LETTER

doi:10.1038/nature10798

Dysfunction of lipid sensor GPR120 leads to obesity inboth mouse and humanAtsuhiko Ichimura1*, Akira Hirasawa1*, Odile Poulain-Godefroy2,3*, Amélie Bonnefond2,3*, Takafumi Hara1, Loı̈c Yengo2,3,
Ikuo Kimura1, Audrey Leloire2,3, Ning Liu1, Keiko Iida1, Hélène Choquet2,3, Philippe Besnard4, Cécile Lecoeur2,3,
Sidonie Vivequin2,3, Kumiko Ayukawa1, Masato Takeuchi1, Kentaro Ozawa1, Maithé Tauber5, Claudio Maffeis6,7,
Anita Morandi2,3,6, Raffaella Buzzetti8, Paul Elliott9, Anneli Pouta10,11, Marjo-Riitta Jarvelin9,10,12, Antje Körner13, Wieland Kiess13,
Marie Pigeyre14,15, Roberto Caiazzo14,16, Wim Van Hul17, Luc Van Gaal18, Fritz Horber19, Beverley Balkau20,21,
Claire Lévy-Marchal22, Konstantinos Rouskas2,3,23, Anastasia Kouvatsi23, Johannes Hebebrand24, Anke Hinney24,
Andre Scherag25, François Pattou14,16, David Meyre2,3,26, Taka-aki Koshimizu27, Isabelle Wolowczuk2,3, Gozoh Tsujimoto1& Philippe Froguel2,3,28

Free fatty acids provide an important energy source as nutrients,
and act as signalling molecules in various cellular processes1–4.
Several G-protein-coupled receptors have been identified as freefatty-acid receptors important in physiology as well as in several
diseases3,5–13. GPR120 (also known as O3FAR1) functions as a
receptor for unsaturated long-chain free fatty acids and has a criticalrole in various physiological homeostasis mechanisms such as
adipogenesis, regulation of appetite and food preference5,6,14–16.
Here we show that GPR120-deficient mice fed a high-fat dietdevelop obesity, glucose intolerance and fatty liver with decreased
adipocyte differentiation and lipogenesis and enhanced hepatic
lipogenesis. Insulin resistance in such mice is associated with
reduced insulin signalling and enhanced inflammation in adipose
tissue. In human, we show that GPR120 expression in adipose tissue
is significantly higher in obese individuals than in lean controls.
GPR120 exon sequencing in obese subjects reveals a deleterious
non-synonymous mutation (p.R270H) that inhibits GPR120 signalling activity. Furthermore, the p.R270H variant increases therisk of obesity in European populations. Overall, this study demonstrates that the lipid sensor GPR120 has a key role in sensing dietary
fat and, therefore, in the control of energy balance in both humans
and rodents.
To investigate the role of GPR120 in metabolism, we examined
GPR120-deficient mice (Supplementary Fig. 1) with respect to lipogenesis, glucose and energy homeostasis. On a normal diet containing 13%
fat, the body weight was similar in both GPR120-deficient and wildtype mice. However, when 5-week-old GPR120-deficient mice were fed
a high-fat diet (HFD) containing 60% fat, their body weight increase
was ,10% higher than that of wild-type mice on a HFD (Fig. 1a). The
difference in HFD-induced body weight gain between wild-type and
GPR120-deficient mice was marked at ,8–10 weeks old and reached aplateau at 13 weeks old. To assess energy expenditure and substrateutilization, we next performed indirect calorimetry on wild-type andmutant mice on a HFD at 9–10 weeks old (Fig. 1b) and 15–16 weeks old(Supplementary Fig. 2a). The young GPR120-deficient mice showeddecreased energy expenditure compared with the young wild-type
mice, particularly during the light/inactive phase (Fig. 1b, left), whereas
older mutant and wild-type mice showed no such a difference(Supplementary Fig. 2a, left). The difference in energy expenditure
between GPR120-deficient and wild-type mice was observed only inthe light phase, indicating that the lack of the GPR120 receptor
primarily affects basal metabolism, especially in young mice. The
decreased energy expenditure might explain the difference we foundin body weight gain between HFD-fed wild-type and mutant young
mice. The lower values of respiratory quotient in mutant mice could be
due to insufficient glucose utility, probably as a result of the decreased
insulin sensitivity. In all experiments, both groups of mice showedsimilar levels of locomotor activity (data not shown).
White adipose tissue (WAT) and liver were substantially heavier in
HFD-fed GPR120-deficient mice (Supplementary Fig. 2b). Plasma
low- and high-density lipoprotein cholesterol levels were significantly
higher in HFD-fed GPR120-deficient mice, and serum alanine aminotransferase levels were substantially increased, indicating abnormalcholesterol metabolism and liver function (Supplementary Table 1).
Microcomputed tomography scanning revealed that 16-week-old
GPR120-deficient mice stored much more fat than did wild type(Fig. 1c). A significant increase in adipocyte size in both epididymal(Fig. 1d) and subcutaneous (Supplementary Fig. 2c) fat was observedin GPR120-deficient mice. Furthermore, the expression of macrophage marker genes (Cd11b (Itgam), Cd68 and F4/80 (Emr1)) andthe number of F4/80-positive cells were markedly enhanced in epididymal tissue from HFD-fed GPR120-deficient mice (Fig. 1e, f).
Moreover, these mice showed liver steatosis and hepatic triglyceride
content was markedly increased (Fig. 1g). Overall, HFD-induced obesityand liver fattiness were more severe in GPR120-deficient mice than inwild type.

1

Department of Genomic Drug Discovery Science, Graduate School of Pharmaceutical Sciences, Kyoto University, 46-29 Yoshida Shimoadachi-cho, Sakyo-ku, Kyoto 606-8501, Japan. 2Centre National dela Recherche Scientifique (CNRS)-Unité mixte de recherche (UMR) 8199, Lille Pasteur Institute, Lille 59000, France. 3Lille Nord de France University, Lille 59000, France. 4Institut National de la Santé et dela Recherche Médicale (Inserm)-UMR U866, Physiologie de la Nutrition, Bourgogne University, AgroSup Dijon, Dijon 21078, France. 5Inserm-U563, Children’s Hospital, Centre Hospitalier Universitaire,
Toulouse 31000, France. 6Regional Centre for Juvenile Diabetes, Obesity and Clinical Nutrition, Verona 37134, Italy. 7Department of Mother and Child, Biology-Genetics, Section of Paediatrics, University of
Verona, Verona 37134, Italy. 8Department of Clinical Sciences, La Sapienza University, Rome 00161, Italy. 9Medical Research Council-HPA Centre for Environment and Health, Department of Epidemiology
and Biostatistics, School of Public Health, St Mary’s campus, Imperial College London, London W2 1PG, UK. 10National Public Health Institute, Biocenter Oulu, University of Oulu, Oulu 90220, Finland.
11
Institute of Clinical Medicine/Obstetrics and Gynecology, University of Oulu, Oulu 90220, Finland. 12Institute of Health Sciences, University of Oulu, Oulu 90220, Finland. 13Center for Pediatric Research,
Department of Women’s & Child Health, University of Leipzig, Leipzig 04317, Germany. 14Inserm-U859, Lille Nord de France University, Lille 59000, France. 15Lille University Hospital, Nutrition, Lille 59000,
France. 16Lille University Hospital, Endocrine Surgery, Lille 59000, France. 17Department of Medical Genetics, University of Antwerp, Antwerp 2610, Belgium. 18Department of Endocrinology, Antwerp
University Hospital, Antwerp 2650, Belgium. 19Department of Surgery and Internal Medicine, Clinic Lindberg, Medical Department, Winterthur 8400, and University of Berne, Berne 3011, Switzerland.
20
Inserm-U780, Centre for research in Epidemiology and Population Health (CRESP), Villejuif 94800, France. 21Paris-Sud 11 University, Orsay 91405, France. 22Inserm-U690, Robert Debré hospital, Paris
75935, France. 23Department of Genetics, Development and Molecular Biology, School of Biology, Aristotle University of Thessaloniki, Thessaloniki 541 24, Greece. 24Department of Child and Adolescent
Psychiatry, University of Duisburg-Essen, Essen 45147, Germany. 25Institute for Medical Informatics, Biometry and Epidemiology, University of Duisburg-Essen, Essen 45122, Germany. 26McMaster
University, Hamilton, Ontario L8S 4L8, Canada. 27Department of Pharmacology, Division of Molecular Pharmacology, Jichi Medical University, Tochigi 329-0498, Japan. 28Department of Genomics ofCommon Disease, School of Public Health, Imperial College London, Hammersmith Hospital, London W12 0NN, UK.
*These authors contributed equally to this work.
3 5 0 | N AT U R E | VO L 4 8 3 | 1 5 M A R C H 2 0 1 2©2012 Macmillan Publishers Limited. All rights reserved

