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Effects of zinc on retinoic acid-induced growth inhibition in human hepatocarcinoma HepG2 cells Ibbitson, Deanna 2012

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  Effects
of
zinc
on
retinoic
acid‐induced
growth
inhibition
in
human
 hepatocarcinoma
HepG2
cells
 
 
  by
 DEANNA
IBBITSON
 B.Sc.,
The
University
of
British
Columbia,
2007
 
 A
THESIS
SUBMITTED
IN
PARTIAL
FULFILLMENT
OF
THE
REQUIREMENTS
FOR
 THE
DEGREE
OF
 
 MASTER
OF
SCIENCE
 in
 THE
FACULTY
OF
GRADUATE
STUDIES
 
(Human
Nutrition)
 
 THE
UNIVERSITY
OF
BRITISH
COLUMBIA
 (Vancouver)
 
 January
2012
 
 
 ©
Deanna
Ibbitson,
2012
  Abstract
 
 Retinoic
acid
(RA),
a
bioactive
metabolite
of
vitamin
A,
inhibits
growth
in
a
variety
 of
 cancer
 cells
 including
 liver
 cancer.
 It
 is
 thought
 that
 this
 function
 of
 RA
 is
 achieved
 by
 modulating
 gene
 expression
 through
 complexing
 with
 retinoic
 acid
 receptors
 (RARs)
 and
 retinoid
 X
 receptors
 (RXRs),
 two
 groups
 of
 zinc‐finger
 proteins.
 
 Zinc
 deficiency
 has
 been
 shown
to
affect
gene
expression
and
to
impair
DNA
binding
ability
of
zinc‐finger
proteins.
 The
hypotheses
of
my
thesis
research
were:

1)
sufficient
cellular
zinc
level
is
important
for
 the
effectiveness
of
RA‐induced
growth
inhibition
in
hepatocarcinoma
HepG2
cells;
and
2)
 the
influence
of
zinc
on
RA‐induced
growth
inhibition
is
through
modulating
expression
or
 function
 of
 RARs
 and
 RXRs,
 which
 in
 turn
 affects
 the
 expression
 of
 their
 target
 genes,
 CYP26a1
and
RARβ.

The
overall
objective
was
to
examine
the
effects
of
zinc
on
RA‐induced
 growth
inhibition
in
HepG2
and
the
possible
mechanisms
involved.

 
 Zinc
manipulation
was
achieved
by
culturing
HepG2
cells
for
6
d
in
low‐zinc
media
 supplemented
 with
 0,
 5,
 and
 10
 µmol/L
 zinc
 to
 mimic
 low‐,
 adequate‐,
 and
 high‐zinc
 conditions.
 Growth
 in
 low‐zinc
 media
 for
 6
 d
 reduced
 total
 cellular
 zinc
 and
 the
 labile
 intracellular
pool
of
zinc
by
29
and
86%,
respectively.

Treating
the
cells
with
35
µM
of
RA
 for
 12
 h
 following
 zinc
 manipulation
 significantly
 reduced
 cell
 proliferation
 in
 all
 zinc‐ treatment
groups
compared
to
their
corresponding
RA
control,
with
the
greatest
reduction
 in
 the
 high‐zinc
 group.
 
 Cell
 cycle
 analysis
 showed
 that
 the
 proportion
 of
 cells
 in
 the
 S‐ phase
 was
 reduced
 by
 RA
 treatment
 at
 24
 and
 72
 h
 at
 all
 zinc
 levels,
 with
 the
 greatest
 reduction
 in
 cells
 cultured
 in
 high‐zinc
 medium.
 Following
 growth
 in
 low‐,
 adequate‐
 and
  
  ii
  high‐zinc
 medium,
 RA
 treatment
 elevated
 the
 abundance
 of
 RARβ
 and
 Cyp26a1
 mRNA
 equally
in
all
zinc‐treatment
groups
compared
to
their
correspondent
RA
controls.

Growth
 in
 low
 zinc
 medium
 increased
 mRNA
 abundance
 of
 RXRα
 while
 RARα,
 RARβ,
 RARγ,
 RXRβ
 and
 RARγ
 were
 not
 affected.
 
 In
 conclusion,
 these
 results
 showed
 that
 increasing
 zinc
 appeared
 to
 sensitize
 HepG2
 cells
 to
 RA‐induced
 growth
 inhibition,
 but
 had
 no
 effect
 on
 RA‐induced
gene
expression
of
CYP26a1
and
RARβ.

 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
  
 
  iii
  Preface
 
 
  This
thesis
was
prepared
in
accordance
to
University
of
British
Columbia
Faculty
of
  Graduate
Studies
requirements.
I
was
responsible
for
performing
all
experiments.
The
 research
design,
interpretation
of
the
results,
and
preparation
of
this
thesis
were
 accomplished
with
the
assistance
and
guidance
of
Dr.
Zhaoming
Xu.

  
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
  
  iv
  Table
of
Contents
 
 Abstract ................................................................................................................................... ii
 Preface ................................................................................................................................... iv
 Table
of
Contents…………………………………………………… ............................................................. v
 List
of
Tables.......................................................................................................................... vii
 List
of
Figures ....................................................................................................................... viii
 List
of
Abbreviations................................................................................................................ x
 Acknowledgements ............................................................................................................... xii
 Introduction ............................................................................................................................1
 
 Chapter
1.
Literature
Review,
Hypothesis,
and
Objectives .............. ..…………………………..……..3
 
 1.1.

Vitamin
A
Nutrition
and
Function .............................................................3
 
 1.2.

Zinc
Nutrition
and
Function ......................................................................9
 
 
 1.2.1.
Zinc–Finger
Proteins……………………………….... …………………………....11
 
 1.3.

Interactions
Between
Zinc
and
Vitamin
A…………… .…………………….……….12
 
 1.4.

