Palmitoyl lactic acid induces adipogenesis and a brown fat-like phenotype in 3T3-L1 preadipocytes

Article abstract of DOI:10.1016/j.bbalip.2018.04.003

Brown adipose tissue is specialized to generate heat by dissipating chemical energy and may provide novel strategies for obesity treatment in humans. Recently, advances in understanding the pharmacological and dietary agents that contribute to the browning of white adipose tissue have been made to alleviate obesity by promoting energy expenditure. Krill oil is widely used as a health supplement in humans. In this study, the components from krill oil that promote adipogenesis of 3T3-L1 cells were screened to reveal palmitoyl lactic acid (PLA) as a promoter of adipogenesis. The PLA-induced adipocytes contained large number of small lipid droplets. Moreover, similar to the peroxisome proliferator-activated receptor (PPAR)γ agonists, pioglitazone and rosiglitazone, PLA significantly enhances adipogenesis in the presence of dexamethasone compared with PLA alone. Treatment with PLA causes a brown fat-like phenotype in 3T3-L1 cells by enhanced expression of various brown/beige cell-specific genes, such as PR domain containing 16 (Prdm16) and peroxisome proliferative activated receptor, gamma, coactivator 1 alpha (Pgc1a), as well as adiponectin gene. The expression profile of the brown/beige cell-specific genes induced by PLA was similar to that of the PPARγ agonist in 3T3-L1 cells. Our findings suggest that PLA induces a brown fat-like phenotype and, thus, likely has therapeutic potential in treating obesity.

Full text of DOI:10.1016/j.bbalip.2018.04.003

Accepted Manuscript
Palmitoyl lactic acid induces adipogenesis and a brown fat-like
phenotype in 3T3-L1 preadipocytes
Yuka Unno, Hirona Yamamoto, Shuto Takatsuki, Yoshinori Sato,
Takefumi Kuranaga, Kazunaga Yazawa, Yasuo Ono, Toshiyuki
Wakimoto
PII:
DOI:
Reference:
S1388-1981(18)30056-8
doi:10.1016/j.bbalip.2018.04.003
BBAMCB 58271
To appear in:
Received date:
Revised date:
Accepted date:
17 October 2017
26 March 2018
8 April 2018
Please cite this article as: Yuka Unno, Hirona Yamamoto, Shuto Takatsuki, Yoshinori Sato,
Takefumi Kuranaga, Kazunaga Yazawa, Yasuo Ono, Toshiyuki Wakimoto , Palmitoyl
lactic acid induces adipogenesis and a brown fat-like phenotype in 3T3-L1 preadipocytes.
The address for the corresponding author was captured as affiliation for all authors. Please
check if appropriate. Bbamcb(2018), doi:10.1016/j.bbalip.2018.04.003
This is a PDF file of an unedited manuscript that has been accepted for publication. As
a service to our customers we are providing this early version of the manuscript. The
manuscript will undergo copyediting, typesetting, and review of the resulting proof before
it is published in its final form. Please note that during the production process errors may
be discovered which could affect the content, and all legal disclaimers that apply to the
journal pertain.

ACCEPTED MANUSCRIPT
For, Biochimica et Biophysica Acta (BBA) - Molecular and Cell Biology of
Lipids, research paper
Title
Palmitoyl lactic acid induces adipogenesis and a brown fat-like phenotype in
3T3-L1 preadipocytes
Authors
Yuka Unno,a＊ Hirona Yamamoto,b Shuto Takatsuki,b Yoshinori Sato,a Takefumi
b
c
a
Kuranaga, Kazunaga Yazawa, Yasuo Ono, ＊ Toshiyuki Wakimoto,b
＊
Affiliation
a, Department of Microbiology and Immunology, Teikyo University School of Medicine,
Tokyo, Japan
b, Faculty of Pharmaceutical Sciences, Hokkaido University, Sapporo, Japan.
c, Integrated Institute for Regulatory Science, Research Organization for Nano and Life
Innovation, Waseda University, Tokyo, Japan.
1