LETTER RESEARCH
Figure 1 | Obesity, hypertrophic adipocytes, accumulation of proinflammatory macrophages and hepatic steatosis in HFD-fed GPR120deficient mice. a, Body weight changes of wild-type (WT) and GPR120deficient mice fed a normal diet (ND) or a HFD (n 5 36–47). b, Indirectcalorimetry in HFD-fed mice. Energy expenditure and respiratory quotient(n 5 4, 5). c, Representative cross-sectional images of wild-type and GPR120deficient mice subjected to microcomputed tomography analysis of the in situ
accumulation of fat. Fat depots are demarcated (green) for illustration.
d, Haematoxylin and eosin (H&E)-stained epididymal WAT and mean area ofadipocytes (n 5 6). Scale bar, 100 mm. e, Relative expression of Cd11b, Cd68 andF4/80 messenger RNA in WAT (n 5 6). a.u., arbitrary units. f, Representative
images of epididymal WAT stained with anti-F4/80 antibody (arrows, F4/80positive cells) and the number of F4/80 cells (n 5 6). Scale bar, 100 mm. g, Oil
Red O-stained liver and hepatic triglyceride content after 24hr fasting (n 5 13).
Scale bar, 50 mm. All data represent mean 6 s.e.m. *P , 0.05 and **P , 0.01
versus the corresponding wild-type value.

b

20
0 ˜
5 6

8 10 12 14
Age (weeks)

16

c

d

23
21

* ** * *

19
17

1

6

HFD

HFD

ND

ND

25

Epididymal WAT,
H&E staining
WT
GPR120–/–

GPR120–/–WT

Dark photoperiod
WT
GPR120–/–

27

12
18
Time (h)

24

**

g

**

30

Liver, Oil Red O staining
GPR120–/–
WT

25
ND

20
15
10

HFD

HFD

ND

F4/80-positive cells (%)

WAT, F4/80 staining
GPR120–/–
WT

5
0

0.82
0.80
0.78
Light Dark

e
*

7
6
5
4
3
2
1
0

ND HFDf

0.84

ND HFD

Cd11b Cd68 F4/80

**

1.0

**

*

0.5

0

NDHFD NDHFD NDHFD

Fig. 3a, b). The level of plasma leptin was significantly higher in HFDfed GPR120-deficient mice than in wild type (Supplementary Fig. 3c).
However, there was no significant difference in terms of plasma
adiponectin level or food intake between the two groups (Supplementary Fig. 3d, e). HFD-fed GPR120-deficient mice showed a marked
increase in the size of islets and KI67 (MKI67)-positive cells, suggesting
adaptive enlargement of the b-cell mass in response to insulin resistance17,18 (Supplementary Fig. 3f, g). Moreover, we observed markedly
reduced peripheral insulin sensitivity in tissues from HFD-fed GPR120deficient mice (Fig. 2c). Insulin was shown to induce the phosphorylation of AKT (AKT1) (on Ser 473) in WAT, liver and skeletal muscle,with similar intensities in wild-type and GPR120-deficient mice on anormal diet (Supplementary Fig. 3h). Consistent with the insulin resistance reported above, HFD-fed GPR120-deficient mice showed loss of
insulin-induced AKT phosphorylation in WAT and the liver.
To determine the molecular basis of the metabolic changes in WAT
and livers of GPR120-deficient mice, we performed gene expression

**

300
250
200
150
100
50

0

ND HFD

Obesity-associated insulin resistance was also more severe in
GPR120-deficient mice. HFD-fed GPR120-deficient mice showed
higher levels of fasting plasma glucose and insulin than did wild type,although these parameters were similar between the two groups on anormal diet (Fig. 2a). HFD-induced insulin resistance, as determinedby an insulin tolerance test, was more prominent in GPR120-deficient
mice than in wild type (Fig. 2b, left, and Supplementary Fig. 3a, b).
A glucose tolerance test further revealed that these mice sufferedfrom impaired glucose metabolism (Fig. 2b, right, and Supplementary

Liver

ND

HFD

ND

HFD

WAT

3

*

*

*

2

*

4

2

3
1

2

1
1
0

0

0
ND

HFD

ND

HFD

ND

HFD

g

h
*
1.2

WT

GPR120–/–TG (μg)/protein (μg)

MEF-derived adipocyte (day 10)

0.8

0.4

0

200

0

0

30 60 90 120
Time (min)IRββ-actin

Scd1

Scd1

100

*

WT GPR120–/–

* **

**
*

2

Insulin
p-AKT
AKT

+

0.08

WT GPR120–/–

i

*

1.4
1.2

25

1.0
0.8

**

0.6

**

15
10

0.4
0.2
0

**

**

5
0

Pparg Fabp4 Srebf1 Scd1

Figure 2 | Impaired glucose metabolism, adipogenesis and lipogenesis in
HFD-fed GPR120-deficient mice. a, Fasting blood glucose and serum insulinlevels (n 5 6–15). b, Plasma glucose during insulin tolerance test (ITT, left) andglucose tolerance test (GTT, right) (n 5 12–14). c, Phosphorylation of AKT(Ser 473) in WAT, liver and skeletal muscle after 24-hr fasting (n 5 6, 7).
NS, not significant. d, Relative mRNA expression of Fabp4 and Scd1 in WAT or
Scd1 in liver (n 5 6). e, Protein expression of IRb, IRS1, IRS2, SCD1 and b-actin
in WAT. f, Protein expression of IRS1, IRS2, SCD1 and b-actin in liver.
g, Oil Red O-staining and triglyceride (TG) content of mouse embryonic

ND HFDWT GPR120–/–

**

*

6

1.8

k

**

5

ND HFD0

ND HFD

0.6

0.6

0

0

0
WT GPR120–/–
WAT/liver ratio l

**
*

0.6

2

**

1.2

1.2

1.2

4

0

**

*

0.10

WAT,
Plasma,non-esterified non-esterified
fatty acidfatty acid
10

**

20

SCD1
β-actinIRS2β-actin

0.20

0

j

GPR120–/–

WT
Liver

IRS1
β-actin

0.5

Liver,
esterified
fatty acid0
GPR120–/–WTf

WT GPR120–/–

**

1.0

GPR120–/–

WT

**

2.0

0

0

0

0
WT GPR120–/–

Skeletal muscle
Insulin – + – +
p-AKT
AKT

+

NS

1
0

–

0.6

1.0

0.04

0.06

+

**

1.2

NS

*

–

NS

SCD1
β-actinIRS2β-actin

0.12

3

–

0.6

IRS1
β-actin

*

+

**

1.2

30 60 90 120
Time (min)
WAT

e

Liver

Fabp4
Relative mRNA expression(a.u.)