The
Role
of
Vitamin
A
in
Cancer……………………… .....……………………………..14
 
 
 1.4.1.

Effect
of
Retinoic
Acid
on
the
Growth
of
HepG2…… .....…..……….17
 
 
 1.4.2.

The
Role
of
Retinoic
Acid
Receptor
Beta
in
Cancer…… ....………...18
 
 
 1.4.3.

CYP26a1
in
Cancer…………………………………………………....……..……..18
 
 1.5.

Role
of
Zinc
in
Cancer…………………………..………………………………………….....19
 
 
 1.5.1.


Zinc
in
Hepatocarcinoma……………………………… ....…………………....20
 
 1.6.

Molecular
Effects
of
Zinc
Deficiency………… ...………………………………………21
 
 1.7.

Summary .................................................................................................25
 
 1.8.

Hypothesis...............................................................................................25
 
 1.9.

Overall
Objectives
and
Specific
Aims ......................................................25
 
 Chapter
 2.
 Effects
 of
 Zinc
 on
 Retinoic
 Acid‐Induced
 Growth
 Inhibition
 in
 Human
 Hepatocarcinoma
HepG2
Cells……… .........................................…………………………………..…....27
 
 2.1
Introduction ..............................................................................................27
 
 2.2
Materials
and
Methods……………………………………………………………………….. .29
 
 
 2.2.1.
Cell
Culture
System………………………………………….......………………….29
 
 
 2.2.2.
Preparation
of
Chelex
100‐Treated
FBS………………… .......…………..29
 
 
 2.2.3.
Zinc
Manipulation…………………………………………………….... ……………30
 
 
 2.2.4.
Quantification
of
the
Labile
Intracellular
Pool
of
Zinc
(LIPZ)........30
 
 
 2.2.5.
Quantification
of
Total
Cellular
Zinc…………………………………… ......32
 
 
 2.2.6.
Assessment
of
Cell
Viability
and
Proliferation……………………........32
 
 
 2.2.7.
Cell
Cycle
Analysis……………………………………………………………….......34
 
 
 2.2.8.
RNA
Extraction
and
Real
Time
Quantitative
PCR………..……… .......35
 
 
 2.2.9.
Protein
Extraction
and
Western
Blot
Analysis…………………….... ….37
 
 
 2.2.10.
Statistical
Analyses………………………………………………………….... …..39
 
 2.3.
Results......................................................................................................39
 
 
 2.3.1.
RA‐Mediated
Reduction
in
Cell
Proliferation… ............................39
  
  v
  
  2.3.2.
 Treatment
 Duration‐Dependent
 Reduction
 in
 Total
 Cellular
 Zinc
Level
and
the
Abundance
of
LIPZ ............................................ …39
 2.3.3.
Dose‐Dependent
Reduction
in
Total
Cellular
Zinc
Level
and
the
 Abundance
of
LIPZ……........................................................................40
 2.3.4.
 Zinc
 Appeared
 to
 Sensitize
 HepG2
 Cells
 to
 RA‐Induced
 reduction
in
Cell
Proliferation……… ................................................ …40
 2.3.5.
Cell
Viability
Was
Unaffected
by
RA
and
Zinc……................. ….42
 2.3.6.
 Growth
 of
 HepG2
 in
 the
 Low‐Zinc
 Medium
 Increased
 mRNA
 Abundance,
 But
 Not
 Protein
 Levels
 of
 the
 RXRα
 Nuclear
 Receptor……… .............................................................................. ……42
 2.3.7.
 Zinc
 Treatment
 Did
 Not
 Affect
 the
 RA‐Induced
 Increase
 in
 mRNA
Abundance
of
Downstream
Targets,
RARβ
and
CYP26a1…....42
 2.4.
Discussion.................................................................................................43
 2.4.1.
Growth
in
Medium
Depleted
of
Zinc
Was
Effective
in
Reducing
 Cellular
Zinc
in
HepG2……… ......................................................... ……43
 2.4.2.
 Zinc
 Appeared
 to
 Sensitize
 HepG2
 Cells
 to
 RA‐Induced
 Reduction
in
Cell
Proliferation………...........................................………45
 2.4.3.
 Growth
 of
 HepG2
 in
 Low
 Zinc
 Media
 Resulted
 in
 Increased
 mRNA
 Abundance,
 But
 Not
 Protein
 Levels
 of
 the
 RXRα
 Nuclear
 Receptor…………….......................................................................... …..47
 2.4.4.
 Zinc
 Treatment
 Did
 Not
 Affect
 the
 RA‐Induced
 Increase
 in
 mRNA
Abundance
of
Downstream
Targets,
RARβ
and
CYP26a1 ......48
 2.4.5.
Summary............................................................................. …..50
  
 Chapter
3.

General
Discussion,
Limitations,
and
Future
Directions .....................................62
 
 3.1
General
Discussion ....................................................................................62
 
 3.2
Limitations.................................................................................................65
 
 3.3
Future
Directions ......................................................................................66
 
 Bibliography………….……………………………………………………………………..…………………….…………….68
 Appendices.………………………………………………………………………………………………………………………79
 
 
 
 
  
  vi
  List
of
Tables
 
 1.1.  Examples
of
genes
affected
by
zinc
status......................................................24
  2.1.
  The
effect
of
retinoic
acid
on
proportion
of
cells
in
G1‐
and
S‐phases
following
 zinc
treatment.................................................................................................51
  A.1.
  
Mineral
supplements
added
to
Chelex‐100
treated
medium........................80
  A.2.
  Primer
sequences
for
real
time
quantitative
PCR ...........................................81
  A.3.
  Western
blot
antibody
dilutions
and
incubation
times ..................................82
  
 
 
 
 
 
 
 
 
 
 
 
  vii
  List
of
Figures
 1.1.

