions in

distilled water. PEI is added to the DNA solution, the solution is vortexed for 10 s and

incubated for 30 min at 37°C. (B) WJ-MSC at 70% of confluence are recovered by

trypsinization and resuspended in DMEM 10% FBS. (C) A few seconds before preparing

the scaffolds, HPCC is mixed with 270 mM CaCl2 solution in a 10:1 proportion (HPCC-Ca).

(D) Complexes are mixed with WJ-MSC into the well plate, then HPCC-Ca solution is

added and mixed gently to avoid bubble formation. Fibrin gel is formed within 5-10

seconds. Then, DMEM 10% FBS and 0.5% tranexamic acid are added to each well (E)

Scaffolds are incubated at 37°C and 5% CO2, and transfection efficiency is evaluated 48 h

later.

A B C D

E

PEI

DNA

PEI/DNA complexes

WJ-MSCs

at 80% of

confluency HPCC

CaCl2 3% w/v

in DMEM

GAM

WJ-MSCs in

DMEM

10% FBS

DMEM 10% FBS and

0.5 % Tranexamic acid



3.3.4 Effect of fibrinogen concentration on the metabolic activity of

WJ-MSC embedded in fibrin scaffolds in the presence of PEI and

PEI/DNA complexes

Fibrin scaffolds with WJ-MSC were prepared following the methodology previously

described. To analyze the effect of fibrinogen concentration on the metabolic activity of WJ-

MSC embedded in fibrin scaffolds in the presence of PEI and PEI/DNA complexes, three

different contents of PEI were loaded into the scaffolds: 5, 10, and 15 µg. Complexes of

PEI/DNA were prepared keeping a PEI/DNA mass ratio of 2 and loaded at the same PEI

contents of 5, 10, and 15 µg, so the DNA content in each scaffold was 2.5, 5, and 7.5 µg,

respectively. After 4 days of culture, the culture medium was removed and fibrin gels were

incubated with 200 µL of Resazurin working solution (1 mL of 440 µM Resazurin sodium

salt, 121519, PanReac AppliChem, Darmstadt, Germany in PBS at pH 7.4 per each 10 mL of

DMEM 10%FBS) at 37°C for 3 h, protected from light. Then, 100 µL of Resazurin solution

was taken from each well and placed in a 96-well plate. Fluorescence intensity (593

emission/535 excitation) was measured and the metabolic activity of cells in HPCC gels was

established by comparison with a standard curve.

3.3.5 Effect of WJ-MSC and PEI/DNA complexes on the rheological

behavior of fibrin gels

Gels of 1 mm thickness and 25mm diameter, containing 18 µg of DNA, 36 µg of PEI,

and 7.2x105 WJ-MSC, were prepared to measure rheological behavior while preserving the

proportions used in the previous assays. Gels were formed by mixing 72 µL of complexes

solution (36 µL of PEI 1 µg/µL at and 36 µL of DNA at 0.5 µg/µL ) with 414 µL of WJ-MSC

in DMEM 10% FBS (1.74x106 cells/mL) and 594 µL of HPCC-Ca (540 µL of HPCC at 6 µg/µL

of fibrinogen and 54 µL of CaCl2 270 mM in DMEM).

HPCC scaffolds without cells and complexes were prepared following the procedure

previously described by exchanging cell solution for DMEM 10% FBS and complexes for

distilled water. In HPCC scaffolds alone, cells and complexes volumes were replaced with



distilled water. The gels were formed on the rheometer plate for 30 min, then the elastic

modulus (Pa) was measured as a function of frequency sweep between 0.1 and 10 Hz.

Measurements of rheological properties were performed using a Bohlin C-VOR 200

rotational rheometer (Malvern Instruments Ltd., England, UK) with a parallel-plate

configuration. The diameter of the rotating plate was 25 mm with a gap thickness of 1.0

mm. All measurements were performed at 37.0 ± 0.1 ºC by means of a Bohlin Peltier

temperature control system fitted to a recirculating water bath. To avoid water evaporation

during the test, the rotor plate was covered with a special solvent trap. HPCC gels were

crosslinked between the plates, as described by Eyrich et al. (2007) [68].

3.3.6 Effect of fibrinogen concentration and DNA content on the

transfection efficiency of HEK cells and WJ-MSC embedded in

fibrin scaffolds

The effect of DNA content on the transfection efficiency of HEK cells and WJ-MSC in

a tridimensional scaffold of HPCC was evaluated. Fibrin gels were prepared as described

in 5.5.3. In separate assays, 4x105 HEK cells and 2x105 WJ-MSC were seeded into scaffolds

of HPCC at three different fibrinogen concentrations (3,2, and 1 µg/µL). Complexes of PEI

and DNA were prepared at five different contents of DNA: 1, 2, 3, 4 and 5 µg/scaffold for

HEK transfection and, 5, 6, 7, 8 and 9 µg of DNA per scaffold for WJ-MSC transfection. The

PEI/DNA mass ratio was kept at 2. The final volume of each scaffold was 240 µL. After gel

formation, 400 µL of DMEM 10% FBS (0.5% v/v tranexamic acid) was added into each well

and cells into the scaffolds were incubated at 37°C and 5% CO2. HEK scaffolds were

incubated for 60 h and WJ-MCSs scaffolds for 48 h.

3.3.7 Statistical analysis

Statistical analyses were performed using Statgraphics Centurion XVI, V. 16.2.04

software. One-way ANOVA was used for analysis of variance with Tukey’s post hoc test

to compare groups. Numerical and graphical results are displayed as the mean ± standard

deviation. Significance was accepted at a level of p < 0.05.



3.4 Results and discussion

Previous studies by other authors have shown that mesenchymal stromal cells from

different sources such as bone marrow [69-71] or adipose-derived [72] are able to proliferate

and differentiate into fibrin scaffolds prepared from commercial products [69-71] or derived

from autologous sources [72].

Likewise, several authors have reported that the concentration of fibrinogen in the

hydrogels affects the internal structure of the gels [73,74] and thus cell growth [75].

Additionally, cell growth and proliferation have been associated with the concentration of

fibrinogen in the hydrogel. In 2003, Bensaïd et al. determined the optimal fibrinogen

concentration (18 mg/mL) to obtain cell proliferation of hMSC from bone marrow in the

resultant fibrin gels prepared using a commercial kit (Tissucol®,Baxter, USA). This

concentration allows good hMSC spreading and proliferation [70]. Later in 2006, Ho et al.

evaluated 10 different fibrin formulations, with variable concentrations of fibrin (50, 34, 35,

17, and 5 mg/mL) and thrombin (1, 125, 167, and 250 U/mL). They found that fibrinogen

concentration, not that of thrombin, has a more dominant impact on hMSC proliferation.

The lowest fibrinogen concentration evaluated (5 mg/mL) was reported to promote the best

proliferation of hMSC [69].

In this study, the use of fibrinogen from an autologous source to prepare gene activated

matrices with Wharton’s Jelly mesenchymal stromal cells (WJ-MSC) was evaluated due to

its low cost, readily obtention and its advantages of safety from transmission of viral

diseases and immunological reactions [76]. For this purpose, human plasma

cryoconcentrated (HPCC) provided by the local blood bank was used to prepare a

homogeneous mixture, which was characterized by blood type, Rhesus factor (Rh) and

fibrinogen content.

3.4.1 Culture of WJ-MSC into HPCC gels

In order to establish the best appropriate culture conditions for WJ-MSC into HPCC

scaffolds, the combined effect of scaffold fibrinogen concentration and thickness on GFP-



WJ-MSC morphology and spreading was evaluated. HPCC gels were prepared of three

different thicknesses (1,2, and 3 mm), keeping constant cell and fibrinogen concentrations.

An increase in gel thickness is related to the appearance of a higher number of rounded

cells in the gel (Figure 3-4). This effect could be attributed to the limitation of transport of

oxygen and nutrients from the medium to the gel and of waste substances and metabolic

products from the interior to the gel, as those are diffusive process [77]. In order to reduce

the diffusional limitations and considering that the increase in the gel depth can hinder the

diffusional process, gels of 1 mm thickness (24-well) were selected for subsequent tests.