0400–

Relative protein expression
normalized with β-actin

0

600

**

Insulinp-AKT
AKT

Relative Scd1 expression(adipose tissue/liver)

50

2.0

800

** **

**

Relative AKT activity(phospho-AKT/total AKT)

100

200

c

C16:1 (mol%)

150

WT
GPR120–/–300

C18:1/C18:0

200

4.0

Blood glucose (mg dl–1)*

HFD, GTTHFD, ITT

Relative protein expression
normalized with β-actin

250

d

b

GPR120–/–Relative mRNA expression
in MEF-derived adipocyte (a.u.)

WT

Fasting insulin (ng ml–1)

Fasting glucose (mg dl–1)

WATa

WT GPR120–/–
WAT,
non-esterified
fatty acid

0.4
0.3

*
**

0.2
0.1
0

0
ND HFDND HFDWT GPR120–/–

m

Liver

Relative mRNA expression(a.u.)**

Respiratory quotient

**

** **

GPR120–/–

WT

Relative mRNA expression (a.u.)

** **

30

**
** **

Adipocyte area (103 μm2)

Body weight (g)40

9–10 weeks of age

Triglyceride (mg g–1 liver)

WT, HFDGPR120–/–, HFD
Energy expenditure(kcal kg–1 h–1)

WT, ND
GPR120–/–, ND

C16:1/C16:0

a

Scd1

**

2.0
1.5
1.0

**

0.5

0
le 7 le 7
hic :1n hic :1n
Ve C16 Ve C16

fibroblast (MEF)-derived adipocyte. Scale bar, 50 mm. h, Relative mRNA
expression in MEF-derived adipocyte (n 5 6). i, The ratio of C18:1 to C18:0 inlivers (n 5 6–8). j, Non-esterified C16:1n7 palmitoleate in WAT and plasma(n 5 4–7). k, The ratio of Scd1 mRNA expression in liver and WAT (n 5 6, 7).
l, The ratio of C16:1 to C16:0 in adipose tissues (n 5 6–8). m, Hepatic Scd1
mRNA expression in mice infused with vehicle or triglyceride:palmitoleate for
6 h (n 5 4, 5). All data represent mean 6 s.e.m. *P , 0.05 and **P , 0.01
versus the corresponding wild-type value.

1 5 M A R C H 2 0 1 2 | VO L 4 8 3 | N AT U R E | 3 5 1©2012 Macmillan Publishers Limited. All rights reserved

RESEARCH LETTER
lean individuals (n 5 14 in each group) were matched for age andgender (Supplementary Table 2). As previously described5,14, we confirmed that GRP120 is barely expressed in liver of either lean or obese
subjects (data not shown). By contrast, we found that GPR120 is well
expressed in the adipose tissue of lean individuals (Fig. 3a). In addition, human obesity is significantly associated with an increase in
GPR120 expression in both subcutaneous and omental adipose tissues(1.8-fold increase; P 5 0.0004 and P 5 0.003, respectively). We also
found that GPR120 expression in subcutaneous adipose tissue
strongly correlates with that in omental adipose tissue (Spearman
analysis; r 5 0.570 and P 5 2.74 3 1028), suggesting a systemic regulation of its expression in humans. Furthermore, we found a positive
correlation between GPR120 expression in both subcutaneous andomental adipose tissues and in subjects’ concentrations of plasma
low-density lipoproteins (on adjustment for age and sex; r 5 0.247,
P 5 0.0115 and r 5 0.255, P 5 0.0118, respectively).
To investigate the contribution of GPR120 to human obesity, the four
GPR120 exons were sequenced in 312 French, non-consanguineous,extremely obese children and adults (Supplementary Table 3). Weab
GPR120 ratio to GAPDH

0.020

6

***
**

0.010
0.005

R270H

4

R67C

3
2
1
0

C
C
M
M
-S
-S
-O
-O
LN
OB
LN
OB
DMSO

c

PMA

NS

**

ALA

1.2

Relative GLP-1 secretion(normalized with PMA)WT5RFU (103)

0.015

0

1

**

3

RFU (103)

0.8
0.6
0.4

100

Vector
WT
R270H
R67C

2

**
1

**

0.2
0

0

e

WT

R270H

R67C

1

10
[ALA] (μM)

WT/WT

PCMV

2×TetO2

FLAG

WT

IRES

HA

WT

WT/R270H

PCMV

2×TetO2

FLAG

WT

IRES

HA

R270H

R270H/WT

PCMV

2×Tet O2

FLAG

R270H

IRES

HA

WT

4

Dox
WT
R270H

2

3
2

WT/R270H

3

4

1

1

R270H/WT

**

2

**
**

1

0

100

WT/WT

Anti-HA

3

RFU (103)

Anti-FLAG

5

RFI

10
[ALA] (μM)