  Structure
of
all‐trans
retinol,
all‐trans
retinal
and
all‐trans
retinoic
acid .........7
  1.2.
  Mechanism
of
retinoic
acid‐induced
gene
transcription ..................................8
  2.1.

  Cell
proliferation
of
HepG2
cells
treated
with
retinoic
acid............................52
  2.2.
  Time‐dependent
 reduction
 in
 total
 cellular
 zinc
 level
 and
 the
 abundance
 of
 the
labile
Intracellular
pool
of
zinc..................................................................53
  2.3.

  Dose‐dependent
 reduction
 in
 total
 cellular
 zinc
 level
 and
 the
 abundance
 of
 the
labile
intracellular
pool
of
zinc..................................................................54
  2.4.

  The
 effect
 of
 retinoic
 acid
 on
 the
 proliferation
 of
 HepG2
 cells
 following
 growth
in
low‐,
adequate‐,
and
high‐zinc
media ............................................55
  2.5.
  The
effect
of
retinoic
acid
on
the
proportion
of
cells
in
the
S‐phase
following
 growth
in
low‐,
adequate‐,
and
high‐zinc
media ............................................56
  2.6.
  The
effect
of
retinoic
acid
on
the
viability
of
HepG2
cells
following
growth
in
 low‐,
adequate‐,
and
high‐zinc
media.............................................................57
  2.7.
  The
 relative
 expression
 of
 RAR
 and
 RXR
 nuclear
 receptor
 mRNA
 in
 HepG2
 following
growth
in
low‐,
adequate‐,
and
high‐zinc
media.............................58
  2.8.

  Western
 blot
 of
 RXRα
 and
 RARα
 following
 growth
 in
 low‐,
 adequate‐,
 and
 high‐zinc
medium............................................................................................59
  2.9.
  The
 relative
 expression
 of
 RARβ
 mRNA
 in
 RA‐treated
 HepG2
 cells
 following
 growth
in
low‐,
adequate‐,
and
high‐
zinc
media ...........................................60
  2.10.
  The
 relative
 expression
 of
 CYP26a1
 mRNA
 in
 RA‐treated
 HepG2
 cells
 following
growth
in
low‐,
adequate‐,
and
high‐zinc
media.............................61
  
  viii
  A.1.
  
  Total
zinc
levels
in
HepG2
and
AML12
cells ....................................................83
  A.2.
  
  Cell
proliferation
of
AML12
cells
treated
with
retinoic
acid ...........................84
  A.3.
  The
 effect
 of
 RA
 on
 viability
 of
 AML12
 cells
 following
 growth
 in
 low‐,
 adequate‐,
and
high‐zinc
media......................................................................85
  A.4.
  The
 effect
 of
 retinoic
 acid
 on
 the
 proliferation
 of
 AML12
 cells
 following
 growth
in
low‐,
adequate‐,
and
high‐zinc
media ............................................86
  
  ix
  List
of
Abbreviations
 
 ADH
 APL
 
 AP‐1
 
 BrdU
 
 CDK
 
 cDNA
 
 CRABPII
 d
 DMSO
 
 DNA
 
 EMSA
 
 FABP
 
 G1
 
 G1/S
 
 G2/M
 
 h
 
 hZIP1
 
 IGFI
 
 IU
 
 LIPZ
 
 MAP
 
 min
 
 mRNA
 
 NFκΒ
 
 Pol
II
 
 PCR
 
 PTC
 
 RA


 
 RAL
 
 RARE
 
 RARα RARβ RARγ RAS
 
 RBP
 
 RDA
 
 RE
 
 RNA
 
 ROL
 
 RXRE
 
  
  
 alcohol
dehydrogenase
 acute
promyelocytic
leukemia
 activator
protein
1
 5‐bromo‐2’‐deoxyuridine
 cyclin‐dependent
kinase
 complementary
DNA
 cellular
retinoic
acid‐binding
protein
II
 day
 dimethyl
sulfoxide
 deoxyribonucleic
acid
 electromobility
shift
assay
 fatty
acid
binding
protein
 gap
1
 gap
1/synthesis
 gap
2/mitosis
 hour
 human
ZRT,
IRT‐like
protein
1
 insulin‐like
growth
factor
I
 international
units
 labile
intracellular
pool
of
zinc
 mitogen‐activated
protein
 minute
 messenger
RNA
 nuclear
factor
κΒ DNA
polymerase
II polymerase
chain
reaction pseudotumor
cerebri
 retinoic
acid
 retinal
 retinoic
acid
response
element
 retinoic
acid
receptor
α
 retinoic
acid
receptor
β
 retinoic
acid
receptor
γ retinoic
acid
syndrome
 retinol
binding
protein
 recommended
dietary
allowance
 retinyl
ester
 ribonucleic
acid
 retinol
 retinoid
X
response
element
  x
  RXRα RXRβ RXRγ S‐phase
 TGF‐β TPEN
 
 ZIP
 
 ZIP4
 
 ZIP14
 
 ZnT
 
 ZnT5
 

 
  retinoid
X
receptor
α
 retinoid
X
receptor
β
 retinoid
X
receptor
γ synthesis
phase
 transforming
growth
factor‐β N,N,N',N'‐tetrakis(2‐pyridylmethyl)ethylenediamine
 ZRT,
IRT‐like
protein
 ZRT,
IRT‐like
protein
4
 ZRT,
IRT‐like
protein
14
 zinc
transporter
 zinc
transporter
5
  
 
 
 
 
 
 
 
 
 
  
  xi
  Acknowledgements
 I
would
like
to
thank
everybody
at
UBC
who
helped
me
throughout
the
process
of
 completing
my
thesis.
Specifically,
I
would
like
to
thank
my
advisory
committee
members:
 Dr.
 Christine
 Scaman
 and
 Dr.
 Tim
 Green,
 my
 supervisor:
 Dr.
 Zhaoming
 Xu,
 my
 lab
 mates:
 Alice
Lin,
Melinda
Bakker,
Wendy
Hempstock,
Jonathan
Lee,
Sylvia
Lymburner,
and
Markus
 Purtzki,
 and
 the
 Lund
 lab.
 Special
 thanks
 goes
 to
 Helen
 Chan
 who
 spent
 countless
 hours
 washing
 dishes,
 autoclaving,
 filling
 pipette
 boxes
 and
 assisting
 with
 experiments,
 always
 with
a
smile
on
her
face.
Last,
but
not
least,
thank
you
to
Daniel
Ahkong
and
my
parents
for
 their
moral
and
emotional
support.

