Effect of cellular folate availability on adipocyte life ...Effect of cellular folate availability on adipocyte life cycle and metabolism 1 Abstract The role of folate in obesity and

Effect of cellular folate availability

on adipocyte life cycle and metabolism

Efeito da disponibilidade celular de folatos

no metabolismo e ciclo de vida de adipócitos

Cláudia Sofia Fonseca Marques

Orientação: Mestre Ana Isabel Gonçalves Faria

Co-orientação: Mestre Elisa Alexandra Mota Ferreira

Trabalho de Investigação

Porto, 2010

i

Ao meu Pai e à minha Mãe

iiiiiiiiii ii Effect of cellular folate availability on adipocyte life cycle and metabolism

iii Effect of cellular folate availability on adipocyte life cycle and metabolism

Agradeço à Professora Doutora Conceição Calhau, à Professora Doutora Elisa

Keating, à Professora Doutora Rosário Monteiro e à Mestre Ana Faria, pois sem

os seus oportunos ensinamentos e sem a sua experiência e sabedoria a

realização deste trabalho não teria sido possível.

iviviviviv iv Effect of cellular folate availability on adipocyte life cycle and metabolism

v Effect of cellular folate availability on adipocyte life cycle and metabolism

Contents

Abbreviations ......................................................................................................... vii

Abstract .................................................................................................................. 1

Resumo .................................................................................................................. 3

Introduction............................................................................................................. 5

Materials and methods ........................................................................................... 9

Cells, chemicals and reagents ............................................................................ 9

Cell culture ........................................................................................................ 10

Sulforhodamine B assay.................................................................................... 11

Methyl-3H-thymidine incorporation into DNA ..................................................... 12

Lipid accumulation in adipocytes ...................................................................... 12

Measurement of 3 H-deoxyglucose uptake ....................................................... 13

Determination of cell viability ............................................................................ 14

Determination of homocysteine released to the culture medium ...................... 14

Statistical analysis ............................................................................................ 15

Results ................................................................................................................. 17

Discussion ............................................................................................................ 25

Conclusion............................................................................................................ 32

References ........................................................................................................... 33

vii Effect of cellular folate availability on adipocyte life cycle and metabolism

Abbreviations

AICAR - 5-amino-4-imidazolecarboxamide ribonucleotide AR - adenosine receptors BMI - body mass index cpm - counts per minute DEX - dexamethasone

DG - deoxyglucose

DHF - dihydrofolate

DHFR - dihydrofolate reductase

DMEM - Dulbecco’s modified Eagle’s medium

DMSO - dimethyl sulfoxide DNA - deoxyribonucleic acid

EDTA - ethylenediaminetetraacetic acid FBS - fetal bovine serum GLUT - glucose transporter HEPES - 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid HSL - hormone-sensitive lipase IBMX - 3-isobutyl-1-methylxanthine LDH - lactate dehydrogenase MTX - methotrexate NADH - nicotinamide adenine dinucleotide NAFLD - nonalcoholic fatty liver disease PBS - phosphate-buffered saline PKA - protein kinase A RFC - reduced folate carrier

viii Effect of cellular folate availability on adipocyte life cycle and metabolism

RNA - ribonucleic acid SAH - S-adenosyl-homocysteine SAHH - S-adenosyl-homocysteine hydrolase SAM - S- adenosyl-methionine SEM - standard error of mean SRB - sulforhodamine B

TAG - triglycerides

TCA - trichloroacetic acid

THF - tetrahydrofolate

Tris - tris(hydroxymethyl)aminomethane TS - thymidylate synthase

1 Effect of cellular folate availability on adipocyte life cycle and metabolism

Abstract

The role of folate in obesity and metabolic syndrome has started to be

investigated but is far from being fully understood. Many studies have drawn

attention to the association of folate status and plasma homocysteine levels, an

established independent risk factor for cardiovascular disease development.

Recently, epidemiologic data shows an inverse association between serum folate

and body mass index. However, there is no straight evidence about the effects of

folate on adipose tissue. In this regard, the aim of this study was to investigate the

effect of cellular folate availability on preadipocyte proliferation, and on adipocyte

adipogenesis and glucose uptake.

3T3-L1 preadipocytes were cultured to evaluate proliferation by

sulforhodamine B staining and methyl-3H-thymidine incorporation, after 24 h or 48

h of treatment with methotrexate (MTX, 0.1 and 10 µM), an inducer of a low

cellular folate status. Preadipocytes were induced to differentiate with an

appropriate adipogenic cocktail in the presence or absence of MTX, and

adipogenesis was determined by measuring lipid accumulation after staining with

oil red O. 3H-Deoxyglucose uptake was determined by liquid scintillation counting.

MTX treatment for 24 h and 48 h reduced culture protein content and

methyl-3H-thymidine incorporation in a time- and concentration-dependent manner

(P < 0.05). At the end of treatment, there was even a lower protein content than in

the beginning of treatment for the highest concentration of MTX (10 µM) used (P <

0.05), revealing cytotoxicity. In adipocytes, MTX treatment increased lipid

accumulation and the effect was much more pronounced for the highest

concentration of MTX. Furthermore, MTX (10 µM) increased basal glucose uptake

(P < 0.05 vs control and MTX 0.1 µM). However, in MTX (10 µM)-treated


adipocytes, insulin-stimulation did not result in an increase of glucose uptake,

contrary to what was observed in untreated adipocytes.

According to the new understanding of obesity therapeutic strategies with

the aim of reducing associated metabolic complications, adipocyte hypertrophy

and dysfunction could be prevented if adipose tissue preserves the ability to recruit

preadipocytes to differentiate, so that fat content can be distributed among the

new adipocytes. Our results showed that, when folate availability was

compromised by MTX treatment, there was a stimulation of preadipocyte

differentiation, but this stimulation was accompanied by a decrease in

preadipocyte number. Furthermore, the results obtained in glucose uptake studies

implying that these cells were resistant to insulin stimulation.

In conclusion, our results suggest that folate deprivation can interfere with

adipocyte proliferation, differentiation and metabolism and promote the

hypertrophic growth of adipocytes, which may contribute to the development of

obesity complications and the metabolic syndrome.

Keywords: Folate; metabolic syndrome; methotrexate; obesity; 3T3-L1.


Resumo

O papel dos folatos na obesidade e na síndrome metabólica começa agora

a ser estudado mas ainda está longe de ser compreendido. Vários estudos têm

focado a sua atenção na associação dos níveis plasmáticos de folatos com os de

homocisteína, um factor de risco independente para o desenvolvimento da

doença cardiovascular. Mais recentemente, estudos epidemiológicos revelaram

uma associação inversa entre os níveis plasmáticos de folatos com o índice de

massa corporal. Contudo, não há evidências directas sobre os efeitos dos folatos

no tecido adiposo. Assim, o presente trabalho teve como objectivo investigar o

efeito da disponibilidade celular de folatos na proliferação dos pré-adipócitos e na

captação de glicose e na adipogénese de adipócitos.

Para tal, usou-se a linha celular de pré-adipócitos 3T3-L1 e avaliou-se a

sua proliferação por coloração das proteínas da cultura com sulforrodamina B

(SRB) e a incorporação de metil-timidina-3H após 24 h ou 48 h de tratamento com

metotrexato (MTX, 0.1 and 10 µM), um indutor de baixos níveis celulares de

folatos. A diferenciação foi induzida nos pré-adipócitos com um cocktail

apropriado de diferenciação na presença ou ausência de MTX e a adipogénese

foi determinada pela quantificação da acumulação lipídica após a coloração com

oil red O. A captação de desoxiglicose-3H pelos adipócitos foi determinada por

cintilometria líquida.