ACCEPTED MANUSCRIPT
＊Corresponding Author
Yuka Unno, Department of Microbiology and Immunology, Teikyo University School
of Medicine, 2-11-1 Kaga, Itabashi, Tokyo 173-8605, Japan.
Phone: +81-3-3964-3632
Fax: +81-3-5375-5284
E-mail: yunno@med.teikyo-u.ac.jp
Yasuo Ono, Department of Microbiology and Immunology, Teikyo University School
of Medicine, 2-11-1 Kaga, Itabashi, Tokyo 173-8605, Japan.
Phone: +81-3-3964-3616
Fax: +81-3-5375-5284
E-mail: onoy@med.teikyo-u.ac.jp
Toshiyuki Wakimoto, Faculty of Pharmaceutical Sciences, Hokkaido University, Kita 12,
Nishi 6, Kita-ku, Sapporo 060-0812, Japan.
Phone: +81-11-706-3239
Fax: +81-11-706-3922
E-mail: wakimoto@pharm.hokudai.ac.jp
2

ACCEPTED MANUSCRIPT
Running title: Palmitoyl lactic acid induces adipogenesis
Keywords: krill oil; beige; adipogenesis; PPARagonist;PRDM16; palmitoyl lactic
acid;
Abstract
Brown adipose tissue is specialized to generate heat by dissipating chemical energy and
may provide novel strategies for the defense against human obesity. Recent advances
have been made in the understanding of pharmacological and dietary agents that
contribute to the browning of white adipose tissue to alleviate obesity by promoting
energy expenditure. Krill oil is widely used as a human health supplement. In this study,
we screened components from krill oil that promote adipogenesis of 3T3-L1 cells
identified palmitoyl lactic acid (PLA). The PLA-induced adipocytes were filled with a
large number of small lipid droplets. Moreover, as with the peroxisome
proliferator-activated receptor (PPAR) agonists pioglitazone and rosiglitazone, PLA
significantly enhanced adipogenesis in the presence of dexamethasone, compared with
treatment with PLA alone. Treatment of PLA causes a brown fat-like phenotype in
3T3-L1 cells via enhanced expression of various brown/beige cells-specific genes, such
3

ACCEPTED MANUSCRIPT
as PR domain containing 16 (Prdm16) and peroxisome proliferative activated receptor,
gamma, coactivator 1 alpha (Pgc1a), as well as adiponectin gene. The expression
profile of the brown/beige cell-specific genes induced by PLA was similar to that by the
PPAR agonist in 3T3-L1 cells. Our findings suggest that PLA induces the brown
fat-like phenotype and thus likely has therapeutic potential for the treatment of obesity.
1. Introduction
Obesity is an important risk factor for cardiovascular and kidney diseases, diabetes,
some cancers, and musculoskeletal disorders [1-5] and has emerged as a leading cause
of staggering health care expenses along with the expanding population of chronically
ill people [1,2]. One of the major causes of obesity is the accumulation of excessive
calories as fat mainly due to the loss of balance between caloric intake and expenditure.
In addition to lifestyle modification to reduce caloric intake and/or increase caloric
expenditure by exercise, it is highly desired to develop new therapies for obesity.
An adipocyte is an endocrine cell that secretes a number of mediators,
including metabolites, cytokines, lipids, and coagulation factors [6]. In general,
adipocytes in mammals are roughly divided into two types: white and brown [7,8].
Brown adipocytes express constitutively high levels of thermogenic genes and contain a
4

ACCEPTED MANUSCRIPT
large amount of uncoupling protein-1 (UCP1), which plays an important role in
thermogenesis [2,7]. Therefore, enhancement of thermogenesis in brown adipocytes is
considered to be an attractive target for anti-obesity therapy [2]. It was revealed that
white adipose tissue harbors brown-like adipocytes [7,8], which are referred to as “beige”
or “brite” (from brown-in-white) adipocytes and are characterized by uncoupled
respiration [7-9]. The beige/brite adipocytes in mice have similar molecular
characteristics to the brown adipocytes of adult humans, such as increased expression of
specific genes, which include tumor necrosis factor receptor superfamily member 9
(Tnfrsf9/CD137), transmembrane protein 26 (Tmem26), and T-box 1 (TBX1); however,
the origin of beige/brite adipocytes remains unclear [2, 8-10]. One possibility is
white-to-brown transdifferentiation, in which “britening” occurs through a direct
interconversion of a white adipocyte into a brown adipocyte phenotype [8]. Another
possibility is that beige/brite adipocytes are recruited via the de novo differentiation of
preadipocytes [11]. An increase in the abundance of beige/brite adipocytes in white
adipose tissue (browning) has been observed in tissue in response to low temperatures.
Notably, such increases have also been observed in white adipose tissue stimulated by
pharmacological and dietary agents [10,11]. For example, capsaicin, berberine, irisin,
monoterpene limonene, curcumin, and chrysin have been reported as natural products
5