 My
 thesis
 research
 project
 was
 supported
 by
 the
 FNH
 Vitamin
 Research
 Fund
 (Z.
 Xu).
 
 
 
 
 
 
 
 
 
  
 
 
  xii
  Introduction
 
 
 
  Zinc
 and
 vitamin
 A
 are
 both
 essential
 nutrients
 for
 humans
 with
 numerous
  overlapping
physiological
roles
such
as
immune
function
and
fetal
development
(Vallee
and
 Falchuk,
 1993;
 Mark
 et
 al.,
 2006).
 Both
 nutrients
 are
 involved
 in
 growth
 regulation
 and
 cellular
 differentiation
 (Blomhoff
 and
 Blomhoff,
 2006;
 Li
 et
 al.,
 2007).
 There
 is
 evidence
 that
 zinc
 is
 required
 for
 the
 function
 of
 vitamin
 A
 at
 the
 whole
 body
 level,
 including
 the
 release
of
vitamin
A
from
the
liver
into
the
blood,
as
well
as
vitamin
A
metabolism
(Smith,
 1980;
 Boron
 et
 al.,
 1988).
 However,
 information
 on
 the
 interactions
 between
 the
 two
 nutrients
at
the
cellular
level
is
much
more
limited.

 
 Vitamin
A,
in
the
form
of
retinoic
acid
(RA),
is
used
clinically
as
a
chemotherapeutic
 agent
for
numerous
types
of
cancer.
Growth
suppression
following
RA
treatment
has
been
 observed
 in
 numerous
 types
 of
 cancerous
 cells
 including
 breast
 cancer,
 liver
 cancer,
 prostate
 cancer,
 non‐small‐cell
 lung
 cancer,
 thyroid
 cancer
 and
 myeloid
 leukemia
 cells
 (Bohnsack
and
Hirschi,
2004,
Nakanishi
et
al.,
2008).
This
 function
 of
RA
is
thought
to
be
 due
to
its
regulatory
role
in
the
expression
of
growth‐regulating
genes
(Schug
et
al.,
2007).
 RA
exerts
its
regulatory
function
in
gene
expression
by
forming
a
complex
with
members
of
 two
families
of
zinc‐finger
nuclear
receptors:
retinoic
acid
receptors
(RARs)
and
retinoid
X
 receptors
(RXRs).
This
RA‐receptor
complex
binds
to
a
specific
DNA
region,
the
retinoic
acid
 response
 element
 (RARE),
 to
 initiate
 the
 expression
 of
 the
 target
 genes
 (Soprano
 and
 Soprano,
2002).


 
  1
  Aberrations
 in
 intracellular
 zinc
 content
 have
 been
 observed
 in
 many
 types
 of
 tumors
 (Franklin
 and
 Costello,
 2009).
 For
 example,
 zinc
 content
 in
 hepatocarcinoma
 is
 markedly
reduced
in
comparison
to
non‐cancerous
liver
cells,
however
the
consequences
 of
 this
 altered
 zinc
 status
 on
 cell
 growth
 and
 response
 to
 treatment
 are
 not
 well
 understood
(Gurusamy
and
Davidson,
2007).

Although
zinc
has
been
shown
to
affect
gene
 expression,
enzyme
function
and
the
ability
of
zinc‐containing
nuclear
receptors
to
bind
to
 DNA,
it
is
not
clear
whether
intracellular
zinc
levels
are
important
for
the
function
of
RA
at
 the
cellular
level
(Satre
et
al.,
2001;
Dieck
et
al.,
2003;
Rana
et
al.,
2008).
 
 Due
to
the
important
role
of
vitamin
A
in
many
physiological
processes
and
in
anti‐ cancer
 treatment,
 it
 is
 important
 to
 understand
 the
 effects
 of
 other
 essential
 nutrients,
 including
zinc,
on
the
function
of
vitamin
A.

 
 
 
 
 
 
 
 
  
 
  
  2
  Chapter
1.
Literature
Review,
Hypothesis,
and
Objectives
 
 
 1.1.
Vitamin
A
Nutrition
and
Function
 Vitamin
 A
 refers
 to
 a
 family
 of
 molecules
 derived
 from
 a
 monocyclic
 parent
 compound
containing
five
carbon‐carbon
double
bonds
and
a
functional
terminal
group
at
 the
terminus
of
the
acyclic
portion
(Blomhoff
and
Blomhoff,
2006).
The
two
major
isomers
 of
 vitamin
 A
 are:
 retinol
 (ROL)
 and
 retinal
 (RAL).
 RAL
 can
 be
 metabolized
 to
 retinoic
 acid
 (RA),
which
is
also
biologically
active.

Vitamin
A
is
stored
in
the
liver
as
retinyl
esters
(RE).
 Foods
of
animal
origin
contain
ROL
or
RE.

Foods
of
plant
origin
contain
carotenoids,
which
 can
 be
 converted
 to
 vitamin
 A
 within
 the
 intestinal
 cell
 during
 digestion.
 Within
 the
 intestinal
cell,
carotenoids
are
converted
to
RAL
and
then
ROL.

