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
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
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.
“Es a fuerza de observación y
reflexión que uno encuentra un
camino”.
Claude Monet
“Si tu intención es describir la
verdad, hazlo con sencillez y la
elegancia déjasela al sastre.”
Albert Einstein
A mis padres
A Pedro
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
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
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.
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
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
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
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
CHAPTER 5: GENERAL DISCUSSION/SUMMARY ................................. 171
CHAPTER 6: CONCLUSIONS ................................................................... 189
ANEXOS
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
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
Introduction
1
CCHHAAPPTTEERR 11
IINNTTRROODDUUCCTTIIOONN
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.
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
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)
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
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).
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.
Introduction
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
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
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
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
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).
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).
Introduction
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
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,
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
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
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
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
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.
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
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
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
Introduction
23
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).
Introduction
24
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,
Introduction
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
Introduction
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).
Introduction
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.
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
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
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
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
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
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
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.
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
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.
Introduction
37
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CCHHAAPPTTEERR 22//CCAAPPÍÍTTUULLOO 22
HHYYPPOOTTHHEESSIISS AANNDD AAIIMMSS//HHIIPPÓÓTTEESSIISS YY OOBBJJEETTIIVVOOSS
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.
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.
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.
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.
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.
CCHHAAPPTTEERR 33//CCAAPPÍÍTTUULLOO 33
MMAATTEERRIIAALL AANNDD MMEETTHHOODDSS//MMAATTEERRIIAALL YY MMÉÉTTOODDOOSS
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).
Material y Métodos
66
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
Material y Métodos
67
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).
Material y Métodos
68
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
Material y Métodos
69
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
Material y Métodos
70
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
Material y Métodos
71
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
Material y Métodos
72
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
Material y Métodos
73
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
Material y Métodos
74
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).
Material y Métodos
75
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
Material y Métodos
76
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
Material y Métodos
77
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
Material y Métodos
78
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).
Material y Métodos
79
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
Material y Métodos
80
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
Material y Métodos
81
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
Material y Métodos
82
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.
Material y Métodos
83
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
Material y Métodos
84
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
Material y Métodos
85
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,
Material y Métodos
86
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
Material y Métodos
87
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
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.
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.
CCHHAAPPTTEERR 44
RREESSUULLTTSS
CCHHAAPPTTEERR 44..11
EEffffeeccttss ooff lliippooiicc aacciidd oonn lliippoollyyssiiss iinn 33TT33--LL11 aaddiippooccyytteess
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.
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
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,
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
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.
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).
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
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).
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
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 (%
)
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-
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
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
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).
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).
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
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.
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
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
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 (
%)
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.
Results (Chapter 4.1)
116
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Results (Chapter 4.2)
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.
Results (Chapter 4.2)
126
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
Results (Chapter 4.2)
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.
Results (Chapter 4.2)
128
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.
Results (Chapter 4.2)
129
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.
Results (Chapter 4.2)
130
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)
Results (Chapter 4.2)
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).
Results (Chapter 4.2)
132
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
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.
Results (Chapter 4.2)
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
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).
Results (Chapter 4.2)
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.
Results (Chapter 4.2)
137
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preserved in cultured primary myotubes from obese donors in response to
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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.
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
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.
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
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,
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-
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.
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.
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).
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).
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
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).
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).
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).
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
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.
Results (Chapter 4.3)
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-
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.
Results (Chapter 4.3)
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.
Results (Chapter 4.3)
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Results (Chapter 4.3)
167
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CCHHAAPPTTEERR 55
GGEENNEERRAALL DDIISSCCUUSSSSIIOONN//SSUUMMMMAARRYY
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.,
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
General discussion/Summary
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
General discussion/Summary
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
General discussion/Summary
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
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
General discussion/Summary
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,
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.
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.
General discussion/Summary
180
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CCHHAAPPTTEERR 66//CCAAPPÍÍTTUULLOO 66
CCOONNCCLLUUSSIIOONNSS//CCOONNCCLLUUSSIIOONNEESS
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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
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
Conclusions/Conclusiones
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.
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.
Conclusions/Conclusiones
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
Conclusions/Conclusiones
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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