O tratamento com metotrexato durante 24 h ou 48 h reduziu o conteúdo

proteico da cultura celular e a incorporação de metil-timidina-3H de uma forma

dependente do tempo e da concentração (P < 0,05). No final do tratamento com

MTX 10 µM, o conteúdo proteico celular era ainda mais baixo do que no início da


experiência (P < 0,05), o que traduziu um efeito citotóxico. Nos adipócitos, o

tratamento com MTX aumentou a acumulação lipídica, sendo este efeito muito

mais pronunciado quando usada a concentração mais elevada do composto. O

tratamento com MTX também aumentou o transporte basal de glicose, contudo, a

estimulação pela insulina não aumentou a captação de glicose nos adipócitos

tratados com MTX, contrariamente ao que se observou nos adipócitos não

tratados.

De acordo com as novas estratégias terapêuticas da obesidade, cujo

objectivo é a redução das suas complicações metabólicas, a hipertrofia e

disfunção dos adipócitos pode ser prevenida se o tecido adiposo preservar a

capacidade de recrutar pré-adipócitos para diferenciar de modo a haver a

distribuição das reservas de gordura entre os novos adipócitos. Os resultados

deste trabalho demonstram que, quando a disponibilidade de folatos está

comprometida pelo tratamento com MTX, há uma estimulação da diferenciação

dos pré-adipócitos. Porém, esta estimulação é acompanhada pela diminuição do

número de pré-adipócitos. As observações efectuadas nas experiências de

transporte de glicose propõem ainda que estas células são resistentes à

estimulação da insulina.

Concluindo, estes resultados sugerem que a privação de folatos pode

interferir com a proliferação, diferenciação e metabolismo dos adipócitos,

promovendo o seu crescimento hipertrófico, o que pode contribuir para o

desenvolvimento de complicações da obesidade e da síndrome metabólica

Palavras-chave: Folatos; metotrexato; obesidade; síndrome metabólica; 3T3-L1.


Introduction

Folate is a generic term for a naturally occurring family of B-group vitamins

composed of an aromatic pteridine ring linked to p-aminobenzoic acid and a

glutamate residue (Figure 1), which typically appears in food in a reduced,

polyglutamyl form(1). The main dietary sources of folate include green leafy

vegetables, like spinach and asparagus, some fruits, and fortified cereals and

cereal products(2). Folic acid is the most common form of folate used in

supplements and in fortified food products because it is highly bioavailable and

Figure 1 – Chemical structures of folate, 5-methyl-tetrahydrofolate and methotrexate. Adapted

from Assaraf et al(1)

.

chemically stable, since it has only one glutamate molecule(3). During their

passage across the intestinal mucosa, folic acid and most dietary folates, after

cleavage to the monoglutamyl form, are metabolized to 5-methyl-tetrahydrofolate

Folate

Methotrexate

5-methyl-tetrahydrofolate


Homocysteine

(Figure 1), the active coenzyme form of folate, to be absorbed in the proximal

jejunum via the reduced folate carrier (RFC)(4).

Figure 2 – Involvement of folate in pyrimidine biosynthesis and the homocysteine remethylation

cycle. DHFR - dihydrofolate reductase, MetSyn - vitamin-B12-dependent methionine synthase,

MTHFR - methylenetetrahydrofolate reductase, SAH - S-adenosyl-homocysteine, SAHH - S-

adenosyl-homocysteine hydrolase, SAM - S- adenosyl-methionine, THF - tetrahydrofolate, TS -

thymidylate synthase. Adapted from Katula et al(5)

.

Cellular folates act as donors and acceptors of methyl groups in the

biosynthesis of nucleotide precursors used for DNA and RNA synthesis, and

provide methyl groups for DNA, RNA and protein methylation(6). Folic acid is

reduced to dihydrofolate (DHF) and tetrahydrofolate (THF) by dihydrofolate

reductase (DHFR), which is converted to 5,10-methyleneTHF(4). This coenzyme is

a substrate for the thymidylate synthase (TS) enzyme in the methylation of

deoxyuridine monophosphate to deoxythymidine monophosphate, which is the

sole de novo source of thymidine and the rate limiting step in DNA synthesis. 5,10-

MethyleneTHF is also used in the production of formylTHF, which is, in turn, used

in de novo purine synthesis(6) by glycinamide ribonucleotide transformylase and 5-


amino-4-imidazolecarboxamide ribonucleotide (AICAR) transformylase enzymes.

In addition, the conversion of 5,10 - methyleneTHF to THF in the folate cycle

provides the methyl group required to convert homocysteine to methionine(7)

(Figure 2).

Folate deficiency can result from an inadequate folic acid intake,

malnutrition, or increased metabolic needs caused by pregnancy, metastatic

cancer or antifolate chemotherapy, e.g. methotrexate (MTX)(4). MTX is actively

transported into mammalian cells by RFC(8). Once in the cytosol, it binds to and

inactivates DHFR, resulting in the depletion of metabolically active intracellular

folates with subsequent inhibition of the synthesis of thymidylate(9). Inhibition of

DHFR causes cessation of the synthesis of purine metabolites which are crucial

for cell proliferation (9).

Low folate status is associated with elevated homocysteine levels(10), an

independent risk factor for cardiovascular disease(11), with coronary heart disease

(independently of total homocysteine levels)(12), with neural tube defects(13) and,

recently, with increased body mass index (BMI)(14). Epidemiological data suggests

that not only folate deficiency is associated with adiposity or body fat percentage,

but also with absolute amounts of central and peripheral fat(15). It has been pointed

as a risk factor for nonalcoholic fatty liver disease (NAFLD)(16) and, therefore, for

metabolic syndrome, since approximately 90% of the patients with NAFLD have

more than one feature of the metabolic syndrome(17). In addition, another study

has shown that weight loss can help to improve the folate status of overweight and

obese women(18). Understanding the mechanisms underlying this association is of

great significance because obesity and its metabolic associated complications are

major health problems(19).


A number of important functions in intermediary metabolism and hormonal

interactions with other tissues have been attributed to adipose tissue, which is no

longer considered as just an energy store(20). Since there is no straight evidence

about the effects of folate on adipose tissue, the aim of this study was to

investigate the effect of cellular folate availability on preadipocyte proliferation, and

on adipocyte adipogenesis and glucose uptake. For this purpose, the 3T3-L1

murine preadipocyte line was chosen due to the ability of these cells to undergo

differentiation into mature adipocytes and because of its wide use and

characterization as a model of this adipose tissue cellular component(21, 22). To

induce a low cellular folate status, cells were treated chronically with different

concentrations of MTX.


Materials and Methods

Cells, chemicals and reagents

The murine 3T3-L1 cell line was obtained from the American Type Culture

Collection (ATCC, CL-173) through LGC Promochem (Spain). L-amethopterin

hydrate (methotrexate), antibiotic-antimycotic solution, dexamethasone (DEX),

Dulbecco’s modified Eagle’s medium (DMEM), 4-(2-hydroxyethyl)-1-

piperazineethanesulfonic acid (HEPES), insulin, 3-isobutyl-1-methylxanthine

(IBMX), β-nicotinamide adenine dinucleotide, reduced disodium salt hydrate (β-

NADH-Na2), oil red O, sodium piruvate, sulforhodamine B (SRB),

tris(hydroxymethyl)aminomethane (Tris), tris(hydroxymethyl)aminomethane

hydrochloride (Tris-HCl), and trypsin-ethylenediaminetetraacetic acid (EDTA)

were obtained from Sigma-Aldrich Chemicals (U.S.A.). Fetal bovine serum (FBS)

was purchased from Gibco (United Kingdom). Dimethyl sulfoxide (DMSO), gelatin,

p-formaldehyde, trichloroacetic acid (TCA) and triton X-100 were obtained from

Merck (Germany). 3H-Deoxyglucose (3H-DG) and methyl-3H-thymidine were

obtained from American Radiolabeled Chemichals (U.S.A), Inc and Amersham

(U.S.A.), respectively.