Upon
release
from
hepatic
 stellate
 cells,
 RE
 is
 hydrolyzed
 by
 hydrolases
 to
 release
 ROL,
 which
 is
 then
 transported
 through
 the
 blood
 to
 target
 tissues.
 The
 ROL
 released
 can
 be
 metabolized
 to
 RA
 via
 RAL
 within
hepatic
and
extrahepatic
cells
(Blomhoff
and
Blomhoff,
2006).
  The
dietary
reference
intakes
for
vitamin
A
for
adults
are:
700
µg/d
(2333
IU/d)
for
 females
and
900
µg/d
(3000
IU/d)
for
males
(Health
Canada,
2010).
Common
food
sources
 of
 pre‐formed
 vitamin
 A
 include
 animal
 sources
 such
 as
 liver,
 kidney,
 fatty
 fish,
 dairy
 products
and
eggs.
Plant
foods
containing
vitamin
A
precursors
(carotenoids)
include
green
 leafy
 and
 orange
 and
 yellow
 vegetables
 and
 orange
 colored
 plants
 (Weber
 and
 Gruve,
 2011).

  
  3
  Vitamin
 A
 is
 important
 for
 most
 forms
 of
 life
 and
 is
 required
 for
 many
 biological
 processes.

The
most
well‐known
and
understood
function
of
vitamin
A
is
its
role
in
night
 vision,
as
it
is
needed
for
the
formation
of
the
photosensitive
visual
pigment
of
the
retina
 (Rando,
 1990;
 Darlow
 and
 Graham,
 2007).
 
 Vitamin
 A
 is
 also
 important
 in
 many
 other
 processes
 including
 immune
 defense
 and
 the
 regulation
 of
 growth
 and
 development
 (Semba,
1998;
Clagette‐Dame
and
Deluca,
2002).
Vitamin
A,
as
ROL,
maintains
the
integrity
 of
skin
and
mucosal
cells,
the
first
defense
against
infection
(McCullough
et
al.,
1999).
RA
is
 involved
 in
 the
 development
 and
 differentiation
 of
 white
 blood
 cells
 (Semba,
 1998).
 RA
 plays
a
key
role
in
embryonic
development
and
tissue
remodeling
in
the
adult
and
it
plays
 essential
roles
in
maintaining
the
integrity
of
epithelial
cells
of
the
respiratory
tract
(Darlow
 and
Graham,
2007;
Osanai
et
al.,
2010;
Donato
et
al.,
2007).
During
fetal
development,
RA
 is
required
for
limb
development
and
formation
of
the
heart,
eyes
and
ears
(Clagette‐Dame
 and
 Deluca,
 2002).
 In
 addition
 to
 its
 role
 in
 normal
 growth
 and
 development,
 RA
 has
 important
 implications
 in
 cancer
 development
 and
 treatment.
 Vitamin
 A
 deficiency
 is
 thought
 to
 increase
 the
 risk
 of
 some
 cancers,
 and
 RA
 is
 administered
 therapeutically
 in
 cancer
patients
(Osanai
et
al.,
2010).
 
 
  RA
exerts
its
effects
on
growth
and
development
by
regulating
gene
expression.
To
  induce
 gene
 transcription,
 RA
 binds
 to
 two
 families
 of
 nuclear
 receptors:
 the
 RARs
 and
 RXRs.
The
metabolites
all‐trans‐RA
and
9‐cis‐RA
are
high
affinity
ligands
for
RARs,
whereas
 only
9‐cis‐RA
has
been
shown
to
bind
to
RXRs
(Soprano
and
Soprano,
2002).



RARs
and
 RXRs
 are
 two
 families
 of
 zinc‐finger
 nuclear
 receptors
 (Mark
 et
 al.,
 2006).
 
 Each
 family
  
  4
  includes
 three
 members:
 the
 RAR‐α,
 ‐β,
 and
 –γ
 and
 RXR‐α,
 ‐β,
 and
 ‐γ.
 
 RXRs
 form
 homodimers
and
bind
to
the
retinoid
X
response
elements
(RXREs)
within
gene
promoters,
 whereas
 RARs
 form
 heterodimers
 with
 RXRs
 and
 bind
 to
 RAREs
 in
 order
 to
 induce
 gene
 transcription
(Mark
et
al.,
2006).

In
the
absence
of
RA,
RXR/RXR
heterodimer
complexes
 bind
 to
 the
 RARE
 as
 well
 as
 a
 corepressor
 complex,
 including
 those
 with
 histone
 deacetylase
activity.
Activation
of
transcription
occurs
when
RA
binds
to
RARs,
altering
the
 RAR/RXR
 interactions
 with
 corepressor
 proteins
 while
 increasing
 binding
 affinity
 to
 coactivator
 proteins,
 including
 those
 with
 histone
 acetyltransferase
 activity
 (Tang
 and
 Gudas,
2011;
Figure
1.3).


 
 Being
fat
soluble,
vitamin
A
requires
numerous
proteins
for
its
absorption,
transport
 through
the
blood,
intracellular
transport,
and
metabolism
(Blomhoff
and
Blomhoff,
2006).
 Transcriptional
activities
of
RA
are
mediated
by
a
small
soluble
protein
termed
cellular
RA‐ binding
protein
II
(CRABPII;
Donato
et
al.,
2007).
CRABPII
shuttles
RA
from
the
cytoplasm
 into
the
nucleus
and
facilitates
the
binding
of
RA
to
RAR
(Sessler
and
Noy,
2005).

Further,
 CRABPII
 has
 been
 shown
 to
 play
 a
 critical
 role
 in
 sensitizing
 tumors
 to
 the
 growth
 suppressive
effects
of
RA
(Manor
et
al.,
2003).



 
 A
large
number
of
genes
are
proven
or
putative
direct
targets
of
RA
and
numerous
 other
genes
are
regulated
by
an
indirect
manner
(Ross
et
al.,
2011).