ACCEPTED MANUSCRIPT
that promote browning [9,12-17]. The activities of beige/brite adipocytes were shown to
reduce metabolic disease in mice and have been correlated with leanness in humans [2].
These lines of evidence have raised the possibility of the discovery of new compounds
that promote browning. Studies to elucidate the molecular mechanism underlying the
ability of such compounds, in addition to known agents, to induce browning may lead to
new therapies for metabolic diseases, including obesity.
Krill oil, extracted from the Antarctic krill Euphasia superba, a shrimp-like
zooplankton, represents a valuable source of omega-3 polyunsaturated fatty acids [18],
in amounts similar to those present in fish oil. Krill oil is widely used as a human health
supplement. Bunea et al. reported that krill oil was significantly more effective than fish
oil for the reduction of glucose, triglycerides, and low-density lipoprotein levels in
humans [19]. Berge et al. showed that dietary supplementation with a krill powder
persistently ameliorated high triglycerides without weight loss in obese men to an extent
similar to that observed following lifestyle-induced weight loss in a previous study of
dyslipidemic overweight/obese men [20]. Moreover, Li et al. demonstrated that krill oil
reduced plasma cholesterol and glucose levels in rats fed a high-cholesterol diet [21].
One possible reason for the beneficial effects unique to krill oil is that omega-3
polyunsaturated fatty acids exist in phospholipid form rather than as a triglyceride in
6

ACCEPTED MANUSCRIPT
contrast to fish oil supplements [22,23]. However, the mechanisms of action of krill oil
remain poorly understood.
In a previous report, krill oil supplementation was shown to modulate key
metabolic pathways, including glucose metabolism, lipid metabolism, and the
mitochondrial respiratory chain [23,24]. Surprisingly, an equimolar dose of
eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA) derived from fish oil did
not modulate key metabolic pathways regulated by krill oil [24], suggesting that krill oil
contains unidentified active compounds other than EPA and DHA that modulate key
metabolic pathways as a signaling molecule. In line with this concept, we screened
components from krill oil that promote adipogenesis and identified palmitoyl lactic acid
(PLA). Moreover, PLA was found to induce a brown fat-like phenotype in 3T3-L1
preadipocytes via enhanced expression of brown/beige cell-specific genes, such as PR
domain containing 16 (Prdm16), cytochrome c oxidase subunit VIIa 1 (Cox7a1),
peroxisome proliferative activated receptor, gamma, coactivator 1 (Pgc1a),
Cbp/p300-interacting transactivator with Glu/Asp-rich carboxy-terminal domain 1
(Cited1), fibroblast growth factor 21 (Fgf21), uncoupling protein 1 (Ucp1), cell
death-inducing DNA fragmentation factor, alpha subunit-like effector A (Cidea), and
transmembrane protein 26 (Tmem26), as well as the proteins PRDM16, PGC-1 and
7

ACCEPTED MANUSCRIPT
UCP1. Our findings suggest that PLA promotes the induction of the brown fat-like
phenotype and thus may have potential therapeutic implications for treatment of obesity.
2. Materials and Methods
2.1. Reagents
Mouse 3T3-L1 cells were obtained from the American Type Culture Collection
(Manassas, VA, USA). Dulbecco’s modified Eagle’s medium (DMEM) with high
glucose (11965-092), 100 mM sodium pyruvate, penicillin-streptomycin (liquid, cat#
15070063), and BODIPY® 493/503 were purchased from Thermo Fisher Scientific, Inc.
(Waltham, MA, USA). Fetal bovine serum (FBS) was obtained from Nichirei
Biosciences, Inc. (Tokyo, Japan). 3-Isobutyl-1-methylxanthine (IBMX), insulin, and
dexamethasone were obtained from Sigma-Aldrich Corporation (St. Louis, MO, USA).
Perilipin (D1D8) XP® rabbit monoclonal antibody (mAb) and PPARγ (C26H12) rabbit
mAb were purchased from Cell Signaling Technology (Beverly, MA, USA). UCP1
(4E5) mouse mAb was purchased from Santa Cruz Biotechnology (Santa Cruz, CA,
USA). Antibodies against PGC1 (NBP1-04676) and PRDM16/MEL1 (NBP1-77096)
were purchased from Novus Biologicals LLC (Littleton, CO, USA). Pioglitazone
hydrochloride was obtained from LKT Laboratories, Inc. (St. Paul, MN, USA).
8