MTX was dissolved in NaOH 0.1 M each time it was used, to obtain MTX

0.1 mM and 10 mM. For all experiments, MTX 0.1 mM, MTX 10 mM or the solvent

were dissolved 1:1000 in incubation media of preadipocytes and adipocytes.


Cell culture

3T3-L1 preadipocytes were cultured at 37ºC in a 5% CO2 humidified

atmosphere in DMEM growth medium (containing in average 35.93 ± 1.505

ng/mL of folic acid), supplemented with 1.5 g/L NaHCO3, 10% (v/v) heat-

inactivated FBS (56°C, 30 min), 100 U/mL penicillin and 100 U/mL streptomycin.

Every other day, when cells got confluent, culture was split at a 1:3 ratio by

incubation with 1 mL 0.25% trypsin-EDTA solution, and subcultured in 21 cm2

polystyrene culture plates (21 cm2, Ø 60 mm, Orange Scientific, Belgium).

To measure proliferation, we used 96-well plates (0.37 cm2/well, Orange

Scientific, Belgium) and 24-well plates (1.65 cm2/well, Orange Scientific, Belgium)

and in differentiation assays, cells were seeded in 24-well plates precoated with

0.2% (w/v) gelatin prepared in phosphate-buffered saline (PBS;140 mM NaCl, 3

mM KCl, 8 mM Na2HPO4, 2 mM KH2PO4, pH 7.4).

Adipocytes were obtained through induction of 3T3-L1 preadipocyte

differentiation (Figure 3). Two days after reaching confluence (day 0 of

differentiation), 3T3-L1 preadipocytes were exposed to IBMX (0.5 mM), DEX (0.25

μM), and insulin (10 μg/mL) in growth medium as described(23). After 2 days, this

differentiation cocktail was removed and cells were maintained in growth medium

containing only insulin (10 μg/mL). This medium was changed every 2-3 days.

Adipocytes were used 12 days after differentiation induction when approximately

80% of control cells were differentiated.


Figure 3 – Representation of differentiation protocol and methotrexate (MTX) treatment of

adipocytes. DMEM, Dulbecco’s modified Eagle’s medium; FBS, fetal bovine serum. Adapted

from Teixeira et al(24).

Sulforhodamine B assay

For sulforhodamine B (SRB) assay, cells were seeded (4 × 103 cells in 100

µL per well) in 96-well plates. After 24 h, MTX was added to the culture in different

final concentrations (0.1 µM and 10 µM) for 24 h or 48 h. At the end of each

experiment, 25 μL of ice-cold 50% (w/v) TCA was added to the culture medium on

each well to fix cells for 1 h at 4°C in the dark. Cells were then gently washed 4

times with distilled water to remove TCA. Plates were air-dried and then stained

for 15 min with 0.4% (w/v) SRB dissolved in 1% acetic acid as described(25).

Excess SRB was removed and cultures were quickly rinsed 5 times with 1% acetic

Proliferation

Day -2

Confluence

Day 0

Differentiation

induction

Day 2 Day 8

Differentiation

Day -4

Seeding

Day 4

Growth medium + 0.25 µM dexamethasone + 0.5 mM

isobuthylmethylxanthine + 10 µg/ml insulin

(differentiation medium)

(Differentiation medium)

Growth medium + 10 µg/ml insulin

Day 6

DMEM + 10% (v/v) FBS + 100

IU/ml penicillin + 100 µg/ml

streptomycin (growth medium)

Day 10 Day 12

Assay

Post-Confluence

MTX treatment


acid to remove unbound dye. After drying, the bound dye was solubilized with 150

μL Tris (10 mM, pH 10.5) and the absorbance was determined at 492 nm with

reference at 650 nm on a plate reader (Thermo Electron Corporation, Multiskan

Ascent, U.S.A.). Before the beginning of treatment (day 0), the absorbance of

control wells containing untreated cells in the 96-well plates was determined and

the proliferative activity was calculated from the ratio of the absorbance readings

between treated and these untreated control wells.

Methyl-3H-thymidine incorporation into DNA

Cells were seeded in 24-well cell culture plates (1 × 104 cells per well) in a

final volume of 500 µL culture medium. After 24 h in culture, cells were treated with

different concentrations of MTX (0.1 and 10 µM) or vehicle (NaOH 0.1 mM)

dissolved in culture medium. After 24 h or 48 h, cells were incubated with 200 µL

of methyl-3H-thymidine (0.5 µCi/well) for 4 h at 37ºC. The medium was removed

and cells were fixed by incubation in 10% TCA for 1 h at 4ºC. Cells were then

washed twice with 10% TCA to remove unbound radioactivity. Plates were air-

dried and cells were lysed with 1 M NaOH (280 µL/well) as described(26). A 250 µL

aliquot of the lysate was neutralized with 5 M HCl prior to the addition of

scintillation fluid. The radioactivity of the samples was quantified in a liquid

scintillation counter and the results are presented in counts per minute (cpm).

Lipid accumulation in adipocytes


Preadipocytes were seeded (1 × 104 cells in 500 μL per well, 24-well cell

culture plates) in wells precoated with 0.2% gelatin. Cells were treated with MTX

(0.1 and 10 µM) during cellular differentiation from day 0 to day 12 (Figure 3).

Differentiation medium was added in the presence of MTX or vehicle (NaOH 0.1

mM) and was replaced two days later by growth medium containing INS and MTX,

which was renewed every 2-3 days. Twelve days after the beginning of

differentiation, the culture medium was removed and cells were washed with PBS

and fixed with p-formaldehyde (3.7% (v/v) in PBS) for 1 h at room temperature.

The cells were stained with 0.3% oil red O solution in isopropanol:distilled water

(3:2) for 15 minutes as described(24). The culture plate was washed 4 times with

distilled water and air-dried. Oil red O in the cells was recovered in 150 μL of

DMSO, and the absorbance measured at 492 nm with reference at 650 nm on a

plate reader (Thermo Electron Corporation, Multiskan Ascent, U.S.A.). Results

were normalized to cell number counted through the trypan blue method.(26)

Measurement of 3H-deoxyglucose uptake

Preadipocytes were seeded, differentiated and treated as detail above. On

day 12 of differentiation, the culture medium was removed and cells were washed

with 0.3 mL glucose-free HEPES buffered saline (140 mM NaCl, 5 mM KCl, 2.5

mM MgSO4, 1 mM CaCl2, 1 mM NaH2PO4.H2O, 4 mM Na2HPO.2H2O, 20 mM

HEPES, pH 7.4) at 37ºC. Cells were then pre-incubated for 30 minutes with 0.3

mL glucose-free HEPES buffered saline in the presence or absence of 0.1 nM

insulin. Glucose uptake was initiated by the addition of 0.3 mL buffer at 37ºC

containing 1 µM 3H-deoxyglucose, according to the procedure described by Faria


et al(27). After 10 minutes, incubation was stopped by washing adipocytes with 0.3

mL ice-cold buffer. The cells were then solubilized with 0.3 mL 0.1% (v/v) Triton X-

100 (in 5 mM Tris-HCl, pH 7.4), and placed overnight at room temperature.

