Download carboxylesterase 1 plays an essential role in non

CARBOXYLESTERASE 1 PLAYS A PROTECTIVE ROLE AGAINST
METABOLIC DISEASE
A dissertation submitted
to Kent State University in partial
fulfillment of the requirements for the
degree of Doctor of Philosophy
By
Jiesi Xu
May 2016
© Copyright
All rights reserved
Except for previously published materials
Dissertation written by
Jiesi Xu
B.S., Jilin University 2008
Ph.D., Kent State University 2016
Approved by
___________________________________ , Chair, Doctoral Dissertation Committee
Dr. Yanqiao Zhang, M.D., Associate Professor, NEOMED
___________________________________ , Member, Doctoral Dissertation Committee
Dr. John Y.L. Chiang Ph.D., Distinguished Professor, NEOMED
___________________________________, Member, Doctoral Dissertation Committee
Dr. Colleen M. Novak, Ph.D., Associate Professor, Kent State University
___________________________________, Member, Doctoral Dissertation Committee
Dr. Min You, Ph.D., Professor, NEOMED
___________________________________, Member, Doctoral Dissertation Committee
Dr. Eric M. Mintz, Ph.D., Professor, Associate dean, Kent State University
Accepted by
___________________________________ , Director, School of Biomedical Sciences
Dr. Ernest Freeman, Ph.D.
___________________________________ , Dean, College of Arts and Sciences
Dr. James L. Blank, Ph.D
ii
TABLE OF CONTENTS
TABLE OF CONTENTS
LIST OF FIGURES ............................................................................................................v
LIST OF ABBREVIATIONS .........................................................................................viii
ACKNOWLEDGEMENTS ...............................................................................................x
ABSTRACT ......................................................................................................................xi
CHAPTER 1 THE ROLE OF CARBOXYLESTERASE 1 IN NON-ALCOHOLIC
FATTY LIVER DISEASE AND CARBOHYDRATE METABOLISM………………...1
1.1 INTRODUCTION……………........................................................................ 1
1.2 METHODS…………........................................................................................7
1.3 RESULTS .......................................................................................................14
1.3.1 Hepatic Carboxylesterase1 is Induced by Glucose and Regulates
Postprandial Glucose Levels................................................................................. 14
1.3.2 Hepatic Carboxylesterase 1 is Essential for Normal and Farnesoid X
Receptor-Controlled Lipid Homeostasis................................................................25
1.4 DISCUSSION……………………..................................................................50
CHPTER 2 THE ROLE OF CARBOXYLESTERASE 1 IN ALCOHOLIC LIVER
DISEASE………………………………...…................................................................... 56
2.1 INTRODUCTION.......................................................................................... 56
2.2 METHODS..................................................................................................... 59
2.3 RESULTS....................................................................................................... 67
iii
2.4 DISCUSSION ................................................................................................ 96
CHAPTER
3
THE
ROLE
OF
CARBOXYLESTERASE
1
IN
ATHEROSCLEROSIS…………………………............................................................100
3.1 INTRODUCTION.........................................................................................100
3.2 METHODS....................................................................................................103
3.3 RESULTS......................................................................................................105
3.4 DISCUSSION ...............................................................................................111
CHAPTER 4 CONCLUSION…….……………............................................................113
REFERENCES…………………………………………………………………………116
iv
LIST OF FIGURES
Figure 1. Hepatic CES1 is regulated by nutritional status……………………………….15
Figure 2. Hepatic CES1 is regulated by glucose but not insulin………………………...18
Figure 3. ACL is required for glucose-induced hepatic CES1 expression………………21
Figure 4. ACL is required for glucose-mediated acetylation of histones (H3, H4) in the
CES1 chromatin………………………………………………………….…….22
Figure 5. CES1 regulates postprandial levels…………………………………………....24
Figure 6. Hepatic expression of CES1 lowers hepatic triglyceride levels and improves
glucose homeostasis………………………………………………………...26,27
Figure 7. Hepatic expression of CES1 selectively regulates gene expression and has no
effect on lipogenesis or VLDL secretion………………………………………30
Figure 8. Hepatic expression of CES1 increases triglyceride hydrolase activity and
activates PPARα……………………………………………………………….32
Figure 9. Loss of hepatic CES1 causes fatty liver and increased plasma cholesterol
level…………………………………………………………………………….35
Figure 10. Loss of hepatic CES1 induces de novo lipogenesis………………………….37
Figure 11. Hepatic CES1 is regulated by FXR…………………………………………..40
Figure 12. CES1 is a direct FXR target gene…………………………………………….43
Figure 13. Essential roles of hepatic CES1 in FXR-regulated lipid homeostasis………..45
Figure 14. Effects of the FXR agonist OCA (INT-747) on lipid and glucose homeostasis
in C57BL/6 mice……………………………………………………………….47
Figure 15. Effects of the FXR agonist OCA on lipid homeostasis in ob/ob mice……….49
v
Figure 16. CES1 and HNF4 expressions are reduced in patients with alcoholic
steatohepatitis and in mouse and mouse primary hepatocytes treated with
ethanol………………………………………………………………………….68
Figure 17. CES1 is regulated by HNF4………………………………………………...71
Figure 18. CES1 is a direct target of HNF4……………………………………………72
Figure 19. Over-expression of hepatic CES1 protects against alcohol-induced triglyceride
accumulation in AML12 cells………………………………………………….74
Figure 20. Hepatic CES1 deficiency alters plasma lipid levels in response to alcohol
challenge……………………………………………………………………….75
Figure 21. Hepatic CES1 deficiency exacerbates alcohol-induced hepatic steatosis……77
Figure
22.
Hepatic
CES1
deficiency
exacerbates
alcohol-induced
liver
inflammation…………………………………………………………………...80
Figure 23. Global deletion of CES1 does not exacerbate alcohol-induced hepatic
steatosis...………………………………………………………………………83
Figure 24. Global deletion of CES1 exacerbates alcohol-induced liver inflammation….85
Figure
25.
Global
deletion
of
CES1
increases
MCD
diet-induced
liver
inflammation………………………………………………………..………….88
Figure 26. CES1 deficiency does not significantly change fibrogenic gene
expressions……………………………………………………………………..88
Figure 27. Global deletion of CES1 does not exacerbate MCD-diet induced
fibrosis….............................................................................................................89
Figure 28. Global deletion of CES1 increases hepatic acetaldehyde level and oxidative
stress……………………………………………………………………………92
vi
Figure 29. Global deletion of CES1 does not change mRNA levels of genes involved in
fatty acid metabolism…………………………………………………………..95
Figure 30. Macrophage cholesterol efflux……………………………………………...102
Figure 31. Global deletion of CES1 results in increased lipid accumulation in
macrophages………………………………………………………………….105
Figure 32. Loss of hepatic CES1 increases lipid contents in ApoE mice……………106
Figure 33. Loss of hepatic CES1 shows atherosclerotic lipid profile…………………..108
Figure 34. Loss of hepatic CES1 aggravates atherosclerosis in ApoE mice…………110
vii
LIST OF ABBREVIATIONS
ABCA1
ATP binding cassette sub-family A1
ABCG5
ATP binding cassette sub-family G5
ACC
acetyl-CoA carboxylase
ACL
ATP citrate lyase
AKT
protein kinase B
ALD
alcoholic liver disease
APOB
apolipoprotein B
CD36
cluster of differentiation 36
CES1
carboxylesterase 1
ChIP
chromatin immunoprecipitation
CPT
carnitine palmitoyltransferase
DGAT
diacylglycerol O-acyltransferase
EMSA
electrophoretic mobility shift assay
FAS
fatty acid synthase
FFA
free fatty acid
FPLC
fast protein liquid chromatography
FXR
farnesoid X receptor
GCK
glucose kinase
G6Pase
glucose-6 phosphatase
HDL
high density lipoprotein
HMGCS
HMG-CoA synthase
viii
HMGCR
HMG-CoA reductase
HNF4
hepatocyte nuclear factor 4
IL-1
interleukin 1 
IL-6
interleukin 6
LDL
low density lipoprotein
L-PK
liver pyruvate kinase
MCP
monocyte chemoattractant protein
MDA
malondialdehyde
MTP
microsomal triglyceride transfer protein
NAFLD
non-alcoholic fatty liver disease
PPAR
peroxisome proliferator-activated receptor 
PDK
pyruvate dehydrogenase kinase
PEPCK
phosphoenolpyruvate carboxykinase
PGC1
peroxisome proliferator-activated receptor  coactivator 1
ROS
reactive oxygen species
SREBP1C
sterol regulatory binding protein 1c
TC
total cholesterol
TNF
tumor necrosis factor 
TG
triglyceride
VLDL
very low density lipoprotein
ix
ACKNOWLEDGEMENTS
I would like to express my gratitude to all the people who gave me help and
support throughout the course of this work. I would especially like to thank my advisor
Dr. Yanqiao Zhang for his invaluable guidance, caring and immense knowledge. I would
also like to thank him for pushing me father than I thought I could go and providing
financial support for my research. I wish to express my sincere thank to Dr. John Chiang,
Dr. Colleen Novak and Dr. Min You for giving me insightful suggestions regarding my
research. Their guidance and encouragements help my research from various perspectives.
I would like to acknowledge and thank Dr. Novak for the help she has given with
CLAMS analysis.
My special thanks go to Dr. Yang Xu and Dr. Preeti Pathak for their selfless,
enormous help. I am so grateful to Dr. Xu for teaching me the techniques that are
important for completing this study; and to Dr. Pathak for lending me her shoulder to cry
on.
This dissertation is dedicated to my mother who is always there for me and
experiences all the peaks and valleys with me during this journey. Even so far away, I
feel her love and care every minute in my life.
x
ABSTRACT
Jiesi Xu
Department of Biomedical Sciences
Kent State University, 2016
Advisor Yanqiao Zhang
Carboxylesterase 1 (CES1) is a phase I drug metabolizing enzyme which is shown
to hydrolyze ester and amide-containing drugs and prodrugs. It also hydrolyzes
triglyceride and cholesteryl ester. The present studies show that glucose induces CES1
expression, and that the induction of CES1 expression, in turn, reduces plasma glucose
level, likely through increasing insulin sensitivity. We then demonstrate that overexpression of hepatic CES1 reduces hepatic triglyceride level and promotes fatty acid
oxidation. In contrast, loss of hepatic CES1 induces hepatic steatosis by increasing sterol
regulatory-element binding protein (SREBP) processing and lipogenesis. We also find
that CES1 is a farnesoid X receptor (FXR) target gene, and that activation of FXR
reduces hepatic and plasma triglyceride levels through, at least in part, inducing CES1.
These results suggest that CES1 plays a protective role against non-alcoholic fatty liver
disease (NAFLD). We further show that CES1 is inhibited by alcohol, and is a hepatocyte
nuclear factor 4 (HNF4) direct target. Using liver specific Ces1 deficient and Ces1
mice together with chronic-binge alcohol feeding, we find that CES1 deficiency
exacerbates alcohol-induced liver inflammation. CES1 deficiency also results in
increased hepatic acetaldehyde level, elevated reactive oxygen species (ROS) level and
enhanced lipid peroxidation, suggesting that CES1 plays a protective role against
xi
alcoholic liver disease (ALD). Lastly, we demonstrate that loss of hepatic CES1
aggravates western diet-induced atherosclerosis in ApoE mice. These exciting results
lead us to conclude that CES1 plays an essential role in lipid and carbohydrate
metabolism, and that it also protects against NAFLD, ALD and atherosclerosis. Targeting
CES1 may be a plausible strategy for treating metabolic disease.
xii
CHAPTER 1: THE ROLE OF CARBOXYLESTERASE 1 IN NON-ALCOHOLIC
FATTY LIVER DISEASE AND CARBOHYDRATE METABOLISM
1.1 INTRODUCTION
Nonalcoholic fatty liver disease (NAFLD), one of the most common liver diseases
worldwide, encompasses a spectrum of liver disorders ranging from simple hepatic
steatosis, nonalcoholic steatohepatitis (NASH), to cirrhosis. NAFLD is often associated
with insulin resistance, obesity, type 2 diabetes and dyslipidemia, and is often a hepatic
manifestation of metabolic disorders [1]. The worldwide prevalence of NAFLD and its
subtype nonalcoholic steatohepatitis (NASH) range from 6.3% to 33% and from 3% to
5% in the general population, respectively [2]. Non alcoholic fatty liver (NAFL) is
defined as the presence of hepatic steatosis with no evidence of hepatocellular injury in
the form of ballooning of the hepatocytes. In contrast, NASH is defined as the presence
of hepatic steatosis and inflammation with hepatocyte injury (ballooning) with or without
fibrosis. Liver fibrosis is characterized by over-stimulation of liver stellate cells and
excessive accumulation of extracellular matrix proteins, thus results in portal
hypertension, cirrhosis and liver failure. Cirrhosis, featured by the development of broad
collagen bands that form nodules, is irreversible and predicted to become the most
common
indication
for liver transplantation. The estimated risk of progression to cirrhosis in patients with
NASH and fibrosis is approximately 20%, compared with only 4% risk of progression in
1
patients with simple hepatic steatosis [3].
Our knowledge of the pathogenesis of NAFLD has greatly advanced. Risks
factors including dietary fat, genetic predisposition, gut microbiome and metabolic
disorders such as type 2 diabetes, obesity and cardiovascular diseases can often
contribute to the development of NAFLD. Hepatic steatosis is a hallmark in NAFLD and
is often associated with disrupted glucose and lipid homeostasis. In human body,
approximately 60% of the triglyceride accumulated in the liver is derived from plasma
non-esterified fatty acid (NEFA) pool, 25% derived from hepatic de novo lipogenesis and
15% derived from dietary intake [3]. Adipose tissue is the largest supplier of fatty acids
and generates multiple signals that alter lipid and glucose metabolism. Free fatty acids
(FFAs), released from adipose tissue, can be used for triglyceride synthesis or broken
down in response to energy demands. Peripheral insulin resistance, a predisposing factor
for hepatic steatosis, enhances adipocyte lipolysis and increases the efflux of free fatty
acids to the plasma NEFA pool [3,4]. Although hepatic de novo synthesis only accounts
for 20% of the lipid pool in the liver, alteration of de novo synthetic gene expressions are
often present in patients with metabolic diseases. Hyperinsulinemia and hyperglycemia
induce sterol regulatory element-binding protein 1c (SREBP-1c) and carbohydrate
response element binding protein (ChREBP) in the liver, subsequently activate lipogenic
genes and increase hepatic de novo lipogenesis [2].
The mechanism underlying the disease progression from simple hepatic steatosis
to NASH, and to more advanced fibrosis and cirrhosis remains elusive. Proinflammatory
cytokines in addition to hepatic steatosis imposes a second “hit” to incite hepatic injury,
through sensitizing kupffer cells and triggering apoptosis and necrosis. Oxidative stress,
2
mitochondria dysfunction, endoplasmic reticulum stress, lipotoxicity and apoptotic
pathways advance liver damage toward fibrosis and cirrhosis [4]. It is reported that less
than 4% of individuals with steatosis progress to cirrhosis, as compared with 20% of
individuals with NASH [3].
Currently, no pharmacological therapy is proven to be effective to alter the nature
history of NAFLD. Thus, weight loss is often recommended to patients to improve liver
biochemistries and histology [5]. The pharmaceutical intervention for NAFLD is directed
toward
correction
of
its
risk
factors.
Insulin-sensitizing
drugs
including
thiazolidinediones, rosiglitazone, pioglitazone, and metformin are shown to improve
insulin sensitivity, serum alanine transaminase (ALT), and histologic features in the
livers of NASH patients without diabetes [6,7,8]. Vitamin E decreases oxidative stress,
which provides a rationale for its use not only in patients with NASH but also those with
liver fibrosis [9]. Pioglitazone combined with vitamin E is more effective to reverse
histologic abnormalities than vitamin E alone [5]. Anti-inflammatory (corticosteroids),
and antioxidants (silymarin, phosphatidylcholine, and s-adenosyl-L-methionine) are
shown to attenuate experimental liver fibrosis [10]. In addition, pentoxifylline
(phosphodiesterase inhibitor), amiloride (Na+/H+ pump inhibitor), S-farnesylthiosalicylic
acid (Ras antagonist), thiazolidinediones (peroxisome proliferator-activated receptor 
ligands), and Renin-angiotensin inhibitors are shown to have antifibrotic effect [10].
