FACULTAD DE FARMACIA -...

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FACULTAD DE FARMACIA Effects of α-lipoic acid on lipid metabolism and mitochondrial biogenesis in adipocytes: study of the molecular mechanisms involved. Efectos del ácido α-lipoico sobre el metabolismo lipídico y la biogénesis mitocondrial en adipocitos: estudio de los mecanismos moleculares implicados. Marta Fernández Galilea Pamplona, 201

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FACULTAD DE FARMACIA

Effects of α-lipoic acid on lipid metabolism and

mitochondrial biogenesis in adipocytes: study of

the molecular mechanisms involved.

Efectos del ácido α-lipoico sobre el metabolismo

lipídico y la biogénesis mitocondrial en

adipocitos: estudio de los mecanismos

moleculares implicados.

Marta Fernández Galilea

Pamplona, 201

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FACULTAD DE FARMACIA

Memoria presentada por Dña. Marta Fernández Galilea para aspirar al grado

de Doctor por la Universidad de Navarra.

Fdo. Marta Fernández Galilea

El presente trabajo ha sido realizado bajo nuestra dirección en el

Departamento de Ciencias de la Alimentación y Fisiología, y autorizamos su

presentación ante el Tribunal que lo ha de juzgar.

Vo B

o Directora V

o B

o Co-directora

María Jesús Moreno Aliaga Carmen Patricia Pérez Matute

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Este trabajo ha sido posible gracias a la financiación de diversas entidades:

Asociación de Amigos de la Universidad de Navarra (beca predoctoral 2010-

2011), Gobierno de Navarra (Ayudas predoctorales de formación del Gobierno

de Navarra. Plan de Formación y de I+D 2010/2011), Ministerio de Educación y

Ciencia (AGL2006-04716/ALI) y Ministerio de Ciencia e Innovación (AGL 2009-

10873/ALI), Proyecto Nutrición, Obesidad y Salud “Línea Especial” Universidad

de Navarra.

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“Es a fuerza de observación y

reflexión que uno encuentra un

camino”.

Claude Monet

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“Si tu intención es describir la

verdad, hazlo con sencillez y la

elegancia déjasela al sastre.”

Albert Einstein

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A mis padres

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A Pedro

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Agradecimientos/Acknowledgements

Una vez terminada la Tesis Doctoral, es difícil emprender la tarea de escribir

los agradecimientos. Es en este momento cuando uno se da cuenta de cuánto

tiene que agradecer a tanta gente. Intentaré resumir en unas líneas la gratitud

que siento a todas las personas que han estado presentes en esta etapa de mi

vida.

En primer lugar quisiera agradecer a la Universidad de Navarra el haberme

formado no solo en el ámbito profesional sino también en el personal, así como

a la Facultad de Farmacia y al Departamento Ciencias de la Alimentación y

Fisiología por darme la oportunidad de realizar el presente trabajo de

investigación.

Quisiera agradecer a la Asociación de Amigos de la Universidad de Navarra

y al Gobierno de Navarra las becas y ayudas recibidas que han hecho posible la

realización de este proyecto.

A mis directoras de tesis, las Doctoras María Jesús Moreno Aliaga y Carmen

Patricia Pérez Matute. A vosotras debo agradeceros en primer lugar la

confianza que depositasteis en mí y expresaros el gran placer que ha sido para

mi trabajar junto a dos investigadoras de vuestra talla. Gracias por vuestro

apoyo y por que habéis estado a mi disposición cuando os he necesitado. En el

plano de lo personal, tengo también mucho que agradeceros, ya que gracias a

vosotras hoy puedo decir que no solo he conseguido finalizar este reto en forma

de Tesis, sino por que de alguna forma, gracias a vosotras pude conocer a la

persona con quien recientemente he comenzado el mejor proyecto mi vida.

Al Profesor Alfredo Martínez ya que sin la ayuda de la “línea especial”

muchos proyectos, este incluido, serían un imposible.

I would like to express my gratitude to Professor Dominique Langin. Thank

you very much for the opportunity to stay in your laboratory and for your

confidence. I will never forget your interest in my research and your help.

Moreover I want to thank to “l’equipe 4” for giving me the great experience I had

in Toulouse. Specially, I want to thank Marianne Houssier the pleasure of

working with you. I will never forget your amazing ability for doing 4 experiments

at the same time. To Lucile Mir, Diane Beuzelin, Emilie Courty and Claire

Estadieu, for making possible the “ladies group”. To Virginie Bourlier and Cedric

Moro, for speaking English with me and make it easier. To Aline, Corinne and

Marie for their expert advice. To Nathalie Vigerie, for your daily smile, for being

so friendly and for your help in different ways. To Etienne Mouisel and Valentin

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Barquissau, thank you for the funny moments inside and outside the lab. Finally,

thank you very much to Jennifer Saussede. I will never be able to put in words

how grateful I am. Thank you for your confidence and for being my friend.

Sincerely, “merci beaucoup”.

Así mismo quisiera expresar mi gratitud a la Dra. Matilde Bustos por sus

buenos consejos, sus muestras de cariño y por toda la ayuda que durante estos

años me ha prestado.

Al personal de administración y servicios de la Universidad de Navarra.

Gracias por que sois un pilar fundamental en el trabajo diario. En especial me

gustaría mencionar a la antigua Gerente de la Facultad de Farmacia Reyes

Saenz y a Gonzalo Flandes, por vuestra paciencia y esas sonrisas que hacen la

tarea más liviana. A los bedeles Gonzalo, Jesús y Enrique, por vuestras

diligencia en el trabajo y por que siempre tenéis una palabra amable.

A todos los miembros que durante estos años han conformado el

Departamento Ciencias de la Alimentación y Fisiología, mis compañeros.

Gracias por que de todos y cada uno de vosotros he aprendido algo. En

especial me gustaría agradecer, a la vicerrectora de Investigación Dª Iciar

Astiasarán, por sus sinceras muestras de afecto y la Decana de la Facultad de

Farmacia Dª Adela López de Cerain por el interés que siempre ha demostrado

en mi persona tanto a nivel personal como a cerca del desarrollo de mi tesis. A

los profesores Fermín Milagro, Ana Barber, Pilar Lostao y a la actual Directora

de Departamento Diana Ansorena el que siempre hayáis tenido la puerta abierta

para escucharme. De igual manera, quisiera mostrar mi más sincera gratitud a

las técnicos de laboratorio Verónica Ciaurriz, Ana Lorente y Asun Redín así

como a las secretarias Paula y Beatriz por todos los momentos compartidos, por

vuestro apoyo tanto científico como personal y por que cada día nos “aguantáis”

con una sonrisa.

A mi equipo. A María Zabala y Miguel L. Yoldi, por que, sin duda alguna,

sois los mejores. Gracias por vuestra ayuda y por compartir y sentir como

vuestro cada experimento, pero sobre todo, gracias por vuestra sincera amistad.

A las personas que ya no están en el equipo, Beatriz Marcos y Silvia Lorente,

gracias por ayudarme en mis primeros pasos de principiante. A las nuevas

incorporaciones, Ana Elsa Huerta y Laura La Iglesia por traernos un soplo de

aire fresco. Finalmente, a todos los que de forma más o menos fugaz habéis

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Agradecimientos/Acknowledgements

pasado por aquí y nos habéis dejado huella como son André, Sonia y Andrea,

nunca olvidaré los momentos que pasamos.

En el plano de lo personal quiero dar las gracias a mis amigos, quienes de

forma paciente habéis permanecido a mi lado con el paso de los años. María

Uría, María Zabala, David, Gabriel, Jon, Miquel, Xisco, Diego y Manuela.

Disfrutar de vuestra amistad me hace ser la persona más afortunada del mundo.

A José Luís Pérez Pastor y la pequeña Victoria. Gracias por vuestra

paciencia y por hacerme un hueco en vuestra casa siempre que ha sido

necesario, incluidos fines de semana y “fiestas de guardar”.

A mi familia. A “los Fernández”, gracias por acogerme cuando llegué a

Pamplona y por todos los fantásticos momentos que hemos compartido a lo

largo de la vida, por vuestro apoyo y por que aunque no nos caractericemos por

ser muy expresivos, “a buen entendedor pocas palabras bastan”. A “los Galilea”.

Gracias por vuestro inestimable apoyo y por ser como sois. Dicen que uno no

elige a la familia, yo, me siento afortunada, ya que si pudiese os elegiría a

vosotros. A mis abuelos Julia y Lorenzo. Gracias por vuestro infinito amor,

vuestra ternura y cariño, por ser un ejemplo de superación y de fuerza de

voluntad y por enseñarme que aunque a veces la vida sea dura no podemos ni

debemos rendirnos. Gracias por hacerme sentir orgullosa de ser vuestra nieta.

Así mismo, me gustaría recordar a mis abuelos Anita y Sixto, quienes en el

transcurso de esta Tesis nos dejaron. Gracias abuela por tu inagotable

paciencia y el inmenso amor que nos dejaste. A ti, abuelo, corazón noble y alma

de poeta, tengo que agradecerte no solo tu amor, sino los valores que nos

enseñaste como son, entre otros, la honestidad y el amor a la familia. Sois la

estrella que nos guía y siempre estaréis en nuestros corazones.

Siguiendo con la familia quiero dar las gracias a mi familia de Segovia. A mis

suegros Chelo y Antonio, gracias por vuestro cariño, por todos los momentos

vividos en “Torre” y Segovia, por hacerme un huequito en vuestras vidas y

sobre todo por preocuparos por mi en estos meses tan difíciles. A mi cuñado

Antonio, por que siempre estas ahí cuando se te necesita demostrando que

eres el mejor hermano mayor. Así mismo me gustaría expresar mi

agradecimiento a todos los tíos y primos, por el cariño que siempre me han

demostrado y en especial al tío Venan, la tía Sole y a “los niños” que nos llenan

de alegría aunque de vez en cuando hagan alguna trastada.

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A mis padres. A vosotros va dedicada esta tesis, y en estos momentos no

encuentro palabras que definan mi agradecimiento. Empezaré por decir que si

he llegado a finalizar una Tesis Doctoral no ha sido sino por la ayuda que me

habéis prestado. Todo lo que soy os lo debo a vosotros y nunca podré pagar

tanto amor y apoyo como he recibido. Gracias por aceptar de buena gana las

llamadas a cualquier hora del día, y de la noche, y por escuchar pacientemente

cualquier problema ya fuese grande o pequeño. Gracias por ser el mejor

ejemplo que se pueda tener, por enseñarme como el sacrificio y el esfuerzo al

final siempre tienen recompensa. Pero, sobre todo quiero agradeceros que

hayáis sabido dejarme cometer mis propios errores y por ayudarme a

solucionarlos.

En último lugar, aunque no en importancia, quiero dar las gracias a mi

marido. Pedro, tu has sido y eres el mejor compañero que se pueda tener,

primero en el laboratorio y ahora en la vida. Quiero darte las gracias por todo el

amor que me regalas a diario, por tu ayuda constante y por que desde que te

conocí ni un solo día ha faltado la felicidad en mi vida.

A todos vosotros, ¡muchas gracias

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INDEX/INDICE

1.1. OBESITY ................................................................................................. 1

1.2. ADIPOSE TISSUE .................................................................................. 2

1.2.1. White adipose tissue ................................................................. 2

1.2.2. Brown adipose tissue ................................................................ 3

1.2.3. Beige adipocytes ........................................................................ 4

1.3. WHITE ADIPOSE TISSUE AS A KEY STORAGE ORGAN .................... 5

1.3.1. Lipolysis in adipocytes .............................................................. 6

1.3.1.1. Adipose Triglyceride Lipase (ATGL) .................................... 6

1.3.1.2. Hormone Sensitive Lipase (HSL) ......................................... 8

1.3.1.3. Perilipin ................................................................................ 9

1.3.1.4. AdPLA .................................................................................. 9

1.3.1.5. Signaling pathways involved in lipolysis ............................. 10

1.3.2. Triglycerides storage: lipogenesis ......................................... 11

1.3.3. Fatty acid esterification ........................................................... 12

1.3.3.1. Diacyl Glycerol Acyl Transferase (DGAT) ......................... 12

1.3.4. De novo lipogenesis ................................................................ 13

1.3.4.1. Acetyl CoA Carboxylase (ACC) ......................................... 14

1.3.4.2. Fatty Acid Synthase (FAS) ................................................. 15

1.3.4.3. Stearoyl CoA Desaturase 1 (SCD1)................................... 15

1.4. MITOCHONDRIA AND OBESITY ......................................................... 16

1.4.1. Mitochondrial biogenesis ........................................................ 16

1.4.2 Mitochondrial function in the regulation of fatty acids

metabolism ................................................................................................. 18

1.5. OBESITY AND OXIDATIVE STRESS ................................................... 19

1.6. α-LIPOIC ACID ...................................................................................... 20

1.6.1. Antiobesity effects of α-LA ...................................................... 21

1.6.1.1. Animal studies ................................................................... 21

1.6.1.2 Human clinical trials ............................................................ 26

1.6.2. Mechanisms of action ...................................................................... 29

1.6.2.1. α-LA reduces food intake and increases energy

expenditure…………………………………..…………………………....29

1.6.2.2. α-LA inhibits adipogenesis ................................................. 29

1.6.2.3. α-LA improves glucose metabolism ................................... 30

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1.6.2.4. Regulation of adipokine production .................................... 31

1.6.2.5. Effects of α-LA on lipid metabolism .................................... 33

CHAPTER 2: HYPOTHESIS AND AIMS/HIPÓTESIS Y OBJETIVOS ......... 57

CAPÍTULO 3: MATERIAL Y MÉTODOS

3.1.-CULTIVO DE LA LÍNEA CELULAR 3T3-L1 .......................................... 65

3.1.1.- Fundamento teórico ................................................................ 65

3.1.2.- Material ..................................................................................... 65

3.1.3.- Procedimiento experimental .................................................. 65

3.1.4.- Tratamientos ............................................................................ 66

3.2.- CULTIVO DE ADIPOCITOS HUMANOS ............................................. 67

3.2.1.- Fundamento teórico ................................................................ 67

3.2.2.- Material ..................................................................................... 68

3.2.3.- Procedimiento experimental .................................................. 68

3.2.4.- Tratamientos ............................................................................ 68

3.3.- MEDIDA DE LA LIPÓLISIS EN ADIPOCITOS 3T3-L1 ......................... 69

3.3.1.-Glicerol ...................................................................................... 70

3.3.2.-Acidos grasos libres ................................................................ 70

3.4.- DETERMINACIÓN DE LA EXPRESIÓN GÉNICA ............................... 70

3.4.1.1.- Procedimiento ................................................................................ 71

3.4.1.-Extracción y cuantificación de RNA ....................................... 71

3.4.2.- Tratamiento con DNAsa y retrotranscripción ....................... 72

3.4.3.- Determinación de los niveles de expresión génica mediante

la técnica q RT-PCR ................................................................................... 72

3.4.3.1.- Procedimiento ................................................................................ 73

3.5.- DETERMINACIÓN DE LOS NIVELES DE PROTEÍNA Y DE

MODIFICACIONES POST-TRANSCRIPCIONALES MEDIANTE WESTERN

BLOT….…………………………………..………………………………………..75

3.5.1.- Preparación de las muestras ................................................. 76

3.5.2.- Electroforesis y electrotransferencia .................................... 77

3.5.3.- Inmunoblot ............................................................................... 77

3.6.- ESTUDIO DE LOS NIVELES DE PGE2 MEDIANTE ELISA ................ 80

3.6.1.- Fundamento básico ................................................................ 80

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3.7.- ESTUDIO DE LOS NIVELES DE cAMP MEDIANTE ELISA ................ 80

3.7.1.- Fundamento básico ................................................................ 80

3.8.- EVALUACIÓN DE LA OXIDACIÓN DE LOS ÁCIDOS GRASOS EN

ADIPOCITOS 3T3-L1 ................................................................................... 81

3.9.- ANÁLISIS DE LA OXIDACIÓN DE LOS ÁCIDOS GRASOS EN

ADIPOCITOS SUBCUTÁNEOS HUMANOS................................................ 81

3.10.- MEDIDA DE LA INCORPORACIÓN DE ÁCIDOS GRASOS A

TRIGLICÉRIDOS EN ADIPOCITOS SUBCUTÁNEOS HUMANOS ............. 83

3.11.- MEDIDA DEL TRANSPORTE DE ACIDOS GRASOS EN

ADIPOCITOS SUBCUTÁNEOS HUMANOS................................................ 84

3.12.- ANÁLISIS DE LA LIPOGÉNESIS DE NOVO EN ADIPOCITOS

SUBCUTÁNEOS HUMANOS ....................................................................... 84

3.13.- CUANTIFICACIÓN DEL CONSUMO DE OXÍGENO EN ADIPOCITOS

SUBCUTÁNEOS HUMANOS ....................................................................... 85

3.14.- ESTUDIO DE LA BIOGÉNESIS MITOCONDRIAL ............................ 86

3.15.- CUANTIFICACIÓN DEL CONTENIDO MITOCONDRIAL MEDIANTE

LA TINCIÓN FLUORESCENTE MITOTRACKER GREEN EN ADIPOCITOS

SUBCUTÁNEOS HUMANOS ....................................................................... 86

3.16.- ANÁLISIS DEL CONTENIDO MITOCONDRIAL MEDIANTE

MICROSOCOPÍA ELECTRÓNICA DE TRANSMISIÓN (TEM) .................... 87

3.17.- DETECCIÓN DE UCP1 MEDIANTE INMUNOFLUORES-

CENCIA………………………………………….……………………………..….87

3.18.- ANÁLISIS ESTADÍSTICO .................................................................. 88

CHAPTER 4: RESULTS

4.1.- EFFECTS OF LIPOIC ACID ON LIPOLYSIS IN 3T3-L1 ADIPOCYTES

95

4.2.- α-LIPOIC ACID REDUCES FATTY ACID ESTERIFICATION AND

LIPOGENESIS IN ADIPOCYTES FROM OVERWEIGHT/OBESE

SUBJECTS ……………….. ...................................................................... 125

4.3.- α-LIPOIC ACID TREATMENT INCREASES MITOCHONDRIAL

BIOGENESIS AND PROMOTES BEIGE ADIPOSE FEATURES IN

SUBCUTANEOUS ADIPOCYTES FROM OVERWEIGHT/OBESE

SUBJECTS ………… ................................................................................. 143

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CHAPTER 5: GENERAL DISCUSSION/SUMMARY ................................. 171

CHAPTER 6: CONCLUSIONS ................................................................... 189

ANEXOS

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ABBREVIATIONS/ABREVIATURAS

α-LA- alpha-lipoic acid

alpha-MG- alpha-methylglucoside

AdPLA- adipose phospholipase A2

ACC- acetyl CoA Carboxylase

AC- adenylate cyclase

Akt- serine-threonine protein kinase Akt;

AMPK- adenosine monophosphate (AMP)-activated protein kinase

ATGL- adipose triglyceride lipase

BAT- brown adipose tissue

BHA- butylated hydroxyanisole

BMI- body mass index

C/EBPα- CCAAT/enhancer-binding protein alpha

C/EBPβ- CCAAT/enhancer-binding protein beta;;

CGI-58- comparative gene identification 58;

CILO- cilostamide

CRP- C reactive protein

CT- computed tomography

DGAT- Diglyceride O-acyltransferase

DGAT2- Diglyceride O-acyltransferase homolog 2

DHLA- dihydrolipoic acid

ERK- extracellular signal-regulated kinase

FA- fatty acids

FAS- fatty acid synthase

FFA- free fatty acids

G0S2- G0/G1 switch gene 2

GPAT-1- glycerol-3-phosphate acyltransferase-1

HSL- hormone-sensitive lipase

IGT- impaired glucose tolerance

IKKbeta- inhibitor of kappaB kinase-beta

IL-6- interleukin 6

i.p- intraperitoneal

IRSs- insulin receptor substrates

JNK- c-Jun NH(2)-terminal kinase

LASY- lipoic acid synthase

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LD- lipid droplet

LPL- lipoprotein lipase

LY- LY294002

MAGL- monoacylglycerol lipase

MAPKs- mitogen-activated protein kinases

MUFA- mono-unsaturated fatty acids

Myf5- myogenic factor 5

NAC- n-acetyl cysteine

NRF1 and NRF2- nuclear respiratory factors 1 and 2

OLEFT- Otsuka Long-Evans Tokushima Fatty

PAT- perilipin, adipophilin, TIP47

PD- PD98059

PDE3B- phosphodiesterase 3B

PET- positron emission tomography

PGC-1α- PPAR gamma coactivator 1 alpha

PGE2- prostaglandin E2

PI3-kinase- phosphatidylinositol 3-kinase

PKA- protein kinase A

PKB- protein kinase B

PKG- protein kinase G

PLA2- phospholipase A2

Plin1- perilipin

PPARγ- peroxixome proliferator-activated receptor gamma

ROS- reactive oxygen species

SAA- serum amyloid A

SCD1- stearoyl CoA Desaturase 1

SFA- saturated fatty acids

SP- SP600125

SREBP-1c- sterol regulatory binding protein-1c

SVF- stroma vascular fraction

TNF-α- Tumour Necrosis Factor α

TFAM- mitochondrial transcription factor A

UCP1- uncoupling protein 1

WAT- white adipose tissue

WHO- world Health Organization

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Introduction

1

CCHHAAPPTTEERR 11

IINNTTRROODDUUCCTTIIOONN

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Introduction

1

1.1. OBESITY

Obesity (defined as body mass index [BMI] > 30 kg/m2) is widely recognised

as a growing public health problem in developed and developing countries

(Holdsworth et al., 2012). The World Health Organization (WHO) predicts that by

2015, approximately 2.3 billion adults will be overweight and more than 700

million will be obese (Simonyi et al., 2012) (Fig. 1).

Obesity can be explained, in biological terms, as the consequence of

prolonged positive energy imbalance (energy intake exceeds energy

expenditure), leading to increased body fat mass. In addition, obesity is a

complex disease caused by a complicated network of contributory components,

such as genomic, epigenomic and also environmental factors such as sedentary

lifestyle and the increased consumption of high-calorie diets (Manco and

Dallapiccola, 2012).

Despite adipose tissue is vital for life as the major source of fatty acids (FA)

in the postprandial state for energy use and heat production, it is well known that

excess adipose tissue accumulation is associated with several comorbidities

such as cardiovascular diseases, type 2 diabetes mellitus, hypertension,

dyslipidaemia, liver steatosis and even various types of cancer. Thus, obesity is

a major determinant of premature mortality and a risk factor for the most

significant causes of death (Wyatt et al., 2006). In this context, it is mandatory to

look for strategies to prevent or to reduce obesity and its associated disorders in

an attempt to improve health as well as to reduce the medical expenses derived.

Fig. 1. Map representing the % of obese people (BMI ≥ 30) in different countries by

World Health Organization, 2013.

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Introduction

2

1.2. ADIPOSE TISSUE

It is well known that mammals have two distinct types of adipose tissue:

white and brown, both different in anatomy and function. However, during the

last years, a third type of adipose tissue has been described (Fig. 2). In fact,

beige or “brite” (also named inducible-brown or brown-like) adipocytes have

been found within certain white adipose depots, exhibiting similar molecular and

functional features as brown adipocytes (Wu et al., 2012).

Fig. 2. Differences between white, beige and brown adipose tissue (Spiegelman, 2013).

1.2.1. White adipose tissue

White adipose tissue (WAT) is a very heterogeneous tissue. It is composed

of several cell types: mature adipocytes and various other small cells (i.e.

preadipocytes, fibroblasts, endothelial cells, histiocytes and macrophages),

usually grouped as the ‘stroma vascular fraction’ (SVF) (Casteilla et al., 2011).

White adipocytes present a unique morphology, having a prominent lipid

droplet (LD) that occupies almost the entire cell volume, therefore, pushing other

intracellular compartments to the cell periphery (Le Lay et al., 2009). WAT

predominantly arise from non myogenic factor 5 (Myf5) lineages and for a long

time it has been considered to be only a passive organ for storage of

accumulated energy (Kopecky et al., 2004; Shan et al., 2013). However, and as

previously mentioned, WAT is an important endocrine/immune organ that

secretes adipokines, including inflammatory cytokines, chemokines, acute phase

proteins and complement-like factors (Medina-Gomez, 2012). These adipokines

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Introduction

3

may act as endocrine factors that regulate many central and peripheral

processes, including appetite, energy metabolism, glucose and lipid metabolism,

inflammatory responses, angiogenesis, blood pressure and reproductive

function. Furthermore, the distribution of body fat appears to be even more

important than the total amount of fat. Visceral adiposity has been strongly

linked to insulin resistance, type 2 diabetes, hypertension and dyslipidemia,

leading to increased risk of cardiovascular disease. However, such associations

seem to be much less consistent in subcutaneous fat mass (Palou et al., 2010).

1.2.2. Brown adipose tissue

The principal cell of brown adipose tissue (BAT) is the brown adipose cell

which contains large numbers of mitochondria that utilize energy substrates and

O2 to generate heat. BAT also contains brown adipose progenitors which are

capable of dividing and differentiate into brown adipose cells. BAT is innervated

by axons from sympathetic neurons that employ the neurotransmitter

norepinephrine (Mattson, 2010). BAT is derived from a Myf5-expressing cell

lineage (Seale et al., 2008).

Thus, BAT is specialized in adaptive thermogenesis in which the uncoupling

protein one (UCP1) plays a key role (Elabd et al., 2009). UCP1 is usually located

in the mitochondrial inner membrane of BAT adipocytes where it uncouples

substrate oxidation by the respiratory chain from ATP synthesis. Substrate

oxidation by the respiratory chain builds up a proton motive force by proton

pumping of the respiratory complexes. UCP1 allows the return of protons into

the matrix without ATP synthesis and thereby dissipates proton motive force as

heat. UCP1-mediated heat production plays an important role in non-shivering

thermogenesis in small rodents, hibernators and human infants (Klaus et al.,

2012). BAT is present throughout the life in rodents but disappears soon after

birth in large mammals. In humans, it is present in newborns to maintain body

temperature in a cold environment. Several recent studies have demonstrated

the existence of brown fat depots in adult humans (Greenhill, 2013; Sacks and

Symonds, 2013). Thus, the application of radiodiagnostic techniques (positron

emission tomography (PET)/computed tomography (CT)), coupled with histology

studies, to healthy humans have identified the presence BAT in humans after

relatively short exposure to mild cold. In fact, BAT has been identified at

supraclavicular, cervical, paraspinal, paraaortic and perirenal regions (Fig. 3)

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Introduction

4

being these BAT depots, metabolically active and it has been suggested that

BAT could contribute to significant energy expenditure upon activation in

humans. Moreover, it has been observed the amount of brown adipose tissue

inversely correlates with body mass index, being triglycerides stored within

brown adipocytes the major fuel for the enhanced metabolic activity of BAT

(Cypess et al., 2009; Richard et al., 2010; Ouellet et al., 2012).

1.2.3. Beige adipocytes

The process by which brown-like adipocytes appear at anatomical sites

characteristic of WAT has been called “browning”. These brown adipocytes are

now named as beige or brite adipocytes. This fact has been observed to occur

after certain stimulus, such as prolonged cold exposure. In contrast to BAT

adipocytes, beige adipocytes are not derived from the myf-5 linage (Wu et al.,

2012). Concerning the origin of these beige adipocytes, there are controversial

hypothesis, One possibility suggest that the brite adipocytes may be recruited de

novo from specific precursor cells within WAT (Wu et al., 2012; Rosenwald et al.,

2013). Alternatively, other possibility is that britening occurs through a direct

interconverison of a white adipocyte into a brown-like phenotype, a process

referred as transdifferentiation (cinti, 2002). In this context, a recent study has

shown bi-directional interconversion of brite and white adipocytes upon

cold/warm stimulation (Rosenwald et al., 2013).

Beige adipocytes have all the morphological and molecular characteristics of

classical brown adipocytes, thus, these are multilocular, express inducible UCP1

having therefore thermogenic characteristics, and have increased mitochondrial

respiratory machinery. However, it has been recently described that beige

adipocytes express several beige adipocyte-specific genes that are not

expressed in classical brown adipocytes such as Tbx1, Tmem26 and CD137

among others (Shan et al., 2013). As mentioned before, beige adipocytes

appearance might be induced by cold exposure, physical activity and several

agents such as the PPARγ activators thiazolidinediones or the recently

discovered miokine irisin, a soluble factor which is a proteolytic fragment of the

type I membrane protein FNDC5 (Bostrom et al., 2012; Ohno et al., 2012).

There are also other master transcription factors such as PGC-1α, C/EBPβ and

PRDM16 that ultimately have been demonstrated to play key roles in this

browning process (Petrovic et al., 2010). Thus, PRDM16 is highly present in

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Introduction

5

brown adipocytes when compared with white adipocytes, and moreover,

suppresses classic white adipocyte genes, and promotes the transcription of

several proteins involved in thermogenesis in WAT including PGC-1α and

UCP1. The enhancement of mitochondrial biogenesis and the brown-like

phenotype within WAT have been proposed as a promising strategy to combat

obesity and its associated disorders (Liu et al., 2009; Bartelt and Heeren, 2013)

Fig. 3. Anatomical sites of brown, white and beige adipocytes in mice and humans (Bartelt

and Heeren, 2013).

1.3. WHITE ADIPOSE TISSUE AS A KEY STORAGE ORGAN

WAT is an outstanding tissue in several aspects. Adipocytes store energy in

the form of triglycerides in situations of energy surplus and they are able to

hydrolyze these triglycerides into free fatty acids (FFA) and glycerol in energy

demand conditions such as stress, exercise and/or fasting (Rajala and Scherer,

2003). Lipid accumulation (lipogenesis) and breakdown (lipolysis) are tightly-

regulated processes in a dynamic equilibrium, responding to the different stimuli

existing during fasting and refeeding. Thus, lipid droplets are highly dynamic

organelles whose metabolic functions can be grouped as 1) catabolic reactions,

which involves hydrolysis, mobilization and further metabolism of triglycerides

and 2) anabolic reactions, including fatty acid synthesis, activation, and

esterification into trigliceride molecules (Ducharme and Bickel, 2008).

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Introduction

6

1.3.1. Lipolysis in adipocytes

During fasting periods triglycerides are hydrolyzed into FFA and glycerol in

order to be used as energetic substrate and therefore to satisfy energy

demands. FFA mobilization occurs through the consecutive action of three

lipases: adipose triglyceride lipase (ATGL), hormone-sensitive lipase (HSL) and

monoacylglycerol lipase (MAGL) (Krintel et al., 2008). ATGL initiates lipolysis by

cleaving the first FFA from triglycerides and then HSL and MAGL act on

diacyglycerol and monoacylglycerol, respectively, releasing two additional FA

and one glycerol molecule (Gaidhu et al., 2010). ATGL and HSL are considered

as the major rate-determining enzymes in adipocyte lipolysis (Miyoshi et al.,

2008). Increased FFA has been related to detrimental metabolic consequences

such as insulin resistance and metabolic syndrome (Ormseth et al., 2013).

However, several studies evidenced that increased lipolysis in adipocytes is

frequently associated with an increased FFA utilization via β-oxidation

suggesting, therefore, that stimulation of lipolysis might be a useful strategy to

prevent or treat obesity (Ahmadian et al., 2009).

The main regulation of the enzymes and proteins involved in lipolysis is

briefly described in the following sections.

Fig. 3: Lipolysis regulation during A) fasting and B) feeding (Ahmadian et al., 2010).

1.3.1.1. Adipose Triglyceride Lipase (ATGL)

ATGL (also called desnutrin and calcium-independent phospholipase A2

[iPLA(2)] zeta) was identified as a 2.0 kb mRNA expressed mainly in adipose

tissue. Atgl over-expression was shown to increase triglyceride hydrolysis.

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7

Several studies have demonstrated that Atgl expression is down-regulated in

ob/ob and db/db obese mice, as well as in diet-induced overweight animals.

(Jenkins et al., 2001; Villena et al., 2004; Zimmermann et al., 2004). In this

sense, further studies revealed that ATGL deficiency caused triglycerides

accumulation in non-adipose tissues such as heart, leading to severe cardiac

insufficiency, or in other tissues, such as testis, kidney or liver. Other

consequences of ATGL deficiency were a severe defect in thermoregulation and

a decrease in oxygen consumption in mice, suggesting a reduction in energy

expenditure. These findings highlight that ATGL plays an important role in

energy homeostasis (Haemmerle et al., 2006; Smirnova et al., 2006).

Furthermore, ATGL has been shown to be regulated by nutritional status. Thus,

Atgl mRNA levels increased during fasting but are donwn-regulated during

refeeding in mice (Villena et al., 2004; Kralisch et al., 2005; Kershaw et al.,

2006). Insulin, an antilipolitic hormone, is also a major ATGL regulator in

adipocytes by inhibiting Atgl mRNA expression both in vitro and in in vivo

models (Kim et al., 2006; Ogasawara et al., 2012).

Different mechanisms have been proposed to be involved in the regulation of

ATGL activity. Thus, several studies have reported post-transcriptional

phosphorylation modifications in ATGL. In this context, it was shown that Ser303

AMP-activated protein kinase (AMPK)-mediated phosphorylation was able to

reduce lipolysis during fasting. However, further studies in vitro found that

murine ATGL activity is activated by post-transcriptional phosphorylation on

Ser406

mediated by both AMPK and β-Adrenergic-mediated cAMP-dependent

protein kinase A (PKA) activation (Narbonne and Roy, 2009; Ahmadian et al.,

2011). ATGL is also known to be regulated by protein-protein interactions. In

adipose tissue, comparative gene identification-58 (cgi-58; also called Abdh5)

encodes a 39-kDa protein of the alpha/beta hydrolase domain subfamily which

binds to intracellular lipid droplets by interaction with perilipin (Subramanian et

al., 2004; Yamaguchi et al., 2004). CGI-58 induces the ATGL hydrolase activity.

It is important to mention that CGI-58 has not a specific lipase activity itself but it

is able to specifically activate ATGL enzyme activity (Lass et al., 2006). Thus,

mutations in CGI-58 result in triglycerides accumulation in several tissues,

leading to serious pathologies including cardiomyopathy and liver steatosis

(Lefevre et al., 2001). However, controversial results have been found in

different studies about the role of CGI-58 on obesity. Thus, in mice fed on a high

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Introduction

8

fat diet, Cgi-58 overexpression did not prevent the development of obesity

(Brown et al., 2010). On the contrary, Cgi-58 knock-down prevented HFD-

induced obesity and insulin resistance (Caviglia et al., 2011). Another protein

directly involved in ATGL regulation is the protein encoded by the G0/G1 switch

gene 2 (G0S2). It selectively down-regulates ATGL enzymatic activity by direct

interaction with the catalytic patatin-like domain of ATGL. G0S2 is highly

expressed in mature adipocytes and in basal conditions binds ATGL regulating

its activity and preventing lipid droplet degradation. When lipolysis is stimulated,

G0S2 gene expression is turned off, therefore, the ratio ATGL/G0S2 and

subsequently ATGL activity increases (Yang et al., 2010). Interestingly, it has

been found that G0S2 does not compete with CGI-58 in binding ATGL.

Moreover, it possesses the capacity to prevent ATGL-mediated lipid droplet

triglycerides turnover in the presence of CGI-58. (Lu et al., 2010; Yang et al.,

2010).

1.3.1.2. Hormone Sensitive Lipase (HSL)

Adipose tissue HSL has been traditionally considered as the key enzyme

catalyzing the rate-limiting step of adipose tissue lipolysis. It is a cytoplasmic

protein with demonstrated hydrolase activity against a wide variety of substrates

including triglycerides, diglycerides, cholesteryl esters, and retinyl esters;

however, the relative hydrolase activity of HSL in vitro is 10-fold greater against

diglycerides than against triglycerides (Raclot et al., 1997). The activity of HSL is

regulated post-translationally by phosphorylation–dephosphorylation reactions.

Activation of its expression by lipolytic molecules (catecholamines, isoproterenol,

glucagon, adrenocorticotropic hormone) involves an increase in the intracellular

concentration of cAMP, which activates PKA. PKA phosphorylates HSL,

resulting in 2- to 100-fold increases in hydrolytic activity. Dephosphorylation is

affected primarily by protein phosphatases (Lobo et al., 2009).

Mouse HSL is phosphorylated in three sites by PKA (Ser563

, Ser659

, and

Ser660

; corresponding to human Ser552

, Ser649

and Ser650

) and at a single site by

extracellular signal-regulated kinase (ERK) (Ser600

; Ser589

in humans) in

response to lipolysis activators (Liu et al., 2011; Lorente-Cebrian et al., 2011).