A
PCR
array
study
in
 mouse
 liver
 shows
 that
 genes
 affected
 by
 RA
 include
 those
 involved
 in
 angiogenesis,
 cell
 differentiation,
 cell
 proliferation,
 cell
 migration
 and
 adhesion,
 hypoxic
 adaptation
 and
  
  5
  apoptosis
 (Mamoon
 et
 al.,
 2008).
 
 
 Many
 of
 these
 genes,
 including
 those
 involved
 in
 cell
 proliferation,
 differentiation
 and
 apoptosis,
 are
 induced
 after
 24
 h
 of
 RA
 treatment,
 indicating
that
they
are
indirect
targets
of
RA
signaling.
The
expression
of
nuclear
receptor,
 RARβ,
 is
 induced
 more
 than
 three
 fold
 after
 only
 three
 hours,
 indicating
 a
 more
 direct
 response
to
RA.

 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
  
  6
  
 
 
 
  
 
 Figure
1.1:

Structure
of
all‐trans
retinol,
all‐trans
retinal,
and
all‐trans
retinoic
acid.

 
 
 
 
 
  
  7
  
 
  
 Figure
 1.2:
 
 Mechanism
 of
 retinoic
 acid‐induced
 gene
 transcription.
 
 Abbreviation:
 Pol
 II,
 DNA
polymerase
II.
 
 
 
 
 
  
  8
  1.2.

Zinc
Nutrition
and
Function
 Zinc
is
an
essential
trace
element
for
humans
and
is
required
for
the
catalytic
and
 structural
 role
 of
 more
 than
 300
 enzymes,
 which
 control
 processes
 including
 DNA
 synthesis,
 normal
 growth,
 brain
 development,
 bone
 formation
 and
 wound
 healing
 (Sandstead,
2003;
Stephanidou
et
al.,
2006).
For
example,
zinc
is
required
for
the
activity
of
 multiple
 enzymes
 required
 for
 DNA
 replication
 and
 transcription
 including
 DNA
 and
 RNA
 polymerases,
 thymidine
 kinase,
 and
 ornithine
 decarboxylase,
 all
 of
 which
 are
 involved
 in
 mitosis
(Mocchegiani
et
al.,
2000).
Zinc
also
provides
structural
stabilization
to
a
multitude
 of
 proteins,
 including
 zinc‐finger
 transcription
 factors,
 which
 regulate
 many
 cellular
 processes
 (Satre
 et
 al.,
 2001;
 Rana
 et
 al.,
 2008).
 
 Some
 of
 the
 zinc‐finger
 proteins
 are
 involved
 in
 DNA
 repair,
 apoptosis,
 cell
 cycle
 progression,
 and
 cell
 proliferation
 and
 differentiation.
 Therefore,
 adequate
 zinc
 status
 is
 essential
 for
 normal
 development
 and
 maintenance
of
the
human
body.

 
 Zinc
 is
 found
 mainly
 in
 foods
 of
 animal
 origin,
 but
 is
 also
 present
 in
 plants
 (Shah,
 2011).
The
recommended
dietary
allowance
(RDA)
for
zinc
is
11
mg
for
men
and
9
mg
for
 women
(IOM,
2000).
Rich
dietary
sources
of
zinc
include
beef
and
lamb,
which
contain
4.1
 and
 3.3
 mg/100g
 tissue,
 respectively
 (McAfee
 et
 al.,
 2010).
 Some
 seafood,
 especially
 oysters,
and
whole
grains
are
also
good
sources
of
zinc
(Simpson
et
al,
2011).
However,
the
 bioavailability
 of
 zinc
 varies
 widely
 (Simpson
 et
 al,
 2011).
 Bioavailability
 from
 unrefined
 cereal
 grains
 and
 legumes
 is
 low
 due
 to
 the
 inhibition
 of
 absorption
 by
 phytate
 (inositol
 hexaphosphate).
 The
 phosphate
 groups
 in
 phytates
 form
 strong
 bonds
 with
 divalent
  
  9
  cations
 including
 zinc
 and
 because
 the
 human
 gastrointestinal
 tract
 lacks
 significant
 phytase
 activity,
 zinc
 bound
 to
 phytate
 is
 not
 available
 for
 absorption
 (Hess
 and
 Brown,
 2009).

 
 While
 severe
 zinc
 deficiency
 is
 rare
 in
 human
 populations,
 mild
 to
 moderate
 deficiency
 is
 extremely
 prevalent,
 with
 about
 two
 billion
 people
 worldwide
 ingesting
 inadequate
amounts
of
zinc
(Brown
et
al.,
2001;
Prasad,
2003;
Song
et
al.,
2009).
Because
 of
its
wide‐ranging
functions,
inadequate
zinc
intake
may
have
severe
consequences.
The
 classical
 zinc
 deficiency
 symptoms
 include
 impaired
 growth,
 wound
 healing,
 and
 reproductive
 function
 in
 men,
 as
 well
 as
 increased
 severity
 of
 a
 variety
 of
 infections
 (Bohnsack
and
Hirschi,
2004;
Hess
et
al.,
2009).
Zinc
deficiency
also
impairs
developmental
 neurogenesis
 and
 evidence
 suggests
 that
 zinc
 deficiency
 may
 also
 impair
 neuronal
 differentiation
(Levenson
and
Morris,
2011;
Corniola
et
al.,
2008).
The
deficiency
symptoms
 of
 zinc
 are
 believed
 to
 occur
 as
 a
 result
 of
 the
 role
 of
 zinc
 in
 enzymes
 and
 transcription
 factors
(Levenson
and
Morris,
2011).
In
addition
to
impaired
growth
and
development,
it
 has
 been
 proposed
 that
 zinc
 deficiency
 could
 increase
 the
 risk
 of
 some
 types
 of
 cancers,
 due
to
the
important
role
that
zinc
plays
in
the
protection
from
oxidation
and
damage
to
 DNA
(Ho,
2004).
 