Radioactivity was afterwards measured by liquid scintillation counting and total

protein amount was determined as described by Bradford to correct results for

total culture proteins(28).

Determination of cell viability

Cell viability was assessed by measuring lactate dehydrogenase (LDH)

activity, as described by Bergmeyer and Bernt(29). Preadipocytes were treated with

MTX or vehicle, 24 h or 48 h after seeding in 24-well cell culture plates. At the end

of each treatment, 50 µL of culture medium were collected to a 96-well culture

plate to determine extracellular LDH activity through the addition of 250 µL of

reagent solution (11.3 mM β-NADH-Na2, 50 mM phospate, 0.63 mM piruvate, pH

7.4). To determine intracellular LDH activity, cells were then washed with 1 mL

PBS and lysed with 0.3 mL 0.1% (v/v) Triton X-100 (in 5 mM Tris-HCl, pH 7.4) for

30 minutes at 37ºC. Lactate dehydrogenase activity was determined by measuring

the oxidation of NADH at a wavelength of 340 nm during the reduction of pyruvate

to lactate. Absorvance values were determined for 2 min, and the rate of NADH

reduction was calculated. Adipocyte viability was also evaluated as well after the

12 days of differentiation and treatment following the same protocol.

Determination of homocysteine released to the culture medium


Homocysteine concentration was measured in culture media before and

after treatments.

After 24 h and 48 h of preadipocyte treatment with vehicle or MTX, the

incubation medium was used to determine total homocysteine released. After 12

days of adipocyte differentiation with MTX or vehicle the incubation medium was

also used for total homocysteine determination. Homocysteine quantification was

performed by Clinical Biochemistry Department of S. João Hospital using an

automatic method that followed the guidelines(30).

The values obtained in the culture media (in µmol/L), after have been

incubated with the cells, were normalized for the available indexes of cell density

in the culture. In the case of preadipocytes, homocysteine concentration were

normalized for methyl-thymidine-3H incorporation, where as homocysteine in

adipocyte-conditioned media was normalized for the number of cells counted by

trypan blue method.

Statistical analysis

Results are presented as arithmetic means and standard error of mean

(SEM) of three independent experiments with measurements at least in triplicate.

Statistical significance of the difference between various groups was evaluated by

one-way analysis of variance (ANOVA test) followed by Bonferroni test (in Graph

Pad Prism 5.0 software, U.S.A.). Student’s t test was used for comparison

between two groups. Whenever variances differed significantly, the correspondent

non-parametric tests were chosen (Kruskal-Wallis followed by Dunn’s multiple

comparison test for comparison of three or more groups and Mann Whitney’s test


for comparison of two groups). Differences between means were considered

significant when P < 0.05.


Results

Preadipocytes were used for proliferation assays and adipocytes for lipid

accumulation and glucose transport studies. Cellular viability was determined both

in preadipocytes and adipocytes after treatment with MTX. The concentrations

chosen for MTX treatment of preadipocytes and adipocytes were based on

previous investigations(31, 32).

3T3-L1 preadipocyte proliferation

Proliferation of 3T3-L1 cells after treatment with different concentrations of

MTX was determined by SRB staining and methyl-3H-thymidine incorporation into

cell culture DNA. In SRB protein staining assay, the dye binds to basic amino

acids of cellular proteins and colorimetric evaluation provides an estimate of total

protein mass which is related to cell number(25). Culture protein content after

treatment was compared with the initial amount of protein (after 24 h of plating)

and this value was considered the baseline. An increase in the number of cells in

comparison to this time point appears graphically as a positive value and a

decrease as a negative value. The percentage control (using values of control

cells grown another 24 h after beginning of treatments) were then calculated and

are displayed in Figure 4.

Cell culture protein content was significantly decreased by 0.1 µM MTX

treatment for 24 h (to 10.33 ± 10.75% of control) and 48 h (to 50.74 ± 10.62% of

control) as compared to control cells (absorvance: 0.071 ± 0.007). For the highest

concentration of MTX used (10 µM) after 24 h or 48 h of treatment, there was even

a lower protein content in the culture than at the beginning of treatment (-31.20 ±


4.880% and -114.6 ± 16.30% of control, respectively). This result indicates that

MTX at 10 µM was cytotoxic to preadipocytes.

Figure 4 - Culture protein content after treatment with methotrexate (MTX). Preadipocytes were

treated with MTX 0.1 µM (MTX0.1), MTX 10 µM (MTX10) or vehicle (C, 0.1 M NaOH) for 24 h or 48

h, 24 h after seeding. At the end of treatment, cells were fixed with trichloroacetic acid and stained

with sulforhodamine B. Bound dye was extracted with Tris solution (10 mM, pH 10.5) and

absorvance was measured at 492 nm with reference at 650 nm. Bars represent means and vertical

lines are one SEM. ***P < 0.0001 vs respective control, #P < 0.05 and

# #P < 0.01 between columns

(n = 24).

The other method used to evaluate cellular proliferation consisted in

measuring the incorporation of labeled thymidine nucleoside into cellular DNA

during cellular cycle S phase. Since DNA synthesis is a requisite for cell

proliferation, incubation of cells with methyl-3H-thymidine allows incorporation of

the tritiated nitrogen base into cells when cells replicate their DNA and divide(26).

Comparatively to the cells treated with vehicle for 24 h (15295 ± 1320 cpm)

and 48 h (21408 ± 2632 cpm), 10 μM MTX reduced methyl-3H-thymidine

incorporation significantly after 24 h (5261 ± 714.2 cpm) or 48 h (7777 ± 1904

cpm) of treatment (Figure 5). On the other hand, 0.1 μM MTX had no effect on

C

MTX

0.1

MTX

10

-200

-100

0

100

200

300

40024 h

48 h

*********

***

# #

# # #

Cu

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on

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methyl-3H-thymidine incorporation after 24 h of treatment but reduced cell

proliferation when cells were incubated 48 h with the compound (19133 ± 2097

cpm).

Figure 5 - Methyl-3H-thymidine incorporation into cellular DNA after treatment with methotrexate

(MTX). Preadipocytes seeded on 24-well plates were treated with MTX 0.1 µM (MTX0.1), MTX 10

µM (MTX10) or vehicle (C, 0.1 M NaOH) for 24 h or 48h, 24 h after seeding. At the end of

treatment, cells were incubated with methyl-3H-thymidine (0.5 µCi/well) at 37ºC, for 4 h, washed

and fixed with trichloroacetic acid, lysed with NaOH and incorporated methyl-3H-thymidine was

measured in cell lysates through liquid cintilometry. Bars represent means and vertical lines are

one SEM. **P < 0.001 and ***P < 0.0001 vs respective control and #P < 0.05 between columns (n =

18).

Lipid accumulation in adipocytes

On the first 4-7 days after preadipocyte differentiation induction, in the

cytoplasm of maturing adipocytes there are multiple small lipid droplets that tend

to coalesce into bigger droplets as differentiation proceeds. At day 12 of

differentiation process, cells were stained with oil red O to assess lipid

accumulation, as an index of the degree of differentiation(33).

C

MTX

0.1

MTX

10

0

10000

20000

3000024 h

48 h

***

***

**

#

Me

thyl-

3H

-th

ym

idin

e i

nc

orp

ora

tio

n

(co

un

ts p

er

min

ute

)


The quantitative spectrophotometric analysis of cellular lipid content

revealed that treatment of adipocytes with MTX resulted in a significant,

concentration-dependent, higher oil red O incorporation (162.5 ± 9.3% of control

and 501.6 ± 17.2% of control respectively) in comparison to vehicle-treated cells

(absorvance: 0.014 ± 0.001 / 104 cells) (Figure 6).