Patients with end-stage liver disease are subject to liver transplantation. However,
transplantation per se does not correct some metabolic syndrome. Thus, weight
management and correction of hyperglycemia and hyperlipidemia become crucial goals
for managing NAFLD.
3
Farnesoid X receptor (FXR) is a key element in regulation of bile acid synthesis,
glucose and lipid metabolism, energy homeostasis and inflammation pathways. It is a
member of the nuclear receptor superfamily and is known to be activated by bile acids
[11]. The beneficial effects of activated FXR have been well documented [12,13,14,15].
It lowers hepatic and plasma triglyceride level and plasma cholesterol level, and
improves insulin sensitivity [14,15].
These beneficial effects of activation of FXR
provide rationales for its use in treating NAFLD. Currently, INT747, a FXR agonist, is
undergoing clinical trial for patients with NASH. The hepatoprotective mechanism of
FXR agonism lies in two hypotheses: 1) activation of hepatic FXR reduces bile acid
synthesis, increases hepatic content of cholesterol, and reduces SREBP1 activity and
triglyceride synthesis [16] 2) activation of intestinal FXR increases fibroblast growth
factor (FGF)15 secretion, which in turn improves hepatic steatosis and insulin sensitivity
[17,18].
In addition, FXR agonism decreases gluconeogenesis, increases glycogen
synthesis and increases insulin sensitivity, which leads to decreases in the hepatic content
of lipids and overall circulating glucose [19]. Interestingly, disruption intestinal FXR
increases glucagon-like peptide-1 (GLP-1), which leads to increased insulin sensitivity
[20]. Activation of FXR also reduces diet-induced weight gain, and enhances energy
expenditure and browning of white adipose tissue [18,21]. More and more FXR target
genes have been reported. Recent studies demonstrate that the hepatoprotective effect of
FXR results from activating its target genes, including small heterodimer partner (SHP)
[16], PPAR, and Apolipoprotein CII (ApoC-II) [23], and FGF15 [18]. To date, the
tissue specific function of FXR in regulation of lipid and glucose metabolism is still
obscure. Our group finds that INT747 enhances reverse cholesterol transport through
4
activation of hepatic FXR but not intestinal FXR and subsequent inhibition of cholesterol
7-hydroxylase (Cyp7A1), reduction of bile acids pool size and reduction of intestinal
cholesterol absorption. In 2014, Gonzalez, et.al. reported that intestine specific Fxr
disruption decreases hepatic triglyceride accumulation, through lowering circulating
ceramides, which in turn downregulate hepatic SREBP1C and subsequently decrease de
novo lipogenesis [24].
Carboxylesterase 1 (CES1) is a drug-metabolizing enzyme that is highly
expressed in the liver but also to a lesser extent in the intestine, macrophages and other
tissues [25]. It is a member of mammalian carboxylesterases family which is localized in
the endoplasmic reticulum and catalyzes the hydrolysis of a variety of ester and amidecontaining chemicals, drugs (including prodrugs), and endogenous compounds (including
acyl-glycerols, long-chain acyl-carnitine and long-chain acyl-coenzyme A ester), to their
respective free acids [26]. Drug-metabolizing enzymes, present predominantly in the liver,
are involved in detoxification and biotransformation of both endogenous and exogenous
compounds to hydrophilic products to facilitate their elimination [27]. CES enzymes
catalyzes phase I drug-metabolizing reactions for angiotensin-converting enzyme
inhibitors (temocapril, cilazapril, quinapril, and imidapril) [28,29,30], anti-tumor drugs
(CPT-11 and Capecitabine) [31,32,33,34,35,36] and narcotics (cocaine, heroin and
meperidine) [37,38]. The gene structures of CES family reveal that the murine CES gene
is located on the minus strand of chromosome 8 at 8C5 in a cluster of six CES genes that
span 260.6kb in total, and that CES families share several common binding sites for
transcription factors in the promoter region, suggesting that orthologous CES genes have
evolutionally conserved transcriptional regulatory patterns [39].
5
The physiological functions of CESs have been well documented. So far, CESs
families have been shown to be involved in metabolizing a number of drugs and prodrugs
such as anti-coagulant [40,41], anti-cancer [33,35,42,43,44], anti-virus[45,46,47] and
anti-hypertensive[48,49,50] drugs. CESs possesses triglyceride (TG) and cholesteryl ester
(CE) hydrolase activity, and acyl-coenzyme A: cholesterol acyltransferase activity
[51,52,53]. Ablation of hepatic CES3 reduces very low-density lipoprotein (VLDL)
assembly, decreases plasma TG and cholesterol (CHOL) levels without affecting glucose
tolerance [54]. However, global ablation of CES3/TGH leads to improved systemic
glucose clearance and insulin sensitivity, decreased pancreatic islet size and decreased
expression of hepatic gluconeogenesis-related genes [55]. CES1 is robustly expressed in
human THP-1 monocytes/macrophages, and loss of macrophagic CES1 enhances
retention of intracellular cholesteryl esters and leads to a “foamy” phenotype, suggesting
that CES1 is involved in the development of atherosclerosis [56]. Over-expression of
human CES1 in the macrophage leads to increased CE hydrolysis [51], elevated
mobilization of cytoplasmic CE [57], enhanced free cholesterol (FC) efflux [58] and
attenuation of atherosclerosis in Ldlrmice [51]. The TG and CE hydrolase property of
CES1 (ESx) makes it an important target in treating NAFLD [52]. In 2012, Lehner et. al.
reported that Ces1mice develop obesity, fatty liver, hyperinsulinemia, and increased
cholymicron production [59,60].
In this chapter, the role of CES1 in regulating NAFLD and carbohydrate
metabolism is delineated.
6
1.2 METHODS
Mice, Diets, and Ligands.
C57BL/6 mice, leptin deficient (ob/ob) mice, leptin receptor deficient (db/db)
mice, and Fxr mice were purchased from the Jackson Laboratories (Bar Harbor, ME).
High fat/high cholesterol (HFHC) diet (40% kcal from fat, 1.5% cholesterol) was
purchased from Research Diets (cat #D12108, New Brunswick, NJ). For FXR ligands
treatment, FXR agonists GW4064 (30mg/kg, twice a day) and OCA (INT-747)
(30mg/kg/d) were administered to Fxr wild type and Fxrmice by gavage. For glucose
treatment, C57BL/6 mice were fasted for 16 h, and 40% glucose (8g/kg) was
administered twice with 3 h interval through oral gavage. Unless otherwise stated, male
mice were used and all mice were fasted for 5-6 hours prior to euthanization. All the
animal studies have been approved by the Institutional Animal Care and Use Committee
at Northeast Ohio Medical University.
Real-Time PCR.
RNA was isolated using TRIzol Reagent (Invitrogen, Carlsbad, CA). Messenger
RNA (mRNA) levels were determined by quantitative reverse-transcription polymerase
chain reaction (qRT-PCR) on a 7500 real-time PCR machine from Applied Biosystems
(Foster City, CA) by using SYBR Green supermix (Roche, Indianapolis, IN). Results
were calculated using Ct values and normalized to 36B4 mRNA level.
Lipid and lipoprotein analysis.
7
Approximately 100 mg liver was homogenized in methanol and lipids were
extracted in chloroform/methanol (2:1 v/v) as described [61]. Hepatic triglyceride and
cholesterol levels were then quantified using Infinity reagents from Thermo Scientific
(Waltham, MA). Hepatic fatty acid profile was quantified using gas chromatography
(GC)-mass spectrometry at the Mouse Metabolic Phenotyping Center (MMPC) of Case
Western Reserve University (Cleveland, OH). Hepatic total free fatty acids and free
cholesterol were quantified using kits from BioVision (Milpitas, CA). Plasma lipid and
glucose levels were also determined using Infinity reagents. Briefly, after 100 mL plasma
was injected, lipoproteins were run at 0.5 mL/min in a buffer containing 0.15 M NaCl,
0.01 M Na2HPO4, 0.1 mM EDTA, pH 7.5, and separated on a Superose 6 10/300 GL
column (GE Healthcare) using the BioLogic DuoFlow QuadTec 10 System (Bio-Rad,
Hercules, CA). A 500-mL sample per fraction was collected.
Adenovirus.
Ad-Ces1-GFP was constructed by cloning mouse Ces1 cDNA into pAd-shuttleIRES-hrGFP vector (Stratagene, CA) as described previously [62]. To generate
adenovirus expressing small hairpin RNA against Ces1 (Ad-shCes1), oligonucleotides
were designed using BLOCK-iT™ RNAi Designer (Invitrogen, CA), annealed, and
ligated to pEnter/U6 vector (Invitrogen, CA). Adenovirus was then generated following
the instructions provided by Invitrogen. Three different shRNA oligonucleotides against
murine Ces1 were designed. The sequences that produced the most inhibitory effect on
endogenous
Ces1
expression
8
are:
5’-
GCTGATTCCAGCAGCTATTGACGAATCAATAGCTGCTGGAATCAGC-3’
(top
strand) and 5’-GCTGATTCCAGCAGCTATTGATTCGTCAATAGCTGCTGGAATCA
GC-3’ (bottom strand). Adenoviruses expressing ChREBP and shAcl have been
described previously [63,64]. All the adenoviruses were grown in 293A cells and purified
by cesium chloride gradient centrifugation. About 1-2x109 plaque formation units (pfu)
of adenoviruses were transfused into each mouse intravenously. Unless otherwise stated,
7 days post infection, mice were fasted for 5-6 h and then euthanized.
Primary hepatocyte isolation.
Mouse primary hepatocytes were isolated as described [65,66]. Mice were
anaesthetized by intraperitoneal injection of 50 mg/kg pentobarbital. The portal vein was
cannulated with a 23-gauge plastic cannula. Mouse livers were perfused with Hank’s
Balanced Salt Solution (HBSS, cat#14170-112, Thermo Fisher Scientific) with 0.19g/L
EDTA. Simultaneously, the inferior vena cava was cut open. Subsequently, livers were
perfused with HBSS, calcium, magnesium buffer (cat#14025092, Thermo Fisher
Scientific) with 0.8mg/mL Collagenase from Clostridium histolyticum type IV (Sigma, St.
Louis, MO). Primary hepatocytes were released and collected in a 50 mL centrifuge tube.
After centrifugation at 50g for 5 minutes and washing with DMEM, cells were cultured
in 6-well plate pre-coated with 0.1% gelatin in a 2 mL of DMEM+10% FBS.
Mutagenesis and Transient Transfection Assays.
The mutant pGL3 promoter-luciferase construct was generated using a
QuickChange Site-directed Mutagenesis kit from Agilent (Santa Clara, CA). HepG2 cells
9
were plated in a 24-well plate and cultured in DMEM containing 10% FBS. Transient
transfections were performed in triplicate as described [67]. Briefly, pGL3-Ces1
luciferase reporter constructs were transfected into HepG2 cells together with plasmids
expressing FXR or RXR, followed by treatment with either vehicle or GW4064 (1 M).
After 36 h, luciferase activities were determined and normalized to -galactosidase
activity. For glucose-induced CES1 expression, pGL3-Ces1 luciferase reporter constructs
were transfected into HepG2 cells, followed by treatment with either 5.5 mM glucose or
27.5 mM glucose. To determine the effect of over-expression of CES1 on PPAR
activity, a CES1-expression plasmid was co-transfected with the 3xPPRE-luc plasmid,
and luciferase activity was determined as described above.
ChIP assay:
Chromatin immunoprecipitation (ChIP) assays were performed as described
previously [68] and following the manufacturer’s instructions (Millipore, Bilerica, MA).
200 mg liver for each sample was used for ChIP assays. Antibodies against acetyl-H3K9,
acetyl-H4 and FXR (Cell Signaling Technology, Danvers, MA) were used to immunoprecipitate chromatin. Non-immune IgG was used as a measure of nonspecific
background in immunoprecipitation. Chromatin purified from 10% sonicated tissue lysate
was used as “input”. Real-time PCR was performed to test the chromatin enrichment on
CES1 promoter region. ChIP assays were performed in triplicates.
Electrophoretic Mobility Shift Assay (EMSA).
10
Oligonucleotides containing the putative FXR response element (FXRE; DR-5; 218 bp) were annealed, and EMSA and competition studies were performed as previously
described [68]. The wild-type oligonucleotide sequence used for EMSA was ATG TAA
GAT GTT CCT TGG TTA GTT TAT GGA CCT CTG TTA TCT GAG AGC TGT CCA
ATG G (top strand). The mutant oligonucleotide sequence was ATG TAA GAT GTT
CCT TAA TTA GTT TAT GCA AAT CTG TTA TCT GAG AGC TGT CCA ATG G
(top strand) (mutation sites are underlined).
Western Blot Assay.
Western blot assays were performed using whole liver lysates [62] or nuclear
lysates of the liver samples as described previously.[69] CES1 antibody was purchased
from Abcam (Cambridge, MA, USA). -actin antibody was from Novus Biologicals
(Littleton, CO). p-AKT(ser473) and AKT were from Millipore (Billerica, MA). SREBP-1
antibody was from Novus Biologicals (Littleton, CO). SREBP-2 antibody was from
Cayman Chemicals (Ann Arbor, MI). Histone antibody was from Cell Signaling
(Beverly, MA).
VLDL Secretion.
C57BL/6J mice were injected intravenously with specific adenoviruses. On day 6,
these mice were fasted overnight, followed by intravenous injection of Tyloxapol (500
mg/kg). Blood was taken at indicated time points and plasma TG levels were determined.
VLDL secretion rate was determined as described [70].
11
Hepatic Lipogenesis.
Mice were fasted for 4 h and then injected intraperitoneally with 2H2O (20-30
l/g). After 4 h, liver and plasma were snap-frozen in liquid nitrogen. The newly
synthesized palmitate, triglycerides, and cholesterol were measured by mass spectrometry
at MMPC of Case Western Reserve University.
TGH Activity Assay.
Cells were lysed in lysis buffer containing 50mM Tris, pH 7.4, 0.25M Sucrose,
and 1mM EDTA, 1 mM dithiothreitol, 50 mM NaF and protease inhibitor cocktail
(Sigma). After spun at 14,000 x g, the supernatant was used for TGH activity assay. For
animals, liver was harvested after a 6-h fast. Liver was homogenized in lysis buffer and
then spun at 800 x g to remove cell debris. The supernatant was then centrifuged at
100,000 × g for 1 h. The cytosolic portion and microsome portion were used for TGH
assay separately.
In brief, 100 l liver extracts (100 μg of protein) were incubated at 37 oC with 100
l substrates containing 0.15 mM cold triolein, 0.32 M [3H]triolein, 10 M egg yolk
lecithin, 100 M sodium taurocholate, 1 mM dithiothreitol, and 50 mM potassium
phosphate
(pH
7.4).
After
1
h,
the
reaction
was
stopped
by
3.75
ml
methanol:chloroform:heptane (10:9:7) and 1 ml of 0.1 M potassium carbonate/0.1M boric
acid. After centrifuge at 800 x g, 1 ml top phase is used for counting radioactivity using a
liquid scintillation counter.
Fatty Acid Oxidation.
12
AML12 cells were cultured in DMEM containing 10% FBS in 12-well dishes and
infected with either Ad-GFP or Ad-Ces1. After 48 h, the media were removed and
washed with 1XPBS. The cells were then cultured in DMEM containing 0.5% fatty acidfree BSA, 0.5 Ci [3H]palmitate and 500 M cold palmitate. Fatty acid oxidation was
performed as described [71]. Briefly, after incubation for 3 h, the supernatant was
collected. 0.1 ml supernatant was added to a round-bottomed Eppendorf tube containing
0.9 ml of charcoal slurry. The samples were left at room temperature for 30 min with
intermittent shaking (at least once every 5 minutes), and then centrifuged at 13,000 rpm
for 15 minutes. 0.2 ml of the supernatant was carefully taken out and added to a
scintillation vial containing 2.8 ml of scintillation liquid and the radioactivity was
determined on a liquid scintillation counter.
Statistical Method:
The data were analyzed statistically using unpaired Student’s t-test (two-tailed)
and ANOVA (for more than two groups), followed by a post hoc Newman-Keuls test.
The data were expressed as mean±SE. Only p<0.05 was considered statistically
significant.
13
1.3 RESULTS
1.3.1 Hepatic Carboxylesterase 1 is Induced by Glucose and Regulates
Postprandial Glucose Levels.