d

1.0Control

T
/W

W
T
R2
70
H

0

H

W

70

R2

W

T/

T/

T
/W
H
70

R2

W

T/

T/

W

T
R2
70
H

0

W

analyses. We identified approximately 700 differentially expressed
genes in WAT between HFD-fed GPR120-deficient and wild-typemice (Supplementary Fig. 4a). Connectivity mapping of these genes
showed that pathways relating to insulin signalling and adipocyte
differentiation were depressed, whereas those related to inflammation
were enhanced in HFD-fed GPR120-deficient mice (Supplementary
Fig. 4b). Quantitative real-time PCR (qRT–PCR) analysis confirmed
thedownregulationofinsulin-signalling-relatedgenes(Insr,Irs1andIrs2),
an adipocyte differentiation marker gene (Fabp4) and a lipogenesisrelated gene (Scd1) in the epididymal fat from HFD-fed GPR120deficient mice (Fig. 2d and Supplementary Fig. 3i). We identified
approximately 100 differentially expressed genes in the liver between
HFD-fed GPR120-deficient and wild-type mice (Supplementary Fig. 5).
Notably, lipogenesis-related genes (Scd1 and Me1) and a fatty acidtransporter gene (Cd36) were significantly upregulated in livers fromGPR120-deficient mice. Quantitative RT–PCR analysis confirmed
upregulation of Scd1 in the liver of GPR120-deficient mice (Fig. 2d).
Western blot analysis further confirmed downregulation of IRb,
IRS1 and SCD1 in adipose tissue of HFD-fed GPR120-deficient mice
(Fig. 2e) but downregulation of IRS1 and IRS2 and upregulation of
SCD1 in their livers (Fig. 2f). Hence, insulin-signalling-related molecules
were downregulated by the lack of GPR120 in both adipose tissue andthe liver. However, the expression of SCD1, the rate-limiting enzyme in
the biosynthesis of mono-unsaturated fatty acids, was downregulated inadipose tissue but upregulated in liver. Furthermore, the expression of
Scd1 and several adipogenic genes14,19 (Pparg, Fabp4 and Srebf1) was
suppressed in mouse-embryonic-fibroblast-derived adipocytes fromGPR120-deficient mice, indicating that GPR120 is required for normal
adipogenesis, as previously reported in differentiating 3T3-L1 adipocytes
depleted of endogenous GPR120 by short interfering RNA14 (Fig. 2g, h).
To determine the effects of altered lipogenesis on lipid composition
in GPR120-deficient mice, we performed lipidomics analysis in WAT,
livers and plasma. Significant changes of major lipid clusters wereobserved (Supplementary Fig. 6). Notably, the hepatic concentrationof oleate (C18:1n9c) was significantly higher in HFD-fed GPR120deficient mice than in wild type. The ratio of C18:1 to C18:0, an
indicator of SCD1 enzyme activity20–23, was markedly enhanced inlivers from HFD-fed GPR120-deficient mice relative to wild type(Fig. 2i). Moreover, the levels of C16:1n7 palmitoleate, which has
recently been recognized as a lipid hormone4, in WAT and plasma
were significantly lower in HFD-fed GPR120-deficient mice than inwild type (Fig. 2j). In particular, lower levels of C16:1n7 palmitoleate
were detected even in WAT of GPR120-deficient mice on a normal
diet (Fig. 2j), which seems to be in good agreement with the suppressed
Scd1 expression and the reduced SCD1 desaturation index20–23 (C16:1/
C16:0; Fig. 2k, l). Lipidomics analysis clearly illustrated dysregulated
lipogenesis in GPR120-deficient mice, and showed the reduced production of lipid hormone C16:1n7 palmitoleate4. To determine
whether the enhanced hepatic lipogenesis in GPR120-deficient mice
was due to the reduced levels of C16:1n7 palmitoleate, we examinedthe effect of C16:1n7 palmitoleate treatment on hepatic Scd1 expression. A 6-h infusion of triglyceride:palmitoleate markedly lowered theenhanced hepatic Scd1 expression in GPR120-deficient mice (Fig. 2m).
The results indicated that the reduced C16:1n7 palmitoleate may
explain the systemic metabolic disorders observed in GPR120deficient mice on a HFD, as palmitoleate has been proposed to be abioactive lipid by which adipose tissue communicates with distantorgans (such as liver) and regulates systemic metabolic homeostasis4.
This study shows that dysfunction of GPR120 can be an underlying
mechanism for diet-associated obesity and obesity-related metabolic
disorders in mouse.
The mouse data prompted us to assess the potential contribution of
GPR120 to the development of obesity and its metabolic complications
in humans. First, the expression levels of GPR120 in both subcutaneousand omental adipose tissues as well as in liver samples were compared
between lean and obese subjects. Normoglycaemic obese patients and

1

10

100

[ALA] (μM)

Figure 3 | GPR120 expression in human obese tissue samples, and effect of
GPR120 variants on [Ca21]i response and GLP-1 secretion. a, GPR120mRNA levels in human subcutaneous (SC) and omental (OM) adipose tissues oflean (LN; n 5 14) and obese (OB; n 5 14) normoglycaemic individuals. Mann–
Whitney analysis, ***P 5 0.0004 and **P 5 0.003. b, ALA-induced [Ca21]i
responses in cells expressing wild-type GPR120 or a p.R67C or p.R270H variant.
c, ALA-induced GLP-1 secretion in NCI-H716 cells expressing a wild-type
GPR120, a p.R67C or a p.R270H receptor. d, Effect of transfection with GPR120
variants on ALA-induced [Ca21]i response in cells stably expressing wild-type
GPR120. e, Effect of co-expression of human GPR120 p.R270H variant withwild-type GPR120 on ALA-induced [Ca21]i response. Top: schematic diagramof constructs. Bottom: expression of wild type and p.R270H (left), andconcentration–[Ca21]i response for ALA in cells expressing wild-type/wildtype, wild-type/R270H or R270H/wild-type receptors (right). **P , 0.01 versus
the corresponding control value. RFI, relative fluorescence intensity; RFU,
relative fluorescence unit. All data show mean 6 s.e.m.

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LETTER RESEARCH
Table 1 | Identified variants in GPR120 exons and association between the p.R67C/rs6186610 or p.R270H non-synonymous variant andobesity
Variant

Missense
p.R67C/rs6186610
variants
p.R270H
Synonymous
p.V38Vvariants
p.S192S
p.V243V
p.S264S

NucleotidechangeChr 10position

MAF1

MAF2
(controls,n 5 7,654)

MAF2(cases,n 5 6,942)

Adjustment: age and gender
OR [95% CI]

P

CRT
GRA
GRA
GRA
CRT
GRA

95,316,666
95,337,031
95,316,581
95,325,846
95,328,938
95,337,014

0.05
0.03
0.00160.00160.00160.0016

0.043
0.013
—
—
—
—

0.055
0.024
—
—
—
—

1.16 [1.02, 1.31]
1.62 [1.31, 2.00]
—
—
—
—

0.022
8.00 3 1026————

Adjustment: age, gender and geography
OR [95% CI]

P1.13 [1.00, 1.28]
0.060
1.58 [1.28, 1.95] 2.17 3 1025
—
—
—
—
—
—
—
—

Variant position was indicated according to human genome build NCBI36/hg18. Association between p.R67C/rs6186610 or p.R270H variant and obesity was assessed by using a logistic regression adjusted forage and gender or for age, gender and geography, under an additive model. Chr, chromosome; MAF1, minor allele frequency in sequencing data set (n 5 312 extremely obese individuals); MAF2, minor allele
frequency in the large obesity case-control genotyping data set; OR, odds ratio; CI, confidence interval.