Figure 6 - Lipid accumulation in adipocytes after treatment with methotrexate (MTX). Cells were

treated with MTX 0.1 µM (MTX0.1), MTX 10 µM (MTX10) or vehicle (C, 0.1 M NaOH) dissolved in

culture medium since the induction of differentiation. At day 12 of the differentiation protocol, cells

were fixed with p-formaldehyde (3.7% (v/v)) and stained with oil red O (0.3% in

isopropanol:destiled water, 3:2). Incorporated dye was extracted with dimethyl sulfoxide and

absorvance was measured at 492 nm with reference at 650 nm. Results were normalized to cell

number counted through the trypan blue method. Bars represent means and vertical lines are one

SEM. **P < 0.001 and ***P < 0.0001 vs control (n = 18).

Measurement of 3H-deoxyglucose uptake

Adipocytes are cells that can express glucose transporters (GLUT) - 1 and

4 in their plasma membrane. GLUT1 transporters have high affinity to glucose and

their action is not insulin-dependent. GLUT4 transporters are stored in intracellular

C

MTX

0.1

MTX

10

0

200

400

600

**

***

Lip

id a

cc

um

ula

tio

n

(% o

f co

ntr

ol)


sites and their translocation to the plasma membrane is determined by insulin

stimulus(34). The effect of MTX treatment on glucose uptake in the presence or

absence of insulin was evaluated by measuring the uptake of 3H-DG (Figure 7).

MTX 0.1 µM treatment had no effect on 3H-DG basal uptake but the highest

concentration of MTX (10 µM) used significantly increased 3H-DG basal uptake (to

167.7 ± 11.1 % of control). Addition of insulin to vehicle-treated cells resulted in a

significant increase of 3H-DG uptake (from 911.5 ± 64.8 fmol/mg of protein to

1122.74 ± 46.05 fmol/mg of protein after insulin treatment). This is in contrast to

what was observed in 0.1 µM MTX- and 10 µM MTX-treated adipocytes

suggesting that MTX-treated cells are resistant to insulin stimulation, at least in

what concerns glucose uptake.

Figure 7 - 3H-Deoxyglucose (

3H-DG) uptake after treatment with methotrexate (MTX) in the

absence (basal) or presence (stimulated) of insulin. Cells were treated with MTX 0.1 µM (MTX0.1),

MTX 10 µM (MTX10) or vehicle (C, 0.1 M NaOH) dissolved in culture medium since the induction

of differentiation. At day 12 of the differentiation protocol, cells were pre-incubated with glucose-

free HEPES buffered saline in the absence or presence of 0.1 nM insulin. Afterwards, cells were

incubated with 3H-deoxyglucose for 10 minutes at 37ºC and

3H-deoxyglucose incorporation was

C

MTX

0.1

MTX

10

0

50

100

150

200Basal

Stimulated#

***

3H

-DG

up

take

(% o

f co

ntr

ol)


measured by liquid scintillation counting. Bars represent means and vertical lines are one SEM.

***P < 0.0001 vs respective control and #P < 0.05 between columns (n = 12).

Determination of cell viability

LDH is a cytoplasmic enzyme which catalyses the conversion of pyruvate to

lactate in the presence of NADH. Its presence in the extracellular medium gives

information about membrane integrity and cell viability. Since released LDH is a

stable enzymatic marker that correlates linearly with cell viability(35) the MTX

cytotoxic effects on preadipocytes and adipocytes were evaluated by measuring

the activity of this enzyme.

In preadipocytes, results suggest that 0.1 µM MTX treatment did not have

cytotoxic effects, since there were no differences between intracellular-to-

extracellular LDH ratio in treated and control cells (4.426 ± 0.417 intracellular-to-

extracellular LDH ratio in control cells) (Figure 8). On the other hand, the highest

concentration of MTX used (10 µM) reduced cell viability significantly after 24 h or

48 h of treatment (to 58.19 ± 2.31% and to 37.18 ± 3.23% of control at 24 h and 48

h, respectively). In adipocytes, MTX treatment, which begun 2 days after cells

reached confluence, did not reduce cell viability in any of the concentrations used,

suggesting that its cytotoxic effects may be restricted to proliferating cells.


Figure 8 - Cell viability in preadipocytes (A) and adipocytes (B) after treatment with methotrexate.

Cells were treated with MTX 0.1 µM (MTX0.1), MTX 10 µM (MTX10) or vehicle (C, 0.1 M NaOH)

24 h or 48 h after seeding (A) or 12 days after differentiation induction (B). Extracellular and

intracellular lactate dehydrogenase activity was determined by measuring the oxidation of NADH at

a wavelength of 340 nm during the reduction of pyruvate to lactate. Bars represent means and

vertical lines are one SEM. ***P < 0.0001 vs respective control (n = 12 and n = 8 on experiments

using preadipocytes and adipocytes, respectively).

Determination of homocysteine released to culture medium

After 24 h, 0.1 or 10 µM MTX treatment significantly increased

homocysteine release to the culture medium (to 131.1 ± 5.7 and to 314.3 ± 15.7%

A

B

C

MTX

0.1

MTX

10

0

50

100

150

200

25024 h

48 h

******

Intr

ac

ell

ula

r-to

-ex

tra

ce

llu

lar

LD

H r

ati

o

(% o

f co

ntr

ol)

C

MTX

0.1

MTX

10

0

50

100

150

Intr

ac

ell

ula

r-to

-ex

tra

ce

llu

lar

LD

H r

ati

o

(% o

f co

ntr

ol)


of control respectively) in comparison with cells treated with vehicle (0.121 ± 0.010

nM/cpm) (Table 1). After 48 h both concentrations of MTX used caused a

significant increase on homocysteine release (to 179.3 ± 2.2 and to 164.8 ± 5.3%

of control in 0.1 µM and 10 µM MTX-treated cells, respectively).

In adition, total homocysteine was also increased from 179 ± 0.0 nM/ 104

cell in control cells to 390.2 ± 0.04 nM/ 104 cell in the culture medium of 10 µM

MTX treated-adipocytes.

Culture medium homocysteine concentration was below the detection level

of the method (< 1 µmol/L).

C MTX 0.1 µM MTX 10 µM

Preadipocytes 24 h 100.00 131.12 314.30 **

Preadipocytes 48 h 129.37 179.33 **,# 164.78 *

,#

Adipocytes 12 days 100.00 120.98 218.35*

Table 1 – Total homocysteine released to the culture medium in % of control. After 24 h and 48 h

of preadipocyte treatment and after 12 days of adipocyte differentiation and treatment with

vehicle (C, 0.1 M NaOH) or MTX, culture medium was collected and used to determine

homocysteine release. Results were normalized for cell number measured by methyl-3H-

thymidine incorporation and trypan blue method on experiments using preadipocytes and

adipocytes, respectively. *P < 0.05 and **P < 0.01 vs respective control and #P < 0.05 vs

preadipocytes at 24 h (n = 2).