Hepatic CES1 is regulated by nutritional status.
To determine whether nutritional status affects CES1 expression, we first
determined hepatic CES1 expression in diabetic mice. Our data indicated that hepatic
Ces1 mRNA (Figure 1A, B) and protein (Figure 1C) levels were significantly induced in
both type 2 diabetic ob/ob mice and db/db mice. In streptozotocin (STZ)-treated mice, a
type 1 diabetic mouse model, hepatic Ces1 mRNA levels were induced by >7 fold
(Figure 1D). Peroxisome proliferator-activated receptor gamma coactivator-1  (PGC-1)
and phosphoenolpyruvate carboxykinase (PEPCK) served as positive controls. To
investigate whether hepatic CES1 expression is affected by a Western diet feeding, we
fed C57BL/6 mice a high fat/high cholesterol (HFHC) diet; the data show that HFHC diet
feeding did not change hepatic Ces1 expression (Figure 1E). ATP-binding cassette (ABC)
transporter A1 (ABCA1) and ABC transporter G5 (ABCG5) served as positive controls.
Finally, we investigated the effect of fasting on hepatic CES1 expression. The data of
Figure 1F show that fasting for 8 or 24 hours caused a reduction in hepatic CES1 protein
levels. Overall, these data indicated that hepatic CES1 is regulated by nutritional status
and glucose may induce hepatic CES1 expression.
14
Figure 1. Hepatic CES1 is regulated by nutritional status. (A-C) Hepatic mRNA
levels in ob/ob (A) and db/db mice (B) mice were determined by qRT-PCR and protein
levels determined by Western blot assays (C) (n=4-6 mice per group). (D) C57BL/6 mice
were treated with either vehicle (0.1 M sodium citrate, pH 4.5) or streptozotocin (STZ)
(50 mg/kg/d) for 5 days. Seven days after STZ treatment, mice were euthanized and
15
hepatic mRNA levels were quantified (n=5 mice per group). (E) Wild-type mice were fed
a chow or high fat/high cholesterol (HFHC) diet (21% fat, 1.5% cholesterol) for 3 weeks
and hepatic mRNA levels were determined (n=8 mice per group). (F) C57BL/6 mice
were fed a chow diet, or fasted for 3, 8, 24 h, or fasted for 24 h followed by refed for 24 h
(n=5 mice per group). Hepatic protein levels were determined. Pgc-1, peroxisome
proliferator-activated receptor gamma coactivator-1. Abca1, ATP-binding
(ABC) transporter A1. Abcg5, ABC transporter G5.
carboxykinase. *p<0.05, **p<0.01
16
cassette
Pepck, phosphoenolpyruvate
Hepatic CES1 is regulated by glucose but not insulin
To test our hypothesis that glucose induces hepatic CES1 expression, saline or
glucose (8g/kg) were administered to C57BL/6 mice twice with 3 hours interval via oral
gavage. Mice were sacrificed 3 hours after second oral gavage and hepatic CES1
expression was determined. Hepatic Ces1 mRNA level (Figure 2A) and protein levels
(Figure 2B, C) were induced by ~2 fold in response to glucose stimulation. In contrast,
insulin did not induce hepatic CES1 expression (Figure 2D).
In mouse primary hepatocytes, high glucose (27.5 mM) induced hepatic Ces1
mRNA expression (Figure 2E), suggesting that glucose can directly regulate CES1
expression. To test whether glucose can directly regulate Ces1 promoter activity,
Transient transfection assays and luciferase reporter assays were performed using a serial
of luciferase reporter constructs with 5’-deletions. The data show that glucose stimulated
Ces1 promoter activity through a region between 75 bp and 150 bp upstream of the
transcription start site (Figure 2F). Collectively, the data of Figure 2 demonstrate that
glucose induces CES1 expression both in vivo and in vitro.
17
Figure 2. Hepatic CES1 is regulated by glucose but not insulin. (A-C) C57BL/6 mice
were fasted for 16 h, followed by gavage with saline or glucose (8g/kg) (n=6 mice per
group). Hepatic mRNA levels (A) and protein levels (B) were determined. Hepatic CES1
protein levels were quantified (C). L-pk serves as a positive control in (A). (D) C57BL/6
mice were fasted for 16 h, followed by i.p. injection of either saline or insulin (0.8
18
units/kg) (n=5 mice per group). After 3 h, mice were euthanized. Srebp-1c serves as a
positive control. (E) Mouse primary hepatocytes were isolated and cultured in dulbecco’s
modified eagle medium (DMEM) plus 10% fetal bovine serum (FBS) overnight,
followed by serum-free fasting for 8 h. Cells were then treated with either normal (5.5
mM) or high (27.5 mM) glucose for additional 24 h prior to quantification of mRNA
levels. Fas serves as a positive control. (F) CES1 promoter-luciferase constructs were
transfected into HepG2 cells, then treated with 5.5 mM or 27.5 mM glucose. After 36 h,
luciferase activity was determined. Srebp-1c, sterol response element binding protein-1c.
L-pk, liver type pyruvate kinase. Fas, fatty acid synthase. RLU, relative luciferase units.
*p<0.05, **p<0.01
19
ACL is required for glucose-induced hepatic CES1 expression
ChREBP is suggested to be the principal mediator of glucose metabolism in the
liver [72,73,74]. Absence of ChREBP leads to the failure of glucose to induce hepatic
glycolytic gene (L-PK) and lipogenic genes (ACC and FAS) [75]. However, overexpression of ChREBP in the liver had no effect on hepatic Ces1 expression (Figure 3A),
suggesting that glucose induces hepatic CES1 expression independent of ChREBP. It has
been shown that ACL is required for glucose-mediated histone acetylation and gene
activation [76,77]. Thus, we investigated the role of ACL in glucose-mediated hepatic
CES1 expression. Adenovirus-mediated expression of Acl shRNA reduced hepatic Acl
mRNA levels by >85% (Fig. 3B). Interestingly, glucose induced hepatic CES1 mRNA
and protein expression in the control mice but not in Acl-deficient mice (Figure 3C-E).
Liver-type pyruvate kinase (L-PK), a gene responsive to glucose stimulation, served as a
positive control (Figure 3F). Hence, the data of Figure 3 indicate that ACL is required for
glucose to induce hepatic CES1 expression.
20
Figure 3. ACL is required for glucose-induced hepatic CES1 expression. (A)
C57BL/6 mice were injected i.v. with adenovirus expressing GFP or ChREBP. After 5
days, hepatic mRNA levels were determined by qPCR (n=7 mice per group). (B-F)
C57BL/6 mice were injected with adenovirus expressing shLacZ or shAcl (n=6 mice per
group). After 5 days, mice were gavaged with either saline or glucose (8g/kg). Hepatic
mRNA levels of Acl (B), Ces1 (C) and L-pk (F) were determined. Hepatic protein levels
21
were determined by Western blot assays (D) and CES1 protein levels quantified (E).
*p<0.05, **p<0.01
ACL is required for glucose-mediated acetylation of histones (H3, H4) in the Ces1
chromatin
Several lines of evidence have shown that glucose may regulate gene expression
via epigenetic modifications [76,78,79,80]. ACL converts glucose-derived citrate to
acetyl-CoA, which subsequently serves as substrate for histone acetyltransferase for
acetylation of H3 and H4 tails. Our data showed that glucose increased the acetylation of
histone 3 and histone 4 in the Ces1 chromatin and these effects were abolished in Acldeficient mice (Figure 4A, B), indicating that ACL is required for glucose-mediated
acetylation of histones (H3 and H4) in the Ces1 chromatin.
Figure 4. ACL is required for glucose-mediated acetylation of histones (H3, H4) in
the CES1 chromatin. (A, B) C57BL/6 mice were treated with glucose as described in
Fig. 3. Liver lysates were used for ChIP assay to determine acetylation of histone 3
(AcH3) (A) and histone 4 (AcH4) (B). *p<0.05, **p<0.01
22
CES1 regulates postprandial glucose levels and insulin sensitivity
Postprandial blood glucose levels are tightly controlled to avoid any unwanted
side effect of glucose. The finding that glucose induces CES1 expression and that CES1
regulates glucose metabolism [1] suggest that hepatic CES1 may regulate postprandial
blood glucose levels. To test this hypothesis, C57BL/6 mice were injected with AdshLacZ and Ad-shCes1. Five days after adenovirus injection, mice were fasted for 16 h
prior to gavage with saline or glucose (8 g/kg). Blood glucose levels were measured 1 h
after gavage. For saline treatment, Ces1-deficient mice had similar blood glucose levels
compared with the control mice (Figure 5A). For glucose treatment, however, Ces1deficient mice had significantly higher blood glucose levels compared to the control mice
(Figure 5A). In the liver, expression of Ces1 shRNA resulted in elevated levels of hepatic
triglycerides (TG) (Figure 5B) and free fatty acids (FFAs) (Figure 5C). Consistent with
the latter data, hepatic Ces1 knockdown reduced the ratio of phospho-AKT (p-AKT) to
total AKT (Figure 5D and 5E). Finally, knockdown of hepatic Ces1 reduced hepatic
Ces1 mRNA levels by>90% (Figure. 5F), and increased hepatic PEPCK and glucose 6phosphatase (G6Pase) expression in saline- but not glucose-treated mice (Figure 5G and
5H). These latter data suggest that hepatic CES1 deficiency may cause hepatic insulin
resistance and that the increase in postprandial glucose levels may not be a result of
uncontrolled hepatic glucose production. Together, the data of Figure 5 demonstrate that
hepatic CES1 plays an important role in regulating postprandial glucose levels.
23
Figure 5. CES1 regulates postprandial levels. (A–H) C57BL/6 mice were injected with
Ad-shLacZ or Ad-shCes1. After 5 days, mice were fasted for 16 h followed by gavage
with saline or glucose (8 g/kg) (n = 6 mice per group). Blood glucose levels were
measured 1 h after gavage using a glucometer (A). Mice were then sacrificed 3 hours
after gavage. Hepatic triglyceride (TG) (B) and free fatty acid (FFA) (C) levels were
analyzed. Hepatic protein levels were assessed by Western blot assays (D) and then the
ratio of p-AKT to total AKT was quantified (E). Hepatic mRNA levels
of Ces1 (F), PEPCK (G) and G6Pase (H) were determined by qRT-PCR. (I) Reciprocal
regulation between plasma glucose and hepatic CES1. Elevated plasma glucose induces
hepatic CES1, which in turn helps lower plasma glucose levels likely via increasing
peripheral insulin sensitivity. AKT, protein kinase B. *p<0.05 **p<0.01.
24
1.3.2 Hepatic Carboxylesterase 1 is Essential for Normal and Farnesoid X ReceptorControlled Lipid Homeostasis.
Over-expression of hepatic CES1 reduces hepatic triglyceride and improves glucose
homeostasis.
We initially examined whether hepatic CES1 plays a role in regulation of hepatic
lipid homeostasis. Adenovirus-mediated over-expression of hepatic Ces1 (Ad-Ces1-GFP)
had no effect on plasma TG and TC levels (data not shown), but significantly lowered
plasma glucose and hepatic TG levels with reduction of ~30% and 60%, respectively
(Figure 6A). Consistently, over-expression of hepatic Ces1 in ob/ob mice reduced plasma
glucose and hepatic TG levels (Figure 6B). H&E and Oil red O staining revealed that
over-expression of hepatic Ces1 reduced lipid accumulation in liver, which is also
evidenced by the appearance of more reddish liver (Figure 6C). The lowered glucose
level led us to determine the role of hepatic CES1 in glucose metabolism. Hepatic
expression of Ces1 improved glucose tolerance in glucose tolerance test (Figure 6D). In
addition, p-AKT/AKT ratio was markedly elevated, suggesting that hepatic Ces1
enhances insulin signaling (Figure 6E). Collectively, over-expression of hepatic Ces1
reduces hepatic lipid accumulation and improves glucose homeostasis.
25
26
Figure 6. Hepatic expression of CES1 lowers hepatic triglyceride levels and
improves glucose homeostasis. (A) C57BL/6 mice were i.v. injected with either AdGFP or Ad-Ces1 (n=7–8 mice per group). After 7 days, mice were fasted for 5 h. Plasma
glucose (left panel) and hepatic TG (right panel) levels were determined. (B–F) ob/ob
mice were i.v. injected with Ad-GFP or Ad-Ces1 (n=5 mice per group). After 7 days and
a 5-h fast, mice were euthanized. Plasma glucose (B, left panel) and hepatic TG levels (B,
right panel) were determined. Representative liver images are shown in (C, top panel)
and representative H&E staining (C, middle panel) or oil red O staining (C, bottom panel)
of the liver sections are shown in (C). Glucose tolerance test (GTT) was performed after a
16 h fast (D). Western blot assays were performed using liver lystates (E) and protein
levels quantified using Image J software (F).
27
Over-expression of hepatic CES1 does not affect fatty acid lipogenesis or very lowdensity lipoprotein (VLDL) secretion.
To determine the mechanism by which CES1 reduces lipid accumulation in the
liver, genes involved in cholesterol, fatty acids and TG synthesis, and fatty acid oxidation
(FAO) were analyzed. Figure 7A showed that mRNA levels of genes in cholesterol
synthesis (Srebp-2, HMG-CoA reductase (Hmgcr) and HMG-CoA synthase (Hmgcs))
were reduced; genes controlling fatty acid and TG synthesis (Srebp-1c, acetyl-CoA
carboxylesterase 1 (Acc1), fatty acid synthase (Fas), acyl-CoA: diacylglycerol
acyltransferase 1 (Dgat1) and Dgat2) were not significantly changed, except that mRNA
level of Acc2, a gene which represses fatty acid oxidation (FAO) was decreased; Ppar 
target genes (fatty acid translocase (Cd36), pyruvate dehydrogenase kinase 4 (Pdk4),
angiopoietin-like protein 4 (Angptl4), carnitine palmitoyltransferase 1b (Cpt1b)) were
markedly
increased.
Interestingly,
genes
involved
in
gluconeogensis
(phosphoenolpyruvate carboxykinase (Pepck) and glucose-6-phosphate (Gpase6) were
not significantly changed in wild-type mice injected with Ad-Ces1 (Figure 7A left panel),
but significantly reduced in ob/ob mice injected with Ad-Ces1 (Figure 7A right panel).
This result is in agreement with enhanced insulin signaling in ob/ob mice injected with
Ad-Ces1 (Figure 6E). Glucokinase (Gck), which facilitates glucose uptake by
phosphorylating glucose to glucose-6-phosphate, is transcriptionally regulated by insulin
[81]. Hepatic expression of Ces1 increased Gck mRNA level by >2 fold (Figure 7A right
panel).
Next, we examined de novo lipogenesis by injection of mice with 2H2O.
Consistent with the gene expression data (Figure 7A), over-expression of hepatic Ces1
28
did not significantly change hepatic de novo biosynthesis of palmitate (Figure 7B) or TG
(Figure 7C), but reduced hepatic cholesterol biosynthesis (Figure 7D). It has been shown
that Ces3 is involved in very low-density lipoprotein (VLDL) assembly in the liver and
affects plasma TG and CHOL levels [54]. Accordingly, we determined whether Ces1 had
similar effect. Interestingly, over-expression of hepatic Ces1 had no effect on the protein
levels of microsomal triglyceride transfer protein (MTP) or apolipoprotein B (ApoB)
(Figure 7E), or VLDL secretion rate (Figure 7F). This result is in line with unchanged
plasma lipid levels in hepatic expression of Ces1 mice. Thus, hepatic expression of CES1
did not affect TG biosynthesis or VLDL secretion.
29
Figure 7. Hepatic expression of CES1 selectively regulates gene expression and has
no effect on lipogenesis or VLDL secretion. (A) Hepatic mRNA levels in wild-type
(left panel) or ob/ob mice (right panel) were determined by qRT-PCR. (B–D) De novo
lipogenesis was determined in mice after injection of 2H2O (n=5 mice per group). The
levels of newly synthesized [2H]palmitate (B), [2H]TG (C) or [2H]cholesterol (D) in the
liver were quantified. (E) Hepatic protein level was assessed by Western blot assays. (F)
VLDL secretion rate was determined (n=6 mice per group).
30
Over-expression of hepatic CES1 increases hepatic TG hydrolysis and stimulates
fatty acid oxidation.