 Within
 the
 cell,
 40%
 of
 zinc
 is
 located
 in
 the
 nucleus,
 50%
 in
 the
 cytoplasm,
 organelles
and
specialized
vesicles
and
the
remainder
in
the
cell
membrane
(Stephanidou
 et
al.,
2006).
While
90%
of
the
total
cellular
zinc
is
tightly
bound
to
proteins,
the
remaining
  
  10
  more
 dynamic
 pool
 of
 zinc
 is
 known
 as
 the
 labile
 intracellular
 pool
 of
 zinc
 (LIPZ).
 
 LIPZ
 consists
of
free
ionic
zinc
within
the
fM
‐
pM
range
and
zinc
bound
to
low
molecular
weight
 ligands
 such
 as
 histidine,
 cysteine,
 aspartate,
 glutamate,
 citrate,
 and
 metallothionein
 (Franklin
and
Costello,
2009).
The
LIPZ
provides
metabolically
 available
zinc
for
processes
 such
 as
 cellular
 signaling,
 second
 messenger
 metabolism,
 and
 the
 function
 of
 enzymes
 (Kambe
et
al.,
2004).

 
 At
 the
 cellular
 and
 molecular
 levels,
 either
 zinc
 deficiency
 or
 excess
 may
 have
 detrimental
 and
 even
 cytotoxic
 effects,
 therefore
 zinc
 homeostasis
 must
 be
 tightly
 maintained
 to
 ensure
 normal
 cellular
 functions
 (Kambe
 et
 al.,
 2004).
 Zinc
 homeostasis
 is
 achieved
 through
 a
 combination
 of
 zinc
 importers,
 zinc
 exporters,
 and
 metallothionein.
 Two
families
of
zinc
transporters
are
the
ZRT,
IRT‐like
protein
(ZIP)
family,
which
consists
of
 14
members
and
controls
the
influx
of
zinc
into
the
cytosol,
and
the
zinc
transporter
(ZnT)
 family,
 which
 consists
 of
 10
 members
 and
 is
 responsible
 for
 transport
 zinc
 out
 of
 the
 cytosol
 (Cousins
 et
 al.,
 2006).
 Zinc
 homeostasis
 is
 also
 maintained
 by
 a
 metal‐binding
 protein
 called
 thionein,
 which
 binds
 to
 zinc
 to
 form
 metallothionein.
 The
 synthesis
 of
 thionein
can
be
induced
by
zinc
and
metallothionein
acts
as
a
reservoir
and
buffer
of
labile
 zinc.
 
 1.2.1.

Zinc‐Finger
Proteins
 Numerous
proteins
contain
loop
structures
stabilized
by
zinc
ions.

This
type
of
loop
 is
 known
 as
 zinc‐finger
 and
 the
 protein
 is
 known
 as
 a
 zinc‐finger
 protein.
 
 Zinc‐finger
  
  11
  proteins
are
involved
in
a
variety
of
physiological
functions,
including
hormone
secretion,
 immune
 defense,
 and
 DNA
 repair
 (Ho
 and
 Ames,
 2002;
 Dieck
 et
 al.,
 2003;
 Song
 et
 al.,
 2009).

Between
3‐10%
of
human
genes
are
thought
to
encode
zinc‐binding
proteins
and
 nearly
half
of
eukaryotic
transcription
factors
bind
zinc
(Loh,
2010).

 
 The
 presence
 of
 zinc
 in
 zinc‐fingers
 is
 essential
 to
 their
 function.
 Zinc‐fingers
 are
 critical
to
proper
folding
of
the
proteins
they
are
found
in
(Klug
and
Schwabe,
1995).
Zinc‐ fingers
 are
 essential
 for
 the
 DNA‐binding
 activities
 of
 the
 transcription
 factors,
 as
 their
 unique
 structure
 allows
 for
 recognition
 and
 binding
 of
 specific
 sequences
 in
 double
 stranded
DNA
(Park
et
al.,
2011).

In
addition
to
binding
to
DNA,
zinc‐fingers
have
also
been
 shown
to
mediate
protein‐protein
interactions
(Gamsjaeger
et
al.,
2006).

 
 1.3.
Interactions
Between
Zinc
and
Vitamin
A
 
 There
 has
 been
 much
 evidence
 of
 an
 interaction
 between
 zinc
 and
 vitamin
 A
 (Christian
 and
 West,
 1998).
 In
 rats,
 zinc
 deficiency
 has
 been
 shown
 to
 impair
 the
 mobilization
 of
 retinol
 from
 the
 liver,
 whereas
 zinc
 repletion
 restores
 plasma
 vitamin
 A
 levels
 (Brown
 et
 al.,
 1976;
 Duncan
 and
 Hurley,
 1978).
 
 In
 human
 studies,
 zinc
 deficiency
 limits
the
ability
of
cirrhotic
patients
to
respond
to
vitamin
A
and
combined
vitamin
A
and
 zinc
deficiencies
are
often
found
in
these
patients
(Russel,
1980).

In
studies
attempting
to
 restore
night
vision
in
Nepalese
pregnant
women,
when
zinc
supplements
were
given
along
 with
vitamin
A
supplements,
the
women
were
four
times
more
likely
to
have
restored
night
  
  12
  vision
 than
 with
 vitamin
 A
 supplementation
 alone,
 if
 they
 were
 zinc
 deficient
 at
 baseline
 (Christian
et
al.,
2001).

 The
mechanisms
involved
in
zinc‐vitamin
A
interactions
are
presently
unclear.