Discussion

Research on the relationship between folate and obesity or the metabolic

syndrome includes only a few studies to date but there is evidence supported by

epidemiological investigations that low levels of plasma folate are associated with

increased BMI(36-38) and body fat percentage(15). Obesity and the associated

metabolic pathologies are the most common and detrimental metabolic

diseases(39). They are associated with a chronic inflammatory response

characterized by abnormal cytokine production and by activation of inflammatory

signaling pathways, which appears to be triggered in adipose tissue(39). Adipose

tissue growth can be a result from an increase in adipocyte number (hyperplasia),

or from an increase in adipocyte volume due the incorporation of triglycerides

(TAG) (hypertrophy), or from both(40). However, it seems that if energy is stored

predominantly through lipogenesis resulting in adipocyte hypertophy instead of

adipogenesis with recruitment and differentiation of new adipocytes and

hyperplasia, adipose tissue responses can be pathological contributing to

metabolic disease(41).

According to the new understanding of obesity therapeutic strategies with

the aim of reducing associated metabolic complications, adipocyte hypertrophy

and dysfunction could be prevented if adipose tissue preserves the ability to recruit

preadipocytes to differentiate, so that fat content can be distributed among the

new adipocytes(42). There is also evidence that the opposite also applies, i.e. when

there is a decrease in the ability to recruit new preadipocytes, obesity

complications may be precipitated(43). Here, we demonstrate that, when folate

availability is compromised by MTX treatment preadipocyte proliferation is

inhibited as shown by methyl-3H-thymidine incorporation and SRB staining


experiments. This decrease in preadipocyte number can limit the recruitment of

these cells to differentiate. In SRB assay, after 24 h or 48 h of 10 µM MTX

treatment, there was a lower protein content in the culture than at the beginning of

treatment. This suggested that MTX at 10 µM concentration was cytotoxic to

preadipocytes. To confirm these results, we determined cell viability after 24 h and

48 h of 10 µM MTX treatment, by LDH activity determination. In fact, 10 µM MTX

treatment caused cell death and reduced cell viability at 24 h and 48 h. Similarly,

Savion et al. suggested that the lower survival rate of embryonic fibroblasts in

response to MTX might be caused, at least partially, by a decrease in cell

proliferation, which might be attributed to the bax protein, an inducer of apoptosis

that enhances the permeability of the mitochondrial membrane and the release of

cell death proteins(32). On the other hand, 0.1 µM MTX did not exert cytotoxic

effects on preadipocytes but did reduce 3T3-L1 fibroblast proliferation measured

by SRB. The results regarding methyl-3H-tymidine incorporation also showed a

reduction in preadipocyte proliferation by MTX in a time- and concentration-

dependent manner. This reduction can be explained through the inhibition of

DHFR and TS by methotrexate, causing cessation of the purine and pyrimidine

synthesis which are important for cell proliferation(44).

Another interesting result from the present work concerns lipid incorporation

into adipocytes measured by oil red O. MTX treatment of adipocytes resulted in a

higher (concentration-dependent) lipid accumulation, suggesting a stimulation of

preadipocyte differentiation when folate availability was compromised. The fact

that low cellular folate availability may impair de novo biosynthesis of purines,

leading to AICAR and adenosine accumulation, may help explain this result. In

vitro and in vivo experimental studies show that MTX increases adenosine release


by fibroblasts(9). MTX inhibits AICAR transformylase, which catalyzes one of the

final stages of de novo purine synthesis. Inhibition of AICAR transformylase leads

to AICAR accumulation. This results, in turn, in a net increase in intra- and

extracellular adenosine(45), since adenosine deaminase, an enzyme responsible

for adenosine degradation, is also inhibited (Figure 9). Adenosine receptors (ARs)

are widely distributed in tissues and modulate numerous physiological

processes(46). A1 ARs are highly expressed in the adipose tissue, and it has been

shown that endogenous adenosine can tonically activate adipose tissue A1 ARs,

causing inhibition of lipolysis(47).

Figure 9 - Methotrexate increases adenosine accumulation via 5-amino-4-imidazolecarboxamide

ribonucleotide (AICAR) transformylase inhibition. THF, tetrahydrofolate; MTX, methotrexate.

Adapted from Dhalla et al(47)

.

Adenosine

AICAR formyl-AICAR

Purines

DNA

10-formylTHF + AICAR transformylase

Adenosine Degradation

AICAR

MTX

Adenosine deaminase


Lipolysis is the process by which TAG are hydrolysed into free fatty acids

and glycerol. Adipocyte lipolysis is activated by catecholamines that bind to GS

protein-coupled receptors activating adenyl cyclase. This stimulation leads to an

increase in intracellular cAMP concentration, resulting in activation of cAMP-

dependent protein kinase A (PKA). PKA phosphorylates hormone-sensitive lipase

(HSL), leading to increased cellular TAG hydrolytic activity(48), and perilipin, the

lipid droplet-coating protein that when activated allows TAG breakdown by HSL.

When adenosine binds to A1 ARs in adipocytes, it inhibits adenylyl cyclase

and cAMP production, causing inhibition of lipolysis and increasing adipocyte TAG

accumulation. This adenosinergic mechanism of action may underline the results

obtained with the oil red O assay. Promoting adenosine accumulation and release

by adipocytes, MTX directly, and also indirectly, through the induction of a low

folate availability, may exert antilipolytic effects leading to increased lipid

accumulation in adipocytes, in a concentration-dependent manner. In the same

line, Lam et al., described that folic acid supplementation has significant

modulatory effects on lipolysis and results in an improvement of both baseline and

beta-adrenoceptor-mediated lipolysis in +db/+db mice(49). Therefore, it is

suggested that folate can contribute to decreased lipid accumulation both by

inhibiting A1 ARs receptor stimulation but also by stimulating adrenoceptor-

mediated lipolysis.

Adipocytes can also re-esterify fatty acids and synthesize TAG from

glucose or other energy sources acquired in the diet through a process called de

novo lipogenesis(50). The results obtained in 3H-DG uptake suggest that 10 µM

MTX treatment stimulates basal glucose uptake in adipocytes (and that 0.1 µM

MTX treatment induces the same trend), probably favoring de novo lipogenesis


and contributing to higher lipid accumulation in these cells. Given the importance

of GLUT1 on adipocyte basal glucose transport, a stimulation of this membrane

transporter by MTX or folate deficiency could explain these results.

Several enzymes involved in adipose tissue lipogenesis, like fatty acid

synthase and acetyl coenzyme A carboxylase, contributing to TAG synthesis from

glucose, are induced by insulin(50). In addition, GLUT4, responsible for increasing

glucose entry to adipocytes after translocation to the plasma membrane, is also

stimulated by insulin. Thus, an increase in 3H-DG uptake after adipocyte

incubation with insulin, was expected. However, incubation with insulin after MTX

treatment did not result in a increased 3H-DG uptake, as opposite to what was

observed in control cells, suggesting that MTX treated adipocytes were resistant to

insulin stimulation. Mechanisms that could explain what cause this resistance to

insulin stimulus, deserve further investigation. The influence of folate on insulin

levels was recently postulated by Setola et al. who reported that prolonged folate

treatment in patients with metabolic syndrome not only decreased homocysteine

levels, but also reduced insulin levels, improving insulin resistance(52). In addition,

Golbahar et al. reported that hyperhomocysteinemia, also associated with low

plasma folate levels, induced insulin resistance in male Sprague Dawley rats(53).

These reports prompted us to determinate homocysteine released to the culture

medium of preadipocytes and adipocytes, before and during MTX treatment.