The increased PPAR target gene expressions indicated that Ces1 overexpression may induce PPAR activity. To test this hypothesis, we co-transfected a
Ces1-expressing plasmid and a luciferase-reporter plasmid containing 3 copies of PPAR
responsive element (PPRE). PPAR activity was markedly induced by over-expression
of CES1 (Figure 8A top penal). Free fatty acids (FFAs) are endogenous ligands for
PPAR [82]. The increased PPAR activity is indicative of increased FFA level as a
result of TG hydrolysis (Figure 8A bottom penal). This result suggests that CES1 has TG
hydrolysis activity. In addition, over-expression of Ces1 significantly increased the
release of [3H]FFAs from [3H]triolein in both COS-7 cells (Figure 8B) and liver (Figure
8C), indicating that CES1 is capable of hydrolyzing TG.
Further study showed that hepatic total fatty acids and a number of long-chain
fatty acids, such as C14:0, C16:0, C16:1, and C18:2, were significantly reduced by
hepatic expression of Ces1 (Figure 8D). Hepatic FFA level was also reduced by ~20%
(Figure 8E). The repression of Acc2 and induction of Cd36, Pdk4, Angptl4, and Cpt1b, all
of which are involved in FAO, suggest that CES1 may regulate FAO. Indeed, overexpression of Ces1 increased FAO (Figure 8F). Thus, hepatic expression of Ces1 induces
triglyceride hydrolysis and increases FAO, leading to reduced hepatic triglyceride level.
31
Figure 8. Hepatic expression of CES1 increases triglyceride hydrolase activity and
activates PPARα (A) HepG2 cells were transfected with a control plasmid or a Ces1expressing plasmid together with a 3xPPRE-Luc reporter plasmid. Luciferase activity
was determined (top panel). In the bottom panel, the diagram shows that CES1
hydrolyzes TG and releases FFAs, which bind to PPARα/RXR complex and then induce
PPARα activity. (B, C) TGH activity was assessed using lysates from COS-7 cells (B) or
the liver (C). (D, E) Hepatic fatty acid profile was determined by GC-mass spectrometry
32
(D) and hepatic FFA levels were quantified (n=8 mice per group) (E). (F) FAO was
performed in the liver cell line AML12 cells that were infected with Ad-GFP or AdCES1 for 48 h, or treated with either vehicle or carnitine (1 mM) (n=3–5 per group).
Carnitine treatment serves as a positive control. Veh, vehicle.
33
Knockdown of hepatic CES1 causes hepatic steatosis and elevated plasma
cholesterol level.
To determine whether hepatic CES1 is required for maintaining normal lipid and
glucose homeostasis, we injected C57BL/6J mice with Ad-shLacZ (control) and AdshCes1. Hepatic Ces1 mRNA level was substantially reduced (Figure 9A). Its protein
levels were also significantly decreased (Figure 9B). Despite the plasma TG level was not
significantly changed, plasma TC level was increased by ~2 folds in mice injected with
shCes1 (Figure 9C). In agreement with increased plasma TC level, hepatic Ces1
deficiency drastically increased VLDL-cholesterol and LDL-cholesterol levels and
slightly reduced HDL-cholesterol level (Figure 9D). The liver of mice injected with
shCes1 showed paler color (Figure 9E, top panel). Oil red O staining confirmed that
knockdown of hepatic Ces1 led to increased accumulation of lipid droplets (Figure 9E,
bottom panel). The latter finding suggests that hepatic Ces1 deficiency results in the
development of fatty liver. Indeed, Hepatic TG level was increased more than 2 folds,
whereas hepatic TC level was unchanged (Figure 9F). Collectively, knockdown of Ces1
results in fatty liver.
34
Figure 9. Loss of hepatic CES1 causes fatty liver and increased plasma cholesterol
level (A–F) C57BL/6 mice were i.v. injected with either Ad-shLacZ or Ad-shCes1 (n=8
mice per group). After a 6-h fast, mice were euthanized. Hepatic mRNA (A) and protein
(B) levels were determined. Plasma TG and total cholesterol (TC) levels were quantified
(C). Plasma cholesterol lipoprotein profile was determined by FPLC (D). Representative
liver images (top panel) and oil red O staining of liver sections (lower panel) are shown
(E). Hepatic TG and TC levels were quantified (F).
35
Hepatic CES1 deficiency results in increased lipogenesis.
Next, we determined whether the increased hepatic TG is caused by increased
lipogenesis. Analysis of hepatic gene expression indicated that many SREBP- or
cholesterol/LXR(liver X receptor)-regulated genes, including Hmgcs, Acc-1, Dgat1,
Dgat2, ATP citrate lyase (Acl), Abca1, and Cd36, were significantly induced (Figure
10A). In agreement with the induction of a number of lipogenic genes, hepatic de novo
lipogenesis of palmitate (Figure 10B), TG (Figure 10C), and cholesterol (Figure 10D),
were induced by 2.1, 2.9 and 2.1 fold, respectively. Consistent with increased mRNA
level of Srebp1c and Srebp2, the protein levels of hepatic mature/nuclear SREBPs
(nSREBPs) were increased in mice injected with shCes1 (Figure 10E), suggesting that
loss of Ces1 results in elevated SREBPs translocation from cytosol to nucleus. SREBP
processing is known to be regulated by the change in intracellular sterol levels and is
sensitive to cholesterol ester (CE) to free cholesterol (FC) ratio; when cellular sterol
levels are low or CE/FC ratio are high, mature SREBPs are increased by inducing
SREBP processing [83]. Given that CES1 has CEH property, we hypothesized that loss
of Ces1 results in a reduction of cholesterol ester hydrolysis and leads to a subsequent
reduction of free cholesterol level, which in turn induces SREBP processing. As
predicted, hepatic FC level was reduced in Ces1-deficient mice (data not shown). The
CE/FC ratio was increased. Thus, loss of hepatic Ces1 increased lipogenesis through
increasing SREBPs processing, which may be accounted, at least in part, by reduced FC
levels.
36
Figure 10. Loss of hepatic CES1 induces de novo lipogenesis (A) Hepatic mRNA
levels were determined by qRT-PCR (n=8 per group). (B–D) De novo lipogenesis was
determined in mice after injection of 2H2O (n=5 per group). The levels of newly
synthesized [2H]palmitate (B), [2H]TG (C), or [2H]cholesterol (D) in the liver were
quantified. (E) Hepatic protein levels were determined by Western blot assays (E, left
panel) and then quantified by using Image J software (E, right panel). nBP-1, nuclear
37
form SREBP-1. nBP-2, nuclear form SREBP-2. (F) Hepatic ratio of CE to FC was
determined (n=8 mice per group). The CE/FC ratio in shLacZ-treated mice was set at 1.
38
Hepatic CES1 is regulated by FXR.
Activation of FXR is shown to be beneficial in regulating lipid and carbohydrate
metabolism. We found that activation of FXR by GW4064, a FXR agonist, reduced
hepatic TG level but did not change TC level (Figure 11A). Activation of FXR induced
hepatic mRNA levels of Shp, an FXR target, and Ces1, but mRNA levels of Ces2 and
Ces3 were unchanged (Figure 11B). Figure 11 C showed that GW4064 treatment
increased protein levels of hepatic CES1, but did not change ApoB and MTP protein
levels, suggesting that activation of FXR does not change VLDL secretion. In addition,
mice were treated with obeticholic acid (OCA, INT-747), a potent and selective FXR
agonist, or cholic acid, the endogenous FXR ligand, hepatic Ces1 mRNA was also
induced (Figure 11D and E). On the other hand, hepatic Ces1 mRNA level was reduced
in Fxr mice (Figure 11F). Therefore, hepatic CES1 expression is regulated by FXR.
39
Figure 11. Hepatic CES1 is regulated by FXR (A–C) C57BL/6 mice were gavaged
with vehicle (0.5% CMC (carboxymethyl cellulose)) or GW4064 (30 mg/kg, twice a day)
for 7 days (n=8 mice per group). Hepatic TG and TC levels were determined (A). Hepatic
mRNA levels were quantified by qRT-PCR (B) and hepatic protein levels were
determined by Western blot assays (C). (D) C57BL/6 mice were gavaged with either
40
0.5% CMC (vehicle) or INT-747 (OCA, 30 mg/kg/d) for 7 days (n=5 mice per group).
Hepatic mRNA levels were determined. Cyp7a1 and Cyp8b1 serve as positive controls.
(E) C57BL/6 mice were fed a chow diet or 0.5% cholic acid (CA) for 7 days. Hepatic
mRNA levels were quantified. (F) Hepatic mRNA levels were quantified by qRT-PCR in
wild-type or Fxr−/− mice (n=8 mice per group).
41
CES1 is a direct FXR target.
Next, we determined how FXR regulated CES1 expression. A luciferase assay
was performed using a serial 5’-deletion construct along with Fxr expressing plasmid.
Figure 12A showed that activation of FXR increased Ces1 promoter activity. Luciferase
and mutagenesis assays revealed that the FXR response element was located in Ces1
promoter region between 300bp and 150bp upstream of transcriptional start site (Figure
12A and B). Electrophoretic mobility shift assays showed that the FXR/RXR complex
bound to this FXRE in vitro and this binding was competed away by cold wild-type
oligonucleotides but not by mutant oligonucleotides (Figure 12C). Lastly, chromatin
immunoprecipitation assay showed that FXR bound to the promoter of Ces1 in the liver.
Akr1b7, a known FXR target gene [84], serves as a positive control (Figure 12 D).
Overall, Figure 11 and Figure 12 suggested that Ces1 is regulated by FXR and its direct
target gene.
42
Figure 12. CES1 is a direct FXR target gene (A, B) Transient transfection assays were
performed using promoter-luciferase constructs containing a serial of 5'-deletions (A) or
mutations (B). (C) EMSA assays were performed using in vitro transcribed/translated
proteins. Wild-type (WT) and mutant (MUT) oligos were used in the competition assays.
(D) Chromatin immunoprecipitation (ChIP) assays were performed using liver lysates
(n=3 per group). Akr1b7 serves as a positive control.
43
Hepatic CES1 is essential for activated FXR to improve lipid homeostasis.
Activation of FXR induced the expression of Ces1. This finding prompted us to
study whether hepatic Ces1 mediated FXR-controlled lipid homeostasis. C57BL/6 mice
were injected i.v. with either Ad-shLacZ or Ad-shCes1, followed by treatment with either
vehicle or OCA for 7 days. As expected, knockdown of hepatic Ces1 increased plasma
cholesterol level by ~4 fold (Figure 13A). Interestingly, OCA treatment reduced plasma
cholesterol levels in the control mice but not in Ces1-deficient mice (Figure 13A).
Similarly, OCA treatment also reduced plasma and hepatic TG levels in the control mice
but not in the Ces1-deficient mice (Figure 13B and C). This result suggested that hepatic
Ces1 is important in FXR-controlled lipid homeostasis.
44
Figure 13. Essential roles of hepatic CES1 in FXR-regulated lipid homeostasis
(A–C) C57BL/6 mice were i.v. injected with Ad-shLacZ or Ad-shCes1. The next day,
these mice were gavaged with either vehicle (0.5% CMC) or INT-747 (OCA, 30 mg/kg/d)
for 7 days (n=8–10 mice per group). After a 5-h fast, mice were euthanized. Plasma total
cholesterol (TC) (A) and TG (B) levels as well as hepatic TG levels (C) were determined.
45
CES1 deficiency alters the lipid profile in mice treated with FXR agonist
Activation of FXR tended to reduce plasma glucose level in control mice
(shLacZ). Similarly, plasma glucose level was reduced in OCA treated Ces1-deficient
mice (Figure 14A). Analysis of plasma by FPLC showed that loss of hepatic Ces1
markedly increased plasma non-HDL-C levels (Figure 14B) and also slightly increased
LDL triglyceride levels (Figure 14C), and these changes were exacerbated following
OCA treatment (Figure 14B and C). Thus, hepatic CES1 is critical for an FXR agonist to
lower plasma TG and cholesterol levels. In addition, OCA did not significantly change
hepatic cholesterol level, whereas it increased cholesterol level in Ces1-deficient mice
(Figure 14D).
46
Figure S7
Figure S7
A
A
B
B
C
C
D
D
Figure 14. Effects of the FXR agonist OCA (INT-747) on lipid and glucose
homeostasis in C57BL/6 mice. C57BL/6 mice were injected i.v. with either Ad-shLacZ
or Ad-shCes1. On the next day, these mice were gavaged with either vehicle or OCA
(n=7-9 mice per group). After 7 days, mice were fasted for 5 h. Plasma glucose levels (A)
and hepatic cholesterol levels (D) were measured. Plasma cholesterol (B) and triglyceride
(C) lipoprotein profiles were determined by FPLC.
47
Ces1 is important in FXR-controlled lipid homeostasis in ob/ob mice.
To further define the role of Ces1 in FXR-controlled lipid homeostasis, ob/ob
mice were injected with Ad-shLacZ or Ad-shCes1, followed by OCA treatment for 7
days. Similar to what we observed in C57BL/6 mice, we found that OCA treatment
lowered plasma cholesterol and TG levels as well as hepatic TG levels in a CES1dependent manner (Figure 15A, B and D), but had no effect on hepatic cholesterol levels
(Figure 15C). Analysis of hepatic gene expression showed that OCA treatment induced
hepatic mRNA levels of Ces1 in the control mice and Shp in both the control and Ces1deficient mice (Figure 15F). Collectively, hepatic CES1 is indispensable for activated
FXR to regulate both plasma lipid and hepatic TG levels.
48
Figure S8
A
B
C
D
E
F
Figure 15. Effects of the FXR agonist OCA on lipid homeostasis in ob/ob mice.
Ob/ob mice were injected i.v. with either Ad-shLacZ or Ad-shCes1. On the next day,
these mice were gavaged with either vehicle or OCA (n=6 per group). After 7 days, mice
were fasted for 5 h. Plasma total cholesterol (TC) (A), plasma triglycerides (TG) (B),
hepatic TC (C) and hepatic TG (D) levels were determined. Hepatic mRNA levels of
Ces1 (E) and Shp (F) were quantified by qRT-PCR. * p<0.05, ** p<0.01. NS, nonspecific.
49
1.4 DISCUSSION
CES1 is highly expressed in the liver. The current studies show that CES1 has
TGH activity; it reduced hepatic TG level via increasing TG hydrolysis and subsequent
fatty acid oxidation (FAO) [52,85]. Lipid homeostasis has a profound impact on insulin
sensitivity and glucose metabolism. Plasma free fatty acid (FFA) levels correlate
negatively to the degree of insulin sensitivity [86,87]. Given the regulatory role of CES1
in TG hydrolysis and FAO, it is not surprising to see that increased hepatic CES1
expression lowers plasma glucose levels and improve insulin sensitivity. In the present
study, we investigate the regulation of hepatic CES1 by glucose and the physiological
role of such regulation. Our data reveal a novel role for hepatic CES1 in postprandial
glucose control.
CES1 regulates the postprandial glucose levels.
Poor control of postprandial glucose is a significant contributor to type 2 diabetes
mellitus. Persistent, moderate increase in postprandial glucose levels is a significant risk
factor for macrovascular complications, and is more indicative of atherosclerosis than
fasting glucose. In light of the risks of postprandial hyperglycemia for vascular events,
tight control of postprandial glucose levels is important for long-term indices of diabetes
control. After the start of a meal, blood glucose levels are increased. The increased blood
glucose levels induce hepatic CES1 expression, which in turn helps lower blood glucose
levels likely by increasing hepatic insulin sensitivity. This conclusion is supported by the
50
findings that increased hepatic CES1 expression lowers plasma glucose levels whereas
loss of hepatic CES1 results in increased postprandial blood glucose levels.
Although our data show that hepatic CES1 is required for regulating postprandial
glucose levels, the underlying mechanism remains to be fully determined. Global
Ces1mice have elevated plasma levels of triglyceride, free cholesterol, FFAs and
insulin [60]. These mice also present insulin resistance, which results from reduced
insulin sensitivity in both skeletal muscle and white adipose tissue [60]. However, CES1
is not expressed in skeletal muscle and its expression level in white adipose tissue is low
(data not shown). Thus, insulin resistance observed in global Ces1mice likely results
from a deficiency in hepatic Ces1. Consistent with the speculation, our data show that
hepatic Ces1 deficiency results in impaired postprandial glucose clearance. Although
hepatic CES1 deficiency may cause hepatic insulin resistance, skeletal muscle and white
adipose tissues are the major organs responsible for plasma glucose clearance. Therefore,
hepatic Ces1 deficiency may affect peripheral insulin sensitivity. To precisely understand
how hepatic CES1 deficiency regulates glucose homeostasis or insulin sensitivity,
heperinsulinemic-euglycemic clamp studies will be needed to help characterize the
underlying mechanism.