ACCEPTED MANUSCRIPT
Rosiglitazone was obtained from Wako Pure Chemical Industries, Ltd. (Osaka, Japan).
Krill oil capsules containing processed krill oil extracted from Antarctic krill
(Euphausia superba) were provided by Sunsho Pharmacetical Co. Ltd. (Shizuoka,
Japan). All reagents were of the highest grade available from commercial sources.
2.2. Isolation of PLA from krill oil
Krill oil (10 g) was dissolved in chloroform and subjected to flash chromatography on a
silica gel column and eluted with a stepwise gradient of methanol in chloroform. The
fractions eluted with 10%–20% methanol were combined and loaded on a Sephadex
LH-20 column (3.2 × 115 cm) and fractionated with 50% methanol in chloroform to
obtain nine fractions (L1–L9). The fractions eluted at a Kd of 1–1.5 were combined (L9,
7.5 mg) and the major components in the fraction were identified to be acyl lactic acids
by nuclear magnetic resonance (NMR) spectroscopy and liquid chromatography–mass
spectrometry (LC–MS) analyses.
2.3. Synthesis of PLA
Benzyl bromide (56.4 mg, 0.33 mmol) and Cs2CO3 (107 mg, 0.33 mmol) were added to
a solution of L-lactic acid (27 mg, 0.3 mmol) in dimethylformamide (1 ml). The
9

ACCEPTED MANUSCRIPT
reaction mixture was stirred for 4 h at room temperature. After the addition of ethyl
acetate, the organic layer was washed with saturated aqueous ammonium chloride and
saturated aqueous sodium chloride. The organic phase was concentrated in vacuo. The
resulting residue was fractionated by silica gel column chromatography to yield benzyl
lactate (39.3 mg, 73% yield).
To synthesize benzyl palmitoyl lactate, the solution of palmitic acid (25.6 mg)
in dichloromethane (5 ml) was treated with 1-ethyl-3-(3-dimethylaminopropyl)
carbodiimide (19 mg) and 4-dimethylaminopyridine (2.4 mg). Benzyl lactate (39 mg)
was then added to the resulting solution and the reaction mixture was stirred overnight
at room temperature and then quenched with saturated aqueous ammonium chloride.
The resulting mixture was extracted with ethyl acetate, and the organic layer was further
washed with saturate aqueous sodium chloride and dried over sodium sulfate. The
organic phase was fractionated by silica gel column chromatography to yield benzyl
palmitoyl lactate (32.8 mg, 55% yield).
To remove the benzyl groups, 10% Pd/C (1 mg) was added to the solution of
benzyl palmitoyl lactate (32.8 mg) in chloroform (5 ml). The resulting solution was
stirred under H2 atmosphere for 40 min at room temperature. The Pd catalyst was
filtered, and the filtrate was concentrated and fractionated by silica gel chromatography
10

ACCEPTED MANUSCRIPT
to yield palmitoyl lactatic acid (7.3 mg, 28% yield).
2.4. Cell culture and treatment
Unless otherwise specified, cell culture and further analyses were conducted at 37°C
under an atmosphere of 5% CO2 and 95% air. 3T3-L1 cells were maintained as
subconfluent cultures in DMEM supplemented with 25 mM glucose, 10% FBS, 1 mM
sodium pyruvate, and 100 U/ml penicillin-100 g/ml streptomycin (medium A). The
3T3-L1 cell line is a widely used model system for the analysis of adipocyte
differentiation. To differentiate into mature adipocytes, these cells were seeded at a
density of 4 × 105 cells/well in 35-mm dishes and then cultured in medium A to
confluency. The cells were incubated for at least 2 days and then cultured for 3 days
with 10 g/ml of insulin, 0.25 g/ml dexamethasone, and 0.5 mM IBMX in medium A
(henceforth called MIX). The cells were further incubated for 4 days in medium A, as
described previously [25]. The differentiated adipocytes were characterized on day 7 by
assessing the intracytoplasmic accumulation of lipid droplets, as indicated by positive
staining with Oil Red O [26].
2.5. Immunofluorescence
11