Moreover, the phosphorylation of both Ser659

and Ser660

by PKA is necessary for

activated lipolysis and the translocation of HSL to the lipidic droplet surface

(Watt et al., 2006). HSL is also phosphorylated on Ser565

(Ser554

in humans) by

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Introduction

9

AMPK in unstimulated adipocytes. The precise role of HSL Ser565

phosphorylation by AMPK remains unclear. Thus, although PKA and AMPK-

mediated phosphorylations are believed to be mutually exclusive, it has been

observed that PKA-mediated activation of AMPK activity is important for

converting a lipolytic signal into an effective lipolytic response (Anthony et al.,

2009; Djouder et al., 2010).

1.3.1.3. Perilipin

Lipid droplets are bounded by a protein-decorated phospholipid monolayer

that envelopes a neutral lipid core. The PAT (Perilipin, adipophilin, TIP47) family

proteins, including perilipin (Plin1), are the best-studied LD-associated proteins

(Bickel et al., 2009). Perilipin is primarily expressed in adipose tissue and

controls adipocyte lipolysis by directly regulating the activity of the lipases

surrounding the droplet (Mottagui-Tabar et al., 2003). Thus, under basal

conditions, perilipin inhibits lipolysis by blocking lipase access to triglycerides

and/or sequestering CGI-58 avoiding the interaction with ATGL, thereby

maintaining a low rate of basal lipolysis (Miyoshi et al., 2006; Granneman et al.,

2009). Upon lipolytic stimulation, PKA phosphorylates perilipin and this facilitates

HSL translocation to the lipid droplet surface and releases the ATGL activator

CGI-58 (Sztalryd et al., 2003; Lass et al., 2006). These perilipin-mediated

actions were highlighted by the use of the perilipin-null mouse. These mice have

chronically elevated basal lipolysis in adipose tissues or primary adipocytes with

increased activity of HSL and ATGL (Zhai et al., 2010). Therefore, as perilipin

null mice are not able to produce a regular response to lipolytic signals, exibiting

a marked reduction in stored adipose mass when fed on a high fat diet, while

showing an increased tendency to develop glucose intolerance and peripheral

insulin resistance (Tansey et al., 2001; Sztalryd et al., 2003).

1.3.1.4. AdPLA

Other molecules have been described to regulate lipolysis. Thus, Duncan et

al., (2008) discovered a novel intracellular phospholipase A2 (PLA2) highly

expressed in adipocytes that releases FA (mainly arachidonic acid) from

phosphatidylcholine in a Ca++

-dependent manner (Wolf, 2009). Arachidonic acid

has been implicated in prostaglandin E2 (PGE2) production, which is known to

down-regulate lipolysis by decreasing cAMP levels, which highlights the

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Introduction

10

importance of AdPLA in adipose tissue and lipolysis (Duncan et al., 2008).

Moreover, further studies revealed that AdPLA-null mice had increased HSL-

phosphorylation, suggesting that HSL phosphorylation through cAMP-mediated

activation of protein PKA is probably a key mediator of increased lipolysis in

these mice (Jaworski et al., 2009).

1.3.1.5. Signaling pathways involved in lipolysis

Adipocyte lipolysis is a tightly regulated process which is of central

importance in the maintenance of whole body energy homeostasis (Koppo et al.,

2012).

Catecholamines are the main hormones involved in lipolysis regulation.

Through β adrenergic receptor stimulation, catecholamines induce lipolysis and

breakdown of FA. These hormones binding to Gs coupled receptors, results in

an increased adenylate cyclase (AC) activity, which leads to an increase in

intracellular cAMP concentrations resulting in activation of cAMP-dependent

PKA (Jaworski et al., 2007). The pro-lipolytic effect of PKA is carried out by both

increasing the HSL intrinsic activity and promoting its access to triglycerides

(Brasaemle et al., 2000; Su et al., 2003).

In contrast, insulin is an important (if not the primary) inhibitor of adipose

tissue lipolysis and FA release into the bloodstream. An increase in insulin

concentration suppresses lipolytic rates and decreases plasma FFA

concentrations, whereas a decrease in insulin concentration leads to

accelerated lipolysis and increased plasma FFA concentrations (Magkos et al.,

2010). Upon insulin binding, the insulin receptor becomes tyrosine

phosphorylated (auto phosphorylation), and recruits insulin receptor substrates

(IRSs) for tyrosine phosphorylation. IRSs bind and activate phosphatidylinositol

3-kinase (PI3-kinase) and other downstream molecules like protein kinase B

(PKB). Then PKB/AKT acts inducing the phosphorylation and activation of

phosphodiesterase 3B (PDE3B) leading to hydrolysis of cAMP and attenuation

of the lipolytic pathway (Buren et al., 2008; Berggreen et al., 2009).

AMPK is a heterotrimeric enzyme consisting of a catalytic α subunit and two

regulatory subunits, β and γ. α-AMPK acts as a fuel sensor that regulates

glucose and lipid homeostasis in adipocytes and its activation leads to numerous

metabolic changes (Hardie, 2007). AMPK contains a serine/threonine protein

kinase catalytic domain that is activated by phosphorylation of the Thr172

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Introduction

11

(Dzamko et al., 2010). AMPK activation has been shown to exert an antilipolytic

effect in WAT (Gaidhu et al., 2010). In vitro studies on purified bovine adipocyte

HSL have shown that AMPK phosphorylates HSL on Ser565

, thereby abolishing

PKA-induced HSL activation (Jocken et al., 2008). In contrast, some studies

show clearly that AMPK activation is required for the maximal increase in

lipolysis induced by isoproterenol. These studies also revealed a mechanism by

which a rise in cAMP levels can lead to an increase in AMPK activity (Yin et al.,

2003; Djouder et al., 2010). Moreover, AMPK has been observed to

phosphorylate ATGL at Ser406

promoting its triglycerides hydrolase activity both

in cultured adipocytes and in in vivo models (Ahmadian et al., 2011).

Other proteins involved in lipolysis regulation are the mitogen-activated

protein kinases (MAPKs). These are serine-threonine kinases that mediate

intracellular signaling associated with a variety of cellular activities including cell

proliferation, differentiation, survival, death, and transformation (Yun et al., 2011;

Choi et al., 2013). In addition, ERK activation causes HSL phosphorylation in the

residue Ser600

, located in the middle of the regulatory module and this

phosphorylation by ERK produces a hydrolytic activity similar to that observed

when HSL is phosphorylated by PKA (Greenberg et al., 2001; Carmen and

Victor, 2006).

Moreover, other signaling pathways have been involved in the control of the

lipolytic response. Thus, Activation of cGMP pathway has also been shown to

promote lipolysis. A downstream effector of cGMP, cGMP-dependent protein

kinase, also called protein kinase G (PKG), was shown to induce perilipin and

HSL phosphorylation and to be at the origin of atrial natriuretic peptide-induced

lipolysis (Sengenes et al., 2005).

1.3.2. Triglycerides storage: lipogenesis

Triglycerides storage in adipocytes can be the result of 1) dietary FA uptake

and subsequent esterification within the cell or 2) de novo FA biosynthesis from

non lipid substrates. This last process accounts less than 25% in human

adipocytes and although, it specially takes place in adipose tissue and liver, it

has been observed to be carried out in almost all cell types (Strawford et al.,

2004).

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Introduction

12

Glucose

Fig. 4. Lipogenesis process in adipocytes (modified from Targett-Adams et al., 2010)

1.3.3. Fatty acid esterification

Triglycerides that are going to be stored in adipose tissue are mainly

acquired from circulating lipoproteins, chylomicrons and VLDL. These

triglycerides require intravascular hydrolysis by lipoprotein lipase (LPL) to

liberate FFA which are then uptaken by adipocytes through specific FA

transporters such as FATP, FABP or CD36 (Febbraio et al., 2001; Garcia-Arcos

et al., 2013). Once the FFA are inside the adipocyte they are esterified into

triglycerides and stored in the lipid droplet.

3.3.1. Diglyceride O-acyltransferase (DGAT)

DGAT catalyzes the final step in triglycerides synthesis by facilitating the

linkage of diacylglycerol with long-chain acyl-CoA FA. DGAT exists in two pri-

mary isoforms: DGAT1 and DGAT2; DGAT1 is most highly expressed in small

intestine and white adipose tissue, whereas DGAT2 is primarily expressed in

liver and white adipose tissue where its expression is insulin-responsive (Choi et

al., 2007).

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13

There is some evidence suggesting that the two enzymes play different roles

in triglycerides metabolism. Dgat1 knockout mice have 50% less triglycerides in

tissues and are protected from diet-induced obesity and insulin resistance

through a mechanism involving increased energy expenditure (at least partly

attributable to increased physical activity) (Smith et al., 2000; Chen et al., 2002).

The study of Dgat2 has been more difficult as homozygous DGAT2 knockout

mice die shortly after birth because of severe lipopenia (90% triglycerides

reduction in Dgat2 null carcasses) and impaired skin barrier function (Stone et

al., 2004).

In fact, genetic deletion of Dgat1 in mice revealed that the knockout mice

have reductions in triglycerides accumulation in many tissues, including adipose

tissue, when fed a high-fat diet (Smith et al., 2000). Dgat1-/-

mice have normal

weight when these are fed with standard chow and resist to weight gain when

fed on a high fat diet. Fat pads were also smaller than those of wild type mice

and the levels of triglycerides were reduced in both WAT and muscle. These

animals where shown to eat more and have increased metabolic rate which was

shown after indirect calorimetry and increased activity, which suggest that the

hyperphagia could be a mechanism to compensate the increase on energy

expenditure (Smith et al., 2000; Chen et al., 2002). It has been also described

that Dgat1 delection decreased insulin resistance because Dgat1–/–

mice

required an approximately 20% higher glucose infusion rate than Dgat1+/+

mice

to maintain euglycemia in hyperinsulinemic-euglycemic clamp studies. They

have also increased leptin sensitivity because the same doses of leptin caused

an additional 3% weight loss in age-matched Dgat1–/–

mice (Chen et al., 2002).

Moreover, curiously, this anti-obesity protection was accompanied by increased

mean and maximal life spans. On the other hand, mice over-expressing Dgat1 in

adipose tissue had larger adipocytes and increased fat depots when fed on a

standard diet and became 20% more obese when these were fed on a high fat

diet; similar effects that were also observed when Dgat1 is overexpressed in

other tissues such as skeletal muscle (Chen et al., 2002; Li et al., 2013).

1.3.4. De novo lipogenesis

De novo lipogenesis is thought to be a relatively minor contributor to whole

body lipid stores in a present-day human consuming a typical high fat diet

(McDevitt et al., 2001). However, pharmacologic or genetic manipulation of

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Introduction

14

enzymes in the lipogenic pathway can have profound metabolic consequences

(Postic and Girard, 2008), suggesting that de novo lipogenesis might serve a

signaling function independent of the generation of lipid stores (Lodhi et al.,

2011). Moreover, recent studies in rodents demonstrated that in obesity de novo

lipogenesis, in WAT is altered and that restoring de novo lipogenesis selectively

in WAT reverts obesity-dependent insulin resistance (Huo et al., 2012).

Taking together, these facts highlight the importance of the enzymes

involved in de novo lipogenesis (Acetyl CoA Carboxylase and Fatty acid

synthase; ACC and FAS respectively) and mono-unsaturated fatty acids (MUFA)

synthesis (Stearoyl CoA Desaturase, SCD1) analyzed in this study.

1.3.4.1. Acetyl CoA Carboxylase (ACC)

ACC is the enzyme that catalyzes the rate-limiting step in FA synthesis in

lipogenic tissues, including WAT, mammary gland, and liver. It catalyzes the

conversion of acetyl-CoA to malonyl-CoA (Peng et al., 2012) (Fig. 4). Malonyl-

CoA is an initial substrate for de novo FA biosynthesis, but it also serves as a

potent inhibitor of carnitine palmitoyl transferase 1 (CPT1), a rate-limiting step for

FA oxidation (McGarry and Brown, 1997). Acc knockout mice, showed similar

weight gain than wild type mice when fed with a standard diet. However under

lipogenic conditions (high fat feeding or similar) lipid accumulation in adipose

tissues of Acc knockout mice was significantly decreased in comparison with the

controls. Regarding ACC regulation, it is well known that activation of AMPK

phosphorylates and inhibits ACC activity and it has been proposed that ACC

inhibition is one of the mechanisms that could explain the ability of AMPK to

inhibit FA synthesis, in response to decreases in energy supply (Zhou et al.,

2001). Thus, studies in isolated rat adipocytes reported that AICAR (an AMPK

activator) inhibited lipogenesis but stimulated phosphorylation of ACC at Ser79

(ACC1) or Ser212

(ACC2). ACC1 is mainly expressed in lipogenic tissues such as

WAT. Isolated rat adipocytes infected with adenoviruses expressing

constitutively active mutant AMPK showed increased phosphorylation of ACC1

Ser79

whereas infection with dominant negative mutant AMPK inhibited AICAR-

stimulated phosphorylation of Ser79

, indicating that ACC1 is certainly regulated

by AMPK in adipocytes (Daval et al., 2005). ACC2 isoform is mainly expressed

in muscle, but its expression in the liver and several other organs is also of

significant importance. Thus, Acc2-/-

mice showed continuous FA oxydation,

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Introduction

15

reduced fat storage and improved insulin sensitivity, although lipogenesis in

these mice was not affected, indicating that ACC2 is not required for progression

of FA synthesis (Abu-Elheiga et al., 2001).

1.3.4.2. Fatty Acid Synthase (FAS)

FAS is an enzyme required for the conversion of acetyl-CoA and malonyl-

CoA to saturated fatty acid (SFA) (Fig. 4). Fas is highly expressed in tissues like

liver, fat, and lactating mammary glands. Interestingly, all human tissues show a

minimal expression of Fas. De novo FA synthesis is essential for embryonic

development (Jayakumar et al., 1995; Chirala et al., 2003). Dysregulation of

FAS has been recognized to be involved in obesity and diabetes for a long time.

Thus, Fas expression in adipose tissue links excess energy intake with

accumulation of body fat predominantly visceral fat, impaired insulin sensitivity

and dysregulation of circulating adipokines (Zeng et al., 2011). The FA synthesis

pathway has become a therapeutic target for ameliorating the adverse effects of

obesity as well as its associated disorders such as type 2 diabetes and

cardiovascular disease. In this context, the use of different FAS inhibitors

revealed several beneficial effects. Thus, among the anti-obesity effects of C75,

a synthetic inhibitor of FAS, it was observed an increase in interscapular brown

adipose tissue, which is an important site for thermogenesis. Cerulenin markedly

improved hepatic function in steatotic ob/ob mice by decreasing serum AST/ALT

levels which was accompanied by fat depletion in the hepatocyte (Cassolla et

al., 2013; Cheng et al., 2013). In this line, mice with Fas deficiency specifically in

adipose tissue manifested increased energy expenditure, increased brown fat-

like adipocytes in subcutaneous adipose tissue, and resistance to diet-induced

obesity. Moreover, mice were protected from deterioration of glucose

homeostasis and hepatic steatosis induced by high-fat diet (Wueest et al., 2010;

Lodhi et al., 2012).

1.3.4.3. Stearoyl CoA Desaturase 1 (SCD1)

SCD1 is an endoplasmic reticulum enzyme which is the rate-limiting enzyme

in the biosynthesis of MUFA. It catalyzes the synthesis of MUFA from SFA,

playing an important role in the synthesis of triglycerides. SCD1 activity

decreases with starvation and insulin deficiency but increases with diets

enriched in SFA (Dobrzyn and Ntambi, 2004). It has been shown that whole

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Introduction

16

body Scd1 knockout mice had reduced body fat and exhibited resistance to diet

induced-obesity. In fact, Scd1−/−

mice have markedly reduced epididymal fat pad

and when fed on a high-fat diet, the livers of the wild-type mice are steatotic,

whereas the Scd1−/−

livers remain healthy. Moreover, Scd1−/−

mice showed an

increase in energy expenditure, oxygen consumption and plasma ketone bodies

which suggests an increase in lipid oxidation. In addition, these mice have an

increased sensitivity to insulin as well as reduced insulin and leptin plasma

levels (Ntambi et al., 2002). However, adipose specific (on WAT and BAT) Scd1

deletion showed no significant differences in body weight or in epididymal or

subcutaneous white adipose weights in both mice fed a standard or a high fat

diet. Food intake and liver weights were not significantly altered, as well as,

fasting glucose and insulin tolerance which suggests that Scd1 deletion from

adipose tissue is insufficient to elicit protection from obesity (Flowers et al.,

2012).

1.4. MITOCHONDRIA AND OBESITY

The mitochondrion is a multifunctional organelle that is responsible for many

cellular functions including energy generation, cell growth (Nuñez et al., 2006),

and cell death (Green and Reed, 1998). Highly dynamic, these organelles

respond to changes in the cellular environment. Several studies have suggested

that mitochondrion plays a major role in influencing individual susceptibility to

obesity. In fact, adipocyte mitochondrial dysfunction has been involved in obesity

and type 2 diabetes (Rong et al., 2007).

Several studies have revealed that mitochondria in obese conditions have

lower energy-generation capacities, less clearly defined inner membranes and

reduced FA oxidation in comparison with lean individuals (Blaak et al., 2006;

Naukkarinen et al., 2013). In this context, it has been suggested that modulating

mitochondrial function, biogenesis and antioxidant status is a nutritional issue of

great interest in the treatment or prevention of several diseases such as obesity

(reviewed by, Perez-Matute et al., 2009).

1.4.1. Mitochondrial biogenesis

Modulation of mitochondrial biogenesis, the formation of mitochondria during

the life cycle of a cell, has been proposed as a therapeutic tool against obesity

(Mustelin et al., 2008). In fact, the abundance of gene transcripts encoding

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Introduction

17

mitochondria proteins decreases with the onset of obesity and it has been

described that in human subcutaneous WAT, the abundance of the

mitochondrial copy number from obese is lower than in those from lean subjects

(Mustelin et al., 2008). Moreover, this lower mitochondrial content, is usually

associated with reduced mitochondrial function and therefore decreased FA β-

oxidation, leading to increased FA accumulation which contributes to the

development of obesity associated comorbidities such as insulin resistance

(Monsenego et al., 2012). For this reason, there is a high level of interest in

developing therapeutic strategies aimed to modulate the regulatory pathways

that increase mitochondrial function and biogenesis in an attempt to prevent or

to treat obesity (Scarpulla et al., 2012).

Mitochondrial mass can be modulated through several stimuli and cellular

pathways including hormones such as thyroid and estrogen or inflammatory

stimuli which requires the orchestrated expression of diverse transcription

activators, including PPAR gamma coactivator 1 alpha (Pgc-1α) (Alvarez-

Guardia et al., 2010; Gleyzer and Scarpulla, 2011; Weitzel and Iwen, 2011;

Ivanova et al., 2013).

Once activated the stimulatory effects of PGC-1α on mitochondrial genes are

achieved through the activation of nuclear respiratory factors 1 and 2 (NRF1 and

NRF2, respectively). These are nuclear respiratory factors which are potent

stimulators of the expression of the mitochondrial transcription factor A (Tfam;

mainly NRF-1). Thus, TFAM translocates to mitochondrial matrix, where it

stimulates mitochondrial DNA replication and mitochondrial gene expression

triggering mitochondrial duplication (Scarpulla, 2006).

PGC-1α activity is known to be regulated by post-translational modification

by the energy sensors: AMPK and Sirtuin 1 (SIRT1) (Fig. 5). SIRT1, a NAD+-

dependent protein deacetylase that has emerged as a key metabolic sensor in

various metabolic tissues, is activated in response to metabolic challenges such

as dietary restriction or exercise due to the increase in cellular NAD+ levels

(Canto and Auwerx, 2009). SIRT1 activates PGC-1α by deacetylation and

thereby drives mitochondrial biogenesis (Scarpulla, 2011). Moreover, PGC-1α is

also activated by another important metabolic sensor, AMPK (Jager et al.,

2007). Thus, restricted nutrient supply increases the cellular AMP/ATP ratio,

which is sensed by AMPK. As a consequence, PGC-1α is activated through

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Introduction

18

AMPK-mediated phosphorylation and stimulates mitochondrial biogenesis and

energy metabolism (Canto and Auwerx, 2009).

Fig. 5: Control of mitochondrial biogenesis by AMPK/SIRT1/PGC-1α signaling pathway

(Brenmoehl and Hoeflich, 2013).

1.4.2 Mitochondrial function in the regulation of FA metabolism

Adipose tissue supplies FFA as fuel to tissues and organs that can oxidize

them in order to obtain energy. However, obesity-mediated increase on lipolysis,

and therefore, FFA release triggers the onset of muscle insulin resistance and

fatty liver by the accumulation of triglycerides on these tissues (Ebbert and

Jensen, 2013). However, mitochondria, through the modulation of FA oxidation

might act buffering the daily flux of circulating FFA as a preventive strategy

against these diseases. AMPK also plays a central role in the regulation of FA

oxidation. AMPK-mediated phosphorylation and inactivation of ACC reduces

malonyl-CoA concentrations and therefore lipogenesis but, increases FA

oxidation (Hardie et al., 2012). In the same line, PGC-1α also mediates an

improvement in mitochondrial function by promoting the

phosphorylation/inactivation and reduced expression of ACC (Lee et al., 2006).

Regarding CPT1, a protein anchored in the outer membrane of mitochondria

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Introduction

19

that catalyses the formation of long chain fatty acyl-carnitine, which is enabled to

traverse the inner mitochondrial membrane and thus committed to β oxidation in

the mitochondria (Nyman et al., 2005; Ji et al., 2008). The essential role of

CPT1A has been inferred by several studies. In patients with unusual

spontaneous mutations in the Cpt1 gene, hepatic deficiency of CPT1 leads to

recurrent episodes of hypoketotic hypoglycaemia, hepatomegaly, seizures and

coma (Fontaine et al., 2012; Roomets et al., 2012). In adipocytes, elevated

expression of Cpt1, protected from FA-induced insulin resistance and

inflammation, and it has been observed that in subcutaneous adipose tissue

CPT1 levels are negatively correlated with body fat percentage (Gao et al.,

2011; Zhang et al., 2013). In addition, mice with deleted Cpt1 are more sensitive

to high-fat diet-induced insulin resistance (Wolfgang et al., 2006; Gao et al.,

2009). Taking together these studies suggest that CPT1 activation might

represent a promising strategy for the prevention and treatment of obesity

related metabolic diseases.

1.5. OBESITY AND OXIDATIVE STRESS

Several studies revealed a significant positive relationship between

increased body mass index (BMI) and oxidative stress. Expanded dysfunctional

adipose tissue in obesity is associated to increased reactive oxygen species

(ROS), which leads to impaired adipogenesis, macrophage recruitment and

activation, the secretion of inflammatory adipokines and the damage of

biological structures (Picklo et al., 2012). Cross-promotion of oxidative stress

and inflammation creates a feed-forward cycle that can initiate and advance

disease progression. Indeed, this oxidative stress-derived inflammation has

been hypothesized to be a major mechanism in the pathogenesis and

progression of obesity-related disorders (Bryan et al. 2013.). Based on this

hypothesis, the use of antioxidants, such as α-lipoic acid (α-LA), could

ameliorate this situation, via suppression of oxidative stress, inflammatory

cytokines and macrophage infiltration, and, in consequence, other obesity-

associated disorders.

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Introduction

20

1.6. α-LIPOIC ACID

α-lipoic acid (5-(1,2-dithiolan-3-yl)-pentanoic acid) or thioctic acid is a natural

occurring compound. α-LA is a small molecule (eight carbons; 206.3 g/mol) that

contains two oxidized or reduced thiol groups, and a single chiral center, which

results in two possible optical isomers: R-α-LA and S-α-LA (Fig. 6). Both α-LA

and its reduced dithiol form, dihydrolipoic acid (DHLA), are powerful

antioxidants. α-LA was first isolated from bovine liver in 1951 (Reed et al., 1951),

and it is present in several nutritional sources; thus, animal tissues such as liver

and kidney contain the highest concentrations of α-LA, but it can also be found

in vegetables such as spinach, broccoli and tomato, brussel sprouts and rice

bran (Packer et al., 2001).

Fig. 6. Molecular structure of R-α-LA, S-α-LA, R-DHA and S-DHA

α-LA is found naturally in mitochondria, where it acts as the coenzyme for

several bioenergetic enzymes such as pyruvate dehydrogenase and α-

ketoglurarate dehydrogenase (Packer et al., 1995). α-LA synthesis was believed

to be carried out only by prokaryotic microorganisms, however, nowadays the

identification of a mouse cDNA encoding a lipoic acid synthase, named LASY

and located in mitochondria, provided evidence that α-LA is also synthesized

from octanoic acid in mammals (Morikawa et al., 2001). R-α-LA is the isomer

that is synthesized by plants and animals and functions as a cofactor for

mitochondrial enzymes in its protein-bound form. However, when synthesized in

a laboratory, α-LA is a racemic mixture of the R- and S- forms, and most studies

are carried out with this mixture of both isomers. Some trials comparing the

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Introduction

21

effects of both α-LA stereoisomers have also been performed (Yaworsky et al.,

2000; Moini et al., 2002).

As mentioned before, it is well known that α-LA possesses antioxidant

properties, thus, it scavenges hydrogen peroxide, single oxygen, hydroxyl

radical, nitric oxide radical, and peroxynitrite, and has the capacity to reduce the

oxidized forms of several important antioxidants, including glutathione, vitamins

C and E. α-LA may also chelate redox-active metals, such as free iron, copper,

manganese and zinc. Together, these properties make α-LA a potentially highly

effective therapeutic antioxidant (Packer et al., 2001; Suh et al., 2004; Suh et al.,

2005). Thus, α-LA has been considered a good candidate for the treatment of

those diseases involving inflammation and oxidative damage. In this context, α-

LA could act as a neuroprotective agent, and has been proposed as a promising

treatment for neurological disorders such as Parkinson’s disease and Alzheimer

by increasing activity of energy producing mitochondrial enzymes and down-

regulating the oxidative stress associated to aging and these related disorders

(Arivazhagan et al., 2001; De Araujo et al., 2011). α-LA could be also an

effective therapy for diabetic neuropathy and multiple sclerosis (Yadav et al.,

2012). It has also been reported that α-LA can improve endothelial function

through a decrease of oxygen-derived free radicals (Xiang et al., 2011). α-LA

causes renal-morphologic improvement in streptozotocin-induced diabetic

nephropathy (Kanter et al., 2010). In end-stage renal disease, patients under

hemodialysis showed a reduction of C reactive protein (CRP) levels after α-LA

supplementation (Nanayakkara and Gaillard, 2010). α-LA supplementation could

be also a potential adjuvant for several oxidative stress associated disorders

such as vascular disease, asthma and/or rheumatoid arthritis or obesity (Shay et

al., 2009).

1.6.1. Antiobesity effects of α-LA

1.6.1.1. Animal studies

Several studies have been carried out aiming to demonstrate the putative

beneficial effects of α-LA on obese animal models. Although, many of them

evidenced body weight loss and/or adiposity lowering effects, others reported

neutral effect (Table 1). The study of Kim et al. Kim et al., (2004a) was the first to

report the anti-obesity effects of α-LA in different experimental models of obesity

in rodents. Thus, dietary supplementation with α-LA (0.25, 0.5 and 1%, w/w of

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Introduction

22

diet) for 2 weeks caused a dose-dependent reduction on food intake and body

weight in rats fed on a standard diet. More importantly, the anorexigenic and

body fat lowering actions of α-LA supplementation (0.5% w/w, 14-28 weeks)

were also observed in models of genetically obesity and diabetes rats such as

the Otsuka Long-Evans Tokushima Fatty (OLETF) rats (Kim et al., 2004; Song

et al., 2005). Interestingly, the anti-obesity properties of α-LA are independent of

leptin or leptin receptor signaling, since the reduction in food intake and fat mass

was also observed in leptin deficient (Lep−/−

) or leptin receptor–deficient (Lepr−/−

)

mice (Kim et al., 2004) as well as in the obese Zucker rats, a recessive trait

(fa/fa) of the leptin receptor (Butler et al., 2009; Yi et al., 2013).

Other studies have provided evidence that dietary supplementation with α-LA

(0.25-0.5% w/w, 8 weeks) is able to prevent the body weight gain and increased

adiposity induced by exposure to a obesogenic environment (high fat diet)

(Prieto-Hontoria et al., 2009; Timmers et al., 2010). The inclusion of pair-fed

groups in some of the previously mentioned studies and the finding that α-LA-

treated animals weighed significantly less than the pair-feds has revealed that α-

LA not only reduce energy intake also promote energy expenditure (Kim et al.,

2004; Prieto-Hontoria et al., 2009). Moreover, α-LA antiobesity effects (200

mg/kg of b.w., 4 weeks) have been also observed in streptozotocin/nicotinamide

C57BL/6J diabetic-induced mice fed on a high fat diet (Chen et al,. 2012).

Other studies have revealed the efficacy of α-LA treatment on body weight

regulation when administered by injection. Thus, intraperitoneal injection of α-LA

(30 mg/kg b.w.) during 8 weeks to ALS/Lt mice, a type 2 diabetes mouse model,

inhibited progression of type 2 diabetes, reducing body weight gain and

adiposity (Mathews et al., 2004). Ovariectomy-induced obese rats mediated by

estrogen deficiency also showed a down-regulation in body weight gain and

adiposity, accompanied by a reduction in food intake after intraperitoneal

treatment with α-LA (200 mg/kg b.w., 7 weeks) (Cheng et al., 2011).

However, other studies reported neutral effects on body weight after α-LA

administration. In this context, the study of Banday et al. (Banday et al., 2007)

showed no effects on body weight or food intake after dietary supplementation

with 0.4% α-LA for 2 weeks in obese zucker rats, although beneficial effects on

parameters associated with obesity onset such as, oxidative stress were

observed in the same study. The study of Cummings et al. (Cummings et al.,

2010) carried out in the type 2 diabetes model UCD-T2DM rats revealed that

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dietary α-LA supplementation (80 mg/kg b.w., 8 weeks) delayed diabetes onset

in fructose-fed animals, without affecting body weight and fat depot weights, or

food intake. Furthermore, no anorexigenic and body lowering effects were found

in obese Zucker rats treated i.p. with α-LA (92 or 30 mg/kg b.w. for 22 and 2

weeks respectively), but a marked down-regulation in muscle triglyceride

accumulation and oxidative damage was observed (Saengsirisuwan et al., 2004;

Muellenbach et al., 2009).

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Table 1. Effects of LA administration on animal obesity models

Species

Durati

on Intervention Effects observed

Beneficial effects on body weight

Kim et al. (Kim et al., 2004a) Male Sprague-

Dawley rats

2

weeks

Dietary administration of standard rat

chow containing α-LA (0.25, 0.5 and 1%,

w/w)

↓ food intake and body weight

OLETF rats 14

weeks

Dietary administration of standard

chow ± α-LA (0.5%, w/w)

↓ body weight and visceral adipose tissue

Lep-/- and Lepr-/-

mice

6 days Dietary administration of standard

chow ± α-LA (0.5%, w/w)

↓ food intake and body weight

Song et al. (Song et al., 2005) OLETF rats 28

weeks

Dietary administration of standard

chow ± α-LA (0.5%, w/w)

↓ body weight

Mathews et al. (Mathews et al., 2004) ALS/Lt diabetic mice 8

weeks

Intraperitoneal (i.p) injection α-LA (30

mg/kg)

↓ body weight and white adipose tissue.

Prieto-Hontoria et al. (Prieto-Hontoria et al.,

2009)

Wistar rats 8

weeks

Dietary administration of standard or

high fat chow ± α-LA (0.25%, w/w). Pair

fed groups included

↓ food intake, body weight, white adipose

tissue, and feed efficiency.

Butler et al. (Butler et al., 2009) Obese male Zucker

rats

5

weeks

Standard chow ± α-LA (2.4 g/kg of

diet).

↓ food intake, visceral adipose tissue and

hypertriglyceridemia

Timmers et al. (Timmers et al., 2010) Male Wistar rats 8

weeks

Low or High fat chow ± α-LA (0.5%,

w/w)

↓ food intake, body weight and white adipose

tissue

Cheng et al. (Cheng et al., 2011) Sprague-Dawley 7 Ovariectomy-induced obesity ± α-LA ↓ food intake, body weight, white adipose

tissue,

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25

rats weeks (200 mg/kg b.w., i.p. injection

Chen et al. (Chen et al., 2012) Streptozotocin/NA-

induced diabetes

C57BL/6J mouse

4

weeks

High fat chow ± α-LA (50 and 200

mg/kg b.w., orally)

↓ visceral adipose tissue and body weight in

mice supplemented with 200 mg/kg α-LA

db/db mice 90

days

Standard chow ± α-LA (200 mg/kg

b.w., orally)

↓ visceral adipose tissue and body weight

Yi et al. (Yi et al., 2013) Obese male Zucker

rats

2

weeks

Dietary administration of standard

chow ± α-LA (3g/kg of diet; approximate

200 mg/kg body weight per day) or Pair

fed group

↓ hypertriglyceridemia and abdominal fat mass.

Neutral effects on body weight

Saengsirisuwan et al. (Saengsirisuwan et al.,

2004)

Obese Zucker rats 2

weeks

α-LA (30 mg/kg)i.p.injection ↓ muscle triglyceride accumulation

Banday et al. (Banday et al., 2007) Lean and obese

male Zucker rats

2

weeks

Standard chow ± α-LA (0.4 %) ↓ oxidative stress and hypertension

Muellenbach et al. (Muellenbach et al., 2009) Obese Zucker rats 22

weeks

α-LA (92 mg/kg)i.p.injection

↓ muscle triglyceride accumulation and

oxidative damage

Cummings et al. (Cummings et al., 2010) UCD-T2DM rats 24

weeks

Dietary administration of standard rat

chow containing or not fructose (20% of

energy) ± α-LA (80 mg/kg b.w.)

↓ oxidative stress

↑ glucose-stimulated insulin secretion

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26

1.6.1.2 Human clinical trials

Although many α-LA-mediated beneficial effects have been reported in

animal models there is little evidence demonstrating the same benefits of α-LA

supplementation on obese subjects. In fact, the existing studies reveal

controversial outcomes (Table 2). Thus, the first study carried out including

overweight/obese subjects with schizophrenia showed that treatment with α-LA

(1200 mg/day) during 12 weeks reduced body weight and BMI, effects that were

accompanied by a decrease in total cholesterol. Moreover, the authors

described a mild to moderate reduction of appetite as the most obvious effect of

α-LA treatment (Kim et al., 2008). A more recent study also suggests that α-LA

is a good candidate for obesity therapy. Thus, Carbonelli et al. (Carbonelli et al.,

2010) studied in 1127 caucasian subjects the effects of α-LA supplementation

(800 mg/day) during four months. The data demonstrate the ability of α-LA to

decrease body weight due to a diminished fat mass and appetite. The

randomization of the volunteers according to sex and BMI revealed that the body

weight lowering effects of α-LA in the group of normoweight subjects (BMI < 25

Kg/m2) were only observed in women. However, in overweight (BMI 25 ≤ BMI ≥

30 Kg/m2) and obese subjects (BMI > 30), the effects were similar for both sex,

and a significant reduction in BMI, weight, waist circumference and appetite was

found. Moreover, this was accompanied by decreases in several inflammatory

parameters such as Interleukin 6 (IL-6), Tumour Necrosis Factor α (TNF-α) or C

reactive protein (CRP).

The study of Koh et al., (2011) aimed to study the efficacy of α-LA as an

adjuvant for body weight loss in subjects following an energy restricted diet. This

randomized, double-blind, placebo-controlled study was carried out in Asian

obese and overweight subjects with hypertension, diabetes mellitus, or

hypercholesterolemia. α-LA (1200 or 1800 mg/day) was administered orally

accompanied by a dietary restriction of 600 kcal/day during 20 weeks. At the end

of the trial, the mean body weight and waist circumference reduction was

significantly greater in the 1800 mg/d α-LA group than in the placebo group.

Furthermore, the amelioration of obesity was accompanied by an improvement

of associated co-morbidities, and those individuals with type 2 diabetes showed

a reduction on haemoglobin-A1c when compared with baseline (Koh et al.,

2011).