Figure 3-4. Effect of scaffold fibrinogen concentration and thickness on GFP-WJ-MSC

morphology and spreading. Fluorescence micrograph images of fibrin hydrogels at (A-C)

3, (D-F) 2, and (G-I) 1 µg of fibrinogen/µL containing 3.6x105 GFP-WJ-MSC/mL, in scaffolds

with different thicknesses: (A,D,G) 1, (B,E,H) 2, and (C,F,I) 3 mm. (J-L) GFP-WJ-MSC

cultured in monolayer were used as controls. Scale bar: 200 µm. Scaffolds were incubated

at 37°C and 5% CO2 for 2 days with DMEM supplemented with 10% FBS and tranexamic

acid (5% v/v).

Additionally, an elongated fibroblast morphology was observed (Figure 3-4J-L),

characteristic of mesenchymal stromal cells [78], as the concentration of fibrinogen in the

gels is decreased; however, this decrease is also related to the formation of less manageable

24

-well

pla

te4

8-w

ell

pla

te9

6-w

ell

pla

te

3 µg/µl 2 µg/µl 1 µg/µl

Scaffold fibrinogen concentration

1 m

m2

mm

3 m

mSca

ffold

thic

kness

0 µg/µl

Monolayer

A

B

C

D

E

F

G

H

I

J

K

L



gels. Consequently, in order to find an optimal cell culture condition, two extreme values

of fibrinogen concentration (low (1 µg/µL) and high (3 µg/µL)) a one intermediate (2 µg/µL)

were selected for the subsequent tests.

3.4.2 Fibrinogen concentration and differentiation media in HPCC

affects GFP-WJ-MSC morphology

Differentiation capacity is one of the most striking features of MSC for use in tissue

engineering. In order to analyze the combined effect of fibrinogen concentration and

differentiation media on morphological changes of the cells embedded in the scaffolds,

three different fibrinogen concentrations (1,2, and 3 µg/µL) were evaluated with WJ-MSC

resuspended in adipogenic, chondrogenic, and osteogenic media. The effect of WJ-MSC

seeding concentration was also evaluated (2,4, and 6x105 cells per scaffold). Morphological

changes of GFP-WJ-MSC grown on HPCC scaffolds, and DMEM, with a final fibrinogen

concentration of 1 µg/µL, on days 1, 10, and 21 of culture, were used as controls (Figure

3-5). Morphological changes in WJ-MSC seeded at 4and 6x105 cells per scaffold, are

presented and evaluated (data not shown).



Figure 3-5. Cell morphology evolution over time of GFP-WJ-MSC embedded in HPCC-

DMEM scaffolds. 2x105 GFP-WJ-MSC were seeded in scaffolds prepared at (A-C) 3, (D-F)

2, and (G-I) 1 µg of fibrinogen/µL. Scaffolds were incubated at 37°Cand 5% CO2 for 21 days

with DMEM supplemented with 10% FBS and tranexamic acid (5% v/v). (J-L) GFP-WJ-MSC

cultured in monolayer were used as controls. The media were changed every 3 days.

Fluorescence images were recorded at (A,D,G,J) day 1, (B,E,H,K) day 10, and (C,F,I,L) day

21. Scale bar: 200 µm.

At day 1, after seeding, morphological differences of GFP-WJ-MSC embedded in fibrin

scaffolds were observed with respect to the monolayer control. The most significant

differences were observed in cells embedded in the gels at the highest concentrations of

fibrinogen evaluated (2, and 3 µg/µL) (Figure 3-5D-F, and A-C, respectively). Figure 3-6

schematically illustrates how cells acquire forms such as elongated spindles. These

elongations become more extensive over time. A slight increase in cell size was also

observed. Regarding the lowest concentration of fibrinogen evaluated (1 µg/µL), an

Day

1D

ay 1

0D

ay 2

1

3 µg/µl 2 µg/µl 1 µg/µl 0 µg/µl


200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

Monolayer

A

B

C

D

E

F

G

H

I

J

K

L



apparent smaller number of cells was observed in days 1, and 10. Cells with wider shapes

were observed in comparison to cells cultured in monolayer (Figure 3-5J-L).

Figure 3-6. Schematic representation of GFP-WJ-MSC morphology embedded in HPCC-

DMEM scaffolds in comparison with monolayer cell culture, elaborated from fluorescence

images of cells from Figure 3-5.

After establishing the morphology acquired by the cells in the gels prepared and

cultured in DMEM, the effect of the differentiation media was evaluated. Cell growth and

morphological changes were analyzed in GFP-WJ-MSC embedded in fibrin scaffolds and

prepared with HPCC and differentiation medium. 2x105 GPF-WJ-MSC were embedded in

HPCC scaffolds prepared at 1 µg/µL of fibrinogen in adipogenic (Figure 3-7D-F),

chondrogenic (Figure 3-7G-I), and osteogenic (Figure 3-7J-L) differentiation media. The

HPCC scaffolds prepared in DMEM and supplemented with 10 %FBS were used as

controls.

Scaffolds were incubated at 37°C, and 5% CO2 for 21 days. Differentiation media

supplemented with tranexamic acid (5% v/v) was changed every 3 days. Fluorescence

images were recorded on days 1, 10, and 21. At day 1, significant differences were observed

between GFP-WJ-MSC cultured in DMEM and those grown in differentiation media

(Figure 3-7A,D,G,J). GFP-WJ-MSC cultured in DMEM were elongated with some bumps

of 500 µm length. This behavior is similar on day 10 of culture. On day 21, very few green

Embedded in HPCC gels Monolayer cell culture



cells were observed in the gel. The presented photograph corresponds to a small portion of

the scaffold in which cells were still visible.

GFP-WJ-MSC growing in adipogenic medium had a polygonal shape (Figure

3-7Figure 3-7D-F). This morphology was more evident on day 10 of cultivation (Figure

3-7D). On day 21, the cells appeared to begin losing fluorescence, and dark patchy areas

were seen on the gel (Figure 3-7F).

Opposite to the morphology acquired in adipogenic media, GFP-WJ-MSC in

chondrogenic medium were more clustered (Figure 3-7Figure 3-7). Cells appear to collect

their cytoplasm and become smaller. Owing to the three-dimensional configuration, it

gives the appearance of being attached but not with an elongated shape. At day 10th, this

initial morphology changed, and the cells began to take a similar shape to those grown in

osteogenic medium on the same day of culture. The difference is that those cells in the

chondrogenic media are less broad and slightly more elongated (Figure 3-7H).

GFP-WJ-MSC cultured in scaffolds with osteogenic medium (Figure 3-7) have

elongated shapes like those cultured in DMEM on day 1 (Figure 3-7A). However, the

appearance of cells with very thin elongated spindles was observed. Other cells are star

shaped. On day 10, the cells appeared to have increased in size and wider shapes were

observed than those of a normal monolayer culture (Figure 3-7K). On day 21, very few cells

were observed in the entire well (Figure 3-7L). Photographs presented in Figure 3-7B, C, F,

I and L correspond to one of the few areas in the well where the cells still had fluorescence.

The appearance of dark spots on the gel and a considerable reduction in fluorescence were

also observed.



Figure 3-7. Cell morphology evolution over time of GFP-WJ-MSC embedded in HPCC

scaffolds. Scaffolds prepared at 1 µg of fibrinogen/µLand 2x105 GFP-WJ-MSC per scaffold

were resuspended in (A-C) DMEM, (D-F) adipogenic, (G-I) chondrogenic, and (J-L)

osteogenic differentiation media. Scaffolds were incubated at 37°C, and 5% CO2 for 21 days.

Differentiation media supplemented with tranexamic acid (5% v/v) was changed every 3

days. Scale bar: 200 µm.