51
CES1 is induced by glucose.
We have previously shown that farnesoid X receptor regulates CES1 expression
[1]. The present study showed that hepatic CES1 is also regulated under physiological
and pathological conditions. Under these latter conditions, hepatic CES1 expression is
altered likely due to the change in plasma glucose levels. Indeed, our data show that
glucose induces CES1 expression both in vitro and in vivo. Consistent with our finding,
very recent data by Xiong et al. also show that glucose induces the expression of CESs in
mouse primary hepatocytes [88].
Several lines of evidence suggest that glucose regulates gene expression through
epigenetic modifications [78] and that nuclear ACL is important for glucose-mediated
histone acetylation [89]. In this study, we show that glucose induces CES1 expression by
epigenetic modifications of the acetylation status (H3 and H4) of the CES1 chromatin in
an ACL-dependent manner. The histone tails interact with a region about 150bp upstream
of transcription start site of CES1. Consistent with this finding, the data from the
luciferase-promoter assays show that this region is required for glucose to induce CES1
promoter activity. Thus, glucose induces CES1 expression via acetylation of H3 and H4
in the CES1 chromatin.
52
Recent reports show that global deletion of CES1 promotes hepatic steatosis,
obesity and hyperlipidemia [60]; intestinal CES1 regulates chylomicron secretion [59].
The TG and CE hydrolase activity of CES1 have also been reported, which highlight its
important role in regulating hepatic lipid homeostasis. In the present studies, we
demonstrate that hepatic CES1 controls lipid and carbohydrate metabolism, reinforcing
its values as a therapeutic target in treating NAFLD. Additionally, we find that FXR
regulates hepatic CES1 expression, and the induction of hepatic CES1 is indispensable
for activated FXR to improve lipid homeostasis.
CES1 plays a key role in regulating hepatic triglyceride and cholesterol levels.
We clearly demonstrated that hepatic CES1 has TG hydrolase activity, suggesting
its role in maintaining hepatic TG homeostasis. Hepatic TG mobilization is little
understood. Previous data suggest that ATGL may play an important role in mobilizing
hepatic TG. However, ATGL expression is low in hepatocytes [90]. CES1, expressed
abundantly in liver, may be one of major enzymes which control hepatic TG levels.
The key finding of this study is that over-expression of hepatic CES1 increases
TG hydrolysis, reduces hepatic TG levels and increases the release of FFA, which in turn
activates PPAR and promotes FAO, leading to reduced FFA levels. The reduced FFA
levels, in turn, lead to increased hepatic insulin sensitivity and reduced plasma glucose
levels, suggesting that hepatic expression of Ces1 also improves carbohydrate
metabolism. On the other hand, loss of hepatic CES1 significantly increases lipogenesis
53
and SREBPs processing. These results suggest that CES1 is required for maintaining
normal lipid homeostasis.
CES1 is important in FXR-controlled lipid homeostasis.
FXR is emerging as a significant target for treatment of fatty liver disease [91].
However, the mechanism underlying FXR-mediated alleviation of hepatic steatosis
remains undetermined. Our study provides compelling evidence demonstrating that CES1
is a direct downstream target of FXR and is necessary for activated FXR to lower plasma
TG and cholesterol levels. Using ob/ob mice, we demonstrate that loss of hepatic CES1
abolishes the effect of activation of FXR on reducing hepatic steatosis, suggesting that
hepatic expression of CES1 is essential for using FXR agonist in treating metabolic
disease. Therefore, hepatic CES1 plays an essential role in FXR-controlled lipid
homeostasis.
In summary, in chapter 1, we proposed a glucose-CES1-glucose (Figure 5I)
reciprocal cascade, in which glucose induces hepatic CES1 expression, which in turn
increases insulin sensitivity and lowers blood glucose levels. This cascade plays an
important role in regulating postprandial glucose levels. Since high levels of postprandial
blood glucose contribute to macrovascular complications, CES1 may be targeted for
prevention of vascular diseases associated with hyperglycemia. In addition to its role in
regulation of carbohydrate metabolism, over-expression of hepatic CES1 is shown to
have beneficial effects on lipid metabolism. In contrast, loss of hepatic CES1 causes
fatty liver and proatherogenic lipid profile. Extensive studies demonstrate that CES1 is a
54
direct target of FXR and is critical for FXR-mediated improvement of lipid homeostasis.
Together, our data suggest that hepatic CES1 is essential in normal and FXR-controlled
lipid and carbohydrate homeostasis, and also suggest that hepatic CES1 may represent a
therapeutic target for treatment of NAFLD.
55
CHAPTER 2. THE ROLE OF CARBOXYLESTERASE 1 IN ALCOHOLIC
FATTY LIVER DISEASE
2.1 INTRODUCTION
Alcoholic liver disease (ALD) is a major cause of chronic liver disease, ranging
from simple steatosis, hepatitis, to more severe forms including fibrosis, cirrhosis and
hepatocellular carcinoma (HCC). Liver cirrhosis is the 12th leading cause of death in the
US, with a total of 36,427 deaths in 2013, 49.8% of which are related to alcohol [92].
Abstinence is an effective strategy for the treatment of ALD, but sustainable lifestyle
changes are difficult for many patients to achieve. Pharmacological treatments, such as
the use of alcohol dehydrogenase inhibitors and anti-craving drugs, do not achieve
satisfactory effect in patients with ALD [92]
Alcohol-induced liver damage is characterized by a reduction of NAD/NADH and
subsequent inhibition of fatty acid oxidation, up-regulation of lipogenesis, and an
increase in inflammation and oxidative stress which promote hepatocyte necrosis and
apoptosis [93,94,95]. Alcohol also increases hepatic free fatty acid (FFA) uptake and
decreases lipolysis, resulting in impaired very low density lipoprotein (VLDL) secretion
[96]. Alcohol-induced liver injury is associated with enhanced lipid peroxidation, protein
carbonyl formation, formation of free radicals, and decrease in glutathione [97].
56
Enzymes,
which
govern
the
metabolism
of
alcohol,
include
alcohol
dehydrogenase, acetaldehyde dehydrogenase and cytochrome P450 system, and abnormal
regulation of these enzymes are associated with the development of ALD. Alcoholinduced hepatic steatosis is prominent in ALD. Alcohol disrupts fatty acid oxidation by
down-regulating peroxisome proliferator-activated receptor (PPAR)-α, and damaging
mitochondrial functions [98,99,100]. Chronic alcohol administration suppresses sirtuin 1
(SIRT1)-AMP-activated kinase (AMPK) axis [101,102,103,104], thus represses rates of
fatty acid oxidation and enhances lipogenesis through modulating PPAR-γ coactivator-α
(PGC-1α)/PPARα and sterol regulatory element-binding protein 1 (SREBP-1)
[101,105,106]. Alcohol increases gut epithelial membrane permeability, thus increases
translocation of lipopolysaccharides (LPS), which in turn elicits Toll-like receptor (TLR)mediated signaling cascade and subsequent increase in productions of cytokines (TNF-α,
IL-1β, and IL-6), resulting in inflammation [107,108]. Furthermore, alcohol metabolites
(acetaldehyde and acetate) [109], free fatty acids [110], and reactive oxygen species
(ROS) [111] contribute to chronic alcohol feeding-induced inflammation. Lipid
homeostasis plays a central role in the pathogenesis of ALD. Many data have shown that
CES1 is essential in regulating lipid homeostasis in liver [1,52,60], intestine [59], and
macrophages [51]. CES1 is also a phase I drug metabolizing enzyme, which can
metabolize xenobiotics. It is plausible to speculate that CES1 may be involved in alcoholinduced hepatotoxicity. In addition, Over-expression of human CES1 increases the
production of fatty acyl ethyl esters (FAEEs) via transesterification of short-chain and
57
long-chain fatty acids with ethanol [112]. FAEEs are toxic byproduct of alcohol abuse.
So far, the role of CES1 in the development of ALD is unknown.
Hepatocyte nuclear factor 4 (HNF4) is a nuclear hormone receptor that is
constitutively active to regulate lipid, glucose and bile acid metabolism. Loss of hepatic
HNF4 causes fatty liver by reducing very low-density lipoprotein (VLDL) secretion.
HNF4 expression is markedly reduced in diabetes, obesity, non-alcoholic fatty liver
disease (NAFLD) and high fat diet (HFD) feeding, likely as a result of increased free
fatty acids, cholesterol and miR-34a expression [113]. A previous study shows that
farnesoid X receptor (FXR) regulates CES1 expression [1]. It is unclear whether HNF4
also regulates CES1 expression.
The present studies determined the role of CES1 in the development of ALD. We
found that alcohol reduces the expressions of CES1 and HNF4 in patients with
alcoholic steatohepatitis and in mice treated with alcohol. HNF4DR1 response element
is identified in Ces1 promoter. Adenovirus-mediated knockdown of hepatic CES1 (AdshCes1) deteriorates alcohol-induced hepatic steatosis and liver inflammation. Compared
with wild-type mice, Ces1 mice does not show aggravated hepatic steatosis in response
to alcohol treatment, whereas Ces1 mice display aggravated liver inflammation upon
ethanol consumption and methionine-choline deficient (MCD) diet feeding. Hepatic
acetaldehyde levels, mitochondrial ROS and hepatic MDA levels are markedly increased
in ethanol-fed Ces1 mice, compared with ethanol-fed wild-type mice.
58
2.2 METHODS
Mice, diets and human ALD samples.
Liver-specific Hnf4 mice were generated by crossing Hnf4fl/fl mice with
albumin-Cre mice (all from the Jackson Laboratory. Bar Harbor, ME). Ces1 mice were
generated by replacing exon 1 with an ACNLacZ cassette as described (30). Detailed
description of the Ces1 mice will be reported elsewhere. Ces1 mice were
backcrossed with C57BL/6 mice for at least 5 generations prior to experimentation. MCD
diets were purchased from Harlan Laboratories (Cat # TD.90262; Madison, WI). LieberDeCarli diets were purchased from Bio-Serv (Flemington, NJ). Human ALD liver
samples were obtained from the Liver Tissue Cell Distribution System at University of
Minnesota. All the animal experiments were approved by the Institutional Animal Care
and Use Committee at Northeast Ohio Medical University (NEOMED) and the use of
human tissue samples were approved by Institutional Review Board at NEOMED.
Chronic-binge alcohol feeding.
C57BL/6J mice (12 weeks old) were administered control Lieber-DeCarli diet
(cat#F1259SP, BioServ) for 5 days. On the 6th day, mice were fed an ethanol LieberDeCarli diet (cat#F1258SP, BioServ) containing 5% (vol/vol) ethanol or pair-fed a
control Lieber-DeCarli diet for 10 days [114]. On the 16th day, mice were gavaged with a
single dose of ethanol (5g/kg body weight) or isocaloric maltose dextrin. For some
studies, on the 6th day, mice were also injected i.v. with either Ad-shLacZ or Ad-shCes1.
59
RNA isolation and quantitative real-time PCR.
Total RNA was isolated using TRIzol Reagent (Life Technologies, NY). mRNA
levels were determined by quantitative reverse-transcription polymerase chain reaction
(qRT-PCR) on a 7500 real-time PCR machine from Applied Biosystems (Foster City,
CA). Relative mRNA levels were calculated using the comparative cycle threshold (Ct)
method and were normalized to the values of 36B4 mRNA levels.
Western blotting.
Tissues were homogenized in ice-cold modified RIPA buffer and protein
concentrations were determined using a Pierce BCA Protein Assay Kit (Thermo
Scientific, IL). Antibodies against mouse CES1, HNF4 and -actin were purchased
from Abcam (ab45957, Cambridge, MA), Santa Cruz Biotechnology (sc6556, Dallas, TX)
and Novus Biologicals (NB600-501, Littleton, CO), respectively.
Transient Transfection and Mutagenesis assays.
Ces1 promoter regions (-1.9 kb, -0.98 kb, -0.3 kb, -0.25 kb and -0.21 kb) were
cloned to pGL3-basic plasmid (Promega, Madison, WI).Transient transfection assay was
performed as described [115]. pGL3-Ces1 luciferase reporter constructs were transfected
into HepG2 cells using lipofetamine 3000 (Invitrogen, CA) along with either pCDNA3 or
pCDNA3-HNF4 plasmid. After 36 hrs, luciferase activities were determined and
normalized to -galactosidase activity. The mutant pGL3 promoter-luciferase construct
(pGL3-Ces1-mut(-1.9k))was generated using a QuickChange Site-directed Mutagenesis
60
kit from Agilent (Santa Clara, CA). Subsequently, the lucifrease activity of the mutant
was tested.
Chromatin immunoprecipitation (ChIP) assay.
ChIP assay was performed following the manufacturer’s instructions (cat#17-295,
Millipore, MA). Antibody against HNF4 was purchased from Santa Cruz
Biotechnology
(sc6556,
Dallas,
TX).
CAGAACACTGAGGTTTGAATTCC
The
primer
(forward)
sequences
were
and
TCACACCGACCTAGAGTTTAAAC (reverse), which amplified a fragment between 250bp and -300bp in the Ces1 gene promoter.
Electrophoretic Mobility Shift Assay (EMSA).
Oligonucleotides were labeled by biotin on the 3’ end following the
manufacturer’s instruction (cat#89818, Thermo Fisher Scientific). Then, EMSA was
performed using a kit purchased from Thermo Fisher Scientific and following the
manufacturer’s instructions (cat# 20148, Thermo Fisher Scientific). The HNF4 protein
was made using TnT T7 Quick Coupled Tanscription/Translation Reactions kit (cat#
L1170, Promega) and pCDNA3-hnf4 plasmid. The wild-type oligonucleotide sequences
used
for
EMSA
were
5’-
CCCTGTCTGAAGGCCTGCTGTGCTACTCTCTGCCTTTGGGAGGCCGACAG-3’
(top strand) and 5’-CTGTCGGCCTCCCAAAGGCAGAGAGTAGCACAGCAGGCCTT
CAGACAGGG-3’ (bottom strand). The mutant oligonucleotide sequences were 5’CCCTGTCTGAAGGCCTGCTGTGTTACTTTTTGTTTTTGGGAGGCCGACAG-3’
61
(top
strand)
and
5’-
CTGTCGGCCTCCCAAAAACAAAAAGTAACACAGCAGGCCTCCAGACAGGG-3’
(bottom strand).
Adenovirus.
Ad-shCes1 was designed using BLOCK-iT RNAi Designer (Invitrogen, CA),
annealed, and ligated to pEnter/U6 vector (Invitrogen, CA). The oligonucleotide
sequences
were
’GCTGATTCCAGCAGTATTGACGAATCAATAGCTGCTGGAATCAGC-3’
5(top
strand)
and
5’-GCTGATTCCAGCAGCTATTGATTCGTCAATAGCTGCTGGAATAGC-3’
(bottom strand). All the adenoviruses were grown in 293A cells and purified by Cesium
chloride gradient centrifugation. About 2x109 plaque formation units (pfu) of adenovirus
was transfused into each mouse intravenously.
Primary hepatocyte isolation.
Mouse primary hepatocytes were isolated as described [65,66]. Mice were
anaesthetized by intraperitoneal injection of 50 mg/kg pentobarbital. The portal vein was
cannulated with a 23-gauge plastic cannula. Mouse livers were perfused with Hank’s
Balanced Salt Solution (HBSS, cat#14170-112, Thermo Fisher Scientific) containing
0.19g/L EDTA. Simultaneously, the inferior vena cava was cut open. Subsequently,
livers were perfused with HBSS, calcium, magnesium buffer (cat#14025092, Thermo
Fisher Scientific) with 0.8mg/mL Collagenase from Clostridium histolyticum type IV
62
(Sigma, St. Louis, MO). Primary hepatocytes were released and collected in a 50 mL
centrifuge tube. After centrifugation at 50g for 5 minutes and washing with DMEM, cells
were cultured in 6-well plate pre-coated with 0.1% gelatin in a 2 mL of DMEM+10%
FBS.