ACCEPTED MANUSCRIPT
3T3-L1 cells were plated in eight-chamber plastic slides (LAB-TEK II, #154534) at a
concentration of 5 × 103 cells/well/0.3 ml of medium A. The cells were differentiated to
adipocytes by the methods described above. At day 7 after stimulation of differentiation,
the cells were fixed with 4% paraformaldehyde at 37°C for 10 min and then subjected to
permeabilization with 0.5% Triton X-100/Tris-buffered saline (TBS). Afterward, the
cells were washed with TBS containing 0.1% Triton X-100 (TBS-T) three times,
blocked with 2% goat serum in TBS-T for 1 h, and incubated with polyclonal
anti-perilipin, anti-PPAR, anti-PRDM16, and anti-PGC1- antibodies (1:100 dilution)
overnight at 4C, followed by three washes with TBS-T. The cells were then incubated
with Alexa Fluor® 594 secondary antibody goat anti-rabbit immunoglobulin G (1:600
dilution). The cells were stained with DAPI (nuclei) and BODIPY® 493/503 lipid stain
(1 μg/ml of phosphate-buffered saline (PBS); Molecular Probes, Eugene, OR, USA) for
30 min. Finally, the cells were washed with TBS-T and treated with mounting agent.
Images were acquired using the Olympus FV10i confocal microscope (Olympus
Corporation, Tokyo, Japan), which has an automated scanning confocal system and is
equipped with a × 600 water immersion objective 1.345 numerical aperture.
.
2.6. AdipoRed™ assays
12

ACCEPTED MANUSCRIPT
Lipid content was measured using a commercially available kit (AdipoRed™; Lonza
Walkersville, MD, USA) according to the manufacturers’ protocol. 3T3-L1 cells were
seeded at a density of 3 × 103 cells/well of 96-well plates with black walls and clear
bottoms and then cultured in medium A to confluency. The cells were treated with 3 M
pioglitazone, 3 M rosiglitazone, or various concentration of PLA for 7 days. The
medium was changed every 3 days. In contrast, the cells were treated with MIX for 3
days and then cultured in fresh medium. On day 7, the cells were fixed with 4%
paraformaldehyde for 30 min and then washed with PBS (pH 7.4), and 200 μl of PBS
was added to the wells. Then, 5 μl of AdipoRed reagent was added to each well. After
20 min, the plates were placed in the ARVO plate reader (PerkinElmer Life and
Analytical Sciences, Boston, MA, USA), and fluorescence was measured with an
excitation wavelength of 485 nm and an emission wavelength of 535 nm.
2.7. BODIPY staining and quantitation
The size and adiposity of the cells were determined using the fluorescent dye
BODIPY® 493/503. Cells were cultured in six-well plates and differentiated for 7 days.
For the fluorescence-activated cell sorting (FACS) experiment, the cells were incubated
with DMEM containing with 0.25 g/ml BODIPY® 493/503 for 30 min at 37°C in the
13

ACCEPTED MANUSCRIPT
dark and then rinsed twice with PBS (pH 7.4) and trypsinized with phenol red-free
0.05% trypsin-ethylenediaminetetraacetic acid (Invitrogen Corporation, Grand Island,
NY, USA) for 3 min at 37°C. The cells were then suspended in 1 mL of PBS and
immediately put on ice. The samples were analyzed with a FACSCanto II (Becton
Dickinson, Franklin Lakes, NJ, USA) using BD FACSDiva software, version 6.0. The
cell size distribution was determined with a forward scatter (FSC) detector.
2.8. Real-time quantitative polymerase chain reaction (qRT-PCR) analysis
qRT-PCR was performed to determine gene expression levels of target genes in mouse
3T3-L1 cells using an ABI7300 RT-PCR System (Applied Biosystems, Foster City, CA,
USA). The qRT-PCR reactions were performed in 10-μl reaction volumes containing the
following components: 1ꢀμl of cDNA solution, 5ꢀμl of SYBR® Green PCR Master Mix
(Applied Biosystems), 300ꢀnM of each primer (qRT-PCR primer set for adipose
(mouse); Hokkaido System Science Co., Ltd., Sapporo, Japan, or specific primers
synthesized by Eurofins Genomics K.K., Tokyo, Japan), and deionized water. The
sequences of all primers are listed in Table 1. The cDNA amplification conditions were
95°C for 2ꢀmin followed by 40 cycles at 95°C for 15ꢀs and 60°C for 60ꢀs. The expression
levels of the target genes were normalized to that of β-actin, as an internal control.
14