3.4.3 Fibrinogen concentration in HPCC scaffolds does not affect the

differentiation of WJ-MSC

Differentiation of GFP-WJ-MSC cultured in HCPP and adipogenic differentiation

media were evaluated by oil red staining. Formation and staining were analyzed in lipid

vacuoles formed by hMSC embedded in the fibrin scaffolds. Three different fibrinogen

concentrations were evaluated: 3 µg/µL (Figure 3-8A-C), 2 µg/µL (Figure 3-8E-G), and 1

µg/µL (Figure 3-8I-K). Cells cultured in a monolayer (Figure 3-8M) were used as control.

Three seeding densities of WJ-MSC were also evaluated (2,4, and 6 x105 cells per scaffold).

Lipid vacuoles staining was observed in all configurations of fibrinogen and cell seeding

density.

ADIPOGENIC CHONDROGENIC

Day

1D

ay 1

0D

ay 2

1

DMEM OSTEOGENIC

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

A

B

C

D

E

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L



Some red staining was observed in gels used as control, which corresponds to 2x105

WJ-MSC cultured in DMEM-HPCC. At 3 and 2 µg/µL of fibrinogen (Figure 3-8D, and H),

some small red dots were observed. However, in the lowest concentration of fibrinogen (1

µg/µL), cells are observed with elongated shape and red stained (Figure 3-8L).

Differentiation of GFP-WJ-MSC cultured in HCPP and chondrogenic differentiation media

were evaluated by staining with Alcian Blue for specific proteoglycans. This was used to

identify cartilage-specific extracellular matrix (ECM) components, produced by hMSC

embedded in fibrin scaffolds (Figure 3-9).

HPCC gels were evaluated at three different fibrinogen concentrations: 3 µg/µL

(Figure 3-9A , B and C), 2 µg/µL (Figure 3-9E, F and G) and 1 µg/µL (Figure 3-9I, J and

K). Cells cultured in monolayer (Figure 3-9M) were used as controls. All HPCC gels, with

and without cells (Figure 3-9N), were stained with Alcian Blue. No difference in staining

between scaffolds was found. In some gels, it was possible to observe the cell morphology.

Seeding densities of WJ-MSC were evaluated between 2 and 6x105 cells per scaffold. At low

seeding densities (2x105), no differences were observed between gel staining at different

fibrinogen concentrations (Figure 3-9A, E and I). In the 4x105 cells, with 3 and 2 µg/µL of

fibrinogen configurations (Figure 3-9B, and F) no differences were observed either.

However, at fibrinogen configuration of 1µg/µL (Figure 3-9J), the appearance of cells with

a rounded shape is notable. This morphology is similar to that obtained by 6x105 cells per

scaffold in all the fibrinogen concentrations evaluated (Figure 3-9C, G and K)

Although rounded, the morphology of the control cells, which correspond to 4x105 WJ-

MSC cultured in DMEM-HPCC, had no dark areas that appeared in the cells of the gels

prepared in chondrogenic media-HPCC (Figure 3-9D,H, and L).

Finally, differentiation of GFP-WJ-MSC cultured in HCPP and osteogenic

differentiation was evaluated using Alizarin Red staining for the detection of mineralized

matrix and calcium deposits (Figure 3-10). HPCC gels at three different fibrinogen

concentrations were evaluated: 3 µg/µL (Figure 3-10A, B and C), 2 µg/µL (Figure 3-10E,

F and G), and 1 µg/µL (Figure 3-10I, J and K). Cells cultured in monolayer (Figure 3-10M)



were used as control. All HPCC gels, with and without cells (Figure 3-10N), were stained

with Alizarin Red.

Figure 3-8. Adipogenic differentiation of GFP-WJ-MSC embedded in HPCC-Adipogenic

media scaffolds. Adipogenic differentiation was evaluated at three cell seeding densities:



control. Scaffolds prepared at (A-D) 3, (E-H) 2, and (I-L) 1 µg of fibrinogen/µL were stained

with Oil Red dye. (M) GFP-WJ-MSC cultured in monolayer and (N) HPCC scaffolds

without cells were used for differentiation control. Adipogenic differentiation media and


Scale bar: 100 µm.

Even though scaffolds prepared with HPCC without cells completely trapped the dye

and their appearance was completely red, notable differences were found in the scaffolds

with cells embedded in osteogenic differentiation media.

2×

10

5



Monocapa

HPCC without cells

100 µm

100 µm

Seedin

g d

ensi

ty (ce

lls/

scaffold

)

6×

10

5D

MEM

2×

10

54

10

5

CONTROL: 2×105 GFP-

WJ-MSCs in a HPCC

scaffold prepared in

DMEM

Monolayer

100 µm

100 µm

100 µm

100 µm

100 µm

100 µm

100 µm

100 µm

100 µm

A

B

C

E

F

G

I

J

K

M

N

D H L



Figure 3-9. Chondrogenic differentiation of GFP-WJ-MSC embedded in HPCC-

chondrogenic media scaffolds. Chondrogenic differentiation was evaluated at three cell

seeding densities: (A,E,I,M) 2, (B,F,J) 4, and (C,G,K) 6x105 GFP-WJ-MSC/scaffold. (D,H,L)

4x105 GFP-WJ-MSC resuspended in DMEM media and cultured in HPCC-DMEM scaffolds

were used as control. Scaffolds prepared at (A-D) 3, (E-H) 2 and (I-L) 1 µg of fibrinogen/µL

were stained with Alcian Blue dye. (M) GFP-WJ-MSC cultured in monolayer and (N) HPCC

scaffolds without cells were used as differentiation controls. Chondrogenic differentiation

media and DMEM, both supplemented with tranexamic acid (5% v/v), were changed every

3 days. Scale bar: 100 µm.

Staining of HPCC scaffold prepared with 3 µg/µL of fibrinogen and 2-4x105 cells

(Figure 3-10A,B), revealed an intense red color like HPCC without cells (Figure 3-10N).

Some areas are dark and not defined cells are observed. In contrast, in scaffolds at different

fibrinogen concentrations prepared with 6x105 cells (Figure 3-10C,G, and K) red grouped

areas were identified in the form of a precipitate inside the scaffold, which probably

suggests the formation of mineralized areas. This behavior is observed with less intensity

2×

10

5



HPCC without cells

100 µm

100 µm

Seedin

g d

ensi

ty (ce

lls/

scaffold

)

6×

10

5D

MEM

4×

10

54×

10

5


WJ-MSCs in a HPCC


DMEM

Monolayer

100 µm

100 µm

100 µm

100 µm

100 µm

100 µm

100 µm

100 µm

100 µm

100 µm

100 µm

100 µm

A

B

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E

F

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I

J

K

M

N

D H L



in gels prepared with 2 and 1 µg/µL of fibrinogen, and 2 and 4x105 cells (Figure 3-10E,I,F,

and J), respectively.

Figure 3-10. Osteogenic differentiation of GFP-WJ-MSC embedded in HPCC-Osteogenic

media scaffolds. Osteogenic differentiation was evaluated at three cell seeding densities:



control. Scaffolds prepared at (A-D) 3, (E-H) 2 and (I-L) 1 µg of fibrinogen/µL were stained

with Alizarin Red dye. (M) GFP-WJ-MSC cultured in monolayer and (N) HPCC scaffolds

without cells were used as differentiation controls. Osteogenic differentiation media and


Scale bar: 100 µm.

The morphology of the control cells, which correspond to 6x105 WJ-MSC cultured in

DMEM-HPCC, is round, and despite being red, staining of DMEM-HPCC scaffolds is less

intense than those where osteogenic differentiation was carried out (Figure 3-10D, H and

L).

2×

10

5



HPCC without cells

100 µm

100 µm

Seedin

g d

ensi

ty (ce

lls/

scaffold

)

6×

10

5D

MEM

6×

10

54×

10

5


WJ-MSCs in a HPCC


DMEM

Monolayer

100 µm

100 µm

100 µm

100 µm

100 µm

100 µm

100 µm

100 µm

100 µm

100 µm

100 µm

100 µm

A

B

C

E

F

G

I

J

K

M

N

D H L



According to the staining results with Oil Red, Alcian Blue, and Alizarin Red for GFP-

WJ-MSC embedded in fibrin scaffolds prepared and cultured with adipogenic,

chondrogenic, and adipogenic media, respectively, it is possible to affirm that fibrin gels

do not interfere with the differentiation ability of WJ-MSC.