Measurement of hepatic acetaldehyde level using High Performance Liquid
Chromotography (HPLC).
The hepatic acetaldehyde level was determined using derivatization with DNPH,
followed by HPLC separation described previously [116]. 100mg liver tissue was
homogenized in 3M perchloric acid. The pH of the solution was adjusted immediately to
4.0 using 2 volumes 3M sodium acetate buffer, pH9.0. After centrifugation (12000 rpm at
4 C for 20 minutes), the supernatant was transferred into an ice-cold tube, followed by
addition of 80-fold molar excess 2,4-dinitrophenylhydrazine (DNPH, Cat# 119266,
Sigma-Aldrich) in 6N HCl. The mixture was then placed on a shaker for 1 hour at room
temperature. Derivatization was stopped with 3 volumes of 3M sodium acetate buffer, pH
9.0. Two volumes of acetonitrile were added to extract AcH-DNP. After centrifugation
(10000g for 5 minutes at 4 C), the organic phase was condensed to 50 L.
The ultra high performance liquid chromatograph (UHPLC) machine was
purchased from Shimadzu Corp. (Columbia, MD). A Restek C18 HPLC column (25cm x
4.6mm i.d.,5m) coupled with an Ultra C18 guard column (10mm x 4mm i.d.) were
purchased from Fisher Scientific (Pittsburgh, PA). The elution program was described
previously [116]. An AcH-DNP standard (Sigma-Aldrich, St.Louis. MO) was used to
63
create a standard curve. The values of the area under the curve of the AcH-DNP peaks
were determined to calculate the concentration of acetaldehyde in each sample.
Malondialdehyde (MDA) assay.
MDA assay was performed as described [115]. Buffer I was made by dissolving
2-Thiobarbituric acid in 10% perchloric acid to a final concentration 0.67%. Assay buffer
was made by adding 20% trichloroacetic acid to buffer I (2:3,v/v), mixed well. 1,1,3,3,tetraethoxypropane was used as a standard. 30mg liver tissue was homogenized with
500L saline. After centrifugation, 100L supernatant was collected and added to 1mL
assay buffer. The reaction mixture was incubated at 95 °C for 30 minutes, cooled down
and centrifuged at 3000rpm for 10 minutes. The absorbance was measured at OD532nm.
Mitochondrial H2O2 assay.
Approximately 50 mg of liver tissue was homogenized in 1 mL of a buffer
containing 25mM Hepes pH7.4, 1mM EDTA, 0.25M sucrose, 2mM MgCl2, 1M
butylated hydroxytoluene (BHT), 1:200 dilution of Sigma P8340 and 1M
diethylenetriaminepentaacetic acid. The homogenate was centrifuged at 500g for 5
minutes to pellet nuclei and debri. Then, the supernatant was centrifuged at 10,000g for
10 minutes at 4°C to obtain Mitochondria. The pellets were resuspended with 150 L
homogenization buffer. About 20L solution was saved for measuring protein
concentration.
Mitochondrial H2O2 was detected using Amplex Red assay as described [117].
The working solution was prepared using 100L of 10mM Amplex Red reagent (Thermo
64
Fisher Scientific), 2L of 1000U/mL horseradish peroxidase (HRP) and 10mL of 50mM
potassium phosphate pH7.7 with 0.5mM diethylenetriaminepentaacetic acid. 50 L of
sample or standard were pipetted into 96 plates, followed by addition of 50 L of the
Amplex Red reagent/HPR working solution. The reaction mixture was incubated at room
temperature for 30 minutes, protected from light. The fluorescence was then measured
with excitation in the range of 530-560 nm and emission at 590nm. Mitochondrial H2O2
levels were normalized to the concentration of protein.
Lipid Analysis.
Plasma triglyceride and cholesterol were measured using Infinity reagent from
Thermo Scientific (Waltham, MA). Plasma and hepatic free fatty acid was measured
using kit from Wako Diagnostics (Richmond, VA). To measure lipids in liver,
approximately 100mg liver tissue was homogenized in methanol and extracted in
chloroform/methanol (2:1 v/v). Hepatic triglyceride and cholesterol levels were then
quantified using Infinity reagents from Thermo Scientific (Waltham, MA).
Plasma Alanine Aminotransferase (ALT) and Aspartate Aminotransferase (AST)
analysis.
Plasma ALT and AST levels were determined using Infinity reagent (Middletown,
VA) following the manufacture’s instruction.
65
Statistical Analysis.
The data were analyzed using unpaired Student t test and ANOVA (GraphPad
Prisim, CA). All values were expressed as meanSEM. Differences were considered
statistically significant at P<0.05.
66
2.3 RESULTS
Alcohol reduces CES1 and HNF4 levels in patients with alcoholic steatohepatitis
and in mice treated with alcohol.
To investigate whether CES1 is associated with the development of ALD, we
investigated the expression of CES1 in patients with alcoholic hepatitis. Hepatic CES1
mRNA level was reduced by 75% (Figure 16A) and protein level decreased by ~ 85%
(Figure 16B). Interestingly, the nuclear receptor HNF4 was also markedly repressed
by >84% in both mRNA (Figure 16A) and protein (Figure16B) levels.
In mice chronically fed a Liber-DeCarli ethanol diet for 10 days, hepatic Ces1 and
Hnf4 mRNA levels were decreased by ~ 50% (Figure 16C). In addition, ethanol
treatment significantly repressed Ces1 and Hnf4 mRNA levels in primary hepatocytes
(Figure 16D). Thus, the data of Figure1 indicate that ethanol inhibits hepatic CES1 and
HNF4 expression in both mice and humans.
67
Figure 16. CES1 and HNF4 expressions are reduced in patients with alcoholic
steatohepatitis and in mice. Liver were collected from normal individuals and patients
with ALD, mRNA (A) and protein (B) levels of CES1 and HNF4 were measured in
patients. C57BL6 mice were fed an ethanol Lieber-DeCarli diet containing 5% ethanol
for 10 days followed by a single dose of ethanol administration (5g/kg), mRNA levels
were determined (C). Mouse primary hepatocytes were treated with 0mM, 50mM and
100mM ethanol for 24 hours. Then, CES1 and HNF4 mRNA levels were tested (D).
*p<0.05, **p<0.01.
68
CES1 is regulated by HNF4and is its direct target.
Then, we determined how CES1 expression was regulated by alcohol. We
previously reported that CES1 is regulated by nuclear receptor farnesoid X receptor. We
then tested whether CES1 is regulated by HNF4. Figure 17 A and B showed that overexpression of HNF4increased mRNA levels of Ces1 in HepG2 cells and mouse
primary hepatocytes with inductions of 2.3 and 2.9 fold, respectively. In mice injected
with Ad-hHNF4, the mRNA level of Ces1 was increased by ~1.8 fold (Figure. 17 C). In
contrast to increased Ces1 mRNA level by over-expression of HNF4, CES1 mRNA
(Figure. 17D) and protein (Figure. 17E) levels were reduced ~50% in liver-specific
Hnf4mice. These data together suggested that HNF4 regulates CES1 expression
both in vitro and in vivo.
Next, we examined whether CES1 was a direct target of HNF4. Promoter
luciferase assays showed that HNF4 induced the luciferase activity by 10 fold, 4 fold
and 6 fold in the 1.9 kb, 0.98 kb and 0.3 kb Ces1 promoters, respectively, but not in the
0.25 kb Ces1 promoter (Figure 18A), suggesting that HNF4 may bind to the response
element(s) located between 0.3 kb and 0.25 kb of the Ces1 promoter. Indeed, there was a
potential DR-1 element (direct repeat separated by one base pair) between 300 bp and
287 bp upstream of the transcription start site (Figure 18 B). Mutation of the DR-1
element abolished the induction of Ces1 promoter activity by HNF4 (Figure 18 A). The
chromatin immunoprecipitation assay showed that HNF4 protein bound to the Ces1
promoter containing the DR-1 element in the liver (Figure 18C). Finally, electrophoretic
mobility shift assays showed that HNF4 protein bound to the DR-1 element and this
69
binding was competed away by the wild-type but not mutant DR-1 oligos (Figure 18D,
left panel). In addition, an HNF4 antibody was able to supershift the DNA/protein
complex (Figure 18D, right panel), indicating that HNF4 binds to the DR-1 element in
vitro. Collectively, the data of Figs. 17 and 18 demonstrate that HNF4 regulates Ces1
expression by binding to a DR-1 element located between 300 bp and 287 bp upstream of
the transcription start site.
70
Figure 17. CES1 is regulated by HNF4. HepG2 cells (A) and mouse primary
hepatocytes (B) were infected with empty adenovirus (Ad-empty) and adenovirus
expressing human HNF4(Ad- hHNF4n=3). After 48h, Ces1 mRNA levels were
analyzed. C57BL6 mice were injected with Ad-empty and Ad-hHNF4n=6). 7 days
later, hepatic Ces1 mRNA levels were tested (C). Hepatic CES1 mRNA (D) and protein
levels (E) were measured in Hnf4fl/fl and L-Hnf4mice (n=5). *p<0.05.
71
Figure 18. CES1 is a direct target of HNF4. Luciferase assay was performed using
pGL3-Ces1 and pGL3-Ces1-mut luciferase reporter constructs together with pCDNA3
and pCDNA3-HNF4 plasmids (n=6). After 36 hrs, luciferase activities were determined
and normalized to -galactosidase activity (A). Wild-type (WT) HNF4 response
element in Ces1 gene promoter was shown in the top. The mutant HNF4 response
element was shown at the bottom (B). Chromatin Immunoprecipitation (ChIP) assay was
performed using liver lysate and chromatin enrichment was determined (C). EMSA
assays were performed using in vitro transcribed/translated proteins. Wild-type (WT) and
mutant (MUT) oligos were used in the competition assays (left panel). Supershift assays
were performed in the presence of an HNF4 antibody (right panel) (D). *p<0.05,
**p<0.01.
72
Hepatic CES1 deficiency exacerbates alcohol-induced hepatic steatosis and
inflammation.
Our initial study showed that over-expression of Ces1 prevented alcohol-induced
lipid accumulation in AML-12 cells (Figure.19). This result coupled with decreased
expressions of CES1 in patients with alcoholic steatohepatitis implicates that CES1 plays
a crucial role in the pathogenesis of ALD. To test this hypothesis, we injected the
C57BL/6 mice with Ad-shLacZ or Ad-shCes1 together with Liber-DeCarli ethanol diet
administration as described previously [114]. Their body weights and food intake were
comparable (Figure. 20A-B). Alcohol treatment led to increased plasma levels of
triglyceride, total cholesterol, free fatty acids (FFA) in wild-type (shLacZ) mice (Figure.
20C-F). These observations were consistent with previous reports [101,114]. Compared
with ethanol-fed mice injected with shLacZ, ethanol-fed mice injected with shCes1
displayed elevated plasma triglyceride level (1.8 fold), slightly reduced plasma total
cholesterol level and unchanged plasma FFA and hepatic total cholesterol levels
(Figure.20C-F).
73
Figure 19. Over-expression of hepatic Ces1 protects against alcohol-induced
triglyceride accumulation in AML12 cells. AML12 cells were cultured in a 6-well
plates and infected with Ad-GFP and Ad-Ces1, followed by treatment with 50mM
ethanol for 24 hours. After 24 hours, cells were collected in 100L PBS buffer. 20L of
cell lysate were used for testing protein concentration. Lipids were extracted using
methanol: chloroform (1:2 v/v). Triglyceride levels were tested using Infinity kit
(Waltham, MA).
74
30
20
10
0
shLacZ
triglyceride(mg/dL)
C
Control
EtOH
250
** **
200
150
*
100
50
0
shLacZ
FFA(M)
1500
1000
shCes1
E
Control
EtOH
*
*
500
0
shLacZ
food intake
shLacZ
shCes1
15
10
5
0
y1 y2 y3 y4 y5 y6 y7 y8 y9 10
da da da da da da da da da ay
d
shCes1
Total cholesterol (mg/mL)
body weight (g)
40
before
after
Lieber DeCarli ethanol diet (mL)
body weight
D
200
**
150
*
Control
EtOH
*
100
50
0
shLacZ
Total cholesterol (g/mg)
A
B
F
Control
EtOH
5
4
3
2
1
0
shLacZ
shCes1
shCes1
shCes1
Figure 20 hepatic CES1 deficiency alters plasma lipid levels in response to alcohol
treatment. C57BL6 mice were subject to alcohol feeding described in figure 5. Their
body weights were shown before and after alcohol feeding (A). Food intake were
recorded (B). Plasma triglyceride, total cholesterol and FFA levels were tested (C-E).
Total cholesterol levels in the liver were determined and normalized to liver tissue weight
(F).
75
Furthermore, we found that knockdown of hepatic Ces1 (shCes1) significantly
increased hepatic TG and FFA levels (Figure. 21 A-B) in ethanol-fed mice. The higher
triglyceride and FFA levels may result from increased lipogenesis. Indeed, knockdown of
Ces1 in ethanol-fed mice markedly increased mRNA levels of lipogenic genes, including
sterol regulatory element-binding protein 1 (Srebp-1), fatty acid synthase (Fas), acetylCoA carboxylase-1 (Acc-1), acetyl-CoA carboxylase-2 (Acc-2), diacylglyceriol Oacyltransferase 2 (Dgat-2), and peroxisome proliferator-activated receptor (Ppar )
(Figure. 21C). Oil Red O staining revealed that ethanol-fed mice injected with shCes1
had a higher degree of hepatic steatosis than those injected with shLacZ (Figure. 21D).
76
Figure 21. Hepatic CES1 deficiency exacerbates alcohol-induced hepatic steatosis.
C57BL/6J mice (12 weeks old) were administered control Lieber-DeCarli diet for 5 days.
On the 6th day, mice were injected with Ad-shLacZ or Ad-shCes1. Then, mice were
either fed an ethanol Lieber-DeCarli diet containing 5% (vol/vol) ethanol or pair-fed a
control Lieber-DeCarli diet for 10 days. On the 16th day, mice were gavaged with a single
dose of ethanol (5g/kg body weight) or isocaloric maltose dextrin. Hepatic triglyceride (A)
and FFA (B) levels were measured. mRNA levels of lipogenic genes, including sterol
regulatory element-binding protein 1 (Srebp-1), fatty acid synthase (Fas), acetyl-CoA
carboxylase-1
(Acc-1),
acetyl-CoA
carboxylase-2
(Acc-2),
diacylglyceriol
O-
acyltransferase 2 (Dgat-2), and peroxisome proliferator-activated receptor (Ppar )
were tested (C). Liver sections of mice injected with Ad-shLacZ or Ad-shCes1 were
subject to Oil Red O staining (D). *p<0.05, **p<0.01, #p<0.05. In figure 4C, * represents
77
comparison between ethanol and control diet. # represents comparison between shCes1
and shLacZ.
78
Next, we determined whether hepatic CES1 deficiency resulted in more liver
injury. Plasma alanine aminotransferase (ALT) and aspartate aminotransferase (AST)
levels were higher in ethanol-fed mice injected with shCes1 (Figure. 22A-B). Compared
with ethanol-fed control mice, Ces1 deficiency increased mRNA levels of tumor necrosis
factor
(Tnf),
chemoattractant
interleukin-1
protein-1

(Mcp-1)
(IL-1), interleukin-6
(Figure.
22C-F).
(IL-6)
and
Interestingly,
monocyte
plasma
-
hydroxybutyrate levels were increased in ethanol-fed wild-type mice, but not in the
ethanol-fed hepatic Ces1 deficient mice (Figure. 22G), suggesting that hepatic CES1
deficiency may cause mitochondrial dysfunction. Collectively, hepatic CES1 deficiency
exacerbated alcohol-induced hepatic steatosis and liver inflammation.
79
Figure 22. Hepatic CES1 deficiency exacerbates alcohol-induced liver inflammation.
Mice were subject to alcohol feeding as described in figure 4. Plasma ALT and AST
80
levels were measured (A and B). mRNA levels of tumor necrosis factor (Tnf) (C),
interleukin-1  (IL-1) (D), interleukin-6 (IL-6) (E) and monocyte chemoattractant
protein-1 (Mcp-1) (F) were determined. Plasma -hydroxybutyrate level was tested (G).
*p<0.05, **p<0.01.