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27

However, not all the studies have reported beneficial actions of α-LA on body

weight loss. Thus, no effects were described after 1 g/day of α-LA

supplementation during 12 weeks on 24 (12 males, 12 females) obese subjects

diagnosed with impaired glucose tolerance (IGT). However, it is important to

point out that, in this study, α-LA only produced a significant decrease in body fat

and in waist and hip circumference when α-LA supplementation was

accompanied by exercise (McNeilly et al., 2011). Another trial carried out in a

cohort of 12 female and 10 male obese with IGT and 6 female and 4 male obese

with normal glucose tolerance subjects of Chinese ethnicity found no significant

differences in BMI and waist circumference among groups were found after 2

weeks of α-LA parenteral administration (600 mg/day). The authors assumed

that it is possible that further differences might occur if the study would be

extended on time (Zhang et al., 2011). Taking together all of these trials, the

effects of α-LA supplementation in obese patients remain unclear and seems to

be highly dependent on several factors such as the length of treatment, dose,

associated diseases or drug prescriptions of the patients. Therefore, more

controlled clinical trials are necessary to better clarify the safety of α-LA

supplementation at higher effective doses or for longer periods of treatment as

therapy for human obesity in different populations.

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Introduction

Table 2. Effects of α-LA on human obese subjects

Study

Design Participants Intervention

Lengh

t of

treatment

Gluco

se

tolerance

Effects observed

α-LA

Kim et al. (Kim et al., 2008) Open

prospective

clinical trial

5

Obese/overweight

Male/female

Schizophrenic patients

under antipsychotic treatment

1200 mg/day

administered orally in

divided doses.

12

weeks

Norma

l

↓ BMI, body weight and total cholesterol

Carbonelli et al. (Carbonelli et al., 2010) Open

prospective

clinical trial

1127 Caucasian,

male/female

(normoweight, overweight

and obese)

800 mg/day by

orally supplementation

16

weeks

Norma

l

↓ BMI, body weight, waist circumference,

appetite, systolic blood pressure.

↓ Inflammatory markers (Erythrocyte

Sedimentation rate, TNF-α and IL-6)

Koh et al. (Koh et al., 2011) Randomize

d, double-blind,

placebo-

controlled trial

228

Asian male/female

obese and overweight

1200 and 1800

mg/day orally

supplemented and

combined with 600

kcal/day restriction

20

weeks

Norma

l and Type

2 diabetes

↓ Body weight and waist circumference

for 1800 mg/d of α-LA.

↓ Hemoglobin-A1c in diabetic subjects.

Zhang et al. (Zhang et al., 2011) Randomize

d, double-blind,

placebo-

controlled trial

32

Asiatic male/female

Obese

600 mg/day

administered

intravenously once

daily

2

weeks

Norma

l/Impaired

none

McNeilli et al. (McNeilly et al., 2011) Balanced,

randomized

controlled trial

24

Caucasian male/female

Obese

1000 mg/day

orally supplemented

12

weeks

Impair

ed

α-LA: none

α-LA + exercise: ↓ Body fat and waist and

hips circumference

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Introduction

29

1.6.2. Mechanisms of action

1.6.2.1. α-LA reduces food intake and increases energy expenditure

The antiobesity effects of α-LA have been partly attributed to its anorexigenic

action, both in animal models as well as in humans (Kim et al., 2004; Prieto-

Hontoria et al., 2009; Carbonelli et al. 2010). Hypothalamic AMPK seems to

function as a major regulator of food intake and energy expenditure (Stark et al.,

2013), and it has been demonstrated that both central (3 µg) and peripheral (75

mg/kg b.w., single dose) administration of α-LA decreases hypothalamic AMPK

activity in the arcuate nucleus of the hypothalamus and causes profound weight

loss in rodents by reducing food intake (Kim et al., 2004). In addition, α-LA also

stimulates whole-body energy expenditure in rodents, which is accompanied by

an enhancement of UCP1 expression in brown adipose tissue (BAT). The

stimulatory effect of α-LA on energy expenditure seems to be mediated by the

central nervous system since the i.c.v. administration of very small amounts of α-

LA (0.3 and 3 µg) was sufficient to produce these effects. Moreover,

hypothalamic AMPK has also a key role in this process since α-LA action on

energy expenditure is abolished by the i.c.v. administration of the AMPK

activator AICAR (Kim et al., 2004). Unlike to the stimulation of glucose uptake by

muscle and adipocytes, α-LA dietary supplementation (0.25% w/w of diet, 8

weeks) significantly decreased intestinal α-methylglucoside (α-MG) absorption

both in lean and obese rats (Prieto-Hontoria et al., 2009), which could contribute

to the lower feed efficiency found in α-LA-treated animals and therefore to its

beneficial effects of on obesity.

In addition to these central effects on food intake and energy expenditure,

some studies have suggested that α-LA (0.75% α-LA in drinking water for one

month) could also promote energy expenditure and loss of body weight acting

peripherally by AMPK activation and fat oxidation in peripheral tissues such as

muscle (Wang et al., 2010). This dual effect of α-LA on hypothalamic and

muscle AMPK has been also described for hormones and cytokines that

regulate body weight such as leptin and CNTF (Minokoshi et al., 2002; Watt et

al., 2006).

1.6.2.2. α-LA inhibits adipogenesis

It has been demonstrated that α-LA exerts anti-adipogenic effects in 3T3-L1

adipocytes (Cho et al., 2003). In fact, α-LA (250-500 µM) is able to inhibit

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Introduction

troglitazone (a PPARγ agonist)-mediated differentiation of 3T3-L1 adipocytes by

lowering the expression levels of adipogenic master transcription factors such as

Pparγ and C/ebpα. α-LA also down-regulated the gene expression of genes

characteristic of the early phase of differentiation (mitotic clonal expansion) such

as c-fos and c-jun, and also decreased activities of aP-1, C/EBPβ and δ, and

CREB. Moreover, it has been described that the activation of MAPKs mediates

the inhibitory effect of α-LA on the adipogenic process. Importantly, the same

study also evidenced that α-LA is able to dedifferenciate 3T3-L1 mature

adipocytes by antagonizyng PPARγ effects (Cho et al., 2003).

1.6.2.3. α-LA improves glucose metabolism

One of the main beneficial effects of α-LA supplementation observed in

obese subjects is the amelioration of impaired glucose tolerance. Indeed, the

ability of α-LA to improve whole body glucose disposal and insulin sensitivity and

to prevent type 2 diabetes development has been observed in several models of

obesity in rodents (Lee et al., 2005; Song et al., 2005). These beneficial actions

of α-LA on glucose metabolism are in part secondary to its body weight lowering

actions and especially by its ability to reduce lipid accumulation in non-adipose

(muscle and pancreas) as well as in adipose tissue (Lee et al., 2005). However,

other studies both in rodents and humans have reported improved glucose

tolerance and insulin sensitivity without significant changes in body weight and

adiposity, suggesting that the mechanisms by which α-LA regulates glucose

utilization in the organism may be multifactorial. Thus, it has been widely

demonstrated that α-LA increase both basal and insulin-stimulated glucose

uptake by skeletal muscle (Eason et al., 2002; Bitar et al., 2004) in several

models of obese diabetic rodents. Lee et al. (Lee et al., 2005) described that α-

LA-induced (0.5% w/w mixed in food for 3 days) increase in insulin-stimulated

glucose uptake is mediated by activation of AMPK. Gupte et al. (Gupte et al.,

2009) described that α-LA (30 mg/kg b.wt., i.p., 6 weeks) reduced

phosphorylation of JNK and reduced the inhibitor of kappaB kinase-beta

(IKKbeta) activity (IkappaBalpha protein levels) in rats fed a high fat diet. This

study also demonstrated that α-LA effectively restored insulin responsiveness

and insulin-stimulated glucose uptake in soleus muscle. Studies in cultured L6

myotubes and 3T3-L1 adipocytes observed that the dose-dependent stimulatory

effect of R-α-LA (1-5 mM) on basal and insulin-stimulated glucose uptake was

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Introduction

31

associated with an intracellular redistribution of GLUT1 and GLUT4 glucose

transporters. It was also dependent on phosphatidylinositol 3-kinase (PI3K)

activity (Estrada et al., 1996; Yaworsky et al., 2000). Other study in adipocytes

has also suggested that R-α-LA (250 µM) modulates glucose uptake by

changing the intracellular redox status. This study also found that all oxidized

forms of α-LA (S-, R-,and racemic α-LA) were able to stimulate glucose uptake,

whereas the reduced form, DHLA, was ineffective (Moini et al., 2002).

The insulin-sensitizing properties of α-LA have been related to its ability to

promote the activation of the insulin signaling pathway at different steps. Thus,

α-LA (100 mg/kg i.p., 4 weeks) has been shown to promote the activation of

IRS1 and PI3K/AKT soleus muscle from rats (Yaworsky et al., 2000; Bitar et al.,

2004). α-LA (300 µM) also induced the activation of p38 MAPK, a protein

previously implicated in insulin-independent glucose uptake in cultured L6

myoblasts (Gupte et al., 2009).

In addition to promoting glucose uptake in several tissues, α-LA might also

facilitate glucose oxidation by acting as a cofactor of several mitochondrial

enzymes. α-LA (2.4 mM) treatment has been shown to increase glucose

oxidation in isolated rat soleus muscle (Dicter et al., 2002). In this context,

Konrad et al. (Konrad et al., 1999) described that α-LA treatment (600 mg, twice

a day orally, 4 weeks) decreases serum lactate and pyruvate concentrations and

improves glucose effectiveness in lean and obese patients with type 2 diabetes

and suggested that increased aerobic glucose oxidation could also contribute to

α-LA-induced amelioration of insulin resistance.

6.2.4. Regulation of adipokine production

Nowadays, it is clear the importance of white adipose tissue-derived factors

(adipokines) in the development of obesity and systemic insulin resistance.

Indeed, adipocytes secrete multiple bioactive peptides, such as leptin,

adiponectin, and apelin that play key roles in the regulation of energy

metabolism and insulin sensitivity (Scherer, 2006). Deregulated adipokine

secretion from the expanded adipose tissue of obese individuals contributes to

the development of obesity-linked disorders including systemic insulin resistance

and metabolic disease (Prieto-Hontoria et al., 2011).

Obesity is usually accompanied by hyperleptinemia, which has been

associated with the development of type 2 diabetes, cardiovascular diseases

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Introduction

and cancer (Prieto-Hontoria et al., 2011). Several studies in obese rodents have

found that α-LA reduces leptin circulating levels in parallel with the reduction of

body weight and fat mass (Huong and Ide, 2008; Cheng et al., 2011 ; Prieto-

Hontoria et al., 2011; Jung et al., 2012), suggesting a potential inhibitory effect of

α-LA on leptin secretion by adipocytes and a potential mechanism that explains

its beneficial actions on obesity and related disorders. In this way, the study of

Prieto-Hontoria et al. (Prieto-Hontoria et al., 2011) corroborated that α-LA

caused a concentration-dependent (1-500 µM) inhibition of leptin secretion and

gene expression mediated at least in part by an inhibition of the transcription

factor Sp1 activity in 3T3-L1 adipocytes.

Adiponectin is an insulin-sensitizing adipokine that regulates glucose and

lipid metabolism and its levels are inversely correlated with adiposity. Some

studies have described an increase in adiponectin circulating levels and mRNA

levels in adipose tissue after dietary supplementation (0.2 - 5 g/kg of diet, 2-16

weeks) with α-LA (Prieto-Hontoria et al., 2012; Huong and Ide, 2008),

suggesting that increased adiponectin could also contribute to the metabolic

actions and insulin-sensitizing properties of α-LA (Prieto-Hontoria et al., 2012).

However, the stimulatory action of α-LA on adiponectin in vivo was not

reproduced in cultured adipocytes (Prieto-Hontoria et al., 2011). Furthermore,

other studies did not find this increase in adiponectin after α-LA (0.08-0.2 g/kg

b.w., 7-8 weeks) supplementation (Cummings et al., 2010; Cheng et al., 2011).

Thus, more studies are needed in this regard.

Apelin is another adipokine up-regulated in obesity and insulin resistance

(Cavallo et al., 2012). However, apelin stimulates glucose utilization and

increases FA oxidation, mitochondrial oxidative capacity, and mitochondrial

biogenesis in muscle of insulin-resistant mice (Dray et al., 2008; Attane et al.,

2012). α-LA (250 µM) has been shown to stimulate apelin secretion in 3T3-L1

adipocytes, although no changes in this adipokine levels have been described

after dietary supplementation with α-LA (0.25% w/w of diet, 8 weeks) in high fat

fed rats (Fernandez-Galilea et al., 2011).

α-LA treatment also modulates different proinflammatory adipocytokines. For

example TNF-α, IL-6 and chemerin were all significantly decreased in human

and rodents plasma levels after dietary supplementation with the antioxidant

(Zhang et al., 2011; Jung et al., 2012 ; Yu et al., 2012), indicating the potential

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Introduction

33

therapeutic role of α-LA in preventing the inflammatory state associated with

obesity and oxidative stress.

1.6.2.5. Effects of α-LA on lipid metabolism

Several studies have evidenced that α-LA is able to prevent/decrease lipid

accumulation not only in adipose tissue but also in other non-adipose key

metabolic organs like skeletal muscle (Lee et al., 2005) and liver (Park et al.,

2008; Huong e Ide, 2008; Valdecantos et al., 2012a) . Thus, different trials in

rodents support the potential protective effect of α-LA supplementation against

the development of nonalcoholic fatty liver associated with a long-term high-fat

diet (Huong and Ide, 2008; Valdecantos et al., 2012a). In this context, the study

of Huong and Ide (Huong and Ide, 2008) revealed that dietary administration of

α-LA (1-5 g/kg of diet for 21 days) in rats, was able to dose-dependently

decrease triglycerides concentration in plasma and liver, in parallel with the

reduction of the activity and mRNA levels of hepatic lipogenic enzymes such as

FAS, ATP-citrate lyase, glucose-6-phosphate dehydrogenase, malic enzyme

and pyruvate kinase. In other study, Park et al. (Park, et al., 2008) have also

demonstrated that that α-LA supplementation (0.5% w/w mixed in food for 3

days) in rats significantly reduced lipid accumulation in liver by decreasing sterol

regulatory binding protein-1c (SREBP-1c), one of the major regulators of the

expression of genes involved in hepatic triglyceride synthesis, in part via AMPK

activation, but also in part by inhibiting the activities of LXR and Sp1, mediators

of insulin-dependent Srebp-1c expression. Moreover, α-LA inhibited the insulin-

stimulated expression of the SREBP-1c target genes such as Acc and fas which

leads to completely prevention of hepatic steatosis. Recently, Valdecantos et al.,

(2012a) also described that α-LA (0.25% w/w of diet during 8 weeks) prevented

hepatic triglyceride accumulation and liver damage in rats fed a high-fat diet

through a down-regulation of lipogenic genes such as Dgat2, a gene directly

involved in triglycerides synthesis in liver. The study of Butler et al. (Butler et al.,

2009) also revealed that feeding Zucker Diabetic Fatty rats ad libitum with α-LA

(2.4 g/kg of diet for 5 weeks) decreased VLDL-triglycerides secretion and the

mRNA levels of key enzymes of triglycerides synthesis, glycerol-3-phosphate

acyltransferase-1 (Gpat-1), and Dgat2, Acc1, Acc2, and Fas. Moreover, it

recently has been described that the reduction of hepatic lipogenic enzymes

(ACC and FAS) after dietary supplementation (3g/kg of diet) in obese zucker

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Introduction

rats was also observed in epididymal fat depot (Yi et al., 2013). Thus, toguether

these results strongly suggested that α-LA inhibits carbohydrate to fat

conversion, triglycerides synthesis and VLDL production.

Apart from the inhibition of lipogenic enzymes above descrited, another

mechanism that could explain the reduction of lipid accumulation in liver muscle

in adipose tissue is the breakdown and elimination of triglycerides existing in

these depots. Lipolysis is a highly regulated complex process, which involves

the co-ordinately participation of several lipases and lipid droplet proteins

(Ahmadian et al., 2009). ATGL and HSL are the major triglyceride lipases in

many tissues such adipose tissue and liver (Reid et al., 2008). Atgl and Hsl

expression are decreased in the obese, insulin-resistant state, suggesting that

insulin resistance is associated with impaired lipolysis (Jocken et al., 2007;

Huijsman et al., 2009). Activation of lipolysis has been proposed as a promising

therapeutic target for the treatment of obesity (Jaworski et al., 2009).

Several studies have demonstrated the lipolytic properties of α-LA both in

vivo and in vitro. Thus, Hamano (Hamano, 2002) observed that dietary α-LA

supplementation (100 mg/kg of diet for 30 days) stimulates rapid lipolytic

response of plasma FFA to clenbuterol injection and FA turnover between

adipose tissue and liver in chickens. In other study, Hamano (Hamano, 2006)

also described that dietary supplementation with α-LA (400 mg/kg of diet for 5

weeks) increased plasma free glycerol, and an increase in the rate of free

glycerol release from abdominal adipose tissue explants was observed when

compared with the non-α-LA supplemented group. Lipolysis is not a process

carried out exclusively in adipocytes (Badin et al., 2012). A recent study

described that α-LA (0.25 to 1.0 mM) decreased intracellular triglycerides

content in fatty liver cell model developed by incubating HepG2 cells in high

glucose, high fat media. This lipid lowering activity was accompanied by an

increase of AMPK phosphorylation and ATGL protein levels and a reduced

FOXO1 phosphorylation, suggesting the increase observed in lipolysis could be

due to the AMPK/FOXO1 pathway (Kuo et al., 2012). Other study has also

shown that the lipid-lowering effect of α-LA was associated with increased ATGL

protein production in liver and skeletal muscle of both STZ/NA-induced and

db/db diabetic mice (Chen et al., 2012). These data suggest that α-LA could also

promote lipolysis on non-adipose tissues; however, control mechanisms seem to

be tissue specific.

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Introduction

35

Increased lipolysis and FFA release from adipose tissue has been

associated with the development of insulin resistance (Ormseth et al., 2011;

2013). However, recent findings have demonstrated that increasing lipolysis in

adipose tissue does not necessarily increase serum FFA levels because

increasing lipolysis in adipose tissue causes a shift within adipocytes and other

metabolic tissues toward increased FFA utilization and energy expenditure and

thus protects against obesity (Zhang et al., 2011).

Therefore, another mechanism that could prevent lipid accumulation by α-LA

supplementation is the increase of the oxidative metabolism. Several studies

support the ability of α-LA to improve mitochondrial function because of its role

as coenzyme for several bioenergetic enzymes (Packer et al., 1995). In this

context, α-LA might be useful to increase overall mitochondrial metabolism.

Thus, promotion of mitochondrial FA β-oxidation might avoid detrimental effects

of fat accumulation on liver or muscle.

Wang et al. (Wang et al., 2010) described that treatment of C2C12 myotubes

with α-LA (1 mM) significantly increased FA β-oxidation, which was associated

with increased phosphorylation of AMPK and expression of Pgc-1α. A more

recent study also support that α-LA (300 µM) increased palmitate β-oxidation

and decreased intracellular triacylglycerol accumulation through induction of

SIRT1 and subsequent activation of AMPK and ACC phosphorylation in C2C12

myotubes (Chen et al., 2012). In liver, it has been shown that dietary

supplementation with α-LA (0.25% w/w of diet, 8 weeks) was able to reverse the

decrease in Cpt1a, Acadl, and Acox1 gene expression, all of them involved in

mitochondrial and peroxisomal β-oxidation induced by high fat feeding

(Valdecantos et al., 2012b). By DNA microarray analysis, Yang et al. (Yang et

al., 2008) showed that α-LA supplementation (0.1% of diet, 6 weeks)

upregulated the expression of genes related to beta-oxidation, such as acyl-

coenzyme A dehydrogenase Acad1, Acox and Cpt1, in liver of high fat-fed mice,

also supporting the ability of α-LA to promote FA oxidation and to prevent the

high-fat diet–induced dyslipidemia.

Mitochondrial dysfunction, including mitochondrial loss and over-production

of oxidants, has been suggested to be involved in the development of metabolic

disorders including obesity and insulin resistance (Rong et al., 2007; Hojlund et

al., 2008). Several studies support that α-LA is able to protect mitochondrial

function and inhibit mitochondrial damage associated to obesity in different

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Introduction

tissues. Thus, Valdecantos et al., (2012b) described that dietary

supplementation with α-LA (0.25% w/w of diet, 8 weeks) induced an increase in

liver mitochondrial copy number in obese high-fat fed rats. This was

accompanied by an increase in mitochondrial antioxidant defenses and a

reduced oxidative damage in mitochondrial DNA through the deacetylation of

Foxo3a and PGC-1β by SIRT1 and SIRT3 respectively (Valdecantos et al. ,

2012b). Other studies in adipocytes have shown that α-LA (100 µM) also

increased the number and mitochondrial mass per cell, the mitochondrial DNA

copy number as well as the protein levels and expression of key transcription

factors involved in mitochondrial biogenesis, including PGC-1α, TFAM and

NRF1 (Shen et al., 2011). Moreover, Wang et al., (2010) observed that α-LA

(0.75% in drinking water for one month) stimulated skeletal muscle mitochondrial

biogenesis with increased phosphorylation of AMPK and PGC-1α levels in aged

mice.

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Introduction

37

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Introduction

53

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Introduction

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Hypothesis and Aims

57

Alpha-lipoic acid (α-LA) is a natural occurring antioxidant compound with

anti-obesity properties in both rodents and humans (Prieto-Hontoria et al., 2009;

Carbonelli et al., 2010). The anti-obesity actions of α-LA have been related to its

ability to reduce food intake and to increase energy expenditure (Kim et al.,

2004). In addition, adipose tissue has emerged as a key target in the body

weight lowering and insulin-sensitizing actions of α-LA (Fernandez-Galilea et al.,

2011; Prieto-Hontoria et al., 2013;).

Adipose tissue is the main energy store (as triglycerides) in the body.

Malfunction of the synthesis or degradation of fat stores within adipocytes is

associated with several prevalent diseases with high public health impact such

as obesity and type 2 diabetes (Lottenberg et al., 2012). In fact, an unbalance

between lipid storage (lipogenesis) and lipid mobilization (lipolysis) occurs in

obesity. Thus, several strategies have been proposed to restore this balance, by

reducing lipid accumulation and/or by promoting lipolysis, in an attempt to

prevent or to treat obesity (Ahmadian et al., 2009; Lodhi et al., 2012).

While white adipose tissue (WAT) functions as an energy storage organ,

brown adipose tissue (BAT) is an energy consumption organ. The principal

function of brown adipocytes is to burn fat in their abundant mitochondria to

generate heat. In this context, the enhancement of mitochondrial biogenesis and

the brown-like phenotype within WAT have been proposed as a promising

strategy to combat obesity and its associated disorders (Liu et al., 2009; Bartelt

and Heeren, 2013).

For all previously described, the hypothesis of the present Doctoral Thesis

was that α-LA beneficial effects on adiposity could be due to its ability to

modulate triglyceride metabolism in white adipocytes and to promote

mitochondrial biogenesis and brown-like phenotype within these white

adipocytes.

The main objective of the present study was to deeply analyze the direct

effects of α-LA on triglycerides storage/mobilization pathways and on

mitochondrial biogenesis and browning-induction in white adipocytes (murine

and human) and to investigate the underlying mechanisms.

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Hypothesis and Aims

58

The specific aims were:

1. To determine the effects of α-LA treatment on lipolysis and on the

regulation of the main lipases and lipid droplet proteins involved in the

lipolytic pathway in cultured white adipocytes.

2. To investigate α-LA actions on FA esterification and de novo

lipogenesis in cultured white adipocytes.

3. To evaluate the actions of α-LA on mitochondrial biogenesis and

to study its ability to induce brown-like features in cultured white

adipocytes.

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Hipótesis y Objetivos

59

El ácido alfa-lipoico (α-LA) es un compuesto antioxidante natural con

propiedades anti-obesidad en roedores y humanos (Carbonelli et al., 2010;

Prieto-Hontoria et al., 2009). Las acciones anti-obesidad del α-LA han sido

asociadas a su capacidad para reducir la ingesta y aumentar el gasto energético

(Kim et al., 2004). Además, el tejido adiposo se ha revelado como un tejido

diana clave de los efectos del ácido lipoico sobre la pérdida de peso y las

propiedades de sensibilización a la insulina (Prieto-Hontoria et al., 2013;

Fernandez-Galilea et al., 2011).

El tejido adiposo es el principal órgano de almacenamiento de energía (en

forma de triglicéridos) del organismo. Disfunciones en la síntesis o degradación

de dichos lípidos almacenados en el adipocito se asocian a enfermedades de

alta prevalencia e impacto en la salud pública como son la obesidad y la

diabetes tipo 2 (Lottenberg et al., 2012). De hecho, en la obesidad se produce

un desequilibrio entre el almacenamiento de triglicéridos (lipogénesis) y la

movilización de los mismos (lipólisis). En este sentido, se han propuesto

diversas estrategias para reestablecer este equilibrio mediante la reducción de

la acumulación de triglicéridos y/o mediante la promoción de la lipólisis en un

intento de prevenir o tratar la obesidad (Ahmadian et al., 2009; Lodhi et al.,

2012).

En contraposición a la función de almacenamiento de energía del tejido

adiposo blanco, el tejido adiposo pardo es un tejido termogénico. Así, la

principal función del adipocito pardo es disipar energía a través de sus

abundantes mitocondrias para generar calor. En este contexto, un aumento de

la biogénesis mitocondrial así como de factores característicos del tejido

adiposo pardo en el tejido adiposo blanco ha sido propuesta como una

estrategia prometedora para combatir la obesidad y sus comorbilidades

asociadas (Liu et al., 2009; Bartelt and Heeren, 2013).

Por todo lo hasta aquí descrito, la hipótesis en la que se fundamenta el

presente trabajo de Tesis Doctoral es que los efectos beneficiosos del α-LA

sobre la adiposidad podrían ser debidos a su capacidad para modular el

metabolismo de los triglicéridos en los adipocitos blancos así como a su posible

habilidad para promover la biogénesis mitocondrial y la inducción de

marcadores y características fenotípicas propias del adipocito pardo en

adipocitos blancos.

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Hipótesis y Objetivos

60

El principal objetivo del presente trabajo fue analizar en profundidad los

efectos directos del α-LA en las rutas de almacenamiento/movilización de

triglicéridos y en la biogénesis mitocondrial y en la inducción de pardeamiento

en adipocitos blancos (murinos y humanos) e investigar los mecanismos

subyacentes.

Los objetivos específicos fueron:

1. Determinar los efectos del tratamiento con α-LA sobre la lipólisis y

la regulación de las principales lipasas y proteínas asociadas a la

gota lipídica involucradas en la ruta lipolítica en adipocitos blancos

en cultivo.

2. Investigar las acciones del α-LA sobre la esterificación de los ácidos

grasos y la lipogenesis de novo en adipocitos blancos en cultivo.

3. Evaluar las acciones del α-LA sobre la biogénesis mitocondrial y

estudiar su capacidad de inducir la aparición de marcadores y

características de un fenotipo pardo en adipocitos blancos en

cultivo.

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Hipótesis y Objetivos

61

REFERENCES/REFERENCIAS

Ahmadian, M., R. E. Duncan, et al. (2009). "The skinny on fat: lipolysis and

fatty acid utilization in adipocytes." Trends Endocrinol Metab 20(9): 424-8.

Bartelt, A. and J. Heeren (2013). "Adipose tissue browning and metabolic

health." Nat Rev Endocrinol.

Carbonelli, M. G., L. Di Renzo, et al. (2010). "Alpha-lipoic acid

supplementation: a tool for obesity therapy?" Curr Pharm Des 16(7): 840-6.

Fernandez-Galilea, M., P. Perez-Matute, et al. (2011). "Effects of lipoic acid on

apelin in 3T3-L1 adipocytes and in high-fat fed rats." J Physiol Biochem

67(3): 479-86.

Kim, M. S., J. Y. Park, et al. (2004a). "Anti-obesity effects of alpha-lipoic acid

mediated by suppression of hypothalamic AMP-activated protein kinase."

Nat Med 10(7): 727-33.

Liu, J., W. Shen, et al. (2009). "Targeting mitochondrial biogenesis for

preventing and treating insulin resistance in diabetes and obesity: Hope

from natural mitochondrial nutrients." Adv Drug Deliv Rev 61(14): 1343-52.

Lodhi, I. J., L. Yin, et al. (2012). "Inhibiting adipose tissue lipogenesis

reprograms thermogenesis and PPARgamma activation to decrease diet-

induced obesity." Cell Metab 16(2): 189-201.

Lottenberg, A. M., S. Afonso Mda, et al. (2012). "The role of dietary fatty acids

in the pathology of metabolic syndrome." J Nutr Biochem 23(9): 1027-40.

Prieto-Hontoria, P. L., P. Perez-Matute, et al. (2013). "Effects of lipoic acid on

AMPK and adiponectin in adipose tissue of low- and high-fat-fed rats." Eur

J Nutr 52(2): 779-87.

Prieto-Hontoria, P. L., P. Perez-Matute, et al. (2009). "Lipoic acid prevents

body weight gain induced by a high fat diet in rats: effects on intestinal

sugar transport." J Physiol Biochem 65(1): 43-50.

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Material y Métodos

65

3.1.-CULTIVO DE LA LÍNEA CELULAR 3T3-L1

3.1.1.- Fundamento teórico

Las fibroblastos de la línea celular murina 3T3-L1 son células capaces de

transformarse en adipocitos en presencia de un cóctel hormonal adecuado y

han sido ampliamente utilizados en los últimos años, tanto para el estudio de la

adipogénesis como de la fisiología del adipocito maduro, incluyendo

aproximaciones a su metabolismo, su función secretora y las vías de

señalización intracelular (Vigilanza et al., 2011; Suzuki et al., 2011; Moreno-

Aliaga et al., 2002).

3.1.2.- Material

Dulbecco´s Modified Eagle Medium (DMEM), High Glucose

(4,5 g/L) (Gibco-Invitrogen Corporation)

Suero Bovino de ternera (CSB) (Gibco-Invitrogen

Corporation)

Suero Fetal Bovino (FBS), Heat Inactivated (Gibco-

Invitrogen Corporation)

Penicilina/Estreptomicina (Gibco-Invitrogen Corporation)

Insulina (Sigma-Aldrich)

Isobutilmetilxantina (IBMX) (Sigma-Aldrich)

Dexametasona (Sigma-Aldrich)

3.1.3.- Procedimiento experimental

La línea celular 3T3-L1 se obtuvo de la American Type Culture Collection

(ATCC, Rockville). Las células se cultivaron en medio DMEM alto en glucosa

(4,5 g glucosa/L) y suplementado con L-glutamina y piruvato. Al medio se le

añadió suero bovino de ternero (CSB) al 10% y estreptomicina/penicilina al 1%.

Cuando las células alcanzaron el 100% de confluencia, se indujo la

diferenciación de los preadipocitos a adipocitos. Para ello, se cultivaron las

células en medio de diferenciación DMEM alto en glucosa suplementado con

10% de suero fetal bovino (FBS), estreptomicina/penicilina al 1%, insulina (10

μg/mL), IBMX (0,5 mM) y dexametasona (1 μM) durante 48 h. Pasado ese

tiempo, se retiró el medio de diferenciación y las células se cultivaron durante

otras 48 h en medio post-diferenciación (DMEM con FBS al 10%,

estreptomicina/penicilina al 1% y 10 μg/mL de insulina).

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A partir del día 4 de postdiferenciación las células se cultivaron con DMEM

suplementado con 10% FBS y estreptomicina/penicilina al 1% hasta el día 7-8

de postdiferenciación en el cual aproximadamente el 100% de las células ya

habían alcanzado tanto la morfología como la funcionalidad típica de adipocitos

maduros, con lo cual pudieron ser utilizadas como modelo in vitro de adipocitos.

Todo el proceso de crecimiento y diferenciación de la línea celular 3T3-L1 se

llevó a cabo en un incubador a 37 ºC y 5% de CO2.

Figura 7: Cultivo y proceso de diferenciación de adipocitos 3T3-L1

3.1.4.- Tratamientos

La tarde previa al tratamiento de las células, el medio de cultivo de post-

diferenciación se cambió a medio DMEM (4,5 g/L) suplementado al 1% con FBS

y estreptomicina/penicilina al 1%. Así, las células se mantuvieron en estas

condiciones durante unas 14-15 h previas a la realización del experimento. A

continuación, se añadieron los medios con los tratamientos correspondientes:

Control (Etanol, vehículo en el que el α-LA fue disuelto) y α-LA (1-500 µM) en

presencia o ausencia de distintos inhibidores o activadores específicos de las

vías de señalización celular a estudiar y a su concentración adecuada. Se

incubaron los adipocitos durante 24-48 h a 37 ºC y 5% CO2 en condiciones de

esterilidad. En el caso de utilizar inhibidores o activadores de diferentes vías de

ADIPOCITOS DIFERENCIADOS

TRATAMIENTOS

DIA 7

PREADIPOCITOS 3T3-L1

DETERMINACIONES

Confluencia Post-confluencia

DMEM + CBS

+ ANTIBIOTICOS

DMEM + FBS

+ ANTIBIOTICOS + IBMX

+ DEXAMETASONA + INSULINA

DMEM + FBS

+ ANTIBIOTICOS + INSULINA

48 h

48 h

DMEM + FBS

+ ANTIBIOTICOS

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señalización, los adipocitos se preincubaron en presencia del compuesto

durante 1 h y a continuación, se añadió el tratamiento correspondiente. Los

adipocitos del grupo control fueron tratados con la misma cantidad del vehículo

utilizado para disolver cada uno de los agentes en estudio (etanol y/o DMSO,

según casos).

Tabla 3: Listado de inhibidores/activadores y sus correspondientes vías de señalización.

Tratamiento Molecula diana Concentración

SP600125

(-) JNK

20 µM

PD98059 (-) MAPK (ERK1/2) 50 µM

LY294002 (-) AKT 50 µM

AICAR (+) AMPK 2 mM

Compound C (-) AMPK 20 µM

H89 (-) PKA 1 µM

Cilostamide (-) PDE3B 2 µM

L798106 (-) EP3-receptor 10 µM

Posteriormente, se tomó muestra del medio de cultivo para las

determinaciones bioquímicas (glicerol, acidos grasos libres y PGE2).

Seguidamente, y tras recoger todo el medio de cultivo, las placas se

congelaron a –80 ºC para su posterior extracción de RNA y análisis de la

expresión génica, o bien se añadió un buffer de lisis para la extracción de

proteínas.

3.2.- CULTIVO DE ADIPOCITOS HUMANOS

3.2.1.- Fundamento teórico

Los preadipocitos subcutáneos humanos se obtuvieron de la compañía Zen-

Bio Inc (NC, USA). Los preadipocitos fueron obtenidos de tejido subcutáneo

adiposo humano, de la región abdominal de mujeres caucásicas con

sobrepeso/obesidad (IMC 25-35 Kg/m2), no diabéticas y con edades

comprendidas entre 25-53 años. Estos preadipocitos son capaces de

transformarse en adipocitos en presencia de un medio de diferenciación

comercial y han sido ampliamente utilizados tanto para el estudio de la

adipogénesis como de la fisiología y metabolismo del adipocito maduro

(Tomlinson et al., 2010; Ortega et al., 2010).

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3.2.2.- Material

Medio de preadipocitos PM-1 (Zen-Bio Inc., NC, USA).

Medio de diferenciación de adipocitos DM-2 (Zen-Bio Inc.,

NC, USA).

Medio de mantenimiento de adipocitos AM-1 (Zen-Bio Inc.,

NC, USA).

3.2.3.- Procedimiento experimental

Los preadipocitos se cultivaron en 1 ml de medio específico, PM-1 (Zen-Bio

Inc., NC, USA). Cuando estos preadipocitos alcanzaron el 100 % de

confluencia, se indujo su diferenciación hasta adipocitos maduros. Para ello, se

cultivaron en 1 ml de medio de diferenciación comercial DM-2, (Zen-Bio Inc.,

NC, USA). Después de 7 días se retiró parte de este medio de diferenciación

(600 µl) y se añadieron 800 µl del medio de mantenimiento de adipocitos AM-1

(Zen-Bio Inc., NC, USA). Tras 2 días de incubación se retiraron 600 µl del medio

y se sustituyeron por 600 µl de AM-1 y se mantuvieron hasta el día 14 de

postdiferenciación en el cual aproximadamente el 100% de las células ya

habían alcanzado la morfología y la funcionalidad típica de adipocitos maduros,

con lo cual pudieron ser utilizadas como modelo in vitro de adipocitos humanos.

Durante todo el proceso de crecimiento y diferenciación, las células fueron

guardadas en un incubador a 37 ºC y 5 % de CO2. Todos los medios contienen

glucosa (3,15 g/L).

3.2.4.- Tratamientos

De forma previa al tratamiento de las células, se cambió el medio de cultivo

de mantenimiento de los adipocitos AM-1. A continuación, se añadieron los

medios con los tratamientos correspondientes: Control (vehículo) y α-LA (1-250

µM) y se incubaron los adipocitos durante 24 h a 37 ºC y 5 % CO2 en

condiciones de esterilidad. Los adipocitos del grupo control fueron tratados con

la misma cantidad del vehículo utilizado para disolver el α-LA (etanol). Así

mismo, en el caso de utilizar inhibidores o activadores de diferentes vías de

señalización, los adipocitos se preincubaron en presencia del compuesto

durante 1 h y a continuación, se añadió el tratamiento correspondiente.