3.4.4 Fibrinogen concentration and culture media affect the inner

morphology of HPCC scaffolds.

Scanning electron microscopy (SEM) analysis of fibrin scaffolds with embedded GFP-

WJ-MSC, after 7 days of culture, revealed similar clot structures with fiber thickness of 100

nm ± 25 at fibrinogen concentrations between 1.5 and 1.9 µg/µL (Figure 3-11A-C).

However, when the fibrinogen concentration increased to 3.3 µg/µL, the formation of a

more compact structure was observed, with few well-defined fibers of 260 nm average

thickness (Figure 3-11D). The structure of the cell-free scaffolds incubated for 7 days in

DMEM supplemented with 10% FBS and tranexamic acid (0.5% v/v) (DMEM-FBS-TA)

(Figure 3-11E) varied significantly compared to the fresh scaffold prepared for SEM

analysis (Figure 3-11A-C). Two important variations were observed. First, the appearance

of a precipitate was observed on the scaffolds incubated for 7 days, which could probably

correspond to other proteins present in human plasma such as albumin and globulin.

Second, the thickness of the fibers in the gel incubated for 7 days was twice the size of the

fibers of the freshly prepared scaffolds (Figure 3-11G).



Figure 3-11. Scanning electron microscopy (SEM) analysis of fibrin scaffolds. (A-D)

HPCC gels with variable fibrinogen concentrations (1.5, 1.7, 1.9, and 3.3 µg/µL) with 5.5

x105 GFP-WJ-MSC per scaffold after 7 days of culture in DMEM supplemented with 10%

FBS and tranexamic acid (0.5 % v/v) (DMEM-FBS-TA). HPCC gels (E) without cells after 7

days of incubation in DMEM-FBS-TA, and (F) without incubation (freshly prepared before

SEM analysis). Scale bar: 10 µm. (G) Average fiber diameter (bar height) of fibrin scaffolds

estimated by image processing using ImageJ software. The error bar corresponds to one

standard deviation (SD). (a,b,c,d) denotes significance (n = 20, p < 0.05) in comparison with all

groups.

1.5

µg/µ

l

1.7

µg/µ

l

1.9

µg/µ

l

3.3

µg/µ

l

6.6

µg/µ

l

6.6

. µg/µ

l

Without in

cubat

ion

A B

C D

E F

G



Figure 3-12. SEM photographs of increasingly smaller sections of the surface of a

scaffold made at a fibrinogen concentration of 6.6 µg/µL. The scaffold was made by

mixing HPCC with calcium, and WJ-MSC in a centrifuge tube of 50 mL to form a spherical

gel (diameter 8 mm) and cultured in DMEM+10%FBS+0.5% tranexamic acid (v/v) for 28

days. Note that the surface of the scaffold is characterized by the presence of a “skin”,

which is indicated in Figure 3-12B by the white arrows. Further magnification reveals the

presence of some sort of low-porosity material (white arrows in Figure 3-12E and F),

covering the most porous fibrin network (Figure 3-12F).

A B C

DEF




zones on the surface of a scaffold made at a fibrinogen concentration of 1.9 µg/µL. The

scaffold was made by mixing HPCC with calcium, DMEM+10%FBS and WJ-MSC, and

cultured for 7 days in DMEM+10%FBS+0.5% tranexamic acid (v/v). Orange-framed sections

(A-F) correspond to a fracture in the “skin” of the scaffold, which allows the visualization

of the fibrin network. Cyan-framed sections (G-J) focus on the structure of the skin. Greater

magnification reveals the radical difference in the structure of the skin vs. fibrin network,

visible at the same magnification in Figure 3-13F, and J.

A B

C

D

E

F

G

H

I

J




zones of the surface of a scaffold made at a fibrinogen concentration of 6.6 µg/µL without

cells. The scaffold was made by mixing 180 µL of HPCC with 10 µL solution of calcium

chloride 3% p/v. After gel formation, 400 µL of DMEM+10%FBS+ 0.5 % tranexamic acid

were added. The scaffold was incubated for 7 days at 37°C, and 5% CO2.

Figure 3-15. SEM photographs of increasingly smaller sections of the surface of a fresh

scaffold made by mixing HPCC with calcium at a final fibrinogen concentration of 6,6

µg/µL, without incubation. Note that the surface is homogeneous and characterized by

well-defined, clean fibers of fibrin.

G

A B C

DEF



Figure 3-16. Graphic representation of fibrin network

3.4.5 Fibrin scaffolds reduce PEI toxicity in WJ-MSC

The effect of fibrinogen, and PEI/DNA concentration on the metabolic activity and

rheological behavior of WJ-MSC embedded in HPCC scaffolds was evaluated (Figure 3-17).

The metabolic activity of 2x105 WJ-MSC cultured in HPCC scaffolds at three different

fibrinogen concentrations was compared with the same seeding conditions in monolayer.

Cells were cultured in DMEM 10% FBS and 5% tranexamic acid for four days. The relative

fluorescence of resazurin reduced to resorufin as an indicator of metabolic activity was

measured as a function of fibrinogen concentration (Figure 3-17).

The metabolic activity of WJ-MSC decreased linearly with fibrinogen content. Since all

HPCC scaffolds were prepared with the same diameter and thickness by keeping a constant

WJ-MSC concentration, this effect could be attributed to a diffusivity restriction imposed

by fibrinogen content into the scaffolds. In Figure 3-11G, it was observed that fibrin fiber

thickness increased with fibrinogen concentration and, consequently, pore size was

reduced. These results suggest that there is a limitation in the diffusion of substances from

the culture medium into the gels as a result of the increased concentration of fibrinogen.

Considering the diffusive restriction imposed by the fibrinogen concentration in the gel,

the effect of the PEI, and PEI/DNA content was analyzed on the metabolic activity of the

WJ-MSC embedded in the fibrin gels (Figure 3-17B-C).



At 1 µg of fibrinogen/µL in the HPCC scaffolds, there were no significant differences

between the metabolic activity of cells with PEI content between 5 and 15 µg/scaffolds. As

the final volume of the scaffold was 240 ul, PEI concentrations between 20-60 ng of PEI/µL

were not cytotoxic to cells in comparison with monolayer cultures, where a concentration

of 12 ng of PEI/µL was lethal for WJ-MSC.

At the same fibrinogen concentration, the effect of DNA complexed with PEI on

metabolic activity was observed. The fluorescence intensity was reduced by 8 ± 4% in

comparison with the control for all complexes with DNA content between 2.5 and 7.5

µg/scaffold. At the highest DNA and PEI content (7.5and 15 µg, respectively), this effect

was reinforced by the increase in fibrinogen concentration into the scaffold. Metabolic

activity was reduced by 10 ± 8% and 27 ± 5% when the fibrinogen concentration increased

to 2 and 3 µg/µL into the scaffold, respectively. Fibrinogen concentrations below 2 µg/µL

generated scaffolds with fibers of 100 nm thickness. When the fibrinogen concentration

increases to 3.3 µg/µL, a compact structure is formed, and the fiber thickness increases 2 or

3 times approximately.



Figure 3-17. Effect of fibrinogen and PEI/DNA concentration on the metabolic activity

and rheological behavior of GFP-WJ-MSC embedded in HPCC scaffolds. 2x105 WJ-MSC

embedded in HPCC scaffolds at different fibrinogen concentrations were treated with 44

E

A

D

B

C



M resazurin in DMEM 10% FBS for 3 h. (A) Relative fluorescence of resazurin reduced to

resorufin as an indicator of metabolic activity was measured as a function of fibrinogen

concentration. The effect of the concentration of PEI (5, 10, and 15 µg/µL), both alone (P)

and complexed with DNA (C), was analyzed on the metabolic activity of WJ-MSC

embedded in HPCC scaffolds made at (B) 1, (C) 2 and (D) 3 µg/µL of fibrinogen. Cells

growing without PEI or DNA in the scaffolds at the proper fibrinogen concentration were

used as controls. In every case, the PEI/DNA mass ratio was maintained at 2. (a,b,c) denotes

significance (n = 3, p < 0.05) in comparison with all groups at the same fibrinogen

concentration. (E) Effect of GFP-WJ-MSC and PEI/DNA complexes on the elastic modulus

of HPCC scaffolds at 3 µg/µL of fibrinogen (strain: 0.1%).