identified only two non-synonymous variants, R270H (minor allele
frequency (MAF), ,3%) and p.R67C/rs6186610 (MAF, ,5%), andfour rare synonymous variants (MAF, ,1%) (Table 1). The two nonsynonymous variants were subsequently genotyped in 6,942 unrelatedobese individuals and 7,654 control subjects, all of European origin(Supplementary Table 4). By using a logistic regression model adjusted
for age and sex, we found that R270H associated with obesity under an
additive model (OR 5 1.62 [1.31, 2.00]95% (odds ratio and 95%
confidence interval), P 5 8.00 3 1026; Table 1); whereas we foundonly a trend for association between p.R67C and obesity (OR 5
1.16 [1.02, 1.31]95%, P 5 0.022; Table 1). It is noteworthy that theseresults were almost the same after adjusting for geographical origin(Table 1).
We then genotyped the p.R270H variant in 1,109 French pedigrees
selected for obesity (n 5 5,045) and in 780 German trios with one
obese child (n 5 2,340). We observed a significant over-transmissionof the p.R270H low-frequency variant to obese offspring in 117
pedigrees or trios where the p.R270H variant was present (transmission, 62%; P 5 0.009; Supplementary Table 5). This family-based study
excludes a hidden population stratification effect as a cause of spurious
association.
We assessed the functional significance of both the p.R67C mutationand the p.R270H mutation in silico using several programs: arginine
residues at positions 67 and 270 presented a high-evolutionaryconservation pattern among mammals and the two amino-acid substitutions were predicted to be potentially damaging (Supplementary
Table 6). To examine the influences of the two non-synonymous variants on GPR120 function in vitro, we assessed each receptor ability tomobilize intracellular calcium (concentration, [Ca21]i) in response to
the endogenous agonist a-linolenic acid (ALA). We found that ALA
induced [Ca21]i responses in T-REx 293 cells expressing either wildtype or p.R67C receptor in a dose-dependent manner, whereas
ALA-induced [Ca21]i responses in cells expressing p.R270H were significantly lower (P 5 1.6 3 1025) than those in cells expressing wild
type at ALA concentrations greater than 10 mM (Fig. 3b). We further
examined the functional ability of the mutated receptors to secrete
GLP-1 (ZGLP1) from human intestinal NCI-H716 cells, as this cellline lacks GPR120 expression and it can secrete GLP-1 in a regulatedmanner5. ALA induced secretion of GLP-1 in NCI-H716 cells expressing either wild-type (P 5 0.004) or p.R67C (P 5 3.2 3 1025) receptor,
but not in NCI-H716 cells expressing p.R270H mutant (P 5 0.96)(Fig. 3c). The transfection efficiencies for the GPR120 variant receptors
were confirmed to be almost the same (data not shown). To examine
the effect of the p.R270H variant on the wild-type receptor signalling,
we analysed the [Ca21]i dose–response curves after the transfection ofan empty vector, a wild-type receptor plasmid or a p.R270H-mutatedplasmid into T-REx 293 cells expressing wild-type GPR120. The transfection of the p.R270H-mutated plasmid suppressed dose–response
curves, and the maximal ALA-induced [Ca21]i response was significantly decreased (P 5 0.004; Fig. 3d).
To assess the effect more quantitatively, we analysed [Ca21]i dose–
response curves in T-REx 293 cells stably expressing bicistronic
wild-type/wild-type, wild-type/p.R270H or p.R270H/wild-type receptors(Fig. 3e, top). Almost equal levels of receptor protein expression in eachcell line were confirmed by flow cytometry analysis (Fig. 3e, bottom
left). Compared with cells expressing wild-type/wild-type receptor,
the [Ca21]i dose–response curves obtained in cells expressing either
wild-type/p.R270H or p.R270H/wild-type receptor were markedly
suppressed, and the maximal ALA-induced [Ca21]i response was significantly decreased (P 5 1.2 3 1025; Fig. 3e, bottom right). Thesefindings suggest that the p.R270H variant that is significantly associated with obesity has an inhibitory effect on GPR120. The
p.R270H mutant lacks the ability to transduce the signal of long-chain
free fatty acids, contrary to the p.R67C mutant, which did not associatewith obesity.
To analyse whether being a p.R270H variant carrier may affect
GPR120 expression in the adipose tissue, we quantified GPR120 expression in samples from both obese p.R270H carriers and obese noncarriers. Two hundred and thirty-eight obese normoglycaemic patientsfrom the Atlas Biologique de l’Obésité Sévère cohort had already been
genotyped for the p.R270H variant. Ten subjects heterozygous for the
p.R270H variant were matched for age, gender and body mass index
with ten non-carrier (wild-type) obese normoglycaemic patients (Supplementary Table 7). The expression of GPR120 was similar between
p.R270H carriers and wild-type subjects, both in subcutaneous andomental adipose tissues (Supplementary Fig. 7a), suggesting that thepresence of the functionally deleterious mutation has no primary or
secondary effect on gene expression in fat depots. The adipogenesis
marker PPARG, the lipogenesis-related factor SCD and the macrophage
marker CD68 were found similarly well expressed in the adipose tissuesof wild-type and p.R270H carrier patients (Supplementary Fig. 7b, c).
Nevertheless, the expression of the fatty-acid-binding protein FABP4
in omental adipose tissue was significantly lower in p.R270H carriers
than in wild-type individuals (28% decrease, P 5 0.043; Supplementary
Fig. 7b).
Our results provide convincing evidence that the lipid sensor GPR120
is involved in obesity in both mice and humans. Given the role of
GPR120 as a physiologic integrator of the environment (especially the
fatty diet), these data provide insight into the molecular mechanisms bywhich the ‘Westernized’ diet may contribute to early-onset obesity andassociated complications including non-alcoholic steatohepatitis. It also
brings some understanding of the metabolic effects of the omega-3 fatty
acids, which are often proposed as food supplements. This may open
novel avenues of research for drug development in the treatment ofobesity, lipid metabolism abnormalities and liver diseases, because
receptors that sense free fatty acids represent attractive drug targets.

METHODS SUMMARY
GPR120-deficient mice were generated by deleting Gpr120 exon 1. All animal
procedures and euthanasia were reviewed by the local animal care committee
approved by local government authorities. Blood analysis, extraction and detection of mRNA and proteins, and immunohistochemical analysis, were performed
following standard protocols as described previously5,24–26. Details of antibodies,primers and probes are given in Methods. The level of significance for the difference between data sets was assessed using Student’s t-test. Analysis of variancefollowed by Tukey’s test was used for multiple comparisons.
1 5 M A R C H 2 0 1 2 | VO L 4 8 3 | N AT U R E | 3 5 3©2012 Macmillan Publishers Limited. All rights reserved