Posteriormente, se tomaron muestras del medio de cultivo para las

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determinaciones bioquímicas. Seguidamente, y tras recoger todo el medio de

cultivo, las placas se congelaron a – 80 ºC para posterior extracción de RNA de

los adipocitos y analizar los niveles de expresión génica, o bien se añadió un

buffer de lisis para la extracción de proteínas.

Figura 8: Cultivo y proceso de diferenciación de los preadipocitos humanos.

3.3.- MEDIDA DE LA LIPÓLISIS EN ADIPOCITOS 3T3-L1

Los niveles de lipólisis fueron evaluados mediante la medida de la cantidad

de glicerol y ácidos grasos libres liberados al medio. La determinación del

glicerol liberado se realizó tras el tratamiento con α-LA (1-500 µM) y tras

diferentes periodos de incubación (1-24 h) mediante el autoanalizador Cobas-

Mira (Roche Diagnostics, Basel, Suiza). Los ácidos grasos libres fueron

cuantificados tras 3 h de tratamiento con LA (1-500 µM) mediante el kit

comercial Lipolysis Assay KIT for Free Fatty Acids Detection (Zen-Bio Inc,

Research Triangle Park, NC). Ambos procedimientos se realizaron siguiendo las

instrucciones del fabricante.

Confluencia Post-confluencia

Medio de diferenciación

DM-2 Medio AM-1 Añadir los dias 7 y 9

ADIPOCITOS DIFERENCIADOS

TRATAMIENTOS

DIA 14

PREADIPOCITOS SUBCUTÁNEOS

HUMANOS

Medio PM-1

24 h h

DETERMINACIONES

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3.3.1.-Glicerol

La determinación del glicerol se realizó mediante un procedimiento

colorimétrico directo que utiliza un sistema cromógeno de quinoneimina en

presencia de las enzimas glicerol quinasa, peroxidasa y glicerol fosfato oxidasa

(Randox, Antrim, United Kingdom).

Los niveles de glicerol liberados al medio se calcularon como el incremento

de glicerol en el medio de cultivo tras la incubación con los diversos agentes en

estudio y se expresó como μmoles/l o como porcentaje sobre el control.

3.3.2.-Acidos grasos libres

La evaluación de los niveles de acidos grasos libres liberados al medio se

realizó mediante una reacción acoplada en la cual el paso inicial es llevado a

cabo por la acil-CoA sintasa (ACS), que produce acil-CoA tiol-esteres a partir de

los acidos grasos libres, ATP, Mg++

y CoA presentes en la reacción. Los acil-

CoA derivados reaccionan con oxígeno en presencia de la acil-CoA oxidasa

(ACOD) para producir peróxido de hidrógeno, el cual en presencia de la enzima

peroxidasa permite la condensación oxidativa del compuesto of 3-metil-N-etil-N-

(β-hidroxietil)-anilina con 4-aminoantipirina, formando así un compuesto de color

morado que puede ser detectado mediante la absorbancia a 550 nm.

3.4.- DETERMINACIÓN DE LA EXPRESIÓN GÉNICA

El análisis de la expresión génica se realizó mediante PCR a tiempo real

(qRT-PCR), tras la extracción del RNA total y su posterior retrotranscripción.

n-(4-antipiril)-3-cloro-5-sulfonato-p-benzoquinoneimina

Glicerol + ATP Glicerol quinasa

Glicerol-3-fosfato + ADP

Glicerol-3-fosfato + O2 Glicerol fosfato oxidasa

H2O2 + Dihidroxiacetona fosfato

H2O2 + ácido 3,5-dicloro-2-hidroxi-benzenosulfónico + 4-aminofenazona Peroxidasa

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Figura 9: Esquema del análisis de la expresión génica.

3.4.1.-Extracción y cuantificación de RNA

La extracción del RNA total se llevó a cabo siguiendo el método del

TRIZOL® (Invitrogen), que consiste en una solución monofásica de fenol e

isotiocianato de guanidina. Durante esta homogeneización, el Trizol mantiene la

integridad del RNA, mientras que rompe las células y disuelve los componentes

celulares. Con este método, se puede partir de pequeñas cantidades de tejidos,

obteniéndose un RNA total libre de contaminación por proteínas y DNA

3.4.1.1.- Procedimiento

Para la realización de la extracción del RNA total se adicionó a las placas de

cultivo 1 ml de Trizol por pocillo, y con ayuda de un rascador y se homogeneizó

el tejido. Posteriormente los homogeneizados se trasvasaron a un tubo

eppendorf de 2 ml y se agitaron vigorosamente durante 1 min, y tras un periodo

de incubación de 5 min a temperatura ambiente se centrifugaron a 12.000 x g

durante 10 min a 4 ºC, para eliminar los principales restos celulares.

A continuación se añadieron 200 µl de cloroformo y se agitaron las

muestras hasta conseguir la completa distribución del cloroformo (Sigma-

Aldrich) sobre la fase del Trizol. Tras esperar 2-3 min, se volvió a centrifugar las

muestras esta vez durante 20 min, para así separar una fase orgánica (fase

inferior donde se encuentra el DNA), una interfase (proteínas) y una fase

acuosa (que supone un 60% y donde se encuentra el RNA).

Tras recoger la fase superior acuosa se le añadió 500 µl de alcohol

isopropílico y 5 µl de Glycoblue (Ambion, Austin, USA) con el fin de identificar

mejor el pellet de RNA que se obtuvo tras una centrifugación a 12.000 x g

durante 20 min a 4 ºC.

Con objeto de lavar el RNA precipitado, se decantó el sobrenadante dejando

intacto el pellet y se añadieron 1,5 ml de 75% etanol-H2O-DEPC centrifugándolo

Homogenización en trizol

Extracción RNA Retrotranscripción

Real time PCR

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posteriormente a 12.000 x g durante 5 min. Tras un nuevo lavado, se retiró todo

el sobrenadante y se dejó secar el pellet a temperatura ambiente durante 2 min.

El RNA fue resuspendido en la solución RNA secure (Ambión, Austin, USA)

utilizando un volumen de 20 µl por muestra, y se incubó durante 10 min en baño

seco a 55-65 ºC para disolver el pellet e inactivar las RNAsas. Se enfrió en hielo

y se procedió a la cuantificación del RNA total presente en cada muestra así

como su grado de pureza respecto a proteínas y sales minerales utilizando el

espectrofotómetro Nanodrop ND1000 (Thermo Scientific, Wilminton, DE, USA) .

3.4.2.- Tratamiento con DNAsa y retrotranscripción

De forma previa a la retrotranscripción se realizó un tratamiento con el

DNAfree kit (Ambion, Austin, USA) con la finalidad de obtener un RNA mas puro

y libre de contaminaciones con DNA genómico.

Para ello, se tomaron 2 µg de RNA, que se incubaron durante 30 min a 37

ºC en presencia de la enzima DNAsa, la cual degrada los restos de DNA

genómico presentes en la muestra.

Seguidamente y tras la inactivación de la enzima con el DNAsa Inactivation

reagent, se procedió a realizar la retrotranscripción de cada una de las

muestras, que consiste en la obtención de DNA complementario (cDNA) a partir

del RNA total obtenido para el posterior análisis de la expresión génica

mediante PCR cuantitativa a tiempo real (qRT-PCR).

Para cada muestra se realizó la retrotranscripción de 2 µg de RNA utilizando

la enzima retrotranscriptasa inversa (M-MLV, Invitrogen) en presencia de la

enzima inhibidora de ribonucleasas (RNasinTM, Promega) en la siguiente

proporción: 4 µl de buffer 5x, 2 µl DTT, 1 µl RNasin y 1 µl M-MLV.

Las muestras se incubaron 10 min a 25 ºC, 60 min a 37 ºC y finalmente 15

min a 70 ºC. Las muestras de cDNA obtenido se alicuotaron y se guardaron a -

80 ºC hasta su posterior utilización.

3.4.3.- Determinación de los niveles de expresión génica mediante la

técnica q RT-PCR

La determinación de los niveles de expresión génica se realizó mediante

PCR a tiempo real, que es un método semi-cuantitativo basado en la actividad

5´exonucleasa de la Taq polimerasa.

Mediante esta técnica, se puede determinar en tiempo real la amplificación

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del gen de estudio utilizándose otro fragmento de DNA (sonda) complementario

a una parte intermedia del DNA que se quiere amplificar. Dicha sonda lleva

acoplada una molécula fluorescente (reporter) y otra molécula que inhibe la

fluorescencia (quencher). De esta forma, cuando la molécula fluorescente es

desplazada por la enzima Taq polimerasa, dicha molécula se libera y emite

fluorescencia al ser iluminada con un láser. La cuantificación de la fluorescencia

emitida durante cada ciclo de PCR será proporcional a la cantidad de DNA que

se está amplificando. El detector fotométrico junto con un programa especial,

monitoriza el incremento en la emisión del fluorocromo. El algoritmo normaliza la

señal a un patrón interno (ΔRn) y automáticamente calcula la línea de corte del

ciclo (Threshold-CT) cuando el ΔRn alcanza diez veces la desviación estándar

de la línea base.

Los datos se obtuvieron como valores CT (ciclo en el cual la señal de

fluorescencia emitida se encuentra considerablemente por encima de los

niveles de amplificación inespecífica y es inversamente proporcional al número

de copias iniciales de la muestra). Después se determinaron los valores de ΔCT

(ΔCT= CT del gen en estudio - CT del gen de referencia) para cada muestra. Los

cambios en la expresión del gen se calcularon por el método de 2-ΔΔ CT

(Perez-

Matute et al., 2009).

Figura 10: Esquema de la determinación de la expresión génica mediante PCR a

tiempo real.

3.4.3.1.- Procedimiento

Inicialmente, se realizó una curva estándar de validación para cada cebador-

sonda con diluciones seriadas de varias muestras para asegurar que el final de

la reacción (tanto para muestras control, como para los distintos tratamientos)

se encontraba en la parte media de la curva exponencial de amplificación. Se

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utilizaron 5,5 µl de cDNA, 4 µl de Taqman Universal PCR Master Mix (Applied

biosystems) y 0,5 µl de cada cebador-sonda por cada muestra. También se

ensayaron diversos genes de referencia (house keeping genes) para normalizar

los datos (18s, ciclofilina, β-actina, Ubiquitina C, y Gliceraldehido 3-fosfato

deshidrogenasa), siendo elegidos aquellos que presentaron una menor

variación entre las distintas muestras y tratamientos. En este trabajo, los genes

en estudio se refirieron a los genes β-actina y/o 18s.

Los reactivos para el análisis de expresión génica de los distintos genes en

estudio, así como los genes de referencia son prediseñados y obtenidos de

Applied Biosystems (Foster City, EEUU), y las condiciones experimentales se

ajustaron a las indicaciones del fabricante. La detección y amplificación de los

genes específicos se llevó a cabo con el sistema de detección de secuencias

ABI PRISM 7900HT (Secuence Detection System, Applied Biosystems).

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Tabla 4: Listado de genes y las correspondientes sondas utilizadas para el análisis de la

expresión génica.

Nombre Sonda Taqman

Ref#

Gen Especie

Lipasa sensible a hormonas (HSL)

Mm00495359_m1

Hsl

Mus musculus

Lipasa de triglicéridos del tejido adiposo

(ATGL)

Mm00503040_m1 ATGL/Pnpla2 Mus musculus

Perilipina Mm00558672_m1 Plin1 Mus musculus

β-actina Mm02619580_g1 Actb Mus musculus

Factor de Transcripción Mitocondrial A Hs01082775_m1 Tfam Homo sapiens

Factor Respiratorio Nuclear 1 Hs00192316_m1 Nrf1 Homo sapiens

Sirtuina 1 Hs01009005_m1 Sirt1 Homo sapiens

PR domain containing 16 Hs00991677_m1 Prdm16 Homo sapiens

Receptor activado por el proliferador de

peroxisomas γ, coactivador 1 alpha (PGC-

1α)

Hs01016719_m1 Ppargc1A

Homo sapiens

Stearoil CoA Desaturasa Hs01682761_m1 Scd1 Homo sapiens

Sintasa de ácidos grasos Hs01005622_m1 Fasn Homo sapiens

Diacil glicerol transferasa 1 Hs00201385_m1 Dgat1 Homo sapiens

Efector DFFA-like inductor de muerte

celular a (CideA)

HS00154455_m1 CideA Homo sapiens

Carnitina palmitoil-transferasa 1a (CPT1α) Hs00912671_m1 Cpt1 Homo sapiens

Acil CoA oxidasa Hs00971639_m1 Acox1 Homo sapiens

T-BOX 1 Hs00962556_m1 Tbx1 Homo sapiens

Fibronectin type III domain-containing

protein 5

Hs00401006_m1 Fndc5 Homo sapiens

18s Hs99999901_s1 18s Homo sapiens

3.5.- DETERMINACIÓN DE LOS NIVELES DE PROTEÍNA Y DE

MODIFICACIONES POST-TRANSCRIPCIONALES MEDIANTE

WESTERN BLOT

El análisis de los niveles de distintas proteínas así como de los niveles de

fosforilación y acetilación se llevó a cabo mediante la técnica Western Blot. La

técnica utiliza un gel de electroforesis (SDS-PAGE) para separar proteínas

desnaturalizadas en función de la longitud del polipéptido. Las proteínas son

transferidas a una membrana, habitualmente de nitrocelulosa o

polivinilidenofluoruro (PVDF), donde son detectadas mediante anticuerpos

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específicos contra la proteína diana. Para el análisis de la fosforilación de

proteínas se utilizaron anticuerpos específicos.

Figura 11: Esquema general de la técnica de Western-Blot

3.5.1.- Preparación de las muestras

Para la obtención de los extractos proteicos de los adipocitos en cultivo

(3T3-L1 y adipocitos procedentes de tejido subcutáneo humano) se adicionaron

250 µL del buffer de lisis que contenía: Tris HCl (pH 7,5) 50 mM; NaCl 150 mM,

EDTA 5 mM; NaF 2 mM; Octilglicina 6 mM; Desoxicolato de Sodio 0,25%;

Ortovanadato de sodio 2 mM, Cóctel inhibidor de proteasas 1 (Sigma-Aldrich)

1%; Tritón X100 y agua de biología molecular o autoclavada. Posteriormente se

procedió a la homogeneización del extracto mediante el uso de un rascador

hasta lograr liberar las células adheridas a la placa obteniéndose un

homogeneizado completo y se incubaron las placas a 4ºC durante 30 min. Todo

el proceso se realizó con la muestra en frío para evitar que las altas

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temperaturas puedan dañar las proteínas. Una vez transcurrido este tiempo, se

procedió a centrifugar 10 min a 13.400 x g a 4 ºC. De los sobrenadantes

obtenidos (extractos proteicos) una alícuota se utilizó para la cuantificación de la

proteína total del extracto mediante la técnica de BCA (Pierce-Thermo Scientific,

Rockford, IL, USA) según las instrucciones del fabricante, y el resto de los

extractos fueron congelados a -80 ºC para su posterior uso.

3.5.2.- Electroforesis y electrotransferencia

Las proteínas se separaron mediante SDS-PAGE, en un gel

desnaturalizante de acrilamida al 12%, a una intensidad de 120 voltios durante

una h (Laemmli, 1970). Se utilizó como marcador de peso molecular el

Prestained SDS-PAGE Standards, Low Range o bien el Precision Plus Protein

Standard, Dual Color (Bio Rad Laboratories).

Las proteínas fueron transferidas desde el gel a una membrana de PVDF

en un tampón que contenía: 25 mM de Tris base, 192 mM de glicina y 20% de

metanol y aplicando una corriente de 400 mA durante un mínimo de 45 min y un

máximo de 2 h. El sistema fue montado tal y como se detalla en la Figura 12.

Figura 12: Sistema de electrotransferencia de proteínas

3.5.3.- Inmunoblot

Una vez transferidas las proteínas a las membranas de PVDF, ésta se

bloqueó durante 2 h a temperatura ambiente en buffer TBS-T (200 mM Tris

base; 1,5 M NaCl; 0,15% Tween 20) con 1% de BSA, con el fin de evitar

uniones inespecíficas.

Esponja Papel de filtro

Membrana de PVDF

Gel de acrilamida

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Tras el bloqueo, las membranas se lavaron con TBS-T tres veces y se

incubaron con los anticuerpos primarios específicos diluidos en buffer TBS-T

con 1% de BSA durante toda la noche a 4 ºC y en agitación.

Para determinar los niveles de acetilación de Pgc-1α se tomaron 200 μg de

los extractos de proteína obtenidos a partir de cultivos de adipocitos humanos,

y fueron diluidos en solución de lisis hasta una concentración final de 1 µg/µl de

proteína. Se añadieron 2 µg de anticuerpo anti-PGC-1α, y se dejó incubando

dos h a 4 ºC, en agitación. Posteriormente se añadieron a las muestras 20 μl de

suspensión de proteína A/G PLUS‐Agarose (Santa Cruz Biotechnology). Tras

incubar la mezcla durante toda la noche a 4 ºC y en agitación moderada, se

centrifugaron los tubos 1 min a 1000 g y a 4 ºC. Se retiró el sobrenadante y se

hicieron 4 lavados de la agarosa y los inmunocomplejos, mediante

centrifugación con 500 μl de solución de PBS, retirando el sobrenadante cada

vez. A continuación, se añadieron sobre la agarosa 40 μl de buffer de carga 2X

(estándar) y se hirvieron las muestras durante 3 min, separando así los

inmunocomplejos de la agarosa. Se centrifugó una vez más, durante 1 min a

1000 g a 4ºC y se recogió el sobrenadante. Finalmente, se cargaron volúmenes

iguales de todas las muestras y se hibridaron con anticuerpos monoclonal

anti‐acetylated‐lysine (Cell Signaling) y anti-PGC-1α (Santa Cruz

Biotechnology).

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Tabla 5: Listado de anticuerpos primarios utilizados.

Anticuerpo Fabricante Referencia Procedencia

HSL Cell signaling technology 4107 Conejo

HSL fosfo ser565

Cell signaling technology 4137 Conejo

HSL fosfo ser563

Cell signaling technology 4139 Conejo

HSL fosfo ser660

Cell signaling technology 4126 Conejo

ATGL (ATGL) Cell signaling technology 2138 Conejo

G0S2 Santa Cruz Biotechnologies Sc-133423 Conejo

CGI58 Santa Cruz Biotechnologies Sc-130934 Conejo

Perilipina Cell signaling technology 9349 Conejo

Fosfo PKA sustrato Cell signaling technology 9624 Conejo

AMPK Cell signaling technology 2532 Conejo

AMPK fosfo Thr172 Cell signaling technology 2535 Conejo

AKT Cell signaling technology 9272 Conejo

AKT fosfo Ser473 Cell signaling technology 4058 Conejo

Anti-MAPK (ERK1/2) Cell signaling technology 9102 Conejo

ERK1/2 fosfo Thr202/Tyr204 Cell signaling technology 4370 Conejo

JNK Cell signaling technology 4668 Conejo

JNK fosfo Thr183/

Tyr185

Cell signaling technology 9252 Conejo

AdPLA Cayman Chemical 10337 Conejo

PGC-1α Santa Cruz Biotechnology SC67286 Conejo

ACC Cell signaling technology 3676 Conejo

ACC fosfo Ser79

Cell signaling technology 3661 Conejo

FAS Cell signaling technology 3180 Conejo

DGAT1 Abcam Ab54037 Conejo

SCD1 Abcam Ab23331 Conejo

SIRT1 Cell signaling technology 2310 Conejo

CPT1α Cell signaling technology 12252 Conejo

ACOX1 Abcam Ab59964 Conejo

Acetylated-Lysine Cell signaling technology 9814 Conejo

CD36 Santa Cruz Biotechnology SC9154 Conejo

UCP1 Abcam Ab10983 Conejo

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3.6.- ESTUDIO DE LOS NIVELES DE PGE2 MEDIANTE ELISA

Los niveles de PGE2 liberados al medio de cultivo por los adipocitos 3T3-L1

fueron cuantificados mediante ELISA empleando el kit “PGE2 Enzyme

Immunoassay kit” (Arbor assays, Ann Arbor, MI, USA).

3.6.1.- Fundamento básico

El kit de ELISA “PGE2 Enzyme Immunoassay kit” está diseñado para el

análisis cuantitativo de la PGE2 presente en suero, plasma, orina, saliva y

medio de cultivo. Para ello las muestras son pipeteadas en una microplaca

cubierta con un anticuerpo para la captura de IgG de ratón. Un anticuerpo

conjugado con peroxidasa es añadido a los estándares y muestras en los

pocillos. La reacción de unión se inicia al añadir un anticuerpo monoclonal anti-

PGE2 a los pocillos. Tras 2 h de incubación la placa es lavada y se añade el

substrato, el cual reacciona con el conjugado unido a la PGE2. Tras una breve

incubación la reacción es parada y la intensidad de color generada se detecta

mediante un lector de placas capaz de medir a 450 nm de longitud de onda.

La concentración de PGE2 en la muestra es calculada tras la interpolación

en una curva estándar de la densidad óptica (DO) obtenida de cada muestra y

la multiplicación por el factor de dilución utilizado.

3.7.- ESTUDIO DE LOS NIVELES DE cAMP MEDIANTE ELISA

Los niveles citoplasmáticos de cAMP por los adipocitos 3T3-L1 fueron

cuantificados mediante ELISA empleando el kit “cAMP Direct EIA kit (Arbor

Assays)”.

7.1.- Fundamento básico

El kit “cAMP Direct EIA kit” esta diseñado para el analisis cuantitativo del

cAMP presente en lisados celulares así como otras muestras tales como

plasma, orina, saliva y medios de cultivo. Para el análisis de las muestras, éstas

son diluidas en un diluyente ácido que contiene aditivos y estabilizantes

especiales para la medida de cAMP. Así, las muestras en presencia de este

buffer, que inactiva las fosfodiesterasas endógenas, son pipeteadas en una

placa recubierta con un anticuerpo que captura IgG de oveja tras añadir una

solución neutralizante. Posteriormente se incorpora a cada pocillo un anticuerpo

conjugado con peroxidasa anti-cAMP. La reacción de unión se inicia al añadir

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un anticuerpo de oveja anti c-AMP en cada pocillo. Tras 2 h de incubación, la

placa es lavada y se añade la solución substrato. El substrato reacciona con el

complejo cAMP-anticuerpo conjugado y tras una breve incubación se para la

reacción y la intensidad de color generado se detecta en un lector de placas a

450 nm. La concentración del cAMP en la muestra es calculada mediante la

interpolación de la DO en una recta patrón y haciendo la corrección por el factor

de dilución adecuado.

3.8.- EVALUACIÓN DE LA OXIDACIÓN DE LOS ÁCIDOS GRASOS

EN ADIPOCITOS 3T3-L1

La oxidación de ácidos grasos en adipocitos murinos 3T3-L1 se determinó

mediante la cuantificación de los productos solubles en medio ácido

previamente descrito por Mercader et al. (2011). Para ello se sembró

preadipocitos 3T3-L1 en placas de 12 pocillos los cuales fueron diferenciados

completamente como se detalla en el apartado 1 (Cultivo de la linea celular 3T3-

L1) hasta la obtención de adipocitos maduros (día 7-8 de diferenciación).

Entonces los adipocitos fueron tratados con α-LA (250 µM) o el vehículo

correspondiente (etanol 0,1%) durante 5 h en presencia de 14

C-ácido palmítico

(Perkin Elmer, Boston, MA). Tras este periodo de incubación los productos

solubles en medio ácido derivados de la oxidación de ácidos grasos fueron

extraídos en 1 ml de ácido perclórico frío (1M) (Panreac, Barcelona, España).

Posteriormente la muestra fue centrifugada durante 10 min a 1800 g y la

radiactividad del sobrenadante fue medida mediante un contador de centelleo

Wallac 1409 (EG&G Company, Turku, Finlandia). Los resultados obtenidos

fueron normalizados por la cantidad total de proteína del extracto medida

mediante el método de BCA (Pierce-Thermo Scientific).

3.9.- ANÁLISIS DE LA OXIDACIÓN DE LOS ÁCIDOS GRASOS EN

ADIPOCITOS SUBCUTÁNEOS HUMANOS

El análisis de la oxidación de ácidos grasos en adipocitos subcutáneos

humanos se realizó mediante la técnica previamente descrita por Bourlier et al.

(2013). La oxidación total del 14

C-ácido palmítico se determinó mediante la

cuantificación tanto del CO2 liberado al medio como de los productos solubles

en medio ácidos. Así, los preadipocitos humanos fueron sembrados en placas

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de 12 pocillos y diferenciados hasta que estos adquirieron las características

típicas de los adipocitos maduros como se describe en el apartado 2 (Cultivo de

adipocitos humanos). En el día 14 de diferenciación, se cambió el medio de

cultivo añadiendo 1 ml de medio AM1 y los adipocitos fueron tratados con α-LA

(100 ó 250 µM) o bien el vehículo. Tras 24 h de incubación se cambió el medio

de cultivo por un buffer que contenía 125 mM NaCl, 5 mM KCl, 2 mM CaCl2,

1.25 mM KH2PO4, 1.25 mM MgSO4•7H2O, 25 mM NaHCO3, 1 mM carnitina

(Sigma), 80 µM final de ácido palmítico no radiactivo (Sigma) y 20 µM de 14

C-

ácido palmítico (Perkin Elmer) y se mantuvo 4 h más en el incubador en

presencia de los tratamientos adecuados. Transcurrido este tiempo, para la

determinación del CO2 producido por las células, se recuperó el medio y se

introdujo en viales de vidrio que contenían un tubo eppendorf con 300 µl de

hidróxido de benzetonio (Sigma) en su interior. Se acidificó el medio con H2SO4

1M (Panreac), se cerraron los viales y se incubó a temperatura ambiente

durante 2 h. Pasado el tiempo de incubación el eppendorf fue recuperado y se

introdujo en un vial de centelleo que contenía 10 ml de líquido de centelleo

EcoScint Ultra (National diagnostics, Charlotte, NC). Tras agitar uniformemente

todos los viales se procedió a medir la radiactividad mediante un contador de

centelleo Wallac 1409 (EG&G Company, Turku, Finlandia). En paralelo, las

células fueron lavadas 2 veces con buffer PBS (Gibco), cosechadas en 500 µl

de buffer STED (0,25 M sacarosa, 10 mM Tris HCl pH 7,4, 1 mM EDTA, 1 mM

DTT) con la ayuda de un rascador y lisadas tras ser sometidas a 2 ciclos de

congelación descongelación en nitrógeno líquido. Para la medida de los

productos solubles en medio ácido, en tubos de vidrio, se mezcló 400 µl de los

extractos STED y 2,5 ml de metanol/cloroformo (1:2) mediante la agitación en

vortex durante 10 min. Posteriormente se añadió 2 ml de KCl/HCl 1M y se agitó

durante 10 min más. Tras este paso se procedió a la centrifugación de las

muestras (1100 g, 10 min, temperatura ambiente) para obtener 2 fases. De la

fase superior acuosa se obtuvieron 2 alícuotas de 500 µl cuya radiactividad fue

medida en viales de centelleo que contenían 4 ml de líquido de centelleo en el

contador Wallac 1409 (EG&G Company). Los resultados obtenidos fueron

normalizados por la cantidad total de proteína del extracto medida mediante el

método de BCA (Pierce-Thermo Scientific). La fase inferior orgánica fue

recogida para la determinación de la incorporación de acidos grasos en

triglicéridos.

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3.10.- MEDIDA DE LA INCORPORACIÓN DE ÁCIDOS GRASOS A

TRIGLICÉRIDOS EN ADIPOCITOS SUBCUTÁNEOS HUMANOS

Partiendo de la fase inferior orgánica que se obtuvo tras la medida de los

productos solubles de la oxidación de ácidos grasos, la cual contiene los lípidos

neutros de los adipocitos subcutáneos humanos, se realizó la medida de la

incorporación del 14

C-ácido palmítico en los triglicéridos presentes en la muestra

mediante cromatografía en capa fina (Thin layer chromatography, TLC).

Para ello la muestra fue evaporada bajo vacío, se resuspendió en 40 µl de

una mezcla metanol/cloroformo (1:2) y se aplicó en una placa de TLC. La placa

(Merck, Darmstadt, Germany) fue situada en una cubeta de desarrollo en

presencia de la fase móvil (heptano:isopropil eter:ácido acético; en una

proporción 60:40:4) y se incubó a temperatura ambiente hasta que la migración

fue completada por la misma. Posteriormente se dejó secar la placa y se

introdujo en otra cubeta que contenía sales de yodo (Panreac, Barcelona,

España) hasta que los lípidos fueron visibles. Las áreas que contenían los

triglicéridos fueron marcadas y tras humedecerlas con H2O destilada se

recuperó la sílice que las contenía, se introdujo en viales de centelleo con 4 ml

de líquido de centelleo EcoScint Ultra (National diagnostics) y se cuantificó la

radiactividad de las mismas mediante el contador Wallac 1409 (EG&G

Company). Al igual que en otras técnicas, los resultados obtenidos fueron

normalizados por la cantidad total de proteína del extracto medida mediante el

método de BCA (Pierce-Thermo Scientific).

Figura 13: Esquema básico de la técnica cromatografía en capa fina.

Frente

Trigliceridos

Acidos grasos libres

1,3 DAG 1,2 DAG

Monoacyl glicerol

Colesterol

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3.11.- MEDIDA DE LA CAPTACIÓN DE ACIDOS GRASOS EN

ADIPOCITOS SUBCUTÁNEOS HUMANOS

El análisis del transporte de ácidos grasos en adipocitos subcutáneos

humanos se llevó a cabo mediante el uso de [1-14

C] ácido 2 bromopalmítico

(Moraveck Biochemicals Inc. Brea, CA, USA), un análogo del ácido palmítico no

metabolizable. Así, el día anterior al experimento los adipocitos fueron tratados

con LA 100 µM, LA 250 µM o bien el vehículo (etanol 0,1%) para las células

control y tras 24 h de incubación en un incubador a 37ºC y 5% de CO2 se

sustituyó el medio de cultivo por un buffer que contenía 125mM NaCl, 5mM KCl,

2mM CaCl2, 1.25mM KH2PO4, 1.25mM MgSO4•7H2Oy 25mM NaHCO3

previamente saturado en O2 y se mantuvo 50 min más en el incubador en

presencia de los tratamientos adecuados. Posteriormente se añadió al medio 1

mM de carnitina (Sigma), 80 µM final de ácido palmítico no radiactivo (Sigma) y

20 µM de [1-14

C] ácido 2 bromopalmítico (Moraveck Biochemicals Inc.) durante

10 min. Transcurrido este tiempo, las células fueron lavadas 2 veces con buffer

PBS (Gibco) frio, cosechadas en 500 µl de buffer STED (0,25 M sacarosa; 10

mM Tris HCl pH 7,4; 1 mM EDTA; 1 mM DTT) con la ayuda de un rascador y

lisadas tras ser sometidas a 2 ciclos de congelación descongelación en

nitrógeno líquido. Con objeto de medir la radiactividad incorporada por los

adipocitos, 400 µl de los lisados fueron añadidos a viales que contenían 4 ml de

líquido de centelleo EcoScint Ultra (National diagnostics) para su análisis en un

contador de centelleo Wallac 1409 (EG&G Company). La radiactividad

observada fue normalizada por la cantidad total de proteína del extracto medida

mediante la técnica de BCA (Pierce-Thermo Scientific).

3.12.- ANÁLISIS DE LA LIPOGÉNESIS DE NOVO EN

ADIPOCITOS SUBCUTÁNEOS HUMANOS

El análisis de la incorporación de glucosa a los triglicéridos presentes en la

muestra se determinó mediante la cuantificación de la incorporación de 14

C-

glucosa en los triglicéridos tras la extracción y separación de los lípidos neutros

almacenados en las células mediante cromatografía en capa fina (Thin layer

chromatography, TLC). Para ello, en el día 14 de diferenciación, los adipocitos

fueron tratados con α-LA (100 ó 250 µM) o bien el vehículo (etanol 0,1%) y tras

24 h de incubación se sustituyó el medio de cultivo por un buffer que contenía

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125 mM NaCl, 5 mM KCl, 2 mM CaCl2, 1.25 mM KH2PO4, 1.25 mM

MgSO4.7H2O, 25 mM NaHCO3, 2 mM de glucosa no radiactiva (Sigma) y 157

µM de 14

C-glucosa (American Radiolabeled Chemicals Inc.; St. Louis, MO) y se

mantuvo 4 h más en el incubador en presencia de los tratamientos adecuados.

Transcurrido este tiempo, las células fueron lavadas 2 veces con buffer PBS

(Gibco), cosechadas en 500 µl de buffer STED (0,25 M sacarosa; 10 mM Tris

HCl pH 7,4; 1 mM EDTA; 1 mM DTT) con la ayuda de un rascador y lisadas

mediante 2 ciclos de congelación descongelación en nitrógeno líquido. Para la

extracción de los lípidos neutros, en tubos de vidrio se distribuyeron 400 µl de

los lisados obtenidos y 2,5 ml de metanol/cloroformo (1:2). La mezcla se agitó

durante 10 min y se centrifugó a 2500 rpm durante 10 min y a temperatura

ambiente. Tras la centrifugación, la fase orgánica inferior obtenida se desecó

bajo vacío y se resuspendió en 40 µl de metanol/cloroformo (1:2). Este volumen

se aplicó en una placa de TLC y se procedió a la separación de los lípidos como

se detalla en el apartado 10 (Medida de la incorporación de ácidos grasos a

triglicéridos en adipocitos subcutáneos humanos). Posteriormente, la fracción

correspondiente a los triglicéridos se recogió en viales para el contador de

centelleo, se añadieron 4 ml de líquido de centelleo EcoScint Ultra (National

diagnostics) y se procedió a la medida de la radiactividad que contenían las

muestras mediante el contador Wallac 1409 (EG&G Company). Los resultados

fueron normalizados por el contenido en proteína del extracto obtenido en

buffer STED y fue medida mediante el método de BCA (Pierce-Thermo

Scientific).

3.13.- CUANTIFICACIÓN DEL CONSUMO DE OXÍGENO EN

ADIPOCITOS SUBCUTÁNEOS HUMANOS.

La medida del consumo de oxígeno en adipocitos subcutáneos humanos se

realizó mediante el BD™ Oxygen Biosensor System (BD biosciences; San José,

CA, USA). Este sistema utiliza un compuesto fluorescente sensible al oxígeno

(tris 1,7-diphenyl-1,10 phenanthroline ruthenium (II) chloride) en una matriz

permeable al gas fijada al fondo de una placa de 96 pocillos, actuando como un

biosensor fluorescente, el cual produce señal a medida que el oxígeno es

consumido en la placa. Para ello, los adipocitos fueron levantados mediante el

uso de 100 µl de tripsina (invitrogen) la cual fue inactivada con 500 µl de un

buffer Hepes fosfato que contiene glucosa 5 mM, 2% de BSA, NaCl 135 mM,

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CaCl2.2H2O 2,2 mM; MgSO4.7H2O 1,25 mM, KH2PO4 0,45 mM, NaH2PO4, 2,17

mM y Hepes 10 mM. Del volumen obtenido en cada suspensión, se procedió a

cargar 200 µl por duplicado en la microplaca para su posterior lectura en un

fluorímetro Polarstar Galaxy (BMG labtech; Ortenberg, Germany) en ciclos de 3

min durante 5 h tras fijar los parámetros óptimos de lectura a 485 nm de

excitación y 630 nm de emisión. Los resultados obtenidos fueron corregidos por

el contenido en proteína de cada pocillo la cual fue medida mediante el método

de BCA (Pierce-Thermo Scientific).

3.14.- ESTUDIO DEL CONTENIDO DE DNA MITOCONDRIAL

El estudio de la biogénesis mitocondrial se basó en el análisis de la

expresión génica por qRT-PCR, descrito anteriormente, de un gen codificado

por el mtDNA y dependiente de la maquinaria genética mitocondrial para su

transcripción y traducción. En este estudio el gen elegido fue el Mtco2 (Applied

Biosystems) que codifica para la subunidad II del Complejo IV de la cadena

transportadora de electrones. La expresión de este gen se corrigió por los

niveles de mRNA de un gen nuclear endógeno, en nuestro caso el 18s,

empleándose este ratio como el número de copias de mtDNA y una

aproximación fiable de la población mitocondrial del tejido {Guo et al., 2011;

Pagel-Langenickel et al., 2008).