Finally, the effect of GFP-WJ-MSC and PEI/DNA complexes on the rheological

behavior of HPCC scaffolds at 3 µg/µL of fibrinogen was evaluated (Figure 3-17E). The

elastic modulus of HPCC scaffolds with cells was 80 Pa. The addition of PEI/DNA

complexes reduced in 20% the elastic modulus. However, in comparison with HPCC

without cells and complexes, there is an increase of 50%. These results suggest that

PEI/DNA complexes slightly weaken the elastic behavior of HPCC, even though this

reduction is compensated by the introduction of WJ-MSC into the gels.

Cell viability experiments were performed to assess the toxicity on WJ-MSC of both

PEI, and PEI-DNA complexes onto the fibrin scaffolds. The analysis was performed on day

4, when cells had already shown an indication of positive transfection. Results showed sets

of treatments with a similar reduction in resorufin fluorescence that depended mainly on

the fibrinogen concentration of the scaffold (block), with smaller variations according to

the other treatments (content of PEI or PEI-DNA). The analysis began by comparing the

results of the control scaffolds, where the only variable was the concentration of fibrinogen.

Figure 3-17A shows a monotonic reduction in resorufin fluorescence of control scaffolds as

fibrinogen concentration increases. A possible interpretation of this result is that fibrinogen

or any other component present in human plasma might cause a reduction in cell viability,

which would seem counterintuitive as human plasma allows the transport of nutrients in

the human body and bovine plasma is used to grow WJ-MSC in vitro. More likely, this

result is due to a simultaneous reduction of both resazurin and resorufin diffusion in and



from inside the scaffold, respectively. Such reduction can be expected as the fibrin fibers

obstruct the movement of the reagent molecules, thus reducing the mean free path

compared to the monolayer culture (identified in Figure 3-17A as the bar with a fibrinogen

concentration of 0 µg/µL). To further examine this possibility, the hypothesis is that more

fibrinogen concentration could create gels with a more entangled and less porous structure,

which would explain these results.

To test this hypothesis, a brief analysis is performed. If all fibrinogen in the HPCC

reacts and is integrated in the fibers conforming the fibrin network, then, the mass of

fibrinogen included in the scaffold can be expressed as shown in equation (3 -1).

𝑚𝑓𝑖𝑏𝑟𝑖𝑛𝑜𝑔𝑒𝑛 = 𝑉𝑠𝑐𝑎𝑓𝑓𝑜𝑙𝑑 ∙ 𝐶𝑓𝑖𝑏𝑟𝑖𝑛𝑜𝑔𝑒𝑛 𝑖𝑛 𝑠𝑐𝑎𝑓𝑓𝑜𝑙𝑑 (3-1)

The packing process of fibrin proto-fibers is assumed to create fibers with the same

mass density (𝜌𝑓𝑖𝑏𝑒𝑟). The diameter of the fibers (𝑑𝑓𝑖𝑏𝑒𝑟) is also assumed to remain constant

through the whole matrix for a certain gelling condition, i.e. the measured diameter

depends only on the fibrinogen concentration, the surrounding environment and the

procedure used to treat the gel for SEM imaging. Under these assumptions, it would be

possible to state that for every fibrinogen concentration in the scaffold

(𝐶𝑓𝑖𝑏𝑟𝑖𝑛𝑜𝑔𝑒𝑛 𝑖𝑛 𝑠𝑐𝑎𝑓𝑓𝑜𝑙𝑑), the whole fibrin matrix could be “decomposed”, and “reassembled”

into a single fiber with a particular diameter and length, as illustrated in Figure 3-16. A

mass balance of fibrinogen could be used to find the total fibrin length (𝐿𝑡𝑜𝑡𝑎𝑙) in every

concentration, as presented in equation (3-2).

𝑚𝑓𝑖𝑏𝑟𝑖𝑛𝑜𝑔𝑒𝑛 = 𝜌𝑓𝑖𝑏𝑒𝑟 ∙𝜋

4∙ 𝑑𝑓𝑖𝑏𝑒𝑟

2 ∙ 𝐿𝑡𝑜𝑡𝑎𝑙 (3-2)

Comparing the results for two different fibrinogen concentrations, equation (3-2)

transforms into (3-3).

𝑚𝑓𝑖𝑏𝑟𝑖𝑛𝑜𝑔𝑒𝑛𝑗

𝑚𝑓𝑖𝑏𝑟𝑖𝑛𝑜𝑔𝑒𝑛𝑘

=𝜌𝑓𝑖𝑏𝑒𝑟 ∙

𝜋4 ∙ 𝑑𝑓𝑖𝑏𝑒𝑟 𝑗

2 ∙ 𝐿𝑡𝑜𝑡𝑎𝑙𝑗

𝜌𝑓𝑖𝑏𝑒𝑟 ∙𝜋4 ∙ 𝑑𝑓𝑖𝑏𝑒𝑟𝑘

2 ∙ 𝐿𝑡𝑜𝑡𝑎𝑙𝑘

(3-3)



=𝑉𝑠𝑐𝑎𝑓𝑓𝑜𝑙𝑑𝑗

∙ 𝐶𝑓𝑖𝑏𝑟𝑖𝑛𝑜𝑔𝑒𝑛 𝑖𝑛 𝑠𝑐𝑎𝑓𝑓𝑜𝑙𝑑𝑗

𝑉𝑠𝑐𝑎𝑓𝑓𝑜𝑙𝑑𝑘∙ 𝐶𝑓𝑖𝑏𝑟𝑖𝑛𝑜𝑔𝑒𝑛 𝑖𝑛 𝑠𝑐𝑎𝑓𝑓𝑜𝑙𝑑𝑘

As the sum of the volume of the reagents used to create the scaffolds in the well plate

remains constant (240 µL) regardless of the fibrinogen concentration, then, equation (3-3)

is reduced to equation (3-4).

𝑑𝑓𝑖𝑏𝑒𝑟 𝑗

2 ∙ 𝐿𝑡𝑜𝑡𝑎𝑙𝑗

𝑑𝑓𝑖𝑏𝑒𝑟𝑘

2 ∙ 𝐿𝑡𝑜𝑡𝑎𝑙𝑘

=𝐶𝑓𝑖𝑏𝑟𝑖𝑛𝑜𝑔𝑒𝑛 𝑖𝑛 𝑠𝑐𝑎𝑓𝑓𝑜𝑙𝑑𝑗

𝐶𝑓𝑖𝑏𝑟𝑖𝑛𝑜𝑔𝑒𝑛 𝑖𝑛 𝑠𝑐𝑎𝑓𝑓𝑜𝑙𝑑𝑘

(3-4)

The ratio of the total surface area of the fibers for two different scaffolds can be

calculated as shown in equation (3-5).