RESEARCH LETTER
In human, GPR120 expression in liver and in both omental and subcutaneous
adipose tissues was assessed by quantitative RT–PCR (Taqman), in lean and obese
subjects from the Atlas Biologique de l’Obésité Sévère cohort. The four GPR120
exons were sequenced in 312 French, extremely obese subjects following a standard
Sanger protocol. The two identified non-synonymous variants (p.R270H and
p.R67C/rs6186610) were subsequently genotyped in a large European obesitycase-control study (ncases 5 6,942, ncontrols 5 7,654), by high-resolution melting
analysis and TaqMan, respectively. The association between obesity status andeach variant was assessed using logistic regression adjusted first for age and genderand then for age, gender and geography origin, under an additive model. The
consequences of the two identified non-synonymous variants for GPR120 function ([Ca21]i response and GLP-1 secretion) were assessed in vitro. The humanstudy protocol was approved by the local ethics committee, and participants fromall of the studies signed an informed consent form.
Full Methods and any associated references are available in the online version of
the paper at www.nature.com/nature.
Received 9 November; accepted 14 December 2011.
Published online 19 February 2012.
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Supplementary Information is linked to the online version of the paper at
www.nature.com/nature.
Acknowledgements We are indebted to all subjects who participated in these studies.
In Japan, the study was supported in part by research grants from the Japan Society for
the Promotion of Science; the Ministry of Education, Culture, Sports, Science andTechnology of Japan; the Japan Science and Technology Agency; and the Funding
Program for World-Leading Innovative R&D on Science and Technology (FIRST
Program), initiated by the Council for Science and Technology Policy. A.I. is a fellow of
the Japan Society for the Promotion of Science. A.B. is a fellow of the EU-funded
EUROCHIP consortium. In France, the study was supported by le Conseil Régional Nord
Pas de Calais/FEDER and the Agence Nationale de la Recherche (Programme deRecherche en Nutrition et Alimentation, SensoFAT). The Northern Finland Birth Cohort
Studies 1986 received financial support from the Academy of Finland, the University
Hospital of Oulu (Finland), the University of Oulu (Finland), the European Commission(EURO-BLCS, Framework 5 award QLG1-CT-2000-01643), and the Medical ResearchCouncil (G0500539, G0600705, PrevMetSyn/SALVE). We thank the ABOS consortiumand the CIC-CCPPRB (Lille CHRU) team for their help in sample handling and clinicaldata collection. We are grateful to M. Deweirder and F. Allegaert for human DNA bankmanagement.
Author Contributions A.I., A. Hirasawa, O.P.-G. and A.B. are equally contributing firstauthors. G.T. and P.F. had the ideas for the mouse and human projects, respectively. A.I.,
A. Hirasawa, A.B., P.F. and G.T. drafted the manuscript. O.P.-G., H.C., D.M. and I.W. edited
the manuscript and contributed to discussions. A. Hirasawa and G.T. designed themouse research. A.I., A. Hirasawa, K.I. and G.T. created Gpr120-mutant mice. A.I., A.
Hirasawa, A. Körner, T.H., I.K., T.-a.K., K.A., M. Takeuchi, K.O., N.L. and G.T. conductedbiochemical and histochemical analyses for the mouse study. A.I. and A. Hirasawa
performed bioinformatic analysis for the mouse study. L.Y. and C.L. performed thestatistical analyses, and A.B. contributed to statistical analyses for the human study.
O.P.-G. and I.W. designed the human expression gene study. A.L. performed the humanexpression gene study. H.C. and S.V. performed GPR120 sequencing and variant
genotyping, respectively. P.B., M. Tauber, C.M., A.M., R.B., P.E., M.-R.J., W.V.H., L.V.G., F.H.,
B.B., C.L.-M., K.R., A. Kouvatsi and F.P. contributed to cohort-study samples andresearched data.
Author Information Microarray data have been deposited in the NCBI Gene Expression
Omnibus under accession number GSE32095. Reprints and permissions information
is available at www.nature.com/reprints. The authors declare no competing financial
interests. Readers are welcome to comment on the online version of this article atwww.nature.com/nature. Correspondence and requests for materials should be
addressed to P.F. (p.froguel@imperial.ac.uk) or G.T. (gtsuji@pharm.kyoto-u.ac.jp).

3 5 4 | N AT U R E | VO L 4 8 3 | 1 5 M A R C H 2 0 1 2©2012 Macmillan Publishers Limited. All rights reserved