3.15.- CUANTIFICACIÓN DEL CONTENIDO MITOCONDRIAL

MEDIANTE MITOTRACKER GREEN EN ADIPOCITOS

SUBCUTÁNEOS HUMANOS.

La tinción de mitocondrias MitoTracker Green FM es un colorante

fluorescente que permite localizar y cuantificar selectivamente las mitocondrias

independientemente de su potencial de membrana. Para analizar el efecto del

tratamiento con α-LA sobre el contenido mitocondrial, en el día 14 de

diferenciación los adipocitos fueron tratados con α-LA (100 y 250 µM) o

vehículo. Tras 24 h de tratamiento se retiró el medio de cultivo, se procedió a

lavar las células 2 veces con PBS (gibco) y se incubaron con el medio de cultivo

AM1 al que se añadió la sonda MitoTracker Green en una concentración de 100

nM durante 30 min. Tras este período de tiempo las células fueron lavadas y se

procedió a la visualización de la fluorescencia emitida mediante el microscopio

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Leica DM IL-EL 6000 (Leica Microsystems; Germany) y se tomaron fotografías

con la cámara Leica DFC 345FX (Leica Microsystems). Además, la

fluorescencia emitida se cuantificó en el fluorímetro Polarstar Galaxy (BMG

labtech) cuyos parámetros óptimos de lectura fueron fijados a 554 nm de

excitación y 576 nm de emisión.

3.16.- ANÁLISIS DEL CONTENIDO MITOCONDRIAL MEDIANTE

MICROSOCOPÍA ELECTRÓNICA DE TRANSMISIÓN (TEM)

El contenido mitocondrial en adipocitos subcutáneos humanos en cultivo fue

evaluado mediante la visualización con un microscopio electrónico de

transmisión de secciones ultrafinas de dichos adipocitos. Para ello, los

adipocitos en cultivo, tras los tratamientos oportunos, fueron prefijados en

glutaraldehido al 4% en buffer cacodilato 0,1M durante 1 h a 4ºC.

Posteriormente dichos adipocitos fueron fijados mediante una solución de

tetróxido de osmio al 1% mediante su incubación durante 1 h a 4 ºC. Tras el

proceso de fijación, se procedió a liberar las células de la placa mediante el uso

de un rascador y se incluyeron en agarosa al 2%. Tras esto las muestras fueron

embebidas en una resina Epoxi de baja viscosidad (SERVA Electrophoresis

GmbH, Heidelberg, Alemania) y cortadas en secciones de 60-70 nm mediante el

ultramicrotomo “Leica ultracut R Ultramicrotome” (Leica Microsystems GmbH,

Wetzlar, Alemania). Los cortes ultra-finos fueron examinados con el

microscópio electrónico “energy filter transmission electron microscope (EFTEM)

Libra 120” (Zeiss GmbH, Oberkochen, Alemania) y las imágenes fueron

obtenidas mediante el software iTEM 5.1 (Olympus Soft Imaging Solutions

GmbH, Münster, Alemania).

3.17.- DETECCIÓN DE UCP1 MEDIANTE INMUNOFLUORES-

CENCIA

La evaluación de los niveles de UCP1 en adipocitos subcutáneos humanos

de sujetos con sobrepeso u obesidad fue realizada mediante la técnica de

inmunofluorescencia. Para ello las células se trataron con α-LA (250 µM) o el

vehículo durante 24 h como se ha descrito anteriormente. Tras este tiempo las

células se lavaron 2 veces con PBS y se fijaron en 500 µl de paraformaldehido

al 4% en PBS pH 7,4 durante 15 min a temperatura ambiente. Transcurrido este

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Material y Métodos

88

tiempo las células se lavaron de nuevo 2 veces con PBS en frío y se

permeabilizaron mediante su incubación durante 10 min con PBS que contenía

0,25% de Tritón X-100. Posteriormente se procedió a bloquear las posibles

uniones inespecíficas y para ello tras lavar de nuevo las células 2 veces con

PBS en frío, se incubaron con BSA al 1% en PBST durante 30 min a

temperatura ambiente. Tras el bloqueo las células se incubaron con el

anticuerpo primario (UCP1, Abcam) durante 1 h a temperatura ambiente.

Posteriormente se lavaron las células 3 veces con PBS y se añadió el

anticuerpo secundario (Alexa fluor® 647; Invitrogen) el cual se incubó en las

mismas condiciones que el anticuerpo primario, 1 h a temperatura ambiente.

Finalmente UCP1 fue detectada mediante el microscopio invertido Leica DM IL-

EL 6000 (Leica Microsystems GmbH).

3.18.- ANÁLISIS ESTADÍSTICO

El análisis estadístico de todas las variables se realizó utilizando el

programa informático GraphPad Prism 5.0 (GraphPad Software Inc, San Diego.

CA, Estados Unidos). Se seleccionaron como estadísticos descriptivos de cada

muestra la media como medida de tendencia central, y el error estándar de la

media como medida de dispersión. El nivel de significación estadístico se situó

en todos los casos en P<0,05.

La normalidad de todas las variables se verificó utilizando el programa

estadístico SPSS para Windows versión 19.0 (SPSS Inc. Chicago, Estados

Unidos) mediante los tests de Kolmogorov-Smirnof y Shapiro Wilk. Cuando las

variables siguieron una distribución normal las comparaciones entre dos grupos

se realizaron mediante un test de t-Student. Las comparaciones entre más de

dos grupos fueron analizadas mediante ANOVA de un factor seguido de un test

a posteriori de Bonferroni. Cuando las muestras no siguieron una distribución

normal se aplicaron los test de Kruskal Wallis o de U-Mann-Whitney.

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Material y Métodos

89

REFERENCIAS

Bourlier, V., C. Saint-Laurent, et al. (2013). "Enhanced glucose metabolism is

preserved in cultured primary myotubes from obese donors in response to

exercise training." J Clin Endocrinol Metab 98(9): 3739-47.

Guo, X. E., C. F. Chen, et al. (2011). "Uncoupling the roles of the SUV3

helicase in maintenance of mitochondrial genome stability and RNA

degradation." J Biol Chem 286(44): 38783-94.

Mercader, J., A. Palou, et al. (2011). "Resveratrol enhances fatty acid

oxidation capacity and reduces resistin and Retinol-Binding Protein 4

expression in white adipocytes." J Nutr Biochem 22(9): 828-34.

Moreno-Aliaga, M. J. and F. Matsumura (2002). "Effects of 1,1,1-trichloro-2,2-

bis(p-chlorophenyl)-ethane (p,p'-DDT) on 3T3-L1 and 3T3-F442A

adipocyte differentiation." Biochem Pharmacol 63(5): 997-1007.

Ortega, F. J., J. M. Moreno-Navarrete, et al. (2010). "MiRNA expression profile

of human subcutaneous adipose and during adipocyte differentiation."

PLoS One 5(2): e9022.

Pagel-Langenickel, I., J. Bao, et al. (2008). "PGC-1alpha integrates insulin

signaling, mitochondrial regulation, and bioenergetic function in skeletal

muscle." J Biol Chem 283(33): 22464-72.

Perez-Matute, P., M. J. Neville, et al. (2009). "Transcriptional control of human

adipose tissue blood flow." Obesity (Silver Spring) 17(4): 681-8.

Suzuki, R., M. Tanaka, et al. (2011). "Anthocyanidins-enriched bilberry extracts

inhibit 3T3-L1 adipocyte differentiation via the insulin pathway." Nutr Metab

(Lond) 8: 14.

Tomlinson, J. J., A. Boudreau, et al. (2010). "Insulin sensitization of human

preadipocytes through glucocorticoid hormone induction of forkhead

transcription factors." Mol Endocrinol 24(1): 104-13.

Vigilanza, P., K. Aquilano, et al. (2011). "Modulation of intracellular glutathione

affects adipogenesis in 3T3-L1 cells." J Cell Physiol 226(8): 2016-24.

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CCHHAAPPTTEERR 44

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CCHHAAPPTTEERR 44..11

EEffffeeccttss ooff lliippooiicc aacciidd oonn lliippoollyyssiiss iinn 33TT33--LL11 aaddiippooccyytteess

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Results (Chapter 4.1)

95

Effects of lipoic acid on lipolysis in 3T3-L1 adipocytes

Marta Fernández-Galilea, Patricia Pérez-Matute, Pedro L Prieto-Hontoria, J

Alfredo Martinez, Maria J Moreno-Aliaga

J Lipid Res. 2012 Nov;53(11):2296-306

ABSTRACT

α-Lipoic acid (α-LA) is a naturally occurring compound with beneficial effects

on obesity. The aim of this study was to evaluate its effects on lipolysis in 3T3-

L1 adipocytes and the mechanisms involved. Our results revealed that α-LA

induced a dose and time-dependent lipolytic action, which was reversed by pre-

treatment with the JNK inhibitor SP600125, the PKA inhibitor H89 and the AMPK

activator AICAR. In contrast, the PI3K/Akt inhibitor LY294002 and the PDE3B

antagonist Cilostamide enhanced α-LA-induced lipolysis. α-LA treatment during

1 h did not modify total protein content of HSL, but significantly increased the

phosphorylation of HSL both at Ser563

and at Ser660

, which was reversed by H89.

α-LA treatment also induced a marked increase in PKA-mediated perilipin

phosphorylation. α-LA did not significantly modify either the protein levels of

ATGL or its activator CGI-58 and inhibitor G0S2. Furthermore, α-LA caused a

significant inhibition of AdPLA protein and mRNA levels in parallel with a

decrease in the amount of PGE2 released and an increase in cAMP content.

Together, these data suggest that the lipolytic actions of α-LA are mainly

mediated by phosphorylation of HSL through cAMP-mediated activation of

protein kinase A probably through the inhibition of AdPLA and PGE2.

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Results (Chapter 4.1)

96

INTRODUCTION

α-Lipoic acid (α-LA) or 1,2-dithiolane-3-pentaenoic acid is a naturally

occurring compound that contains two thiol groups with diverse beneficial effects

on health. The biological effects of α-LA were primarily associated with its

antioxidant properties. In fact, α-LA is able to directly scavenge reactive oxygen

species (ROS) and regenerate endogenous antioxidants such as glutathione,

and vitamins E and C (Scholich et al., 1989; Han et al., 1995). Moreover, several

studies have described potential beneficial effects of α-LA on obesity and

associated commorbidities such as insulin resistance, type 2 diabetes or fatty

liver diseases. Thus, in rodents α-LA has been shown to cause profound weight

loss by reducing food intake and enhancing energy expenditure (Kim et al.,

2004) as well as by inducing a reduction on intestinal sugar absorption (Prieto-

Hontoria et al., 2009). More recently, two clinical trials in humans reported that

α-LA caused significant reductions of body weight, body mass index (BMI),

blood pressure and abdominal circumference in obese subjects (Carbonelli et

al., 2010; Koh et al., 2011). α-LA also improved insulin sensitivity and plasma

lipid profile possibly through amelioration of oxidative stress and chronic

inflammatory status in obese patients with impaired glucose tolerance (Zhang et

al., 2011). Previous studies provided strong evidences that α-LA is able to

deeply affect adipose tissue development and function by the inhibition of

adipogenesis (Cho et al., 2003), the regulation of the secretion of several

adipokines such as leptin (Prieto-Hontoria et al., 2011) and apelin (Fernandez-

Galilea et al., 2011) and also by the promotion of mitochondrial biogenesis

(Shen et al., 2011).

In this context, previous studies suggested that α-LA seems to stimulate the

lipolytic response in an in vitro model of broiler chicken adipocytes (Hamano,

2006). However, the molecular mechanisms that mediate these effects remain

unclear. Lipolysis is a complex process highly regulated, which involves the co-

ordinately participation of several lipases and lipid droplet (LD) proteins

(Ahmadian et al., 2009). Thus, the lipolytic process occurs through the

consecutive action of three lipases: adipose triglyceride lipase (ATGL/desnutrin),

hormone-sensitive lipase (HSL) and monoacylglycerol lipase (MAGL) (Lass et

al., 2011). ATGL exhibits high substrate specificity for triacyl glycerol (TAG)

(Villena et al., 2004). Lipase activity of ATGL largely depends on its coactivation

by CGI-58, while G0S2 acts as an inhibitor of ATGL activity and ATGL-mediated

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Results (Chapter 4.1)

97

lipolysis (Lu et al., 2010). Recently it has been shown that ATGL is

phosphorylated by AMPK at Ser406

, increasing TAG hydrolase activity

(Ahmadian et al., 2011).

The activity of HSL is well known to be regulated post-transcriptionally by

reversible phosphorylation. In murine adipocytes PKA phosphorylates HSL at

several serine residues (563, 659, and 660) resulting in increased translocation

of HSL to the lipid droplet surface and increased lipolytic activity (Watt et al.,

2006). Furthermore, AMP-activated protein kinase (AMPK) phosphorylates HSL

at Ser565

, which prevents phosphorylation induced by PKA (Anthony et al., 2009;

Gaidhu et al., 2009). Activation of phosphodiesterase 3B (PDE3B) via the Akt-

mediated phosphorylation of Ser273

attenuates PKA activity and thereby HSL

activation and lipolysis (Degerman et al., 1998; Kitamura et al., 1999). In

addition to the PKA-mediated phosphorylation, HSL may be also phosphorylated

by other kinases such as ERK1/2, which activates HSL by phosphorylation on

Ser600

(Greenberg et al., 2001). It has been also suggested that JNK could play

a role in the regulation of lipolysis based on the fact that silencing of Jnk1 and

Jnk2 accelerates basal lipolysis in mouse adipocytes (Rozo et al., 2008).

Protein trafficking and specific protein-protein interactions at the surface of

lipid droplets are also key factors in the regulation of lipolysis. Perilipin A is a

lipid droplet scaffold protein that plays a central role in orchestrating interactions

among lipolytic effector proteins (Granneman et al., 2009). Under basal

conditions, perilipin restricts the access of cytosolic lipases to LD, thereby

maintaining a low rate of basal lipolysis. However, the phosphorylation of

perilipin by PKA results in perilipin conformational changes that expose LD

stores and facilitates the translocation of phosphorylated HSL to the LD, thereby

increasing the lipolytic process (Miyoshi et al., 2007).

Recently, a novel intracellular adipose-specific phospholipase A2 (AdPLA)

has been identified (Duncan et al., 2008). It was suggested that AdPLA could be

another mediator in the regulation of lipolysis by generating arachidonic acid for

the production of prostaglandins (Jaworski et al., 2009). In fact, AdPLA null

mice exhibited reduced adipose tissue prostaglandin E2 (PGE2) production, and

augmented HSL-phosphorylation leading to increased lipolysis, supporting that

AdPLA is a major regulator of adipocyte lipolysis by regulating PGE2 abundance

(Jaworski et al., 2009).

Previous studies have demonstrated the ability of α-LA to modulate ERK,

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Results (Chapter 4.1)

98

JNK and Akt signaling pathways (Cho et al., 2003; Min et al., 2010; Prieto-

Hontoria et al., 2011), as well as AMPK activity (Packer and Cadenas, 2008;

Cheng et al., 2011) in different cell types. Moreover, α-LA stimulates cAMP

production in purified human NK cells (Salinthone et al., 2011) and modulates

the production of PGE2 in osteoblasts (Ha et al., 2006).

Based on these previous findings, we hypothesized that α-LA could be a key

regulator of lipolysis in mammals through modulation of lipases and lipid droplet

proteins activities. Therefore, the aim of this study was to characterize the

lipolytic action of α-LA in cultured adipocytes and to elucidate the molecular

mechanisms and signaling pathways involved.

MATERIAL AND METHODS

Cell culture and differentiation of 3T3-L1 cells

Murine 3T3-L1 cells (American Type Culture Collection, Rockville, MD, USA)

were cultured in Dulbecco´s modified Eagle´s medium (DMEM) containing 25

mM glucose, 10% calf bovine serum (Invitrogen, CA, USA) and 1% penicillin and

streptomycin (Gibco, Invitrogen Corporation, CA, USA) and were maintained in

an incubator set up to 37 ºC and 5% of CO2. At confluence pre-adipocytes were

induced to differentiate into adipocytes by culturing them for 48 h in DMEM

containing 10% fetal bovine serum (FBS) (Invitrogen), antibiotics and

supplemented with dexamethasone (1 mM; Sigma, St. Louis, MO, USA),

isobutylmethylxantine (0.5 mM; Sigma) and insulin (10 mg/mL; Sigma). Then,

cells were cultured with 10% FBS and insulin for 48 h. After that, media was

replaced with 10% FBS in DMEM and antibiotics, but without insulin and this

media was changed every 2 days up to day 8 post confluence, when cells were

completely differentiated to adipocytes (Lorente-Cebrian et al., 2009; Lorente-

Cebrian et al., 2010).

Treatments

α-LA (Sigma) was dissolved in ethanol. The inhibitors SP600125 (SP)

(Calbiochem, San Diego, CA, USA), PD98059 (PD) (Sigma), H89 (Santa Cruz,

Santa Cruz, CA, USA), LY294002 (LY) (Sigma), Cilostamide (CILO) (Sigma),

and L798106 (Tocris, Ellisville, MO) were dissolved in DMSO. The AMPK

activator AICAR (Sigma) was dissolved in ultrapurified water. All compounds

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Results (Chapter 4.1)

99

were prepared as 1000x stock solutions and added to the culture medium.

Control cells were treated with the same amount of the corresponding vehicle.

Prior to the addition of the appropriate treatments, fully differentiated 3T3-L1

adipocytes were serum starved for at least 4 h (by switching to DMEM

containing 2-2.5% FA free-BSA or to DMEM with 1% FBS) and then treated with

or without α-LA (1-500 µM) during different time intervals (30 min to 24 h). In

order to analyze the signaling pathways involved in α-LA-actions, adipocytes

were pre-incubated for 1h with the selective inhibitors or activators (20 µM SP,

50 µM PD, 1 µM H89, 50 µM LY, 2 µM CILO, 10 µM L798106 and 2 mmol/l

AICAR).

Lipolysis measurement

Lipolysis was evaluated by measuring the amount of glycerol and FFA

released to the media. Glycerol was determined after 1 to 24 h of α-LA treatment

using an autoanalyser following the manufacturer instructions (Cobas-Mira,

Roche Diagnostics, Basel, Swiss). FFA were quantified after 3 h of α-LA

treatment by using the Lipolysis Assay KIT for Free Fatty Acids Detection (Zen-

Bio Inc, Research Triangle Park, NC) according to the manufacturer’s

instructions.

Analysis of mRNA levels

Total RNA was extracted from 3T3-L1 cells using TRIzol® reagent

(Invitrogen) according to the manufacturer’s instructions. RNA concentrations

and quality were measured by Nanodrop Spectrophotometer ND1000 (Thermo

Scientific, Wilminton, DE, USA). RNA was then incubated with the RNAse-free

kit DNAse (Ambion, Austin, TX, USA) for 30 min at 37 ºC. RNA (2 µg) was

reverse-transcribed to cDNA using MMLV (Moloney Murine Leukaemia Virus)

reverse transcriptase (Invitrogen). For the real time quantitative polymerase

chain reaction analysis, 4.5 µl of 1/100 or 1/50 dilution of cDNA per reaction

were used in a final reaction volume of 10 µl.

Atgl, Hsl, Plin1, AdPLA, Pparγ, C/ebpα and C/ebpβ mRNA levels were

determined using predesigned Taqman® Assays-on-Demand (Applied

Biosystems, Foster City, CA, USA). Taqman Universal Master Mix was also

provided by Applied Biosystems. The reaction conditions were followed

according to manufacturer’s instructions.

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Results (Chapter 4.1)

100

Amplification and detection of specific products were performed using the

ABI PRISM 7900HT Fast System Sequence Detection System (Applied

Biosystems).

All mRNA levels were normalized by the housekeeping gene β-Actin

obtained from Applied Biosystems. Samples were analyzed in duplicate. Ct

values (the cycle where the emitted fluorescence signal is significantly above

background levels and is inversely proportional to the initial template copy

number) were generated by the ABI software. Finally, the relative expression

level of each gene was calculated as 2-ΔΔCt

(Perez-Matute et al., 2009).

Western blot analyses

Western blot analyses were performed in 8 days post-differentiation

adipocytes. Cells were incubated in serum-free DMEM overnight and then with

or without the appropriate treatment. Lysates were obtained by the addition of a

buffer containing: 2 mM Tris HCl (pH 8); 137 mM NaCl; 2 mM EDTA; 1%

protease inhibitor cocktail 1 (Sigma); 1 mM Sodium orthovanadate and 1 mM

PMSF. Protein extracts were collected after sample centrifugation. Proteins were

quantified with the BCA method according to the supplier´s instructions (Pierce-

Thermo Scientific, Rockford, IL, USA). Total proteins were resolved in SDS-

PAGE minigels and electroblotted onto PVDF membranes (GE Healthcare

Europe GmbH, Barcelona, Spain). The membranes were blocked and incubated

with specific antibodies against ATGL, HSL, HSL phospho Ser565

, HSL phospho

Ser563

, HSL phospho Ser660

, perilipin, phospho (ser/thr) PKA substrate (p-

perilipin), AMPK, AMPK phospho Thr172

, AKT, AKT phospho Ser473

, MAPK

(ERK1/2), ERK1/2 phospho Thr202/204

, JNK and JNK phospho Thr183/

Tyr185

(from

Cell Signaling Technologies, Beverly, MA, USA), AdPLA (from Cayman

Chemical, Ann Arbor, MI, USA), CGI-58, G0S2 (from Santa Cruz) and Actin

(from Sigma). Secondary antibody was horseradish peroxidase goat anti-rabbit

IgG-HRP (Bio Rad Laboratories). The immunoreactive proteins were detected

with enhanced chemiluminescence (Pierce Biotechnology, Rockford, Illinois,

USA). Band intensities were quantified using a GS-800 calibrated densitometer

(Bio Rad Laboratories).

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Results (Chapter 4.1)

101

Fatty acid oxidation determination

FA oxidation to acid-soluble metabolites (ASM) was measured with

radiolabeled 14

C- palmitate (Perkin Elmer, Boston, MA) in mature 3T3-L1

adipocytes as previously described (Mercader et al., 2011). ASM were extracted

by addition of 1 ml cold 1 M HClO4 (Panreac, Barcelona, Spain). After

centrifugation (10 min, 1800 g), radioactivity in the supernatant was measured

by scintillation counting by using a Wallac 1409 liquid scintillation counter

(EG&G Company, Turku, Finland). Protein content in parallel cultures of vehicle-

and α-LA-treated cells was analyzed using a BCA method.

ELISA assays

Prostaglandin E2 (PGE2) concentration in the media was determined after

24 h of α-LA (250 µM) treatment by using a PGE2 Enzyme Immunoassay kit

(Arbor assays, Ann Arbor, MI, USA). The amount of intracellular cAMP was

quantified after 1 and 24 h of α-LA (250 µM) treatment by using the cAMP Direct

EIA kit (Arbor Assays).

Data analysis

Data are expressed as mean ± standard errors (SE). Differences were set up

as statistically significant at P<0.05. Comparisons between the values for

different variables were analyzed by one-way ANOVA, followed by Bonferroni

post hoc tests, or by Student’s t test or U-Mann Whitney once the normality with

the Kolmogorov-Smirnoff and Shapiro-Wilk tests was screened. SPSS 19.0

version for Windows (SPSS, Chicago, IL, USA) and GraphPad Prism 5.0

(Graph-Pad Software INC. San Diego, CA, USA) were used for the statistical

analysis.

RESULTS

Effects of α-LA on lipolysis in 3T3-L1 adipocytes

A dose-dependent significant increase in the amount of glycerol released into

the media was observed in those adipocytes treated with α-LA (250-500 µM,

P<0.01 and P<0.001) for 24 h (Fig. 1A). Moreover, the lipolytic effect of α-LA

was time-dependent. Thus, the significant increase in glycerol release was

observed after 1 h of treatment (P<0.05) and it became more prominent after 3

and 6 h of treatment (250-500 µM, (P<0.001) (Fig. 1B). Furthermore, α-LA lso

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Results (Chapter 4.1)

102

induced a concentration-dependent increase in the amount of FFA released

after 3 h treatment (100-500 µM, P<0.001) (Fig. 1C).

0

500

1000

1500

**

***

Gly

ce

rol

(µm

ol/

L)

0 1 10 100 250 500 LA (µM)

A

C

0

20

40

60

80

100

***

***

***

0 1 10 100 250 500 LA (µM)

FF

A (

µm

ol/

L)

Length of treatment (h)

0 1 2 3 4 5 6 780

100

120

140

160

180

***

***

***

***

*

*Gly

ce

rol

(%)

Basal

LA 250 µM

LA 500 µM

B

Fig. 1. α-LA stimulates lipolysis in 3T3-L1 adipocytes. Mature 3T3-L1 adipocytes were treated

with α-LA (0-500 µM) for the indicated times (1, 3, 6 or 24 h). A: Lipolysis was assessed by the amount of

glycerol released into media in adipocytes treated for 24 h. B: Time-dependent effects of α-LA (250 and 500

µM) on glycerol release. C: Concentration-dependent effects of α-LA on FA release in adipocytes treated

during 3 h. Data are expressed as mean ± S.E. of 6 independent experiments. *P <0.05, **P<0.01 and

***P<0.001 vs. Control (vehicle-treated cells).

We also tested the effects of α-LA on isoproterenol-induced lipolysis and the

data revealed that α-LA did not have any additional effect on the lipolytic effect

of isoproterenol (Supplementary Fig. I).

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Results (Chapter 4.1)

103

Supplementary Fig. I. α-LA stimulates basal but not isoproterenol-induced lipolysis. Differentiated

3T3-L1 adipocytes were treated with α-LA (250 μM) in the absence or presence of isoproterenol (10-6 M) for

24 h. Lipolysis was estimated by measuring the amount of glycerol released into media. Data are means ±

SE of 4 independent experiments. ***P<0.001 vs. Control (vehicle-treated cells).

In order to rule out if the lipolytic effect of α-LA was caused by a global down-

regulation of adipocyte differentiation markers, Pparγ, C/ebpα and C/ebpβ gene

expression levels were analyzed after 24 h of α-LA (250 µM) treatment and no

differences were observed when compared with control cells (Supplementary

Fig. II).

Supplementary Fig. II. α-LA treatment does not modify adipocyte differentiation markers in

mature 3T3-L1 adipocytes. Differentiated 3T3-L1 adipocytes were treated with α-LA (250 μM) for 24 h.

mRNA levels of several adipogenic factors (Pparγ, C/ebpα and C/ebpβ) were determined by RT-PCR. Data

are means ± SE of 5 independent experiments.

Moreover, to test if the lipolytic actions of α-LA were also shared by other

molecules with antioxidant properties, the effects of vitamin C, resveratrol, N-

acetyl cysteine (NAC) and butylhydroxyanisole (BHA) on glycerol release were

evaluated. The data showed that, at the concentration tested, resveratrol and

BHA, but not NAC or Vitamin C were able to stimulate lipolysis in 3T3-L1

adipocytes (Supplementary Fig. III).

Control LA ISO ISO+LA0

50

100

150

200*** ***

***

Gly

cero

l (%

)

0.0

0.5

1.0

1.5

2.0

Ppar C/ebp C/ebp

mR

NA

/ A

cti

n

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Results (Chapter 4.1)

104

Supplementary Fig. III. Differential effects on lipolysis of several antioxidants. The amount of

glycerol released into media was determined in fully differentiated 3T3-L1 adipocytes treated with α-LA (250

μM), Vit C (250 μM), resveratrol (50 μM), NAC (20 mM) and BHA (10 mM) during 24 h. Data are means ± SE

of at least 3 independent experiments. **P <0.01 and ***P <0.001 vs. Control (vehicle-treated cells).

Signaling pathways involved in the lipolytic actions of α-LA

In order to evaluate the ability of α-LA to modify some signaling pathways

involved in the regulation of lipolysis, the phosphorylation levels of JNK, ERK1/2,

AMPK and PI3K/AKT were analyzed both after a short (30 min-1h) and long-

term treatment (24 h).

No effects were observed in JNK Thr183

/Tyr185

phosphorylation after 30 min

of treatment with α-LA (250 µM), while a significant (P<0.05) reduction of JNK

phosphorylation was observed after 24 h of treatment (Fig. 2A). In contrast, the

significant increase on ERK1/2 Thr202

/Tyr204

phosphorylation (P<0.01) induced

by α-LA (250 µM) after 1 h was reversed to basal levels after 24 h of treatment

(Fig. 2B). Moreover, the stimulatory effect (P<0.05) of α-LA on AMPK Thr172

phosphorylation was only observed in long-term treated (24 h) adipocytes (Fig.

2C). Regarding the PI3K/AKT signaling pathway, α-LA (250 µM) treatment

caused a significant inhibition of AKT Ser473

phosphorylation both at short

(P<0.05) and long-term (P<0.01) treatments (Fig. 2D).

For a better understanding of the potential signaling pathways involved in the

lipolytic action of α-LA, the effects of specific kinase inhibitors or activators on α-

LA-induced glycerol release were studied. Basal lipolysis was significantly

enhanced by the PI3K/Akt inhibitor LY294002 (P<0.001) and the PDE3B

antagonist Cilostamide (P<0.01), and decreased by the PKA inhibitor H89 and

the AMPK activator AICAR (P<0.001). Interestingly, our data revealed that the

lipolytic actions of α-LA were reversed by pre-treatment with the JNK inhibitor

SP600125 (P<0.01), the PKA inhibitor H89 and the AMPK activator AICAR

Control LA Vit C Resv NAC BHA0

50

100

150

200

****

***

Gly

cero

l (%

)

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Results (Chapter 4.1)

105

(P<0.001). Interestingly, the stimulatory effects of α-LA on lipolysis were

significantly enhanced (P<0.01 and P<0.001) in adipocytes treated with the

PI3K/AKT inhibitor LY294002 and the PDE3B antagonist Cilostamide (Fig. 2E).

Basal SP AICAR PD H89 LY CILO

0

50

100

150

200

250

***

##b###c

***

#

***#b

***

###c

##c

**

Gly

ce

rol (%

)

0.0

0.5

1.0

1.5

2.0

*

0

1

2

3

4

5 **

P-E

RK

1/2

/ER

K1

/2

P-J

NK

/JN

K

1h 24h 1h 24h

30 min 24h 1h 24h

LA 250 µM - + - + LA 250 µM - + - +

0.0

0.5

1.0

1.5

**

*

P-A

KT

/AK

T

1h 24h

1h 24h1h 24h

0.0

0.5

1.0

1.5

2.0 *

1h 24h

P-A

MP

K/A

MP

K

A

E

B

C D

p-JNK

JNK

p-ERK1/2

ERK1/2

AMPK

p-AMPK p-AKT

AKT

LA 250 µM - + - + LA 250 µM - + - +

Fig. 2. Signaling pathways involved in the lipolytic effects of α-LA. A-D: Effects of α-LA on the

phosphorylation of (A) JNK, (B) ERK1/2, (C) AMPK and (D) PI3K/AKT. Band intensities for each

phosphorylated species were normalized to their respective total fractions. E: Effects of α-LA treatment

during 24 h on glycerol release in the presence or absence of the JNK inhibitor SP600125 (SP), the AMPK

activator AICAR, the ERK1/2 inhibitor PD98059 (PD), the PKA inhibitor H89, the PI3K/AKT inhibitor

LY294002 (LY) and the PDE3B inhibitor Cilostamide (CILO). Data are expressed as mean ± S.E. of at least 3

independent experiments. *P <0.05, **P<0.01 and ***P<0.001 vs. Basal Control (vehicle-treated cells)

#P<0.05, ##P<0.01 and ###P<0.001 vs. respective control, bP<0.01 and cP<0.001 vs. basal α-LA-treated

adipocytes.

Effects of α-LA treatment on HSL, ATGL, Perilipin, CGI-58 and G0S2

levels

In contrast to the α-LA lipolytic actions, a significant (P<0.05) decrease on

total protein content of the two main lipases ATGL and HSL was observed in α-

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Results (Chapter 4.1)

106

LA-treated (250 µM for 24 h) adipocytes (Fig. 3A). Accordingly, gene expression

levels of both Atgl and Hsl were also significantly downregulated (P<0.05) by α-

LA treatment for 24 h (Fig. 3B). Perilipin mRNA levels were also reduced in α-

LA-treated adipocytes while no changes in perilipin protein content were

observed (Fig. 3A and 3B).

Basal SP AICAR PD H89 LY0

1

2

3

4

* *

a

**

**

a##

Per

ilip

in m

RN

A/

Act

in

Basal SP AICAR PD H89 LY0.0

0.5

1.0

1.5

2.0

2.5

*

***

#a a

**##

Atg

l mR

NA

/A

ctin

Basal SP AICAR PD H89 LY0.0

0.5

1.0

1.5

2.0

2.5

**

**

##

* a

**

*

Hsl

mR

NA

/ A

ctin

0.0

0.5

1.0

1.5

* *

0.0

0.5

1.0

1.5

*

*

*

Pro

tein

/Ac

tin

ATGL HSL Perilipin

ATGL HSL PERILIPIN

- + - + - + LA 250 µM

A

C

D

E

ACTIN

Atgl Hsl Perilipin

mR

NA

/βA

cti

n

B

Fig. 3. Long-term α-LA treatment downregulates total HSL, ATGL and perilipin transcripts. The

effects of α-LA (250 µM) on total ATGL, HSL and perilipin protein (A) and mRNA (B) levels were assessed in

3T3-L1 adipocytes after 24 h of treatment. C-E: Effects of the JNK inhibitor SP600125 (SP), the ERK1/2

inhibitor PD98059 (PD), the PKA inhibitor H89, the AMPK activator AICAR and the PI3K/AKT inhibitor

LY294002 (LY) on (C) Atgl, (D) Hsl and (E) Perilipin mRNA levels in control and α-LA-treated 3T3-L1

adipocytes. Data are expressed as mean ± S.E. of at least 3 independent experiments. *P<0.05, **P<0.01

and ***P<0.01 vs. Basal Control (vehicle-treated cells). #P<0.05, ##P<0.01 vs. respective control. aP<0.05 vs.

basal α-LA-treated adipocytes.

The inhibitory actions of α-LA treatment on Atgl gene expression were not

observed in presence of the JNK inhibitor SP600125 and ERK1/2 inhibitor

PD98059 (P<0.05) (Fig. 3C). Similarly, the inhibition of the ERK1/2 signaling

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Results (Chapter 4.1)

107

pathway was able to reverse the down-regulation of Hsl and Perilipin gene

expression observed after α-LA treatment (P<0.05) (Fig. 3D and E).

HSL activity is regulated by reversible phosphorylation in serine residues.

PKA phosphorylates HSL at Ser563

, and Ser660

, which stimulates HSL activity.

Thus, to better elucidate the mechanisms underlying the lipolytic actions of α-LA,

we next investigated the effects of α-LA on HSL phosphorylation both in Ser563

and Ser660

. α-LA treatment (250 µM) during 1 h did not modify total protein

content of HSL, but significantly increased (P<0.05) the phosphorylation of HSL

both at Ser563

(Fig. 4A) and at Ser660

(Fig. 4B).

However, α-LA did not modify the AMPK-induced phosphorylation of HSL at

Ser565

(Supplementary Fig. IV).

Supplementary Fig. IV. α-LA does not phosphorylate HSL at Ser565. Representative Western blots

for Ser565-phosphorylated HSL and total HSL in differentiated 3T3-L1 adipocytes treated with α-LA (250 µM)

for 1 h. Band intensities were normalized to total HSL.

These data suggest that α-LA stimulates lipolysis by increasing PKA activity.

Perilipin phosphorylation is also PKA-dependent. Using a perilipin-specific

antibody and a phospho-PKA-motif-specific substrate antibody, we found that α-

LA treatment induced a marked increase (P<0.01) in PKA-mediated perilipin

phosphorylation (Fig. 4C). In fact, the α-LA-induced phosphorylation of HSL at

Ser563

and Ser660

as well as of perilipin was dramatically blunted in the presence

of the PKA inhibitor H89. We also found that AMPK activation disrupted the α-

LA-induced phosphorylation of HSL at both Ser563

and Ser660

(Fig. 4A and 4B),

without modifying the p-PKA substrate/perilipin ratio (Fig. 4C). Interestingly, the

inhibition of the JNK pathway induced a significant increase in the

phosphorylation of HSL at Ser660

, both in the absence and presence of α-LA, and

in PKA-mediated perilipin phosphorylation (Fig. 4B and 4C). Morover, the

ERK1/2 inhibitor PD98059 prevented the α-LA-induced phosphorylation of HSL

at Ser563

without modifying α-LA-effects on Ser660

and p-PKA substrate/perilipin

Control LA0.0

0.5

1.0

1.5

p-H

SL

56

5/H

SL

+ -

P-HSL565

LA 250 µM

P-HSL565

HSL

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Results (Chapter 4.1)

108

ratio. All these data suggest that α-LA stimulates lipolysis mainly through the

PKA-mediated phosporylation of perilipin and HSL.