ACCEPTED MANUSCRIPT
Relative expression levels were calculated using the 2−ΔΔCT method. qRT-PCR analysis
was performed in triplicate with a mean of at least three separate experiments.
Table. 1
2.9. Immunoblot analysis
Cells were cultured in six-well plates, differentiated for 7 days, and then trypsinized
with phenol red-free 0.05% trypsin-ethylenediaminetetraacetic acid for 3 min at 37°C.
According to the manufacturer’s instructions of a nuclear extraction kit (Cayman
15

ACCEPTED MANUSCRIPT
Chemicals, Ann Arbor, MI, USA), the detached cells were washed twice with ice-cold
PBS, homogenized in 500ꢀμl of ice-cold complete hypotonic buffer for 15 min,
combined with 100 μl of NP-40, and then centrifuged at 14,000 × g for 30 s at 4°C. The
supernatant containing the cytoplasmic protein fraction was used for determination of
PPAR. The remaining nuclear pellet was resuspended in 100ꢀμl of ice-cold complete
nuclear extract buffer for 30 min and then centrifuged at 14,000 × g for 15 min at 4°C.
The supernatant containing the nuclear fraction was used for determination of PPAR.
For immunoblot analyses of UCP1, cells were lysed in radioimmunoprecipitation assay
buffer. The protein concentration of the supernatant was determined with a
bicinchoninic acid protein assay kit (Takara Bio Inc., Otsu, Japan). Each protein sample
(5 g) was separated by 10% sodium dodecyl sulfate–polyacrylamide gel
electrophoresis and transferred to a polyvinylidene difluoride (PVDF) filter membrane
(Millipore, Billerica, MA), which was subjected to immunoblot analyses with each
primary antibody (anti-PPAR antibody or anti-UCP1 antibody) and horseradish
peroxide-conjugated secondary antibody. The ECL Prime Western Blotting Detection
Reagent kit (GE Healthcare Life Sciences, Marlborough, MA, USA) was used for
visualization. The intensity of each immunoreactive band was measured with Image
Gauge software and an Amersham Imager 600 system (GE Healthcare Life Sciences).
16

ACCEPTED MANUSCRIPT
2.10. Statistical Analysis
The results are expressed as the mean ± standard error of mean (SEM). Prism 6 software
(Graph Pad Software, Inc., San Diego, CA, USA) was used to perform all statistical
analyses. Comparisons between groups were performed by one-way analysis of
variance and the Dunnett’s post-hoc test. Differences were considered significant at P <
0.05.
3. Results
3.1. Identification of active components that induce adipogenesis
To identify the active constituents responsible for the adipogenesis in 3T3-L1 cells, krill
oil was fractionated by silica gel chromatography and Sephadex LH-20 gel filtration,
which resulted in nine fractions (L1–L9). Fraction L9, which was eluted in the latter
part by Sephadex LH-20 column chromatography, showed the most potent adipogenic
activity, as indicated by positive staining with Oil Red O (Fig. 1A). Under these
conditions, fraction L1 exhibited cytotoxicity. Based on the 1H NMR spectrum, the
active compounds contain a common structural unit, lactic acid, of which hydroxyl
group is further acylated with several different fatty acids. The major ion peaks detected
by LC–MS analysis were consistent with the molecular weights of acyl lactic acids
17

ACCEPTED MANUSCRIPT
having different acyl unit compositions, including palmitic acid, oleic acid, EPA, and
DHA. Furthermore, PLA was chemically synthesized from commercially available
palmitic acid and L-lactic acid. The LC–MS analysis confirmed that the major
component of active fraction was PLA (Fig. 1B).
Figure 1. Adipogenic activity of each fraction and palmitoyl lactic acid
18