81
Global deletion of CES1 does not change alcohol-induced hepatic steatosis, but
exacerbates liver inflammation.
To test whether global deletion of CES1 exacerbates alcohol-induced liver injury,
Ces1 and Ces1 mice were administered a Lieber-DeCarbli ethanol diet containing
5% ethanol for 10 days followed by a single dose of ethanol (3g/kg) administration as
described [114]. Protein expressions of hepatic CES1 were completely abolished in
Ces1 mice (Figure. 23A). The chronic-binge feeding did not affect their body weights
(Figure. 23B). Compared with ethanol-fed Ces1mice, ethanol-fed Ces1mice
displayed moderately higher plasma triglyceride (~35%) and mildly lower total
cholesterol (~20%) levels (Figure. 23C-D). Surprisingly, hepatic triglyceride and hepatic
total cholesterol levels were comparable (Figure. 23E-F).
82
B
body weight
before
after
body weight (g)
40
30
20
10
250
200
150
Control
EtOH
*
**
100
50
0
Ces1
E
Triglyceride(g/mg)
*
Control
EtOH
40
30
Ces1
*
*
20
10
0
Ces1
Ces1
Total cholesterol (g/mg)
Triglyceride(mg/dL)
C
Total cholesterol (mg/mL)
0
Ces1
D
Ces1
Control
EtOH
150
100
10
50
0
Ces1
F
*
8
Ces1
Control
EtOH
6
4
2
0
Ces1
Ces1
Figure 23. global deletion of Ces1 does not exacerbate alcohol-induced hepatic
steatosis. Wild-type and Ces1 mice were subject to alcohol feeding described in the
legend of figure 6. The protein levels of hepatic CES1 was shown in (A). Body weights
were shown before and after alcohol feeding (B). Plasma triglyceride (C) and total
cholesterol (D) levels were tested. Hepatic triglyceride (E) and total cholesterol (F)
levels were determined.
83
Furthermore, ethanol feeding significantly increased plasma levels of ALT
(Figure. 24A) and AST (Figure. 24B) in both Ces1+/+ and Ces1 mice, and such
increases were further enhanced by Ces1 deficiency. In addition, ethanol significantly
induced hepatic the mRNA levels of Tnf (Figure. 24C), IL-1 (Figure. 24D), IL-6
(Figure. 24E) and Mcp-1 (Figure. 24F) in Ces1 mice, and the induction of Tnf, Il1
and Il6 was not significant in Ces1 mice (Figure 24C-24D). Importantly, Ces1
deficiency significantly potentiated ethanol-induced Tnf, Il1, Il6 and Mcp1 mRNA
levels in the liver (Figure 24C-24F). Overall, figure 23 and 24 demonstrate that global
deletion of CES1 exacerbates alcohol-induced liver inflammation, but does not change
hepatic steatosis.
84
Figure 24. Global deletion of CES1 exacerbates alcohol-induced liver inflammation.
Ces1 and Ces1 mice were administered control Lieber-DeCarli diet for 5 days,
followed by ethanol diet or pair-fed control diet administration for 10 days and a single
dose of ethanol (3g/kg body weight) gavage on the 16th day. Plasma ALT (A) and AST
(B) levels were measured. mRNA levels of Tnf (C), IL-1 (D), IL-6 (E) and Mcp-1(F)
were tested. *p<0.05, **p<0.01.
85
Global deletion of CES1 increases MCD diet-induced liver inflammation.
We then tested whether global deletion of CES1 affects MCD diet-induced liver
injury. MCD diet results in liver injury similar to human nonalcoholic steatohepatitis
(NASH). There was a 21% increase in hepatic TG level but unchanged hepatic
cholesterol level (Figure. 25A). Fat mass of Ces1 mice was higher (5.03%) than that in
wild-type mice (3.07%) (Figure. 25B). Plasma ALT level and mRNA levels of TnfIL1IL-6, and transforming growth factor  (Tgf) were potentiated in MCD diet-fed
Ces1 mice (Figure. 25C-G), suggesting that CES1 deficiency aggravates MCD dietinduced liver inflammatory response and liver injury.
Global deletion of CES1 does not change MCD diet-induced liver fibrosis.
We further determined MCD diet-induced liver fibrosis in wild type and Ces1
mice. Ces1 deficiency did not potentiate the expressions of figrogenic genes (Figure. 26),
neither did it aggravate MCD diet-induced fibrosis, which is shown by unchanged degree
of fibrosis in Sirius red stained liver sections (Figure. 27).
86
Figure 25. Global deletion of CES1 aggravates MCD diet-induced liver injury.
Ces1 and Ces1 mice were fed a MCD diet for 4 weeks. At the end of MCD diet
87
feeding, hepatic lipids were tested (A). Fat and lean mass were measured using EchoMRI
(B). Plasma ALT level (C) and mRNA levels of Tnf (D), IL-1 (E), IL-6 (F) and
TGF(G) were determined. *p<0.05, **p<0.01.
Figure 26. CES1 deficiency does not change fibrogenic gene expressions. Ces1 and
Ces1 mice were fed an MCD diet for 4 weeks. mRNA levels of -SMA ( -smooth
muscle actin), TGF-, Col1 (collagen 1) and TIMP (tissue inhibitor of
metalloproteinase) were determined.
88
Figure 27. Global deletion of CES1 does not exacerbate MCD diet-induced fibrosis.
Sirius Red staining was performed using the livers collected from MCD diet-fed mice.
89
Global deletion of CES1 results in increased hepatic acetaldehyde level, elevated
oxidative stress and mitochondrial dysfunction
To define how Ces1 deficiency aggravates ethanol-induced liver injury, we
analyzed acetaldehyde level in the liver. Acetaldehyde is a reactive compound, and is
highly toxic to hepatocytes because it can sensitize cells to oxidative stress or other
damaging signals, ultimately lead to mitochondria damage and cell death [92,118]. At the
basal level, hepatic acetaldehyde level was increased by 2.2 fold in Ces1 mice than in
Ces1 mice (Figure 28A). Upon ethanol feeding, hepatic acetaldehyde level was
increased by 1.7 fold in Ces1 mice, 2 fold in Ces1mice and 2.4 fold in Ces1 mice
than in Ces1 mice (Figure 28A). Consistent with this observation, the mRNA level of
acetaldehyde dehydrogenase 2 (Aldh2) was reduced by 50% in Ces1 mice (p<0.05; data
not shown).
Oxidative stress is an important factor in the pathogenesis of ALD. Mitochondrial
reactive oxygen species (ROS) triggers proinflammatory cytokine production, damages
mitochondrial DNA and promotes lipid peroxidation [119,120,121]. We examined
whether the aggravated liver inflammation in Ces1 mice was associated with increased
ROS. Indeed, figure 28B showed that mitochondrial H2O2 level was increased more than
3 fold in ethanol-fed Ces1 mice compared with ethanol-fed wild-type mice.
Consistently, malondialdehyde (MDA) level was higher (2 fold) in the livers of ethanolfed Ces1 mice (Figure. 28C). The increased production of ROS is a crucial cause of
mitochondrial DNA mutation, degradation and mitochondria dysfunction [122]. We then
tested mRNA levels of genes which encode mitochondrial enzymes, including
90
cytochrome oxidase subunit I (Cox1), cytochrome b (Cyt b), NADH dehydrogenase,
subunit (Nd1), and ATP synthase F0 subunit 6 (Atp6), all of which were significantly
down-regulated (Figure. 28D). Thus, global deletion of CES1 led to increased alcoholinduced ROS production, increased MDA and acetaldehyde formation, possibly resulting
in mitochondrial dysfunction.
91
B
**
*
200
150
*
100
**
50
0
Ces1
*
C
Relative mRNA
MDA(mol/g)
*
2
0
Ces1
E
400
Control
EtOH
300
*
200
100
0
Ces1
100
Ces1
* *
50
0
Ces1
Ces1
Ces1+/+ , EtOH
Ces1-/-, EtOH
1.0
*
*
0.5
*
0.0
Ces1
*
Control
EtOH
D
1.5
**
Control
EtOH
4
150
Ces1
6
FFA(nmol/mg protein)
H2O2 (M/mg protein)
250
Control
EtOH
Cox1
-hydroxybutyrate(M)
Acetaldehyde (nmol/g)
A
Cytb
Nd1
F
1000
800
Atp6
Control
EtOH
*
*
600
400
200
0
Ces1
Ces1
Figure 28. Global deletion of CES1 increases hepatic acetaldehyde level and
oxidative stress. Ces1 and Ces1 mice were subject to alcohol feeding as described in
the legend of figure 24. Hepatic acetaldehyde levels were measured using HPLC (A).
Mitochondrial H2O2 levels were measured using liver lysate and normalized to protein
concentration (B). MDA assay was performed using liver lysate and normalized to liver
92
weight (C). mRNA levels of mitochondrial genes were tested in ethanol-fed Ces1 and
Ces1 mice (D). -hydroxybutyrate (E) and FFA (F) levels were determined. *p<0.05,
**p<0.01.
93
Global deletion of CES1 inhibits alcohol-induced elevation of -hydroxybutyrate,
but it enhances hepatic FFA level.
Ethanol feeding significantly increases acetate level and therefore the synthesis of
-hydroxybutyrate (-HB). On the other hand, elevated mitochondrial FAO is also
accompanied by increased -HB synthesis. As such, plasma -HB level may reflect
mitochondrial functions under normal conditions. As shown in Figure 28F, ethanol
significantly increased plasma -HB level in Ces1 mice but not in Ces1 mice. In
addition, ethanol induced a 2.5-fold increase in hepatic free fatty acid (FFA) level in
Ces1 mice but not in Ces1 mice (Figure 28E). The increase in hepatic FFA level was
not accompanied by any change in genes involved in fatty acid synthesis (Srebp1c, Fas,
Acc1, Acc2), transport (CD36, Fabp1) or oxidation (Ppar, Cpt1, Mcad, Acox1, and
Acox2) (Figure 29). These data suggest that Ces1 mice may have an impaired
mitochondrial function.
94
Figure 29. global deletion of CES1 does not change mRNA levels of genes involved
in fatty acid metabolism. mRNA levels of genes were tested in ethanol-fed wild-type
and Ces1 mice. These genes are sterol regulatory-element binding protein 1c(Srebp1c),
fatty acid synthase (Fas), acetyl-CoA carboxylase 1 (Acc1), acetyl-CoA carboxylase 2
(Acc2), cluster of differentiation 36 (CD36), fatty acid binding protein 1 (Fabp1),
peroxisome proliferator activated receptor  (Ppar ), carnitine palmitoyltransferase 1-a
(Cpt1-), medium-chain acyl coenzyme A dehydrogenase (Mcad1), peroxisomal acylcoenzyme A oxidase 1 (Acox1), peroxisomal acyl-coenzyme A oxidase 2 (Acox2).
95
2.4 DISCUSSION
In the current study, we find that mRNA and protein expressions of CES1 and
HNF4 are markedly reduced in patients with alcoholic steatohepatitis and in mice
treated with alcohol. We clearly demonstrate that CES1 is transcriptionally regulated by
HNF4 and that it is a direct HNF4 target. Then, using chronic-binge alcohol feeding,
we find that hepatic CES1 deficiency exacerbates hepatic steatosis and inflammation in
response to alcohol challenge. In addition, global deletion of CES1 does not change
alcohol-induced hepatic steatosis, but it exacerbates alcohol and MCD diet-induced liver
inflammation, likely through increased hepatic acetaldehyde level, elevated oxidative
stress and enhanced lipid peroxidation.
Alcohol suppresses CES1 expression through, at least in part, repressing HNF4.
HNF4 is known to orchestrate a variety of genes associated with lipid, glucose
and bile acids metabolism, differentiation and morphogenesis [123]. Our results suggest
that ethanol inhibits CES1 expression through, at least in part, repressing HNF4.
However, how ethanol suppresses HNF4 expression is unknown. It is worth to note that
transcriptional activity of HNF4 is inhibited by unsaturated fatty acyl-CoAs with
greatest inhibition by C18:3, -3, and C20:5, -3 [124]. In addition, HNF4 protein
level is reduced by alcohol treatment in Sprague Dawley rats, but it is restored in alcohol
combined with saturated fatty acids treatment [125]. Extensive studies show that HNF4
protein level is decreased by unsaturated linoleic acid in HepG2 cells and primary
hepatocytes [125,126]. On the other hand, it is known that alcohol feeding alters fatty
96
acid composition. In absolute terms, alcohol feeding significantly increases hepatic levels
of monounsaturated FA (MUFA) and polyunsaturated FA (PUFA), but it does not change
hepatic saturated FA (SFA) levels [127]. Accordingly, we hypothesize that the increased
level of unsaturated fatty acids as a result of alcohol administration may partly contribute
to the suppression of HNF4. Given that tissue specific loss of HNF4in the liver results
in increased lipid deposits in hepatocytes [128], the suppression of HNF4CES1 axis
may have profound impact on the pathogenesis of alcohol-induced hepatic steatosis.
In addition, under the conditions of common metabolic stress (diabetes, obesity
and NAFLD, HFD feeding), elevated cholesterol, p53 and miR-34a orchestrate to inhibit
HNF4 expression. Thus, it is also possible that other unknown mechanism(s) are
involved in inhibiting HNF4 expression.
Global CES1 deficiency does not exacerbate alcohol-induced hepatic steatosis.
An unexpected finding in this study is that unlike ethanol-fed mice injected with
shCes1 which had increased ethanol-induced hepatic steatosis, ethanol-fed Ces1 mice
do not display aggravated hepatic steatosis. It is evident that CES1 also expresses in the
intestine; intestinal CES1 regulates chylomicron assembly [59]. We speculate that the
intestinal fat absorption may be diminished in Ces1 mice. Thus, in Ces1 mice, the
net effect of the absence of intestinal and hepatic CES1 on lipid homeostasis results in
unchanged hepatic triglyceride and cholesterol levels, compared with those in Ces1
mice. In addition, our previous report shows that loss of hepatic CES1 increases
lipogensis [1]. Our present study further proves that lipogenic gene expression levels are
97
elevated by the loss of hepatic CES1 and alcohol feeding. This finding suggests that
hepatic CES1 plays a protective role against ALD.
Global deletion of CES1 exacerbates alcohol-induced liver inflammation.
In the current study, global deletion of Ces1 aggravates ethanol-induced liver
inflammation without affecting hepatic steatosis. Although hepatic steatosis appears to
correlate with more severe forms of liver diseases which require the imposition of second
“hit” to incite hepatic inflammation and fibrosis, it has been well documented that
hepatocyte triglycerides per se are not hepatotoxic [129]. Our results support this point of
view. On the other hand, our results show that the hepatic level of FFA, a direct
hepatotoxic substance, is higher in ethanol-fed Ces1 mice than that in ethanol-fed wildtype mice. Mitochondrial gene expressions are reduced. This may be indicative of
mitochondrial dysfunction, which in turn affects mitochondrial  oxidation, resulting in
decreased FFA disposal and increased ROS production. Indeed, hepatic FAA,
mitochondrial H2O2, MDA levels are significantly elevated in ethanol-fed Ces1 mice.
Furthermore, hepatic acetaldehyde, a reactive hepatotoxic compound, is markedly
increased in ethanol-fed Ces1 mice. In addition to acetaldehyde, increased productions
of fatty acid ethyl ester (FAEE), palmitic acid are also associated with the pathogenesis
of ALD. It is shown that CES1 is involved in the synthesis of FAEE [130]. Thus, we will
not rule out other contributing factors to the aggravated liver injury in Ces1 mice. In
the present study, we find that -hydroxybutyrate is not induced by alcohol in Ces1
mice. -hydroxybutyrate is one of the ketone bodies which are synthesized from acetylCoA; it is increased in patients with excessive alcohol consumption [131]. Alcohol-
98
induced increase in -hydroxybutyrate level may be attributed to 1) increased acetate
which in turn becomes a source of acetyl-CoA [131] and 2) the increased ratio of
NADH/NAD+ which inhibits TCA cycle and promotes ketone body production. The
unchanged level of -hydroxybutyrate in response to alcohol challenge in Ces1 mice
remains unknown. We speculate that either the production of acetyl-CoA is reduced or
TCA cycle remains intact. All together, the elevated ROS, MDA and hepatic
acetaldehyde levels accompanied by increased hepatic FFA level are accountable for the
aggravated liver inflammation in ethanol-fed Ces1mice.