𝐴𝑓𝑖𝑏𝑒𝑟𝑠 𝑖𝑛 𝑠𝑐𝑎𝑓𝑓𝑜𝑙𝑑𝑗

𝐴𝑓𝑖𝑏𝑒𝑟𝑠 𝑖𝑛 𝑠𝑐𝑎𝑓𝑓𝑜𝑙𝑑𝑘

=𝜋 ∙ 𝑑𝑓𝑖𝑏𝑒𝑟𝑗

∙ 𝐿𝑡𝑜𝑡𝑎𝑙𝑗

𝜋 ∙ 𝑑𝑓𝑖𝑏𝑒𝑟𝑘∙ 𝐿𝑡𝑜𝑡𝑎𝑙𝑘

(3-5)

Equations (3-4) and (3-5) can be combined to render equation (3-6):



=𝑑𝑓𝑖𝑏𝑒𝑟𝑗

∙ 𝐿𝑡𝑜𝑡𝑎𝑙𝑗

𝑑𝑓𝑖𝑏𝑒𝑟𝑘∙ 𝐿𝑡𝑜𝑡𝑎𝑙𝑘

=𝑑𝑓𝑖𝑏𝑒𝑟𝑗

𝑑𝑓𝑖𝑏𝑒𝑟 𝑘

∙ (𝐶𝑓𝑖𝑏𝑟𝑖𝑛𝑜𝑔𝑒𝑛 𝑖𝑛 𝑠𝑐𝑎𝑓𝑓𝑜𝑙𝑑𝑗


∙𝑑𝑓𝑖𝑏𝑒𝑟𝑘

2

𝑑𝑓𝑖𝑏𝑒𝑟 𝑗

2 )



∙𝑑𝑓𝑖𝑏𝑒𝑟 𝑘

𝑑𝑓𝑖𝑏𝑒𝑟𝑗

(3-6)

According to Figure 3-11G, there is a linear relationship between the fiber diameter

and the concentration of fibrinogen in the scaffold, given by equation (3-7), where 𝑎 =

64,76 𝑛𝑚/(𝜇𝑔

𝜇𝑙) . Such relationship can be extrapolated to pass through the origin, implying

that the theoretical formation of gel fibers with a measurable diameter would happen at



any concentration of fibrinogen different than zero. This is consistent with the experimental

results of this work, as it has been possible to set gels at concentrations as low as 0,3 µg/µL.

𝑑𝑓𝑖𝑏𝑒𝑟 = 𝑎 ∙ 𝐶𝑓𝑖𝑏𝑟𝑖𝑛𝑜𝑔𝑒𝑛 𝑖𝑛 𝑠𝑐𝑎𝑓𝑓𝑜𝑙𝑑 (3-7)

Then, from equations (3-6) and (3-7), the ratio of areas of fibers in the scaffold with

different fibrinogen concentrations can be expressed as:





∙𝑎 ∙ 𝐶𝑓𝑖𝑏𝑟𝑖𝑛𝑜𝑔𝑒𝑛 𝑖𝑛 𝑠𝑐𝑎𝑓𝑓𝑜𝑙𝑑𝑘

𝑎 ∙ 𝐶𝑓𝑖𝑏𝑟𝑖𝑛𝑜𝑔𝑒𝑛 𝑖𝑛 𝑠𝑐𝑎𝑓𝑓𝑜𝑙𝑑𝑗



= 1

(3-8)

The obstructing area is the projected area of the cylindrical fibrin fiber perpendicular

to the path of a particular diffusing molecule. Although Equation (3-8) describes the ratio

of the surface area of fibrin fiber in two scaffolds, the obstructing area is proportional to the

surface area. Therefore, the proportionality constant is cancelled in equation (3-8) and the

same equation describes the ratio of reduction in the diffusion area within two different

fibrin gels. Consequently, the area obstructing the diffusion of reagents into or out of the

fibrin gels should remain the same, regardless of the fibrinogen concentration.

From the results of the previous analysis, the diffusion area occupied by the fibers is

expected to be the same in the analyzed range of concentrations. In consequence, the

linearly reduced diffusion as the fibrinogen concentration increases should have another

explanation.

From the images in Figure 3-12 and Figure 3-13, it is possible to observe that, in

addition to the fibrin matrix, the surface of scaffolds maintained in culture presents some

sort of additional layer. Such layer might correspond to a different, denser structure of the

same fibrin formed in response to air during polymerization. This phenomenon has been

reported before by Weiss et al. (2017) [79]. However, SEM photographs show the presence



on the surface of what appears to be other types of material, probably other types of

precipitated proteins such as albumin or globulin [80] present in human plasma. Those

protein deposits are unlikely caused by the cells, as they are also observable in scaffolds

without including WJ-MSC left in incubation for over a week (Figure 3-14). Besides, the

deposits are not present in freshly prepared scaffolds, as evidenced in Figure 3-15.

A visual examination of Figure 3-12 and Figure 3-13 allows establishing that the

porosity of the observed layer is much smaller than the porosity of the fibrin network.

Therefore, it is likely that the diffusion of reagents entering or leaving the scaffold will be

further diminished. As the fibrinogen in the scaffolds comes from human plasma, larger

concentrations of fibrinogen produce larger concentrations of plasma, hence larger

concentrations of other proteins. As a result, larger fibrinogen in the scaffold corresponds

to thicker deposits of proteins, smaller diffusion of resazurin toward cells, and of resorufin

from cells out of the scaffold, which agrees with the lower fluorescence results observed in

Figure 3-17A to D.

According to the previous discussion, the results from scaffolds with different

fibrinogen concentrations cannot be directly compared. Consequently, the results for every

concentration were arranged in a different panel in Figure 3-17 and compared with their

respective control scaffold without any PEI or PEI-DNA complex.

The only statistically significant difference in comparison with the control appears to

exist when 15 µg of PEI-DNA complexes are included in scaffolds with 3 µg/µL of

fibrinogen. In this case, an average reduction in cell viability to 73 % was observed in

culture at day 4 with respect to the control at the same fibrinogen concentration. A similar

reduction in cell viability of PEI-DNA complexes at early time points in fibrin scaffolds was

reported by Saul et al. (2007), who attributed the phenomenon to the immediate access of

cells to the complexes. [81].



3.4.6 PEI/DNA nanoparticles characterization

Complexes formed by mixing DNA with PEI at different PEI/DNA mass ratios were

evaluated by agarose gel electrophoresis (Figure 3-18). PEI and DNA mass ratios equal or

greater than 0.25 do not allow band displacement, which suggests the formation of

complexes of DNA and PEI.

Figure 3-18. Agarose gel electrophoresis of PEI/DNA complexes at different PEI/DNA

mass ratios. Non-displacement in the band indicates the formation of complexes between

PEI and DNA. For this assay, 100 ng of DNA were mixed with PEI at different

concentrations to establish the PEI/DNA mass ratio at which complex formation occurs.

3.4.7 PEI and DNA content regulate transfection efficiency in HPCC

scaffolds

In order to evaluate the effect of fibrinogen and DNA content on the transfection

efficiency of HEK cells and WJ-MSC into the HPCC scaffolds, different contents of

complexes of PEI and DNA at the same mass ratio of 2 were embedded in fibrin scaffolds

at three different fibrinogen concentrations (1, 2, and 3 ug/ul). 4x105 HEK cells and 2x105

WJ-MSC per scaffold were employed. Successfully transfected cells showed characteristic

green fluorescence due to the expression of the gene, and formation of the GFP protein.

500 0 12.5 25 50 100 150 200 300 400 500 0 500

ng of PEI in complexes Only

PEI

MW

marker

MW

marker

Only

PEI

bp

1500

100

bp

1500

100



Transfection efficiency was estimated based on GFP expression by fluorescence

microscopy. Scaffolds with transfected cells were analyzed by fluorescence microscopy.

Briefly, 108 photos of each scaffold were taken under the same magnification (4X) to

reconstruct the entire gel.

HEK cells transfection was evaluated at 5 different DNA contents (1, 2, 3, 4, and 5 g

per scaffold). The results of transfection after 24 h were presented in Figure 3-19. It was

observed that the transfection efficiency of HEK increased with DNA content in the

scaffold, but there is no apparent relationship between fibrinogen concentration into the

scaffold and HEK transfection (Figure 3-19). The number of transfected cells increased with

DNA content into the scaffold 60 h after transfection (Figure 3-20); however, at this time

strong differences were observed when fibrinogen content varied. Except for 4 µg of DNA

content, when the maximum value of transfection was reached, at every content of DNA

evaluated, transfection efficiency of HEK cells increased with fibrinogen content. Similarly,

a relationship between post-transfection time and transfection efficiency was observed,

finding more transfected cells at 60 h in comparison with 24 h post-transfection.