Basal SP AICAR PD H890

1

2

3

*

a***

ba

p-H

SL

56

3/H

SL

Basal SP AICAR PD H890

1

2

3

4

*

*

* b

#

p-P

KA

su

bstr

ate

/peri

lip

in

P-HSL563

HSL

HSL

P-HSL660

P-PKA substrate

Perilipin

Basal SP AICAR PD H890

1

2

3

4

*

*

a

***c*

a

p-H

SL

660/H

SL

A

C

Basal

LA 250 µM - + - + - + - + - +

SP AICAR PD H89

BBasal

LA 250 µM - + - + - + - + - +

SP AICAR PD H89

Basal

LA 250 µM - + - + - + - + - +

SP AICAR PD H89

Fig. 4. α-LA stimulates PKA-mediated phosporylation of HSL and perilipin. A-B: Representative

Western blots for (A) Ser563-phosphorylated HSL and (B) Ser660-phosphorylated HSL in differentiated 3T3-L1

adipocytes treated with α-LA (250 µM) for 1 h in the presence or absence of the JNK inhibitor SP600125

(SP), the AMPK activator AICAR, the ERK1/2 inhibitor PD98059 (PD) and the PKA inhibitor H89. Band

intensities were normalized to total HSL. C: Adipocyte lysates were also immunoblotted using a phospho-

PKA-motif-specific antibody and then the blots were stripped and reprobed with antiperilipin antibodies to

detect native perilipins. The density of the protein bands was quantified and the data (mean ± S.E.) were

expressed as p-PKA substrate/perilipin ratio. (n ≥ 3 independent experiments). *P<0.05 and ***P<0.001 vs.

Basal Control (vehicle-treated cells). #P<0.05 vs. respective control. aP<0.05, bP<0.01 and cP<0.001 vs. basal

α-LA-treated cells.

However, α-LA treatment during 1 h did not significantly modify the protein

levels of ATGL. Neither CGI-58 nor G0S2, the activator and inhibitor of ATGL

activity respectively, were significantly altered after 1 or 24 h of α-LA treatment

(Fig. 5A and B respectively).

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Results (Chapter 4.1)

109

ATGL CGI-58 G0S20.0

0.5

1.0

1.5

2.0

Pro

tein

/Ac

tin

ATGL

CGI-58

ACTIN

LA (250 µM) - +

G0S2

1h

A

LA (250 µM) - +

24h

0.0

0.5

1.0

1.5

ATGL CGI-58

*

G0S2

Pro

tein

/Ac

tin

ATGL

CGI-58

ACTIN

G0S2

B

Fig. 5. α-LA does not modify the levels of the ATGL co-activator CGI-58 and ATGL inhibitor

G0S2. A-B: Lysates from 3T3-L1 adipocytes treated with α-LA (250 µM) for 1 h (A) and 24 h (B) were

immunoblotted for ATGL, CGI-58, G0S2 and Actin antibody. Band intensities for ATGL, CGI-58 and G0S2

were normalized to Actin. Data are expressed as mean ± S.E. of at least 5 independent experiments.

*P<0.05 vs. Control (vehicle-treated cells).

Effects of α-LA on AdPLA levels, and on PGE2 and cAMP production

AdPLA has been described as the major phospholipase A2 in adipose tissue

with a key role in the regulation of lipolysis through the modulation of PGE2

levels. As shown in Fig. 6A, α-LA treatment during 1 and 24 h (250 µM) caused

a significant inhibition (P<0.05) of AdPLA protein content as well as on mRNA

levels (Figure not shown).

We next aimed to evaluate the effects of α-LA on the major AdPLA product,

PGE2, which binds the Gαi-coupled receptor EP3, and down-regulates lipolysis

by inhibiting cAMP production. Our data showed that the amount of PGE2

released to the media was significantly reduced in α-LA-treated adipocytes at 24

h of treatment (P<0.05) (Fig. 6B) and also at shorter (4 and 8 h) periods of

treatment (data not shown). In parallel, a significant increase in cAMP levels was

found in α-LA-treated adipocytes for 1 and 24 h (Fig. 6C). Moreover, the lipolytic

effect of α-LA was partially reversed by co-treatment with PGE2, an effect that

was not observed in the presence of the EP3-receptor antagonist L798106 (Fig.

6D).

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Results (Chapter 4.1)

110

0

50

100

150

200

1 h 24 h

***

cA

MP

(% o

f C

on

tro

l)

Control LA0

500

1000

1500

*

Pro

sta

gla

nd

in E

2

(pg

/ml)

0.0

0.5

1.0

1.5

*

*

1 h 24 h

Ad

PL

A/A

CT

IN

AdPLA- + - + LA 250 µM

A

1 h

Actin

24 h

B C

Contr

olLA

2

PGE

+ L

7810

6

2

PGE

2

LA +

PGE

+ L

7810

6

2

LA +

PGE

0

50

100

150

200

250

**

a

**

#

***

Gly

ce

rol (%

)D

Fig. 6. α-LA reduces AdPLA levels and PGE2 secretion and increases intracellular cAMP levels in

3T3-L1 adipocytes. A: AdPLA protein levels at 1 and 24 h of treatment with α-LA (250 µM). B: PGE2

released to the media in 3T3-L1 adipocytes treated with α-LA (250 µM) during 24 h. C: Intracellular cAMP

levels at 1 and 24 h of treatment with α-LA (250 µM). D: Effects of PGE2 (0.5 ng/ml) on the lipolytic action of

α-LA (250 µM) in the presence or absence of the EP3 antagonist L78106 (10 µM). Data are expressed as

mean ± S.E. of at least 3 independent experiments. *P<0.05, **P<0.01 and ***P<0.01 vs. Control (vehicle-

treated cells). #P<0.05 vs. PGE2-treated cells. aP<0.05 vs. basal α-LA-treated cells.

DISCUSSION

Our current data demonstrate the lipolytic action of α-LA in cultured

adipocytes in a concentration and time-dependent manner. It is important to note

that the doses able to induce lipolysis were similar to those that inhibited

adipogenesis in 3T3-L1 preadipocytes (Cho et al., 2003) and no toxicity was

observed. Previous studies in broiler chickens also support the lipolytic action of

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Results (Chapter 4.1)

111

α-LA both in vitro and in vivo models (Hamano, 2002; Hamano, 2006). However,

the mechanisms of action remain uncertain. In the present study we tested if the

lipolytic effects of α-LA were shared by other compounds with antioxidant

properties. Our data revealed that resveratrol and BHA, but not Vitamin C or

NAC were able to stimulate lipolysis, suggesting that the lipolytic actions seem

to be independent of the antioxidant capacities.

Moreover, our data showed that despite the stimulatory effects of α-LA on

lipolysis, both Hsl and Atgl gene expression and protein levels were inhibited

after 24 h of α-LA treatment, together with a decrease in Perilipin mRNA levels.

These effects of α-LA on Hsl, Atgl and Perilipin were reversed by the presence

of the ERK1/2 inhibitor PD98059 in the media. A down-regulation of Hsl, Atgl

and Perilipin gene expression together with increased lipolysis has also been

described after TNF-α treatment in adipocytes (Ryden et al., 2004; Kim et al.,

2006; Kralisch et al., 2008). Moreover, it was observed that the administration of

Trecadrine, a beta-3 adrenergic agonist that stimulates lipolysis (Moreno-Aliaga

et al., 2002), induced a decrease in Hsl mRNA levels in abdominal WAT,

whereas an increase in HSL activity was observed (Berraondo and Martinez,

2000). Furthermore, a recent study reported that serum amyloid A (SAA) also

stimulated lipolysis in parallel with a reduced HSL protein content. However,

SAA caused a significant increase of PKA-mediated HSL phosphorylation (Liu et

al., 2011), suggesting opposite trends in HSL expression and activity. In this

context, the mechanisms controlling HSL activity have been thoroughly studied,

showing that reversible phosphorylation at several serine sites is a hallmark of

HSL regulation. Indeed, HSL is activated by PKA-induced phosphorylation at

Ser563

and Ser660

. Moreover, the lipid droplet protein perilipin is also

phosphorylated by PKA, and upon phosphorylation, perilipin shifts to the

cytoplasm and the accessibility of HSL to the lipid surface is promoted and the

lipolysis enhanced (Holm, 2003; Shen et al., 2009; Xu et al., 2009). The results

of our study suggest a key role of PKA-induced lipolysis in the lipolytic actions of

α-LA because of i) α-LA increased HSL phosphorylation both at Ser563

and

Ser660

; ii) PKA-induced perilipin phosphorylation was increased by α-LA

treatment; iii) the PKA inhibitor H89 completely blunted the lipolytic action of α-

LA as well as the α-LA-induced phosphorylation of phospho-PKA substrates.

Taking together, these data suggest an important role of PKA-mediated

phosporylation of perilipin and HSL in the lipolytic effect of α-LA.

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Results (Chapter 4.1)

112

ATGL plays a governing role in both basal and adrenergically stimulated

TAG breakdown in adipocytes (Lass et al., 2011). However, our data suggest

that ATGL activation is not importantly involved in the lipolytic action of α-LA as

concluded from the findings that no significant changes were observed either on

the levels of the ATGL co-activator protein called CGI-58 or the inhibitory protein

G0S2 (Lu et al., 2010; Yang et al., 2010).

PI3K/AKT is a major player of insulin action and its activation increases

PDE3B activity, and hydrolysis of cAMP leading to a net dephosphorylation of

HSL and inhibition of lipolysis (Ridderstrale, 2005). In our experimental cell

model, α-LA inhibits AKT phosphorylation both at 30 min and 24 h of treatment,

and both the PI3K/AKT inhibitor LY294002 and the PDE3B antagonist

Cilostamide potentiated the stimulatory effects of α-LA on basal lipolysis.

Therefore, the present results suggest that the lipolytic effects of α-LA could be

mediated by decreasing AKT activation, which might increase cAMP, and

lipolysis mediated by HSL and perilipin activation.

Mitogen-activated protein (MAP) kinases are serine/threonine-specific

protein kinases that regulate various cellular activities, including lipolysis.

Regarding the role of JNK activation in the regulation of lipolysis, it was

described that JNK1/JNK2-deficiency drastically enhanced basal lipolysis (Rozo

et al., 2008). In this context, our data show that incubation with the JNK inhibitor

SP600125 (2 h) stimulates the phosphorylation of HSL at Ser563

and Ser660

as

well as phospho-PKA substrate/perilipin ratio, supporting the idea that JNK

inhibition leads to increased lipolysis. However, our current data and previous

studies show that the amount of glycerol released into the media is not modified

or even reduced by longer-term incubation with SP600125 (Ryden et al., 2004;

Deng et al., 2012), suggesting that the effects of JNK inhibition on lipolysis might

be time-dependent. Our results demonstrated that α-LA induced a time-

dependent inhibition of JNK phosphorylation, which might suggest the

involvement of this pathway in the lipolytic actions of α-LA Thus, preincubation

with SP600125 for 1 h potentiated the phosphorylation of HSL at Ser660

observed after 1 h of treatment with α-LA. However, co-incubation with the JNK

inhibitor SP600125 partially reversed the stimulatory effect on lipolysis and the

inhibition induced by α-LA on ATGL gene expression after 24 h of treatment,

suggesting that the involvement of JNK on α-LA-induced lipolysis is complex and

seems also to be time-dependent. On the other hand, the fact that pretreatment

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Results (Chapter 4.1)

113

with the ERK1/2 inhibitor PD98059 reversed the downregulation of Hsl, Atgl and

Perilipin gene expression induced by α-LA treatment during 24 h might suggest

the involvement of this pathway in α-LA-induced lipolysis. However, our data

evidenced that ERK1/2 phosphorylation is not affected by α-LA after 24 h of

treatment and that pretreatment with PD98059 was not able to reverse the

lipolytic action of α-LA, arguing against the involvement of this pathway.

AMPK has been also involved in the regulation of lipolysis (Hardie, 2008;

McGee and Hargreaves, 2010). Thus, it has been reported that phosphorylation

of HSL at Ser565

by AMPK prevents activation by PKA, inhibiting lipolysis (Dagon

et al., 2006; Boon et al., 2008; Anthony et al., 2009). Moreover, the negative

regulation of AMPK activity by PKA has been shown to be important for

converting a lipolytic signal into an effective lipolytic response (Djouder et al.,

2010). However, it has been recently reported that ATGL is

phosphorylated/activated by AMPK to increase lipolysis (Ahmadian et al., 2011).

Thus, the effects described for AMPK activators on lipolysis are controversial

showing both inhibiton (Bourron et al., 2010; Lorente-Cebrian et al., 2012) and

activation of lipolysis (Gaidhu et al., 2009; Ahmadian et al.), and it has been

suggested that the effects of AMPK activation on lipolysis might be time-

dependent (Yin et al., 2003). Our present data show that α-LA treatment

stimulates AMPK phosphorylation and promotes lipolysis. However, the lipolytic

effects of α-LA were already observed after 1 h of treatment when AMPK

phosphorylation was not induced, suggesting that AMPK is not involved in the

short-term lipolytic effects of α-LA. On the contrary, the presence of the AMPK

activator AICAR inhibited α-LA-stimulated lipolysis at 24 h of treatment,

according with the remarkable increase of AMPK phosphorylation observed at

this period of time. Taking together, these data suggest that the lipolytic action of

α-LA is not mediated by the activation of AMPK in the first stages but it could

contribute to the regulation of the long-term lipolytic effects of α-LA.

Recently it has been described and functionally characterized a new

adipocyte phospholipase A2 called AdPLA (Duncan et al., 2008). Afterwards, it

was demonstrated that AdPLA ablation increased lipolysis by reducing PGE2

levels and thereby stimulating cAMP and phosphorylation of HSL through cAMP-

mediated activation of PKA (Jaworski et al., 2009). Our results showed for the

first time that AdPLA is down-regulated by α-LA treatment as well as PGE2

levels, accompanied by an increase in cAMP levels, which could also contribute

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Results (Chapter 4.1)

114

to the increased phosphorylation of HSL at Ser563

and Ser660

and thereby to the

lipolytic effects of α-LA. In support of this, our data revealed that co-incubation

with PGE2 was able to partially reverse the stimulatory effect of α-LA on lipolysis,

while this effect of PGE2 was not observed in the presence of an EP3

antagonist.

All these data suggest that the ability of α-LA to stimulate lipolysis in

adipocytes could also contribute to its antiobesity properties. It is important to

take into consideration that increased lipolysis and FFA release from adipose

tissue has been associated with the development of insulin resistance (Ormseth

et al., 2011). However, recent findings have demonstrated that, surprisingly,

increasing lipolysis in adipose tissue does not necessarily increase serum FFA

levels because increasing lipolysis in adipose tissue causes a shift within

adipocytes towards increased FA utilization and energy expenditure and thus

protects against obesity. Therefore, it has been suggested an activation of

lipolysis may be a promising therapeutic target for the treatment of obesity

(Ahmadian et al., 2009; Ahmadian et al., 2010). In this context, we and others

have demonstrated that dietary supplementation with α-LA reduces weight loss

and fat mass without increasing circulating FFA and improves insulin resistance

both in rodents (Park et al., 2008; El Midaoui et al., 2011; Fernandez-Galilea et

al., 2011) and humans (Zhang et al., 2011), and as previously suggested, this

could be associated to α-LA-induced FA oxidation. In this context, our

experimental data support the notion about the ability of α-LA to promote FA

oxidation in 3T3-L1 adipocytes (Supplementary Fig. V).

Supplementary Fig. V. α-LA increases palmitate oxidation to acid-soluble metabolites.

Fatty acid oxidation was estimated as 14C-labeled palmitate oxidation to acid-soluble metabolites

(ASM) in 3T3-L1 adipocytes treated for 6 h with or without LA (250 µM) in DMEM containing 2.5%

BSA, 200 μM L-carnitine, 200 μM cold palmitic acid and 200 μM [14C(U)] palmitate (0.1 μCi/mL).

The value of a vehicle control was set at 100% and the relative value was presented as fold

induction with respect to that of the vehicle control. Data are means ± SE of 6 independent

experiments. ** P <0.01.

Control LA0

50

100

150

**

Pa

lmit

ate

ox

ida

tio

n

AS

M R

ele

as

e (

%)

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Results (Chapter 4.1)

115

A recent study have also evidenced that α-LA subsequently increased AMPK

and ACC phosphorylation, leading to increased palmitate β-oxidation in

myotubes (Chen et al., 2012). Moreover, studies of our group also have shown

that α-LA supplementation prevents the downregulation of genes involved in

mitochondrial and peroxisomal β-oxidation in liver of high fat-induced obese rats

(Valdecantos et al., 2012).

In summary, the present data demonstrate the ability of α-LA to stimulate

lipolysis in 3T3-L1 adipocytes and suggest that these lipolytic actions of α-LA are

mainly mediated by the phosphorylation of HSL through cAMP-mediated

activation of PKA probably through the inhibition of AdPLA and PGE2.

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CCHHAAPPTTEERR 44..22

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125

α-Lipoic acid reduces fatty acid esterification and

lipogenesis in adipocytes from overweight/obese subjects

Marta Fernández-Galilea, Patricia Pérez-Matute, Pedro L. Prieto-Hontoria,

Marianne Houssier, J. Alfredo Martínez, Dominique Langin, María J. Moreno-

Aliaga.

ABSTRACT

Objective: α-Lipoic acid (α-LA) is a natural occurring antioxidant with

beneficial effects on obesity. The aim of this study was to investigate the

putative effects of α-LA on triglyceride accumulation and lipogenesis in

subcutaneous adipocytes from overweight/obese subjects and to determine the

potential mechanisms involved.

Design and Methods: Fully differentiated human subcutaneous adipocytes

were treated with α-LA (100 and 250 µM) during 24 h for studying triglyceride

content, de novo lipogenesis, and levels of key lipogenic enzymes. The

involvement of AMPK activation was also evaluated.

Results: α-LA down-regulated triglyceride content by inhibiting FA

esterification and de novo lipogenesis. These effects were mediated by

reduction in FAS, SCD1 and DGAT1 protein levels. Interestingly, α-LA increased

AMPK and ACC phosphorylation, while the presence of the AMPK inhibitor

Compound C reversed the inhibition observed on FAS protein levels.

Conclusions: α-LA down-regulates key lipogenic enzymes, inhibiting

lipogenesis and reducing triglyceride accumulation through the activation of

AMPK signaling pathway in human subcutaneous adipocytes from

overweight/obese subjects.

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INTRODUCTION

Alpha-lipoic acid (5-(1,2-dithiolan-3-yl)-pentanoic acid; α-LA) is a natural

occurring antioxidant compound (Packer et al., 2001) with anti-obesity properties

both in rodents and humans (Carbonelli et al., 2010; Prieto-Hontoria et al.,

2009). Thus, recent clinical trials in overweight/obese humans have

demonstrated that α-LA reduced body weight, fat mass and BMI, which was

accompanied by a decrease in total cholesterol (Kim et al., 2008), inflammatory

markers such as IL-6, TNF-α or CRP (Carbonelli et al., 2010), and haemoglobin-

A1c in obese individuals with type 2 diabetes (Koh et al., 2011).

The antiobesity actions of α-LA have been related to its ability to reduce food

intake and to increase energy expenditure (Kim and Lee, 2005). Furthermore,

several studies have revealed that adipose tissue is a key target in the body

weight lowering and insulin-sensitizing actions of α-LA (Fernandez-Galilea et al.,

2011; Prieto-Hontoria et al., 2009). Thus, in a previous study, our group

demonstrated that α-LA stimulates lipolysis rates in adipocytes, which could

contribute to adiposity reduction (Fernandez-Galilea et al., 2012). Other

mechanism that could also be involved in α-LA fat mass lowering properties is

the modulation of triglyceride accumulation in adipocytes. Traditionally a minor

contribution to whole body lipid stores in human diet-induced obesity has been

attributed to adipose tissue lipogenesis, however, in the setting of positive

energy balance, adipose triglycerides are mainly acquired from circulating

lipoproteins, thus dietary chylomicron triglycerides and endogenously produced

VLDL triglycerides require are hydrolizated by lipoprotein lipase to liberate FFA

which are then acquired by adipocytes further esterified (Garcia-Arcos et al.,

2012; Letexier et al., 2003; Febbraio et al., 2001). However, recent studies have

demonstrated that the inhibition of related lipogenic enzymes could be an

effective strategy against obesity and diabetes conditions, having a signaling

function beyond the generation of lipid stores (Lodhi et al., 2012). In fact, it has

been reported that de novo lipogenesis not only contributes to the rapid recovery

of fat in adipose tissue, but also acts as a glucose sink that allows glycaemia to

be maintained within the range of physiological values (Marcelino et al., 2013).

Thus, mice with targeted deletion of adipose tissue Fas showed increased

energy expenditure and decreased adiposity, which was accompanied by an

enhanced insulin sensitivity and glucose tolerance when fed on a high fat diet

(Lodhi et al., 2012). Although α-LA have revealed important anti-lipogenic

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127

actions in liver in rodents (Chen et al., 2012; Valdecantos et al., 2012), there is

no information available about the effects of α-LA on lipogenesis in human

adipocytes. Thus, the aim of this study was to evaluate the putative effects of α-

LA on triglyceride accumulation and lipogenesis in subcutaneous adipocytes

from overweight/obese subjects, and the potential underlying mechanisms were

also examined.

MATERIALS AND METHODS

Cell culture and differentiation of human subcutaneous preadipocytes

Commercially available cryopreserved human subcutaneous preadipocytes

from non-diabetic overweight-obese female donors (BMI: 26.85-33.37 kg/m2)

were purchased from Zen-Bio Inc. (Research Triangle Park, NC) and

differentiated according to manufacturer’s instructions. Briefly, cryopreserved

preadipocytes were plated in 12 wells plates (Nunc A/S; Roskilde, Denmark) at

40,000 cells/cm2 and cultured in an incubator set up to 37 ºC in a humidified 5%

CO2 atmosphere in preadipocyte medium (PM-1; DMEM/Ham’s F-12 medium,

HEPES, FBS, penicillin, streptomycin, amphotericin B; Zen-Bio). Cells were fed

every other day with 1 ml of PM-1 until confluent. To induce differentiation, PM-1

medium was replaced with 1 ml of differentiation medium (DM2; Zen-Bio)

including biotin, pantothenate, human insulin, dexamethasone,

isobutylmethylxanthine, and a PPARγ agonist (days 0–7). After 7 days, 600 µl of

DM-2 medium were removed and 800 µl of adipocyte medium (AM1; Zen-Bio),

which included PM-1, biotin, pantothenate, human insulin, and dexamethasone

was added. Cells were incubated for additional 2 days, and 800 µl of media were

replaced by 800 µl of fresh AM1. By day 14 of incubation, cells contained large

lipid droplets and were considered mature adipocytes.

Treatments

Before treatments, cell media was removed and replaced with 1 ml of fresh

AM1. α-LA (Sigma; St. Louis, MO) was dissolved in ethanol and the selective

AMPK antagonist Compound C in DMSO. 1000x stocks were prepared and 1

µl/ml of media was added. When Compound C (Calbiochem; San Diego, CA)

(20 µM) was used, cells were pre-incubated during 1 h. Control cells were

treated with the same amount of the corresponding vehicle.

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Triglyceride Measurement

Adipocyte lysates were obtained by the addition of a buffer containing: 2 mM

Tris HCl (pH 8); 137 mM NaCl; 2 mM EDTA; 1% protease inhibitor cocktail 1

(Sigma); 1 mM Sodium orthovanadate and 1 mM PMSF. Triglyceride content

was evaluated after 24 h of treatment by using the Triglycerides Kit (Sigma)

following manufacturer’s instructions.

Fatty acid incorporation to triglycerides

FA incorporation to triglycerides in human subcutaneous adipocytes was

carried out as previously described (Bourlier et al., 2013). Briefly, differentiated

human adipocytes were treated during 24 h with or without α-LA in AM-1, then

cells were washed with PBS and incubated for 4 h in Krebs-Ringer buffer without

glucose, containing 125 mM NaCl, 5 mM KCl, 2 mM CaCl2, 1.25 mM KH2PO4,

1.25 mM MgSO4.7H2O, 25 mM NaHCO3 and 3% FA-free BSA pH 7.8; in

addition, 2 mM L-carnitine, 80 μM palmitic acid (Sigma), and 20 μM 14

C-

palmitatic acid (58 μCi/μmol, Perkin Elmer; Waltham, MA) was added. Then,

cells were washed with PBS and scraped in cold buffer (0.25 M sucrose; 10 mM

Tris HCl; 1 mM EDTA; 1 mM dithiothreitol, pH 7.4). Neutral lipids were extracted

by adding 5 vol chloroform/methanol (2:1) and 0.4 vol 1 M KCl/HCl. Lipids were

separated by thin-layer chromatography to measure labeled palmitate

incorporation into triglycerides using heptane-isopropylether-acetic acid

(60:40:4, v/v/v) as developing solvent. Results were normalized to total protein

content of cell extracts.

Glucose incorporation into triglycerides

For glucose incorporation into triglycerides, cells were incubated for 4 h in

Krebs-Ringer buffer containing 3% BSA, 10 mM HEPES, 2 mM glucose, and 0.5

μCi/ml 14

C-D-glucose (56.3 mCi/mmol; American radiolabeled chemicals, St.

Louis, MO). After 4 h of incubation, cells were washed with cold PBS and then

scraped in cold buffer (0.25 M sucrose; 10 mM Tris HCl; 1 mM EDTA; 1 mM

dithiothreitol, pH 7.4). Neutral lipids were extracted by adding 5 vol

chloroform/methanol (2:1). Lipids were separated by thin-layer chromatography

to measure labeled glucose incorporation into triglycerides using heptane-

isopropylether-acetic acid (60:40:4, v/v/v) as developing solvent. The results

were normalized to total protein content of cell lysates.

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Western blot analysis

Western blot analysis was performed in 14 days post-differentiation

adipocytes. Cells were incubated in AM-1 with or without the appropriate

treatment during 24 h. Lysates were then obtained by the addition of a buffer

containing: 2 mM Tris HCl (pH 8); 137 mM NaCl; 2 mM EDTA; 1% protease

inhibitor cocktail 1 (Sigma); 1 mM Sodium orthovanadate and 1 mM PMSF.

Protein extracts were collected after sample centrifugation. Proteins were

quantified with the BCA method (Pierce-Thermo Scientific, Rockford, IL)

according to the supplier´s instructions. Total proteins were resolved in SDS-

PAGE minigels and electroblotted onto PVDF membranes (GE Healthcare

Europe GmbH, Barcelona, Spain). The membranes were blocked and incubated

with specific antibodies against FAS, AMPK, phospho-Thr172

AMPK, ACC,

phospho-Ser79

ACC (Cell signaling, Beverly, MA); SCD1, DGAT1 (Abcam,

Cambridge, UK); and Actin (Sigma). Secondary antibody was horseradish

peroxidase goat anti-rabbit IgG-HRP (Bio Rad Laboratories, Hercules, CA). The

immunoreactive proteins were detected with enhanced chemiluminescence

(Pierce Biotechnology, Rockford, Illinois, USA). Band intensities were quantified

using a GS-800 calibrated densitometer (Bio Rad Laboratories). In some cases,

infrared fluorescent secondary antibodies (Cell signaling) were used and

quantitated using an Odyssey scanner (LI-COR Biosciences, Lincoln, USA).

Data analysis

Data are expressed as mean with standard errors (SE). Differences were set

up as statistically significant at P<0.05. Comparisons between the values for

different variables were analysed by one-way ANOVA, followed by Bonferroni

post hoc tests, or by Student’s t test or U-Mann Whitney as appropriate after

testing the normality with the Kolmogorov-Smirnoff and Shapiro-Wilk tests.

SPSS 19.0 version for Windows (SPSS, Chicago, IL) and GraphPad Prism 5.0

(Graph-Pad Software INC., San Diego, CA) were used for the statistical

analysis.

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RESULTS

α-LA reduces triglyceride content in human subcutaneous adipocytes

A significant decrease on triglyceride content was found in abdominal

subcutaneous adipocytes from overweight/obese subjects after 24 h of

treatment with α-LA (100 and 250 µM; P<0.05) (Fig. 1).

Fig. 1. α-LA reduces triglyceride accumulation in human subcutaneous adipocytes

from overweight/obese subjects. Mature adipocytes were treated with α-LA (100 and 250 µM)

during 24 h; cells were lysed and intracellular triglyceride content was assayed. Data are

expressed as mean ± S.E. of 6 independent experiments. *p<0.05 vs. Control (vehicle-treated

cells).

α-LA inhibits lipogenesis in human subcutaneous adipocytes

To evaluate if the α-LA triglyceride-lowering effects were caused by an

inhibition of lipogenesis, both FA esterification and glucose incorporation into

triglycerides were evaluated. Twenty-four h of treatment with α-LA (100 and 250

µM) induced an inhibition of 14

C-palmitic acid incorporation into triglycerides

(P<0.05 and P<0.01 respectively; Fig. 2A). Moreover, α-LA also caused a

significant reduction in de novo lipogenesis, was analysed by the 14

C-glucose

incorporation into triglycerides (P<0.05; Fig. 2B).

Control -LA 100 -LA 250 0

50

100

150

**

Tri

gly

ceri

des

(% o

f C

on

tro

l)

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131

Control -LA 100 -LA 250 0

1

2

3

4

*

**

14C

- Pa

lmit

ate

In

co

rpo

rati

on

into

tri

gly

ce

rid

es

(nm

ol/h

/mg

)

Control -LA 100 -LA 250 0

5

10

15

20

* *

14C

-Glu

co

se

in

co

rpo

rati

on

into

tri

gly

ce

rid

es

(nm

ol/h

/mg

)

B

A

Fig. 2. α-LA inhibits lipogenesis in human subcutaneous adipocytes from

overweight/obese subjects. Effects of α-LA on A: FA esterification, evaluated by measuring

labeled palmitate incorporation into triglycerides, and B: De novo lipogenesis assessed after the

fate of [14

C]glucose into triglycerides. Data are expressed as mean ± S.E. of at least 4

independent experiments. *P<0.05 and **P<0.01 vs. Control (vehicle-treated cells).

α-LA represses lipogenic enzymes in human subcutaneous adipocytes

For a better understanding of the mechanisms involved in the inhibition of

triglyceride accumulation induced by α-LA, several key enzymes of the lipogenic

pathway were analyzed. Protein levels of DGAT1, one of the acyltransferases

responsible for the esterification of triglycerides in adipose tissue were

diminished following treatment with α-LA (250 µM, P<0.01) (Fig. 3). Moreover, α-

LA (100 and 250 µM) caused a dose-dependent inhibition of FAS (P<0.01) and

SCD1 protein levels (P<0.01 - P<0.001), the two rate-limiting enzymes in de

novo lipogenesis (Fig. 3).

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FAS DGAT1 SCD10.0

0.5

1.0

1.5

****** **

***

Pro

tein

/Ac

tin

(Arb

itra

ryU

nit

s)

LA 250 µM

LA 100 µM

FAS SCD1DGAT1

ACTIN

- - +

-+-

- - +

-+-

- - +

-+-

Fig. 3. α-LA down-regulates lipogenic enzymes in human subcutaneous adipocytes

from overweight/obese subjects. The effects of α-LA on DGAT1, FAS and SCD1 protein levels

were analyzed after 24 h of α-LA (100 and 250 µM) treatment. Data are expressed as mean ±

S.E. of at least 4 independent experiments. **P<0.01 and ***P<0.001 vs. Control (vehicle-treated

cells)

Activation of AMPK signaling pathway is involved in the anti-lipogenic

actions of α-LA in human subcutaneous adipocytes

AMPK is an enzyme involved in cellular energy homeostasis that switches off

pathways which consume energy such as lipogenesis (Daval et al., 2006).

Because of that, we tested the effects of α-LA on AMPK pathway. The results

revealed that 250 µM α-LA significantly increased (P<0.05) AMPK

phosphorylation (Fig. 4A). One of the first proteins identified as a target of AMPK

is acetyl-CoA carboxylase (ACC), a key enzyme of the lipogenic pathway. In

fact, AMPK phosphorylates and subsequently inhibits ACC (Daval et al., 2006).

In this context, in parallel with the activation of AMPK, α-LA increased ACC

phosphorylation on Ser79

, being significant at 100 and 250 µM (P<0.01; Fig. 4B).

Furthermore, to better characterize the involvement of AMPK activation on the

effects of α-LA on lipogenesis, we assessed the actions of Compound C, a

selective inhibitor of AMPK, on lipogenic enzymes. Thus, AMPK inhibition

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Results (Chapter 4.2)

133

reversed the inhibitory effect of α-LA on FAS protein levels (P<0.05; Fig. 4C).

Although a similar tendency was observed on SCD1 and DGAT1, no statistical

significance was reached (data not shown).

Control -LA 100 -LA 2500

2

4

6

**

**p

AC

C/A

CC

Control -LA 100 -LA 2500

1

2

3

*

pA

MP

K/A

MP

K

P-ACC

ACC

LA 250 µM

LA 100 µM

- - +

-+-

P-AMPK

AMPK

LA 250 µM

LA 100 µM

- - +

-+-

B

A

CC 20 µM

LA 250 µM

- - +

++-

FAS

ACTIN

0.0

0.5

1.0

1.5

**

a

- + +

- - +

-LA (250 M)

CC (20 M)

FA

S/A

ctin

C

Fig. 4. AMPK signaling pathway is involved in α-LA anti-lipogenic effects in human

subcutaneous adipocytes from overweight and obese subjects. A-B: Effects of α-LA (100

and/or 250 µM) treatment during 24 h on (A) AMPK and (B) ACC phosphorylation. C: Effects of

α-LA on FAS protein levels in the presence or absence of the AMPK selective inhibitor

Compound C (CC). Data are expressed as mean ± S.E. of at least 4 independent experiments.

*P<0.05 and **P<0.01 vs. Control (vehicle-treated cells); aP<0.05 vs. α-LA-treated adipocytes.

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134

DISCUSSION

Our data provide novel evidence concerning the ability of α-LA to decrease

triglyceride content in human subcutaneous adipocytes obtained from the

abdominal region of overweight/obese subjects. Adipose tissue triglyceride

content is determined by the balance between triglyceride synthesis

(lipogenesis) and hydrolysis (lipolysis). We have previously reported the ability of

α-LA to promote lipolysis in adipocytes (Fernandez-Galilea et al., 2012), and the

current data demonstrated that α-LA is also able to reduce triglyceride synthesis

in adipocytes from overweight/obese subjects. Lipogenesis occurs either as a

consequence of esterification of FA with glycerol or de novo synthesis of FA.

Concerning FA incorporation into triglycerides, DGAT1 catalyzes the final

acylation step to yield triglycerides. The key role of this enzyme in triglyceride

accumulation in adipose tissue was clearly demonstrated by the fact that

DGAT1-deficient mice have reduced adiposity and are resistant to diet-induced

obesity (Smith et al., 2000). Furthermore, DGAT activity has also been involved

in the regulation of FFA uptake/storage in adipose tissue in humans (Hou et al.,

2009). Therefore, pharmacological inhibition of DGAT1 has been proposed as a

feasible therapeutic strategy for human obesity and type 2 diabetes. Our data

show for the first time the ability of α-LA to decrease DGAT1 protein content in

adipocytes from overweight/obese subjects, suggesting that DGAT1 inhibition

could contribute to the anti-obesity properties of α-LA.