LETTER RESEARCHMETHODS
Generation and genotyping of GPR120-deficient mice. GPR120-deficient miceon a mixed C57Bl/6/129 background were generated by homologous recombination. Exon 1 of the Gpr120 gene was replaced with a PGK-neo cassette (Supplementary Fig. 1).
Animals. Mice were housed under a 12-hr light–dark cycle and given regularchow, MF (Oriental Yeast Co.). For HFD studies, 5-week-old male mice wereplaced on a 58Y1 diet (PMI Nutrition International) for a total period of 11 weeks.
The methods used for animal care and experimental procedures were approved by
the Animal Care Committee of Kyoto University.
Indirect calorimetry. Twenty-four-hour energy expenditure and respiratory
quotient (RQ) were measured by indirect calorimetry, using an open-circuit
calorimeter system (MK-5000RQ, Muromachi Kikai Co.). Respiratory quotientis the ratio of carbon dioxide production to oxygen consumption (VO2). Energy
expenditure was calculated as the product of the calorific value of oxygen(3.815 1 1.232RQ) and VO2. Locomotor activity was measured by using an
infrared-ray passive sensor system (Supermex, Muromachi Kikai Co.).
Histology and immunohistochemistry. Epididymal adipose and pancreatic tissues
were fixed in 10% neutral-buffered formalin, embedded in paraffin, and sectioned at
5 mm. H&E staining was performed using standard techniques. To measure thediameter of adipocytes and the area of pancreatic islets, the diameters of 100 cells
from five sections from each group were measured using NIH IMAGE software. Morethan 10 fields were examined, islet area was traced and total islet area was calculated
and expressed as the average score. Liver tissues were embedded in OCT compound(Sakura Finetech) and snap-frozen in liquid nitrogen. Tissue sections were stainedwith Oil Red O (Sigma-Aldrich) for lipid deposition using standard methods.
Triglyceride content assay. To determine the triglyceride content of liver, tissue
was homogenized with 1/2.5/1.25 (vol/vol) 0.5 M acetic acid/methanol/
chloroform. The mixture was shaken and 1.25 volumes of chloroform added.
The mixture was shaken overnight and then 1.25 volumes of 0.5 M acetic acidadded. After centrifugation at 1,500g for 10 min, the organic layer was collected,
dried and resuspended in 100% isopropyl alcohol. Measurements were conductedusing Triglyceride E-test Wako (Wako).
Glucose tolerance and insulin tolerance tests. Glucose tolerance assays were
performed on 24-hr-fasted mice. After baseline glucose values were individually
established using One Touch Ultra (LifeScan), each mouse was given an intraperitoneal injection of 1.5 mg glucose per gram of body weight. Insulin tolerance
was conducted using the same glucometer. After baseline glucose values were
established, mice were given human insulin (0.75 mU g21 intraperitoneal;
Sigma-Aldrich). Clearance of plasma glucose was subsequently monitored at 15,30, 60, 90 and 120 min post-injection.
Immunoblot analysis. For insulin stimulation, 5 U insulin (Sigma-Aldrich) was
injected through the inferior vena cava. Five minutes later, samples of liver, skeletal
muscle or WAT were dissected and immediately frozen in liquid nitrogen. Immunoblot analysis were performed as described previously5,24,25. Anti-IRS1 (Millipore),anti-IRS2 (Millipore), anti-SCD1 (Santa Cruz Biotechnology), anti-IRb anti-AKT(Cell Signaling Technology), anti-p-AKT (Cell Signaling Technology) and anti-bactin (Sigma-Aldrich) antibodies were used as the primary antibodies.
Mouse gene expression analysis. Total RNA was extracted from tissue or cells
using ISOGEN (Nippon Gene). Quantitative RT–PCR and microarray analysis
were performed as described previously24,26. Briefly, genome-wide mRNA expression profiles were obtained by microarray analysis with the Affymetrix GeneChip
Mouse 430 2.0 Array, according to the manufacturer’s instructions. We used therobust multi-array analysis expression measure that represents the log-transformationof intensities (background corrected and normalized) from the GeneChips27.
Functional associations between differentially expressed genes were analysed using
Ingenuity Pathways Analysis (version 4.0, Ingenuity Systems).
Microcomputed tomography scanning. Images were obtained using a microcomputed tomography system (SHIMADZU ClairvivoCT) with a high-resolution
flat-panel detector. The maximum resolution of this modality was less than 40 mm.
The scanner was assumed to have a cylindrical field of view of 65.3 mm in section
view and of 300 mm in transaxial view. The X-ray source was biased at 60 keV withthe anode current set to 160 mA. Computed tomography images were analysedwith OSIRIX software (http://www.osirix-viewer.com/).
Fatty acid composition of epididymal WAT, liver and plasma. Esterified andnon-esterified fatty acid composition was measured by gas chromatography.
Briefly, to analyse esterified fatty acid, samples of epididymal adipose tissue(20–25 mg), liver (25–30 mg) and plasma (100 ml) were snap-frozen in liquid
nitrogen and homogenized in 4 ml of 0.5 N KOH-methanol. Samples were thenboiled at 100 uC for 30 min to hydrolyse. Total lipids in each sample homogenate
were then extracted with hexane, followed by trans-esterification of fatty acids
using boron trifluoride-methanol at 100 uC for 15 min. Methylated fatty acids werethen extracted with hexane and analysed using a GC-2010AF gas chromatograph(SHIMADZU). For the analysis of non-esterified fatty acid, samples of epididymal
adipose tissue (10–15 mg), liver (10–15 mg) and plasma (100 ml) were snapfrozen in liquid nitrogen and homogenized in a mixture of 1.2 ml water, 3 ml
methanol and 1.5 ml chloroform. Total lipids in each sample homogenate wereextracted with a mixture of 1.2 ml of water and 1.2 ml of chloroform, followed bysilylation of fatty acids using N,O-bis(trimethylsilyl)trifluoroacetamide with 1%
trimethylchlorosilane at 100 uC for 60 min. Silylated fatty acids were then extractedwith hexane and analysed using a GC-2010AF gas chromatograph (SHIMADZU).
Mouse embryonic fibroblast adipogenesis assay. To prepare MEFs, we minced
13.5-d-post-coital mouse embryos and digested them with trypsin. Cells werecollected and cultured in modified Eagle’s medium (a-MEM; supplemented with
10% fetal bovine serum (FBS), 50 U ml21 penicillin and 50 mg ml21 streptomycin).
We induced confluent MEFs to undergo adipogenic differentiation by incubatingthem first for 2 d with 10 mg ml21 insulin, 250 nM dexamethasone and 0.5 mM
isobutylmethylxanthine (Sigma-Aldrich). We measured cellular triglyceride content with Triglyceride E-test Wako (Wako).
Lipid infusion. Intralipid solution with 2 mM triglycerides:palmitoleate was prepared using a previously described protocol4. Briefly, lipids were dissolved in asolvent containing 5% glycerol and 0.72% phosphocholine in 0.9% saline andsonicated repeatedly. Lipids stayed in suspension for one week and had to bevortexed well before loading the syringe and tubing to prevent clogging. Before
lipid infusion, mice were anaesthetized and an indwelling catheter was inserted in
the left internal jugular vein. After overnight fasting, lipids were infused at a rate of
500 ml kg21 h21 for 6 h. At the end of the infusion, tissues were collected.
Statistical analysis of the GPR120-deficient mouse study. The level of significance for the difference between data sets was assessed using Student’s t-test.
Analysis of variance followed by Tukey’s test was used for multiple comparisons.
Data were expressed as mean 6 s.e.m. P , 0.05 was considered to be statisticallysignificant.
Human study population. The study protocol was approved by all local ethics
committees and informed consent was obtained from each subject before participation in the study, in accordance with the Declaration of Helsinki principles.
For children younger than 18 years, an oral consent was obtained and parents
provided written informed consent. All subjects were of European origin.
Human gene expression analysis. We used liver, subcutaneous and omental
adipose tissue samples from the Atlas Biologique de l’Obésité Sévère’ (ABOS)
cohort (ClinicalGov NCT01129297), a cohort studied in the Département deChirurgie Générale et Endocrinienne28 (Lille CHRU). Total RNA was extracted
from the tissues using an RNeasy protect Mini Kit (QIAGEN) and quantified byabsorbance at 260 nm and 280 nm in a PerkinElmer spectrophotometer. Human
GPR120, FABP4, PPARG, CD68 and SCD mRNA levels were quantified by reverse
transcription reaction followed by qRT–PCR. Quantitative assessment of humanmRNA expression was performed using TaqMan Gene Expression Assays(Hs01111664_m1: GPR120 and Hs99999905_m1: GAPDH; Hs00609791_m1:
FABP4; Hs00234592_m1: PPARG; Hs00154355_m1: CD68; Hs01682761_m1:
SCD; Applied Biosystems) with an Applied Biosystems 7900HT Fast Real-Time
PCR System. As an internal control for potential housekeeper reference variability,
gene transcript levels were normalized to GAPDH reference housekeeper transcript level. The mean of the triplicate cycle thresholds of the target was normalized
to the mean of triplicate cycle thresholds of the reference internal housekeeper genes
using the formula 2CTGAPDH {CTtarget , which yielded a relative target-to-reference transcript concentration value as a fraction of reference transcript. Samples for which
the cycle threshold was above 35 were excluded from the analysis.
GPR120 exon sequencing. We sequenced the four GPR120 exons in 312 obese
patients including 121 French obese adults and 191 French obese children who
were recruited by the CNRS-UMR8199 unit and the Department of Nutrition of
Paris Hotel Dieu Hospital. GPR120 is located on human chromosome 10q23.33
and encodes a 377-amino-acid protein (NCBI NM_181745.3 and NP_859529).
PCR conditions and primer sequences are available on request. Fragments werebidirectionally sequenced using the automated 3730xl DNA Analyzer (Applied
Biosystems). Electrophoregram reads were assembled and analysed using the
VARIANT REPORTER software (Applied Biosystems). The locations of the variants are displayed by base numbers counting from the ATG translation initiation
codon following the Human Genome Variation Society nomenclature for thedescription of sequence variations. The positions of mutations were indicatedby reference to the human genome build NCBI36/hg18.
Genotyping of the p.R270H and p.R67C/rs6186610 variants. We genotyped
the two non-synonymous variants in 6,942 unrelated obese subjects and in 7,654
control subjects, all of European descent. Genotyped populations are described in
Supplementary Table 4. The set of obese subjects included 516 unrelated Frenchobese children who were recruited by the CNRS-UMR8199 unit or Toulouse
Children’s Hospital29; 332 Italian obese children from Verona30 or Rome31; 170
Finnish obese adolescents from the Northern Finland Birth Cohort 198632©2012 Macmillan Publishers Limited. All rights reserved