The variation in transfection efficiency of WJ-MSC with fibrinogen concentration and

DNA content is presented in Figure 3-21 and Figure 3-22. The same methodology

employed to quantify HEK transfection was used to establish the best transfection

conditions for WJ-MSC in fibrin scaffolds. Transfection efficiency of WJ-MSC was strongly

dependent on DNA content; however, the efficiency decreased with the augment in

fibrinogen concentration into the scaffold. This effect is evident mostly at DNA contents in

the scaffold below 7 µg. An increase in the complex content of the scaffold is translated into

an increase in transfection efficiency. Figure 3-23B reveals that there is a combined effect of

fibrinogen concentration in the scaffold, and DNA and PEI content in transfection

efficiency.

Since all the tests were performed at the same PEI /DNA mass ratio, it is possible to

attribute the observed effects to the number of complexes available in the scaffold. A lower

DNA content indicates a lower number of available complexes.




tridimensional scaffold of HPCC at 24 h. 4x105 HEK cells were seeded in scaffolds of

HPCC. Three different fibrinogen concentrations were evaluated: (A-E) 3 µg/µL, (F-J) 2

µg/µL and (K-O) 1 µg/µL. Complexes of PEI and DNA were prepared at five different

contents of DNA: (A, F and K) 1 µg, (B, G and L) 2 µg, (C, H and M) 3 µg, (D, I and N) 4 µg

and (E, J and O) 5 µg/scaffold. In every case, the PEI/DNA mass ratio was maintained at 2.

The final volume of each scaffold was 240 µL, scale bar: 500 µm



1 µ

g2

µg

3 µ

g4

µg

5 µ

g

DN

A c

onte

nt/

scaffold

A

B

C

D

E

F

G

H

I

J

K

L

M

N

O





using Gen5TM software from Biotek®. 4x105 HEK cells were seeded in scaffolds of HPCC.

Three different fibrinogen concentrations were evaluated (A-E) 3 µg/µL, (F-J) 2 µg/µL, and


DNA: (A,F, and K) 1 µg, (B,G, and L) 2 µg, (C,H, and M) 3 µg, (D,I, and N) 4 µg, and (E, J,


volume of each scaffold: 240 µL. Scale bar: 5 mm



1 µ

g2

µg

3 µ

g4

µg

5 µ

g

DN

A c

onte

nt/

scaffold

A

B

C

D

E

F

G

H

I

J

K

L

M

N

O

5 mm




tridimensional scaffold of HPCC at 48 h. 2x105 WJ-MSC were seeded in scaffolds of HPCC.





volume of each scaffold: 240 µL. Scale bar: 200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm

200 µm



5 µ

g6

µg

7 µ

g8

µg

9 µ

g

DN

A c

once

ntr

ation/s

caffold

A

B

C

D

E

F

G

H

I

J

K

L

M

N

O





using Gen5TM software from Biotek®. 2x105 WJ-MSC were seeded in scaffolds of HPCC.


(K-O) 1 µg/µL. Complexes of PEI and DNA were prepared at five different contents of



volume of each scaffold: 240 µL. Scale bar: 5 mm



5 µ

g6

µg

7 µ

g8

µg

9 µ

g

DN

A c

onte

nt/

scaffold

A

B

C

D

E

F

G

H

I

J

K

L

M

N

O

5 mm



Figure 3-23. Effect of DNA content on the transfection efficiency of HEK cells, and WJ-

MSC in a tridimensional scaffold of HPCC. Quantitative analysis. 108 photos of each gel

at 4X were taken to reconstruct the conformation of the entire well. Images were processed

and analyzed using ImageJ software to establish the percentage of the fluorescent green

area into the well. Transfection efficiency was defined as the percentage of green area with

respect to the total area of the well. Effect of scaffold DNA content (PEI/DNA mass ratio of

2) was studied on transfection efficiency of (A) HEK cells and (B) WJ-MSC. Final volume

of each scaffold: 240 µL.

3.5 Discussion

Fibrinogen is a soluble glycoprotein, specifically a clotting factor (factor I), present in

blood. The main role of fibrinogen is its polymerization into fibrin clots [82], which are the

end result of the proteolytic activation of clotting factors [83]. Fibrin scaffolds have been

used in tissue engineering and regenerative medicine [84] to accelerate or promote wound

repair [85] in cardiovascular tissue engineering [86-88], cartilage engineering [68,89], among

others, due to their biocompatibility and biodegradability.

Additionally, fibrin is widely used as a glue in surgery because of its hemostatic and

adhesive properties [90,91]. It is the only agent currently approved as a hemostat, sealant,

and adhesive by the Food and Drug Administration (FDA) [92].

Several studies have incorporated the use of fibrin as a delivery system for growth

factors[93,94], drugs (anticancer agents)[95,96], antibiotics[97] or genes[81,98]. Gene release

has sparked special interest in tissue engineering. Both autologous[99] and heterologous

A B



fibrin formulations [100] have been used in gene delivery. In the present work, autologous

fibrin from human plasma was used as a gene delivery system for the transfection of WJ-

MCSs.

For WJ-MSC, our results showed that a decrease of the fibrinogen concentration in the

scaffold produces a greater gene transfer efficiency at the same DNA content. However,

the opposite effect was observed for the HEK cell line. The three-dimensional configuration

of scaffolds, where the complexes are immobilized simultaneously with the cells, creates

an environment where the cells are surrounded by the complexes, but their access to them

depends on the growth and adhesion of the cells to the scaffold and its corresponding

degradation. Additionally, it was found that fibrin gels reduce the cytotoxic effects of PEI,

which was used as a transfection agent. By reducing toxicity, it is possible to increase

transfection efficiency in comparison with the same conditions in a two-dimensional

culture. Our results demonstrate that a three-fold increase in the PEI dose into the

scaffolds/cell does not affect cell viability relative to monolayer transfection conditions.

Andreadis et al. (2009) attributed this increase in cell viability to the specific and

immobile location of complexes within the fibrin fibers, so that cells are only exposed to

the complex as fibrin fibers degrade. This differs from traditional transfection

methodologies where cells are exposed to high concentrations of gene delivery vehicles,

which are generally toxic to cells [101-103].

Similar to transfection using traditional two-dimensional (monolayer) cultures, there

is a critical level at which PEI is cytotoxic. This level is dependent on the cell type. In our

monolayer assays, the onset of PEI cytotoxicity was 22ng/µL for 1x105 HEK and 0.8ng/µL

for 4.5x104 WJ-MSC. In three-dimensional transfection assays, a transfection peak for HEK

was reached when the DNA content was 4 µg and PEI was 8 µg. Above that values the

transfection efficiency decreased with the fibrinogen concentration in the gel. The lower

the fibrinogen concentration in the gel, the more particles are exposed faster and therefore

the efficiency of transfection is decreased since it begins to be toxic to cells.

For WJ-MSC, this critical point does not appear to be within the evaluated conditions

because an increase in transfection is observed when increasing DNA content and



decreasing fibrinogen concentration. Therefore, in subsequent tests it is necessary to

evaluate a higher content of DNA in the scaffold in order to find the value that maximizes

the efficiency of transfection. According results presented by Andreadis (2009) [104]and

Tabata (2010), transfection conditions, as in monolayer culture, are dependent on cell type

and scaffold interaction [105,106].

It is possible that primary cells are more sensitive to matrix interactions, as they

proliferated quickly in 3D scaffolds, which means that the 3D environment acts as a more

powerful adjuvant for these cells [105].

One of the unexpected results obtained was the inverse relationship between the

transfection efficiency of HEK with respect to WJ-MSC. The transfection efficiency of HEKs

was favored by the highest concentration of fibrinogen in the scaffold evaluated, while the

best results for WJ-MSC were obtained at the lowest concentrations of fibrinogen in the

scaffold.

In WJ-MSC, an increase in transfection efficiency with a decrease in concentration can

be explained by the size of the WJ-MSC and the better colonization of the scaffold. Due to

these conditions with degradation of fibrin, the complex is more available. It is also possible

that WJ-MSC degrade fibrin faster.