De novo lipogenesis is involved in FA biosynthesis and in the regulation of

the triglyceride storage capacity of adipose tissue (Garrido-Sanchez et al.,

2012). Indeed, it has been suggested that de novo lipogenesis may account for

up to 20% of lipid turnover within adipose tissue (Strawford et al., 2004). Our

present data clearly show that the intracellular lipid-lowering effects of α-LA are

associated with suppressed de novo lipogenesis, which could be triggered in

part, by the inhibition of some of the main enzymes regulating this pathway such

as FAS and SCD1. FAS catalyzes the first committed step in de novo

lipogenesis, and adipose tissue FAS has been implicated in obesity and insulin

resistance in humans (Roberts et al., 2009). Here, we demonstrate that α-LA

diminishes FAS protein level. Targeted deletion of Fas in adipose tissue has

been shown to decrease adiposity in mice, which are resistant to diet-induced

obesity (Lodhi et al., 2012). SCD1 is a key enzyme involved in the control of de

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Results (Chapter 4.2)

135

novo lipogenesis by catalyzing the rate-limiting step in the synthesis of MUFA

(Dobrzyn, 2012). SCD1 deficiency reduces lipogenesis and protects mice from

diet-induced obesity (Kim et al., 2011). We have found that α-LA treatment also

caused a strong inhibition in SCD1 protein level in our model of human

adipocytes. A recent study has shown that the inhibition of SCD1 (induced by

sterculic acid or by conjugated linoleic acid (CLA) reduces de novo lipogenesis

and down-regulates lipogenic genes such as Acc or Fas in primary bovine

adipocytes (Kadegowda et al., 2013). Our data revealed that in addition to FAS

and SCD1 decrease, α-LA also reduced ACC protein levels, a key enzyme of

the lipogenic pathway which mediates the initial step of the FA synthesis. ACC

activity is regulated mainly by phosphorylation, which causes enzyme

inactivation (Kim, 1997). AMPK, a major cellular regulator of lipid metabolism,

has been shown to phosphorylate and inactivate ACC in adipocytes (Peng et al.,

2012). In this way, several of our findings suggest that AMPK mediates the α-LA

inhibitory effects on lipogenic enzymes in human adipocytes from

overweight/obese subjects. Thus, an increase in AMPK phosphorylation was

observed after α-LA treatment, which was accompanied by the subsequent

increase of ACC phosphorylation. In addition, the use of the AMPK antagonist

Compound C reversed the α-LA-mediated down-regulation observed in FAS

protein levels. These results are in agreement with the observations of Chen et

al. (Chen et al., 2012), reporting that FAS inhibition is an important consequence

of AMPK activation mediated by α-LA in C2C12 myotubes.

While in vitro models have limitations, several studies have previously

reported the ability of similar concentrations of α-LA (100-250 µM) to regulate

glucose and lipid metabolism (Moini et al., 2002; Shen et al., 2011; Fernandez-

Galilea et al., 2012), mitochondrial biogenesis (Shen et al., 2011) and adipokine

secretion (Fernandez-Galilea et al., 2011; Prieto-Hontoria et al., 2011.) in murine

adipocytes. Because therapeutic concentrations of α-LA fall within this

micromolar range (Maddux et al., 2001; Carlson et al., 2007), it is possible that

the effects of α-LA in vitro are linked to its therapeutic effect in vivo. In this

context, several studies in rodents have demonstrated that dietary

supplementation with α-LA reduced lipogenesis in liver and muscle in part by

AMPK-dependent pathways (Park et al., 2008; Chen et al., 2012; Valdecantos et

al., 2012).

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136

Taking together, the present study demonstrates the ability of α-LA to down-

regulate key lipogenic enzymes, inhibiting both de novo lipogenesis and FA

esterification and reducing triglyceride accumulation in subcutaneous adipocytes

from overweight/obese subjects through the activation of AMPK signaling

pathway. These data suggest that the inhibition of adipose tissue lipogenesis

could also contribute to the anti-obesity actions of α-LA in humans.

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137

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(AMPK)-dependent and AMPK-independent pathways." Hepatology 48(5):

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Roberts, R., L. Hodson, et al. (2009). "Markers of de novo lipogenesis in

adipose tissue: associations with small adipocytes and insulin sensitivity in

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Shen, W., J. Hao, et al. (2011). "Lipoamide or lipoic acid stimulates

mitochondrial biogenesis in 3T3-L1 adipocytes via the endothelial NO

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162(5): 1213-24.

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Strawford, A., F. Antelo, et al. (2004). "Adipose tissue triglyceride turnover, de

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Valdecantos, M. P., P. Perez-Matute, et al. (2012). "Lipoic acid administration

prevents nonalcoholic steatosis linked to long-term high-fat feeding by

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CCHHAAPPTTEERR 44..33

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Results (Chapter 4.3)

143

α-Lipoic acid treatment increases mitochondrial biogenesis

and promotes beige adipose features in subcutaneous

adipocytes from overweight/obese subjects.

Marta Fernández-Galilea, Patricia Pérez-Matute, Pedro L Prieto-Hontoria;

Marianne Houssier, Marián Burell; J Alfredo Martínez, Dominique Langin, María

J Moreno-Aliaga.

ABSTRACT

Background/Objectives: α-Lipoic acid (α-LA) is a natural occurring

antioxidant with beneficial anti-obesity properties. The aim of this study was to

investigate the putative effects of α-LA on mitochondrial biogenesis and the

acquirement of brown-like characteristics by subcutaneous adipocytes from

overweight/obese subjects and to determine the mechanisms involved.

Methods: Fully differentiated human subcutaneous adipocytes were treated

with α-LA (100 and 250 µM) during 24 h for studies of mitochondrial content and

morphology, mtDNA copy number, FA oxidation enzymes and brown/beige

characteristic genes. The involvement of the SIRT1/PGC-1α pathway was also

evaluated.

Results: α-LA increased mitochondrial content in cultured human adipocytes

as revealed by TEM images and by mitotracker green labeling. Moreover, an

enhancement in mtDNA content was observed. This increase was accompanied

by an up-regulation of SIRT1 protein levels, a decrease in PGC-1α acetylation

and up-regulation of Nrf1 and Tfam transcription factors. Enhanced oxygen

consumption and FA-oxidation enzymes (CPT1 and ACOX) were also observed.

Mitochondria from α-LA-treated adipocytes exhibited some morphological

characteristics of brown mitochondria, and α-LA also induced up-regulation of

some brown/beige adipocytes markers such as Cidea, UCP1 and Tbx1.

Moreover, α-LA up-regulated Prdm16 as well as Fndc5 mRNA levels and irisin

release from adipocytes.

Conclusions: All of these facts suggest the ability of α-LA to promote

mitochondrial biogenesis and brown-like remodelling in cultured white

subcutaneous adipocytes.

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Results (Chapter 4.3)

INTRODUCTION

Mitochondrial dysfunction in adipocytes has been associated with the

development of obesity and type 2 diabetes (Rong et al., 2007). In fact, the

abundance of the mitochondrial copy number in adipocytes from obese is lower

than in those from lean subjects (Mustelin et al., 2008). This lower mitochondrial

content is usually associated with reduced mitochondrial function and, therefore,

decreased fatty acid β-oxidation leading to increased fatty acid accumulation,

which contributes to the development of obesity-associated comorbidities such

as insulin resistance and dislipidemia (Monsenego et al., 2012; Bach et al.,

2013). For this reason, there is a high level of interest in developing therapeutic

strategies aimed to modulate the regulatory pathways that increase

mitochondrial function and biogenesis in an attempt to prevent or treat these

disorders related to mitochondrial dysfunction (Scarpulla et al., 2012). In this

context, mitochondrial biogenesis is a complex process requiring the

coordinated expression and assembly of many proteins encoded by both nuclear

and mitochondrial genomes (Calvo et al., 2006; Scarpulla 2008). Sirtuin 1

(SIRT1)-mediated activation of peroxisome proliferator-activated receptor-

gamma coactivator-1alpha (PGC-1α) is one of the pathways that are particularly

important for mitochondrial biogenesis. Several studies have postulated that

PGC-1α could be a target to prevent and reverse insulin resistance, obesity and

diabetes (Choo et al., 2006; Flachs et al., 2005). This family of regulated

coactivators plays an important role through their interactions with transcription

factors such as nuclear respiratory factor-1 (NRF1), which coordinates the

transcriptional control of nuclear and mitochondrial genomes and directly

activates mitochondrial transcription factor A (TFAM) (Puigserver 2005; Gleyzer

et al., 2005), a core component of the mitochondrial transcription machinery (Shi

et al., 2012). Moreover, PGC-1α is related to the switch from white to brown-like

(brite or beige) inducible adipocytes (Harms and Seale 2013). Thus, Positive

regulatory domain containing 16 (PRDM16), a transcription factor that appears

to control the fate of brown adipose tissue development, binds to PGC-1α and

allows the activation of brown fat-specific genes triggering browning of

adipocytes, which constitutes a novel strategy against obesity (Seale et al.,

2007; Spiegelman, 2013).

Alpha-lipoic acid (5-(1,2-dithiolan-3-yl)-pentanoic acid; α-LA) is an antioxidant

compound (Packer et al., 1995) with demonstrated anti-obesity properties both

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Results (Chapter 4.3)

145

in rodents and humans (Prieto-Hontoria et al., 2009; Carbonelli et al., 2010). In

addition to the body lowering actions of α-LA, beneficial effects on insulin

sensitivity, glucose and lipid metabolism have been described in humans

(Carbonelli et al., 2010; Koh et al., 2011). Several studies have revealed that

white adipose tissue is a target for α-LA therapeutics actions, by regulating key

metabolic pathways such as lipolysis (Fernandez-Galilea et al., 2012), and the

secretion of important adipokines that controls body weight and insulin sensitivity

(Prieto-Hontoria et al., 2011; Prieto-Hontoria et al., 2013; Prieto-Hontoria et al.,

2013; Fernandez-Galilea et al., 2011). Several trials in rodents and murine cells

have suggested that the beneficial actions of α-LA could be also related to its

ability to promote mitochondrial biogenesis in different metabolic tissues such as

liver (Valdecantos et al., 2012). However, there is no information available

regarding the effects of α-LA on mitochondrial biogenesis in human adipocytes

in obesity conditions. Thus, we aimed to evaluate the effects of α-LA on

mitochondrial biogenesis and on the induction of brown-like features in human

subcutaneous adipocytes obtained from overweight/obese subjects. The

molecular mechanisms underlying these effects were also investigated in the

present study.

MATERIAL AND METHODS

Cell culture and differentiation of human subcutaneous preadipocytes

Commercially available cryopreserved human subcutaneous preadipocytes

from non-diabetic overweight-obese female donors (BMI: 26.85-33.37 kg/m2)

were purchased from Zen-Bio Inc. (Research Triangle Park, NC) and

differentiated according to manufacturer’s instructions. Fourteen days after the

induction of differentiation, cells contained large lipid droplets and were

considered mature adipocytes.

Treatments

Before treatment, cell media was removed and replaced with 1 ml of fresh

AM1. α-LA (Sigma; St. Louis, MO) was dissolved in ethanol. 1000x stocks were

prepared and 1 µl/ml of media was added.

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Results (Chapter 4.3)

Transmission Electron Microscopy (TEM)

Mitochondria of differentiated human subcutaneous adipocytes were

examined by TEM. The cultured adipocytes were prefixed during 1 h at 4 ºC in

4% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.2) and postfixed for

2.5 hr at 4 ºC in 1% phosphate buffered osmium tetroxide.After fixation cells

were detached by using a cell scraper and pre-embedded in 2% agarose. Then

samples were embedded in a low viscosity Epoxy resin (SERVA Electrophoresis

GmbH, Heidelberg, Germany) and sectioned to a thickness of 50 nm with Leica

Ultracut R Ultramicrotome (Leica Microsystems GmbH, Wetzlar, Germany).

Finally, Ultrathin sections were double stained in uranyl acetate and lead

hydroxide and examined with an energy filtered transmission electron

microscope (EFTEM) Libra 120 (Zeiss GmbH, Oberkochen, Germany). Images

were acquired by using the software iTEM 5.1 (Olympus Soft Imaging Solutions

GmbH, Münster, Germany).

Analysis of mRNA levels

Total RNA was extracted from fully differentiated human subcutaneous

adipocytes by using TRIzol® reagent (Invitrogen; Carlsbad, CA) according to the

manufacturer’s instructions. RNA concentrations and quality were measured by

Nanodrop Spectrophotometer ND1000 (Thermo Scientific, Wilminton, DE). RNA

was then incubated with the RNase-free kit DNase (Ambion, Austin, TX) for 30

min at 37 ºC. RNA (1 µg) was reverse-transcribed to cDNA using Moloney

Murine Leukaemia Virus MMLV reverse transcriptase (Invitrogen). For the real

time quantitative polymerase chain reaction analysis, 4.5 µl of 1/5, 1/50 or

1/10,000 dilution of cDNA per reaction were used in a final reaction volume of 10

µl.

Tfam, Nrf1, Prdm16, Cidea, Fndc5, Ctp-1 and Acox mRNA levels were

determined using predesigned Taqman® Assays-on-Demand and Taqman

Universal Master Mix (Applied Biosystems, Foster City, CA). The reaction

conditions were followed according to manufacturer’s instructions. Amplification

and detection of specific products were performed using the ABI PRISM 7900HT

Fast System Sequence Detection System (Applied Biosystems).

All mRNA levels were normalized by the housekeeping gene 18s obtained

from Applied Biosystems. Samples were analyzed in duplicate. Ct values (the

cycle where the emitted fluorescence signal is significantly above background

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Results (Chapter 4.3)

147

levels and is inversely proportional to the initial template copy number) were

generated by the ABI software. Finally, the relative expression level of each

gene was calculated as 2-ΔΔCt

(Perez-Matute et al., 2009).

Analysis of mitochondrial DNA content

The amount of mitochondrial DNA (mtDNA) was quantified by real-time PCR

as described previously(Reiling et al., 2010). Briefly, the relative amount of

mtDNA was quantified by comparison of a mitochondrial target, the Mtco2 gene,

with a nuclear target, the 18s gene. Quantitative real-time PCR was performed

using the Applied Biosystems 7900HT (Applied Biosystems) as described

above. For quantification, a ratio between mtDNA (MTCO2) and 18s was

calculated, and used as mtDNA content.

Analysis of mitochondrial content by MitoTracker green staining.

Mitochondria were labelled using the mitochondria-specific dye MitoTracker

Green (Molecular Probes, Life Technologies Ltd, Paisley, UK) according to

manufacturer’s protocol. The final dye concentration was 100 nmol/l and the

incubation time was 30 min prior to visualization. Fluorescent microscopy was

performed on living cells with a Leica DM IL-EL 6000 (Leica Microsystems

GmbH) inverted microscope. For fluorescence intensity quantification a Polarstar

Galaxy fluorimeter (BMG labtech) set up to 554 nm excitation and 576 nm

emission wavelengths was used.

Western blot analysis

Western blot analyses were performed in 14 days post-differentiation

adipocytes. Cells were incubated in serum-free DMEM overnight and then with

or without the appropriate treatment. Lysates were obtained by the addition of a

buffer containing: 2 mM Tris HCl (pH 8); 137 mM NaCl; 2 mM EDTA; 1%

protease inhibitor cocktail 1 (Sigma); 1 mM Sodium orthovanadate and 1 mM

PMSF. Protein extracts were collected after sample centrifugation and quantified

with the BCA method (Pierce-Thermo Scientific, Rockford, IL, USA) according to

the supplier´s instructions. Total proteins were resolved in SDS-PAGE minigels

and electroblotted onto PVDF membranes (GE Healthcare Europe GmbH,

Barcelona, Spain). The membranes were blocked and incubated with specific

antibodies against SIRT1, CPT-1 (cell signaling, Beverly, MA); ACOX (Abcam,

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Results (Chapter 4.3)

Cambridge, UK); CD36 (Santa Cruz biotechnology, Dallas, TX) and Actin

(Sigma). Secondary antibody was horseradish peroxidase goat anti-rabbit IgG-

HRP (Bio Rad Laboratories, Hercules, CA), detected with enhanced

chemiluminescence (Pierce Biotechnology, Rockford, Illinois, USA). Band

intensities were quantified using a GS-800 calibrated densitometer (Bio Rad

Laboratories). In some cases, infrared fluorescent secondary antibodies (Cell

signaling) were used and quantitated using an Odyssey® Sa infrared imaging

system (LI-COR Biosciences, Lincoln, USA).

Immunoprecipitation

For acetylation analysis, 2 μg of anti-PGC-1α (Santa Cruz Biotechnology)

antibody was added to 200 µg of protein extracts (1 μg/μl) and incubated for 2 h

at 4 ºC. After the addition of 20 μl of protein A/G PLUS-Agarose (Santa Cruz

Biotechnology), incubation at 4°C with shaking was carried out overnight. The

A/G PLUSAgarose-beads were pelleted by centrifugation at 12,000 g for 1 min

at 4°C and washed four times with PBS at 4°C to remove non-adsorbed

proteins. After the final wash, protein was released from the beads by treatment

at 95°C for 7 min in 2x sample buffer (Invitrogen), resolved on 12% SDS-PAGE

gels and transferred to PVDF membranes. The membranes were blocked,

incubated with specific antibodies against anti-acetylated Lys (Cell Signalling)

and anti-PGC-1α (Santa Cruz Biotechnology) and detected as described above.

Oxygen consumption analysis

Oxygen consumption in fully differentiated human subcutaneous adipocytes

was carried out by the use of the BD™ Oxygen Biosensor System (BD

biosciences; San Jose, CA). Thus, adipocytes were detached from the plate by

using a tripsine solution (Invitrogen) and placed in the microplate provided.

Oxygen consumption was determined in a Polarstar Galaxy fluorimeter (BMG

labtech; Ortenberg, Germany) every 3 min cycle. Optical Parameters were set

up at excitation 485 nm and emission 630 nm wavelenght.

Fatty acid metabolism measurements

Triglycerides-derived FA oxidation was measured by the sum of 14

CO2 and

14C-ASM (acid soluble metabolites) as previously described (Bourlier et al.,

2013). Briefly, differentiated human adipocytes were incubated for 4 h in Krebs-

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Results (Chapter 4.3)

149

Ringer buffer without glucose, containing 125 mM NaCl, 5 mM KCl, 2 mM CaCl2,

1.25 mM KH2PO4, 1.25 mM MgSO4.7H2O, 25 mM NaHCO3 and 3% fatty acid-

free BSA pH 7.8, 2 mM L-carnitine, 80 μM palmitic acid (Sigma), and 20 μM 14

C-

palmitic acid (58 μCi/μmol, Perkin Elmer; Waltham, MA). Medium was

transferred to a glass vial with a central well containing benzethonium hydroxide

(Sigma). 14

CO2 was liberated by acidification with 1 M H2SO4 and collected

during 2 h in the central well. 14

CO2 was measured by scintillation counting in a

scintillation counter Wallac 1409 (EG&G Company, Turku, Finland). Cells were

washed and then scraped in cold buffer (0.25 M sucrose; 10 mM Tris HCl; 1 mM

EDTA; 1 mM dithiothreitol, pH 7.4). Neutral lipids and ASM were separated by

adding 5 vol chloroform/methanol (2:1) and 0.4 vol 1 M KCl/HCl. Specific activity

was measured and used to calculate total oxidation as equivalent of oxidized

palmitic acid. Results were normalized to total protein content of cell extracts.

For free fatty acids uptake cells were incubated in Kreb-Ringer buffer as

described above during 50 min and after this period of time, cold palmitic acid,

carnitine and the non metabolizable analogue 2Bromo-palmitic acid (Moraveck

Biochemicals, Brea, CA) were added to the media in a final concentration of 2

mM L-carnitine, 80 μM palmitic acid (Sigma), and 20 μM 14

C-2Bromo-palmitic

acid and incubated during 10 min. Culture plates were then put on ice and rinsed

twice with cold PBS. Cells were scraped in 0.05 M NaOH, and 14

C-2Bromo-

palmitic acid uptake was measured by liquid scintillation counting of cell lysate.

The results were normalized to total protein content of cell lysates.

Immunofluorescence

For uncoupling protein 1 (UCP1) immunofluorescence evaluation, after

treatment (α-LA 250 µM or vehicle) the adipocytes were washed twice in PBS

and fixed in 4% paraformaldehyde in PBS pH 7.4 for 15 min at room

temperature. Then cells were permeabilized by incubating the samples for 10

min with PBS containing 0.25% Triton X-100 and unspecific binding was blocked

by incubating with 1% BSA in PBST for 30 min. After that cells were incubated

with UCP1 antibody (Abcam) in 1% BSA in PBST for 1 h at room temperature

and detected with the fluorescent secondary antibody Alexa fluor® 647

(Invitrogen). Finally, cells were incubated with Dapi during 1 min for nuclei

staining and observed with a Leica DM IL-EL 6000 (Leica Microsystems GmbH)

inverted microscope.

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Results (Chapter 4.3)

Data analysis

Data are expressed as mean ± standard errors (SE). Differences were set up

as statistically significant at P<0.05. Comparisons between the values for

different variables were analyzed by one-way ANOVA, followed by Bonferroni

post hoc tests, or by Student’s t test or U-Mann Whitney once the normality with

the Kolmogorov-Smirnoff and Shapiro-Wilk tests was screened. SPSS 19.0

version for Windows (SPSS, Chicago, IL, USA) and GraphPad Prism 5.0

(Graph-Pad Software INC. San Diego, CA, USA) were used for the statistical

analysis.

RESULTS

Effects of α-LA on mitochondrial biogenesis in adipocytes from

overweight/obese subjects

In order to evaluate the putative effects of α-LA on mitochondria, several

parameters were evaluated. TEM images of mature adipocytes treated during 24

h with α-LA (250 µM) showed an increase in the number of mitochondria when

compared with vehicle treated cells (Fig. 1).

Fig. 1. Representative TEM images of mature subcutaneous adipocytes from

overweight/obese subjects treated with α-LA (250 µM) or vehicle (Control cells) during 24

h. A-B: TEM images of Control adipocytes at (A) x1600 and (B) x8000, showing low mitochondrial

content. C-D: TEM images of α-LA-treated adipocytes at (C) x1600 and (D) x8000, exhibiting

increased mitochondrial mass.

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Results (Chapter 4.3)

151

Moreover, we ensured the effects of α-LA on mitochondria by using

mitotracker green probes, which stains mitochondrial matrix proteins

independently of the mitochondrial membrane potential and mainly reflects the

mitochondrial mass. Fluorescence microscopy observations and fluorimetric

measurements showed that mitochondrial mass was increased (P<0.05) after 24

h of α-LA (250 µM) treatment (Fig. 2A). In agreement with this, the analysis of

mtDNA content suggested that α-LA produced an increase in mitochondrial DNA

copy number (P<0.05) at the highest concentration (250 µM) assayed (Fig. 2B).

All these data suggest the ability of α-LA to up-regulate mitochondrial biogenesis

in subcutaneous human adipocytes.

0 100 250 0

20

40

60

80*

-LA (M)

Mit

otr

acker

gre

en

(RF

U/p

rote

in)x

10

3

Control LA 250 µM

0 100 250 0

1

2

3

4**

-LA (M)

Rela

tive m

tDN

A level

(Arb

itra

ry U

nits)

A)

B)

Fig. 2. Effects of α-LA treatment on mitochondrial content and mitochondrial DNA

(mtDNA) quantity in mature subcutaneous adipocytes from overweight/obese subjects. A:

Top panel: Live cell imaging of mitochondria with MitoTracker Green. Bottom panel: Fluorescence

intensity quantification after MitoTracker labeling shows that α-LA-treated adipocytes (250 µM for

24 h) contain more mitochondria. B: α-LA enhances mitochondrial DNA (mtDNA) quantity,

calculated by comparison of the levels of subunit 2 of COX (MTCO2), an mtDNA-encoded

mitochondrial protein and the levels of nuclear 18s gene determined by real-time PCR. Results

were normalized by the mean value for the Control cells set to 1.0 unit. Data are means ± S.E.M.

of at least 4 independent experiments. *P <0.05 and **P<0.01 vs. Control (vehicle-treated cells).

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Results (Chapter 4.3)

Effects of α-LA on SIRT1 and PGC-1α in adipocytes from

overweight/obese subjects.

Because SIRT1 and PGC-1α are considered master regulator of

mitochondrial biogenesis (Philp and Schenk 2013), the effects of α-LA on

SIRT1/PGC-1α pathway were tested. Interestingly, an increase on both Sirt1

mRNA and SIRT protein levels (P<0.05) was observed in 250 µM α-LA-treated

adipocytes (Fig. 3A). It is well known that SIRT1 activates PGC-1α by

deacetylation (Lagouge et al., 2006). In this context, our data revealed that the

increase in SIRT1 deacetilase activity induced by α-LA was reflected in a

decreased PGC-1α acetylation levels (P<0.05) (Fig. 3B). PGC-1α integrates and

coordinates the activity of multiple transcription factors that control mitochondrial

biogenesis such as NRF-1, which stimulates the synthesis of TFAM, a final

effector activating the duplication of mitochondrial DNA molecules (Vina et al.,

2009). Interestingly, a significant increase in both Nrf1 (P<0.05) and Tfam

(P<0.05) mRNA levels was observed in adipocytes after 24 h of treatment with

α-LA at the concentration of 250 µM (Fig.3C).

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Results (Chapter 4.3)

153

0 100 2500.0

0.5

1.0

1.5

**

-LA (M)P

GC

-1

ac

ety

lati

on

0 100 2500.0

0.5

1.0

1.5

2.0 *

-LA (M)

Sir

t1 m

RN

A(A

rbit

rary

Un

its)

0 100 2500.0

0.5

1.0

1.5

2.0

*

-LA (M)

SIR

T1/A

cti

n

SIRT1

Actin

α-LA 250 µM

α-LA 100 µM

- - +

-+-

WB: Acetyl-Lys

IP: PGC-1α

WB: PGC1α

α-LA 250 µM

α-LA 100 µM

- - +

-+-

B)

A)

NRF1 TFAM0.0

0.5

1.0

1.5

2.0 **

Control

-LA 100 M

-LA 250 M

mR

NA

exp

ressio

n

(Arb

itra

ry U

nits)

C)

Fig. 3. Effects of α-LA treatment on SIRT1/PGC-1α signaling pathway and

mitochondrial biogenesis related genes in subcutaneous adipocytes from

overweight/obese subjects. Fully differentiated adipocytes were incubated with α-LA (100 and

250 µM) for 24 h. A: Up-regulation of Sirt1 mRNA and protein levels after α-LA treatment. B:

PGC-1α deacetylation is increased in α-LA-treated adipocytes. C: Up-regulation of PGC-1α-

related nuclear factors Nrf1 and Tfam mRNA levels after incubation with α-LA. Data are means ±

S.E.M. of at least 4 independent experiments. *P <0.05 vs. Control (vehicle-treated cells).

Effects of α-LA on oxygen consumption and fatty acid oxidation in

adipocytes from overweight/obese subjects.

We next evaluated whether increased mitochondrial biogenesis was

accompanied by changes in oxygen consumption. Interestingly, a significant

increase in oxygen consumption was found in adipocytes treated with α-LA (100

and 250 µM; P<0.01) in accordance to the increase observed in mitochondrial

mass (Fig. 4A).

The effects of α-LA on CPT-1 and ACOX, two rate limiting enzymes involved

in fatty acid oxidation were also tested. Our data revealed that α-LA (100 and

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Results (Chapter 4.3)

250 µM) increased both CPT-1 (P<0.05 and P<0.001 respectively) and ACOX

(P<0.05) gene expression and protein levels (Fig. 4B). Paradoxically, no effects

on 14

C-palmitic acid oxidation were found at any of the concentrations used (Fig.

4C).

0 100 2500

2

4

6

8

10 ****

-LA (M)

Oxyg

en

Co

nsu

mp

tio

n

(AU

C)x

10

3

Cpt1 Acox0

1

2

3

*

***

* *

-LA 100 M

Contol

-LA 250 M

mR

NA

exp

ressio

n(A

rbit

rary

Un

its)

A)

CTP1 ACOX0

1

2

3

4

5

***

* *

Pro

tein

/Acti

n(A

rbit

rary

Un

its)

CPT1 ACOX

ACTIN

LA 250 µM

LA 100 µM

- - +

-+-

- - +

-+-

B)

C)

0 100 2500.0

0.1

0.2

0.3

0.4

-LA (M)

Pa

lmit

ate

oxid

ati

on

(nm

ol/m

g/h

)

Fig. 4. Effects of α-LA on oxygen consumption and fatty acid oxidation in

subcutaneous adipocytes from overweight/obese subjects. Mature adipocytes were

incubated with α-LA (100 and 250 µM) for 24 h. A: Oxygen consumption, measured in a BD

Oxygen Biosensor System plate, is increased in α-LA-treated adipocytes. Data are presented as

area under curve (AUC) of RFU (relative fluorescent units). B: Cpt-1 and Acox mRNA (top panel)

and protein (bottom panel) levels increases after α-LA treatment. C: Palmitate oxidation rates in

control and α-LA-treated adipocytes. Data are means ± S.E.M. of at least 4 independent

experiments. *P <0.05 **P<0.01 and ***P<0.001 vs. Control (vehicle-treated cells)

This apparent controversial finding led us to test if α-LA was affecting

exogenous FFA uptake by adipocytes. For this purpose, the uptake of 14

C-2-

bromopalmitic acid, a non metabolizable analogue of palmitic acid, was

evaluated. Thus, an inhibition of 14

C-2-bromopalmitic acid uptake was observed

after 24 h of treatment with α-LA (100 and 250 µM; P<0.05 and P<0.01

respectively) (Fig. 5A). Moreover, a decrease in the fatty acid translocase CD36

protein content was also observed after α-LA treatment (100 and 250 µM,

P<0.05) (Fig. 5B).

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Results (Chapter 4.3)

155

0 100 2500.0

0.5

1.0

1.5

***

-LA (M)

2-B

rom

op

alm

itic

acid

Up

take (

nm

ol/m

g/m

in)

0 100 2500.0

0.5

1.0

1.5

**

-LA (M)

CD

36/A

cti

n

A)

B)

CD36

ACTIN

α-LA 250 µM

α-LA 100 µM

- - +

-+-

Fig. 5. Effects of α-LA on fatty acids uptake in subcutaneous adipocytes from

overweight/obese subjects. Fully differentiated adipocytes were incubated with α-LA (100 and

250 µM) for 24 h. A: 2-Bromopalmitate uptake decreases after α-LA treatment. B: Down-

regulation of the fatty acid traslocase CD36 in α-LA-treated adipocytes. Data are means ± S.E.M.

of at least 4 independent experiments. *P <0.05 and **P<0.001 vs. Control (vehicle-treated cells).

Effects of α-LA on the induction of brown-like features in subcutaneous

adipocytes from overweight/obese subjects.

As the increase in mitochondrial mass and function is a hallmark of white to

brown switch in adipocytes (Barneda et al., 2013), we next evaluated if α-LA was

able to induce other brown-like features in white adipocytes. In this context, TEM

images demonstrated that α-LA treatment (250 µM) produced remarkable

differences on mitochondrial morphology, adipocytes treated with α-LA showing

brown-like mitochondria (Fig. 6A). To further investigate the ability of α-LA to

promote browning features in subcutaneous adipocytes, we next evaluated the

effects of α-LA on brown and beige adipocytes specific markers. The gene

expression analysis carried out revealed that α-LA treatment (250 µM) increased

(P<0.01) the mRNA levels of Prdm16, Cidea and Fndc5 (Fig. 6B). Moreover, the

mRNA levels of Tbx1, a beige adipocyte specific marker was also significantly

up-regulated (P<0.05) in α-LA-treated cells at both concentrations assayed (100

and 250 µM) (Fig. 6B).

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Results (Chapter 4.3)

PRDM16 CIDEA FNDC5 TBX10

1

2

3 ** ** ** Control

-LA 100 M

-LA 250 M* *

mR

NA

exp

res

sio

n(A

rbit

rary

Un

its)

B)

Control α-LA 250 µMA)

Fig. 6. Evaluation of the acquisition of beige adipocytes features after α-LA treatment

in subcutaneous white adipocytes from overweight/obese subjects. Differentiated

adipocytes were incubated with α-LA (100 and 250 µM) for 24 h. A: Representative TEM images

showing differenceses in mitochondrial morphology between control (left panels) and α-LA-

treated (right panels) adipocytes. B: Up-regulation of brown/beige adipocytes-related genes

Prdm16, Cidea, Fndc5 and Tbx1 mRNA levels in α-LA-treated adipocytes. Data are means ±

S.E.M. of at least 4 independent experiments. *P <0.05 and **P<0.001 vs. Control (vehicle-

treated cells).

Furthermore, the production of irisin, the protein encoded by Fndc5 gene,

was also increased (P<0.001) after 24 h of incubation with α-LA (250 µM)

(Supplemental Fig. I).

0 100 2500

2

4

6

8

10**

-LA (M)

Iris

in (

g/m

l)

Supplemental Fig I. α-LA increases the amount of irisin released into media in

subcutaneous adipocytes from overweight/obese subjects. Mature adipocytes were treated with

α-LA (100 and 250 µM) for 24 h. Data are means ± S.E.M. of at least 4 independent experiments.

**P<0.001 vs. Control (vehicle-treated cells).

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Results (Chapter 4.3)

157

Control

α-LA (250 µM)

UCP1 DAPI MERGE

FIGURE 7

A)

B)

C)

D)

E)

F)

In addition, immunofluorescence analysis revealed an induction of UCP1 in

α-LA (250 µM) treated adipocytes (Fig. 7).

Fig. 7. Effects of α-LA on UCP1 expression in subcutaneous adipocytes from

overweight/obese subjects. Differentiated adipocytes were incubated with α-LA (250 µM) or

vehicle for 24 h. A-B: UCP1 immunofluorescence-staining. C-D: Dapi nuclei staining. E-F: merged

images.

DISCUSSION

Mitochondrial number and oxygen consumption are reduced in genetic and

high fat-induced obesity (Rong et al., 2007) and this mitochondrial dysfunction in

mature adipocytes has been linked to defects in fatty acid oxidation (Gao et al.,

2010), secretion of adipokines (Koh et al., 2007), and dysregulation of glucose

homeostasis (Sutherland et al., 2008). Previous studies in murine 3T3-L1

adipocytes showed that α-LA treatment, both alone (Shen et al., 2011) or in

combination with acetyl-L-carnitine (Shen et al., 2008) promoted mitochondrial

biogenesis. Importantly, our current study demonstrates that α-LA is also able to

increase mitochondrial biogenesis in adipocytes in overweight/obese condition in

humans, as revealed by the higher mitochondrial number per cell observed with

TEM imaging, mitotracker green staining and the raise in mitochondrial DNA

content. These effects seem to be secondary to α-LA-mediated up-regulation of

Nrf1 and Tfam levels caused, at least in part, by SIRT1-induced deacetilation,

and therefore activation, of PGC-1α. In this context, it is known that SIRT1

activation improves mitochondrial function and exerts some protective effects

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Results (Chapter 4.3)

against diet-induced obesity by deacetilating PGC-1α which is a master

regulator of mitochondrial biogenesis (Seale et al., 2007). Thus, PGC-1α

integrates and coordinates the activity of multiple transcription factors, such as

NRF1 and 2, which control the transcription of key mitochondrial proteins

including mitochondrial transcriptional factor, as TFAM that is essential for

mitochondrial replication and transcription (Santos and Kowluru 2011). In

support of our study, previous trials have observed that SIRT1 is a target for α-

LA. In fact, it was described that α-LA increases mitochondrial copy number in

the liver of rats fed on a high fat diet and it also improves mitochondrial function

in C2C12 myotubes by increasing SIRT1 deacetylase (Valdecantos et al., 2012;

Chen et al., 2012).

The current data also suggest that α-LA not only increase mitochondrial

content, but also mitochondrial function. In fact, an increase in oxygen

consumption and in CPT-1, which is the key regulatory enzyme of mitochondrial

long-chain fatty acid oxidation, was observed. Moreover, α-LA treatment also

increased ACOX, which catalyzes the initial and rate-determining step of the

peroxisomal fatty acid β-oxidation pathway. These facts are consistent with an

increase in fatty acid oxidation machinery in α-LA-treated adipocytes. However,

surprisingly, the oxidation rate of exogenous palmitate was not affected after 24

h of α-LA treatment. Other studies carried out in murine 3T3-L1 adipocytes also

revealed a lack of effect of α-LA (10 µM) on palmitate oxidation and only a

moderate increase in palmitate oxidation to acid-soluble metabolites was

observed at high concentrations (250 µM) (Shen et al., 2008; Fernandez-Galilea

et al., 2012). Due to the strong up-regulation observed in key rate-limiting

enzymes involved in fatty acid oxidation, the lack of effects found when

measuring palmitate oxidation could suggest interferences in exogenous fatty

acid entry into the adipocyte. In fact, the measurement of 14

C-2-bromopalmitic

acid, a non metabolizable analogue of palmitic acid, uptake evidenced an α-LA-

mediated inhibition of fatty acids transport, in parallel with a reduction of the

levels of FAT/CD36, a translocase involved in fatty acid uptake, binding and

transport(Yang et al., 2007). A recent study of our group have demonstrated that

α-LA is a potent lipolytic agent (Fernandez-Galilea et al., 2012), and therefore, α-

LA-treated adipocytes have increased amount of endogenous of free fatty acids

to be oxidized, which is in accordance with the up-regulation of the main

enzymes involved in this process.