RESEARCH LETTER(NFBC1986); 1,164 unrelated French obese adults from the ABOS cohort28 or
recruited by the CNRS-UMR8199 unit and the Department of Nutrition of
Paris Hotel Dieu Hospital29; 2,514 Belgian obese patients from the outpatient
obesity clinic at Antwerp University Hospital33; 1,736 Swiss obese subjects who
were recruited after gastric surgery in Zurich34; and 510 Greek obese subjects
recruited in the Hippokration Hospital of Thessaloniki or in the Second
Department of Internal Medicine of the Hospital of Alexandroupolis35. The setof control subjects included 422 Italian lean children from Verona30; 4,639 Finnish
lean adolescents from the NFBC1986 cohort32; 1,976 French lean adults from the
D.E.S.I.R (Data from the Epidemiological Study on the Insulin Resistancesyndrome) prospective study36 and from the Haguenau study37; 148 Belgian leansubjects from Antwerp Hospital33; and 469 Greek lean individuals recruited in
medical examination centres in Thessaloniki35. The 1,109 French pedigrees
selected for obesity were recruited by the CNRS-UMR819938, and the 780
German childhood obesity trios were recruited at the Universities of Marburg
and Essen39. The p.R270H variant was genotyped using the LightCycler 480
High Resolution Melting (HRM) Master kit (Roche), following the manufacturer’s
protocol. Genotyping of p.R67C/rs6186610 was performed using a custom TaqMan
assay according to the manufacturer’s instructions (Applied Biosystems). Allelicdiscrimination was performed using an Applied Biosystems 7900HT Fast RealTime PCR System and SDS 2.3 software. For both variants, genotype success rate
was at least 95% and no deviation (P . 0.05) from Hardy–Weinberg equilibrium
was observed in any of the examined populations.
Phenotyping. The 90th and 97th body mass index (BMI) percentiles were used as
thresholds for childhood overweight and obesity, respectively, according to therecommendations of the European Childhood Obesity Group study in local reference
populations40,41. Adult subjects were defined as normal (BMI , 25 kg m22), overweight (25 # BMI , 30 kg m22) and obese (BMI $ 30 kg m22) according to theInternational Obesity Task Force recommendations.
In silico analysis of both p.R270H and p.R67C variants. Phylogenetic conservation of the part of GPR120 containing each non-synonymous variant wasassessed using the UCSC genome browser (Vertebrate Multiz Alignment &Conservation), based on a phylogenetic hidden Markov model, phastCons42. To
predict the possible effect of both amino-acid substitutions on the structure andfunction of GPR120, we used several programs: the POLYPHEN (polymorphism
phenotyping) web-based program43,44; PANTHER45 (protein analysis throughevolutionary relationships); the SIFT (sorting intolerant from tolerant) algorithm46; the SNAP (screening for non-acceptable polymorphisms) software47;
and the PMUT web-based program48.
Plasmid construction. A FLAG-human GPR120/pcDNA5/FRT/TO plasmid wasconstructed by ligating GPR120 complementary DNA into the multicloning site of
the mammalian expression vector pcDNA5/FRT/TO (Invitrogen) with the aminoterminal FLAG tag. The point mutation for constructing the FLAG-human GPR120
p.R67C/pcDNA5/FRT/TO and FLAG-GPR120 p.R270H/pcDNA5/FRT/TO
plasmids was carried out using the following primers: p.R67C (sense: 59ggtgctggtggcgtgccgacgacgcc-39; antisense: 59-ggcgtcgtcggcacgccaccagcacc-39) andp.R270H (sense: 59-agccaccagatccacgtgtcccagcaggac-39; anti-sense: 59-gtcctgctggga
cacgtggatctggtggct-39). All constructs were confirmed by DNA sequencing.
Cell lines and cell culture. Flp-In T-REx-293 (T-REx 293) cells (Invitrogen) wereused to develop inducible and stable cell lines expressing GPR120 (wild type,
p.R270H or p.R67C). T-REx 293 cells were routinely cultured in Dulbecco’s modified
Eagle’s medium (DMEM; Sigma) supplemented with 10% FBS, 100 mg ml21 Zeocin(Invitrogen) and 10 mg ml21 blasticidin S (Funakoshi). T-REx 293 cells were transfected with FLAG–GPR120 (wild type, p.R270H or p.R67C)/pcDNA5/FRT/TO
using Lipofectamine reagent (Invitrogen) and selected with DMEM, which had
been supplemented with 10% FBS, 10 mg ml21 blasticidin S and 100 mg ml21
hygromycin B (Gibco BRL). GPR120 protein expression was induced by adding
10 mg ml21 of doxycycline hyclate (Dox; Sigma) for 48 h. Human NCI-H716 cells
were obtained from the American Type Culture Collection (Manassas). Cells weregrown in suspension in Roswell Park Memorial Institute 1640 medium supplemented with 10% FBS, 100 IU ml21 penicillin and 100 mg ml21 streptomycin.
[Ca21]i response analysis. Cells were seeded at a density of 2 3 105 cells per well on
collagen-coated 96-well plates, incubated at 37 uC for 21 h and then incubated in
Hanks’ Balanced Salt Solution (HBSS, pH 7.4) containing Calcium Assay Kit
Component A (Molecular Devices) for 1 h at 20 uC. ALA used in the fluorometric
imaging plate reader (FLIPR, Molecular Devices) assay were dissolved in HBSS containing 1% DMSO and prepared in another set of 96-well plates. These plates were seton the FLIPR, and mobilization of [Ca21]i evoked by agonists was monitored.
Transfection. One million cells were seeded into a 3.5-cm-diameter dish before
transfection. NCI-H716 cells were transfected with 5 mg of each plasmid using
Lipofectamine 2000 (Invitrogen) according to the manufacturer’s protocol. At
24 h post-transfection, transfection of each FLAG-tagged construct was confirmedby anti-FLAG FACS analysis. Then the cells were reseeded in 24-well culture plates

coated with Matrigel matrix (BD Biosciences) at a density of approximately
3 3 105 cells per well for the secretion studies. To test the effect of variant receptors
on the ALA-induced [Ca21]i response of the wild-type receptor, 2 3 107 T-REx
293 cells expressing Dox-inducible FLAG–GPR120 wild type were seeded into a
15-cm-diameter dish before transfection. Cells were then transfected with 32 mg ofeach plasmid (empty vector, wild type and p.R270H GPR120) using
Lipofectamine 2000 (Invitrogen) according to the manufacturer’s protocol. At
24 h post-transfection, cells were reseeded at a density of 2 3 105 cells per wellon collagen-coated 96-well plates and treated with 10 mg ml21 of Dox, and at 48 h
post-transfection, ALA-induced [Ca21]i response was monitored.
GLP-1 secretion analysis. Cells were serum-starved with FBS-free DMEM for 3 h,washed with HBSS and incubated for 2 h at 37 uC in 0.3 ml FBS-free DMEM
containing DMSO (negative control), 1 mM phorbol 12-myristate 13-acetate(positive control) or 100 mM ALA. Supernatants were collected and the active
GLP-1 concentration in the supernatant was determined by enzyme immunoassay
using GLP-1 (Active) ELISA Kit (Millipore).
Flow cytometry analysis. Anti-FLAG (Sigma) and anti-HA (Roche) antibodies
were used for staining. Data were acquired and analysed on FACSCalibur with
CELLQUEST software (Becton Dickinson).
Statistical analysis of human study. We assessed the effect of both nonsynonymous variants (p.R270H and p.R67C) on obesity using a logistic regression
adjusted first for age and gender and then for age, gender and geography, under an
additive model, using the software R (version 2.12). Adjustment for geography wasachieved to reflect a north–south gradient between the six different countries oforigin of the study participants. An ordinal variable was created and coded: 1 for
Finland, 2 for Belgium, 3 for France and Switzerland, 4 for Italy and 5 for Greece.
This variable was added as a covariate in the logistic regression model.
Data analysis for the [Ca21]i response was performed using IGOR PRO(WaveMetrics). Significant differences between expression among wild-type andheterozygous groups, and among lean and obese wild-type subjects, were assessedusing non-parametric Mann–Whitney analysis (GRAPHPAD PRISM 5 software).
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