CES1 has the ability to hydrolyze amide or ester bonds. Our previous studies
show that TG is a substrate for CES1 [1]. Defective TG hydrolase activity in Ces1deficient hepatocytes is responsible for elevated TG accumulation in the liver. Ethanol is
unlikely to be a direct substrate of CES1. However, the metabolites of CES1 may
negatively affect alcohol metabolism and mitochondrial functions, leading to
accumulation of toxic substances, such as acetaldehyde and reactive oxygen species.
These toxic substances are sufficient to cause damages on mitochondria and cell
membranes, eventually resulting in liver inflammation and damage. One of our future
directions will be to determine which metabolite(s) cause elevation of these toxic
substances.
In summary, we have presented novel evidence that Ces1 deficiency exacerbates
alcohol-induced liver injury. Our findings further highlight that HNF4CES1 axis may
play an important role in the pathogenesis of ALD. The present studies identify CES1 as
a potential therapeutic target for treating ALD.
99
CHAPTER 3: THE ROLE OF CARBOXYLESTERASE 1 IN
ATHEROSCLEROSIS.
3.1 INTRODUCTION
Atherosclerosis-associated cardiovascular disease (CVD) is a leading cause of
death in the developed countries. It is characterized by increased lipid accumulation in
intima of the arterial walls, accompanied with chronic inflammation of arterial
endothelium [132]. Risk factors for this disease include obesity, dyslipidemia and type 2
diabetes; and keeping theses metabolic disorders under control is an effective strategy for
reducing cardiac events associated with atherosclerosis. High plasma cholesterol level is
a hallmark and a direct cause of atherosclerosis. Current pharmacological intervention
concentrates on reducing high cholesterol levels in patients, through enhancing
cholesterol efflux from macrophages, promoting reverse cholesterol transport (RCT),
enhancing plasma HDL level, and reducing plasma LDL level.
Atheromatous plaques, formed from fat, cholesterol, calcium and other substances,
build up in the inner lining of the arteries and impede blood flow. Life threatening events
such as myocardial infarction or stroke result from the rupture or ulceration of an
“unstable” plaque [133]. Inflammation and hypercholesterolemia are important features
of atherosclerosis. LDLs, trapped in the subendothelial space, are subject to oxidative
modification; and the oxidized lipids triggers monocyte recruitment to endothelial layer
of the artery [134]. Monocytes will then penetrate into the tunica intima and acquire
characteristics of the tissue macrophage. Furthermore, Foam cells, derived from
100
macrophages as a result of accumulation of lipid droplets, will secrete pro-inflammatory
cytokines to amplify the local inflammatory response in the lesion [135]. The progression
of lesion induces arterial calcification and predisposes to plaque rupture at sites of
monocytic infiltration. Oxidized LDL induces endothelial cells and monocytes to express
high levels of tissue factor in the lesion, and results in thrombosis, which is the proximate
cause of the clinical event [136].
Macrophage cholesterol efflux is shown to have antiatherosclerotic effect. Free
cholesterol efflux from macrophage foam cells requires ATP-binding cassette transporter
A1 (ABCA1) and ATP-binding cassette transporter G1 (ABCG1), which mediate free
cholesterol (FC) transportation to cholesterol-deficient and phospholipid depleted
apolipoprotien (apo)A1 and HDL in plasma, respectively [137] (Figure.30). Another
efflux pathway is mediated by the scavenger receptor class B type 1(SRB1), which
effluxes FC to the mature HDL. In addition, SRB1 also mediates cholesterol uptake in
macrophage. In macrophage, cholesteryl esters, accumulation of which causes “foamy”
characteristic of macrophage, are hydrolyzed to free cholesterol, which is an
antiatherosclerotic process in the body [137] (Figure.30).
101
Figure 30. macrophage cholesterol efflux. (picture is adapted from Rader, D et al.
Molecular regulation of HDL metabolism and function: implications for novel therapies.
J Clin Invest. 2006; 116(12))[138]
The antiatheroclerotic role of CESs has been explored. Over-expression of
macrophage human CES1, an equivalent form of mouse Ces3, leads to increases in
cholesteryl ester (CE) hydrolysis [51], mobilization of cytoplasmic CE [57], free
cholesterol (FC) efflux [58] and attenuation of atherosclerosis in Ldlr mice [51]. Overexpression of hepatic human CES1 has anti-atherogenic effects, through enhancing the
elimination of cholesterol into bile [139].
The studies in chapter 3 show the role of hepatic Ces1 in the development of
atherosclerosis. We find that hepatic CES1 deficiency aggravates western diet-induced
atherosclerosis in ApoE mice. This may result from increased cholesterol synthesis and
increased plasma cholesterol level in hepatic Ces1 deficient ApoE mice.
102
3.2 METHODS
Mice and diet.
8 week old ApoE mice were purchased from Jackson Laboratory (Bar Harbor,
Maine). ApoEmice were fed a western diet containing 0.21% cholesterol and 41% fat
from Research diets (New Burnswick, NJ) for 1 week, followed by adenovirus injection
with either shLacZ (n=7 per group) or shCes1 (n=7 per group). Mice were given a
western diet for another 3 weeks. 3 weeks after adenovirus injection, mice were
euthanized and their hearts and aortas were isolated as described [140,141].
Aorta isolation.
The aorta, including the ascending arch, thoracic, and abdominal segments, were
dissected, gently cleaned of the adventitia, and stained with Oil Red O as described
previously [140,141]. In addition, the aortic roots were collected from the base of the heart
including the atria and embedded in optimal cutting temperature compound. Sections (5
μm) were obtained every 50 μm from the base of the aortic leaflets to 400 μm above.
After staining with Oil Red O, images were captured with a microscope, and the lesion
area for each aortic ring was analyzed with Image J software [142].
Peritoneal macrophage isolation.
Peritoneal macrophages were isolated according to previous report [143].
C57BL/6J mice were injected with 1mL of 3% (w/v) brewer thioglycollate medium into
their peritoneal cavity. 3-5 days later, mice were euthanized. The outer skin of the
103
peritoneum was cut with gentle and pulled back to expose the inner skin lining the
peritoneal cavity. DMEM medium with 10%FBS was injected into the peritoneal cavity
using a 21.5g needle, followed by gently massage the peritoneum to dislodge any
attached cells into the medium. The cells were collected into a 5mL syringe. DMEM
medium was injected 1-2 more time to retrieve more cells. Cell suspension was spun at
1500rpm, resuspended and cultured in 6-well plates.
Lipid Analysis.
Plasma triglyceride and cholesterol were measured using Infinity reagent from
Thermo Scientific (Waltham, MA). To measure lipids in liver, approximately 100mg
liver tissue was homogenized in methanol and extracted in chloroform/methanol (2:1 v/v).
Hepatic triglyceride and cholesterol levels were then quantified using Infinity reagents
from Thermo Scientific (Waltham, MA).
Statistical Analysis.
The data were analyzed using unpaired Student t test and ANOVA (GraphPad
Prisim, CA). All values were expressed as meanSEM. Differences were considered
statistically significant at P<0.05.
104
3.3 RESULTS
The macrophages of Ces1 mice accumulate more lipids.
Our interest in studying the role of CES1 in atherosclerosis originates from
previous studies of macrophage CESs which are shown to protect against atherosclerosis.
In this regard, we fed wild type and Ces1 mice a western diet for 8 weeks and their
macrophages were isolated. Oil red O staining revealed that more lipids accumulated in
Ces1 deficient macrophages (Figure 31A). Macrophagic total cholesterol levels were
higher in Ces1 mice (Figure 31B). Thus, Ces1 deficiency increased total cholesterol
levels in macrophages.
Figure 31.
Global deletion of CES1 results in increased lipid accumulation in
macrophages. Oil red O staining of peritoneal macrophages of wild type (left penal) and
Ces1 mice (right penal) fed a western diet for 8 weeks (A). Total cholesterol levels
were measured and normalized to protein concentration (B).
105
Loss of hepatic CES1 increases plasma and hepatic lipids in ApoE mice.
To investigate whether hepatic Ces1 plays a role in atherosclerosis, ApoEmice
were fed a western diet for 1 week. Then mice were then injected with either Ad-shLacZ
or Ad-shCes1. Mice continued receiving western diet for another 3 weeks. 21 days after
adenovirus injection, plasma, liver and aorta were collected. Figure 32 showed that
plasma triglyceride, cholesterol levels were higher in Ces1 deficient ApoE mice (Figure.
32 A and B). In addition, hepatic triglyceride and cholesterol levels were higher in
ApoEmice injected with shCes1 (Fig. 32 C and D).
Figure 32. Loss of hepatic CES1 increases lipid contents in ApoE mice. ApoE
mice were fed a western diet for 1 week, followed by adenovirus injection of shLacZ or
shCes1. After virus injection, mice received a western diet for another 3 weeks. Plasma
106
triglyceride (A) and plasma cholesterol (B) were measured. Hepatic triglyceride (C) and
total cholesterol (D) were determined. *p<0.05, **p<0.01.
Loss of hepatic CES1 shows atherosclerotic lipid profile.
Plasma HLD-C and LDL-C are significant factors for determining the
atherosclerosis. Next, we determined the lipoprotein levels using fast protein liquid
chromatography (FPLC). Figure 33 showed that VLDL-TG was increased in ApoE
mice injected with shCes1 (Figure. 33A). Similarly, VLDL-C and LDL-C were higher in
those mice (Figure. 33B). Thus, these data suggest that hepatic Ces1 deficiency has
atherosclerotic lipid profile.
107
(A)
Triglyceride
(g/fraction)
40
sh-LacZ
sh-Ces1
30
20
10
30
27
24
21
18
15
12
9
6
3
0
0
Fraction
(B)
Cholesterol
(g/fraction)
150
sh-LacZ
sh-Ces1
100
50
27
24
21
18
15
12
9
6
3
0
0
Fraction
Figure 33. Loss of hepatic CES1 shows atherosclerotic lipid profile. ApoE mice
were subject to adenovirus injection and western diet feeding as described in the legend
of Figure 32. Plasma lipoprotein levels were measured using FPLC (A and B).
108
Loss of hepatic CES1 aggravates atherosclerosis in ApoE mice.
To further determine whether loss of hepatic Ces1 affects the development of
atherosclerosis, we isolated the aortas and stained them with Oil Red O. Oil red O
staining revealed that loss of hepatic Ces1 increased lipid accumulations in the arterial
walls (Figure. 34 A). A further quantification confirmed that the aorta lesion areas were
larger in Ces1 deficient ApoE mice than those in their control counterparts (Figure.
34B). The lesion size of aortic root in Ces1 deficient ApoE mice is larger than that in
control mice (Figure 34C and D). Thus, Ces1 deficiency aggravates atherosclerosis.
109
A
B
10
shLacZ
Lesion area%
8
shCes1
6
4
2
0
shLacZ
shLacZ
shCes1
D
Heart lesion (mm2/section)
C
shCes1
1.010 5
8.010 4
6.010 4
4.010 4
2.010 4
0
shLacZ
shCes1
Figure 34. Loss of hepatic CES1 aggravates atherosclerosis in ApoE mice. ApoE
mice were subject to western diet feeding and adenovirus injection as described in the
legend of Figure 32. Aortas of control mice (A, top penal) and Ces1 deficient mice (A,
bottom penal) were isolated and stained with Oil red O. The lesion areas were analyzed
using commercially available software and were shown in (B). Aortic root of control (top
penal) and Ces1 deficient mice (bottom penal) was stained by oil red O (C) and lesion
areas were quantified (D).
110
3.4 DISUSSION
In chapter 3, we studied the role of CES1 in the development of atherosclerosis.
CES3, a murine form of human CES1, has been shown to promote macrophage
cholesterol efflux and cholesterol elimination in liver. In the present study, we showed
that hepatic Ces1 deficiency increases plasma and hepatic lipids levels, and increases
VLDL-TG, VLDL-C and LDL-C levels, all of which directly result in atherosclerosis.
We have previously shown that hepatic Ces1 deficiency results in increased lipogenesis
and enhanced VLDL secretion, both of which may contribute to the increased plasma
lipid levels, eventually lead to atherosclerosis (Figure 32). Collectively, these data
suggest that hepatic Ces1 deficiency alters systemic lipid homeostasis, likely via
enhancing lipogenic genes expressions.
Despite hepatic Ces1 deficient mice show striking changes in lipid homeostasis,
global deletion of Ces1 does not significantly change plasma and hepatic lipid levels in
response to western or high fat diet challenge, except that plasma cholesterol level is
slightly decreased (~20%) (data not shown). More tissue-specific functional studies are
needed to clearly understand the role of Ces1 in regulation of lipid metabolism. Since the
lipid phenotype in Ces1 mice is different from that in hepatic Ces1 deficient mice, we
generated Ces1Ldlrmice and these mice are fed a western diet to induce
atherosclerosis. The double knockout mice will help us understand the role of
macrophagic and hepatic Ces1 in the development of atherosclerosis.
111
In summary, using adenovirus gene delivery system, we find that loss of hepatic
Ces1 aggravates atherosclerosis. However, more data have to be collected from
Ces1Ldlrmice to clearly understand the role of Ces1 in regulation of atherosclerosis.
112
CHAPTER 4: CONCLUSION
CES1 is a phase I drug metabolizing enzyme which is shown to be involved in
biotransformation and detoxification of a variety of drugs and prodrugs, including
angiotensin-converting enzyme inhibitors, necrotic, anti-tumor and anti-virus drugs.
Given its nature of hydrolyzing ester and amide bond, CES1 is reported to have
triglyceride hydrolase and cholesteryl ester hydrolase activity. Previous in vitro studies
show that CES1 hydrolyzes triglyceride and promotes fatty acid oxidation.
Our studies focus on in vivo elucidation and delineation of CES1 in regulation of
lipid and carbohydrate metabolism. First, we find that in diabetic mouse models, CES1
expression is significantly increased; and we further demonstrate that CES1 is induced by
glucose. The induction of CES1 expression, in turn, reduces plasma glucose levels,
through increasing insulin sensitivity. Second, we clearly demonstrate that overexpression of hepatic CES1 protects against NAFLD, through reducing hepatic
triglyceride and promoting fatty acid oxidation. In contrast, knockdown of hepatic CES1
induces hepatic steatosis by increasing SREBP processing and lipogenesis. We also find
that CES1 is a farnesoid X receptor (FXR) target gene, and that activation of FXR
reduces hepatic and plasma triglyceride levels through, at least in part, inducing CES1.
Third, we show that CES1 is inhibited by alcohol, and that CES1 deficiency exacerbates
alcohol-induced liver injury, possibly resulting from increased hepatic acetaldehyde
113
level, elevated ROS level and enhanced lipid peroxidation, suggesting that CES1 plays a
protective role against ALD. We also find that CES1 is a direct target of HNF4, and
HNF4-CES1 axis may have a profound impact on the development of ALD. Lastly, we
demonstrate that loss of hepatic CES1 aggravates western diet-induced atherosclerosis in
ApoE mice. These exciting data lead us to conclude that CES1 plays an essential role in
regulating lipid and carbohydrate metabolism, and also protects against NAFLD, ALD
and atherosclerosis.
One of our future directions will focus on studying how CES1 regulates insulin
sensitivity. Different from deletion of hepatic CES1 which causes insulin resistance,
global deletion of CES1 does not change insulin sensitivity (data not shown). How CES1
affects systemic insulin sensitivity and whether CES1 in adipose tissue or muscle is
involved in the regulation of insulin signaling will be our future focuses. Furthermore,
Ces1 mice display reduced plasma cholesterol level (~20%) compared to wild type
mice (data not shown). Therefore, we will investigate whether the fat absorption is
changed in Ces1 deficient mice. In addition, we plan to use liquid chromatography-mass
spectrometry (LC-MS) to study the endogenous substrates and metabolomics of hepatic
CES1 in mice treated with alcohol or western diet. This research will help us understand
how CES1 deficiency causes liver injury. Lastly, we are interested in investigating the
pathway mediated by CES1 in regulation of the development of atherosclerosis. We plan
to use macrophage and liver specific Ces1 deficient Ldlrmice as well as global Ces1
deficient Ldlrmice to study the underlying mechanism.
113
In conclusion, the present studies underscore CES1 as a potential therapeutic
target in treating metabolic disorders. We propose that manipulation of CES1 expression
is a plausible strategy for managing metabolic disease.
115
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