ACCEPTED MANUSCRIPT
3.2. PLA-induced differentiation into adipocytes
To investigate differentiation of adipocytes by PLA, 3T3-L1 cells were treated with 100
M PLA for 7 days. The PLA-treated cells were accumulated in intracytoplasmic lipid
droplets that were positively stained with Oil Red O (Fig. 2A). The lipid accumulation
was significantly increased by PLA in a dose-dependent manner (Fig. 2B, E, and F).
Conversely, both 100 M lactose and 100 M palmitic acid did not affect accumulation
in 3T3-L1 cells (Fig. 2A). We then examined the effect of PLA on typical expression
patterns of adipocyte markers, including perilipin and fatty acid-binding protein 4
(FABP4; Fig. 2C, D, and 4D). PLA significantly enhanced perilipin expression at both
the mRNA and protein levels (Fig. 2C and D) and enhanced FABP4 expression at the
mRNA level (Fig. 4D). Under these conditions, PLA used for stimulation had no effect
on cytotoxicity, as assessed by the WST-8 cell proliferation assay (Supplementary Fig.
1A). FACS analysis showed that the population of control cells (Fig. 2E) exhibited a
wide heterogeneity in size and low BODIPY fluorescence (Q3 and Q4), consistent with
the undifferentiated state. Upon differentiation by MIX and PLA treatment, the
population of small cells with high BODIPY fluorescence (Q1) increased in a
dose-dependent manner (Fig. 2E and F). PLA treatment did not have a noticeable effect
19

ACCEPTED MANUSCRIPT
on the appearance of Q2 cells. Thus, PLA promoted fat accumulation without increasing
cell size, which is thought to be controlled by a mechanism independent of fat
accumulation. Conversely, cell populations in the undifferentiated state (Q3 and Q4)
were reduced by PLA and MIX treatment, and a particularly large size cell population
(Q4) was significantly reduced. These results indicate that PLA promoted the
accumulation of intracellular lipid droplets of preadipocytes and induced the expression
of the adipogenesis markers perilipin and FABP4. PLA showed potent 3T3-L1
differentiation activity.
20

ACCEPTED MANUSCRIPT
Figure 2. Effect of PLA on adipogenesis of 3T3-L1 preadipocytes
3.3. Like the PPAR agonist, PLA promoted adipogenesis in the presence of
dexamethasone
21

ACCEPTED MANUSCRIPT
Dexamethasone, IBMX, and insulin, are insufficient to cause differentiation
independently, although the mixture of these regents accelerate the differentiation of
3T3-L1 cells (Fig. 4A) [27]. Previous reports demonstrated that a supply of an
exogenous PPAR-agonist accelerated differentiation in combination with
dexamethasone [28]. The promotion of adipogenesis with dexamethasone is a unique
action of the PPAR-agonist [28]. The identification of agents that accelerate
differentiation, provides insights to the biochemical pathways that may function during
adipocyte differentiation [27]. In this study, PLA significantly enhanced the expression
of PPARat both the mRNA and protein levels, and PLA-enhanced PPAR accumulated
in the nucleus (Fig. 3A–D). Thus, we next tested the ability of PLA and the
PPAR-agonists pioglitazone and rosiglitazone to induce adipocyte differentiation in the
system. As demonstrated in Fig. 4A, neutral lipid accumulation was increased by
1.9-fold by treatment with 100 M PLA alone, while accumulation was increased by
5.4-fold by 100 M PLA in the presence of dexamethasone. Furthermore, the
population of adipocytes was analyzed by FACS experiment. When treated with PLA in
the presence of dexamethasone, the population of differentiated adipocytes increased by
1.9-fold, compared with treatment with PLA alone (Fig. 4C). The PLA-induced
adipogenesis in the presence of dexamethasone was phenotypically similar to 3T3-L1
22