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159

Moreover, the inhibition of exogenous fatty acids uptake observed in α-LA-

treated cells could also be an adaptive mechanism to counteract the overload in

endogenous free fatty acids as a consequence of increased lipolysis. Taking

together, we further hypothesize that the absence of effects observed on

exogenous palmitate oxidation could be the result of an inhibition of exogenous

fatty acids uptake, and, therefore, we cannot rule out the ability of α-LA to

increase fatty acid oxidation based only on this observation. In this way, several

studies have demonstrated that α-LA increases fatty acid oxidation in non-

adipose cells both in vitro and in vivo in rodents. Thus, in C2C12 myotubes, α-LA

produced an increase in palmitate oxidation using a range of concentrations (30-

1,000 µM) similar to the dose used in the present study (Chen et al., 2012).

Moreover, dietary administration of α-LA increased in liver of high fat-fed mice

and rats, the expression of genes related to beta-oxidation, such as acyl-

coenzyme A dehydrogenase, Cpt-1 and Acox (Yang et al., 2008; Valdecantos et

al., 2012; Kim and Miura, 2004).

Our TEM studies revealed that α-LA not only increase mitochondrial mass

but also induced changes in mitochondrial morphology. In fact, mitochondria

from α-LA-treated adipocytes exhibit some morphological characteristics of

brown mitochondria, being larger and with numerous transverse cristae

(Barbatelli et al., 2010) in comparison with untreated adipocytes. Moreover,

Cidea which is considered a brown adipocyte marker, was up-regulated in α-LA-

treated adipocytes (Barneda et al., 2013). Taking together, the increase in

mitochondrial biogenesis, fatty acid oxidation machinery and the changes

observed in the mitochondrial morphology suggest that α-LA might induce a

brown-like phenotype within white subcutaneous adipocytes. Recently, it has

been established that inducible brown adipocytes (also called beige, brown-in-

white, or brite adipocytes), are phenotypically distinct from both white and brown

adipocytes. In this context, TBX1 has been identified as a reliable marker for

beige adipocytes in humans (Jespersen et al., 2013). Our data showed that α-LA

treatment increased Tbx1 mRNA levels suggesting that α-LA might promote

transdifferentiation of mature white adipocytes into brite adipocytes. Moreover, it

has been described that human beige adipocytes initially expressed low levels of

UCP1, but expression of UCP1 can be induced after mimicking cold exposure or

by different treatments such as β-3 adrenergic and PPARγ agonists (Bartelt and

Heeren 2013; Ohno et al., 2012). Interestingly, our results demonstrated that α-

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Results (Chapter 4.3)

LA is also able to induce UCP1 in subcutaneous adipocytes of overweight/obese

subjects. Browning of WAT can be brought about by transcriptional modulation

through PRDM16 (Seale et al., 2011) and PGC-1α (Tiraby et al., 2003). In fact,

in primary human subcutaneous white fat, adenovirus-mediated expression of

PGC-1α was described to lead to a brown-fat phenotype and increased

respiratory chain proteins and fatty acid oxidation enzymes (Tiraby et al., 2003).

Transgenic expression of PRDM16 in fat tissue also induces the brown-like cells

in subcutaneous WAT(Seale et al., 2011), and PRDM16 has been described as

a required and sufficient molecule to promote brown features in white adipose

tissue (Seale et al., 2007; Seale et al., 2008). PRDM16 directly binds to PGC-1α

allowing the activation of other brown fat-specific genes (Seale et al., 2007). In

this context, our data have revealed that the up-regulation of Prdm16 observed

after α-LA treatment is accompanied by the activation of PGC-1α, which might

promote the white to brite transdifferentiation.

Recently, it has been shown that PGC-1α stimulates the expression of

Fndc5, which encodes for irisin, a polypeptide hormone cleaved and released by

muscle (Bostrom et al., 2012) and adipose tissue (Moreno-Navarrete et al.,

2013). Irisin induces the browning of subcutaneous white adipose tissue

(Bostrom et al., 2012) in mice. In this line, our data revealed that α-LA up-

regulates Fndc5 mRNA levels and also irisin release from adipocytes, which

could also contribute to the browning properties of α-LA. However, it is important

to mention that a recent study has jeopardized the role of Fndc5 and irisin as

inductor of brite adipocytes differentiation in humans (Raschke et al., 2013).

In summary, the present data revealed the ability of α-LA to induce a

remodelling of white subcutaneous adipocytes from overweight/obese subjects,

characterized by increased mitochondrial biogenesis and fatty acid oxidation

enzymes, and accompanied by the acquirement of beige adipocytes features, in

part mediated through SIRT1/PGC-1α pathway and by the induction of irisin. All

of these facts suggest that the brown-like remodelling induced by α-LA in white

adipocytes might also contribute to the anti-obesity properties of α-LA and could

help to improve metabolic health.

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161

ACKNOWLEDGEMENTS

This study was supported by a grant from Ministerio de Ciencia e Innovación

of Spain (AGL 2009-10873/ALI) and by Línea Especial de Investigación

“Nutrición, Obesidad y Salud”, University of Navarra-Spain LE/97. M.

Fernández-Galilea was supported by a predoctoral grant from Navarra

Goverment. Authors thank María Zabala-Navó for excellent technical assistance.

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Results (Chapter 4.3)

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CCHHAAPPTTEERR 55

GGEENNEERRAALL DDIISSCCUUSSSSIIOONN//SSUUMMMMAARRYY

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General discussion/Summary

171

GENERAL DISCUSSION/SUMMARY

Human white adipose tissue plays a pivotal role in maintaining whole-body

energy homeostasis by storing triglycerides when energy surplus, releasing free

FA as fuel during energy shortage, and secreting adipokines that are important

for regulating lipid and glucose metabolism (Feng et al., 2013). Nowadays, the

role of adipose tissue as a key regulator of whole-body lipid and glucose

homeostasis is well established based on extensive experimental evidence.

Thus, dysfunctions in adipose tissue metabolism have a direct impact on lipid

and glucose homeostasis. Indeed, the combination of hyperphagia and adipose

dysfunction seems to underline important metabolic pathologies such as insulin

resistance, type 2 diabetes and cardiovascular diseases (Guilherme et al.,

2008). In this context, several studies have suggested that the development of

obesity and its complications is related to modifications in lipid turnover in white

adipose tissue (Klop et al., 2013). In fact, the hypertrophy of adipocytes as a

consequence of an excess in triglyceride storage is associated with the

development of intracellular abnormalities of adipocyte function, including

alteration of the production of adipokines, and mitochondrial function among

others. Finally, these metabolic dysfuntions of adipose tissue in obesity could

contribute to later impairment of insulin action. In this context, it is well known

that obese patients with smaller-size adipocytes exhibit better lipid profiles and

insulin sensitivity, while those with adipocyte hypertrophy in subcutaneous

adipose tissue exhibit more adverse metabolic profiles (Farnier et al., 2003;

Lafontan, 2013). Moreover, the ability of hyperthophic adipocytes to function as

endocrine cells and secrete important adipokines involved in metabolic

regulation is altered and this could also contribute to the development of the

metabolic disorders associated to obesity (Prieto-Hontoria et al., 2011).

Therefore, it is important to find strategies to control adipocyte size through the

balance between triglycerides storage and break-down in an attempt to prevent

or to treat obesity and its associated co-morbidities (Ahmadian et al., 2009;

Lodhi et al., 2012).

Alpha-lipoic acid (α-LA) is a natural occurring antioxidant compound with

anti-obesity properties in both rodents and humans (Prieto-Hontoria et al., 2009;

Carbonelli et al., 2010). The anti-obesity actions of α-LA have been related to its

ability to reduce food intake and to increase energy expenditure (Kim et al.,

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General discussion/Summary

172

2004). In addition, adipose tissue has emerged as a key target in the body

weight lowering and insulin-sensitizing actions of α-LA. In fact, supplementation

with α-LA is able to regulate the adipose tissue inflammation associated with

obesity and the secretion of key adipokines involved in the regulation of body

weight and energy metabolism (Prieto-Hontoria et al., 2009; Deiuliis et al., 2011;

Prieto-Hontoria et al., 2011; Pashaj et al., 2013; Prieto-Hontoria et al., 2013).

The present study demonstrates that α-LA is also able to regulate the main

metabolic pathways involved in adipocyte triglycerides/FFA metabolism, which

controls adipocyte lipid content. In this context, one of the main outcomes of the

present study was the observation that α-LA stimulates lipolysis by the

regulation of some of the main lipases and lipid droplet proteins involved in the

control of this pathway in adipocytes. Thus, our data suggest that both HSL and

perilipin are targets of α-LA. Moreover, a key role of PKA activation in the

lipolytic actions of α-LA was evidenced based on the increase in PKA-mediated

phosphorylation of HSL at Ser563

and Ser660

, as well as, on PKA-induced

phosphorylation of perilipin in α-LA-treated adipocytes. In addition, the PKA

inhibitor H89 completely blunted the lipolytic action of α-LA and the α-LA-

induced phosphorylation of PKA substrates, perilipin and HSL. However, the

present data suggest that ATGL activation is not a determinant factor involved in

α-LA-mediated lipolytic actions as concluded from the findings that no significant

changes were observed neither on the levels of the ATGL co-activator protein

called CGI-58 or the inhibitory protein G0S2.

Few years ago, a new adipocyte phospholipase A2 called AdPLA was

identified and functionally characterized (Duncan et al., 2008). Afterwards, it was

demonstrated that AdPLA ablation increased lipolysis by reducing PGE2 levels

and thereby stimulating cAMP and phosphorylation of HSL through cAMP-

mediated activation of PKA (Jaworski et al., 2009). Our results demonstrated for

the first time that α-LA treatment down-regulated AdPLA as well as PGE2 levels

accompanied by an increase in cAMP levels, which could also contribute to the

increased PKA-mediated phosphorylation of perilipin and HSL at Ser563

and

Ser660

and, thereby, contribute to the lipolytic effects of α-LA.

PI3K/AKT pathway has also been involved in the regulation of lipolysis. In

fact, PI3K/AKT activation increases PDE3B activity, and hydrolysis of cAMP

leading to a net dephosphorylation of HSL and inhibition of PKA-mediated

lipolysis (Ridderstrale, 2005). In our experimental cell model, α-LA inhibits AKT

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173

phosphorylation and both the PI3K/AKT inhibitor LY294002 and the PDE3B

antagonist Cilostamide potentiated the stimulatory effects of α-LA on basal

lipolysis. Therefore, the present results suggest that the lipolytic effects of α-LA

could also be mediated by decreasing AKT activation, which might increase

cAMP and lipolysis mediated by HSL and perilipin activation.

AMPK has been also involved in the regulation of lipolysis (Hardie, 2008;

McGee and Hargreaves, 2010). Thus, it has been reported that phosphorylation

of HSL at Ser565

by AMPK prevents activation by PKA, inhibiting lipolysis (Dagon

et al., 2006; Boon et al., 2008; Anthony et al., 2009). However, the role of AMPK

activation in the regulation of lipolysis is complex, since it has been recently

reported that ATGL is phosphorylated and activated by AMPK to increase

lipolysis (Ahmadian et al., 2011). Thus, the effects described for AMPK

activators on lipolysis are controversial, showing inhibition (Bourron et al., 2010;

Lorente-Cebrian et al., 2012) and activation of lipolysis (Gaidhu et al., 2009;

Ahmadian et al., 2011). In addition, it has been suggested that the effects of

AMPK activation on lipolysis might be time-dependent (Yin et al., 2003).

However, the lipolytic effects of α-LA were already observed after 1 h of

treatment when AMPK phosphorylation was not induced, suggesting that AMPK

is not involved in the short-term lipolytic effects of α-LA. On the contrary, the

presence of the AMPK activator AICAR inhibited LA-stimulated lipolysis at 24 h

of treatment, according with the remarkable increase of AMPK phosphorylation

observed at this time. Taken together, these data suggest that the lipolytic action

of LA is not mediated by the activation of AMPK in the first stages but could

contribute to the regulation of the long-term lipolytic effects of LA.

The present data also revealed that the JNK and ERK1/2 pathways are not

likely to be main determinants for α-LA-stimulated-lipolysis in adipocytes.

Moreover, the lipolytic effect of α-LA was not caused by a global down-regulation

of adipocyte differentiation, since mRNA levels of some of the main adipogenic

transcription factors Pparγ, C/ebpα and C/ebpβ were not significantly altered in

α-LA-treated adipocytes. On the other hand, the lipolytic actions of α-LA seem to

be independent of its antioxidant capacities, since other antioxidants such as

Vitamin C or NAC were not able to stimulate lipolysis.

Interestingly, our data demonstrated that α-LA inhibits FA esterification in

overweight/obese fully differentiated adipocytes. DGAT1 catalyzes the final

acylation step to yield triglycerides. Our data show the ability of α-LA to

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174

decrease DGAT1 protein content in adipocytes from overweight/obese subjects,

suggesting that DGAT1 inhibition could contribute to the reduction in triglyceride

content observed in α-LA-treated adipocytes. Enlarged adipocytes are

considered to arise from increased fat deposition not only as a consequence of

esterification of FFA but also by de novo lipogenesis (Lafontan, 2013). While the

lipogenic capacity of adipose tissue in humans is believed to be much smaller

than in liver, some studies have suggested that adipose tissue may account for

up to 40% of whole-body lipogenesis (Schwarz et al., 2000). High lipogenic rates

in adipose tissue may contribute to the development of obesity (Jeyakumar et

al., 2009). In this context, several studies in rodents have demonstrated that

dietary supplementation with α-LA reduced lipogenesis in liver and muscle,

however there were no studies addressing the effects of α-LA on lipogenesis in

human adipose tissue (Park et al., 2008; Chen et al., 2012; Valdecantos et al.,

2012).

Our present data clearly show that the intracellular lipid-lowering effects of α-

LA are associated with suppressed de novo lipogenesis, which could be

triggered, in part, by the inhibition of some of the main enzymes regulating this

pathway such as FAS and SCD1. FAS catalyzes the first committed step in de

novo lipogenesis, and adipose tissue FAS has been implicated in obesity and

insulin resistance in humans (Roberts et al., 2009). Here, we demonstrate that

α-LA diminishes FAS protein level. Targeted deletion of Fas in adipose tissue

has been shown to decrease adiposity in mice, which are resistant to diet-

induced obesity (Lodhi et al., 2012). SCD1 is a key enzyme involved in the

control of de novo lipogenesis by catalyzing the rate-limiting step in the

synthesis of MUFA (Dobrzyn, 2012). Scd1 deficiency reduces lipogenesis and

protects mice from diet-induced obesity (Kim et al., 2011). We have found that α-

LA treatment also caused a strong inhibition in SCD1 protein level in our model

of human adipocytes. In addition to FAS and SCD1 decrease, α-LA also reduced

ACC protein levels, a key enzyme of the lipogenic pathway which mediates the

initial step of the FA synthesis. ACC activity is regulated mainly by

phosphorylation, which causes enzyme inactivation (Kim, 1997). In fact, AMPK

activation phosphorylates and inactivates ACC in adipocytes (Peng et al., 2012).

In agreement with the study of Chen et al. (Chen et al., 2012) reporting that FAS

inhibition is an important consequence of AMPK activation mediated by α-LA in

C2C12 myotubes, several of our findings suggest that AMPK mediates the α-LA

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175

inhibitory effects on lipogenic enzymes in human adipocytes from

overweight/obese subjects. Thus, an increase in AMPK phosphorylation was

observed after α-LA treatment, which was accompanied by the subsequent

increase of ACC phosphorylation. In addition, the use of the AMPK antagonist

Compound C reversed the α-LA-mediated down-regulation observed in FAS

protein levels.

AMPK is an activator of FA oxidation in several tissues (Ceddia, 2013) and

this action is also mediated by the phosphorylation of ACC and the subsequent

formation of malonyl-CoA, which has two important functions: 1) it serves as the

essential substrate for FA synthesis as we previously described, and 2) it also

blocks CPT1, a rate-limiting step for the entry of FA from the cytosol into the

mitochondria, where they undergo oxidative metabolism. Through these effects,

malonyl-CoA is a key control point for fat metabolism in adipose and other

peripheral tissues (Brownsey et al., 2006). In this context, the current study

shows that α-LA-induced phosphorylation and inhibition of ACC, is accompanied

in parallel by an increase CPT1. In addition, α-LA treatment also increased

ACOX, which catalyzes the initial and rate-determining step of the peroxisomal

FA β-oxidation pathway. These facts together with the observation that α-LA-

treated adipocytes exhibit increased oxygen consumption suggest that α-LA

might promote oxidation of FA within adipocytes. However, surprisingly, the

oxidation rate of exogenous palmitate was not affected after 24 h of α-LA

treatment. This could be explained by the fact that α-LA decreases the uptake of

free FA by adipocytes, reducing their availability for oxidation. In this context,

Gaidhu et al. (2006) observed that adipocytes treated with the AMPK activator

AICAR exhibited a reduction in FA oxidation probably as a consequence of a

potent suppression of FA uptake.

In the present study, α-LA was revealed as a potent lipolytic agent, and

therefore, α-LA-treated adipocytes have increased amount of endogenous FFA

to be oxidized, which is in accordance with the upregulation of the main

enzymes involved in thi process. Moreover, the inhibition of exogenous FFA

uptake observed in α-LA-treated cells could also be an adaptive mechanism to

counteract the overload in endogenous FFA as a consequence of increased

lipolysis. Taking together these data we further hypothesize that the absence of

effects observed on exogenous palmitate oxidation observed in subcutaneous

adipocytes from overweight/obese subjects could be the result of an inhibition on

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General discussion/Summary

176

exogenous FA uptake, and therefore we cannot rule out the ability of α-LA to

increase FA oxidation based only on this observation. Another important issue to

take into consideration is that increased lipolysis and FFA released after the

stimulation of lipolysis may be re-esterified or oxidized within adipocytes, or they

are released from adipocytes and exported to other tissues. It is important to

take into account that increased FFA release from adipose tissue under lipolytic

conditions has been associated with the development of insulin resistance

(Ormseth et al., 2011). However, recent findings have demonstrated that

increasing lipolysis in adipose tissue does not necessarily increase serum FFA

levels because increasing lipolysis in adipose tissue causes a shift within

adipocytes toward increased FA utilization and energy expenditure and thus

protects against obesity. Therefore, it has been suggested that an activation of

lipolysis may be a promising therapeutic target for the treatment of obesity

(Ahmadian et al., 2009; Ahmadian et al., 2010). In this context, we and others

have demonstrated that dietary supplementation with α-LA reduces weight loss

and fat mass without increasing circulating FFA and improves insulin resistance

in rodents (Park et al., 2008; El Midaoui et al., 2011; Fernandez-Galilea et al.,

2011) and in humans (Zhang et al., 2011), and, as previously suggested, this

could be associated with α-LA-induced FA oxidation. Moreover, several studies

have demonstrated that α-LA also increases FA oxidation in non-adipose cells

both in vitro and in vivo in rodents. Thus, in C2C12 myotubes α-LA produced an

increase in palmitate oxidation using a range of concentrations (30-1,000 µM)

similar to the dose used in the present study (Chen et al., 2012). Moreover,

dietary administration of α-LA increased in liver of high fat-fed mice and rats, the

expression of genes related to beta-oxidation, such as acyl-coenzyme A

dehydrogenase, Cpt1 and Acox (Valdecantos et al., 2012; Kim et al., 2004).

AMPK has been suggested not only to promote rapid metabolic changes in

cells, but also to increase the expression and activity of transcription factors that

induce mitochondrial biogenesis (Zong et al., 2002; Jager et al., 2007). Thus,

under conditions of prolonged AMPK activation, WAT metabolism could be

remodeled towards oxidation rather than storage (Ceddia, 2013). In this line,

previous studies in murine 3T3-L1 adipocytes showed that α-LA treatment, both

alone (Shen et al., 2011) or in combination with acetyl-L-carnitine (Shen et al.,

2008) promoted mitochondrial biogenesis. Importantly, our current study

demonstrates that α-LA is able to increase mitochondrial biogenesis in

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177

adipocytes in overweight/obese condition in humans, as revealed by the higher

mitochondrial mass content observed with TEM imaging, mitotracker green

staining and the raise in mitochondrial DNA content. These effects seem to be

secondary to α-LA-mediated upregulation of Nrf1 and Tfam levels caused, at

least in part, by activation of PGC-1α, a master regulator of mitochondrial

biogenesis. It is well established that AMPK is able to activate PGC-1α (Jager et

al., 2007), and therefore the stimulatory effects of α-LA on mitochondrial

biogenesis could be in part secondary to α-LA-induced AMPK activation. α-LA

also increased the levels of SIRT1, which is also able to promote mitochondrial

biogenesis through deacetylation and activation of PGC-1α (Rodgers et al.,

2005; Gerhart-Hines et al., 2007). In support of our study, previous trials have

observed that SIRT1 is a target for α-LA. In addittion, α-LA increases

mitochondrial copy number in the liver of rats fed on a high fat diet and it also

stimulates mitochondrial function in C2C12 myotubes by increasing SIRT1

deacetylase (Chen et al., 2012; Valdecantos et al., 2012). Finally, PGC-1α

integrates and coordinates the activity of multiple transcription factors, such as

NRF1 and 2, which control the transcription of key mitochondrial proteins

including mitochondrial transcriptional factor, like TFAM that is essential for

mitochondrial replication and transcription (Santos and Kowluru, 2011).

While white adipose tissue functions as an energy storage organ, brown

adipose tissue (BAT) is an energy consumption organ. The principal function of

brown adipocytes is to burn fat in their abundant mitochondria to generate heat.

Another recently discovered cell type is the beige (also called inducible brown,

brown-in-white, or brite) adipocyte. The accumulation of beige adipocytes in

WAT is often referred to as ‘browning’ of WAT (Bartelt and Heeren, 2013). In this

context, the enhancement of mitochondrial biogenesis and the brown-like

phenotype within WAT have been proposed as a promising strategy to combat

obesity and its associated disorders (Liu et al., 2009; Bartelt and Heeren, 2013).

Our TEM studies revealed that α-LA not only increase mitochondrial mass

but also induced changes in mitochondrial morphology. In fact, mitochondria

from α-LA-treated adipocytes exhibit some morphological characteristics of

brown mitochondria, being larger and with numerous transverse cristae

(Barbatelli et al., 2010) in comparison with the untreated adipocytes. These data

suggest the ability of α-LA to induce a remodelling of white adipocytes,

characterized by increased mitochondrial biogenesis and FA oxidation enzymes,

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General discussion/Summary

178

and accompanied by the acquirement of brown-like characteristics. In this

context, it is known that increased clearance and utilization of nutrients by brown

and beige adipocytes could reduce the excess of triglycerides and confer

beneficial metabolic effects or protection from obesity, highlighting the important

anti-obesity role of the acquisition of beige features by white adipocytes.

Among these brown-like characteristics, Cidea, considered a brown

adipocyte marker, was upregulated in α-LA treated adipocytes (Barneda et al.,

2013). Taking together, the increase in mitochondrial biogenesis and FA

oxidation machinery and the changes observed in the mitochondrial morphology

suggest that α-LA might induce a brown-like phenotype within white

subcutaneous adipocytes. Recently, it has been established that inducible brown

adipocytes are phenotypically distinct from both white and brown adipocytes. In

this context, TBX1 has been identified as a reliable marker for beige adipocytes

in humans (Jespersen et al., 2013). Our data showed that α-LA treatment

increased Tbx1 mRNA levels suggesting that α-LA might promote

transdifferentiation of mature white adipocytes into brite adipocytes. Moreover, it

has been described that human beige adipocytes initially expressed low levels of

UCP1, but expression of UCP1 can be induced after mimicking cold exposure or

by different treatments such as β-3 adrenergic and PPARγ agonists (Bartelt and

Heeren 2013). Interestingly, our results demonstrated that α-LA is also able to

induce UCP1 in subcutaneous adipocytes of overweight/obese subjects.

Browning of WAT can be caused by transcriptional modulation through

PRDM16 (Seale et al., 2011) and PGC-1α (Tiraby et al., 2003). In fact, in

primary human subcutaneous adipocytes, adenovirus-mediated expression of

Pgc-1α was described to lead to a brown-fat phenotype and increased

respiratory chain proteins and FA oxidation enzymes (Tiraby et al., 2003).

Transgenic expression of Prdm16 in subcutaneous fat tissue also induces the

brown-like cells (Seale et al., 2011), and PRDM16 has been described as a

required and sufficient molecule to promote brown features in white adipose

tissue (Seale et al., 2007; Seale et al., 2008). PRDM16 directly binds to PGC-1α

allowing the activation of other brown fat-specific genes (Seale et al., 2007). In

this context, our data have revealed that the up-regulation of PRDM16 observed

after α-LA treatment is accompanied by the activation of PGC-1α, which might

promote the white to brite transdifferentiation.

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General discussion/Summary

179

Recently, it has been shown that PGC1-α stimulates the expression of

Fndc5, which encodes for irisin, a polypeptide hormone cleaved and released by

muscle (Bostrom et al., 2012) and adipose tissue (Moreno-Navarrete et al.,

2013). Irisin induces the browning of subcutaneous white adipose tissue

(Bostrom et al., 2012). In this line, our data revealed that α-LA up-regulates

Fndc5 mRNA levels and also irisin release from adipocytes, which could also

contribute to the browning properties of α-LA in white adipocytes. Although

controversial effects of irisin on browning process in humans have been

observed (Raschke et al., 2013), these findings might corroborate the previoulsy

described anti-obesity properties of α-LA.

Recently, it has been suggested that inhibiting adipose tissue lipogenesis

reprograms subcutaneous adipose tissue thermogenesis. In fact, this study

demonstrated that mice lacking FAS in adult adipose tissue manifested

increased brown fat-like (brite) adipocytes in subcutaneous adipose tissue,

characterized by increased expression of PRDM16, Cidea and PGC-1α as well

as CPT1 and ACOX (Lodhi et al., 2012). In this context, these findings suggest

that the inhibition induced by α-LA on FAS and de novo lipogenesis might also

contritube to the ability of α-LA to promote beige adipocytes features.

In summary, our study demonstrate that α-LA: 1) Inhibits FA uptake (through

CD36 downregulation); 2) Reduces FA esterification into triglycerides (through

DGAT1 reduction); 3) Decreases de novo lipogenesis (induced by inhibition of

FAS, SCD1 and ACC mediated by AMPK activation); 4) Stimulates lipolysis

(mainly mediated by the phosphorylation of HSL through cAMP-mediated

activation of PKA, probably through the inhibition of AdPLA and PGE2; 5)

Increases FA oxidation machinery (CPT1 and ACOX) in white adipocytes; 6)

Promotes mitochondrial biogenesis by the activation of PGC-1α mediated by

SIRT1 and AMPK activation; and 7) Induces a brown-like remodelling in white

subcutaneous adipocytes from overweight/obese subjects.

Taking together all of these facts suggest that α-LA remodels adipocyte

metabolism towards oxidation rather than storage, which might convert white

adipocytes into a more effective ‘‘fat burning machines’’, and this could

contribute to the anti-obesity properties of α-LA and could help to improve

patients’ metabolic state and health.

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General discussion/Summary

180

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CCHHAAPPTTEERR 66//CCAAPPÍÍTTUULLOO 66

CCOONNCCLLUUSSIIOONNSS//CCOONNCCLLUUSSIIOONNEESS

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Conclusions/Conclusiones

189

CONCLUSIONS

1. α-LA significantly stimulates lipolysis in 3T3-L1 adipocytes. These

lipolytic actions are mainly mediated by phosphorylation of the lipid

droplet coat protein “perilipin” and “hormone sensitive lipase” (HSL) at

Ser563

and at Ser660

. However, α-LA did not significantly modify the

expression of other key lipase, ATGL, or its activator CGI-58 and inhibitor

G0S2.

2. α-LA also caused a significant inhibition of a novel intracellular

adipose-specific phospholipase A2 (AdPLA) in parallel with a decrease in

the amount of PGE2 released and an increase in intracellular cAMP

content. Moreover, the PKA inhibitor H89 completely blunted the lipolytic

action of α-LA as well as the α-LA-induced phosphorylation of phospho-

PKA substrates perilipin and HSL. Taken together these data suggest

that the lipolytic effects of α-LA are mediated by a cAMP-mediated

activation of PKA possibly caused by the inhibition of AdPLA and PGE2.

3. α-LA inhibits AKT phosphorylation and both the PI3K/AKT inhibitor

LY294002 and the PDE3B antagonist Cilostamide potentiated the

stimulatory effects of α-LA on basal lipolysis, suggesting that the lipolytic

effects of α-LA could also be explain by an inhibition of AKT signaling

pathway, which might also increase cAMP, and, therefore, induce the

activation of PKA and the subsequent events previously described.

4. α-LA treatment reduces triglyceride accumulation in cultured

subcutaneous adipocytes from overweight/obese subjects by inhibiting

both de novo lipogenesis and FA esterification. These effects are

mediated by the reduction observed in the protein levels of key enzymes

regulating these processes FAS, SCD1 and DGAT1.

5. α-LA increases AMPK phosphorylation, which is accompanied by

the subsequent increase in ACC phosphorylation and inactivation.

Moreover, the AMPK inhibitor Compound C reversed the α-LA-mediated

down-regulation observed in FAS protein levels, suggesting that AMPK

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Conclusions/Conclusiones

190

might mediate, at least in part, the α-LA inhibitory effects observed on the

main lipogenic enzymes.

6. Human subcutaneous adipocytes from overweight/obese subjects

treated with α-LA exhibits an increase in mitochondrial mass and mtDNA

content. These actions are accompanied by an increase in SIRT1 protein

levels and by the activation of PGC-1α by deacetylation, as well as by an

increase in the gene expression of the transcriptional factors Nrf1 and

Tfam involved in mitochondrial biogenesis. These facts suggest that α-LA

promotes mitochondrial biogenesis through the activation of the

SIRT1/PGC-1α signaling pathway.

7. α-LA also enhances oxygen consumption and the protein levels of

CPT1 and ACOX, two rate limiting enzymes involved in FA oxidation.

However, α-LA reduces FA uptake and the FA traslocase CD36 protein

content in subcutaneous adipocytes from overweight/obese subjects.

8. Mitochondria from α-LA-treated white adipocytes exhibit

morphological characteristics of brown adipocytes mitochondria.

Moreover, α-LA treatment up-regulates several brown/beige-adipocytes

distinctive genes and proteins such as Cidea, Tbx1 and UCP1 suggesting

the ability of α-LA to induce the acquirement of beige features in

subcutaneous adipocytes from overweight/obese subjects. These

potential browning properties of α-LA could be mediated by the up-

regulation of Prdm16 gene, a transcription factor that promotes brown

features in white adipose tissue, and by Fndc5 mRNA and irisin release,

which is also able to induce browning of subcutaneous white adipose

tissue.

GENERAL CONCLUSION

The current data show the ability of α-LA to modulate lipid metabolism in

white adipocytes. Our study demonstrate that the anti-adiposity actions

previously described for α-LA could be due, at least in part, by inhibitory actions

on FA uptake and esterification into triglycerides, by the reduction observed in

de novo lipogenesis and, finally, by the stimulation of lipolysis and FA oxidation

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191

machinery in white adipocytes. Moreover, the capability of α-LA to induce

mitochondrial biogenesis and brown-like remodelling in white subcutaneous

adipocytes from overweight/obese subjects, suggest that these mechanisms

might also contribute to the anti-obesity properties of α-LA and could help to

improve patients’ metabolic state and health.

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Conclusions/Conclusiones

192

CONCLUSIONES

1. El α-LA estimula significativamente la lipólisis en adipocitos 3T3-

L1. Dichas acciones lipolíticas son mediadas por la fosforilación de

perilipina, una proteína que recubre la gota lipídica, y de la lipasa

sensible a hormonas (HSL) en los residuos Ser563

y Ser660

. Sin embargo,

el α-LA no modificó significativamente la expresión de otra lipasa clave

como es la ATGL ni a su proteína activadora CGI-58 ni a la inhibidora

G0S2.

2. El α-LA causó una inhibición significativa de la recientemente

descrita fosfolipasa intracelular A2, específica del tejido adiposo

(AdPLA). Así mismo, produjo una reducción en los niveles de PGE2

liberados al medio de cultivo y un aumento de los niveles de cAMP

intracelular. Además, el inhibidor de PKA H89 revirtió completamente las

acciones lipolíticas del α-LA así como la fosforilación inducida por el α-LA

de los substratos de PKA, perilipina y HSL. En conjunto, estos datos

sugieren que los efectos lipolíticos del α-LA están mediados por la

activación de PKA inducida, a su vez, por el incremento de cAMP,

posiblemente causado por la inhibición de AdPLA y PGE2.

3. El α-LA inhibe la fosforilación de AKT mientras que el inhibidor de

PI3K/AKT, LY294002, así como el antagonista de la PDE3B,

Cilostamide, potenciaron los efectos estimulantes del α-LA en la lipólisis

basal, lo cual sugiere que los efectos lipolíticos del α-LA podrían estar

mediados por una reducción en la activación de AKT, lo cual podría

aumentar los niveles de cAMP y, esto, a su vez, activar a PKA y a toda la

cascada de eventos señalados anteriormente.

4. El tratamiento con α-LA de adipocitos subcutáneos procedentes

de individuos con sobrepeso u obesidad reduce la acumulación de

triglicéridos mediante la inhibición de la lipogenesis de novo así como de

la esterificación de ácidos grasos. Estos efectos están mediados por la

reducción observada en los niveles de las principales enzimas

implicadas en estos procesos como son la FAS, SCD1 y DGAT1.

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193

5. El α-LA aumenta la fosforilación de AMPK, lo cual conlleva un

aumento en la fosforilación e inactivación de ACC. El inhibidor de AMPK,

Compound C, revirtió los efectos inhibitorios del α-LA sobre los niveles

de FAS, lo cual sugiere que AMPK podría mediar, al menos en parte, las

acciones del α-LA sobre la activación de las enzimas lipogénicas.

6. El tratamiento con α-LA de adipocitos subcutáneos procedentes

de individuos con sobrepeso u obesidad indujo un incremento de la masa

mitocondrial y del contenido en DNAmt. Estas acciones fueron

acompañadas por un aumento de SIRT1 y la activación, mediante

desacetilación, de PGC-1α junto con un incremento en la expresión de

dos factores de transcripción implicados en la biogénesis mitocondrial

Nrf1 y Tfam. Nuestros datos sugieren, por tanto, que el α-LA promueve la

biogénesis mitocondrial mediante la activación de la ruta SIRT1/ PGC-

1α.

7. El α-LA aumentó el consumo de oxígeno y los niveles de CPT1 y

ACOX, dos enzimas claves de la oxidación de ácidos grasos. Sin

embargo, este ácido redujo la captación de ácidos grasos así como los

niveles de la proteína traslocasa CD36 en adipocitos subcutáneos de

sujetos con sobrepeso u obesidad.

8. Las mitocondrias de los adipocitos blancos tratados con α-LA

mostraron características morfológicas similares a las que se encuentran

en adipocitos de tipo pardo. Además, el tratamiento con α-LA indujo un

aumento en los niveles de expresión de algunos genes y proteínas

característicos del adipocito pardo o beige como son el Cidea, Tbx1 y

UCP1 lo cual sugiere la capacidad del α-LA de inducir la adquisición de

un fenotipo beige en adipocitos subcutáneos de sujetos con sobrepeso u

obesidad. Este “pardeamiento” inducido por el α-LA podría estar mediado

por un aumento de la expresión génica de Prdm16, un factor de

transcripción que promueve la aparición de características de adipocitos

pardos en los adipocitos blancos, así como por el incremento observado

en los niveles de mRNA de Fndc5 y de la liberación de irisina, los cuales

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194

son también capaces de inducir el “pardeamiento” en tejido adiposo

blanco subcutáneo.

CONCLUSIÓN GENERAL

Nuestros datos muestran la capacidad del α-LA de modular el metabolismo

lipídico en adipocitos blancos. De hecho, el presente estudio sugiere que las

propiedades anti-adiposidad previamente descritas para este ácido podrían

estar mediadas, al menos en parte, por sus acciones inhibitorias en la captación

de ácidos grasos y posterior esterificación a triglicéridos, por la reducción de la

lipogenesis de novo y, finalmente, por sus efectos estimulantes sobre la lipólisis

y sobre enzimas implicados en la oxidación de ácidos grasos en adipocitos

blancos. Además, la capacidad del α-LA de promover la biogénesis mitocondrial

e inducir características fenotípicas de adipocitos pardos/beige en los adipocitos

subcutáneos blancos de sujetos con sobrepeso u obesidad, sugieren que estos

mecanismos también podrían contribuir a las propiedades anti-obesidad del α-

LA y, por tanto, podrían mejorar el estado metabólico y, en consecuencia, la

salud de estos individuos.

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AANNEEXXOOSS

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