WJ-MSC are less but larger, in comparison with HEK and, in monolayer, they have

shown to grow better when cultured in platelet lysate. Within the fibrin gel, cells have

access to the same more concentrated factors, so they must be proliferating better than in

monolayer. This can be corroborated with the test where the seeding density was varied,

and it was observed that the WJ-MSC colonized the fibrin gel very quickly and that they

ran out of space. Therefore, seeding density was selected as the starting concentration for

the transfection in gels. Preliminary tests also showed that when fewer cells were seeded

(50,000), the cells were widely spaced from each other, thus colonization of the scaffold

slowed down, and the cells began to lengthen too much, perhaps because they were looking

for other adjacent cells. Several authors have pointed out the effect of fibrinolytic inhibitors

on gene transfer. Future tests should consider the variation of tranexamic acid, which in



this test was 0.5% v/v (mg/mL), in order to study the effect of fiber degradation on

transfection efficiency.

Our methodology includes mixing the cells with the complexes before the formation

of the gel with HPCC-Ca. This interaction facilitates that, once in the scaffold, the distance

between the cells and the complexes is minimal, therefore, the absorption of the complexes

by the cells is easier. Contrary to the results reported by Andreadis (2010) for the

transfection of HEK with lipofectamine in fibrin scaffolds, we found that transfection is

favored by an increase in the concentration of fibrinogen in the gel. It is necessary to

evaluate the differences between lipofectamine and PEI at different concentrations of

fibrinogen in order to establish whether the observed effect is due to the content of

fibrinogen in the scaffold or to the mechanism of the transfection agent and its interaction

with the scaffold [104].

Another finding is related to the toxicity of PEI alone and PEI complexed to DNA in

fibrin scaffolds. It was observed that the toxicity of PEI and of the complexes was

proportional to their final content in the scaffold. However, it was also found that the

toxicity augmented with the increase in fibrin concentration. This analysis was made by

normalizing the results with respect to the cells in the scaffold without treatment to

eliminate the diffusional effects, which, as previously demonstrated, are a linear function

with the concentration of fibrinogen in the scaffold. This means that a higher concentration

of fibrinogen in the scaffold translates into less diffusion from the culture medium into the

gel and vice versa. A higher concentration of fibrinogen implies a closer contact between

the cells and the complex, which in turn can translate into greater transfection efficiency,

but also into greater toxicity. The complexes are approximately 100 nm in size, PEI alone

does not form particles and is in solution. It is likely that when PEI is complexed with DNA,

there will be "spots" with higher concentration of PEI and, therefore, the complexes are

more toxic. This increased toxicity can also be associated with exogenous DNA content,

which has also shown to affect cell viability [107].

Previous research has shown that the rheological properties of fibrin gels are a function

of the concentration of fibrinogen and thrombin. In our work, these properties are



improved with the presence of cells, but reduced with the presence of the complexes. The

configuration gel + cells + complex have an intermediate value of the elastic modulus

between that of the gel alone and the gel with cells.

Monolayer PEI transfection assays demonstrated that transfection does not affect the

immunomodulatory properties, immunophenotype, and differentiability of WJ-MSC.

Further testing is required to evaluate these same characteristics, especially the

differentiation capacity in the scaffold, in order to evaluate whether it is improved or not

by introducing genes that promote cell differentiation towards a specific lineage. When

using a three-dimensional transfection strategy in fibrin scaffolds, complexes become

trapped within the network and can be released with the degradation of the fibrin fibers,

which favors transfection and reduces cytotoxicity associated with non-viral vectors.

Additionally, three-dimensional transfection can increase the number of transfected cells

per unit area, compared with two-dimensional transfection, where there are limitations in

cell growth and expansion due to inhibition by cell-cell contact. Our results suggest that

human plasma-based (autologous) fibrin hydrogels containing PEI / DNA complexes and

WJ-MSC can be used as platforms for localized and targeted gene delivery to enhance the

potential therapeutic use of MSC in tissue engineering.

3.6 Conclusions

PEI/DNA complexes embedded in human plasma scaffolds allow the efficient gene

release, minimizing cytotoxicity compared with traditional two-dimensional transfection

methodology. Fibrinogen content into the scaffolds regulates both the diameter of the fibrin

fibers and the pore size, which in turn influence both the mechanisms of nutrient transport

from the culture medium to the interior of the gels and the transfection efficiency of

complexes embedded in scaffolds. The methodology developed to elaborate scaffolds from

human plasma, in which the cells are in close contact with the complexes when forming

the gel, favors the interactions between the cells and the complexes, which may translate

into greater transfection efficiency.



3.7 Supplementary material

Supplementary Figure 3-S1. Fibrinogen content in cryoconcentrated human plasma by the

Clauss technique.

Supplementary figure 3-S2. Detailed concentrations used to prepare HPCC scaffolds with

MSC.



Author Contributions: Conceptualization, methodology and validation, A.I.R.-M., G.S. and R.D.G.-

S.; formal analysis, A.I.R.-M.; investigation, A.I.R.-M., I.S., L.S.; resources, G.A.S., B.C. and R.D.G.-

S.; writing—original draft preparation, A.R.; writing—review and editing, A.I.R.-M., R.D.G.-S.;

visualization, A.I.R.-M.; supervision, project administration and funding acquisition G.A.S., R.D.G.-

S. and B.C. All authors have read and agreed to the published version of the manuscript.

Funding: This research was funded by Universidad Nacional de Colombia under Grant

203010026990, the Ministry of Science, Technology, and Innovation (MINCIENCIAS) through the

Doctoral Scholarship Program 567-2012 and grant BPIN2012000100186 from Fondo de Ciencia,

Tecnología e Innovación, Sistema General de Regalías, Colombia.

Acknowledgments: A.I.R.-M. thanks Valerie Dorsand for her help in TOP10® transformation and

plasmid extraction and thanks to Alejandra García Herrera who provided language help and writing

assistance of the article.

Conflicts of Interest: The authors declare no conflict of interest.

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4. Conclusions and recommendations

4.1 Conclusions

In this work, the effect of polyethylenimine (PEI) in the transfection of mesenchymal

stem/stromal cells from Wharton's Jelly (WJ-MSC) into a tridimensional scaffold of human

plasma cryoconcentrated was evaluated. This work was divided in two big steps. The first

step evaluates the efficacy of PEI as a non-viral gene delivery system that can be optimized

for gene therapy in WJ-MSC. Our results showed that WJ-MSC transfected with PEI

retained their morphology, plastic adherence, immunophenotype, immunomodulatory

function, and multi-lineage differentiation potential. In the second step, once the improved

methodology for transfection of WJ-MSC based on PEI was standardized, the evaluation

into a three-dimensional scaffold that emulates the characteristics of the extracellular

matrix was carried out.

For this purpose, WJ-MSC mixed with PEI/DNA complexes at several concentrations

were embedded in scaffolds of human plasma cryoconcentrated (HPCC) at different

fibrinogen concentrations. Fibrinogen content in the HPCC scaffolds regulates both the

diameter of the fibrin fibers and the pore size of the gels, which in turn influence both the

mechanisms of nutrient transport from the culture medium to the interior of the gels and

the transfection efficiency of complexes embedded in scaffolds. PEI/DNA complexes

embedded in HPCC scaffolds allow efficient gene release and minimize cytotoxicity

compared with traditional two-dimensional transfection methodology.



4.2 Recommendations

In this work, a gene-activated HPCC matrix combined with WJ-MSC was developed,

with potential use for tissue engineering. Our results open new research horizons. We

recommend changing the GFP reporter gene to a therapeutic one, in order to evaluate the

expression of the protein over time and the effect on WJ-MSC. Additionally, it is necessary

to characterize the different growth factors and cytokines present in human plasma and to

evaluate the effect of the concentration of the anti-fibrinolytic agent (thrombin) on the

transfection efficiency of WJ-MSC.

Finally, the most promising path for tissue engineering is to combine the gene-

activated matrix developed in this work with other porous scaffolds and evaluate the

release of therapeutic genes in other models, for example chondrogenic or osteogenic.

Modificación génica no viral de células madre ...

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