Homocysteine is a sulphur-containing amino acid derived from the

conversion of methionine to cysteine and has been regarded as an independent

risk factor for atherosclerotic cardiovascular, since elevated plasma levels (> 15

µmol/L) are associated with the pathogenesis of atherosclerosis and

cardiovascular disease(54). Folate deficiency can increase plasma homocysteine


levels, since the conversion of 5-methylTHF into THF provides the methyl group

required for homocysteine metabolism through the remethylation pathway(10). This

is well demonstrated by our results, showing that when folate availability was

compromised by MTX treatment, homocysteine in the culture medium of

preadipocytes was increased at 24 h and at 48 h after treatment for both

concentrations of MTX used. After 12 days of differentiation under folate

deprivation due the presence of MTX, adipocytes released increased amounts of

homocysteine to the culture medium in a concentration-dependent manner.

Riederer et al. had already found that progressive differentiation of 3T3-L1 cells

resulted in increased homocysteine concentration in cell culture supernatants(55).

They attributed this finding to N-nicotinamide metyltransferase, which catalyses

the reaction of nicotinamide to methyl-nicotinamide using the methyl group

provided by S-adenosyl-methionine (SAM) conversion into S-adenosyl-

homocysteine (SAH) (Figure 2). SAH is then converted into homocysteine by S-

adenosyl-homocysteine hydrolase, increasing through this pathway the release of

the atherogenic metabolite in the culture medium(55). In our study, N-nicotinamide

metyltransferase of adipocytes could have used the nicotinamide present in the

culture medium (4 mg/L) to produce methyl-nicotinamide and thus release

homocysteine. Since homocysteine metabolism is compromised in MTX-treated

cells due to folate deprivation, differences between homocysteine concentration in

culture media of treated and untreated cells could be explained. Accordingly, we

show that preadipocytes and adipocytes can be a source of homocysteine and

that the release of this metabolite is dependent of folate status. However, further

investigation is necessary to understand the relationship between adipocyte de


novo lipogenesis, insulin resistance and homocysteine to understand the role of

folate in these processes.


Conclusion

In our study, we present the possibility that low cellular folate availability

may result in a higher accumulation of lipids in adipocytes which may denote that

there was an increase of the differentiation of preadipocytes into adipocytes. This

can be due either to inhibition of lipolysis or to stimulation of de novo lipogenesis,

two important processes involved on the regulation of intracellular lipid content.

Furthermore, we show that there was a simultaneous decrease in preadipocyte

proliferation when folate availability is compromised, which can contribute to

adipocyte hypertrophy and dysfunction, since fewer cells would be present to

accommodate nutrient surplus. In addition, we demonstrate that folate deprivation

can make adipocytes resistant to insulin stimulation, as demonstrated in glucose

transport studies. It can also increase homocysteine released by these cells, which

supports the idea that the adipose tissue may be a source of homocysteine,

especially when there is low folate availability.

In conclusion, our results suggest that folate deprivation can interfere with

adipocyte proliferation, differentiation and metabolism and promote the

hypertrophic growth of adipocytes, which may contribute to the development of

obesity complications and the metabolic syndrome.


References

1. Assaraf YG. Molecular basis of antifolate resistance. Cancer Metastasis

Rev. 2007; 26(1):153-81.

2. Rampersaud GC, Kauwell GPA, Bailey LB. Folate: A Key to Optimizing

Health and Reducing Disease Risk in the Elderly. J Am Coll Nutr. 2003; 22(1):1-8.

3. Shane B. Folate and vitamin B12 metabolism: overview and interaction with

riboflavin, vitamin B6, and polymorphisms. Food And Nutrition Bulletin. 2008; 29(2

Suppl):S5.

4. Depeint F, Bruce WR, Shangari N, Mehta R, O'Brien PJ. Mitochondrial

function and toxicity: Role of B vitamins on the one-carbon transfer pathways.

Chemico-Biological Interactions. 2006; 163(1-2):113-32.

5. Katula KS, Heinloth AN, Paules RS. Folate deficiency in normal human

fibroblasts leads to altered expression of genes primarily linked to cell signaling,

the cytoskeleton and extracellular matrix. The Journal of Nutritional Biochemistry.

2007; 18(8):541-52.

6. Hubner RA, Houlston RS. Folate and colorectal cancer prevention. Br J

Cancer. 2009; 100(2):233-9.

7. Gregorio V-M, Michelle MM, John MS. Cobalamin, folic acid, and

homocysteine. Nutrition Reviews. 2009; 67(s1):S69-S72.

8. Matherly L, Hou Z, Deng Y. Human reduced folate carrier: translation of

basic biology to cancer etiology and therapy. Cancer and Metastasis Reviews.

2007; 26(1):111-28.

9. Swierkot J, Szechinski J. Methotrexate in rheumatoid arthritis. Pharmacol

Rep. 2006; 58(4):473-92.


10. Wierzbicki AS. Homocysteine and cardiovascular disease: a review of the

evidence. Diabetes and Vascular Disease Research. 2007; 4(2):143-49.

11. Ntaios G, Savopoulos C, Grekas D, Hatzitolios A. The controversial role of

B-vitamins in cardiovascular risk: An update. Archives of Cardiovascular Diseases.

2009; 102(12):847-54.

12. Onat A, Hergenç G, Küçükdurmaz Z, Can G, Ayhan E, Bulur S. Serum

folate is associated with coronary heart disease independently of homocysteine in

Turkish men. Clinical Nutrition. 2008; 27(5):732-39.

13. Wolff T, Witkop CT, Miller T, Syed SB. Folic Acid Supplementation for the

Prevention of Neural Tube Defects: An Update of the Evidence for the U.S.

Preventive Services Task Force. Annals of Internal Medicine. 2009; 150(9):632-

39.

14. Mojtabai R. Body mass index and serum folate in childbearing age women.

Eur J Epidemiol. 2004; 19(11):1029-36.

15. Mahabir S, Ettinger S, Johnson L, Baer DJ, Clevidence BA, Hartman TJ, et

al. Measures of adiposity and body fat distribution in relation to serum folate levels

in postmenopausal women in a feeding study. Eur J Clin Nutr. 2007; 62(5):644-50.

16. Hirsch S, Poniachick J, Avendano M, Csendes A, Burdiles P, Smok G, et al.

Serum folate and homocysteine levels in obese females with non-alcoholic fatty

liver. Nutrition. 2005; 21(2):137-41.

17. Almeda-Valdes P, Cuevas-Ramos D, Aguilar-Salinas CA. Metabolic

syndrome and non-alcoholic fatty liver disease. Ann Hepatol. 2009; 8 Suppl 1:S18-

24.

18. Ortega RM, Oacute, Pez-Sobaler AM, Andr, Eacute, S P, et al. Folate

Status in Young Overweight and Obese Women: Changes Associated with Weight


Reduction and Increased Folate Intake. Journal of Nutritional Science and

Vitaminology. 2009; 55(2):149-55.

19. Stein CJ, Colditz GA. The Epidemic of Obesity. Journal of Clinical

Endocrinology and Metabolism. 2004; 89(6):2522-25.

20. Galic S, Oakhill JS, Steinberg GR. Adipose tissue as an endocrine organ.

Molecular and Cellular Endocrinology. 2009; 316(2):129-39.

21. Okada T, Nakai M, Maeda H, Hosokawa M, Sashima T, Miyashita K.

Suppressive Effect of Neoxanthin on the Differentiation of 3T3-L1 Adipose Cells.

Journal of Oleo Science. 2008; 57(6):345-51.

22. Do G-M, Choi M-S, Kim H-J, Woo M-N, Lee M-K, Jeon S-M. Soy pinitol acts

partly as an insulin sensitizer or insulin mediator in 3T3-L1 preadipocytes. Genes

& Nutrition. 2008; 2(4):359-64.