ACCEPTED MANUSCRIPT
cells induced by pioglitazone or rosiglitazone (Fig. 4A–C). Moreover, neutral lipid
accumulation was not changed by PLA or the PPAR-agonists in the presence of IBMX
but was increased by both PLA and PPAR-ligand in the presence of insulin (Fig. 4A
and B). When treated with both PLA and the PPAR-agonist in the presence of
dexamethasone, the expression of FABP4, a typical adipocyte marker, significantly
increased, compared with alone treatment (Fig. 4D). PPARγ agonists, such as
thiazolidinediones (TZDs), are known to promote adipogenesis and induce browning in
vivo [29-31]. Thus, we next examined whether PLA affects the brown fat-like
phenotype in 3T3-L1 cells.
23

ACCEPTED MANUSCRIPT
Figure 3. PLA significantly increased PPAR
24

ACCEPTED MANUSCRIPT
Figure 4. PLA significantly increased the adipogenesis in the presence of
dexamethasone
3.4. PLA-induced a brown fat-like phenotype in 3T3-L1 adipocytes
We next conducted a comparative analysis of the expression levels of nine brown/beige
cell-specific genes in 3T3-L1 adipocytes treated with PLA and TZDs. As shown in Fig
25

ACCEPTED MANUSCRIPT
5A, PLA significantly increased the key brown fat markers (PRDM16, PGC-1, and
UCP1) at the mRNA level in 3T3-L1 adipocytes treated with PLA for 7 days, compared
with untreated control cells. Treatment of cells with 100 M PLA produced a 32.8-fold
induction of Prdm16, a 3.5-fold induction of Pgc1a, and a 1.7-fold induction of Ucp1.
TZDs also increased the expression levels of the brown fat markers Prdm16 (93.0-fold
for pioglitazone and 79.9-fold for rosiglitazone), Pgc1a (6.7-fold for pioglitazone and
5.9-fold for rosiglitazone), and Ucp1 (1.5-fold for pioglitazone, 1.3-fold
for rosiglitazone) in 3T3-L1 adipocytes. In contrast, none of these three brown fat
markers were increased in 3T3-L1 white adipocytes induced by MIX (data not shown).
PLA enhanced expression of other key brown/beige cell-specific genes, Cox7a1, Cited1,
Fgf21, Cidea, and Tmem26 (Fig. 5A). The PLA-induced expression profile of the
brown/beige adipocyte-specific genes was similar to that by the TZDs except for
Tmem26 in 3T3-L1 adipocytes (Fig. 5A). These findings have raised the possibility that
PLA promotes the brown fat-like phenotype in 3T3-L1 preadipocytes, like the PPAR
agonist. This notion is supported by the enhanced expression of the key brown cell
marker proteins PRDM16, PGC-1 and UCP1 upon PLA treatment, which was further
confirmed at the cellular level by immunoblotting and immunostaining (Fig. 5B and C).
26

ACCEPTED MANUSCRIPT
Figure 5. Effect of PLA on expression of brown/beige cell-specific genes in
3T3-L1 cells
27

ACCEPTED MANUSCRIPT
Figure 6. PLA induces lipid accumulation in the different manner from
MIX and works antagonistically with MIX in 3T3-L1 cells
28

ACCEPTED MANUSCRIPT
3.5. PLA promoted adiponectin expression and reduced lipid accumulation
PPAR/retinoid X receptors have been shown to directly bind to a functional
PPAR-responsive element in the human adiponectin promoter and to increase promoter
activity [29]. Furthermore, PPAR agonists, such as TZDs, increased adiponectin
synthesis in cultured 3T3-L1 adipocytes, which was associated with increases in
mtDNA content and fatty acid oxidation [33]. In a more recent study of krill oil, serum
adiponectin was increased in animals fed krill oil [34]. Thus, we next examined the
effect of PLA on adiponectin expression at the mRNA level in 3T3-L1 cells. After
treatment of the cells with 100 M PLA for 7 days, Adiponectin expression was
significantly increased (17.7-fold in PLA-treated cells vs. 2.8-fold in MIX-treated cells).
TZDs also increased Adiponectin expression by 19.6-fold in pioglitazone-treated cells
and 15.8-fold in rosiglitazone-treated cells (Fig. 5D). Moreover, total lipid content
significantly decreased in cells treated with both MIX and PLA (MIX-plus-PLA cells),
as compared with those treated MIX only (Fig. 6A). FACS analysis also showed that the
population of cells with high fat content was decreased by treatment with MIX plus
PLA, as compared to the MIX alone, although single PLA treatment increased the
29