23. Horie T, Ono K, Kinoshita M, Nishi H, Nagao K, Kawamura T, et al. TG-

interacting factor is required for the differentiation of preadipocytes. J Lipid Res

2008; 49(6):1224-34.

24. Teixeira D, Pestana D, Faria A, Calhau C, Azevedo I, Monteiro R.

Modulation of Adipocyte Biology by Delta(9)-Tetrahydrocannabinol. Obesity (Silver

Spring). Epub 2010;

25. Papazisis KT, Geromichalos GD, Dimitriadis KA, Kortsaris AH. Optimization

of the sulforhodamine B colorimetric assay. Journal of Immunological Methods.

1997; 208(2):151-58.

26. Miranda CL, Stevens JF, Helmrich A, Henderson MC, Rodriguez RJ, Yang

YH, et al. Antiproliferative and cytotoxic effects of prenylated flavonoids from hops

(Humulus lupulus) in human cancer cell lines. Food Chem Toxicol. 1999;

37(4):271-85.


27. Faria A, Pestana D, Azevedo J, Martel F, de Freitas V, Azevedo I, et al.

Absorption of anthocyanins through intestinal epithelial cells - Putative involvement

of GLUT2. Mol Nutr Food Res. 2009; 53(11):1430-7.

28. Bradford MM. A rapid and sensitive method for the quantitation of

microgram quantities of protein utilizing the principle of protein-dye binding. Anal

Biochem. 1976; 72:248-54.

29. Bergmeyer HU, Bernt E. Lactate dehydrogenase. In: Bergmeyer, H.U., ed.

Methods in Enzimatic Analysis. New York: Academic Press.1974. pp 574-9.

30. Recommended methods for the determination of four enzymes in blood.

Scand J Clin Lab Invest. 1974; 33(4):291-306.

31. Brengauz-Breitmann M, Friedman E, Savion S, Torchinsky A, Fein A, Toder

V. Involvement of NF-[kappa]B in the response of embryonic cells to Methotrexate.

Reproductive Toxicology. 2006; 22(3):469-78.

32. Savion S, Shtelman E, Orenstein H, Torchinsky A, Fein A, Toder V. Bax-

associated mechanisms underlying the response of embryonic cells to

methotrexate. Toxicology in Vitro. 2009; 23(6):1062-68.

33. Manickam E, Sinclair A, Cameron-Smith D. Suppressive actions of

eicosapentaenoic acid on lipid droplet formation in 3T3-L1 adipocytes. Lipids in

Health and Disease. 2010; 9(1):57.

34. Huang S, Czech MP. The GLUT4 Glucose Transporter. Cell Metabolism.

2007; 5(4):237-52.

35. Monteiro R, Calhau C, Martel F, Faria A, Mateus N, Azevedo I. Modulation

of MPP+ uptake by tea and some of its components in Caco-2 cells. Naunyn

Schmiedebergs Arch Pharmacol. 2005; 372(2):147-52.


36. Casanueva E, Drijanski A, Fernández-Gaxiola AC, Meza C, Pfeffer F.

Folate deficiency is associated with obesity and anemia in Mexican urban women.

Nutrition Research. 2000; 20(10):1389-94.

37. Karatela RA, Sainani GS. Plasma homocysteine in obese, overweight and

normal weight hypertensives and normotensives. Indian Heart J. 2009; 61(2):156-

9.

38. Gallistl S, Sudi K, Mangge H, Erwa W, Borkenstein M. Insulin is an

independent correlate of plasma homocysteine levels in obese children and

adolescents. Diabetes Care. 2000; 23(9):1348-52.

39. Wellen KE, Hotamisligil GS. Obesity-induced inflammatory changes in

adipose tissue. J Clin Invest. 2003; 112(12):1785-8.

40. de Ferranti S, Mozaffarian D. The perfect storm: obesity, adipocyte

dysfunction, and metabolic consequences. Clin Chem. 2008; 54(6):945-55.

41. Bays HE, Gonzalez-Campoy JM, Bray GA, Kitabchi AE, Bergman DA,

Schorr AB, et al. Pathogenic potential of adipose tissue and metabolic

consequences of adipocyte hypertrophy and increased visceral adiposity. Expert

Rev Cardiovasc Ther. 2008; 6(3):343-68.

42. Sethi JK, Vidal-Puig AJ. Thematic review series: Adipocyte Biology.

Adipose tissue function and plasticity orchestrate nutritional adaptation. J Lipid

Res. 2007; 48(6):1253-62.

43. Tchoukalova YD, Koutsari C, Karpyak MV, Votruba SB, Wendland E,

Jensen MD. Subcutaneous adipocyte size and body fat distribution. Am J Clin

Nutr. 2008; 87(1):56-63.

44. Joel MK. Toward a better understanding of methotrexate. Arthritis and

Rheumatism. 2004; 50(5):1370-82.


45. Bangert CA, Costner MI. Methotrexate in dermatology. Dermatol Ther.

2007; 20(4):216-28.

46. Hutchinson SA, Scammells PJ. A(1) adenosine receptor agonists: medicinal

chemistry and therapeutic potential. Curr Pharm Des. 2004; 10(17):2021-39.

47. Dhalla AK, Chisholm JW, Reaven GM, Belardinelli L. A1 adenosine

receptor: role in diabetes and obesity. Handb Exp Pharmacol. 2009; (193):271-95.

48. Jaworski K, Sarkadi-Nagy E, Duncan RE, Ahmadian M, Sul HS. Regulation

of triglyceride metabolism. IV. Hormonal regulation of lipolysis in adipose tissue.

Am J Physiol Gastrointest Liver Physiol. 2007; 293(1):G1-4.

49. Lam TY, Seto SW, Au AL, Poon CC, Li RW, Lam HY, et al. Folic acid

supplementation modifies beta-adrenoceptor-mediated in vitro lipolysis of

obese/diabetic (+db/+db) mice. Exp Biol Med (Maywood). 2009; 234(9):1047-55.

50. Vazquez-Vela ME, Torres N, Tovar AR. White adipose tissue as endocrine

organ and its role in obesity. Arch Med Res. 2008; 39(8):715-28.

51. Lee H, Kim HJ, Kim J-m, Chang N. Effects of dietary folic acid

supplementation on cerebrovascular endothelial dysfunction in rats with induced

hyperhomocysteinemia. Brain Research. 2004; 996(2):139-47.

52. Setola E, Monti LD, Galluccio E, Palloshi A, Fragasso G, Paroni R, et al.

Insulin resistance and endothelial function are improved after folate and vitamin

B12 therapy in patients with metabolic syndrome: relationship between

homocysteine levels and hyperinsulinemia. Eur J Endocrinol. 2004; 151(4):483-89.

53. Golbahar J, Aminzadeh MA, Kassab SE, Omrani GR.

Hyperhomocysteinemia induces insulin resistance in male Sprague-Dawley rats.

Diabetes Research and Clinical Practice. 2007; 76(1):1-5.


54. Ntaios G, Savopoulos C, Grekas D, Hatzitolios A. The controversial role of

B-vitamins in cardiovascular risk: An update. Arch Cardiovasc Dis. 2009;

102(12):847-54.

55. Riederer M, Erwa W, Zimmermann R, Frank S, Zechner R. Adipose tissue

as a source of nicotinamide N-methyltransferase and homocysteine.

Atherosclerosis. 2009; 204(2):412-7.

Effect of cellular folate availability on adipocyte life ...Effect of cellular folate availability on adipocyte life cycle and metabolism 1 Abstract The role of folate in obesity and

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