One
 theory
 for
 the
 interaction
 between
 zinc
 and
 vitamin
 A
 is
 that
 zinc
 deficiency
 impairs
 the
 synthesis
of
retinol
binding
protein
(RBP),
which
in
turn
inhibits
the
release
of
ROL
from
the
 liver
into
the
blood
(Smith,
1980).

However,
in
HepG2
cells,
culture
in
low
zinc
media
has
 been
 shown
 to
 increase
 the
 expression
 of
 RBP
 (Satre
 et
 al.,
 2001).
 
 It
 has
 also
 been
 suggested
that
zinc
deficiency
may
interfere
with
the
metabolism
of
vitamin
A
by
affecting
 the
activity
of
enzymes
involved.

In
rats,
zinc
deficiency
decreases
the
activity
of
alcohol
 dehydrogenase
 (ADH),
 a
 zinc‐dependent
 enzyme
 that
 catalyzes
 the
 conversion
 of
 ROL
 to
 RAL,
whereas
the
activity
of
retinal
oxidase,
an
enzyme
that
converts
RAL
to
RA,
increases
 (Boron
et
al.,
1988).



 
 It
is
clear
that
zinc
is
required
for
vitamin
A
function
at
the
whole‐body
level,
but
in
 vitro
evidence
of
a
zinc‐vitamin
A
interaction
is
sparse,
resulting
in
limited
information
on
a
 zinc‐vitamin
 A
 interaction
 at
 the
 cellular
 level.
 Interestingly,
 both
 zinc
 and
 vitamin
 A
 are
 required
 for
 neuronal
 differentiation
 (Levenson
 and
 Morris,
 2011).
 Recently,
 one
 study
 assessing
the
effects
of
zinc
status
on
RA‐induced
differentiation
of
human
neuronal
cells
 found
 that
 zinc
 deficiency
 impairs
 RA‐induced
 differentiation
 in‐vitro
 in
 a
 zinc
 concentration‐dependent
manner
(0.4
–
2.5
µM
zinc),
but
the
reasons
for
this
dependence
 are
unknown
(Gower‐Winters,
2008).

 
  
  13
  1.4.
The
Role
of
Vitamin
A
in
Cancer
 Vitamin
 A
 also
 plays
 key
 roles
 in
 the
 development,
 progression,
 prevention,
 and
 treatment
of
cancer.
Early
studies
on
animals
have
shown
an
association
between
vitamin
 A
 deficiency
 and
 carcinogenesis
 when
 it
 was
 discovered
 that
 vitamin
 A
 is
 required
 for
 normal
epithelial
cell
morphology
in
rats
(Walbach
and
Howe,
1925).
More
recently,
it
has
 been
shown
that
vitamin
A
treatment
can
reduce
the
incidence
of
second
primary
tumors
 in
 patients
 with
 prior
 lung,
 head
 and
 neck,
 and
 liver
 cancer
 (Dragnev,
 et
 al.,
 2000).
 Epidemiological
studies
show
an
association
between
vitamin
A
and
risk
of
some
cancers.
 For
 example,
 high
 intake
 or
 high
 circulating
 vitamin
 A
 (as
 ROL)
 levels
 correlates
 with
 a
 decreased
 risk
 of
 developing
 breast
 cancer,
 but
 an
 increased
 risk
 of
 prostate
 and
 gastric
 cancer
 (Fulan
 et
 al.,
 2011;
 Miyazaki
 et
 al.,
 2011;
 Mondul
 et
 al.,
 2011)
 In
 addition,
 RA
 displays
 distinct
 anti‐carcinogenic
 activities
 and
 is
 currently
 used
 in
 treatment
 or
 is
 being
 tested
in
clinical
trials
as
a
preventative
and
therapeutic
agent
in
several
types
of
cancer,
 including
 breast
 cancer,
 liver
 cancer,
 prostate
 cancer,
 non‐small‐cell
 lung
 cancer,
 thyroid
 cancer,
and
myeloid
leukemias
(Bohnsack
and
Hirschi,
2004,
Nakanishi
et
al.,
2008;
Donato
 et
al.,
2007).

 
 RA
can
exert
its
effects
by
directly
or
indirectly
influencing
the
expression
of
genes
 involved
 in
 cellular
 growth
 (Alisi
 et
 al.,
 2003).
 
 Most
 often,
 treatment
 with
 RA
 causes
 inhibition
 of
 cell
 growth
 in
 many
 RA‐sensitive
 tumor
 cells,
 with
 signaling
 through
 RARα
 (Fitzgerald
 et
 al.,
 1997).
 
 In
 these
 cases,
 transcriptional
 activation
 of
 RAR
 may
 trigger
 differentiation,
apoptosis,
and
cell‐cycle
arrest,
depending
on
the
type
of
cancer
(Schug
et
  
  14
  al.,
 2007).
 
 For
 example,
 H157
 human
 squamous
 cell
 carcinoma
 cells
 respond
 to
 RA
 treatment
by
inhibition
of
cell
proliferation
(Sun
et
al.,
2000).

In
MCF7
breast
cancer
cells
 and
other
estrogen
receptor‐positive
breast
cancer
cell
lines,
RA
treatment
inhibits
growth
 and
induces
apoptosis
(Liu
et
al.,
1997;
Estner
et
al.,
1998).

 
 Numerous
proteins
and
signaling
cascades
are
thought
to
be
involved
in
the
effect
 of
 RA
 on
 growth.
 
 For
 example,
 in
 HepG2
 cells,
 treatment
with
 RA
 induces
 growth
 arrest
 and
differentiation,
which
is
thought
to
be
due
to
the
influence
of
RA
on
the
activities
of
 cyclin‐CDK
complexes
involved
in
the
regulation
of
G1/S
transition
and
S‐phase
progression
 (Alisi
 et
 al.,
 2003).
 RA
 has
 been
 shown
 to
 cause
 growth
 inhibition
 in
 liver
 and
 pancreatic
 cancer
 cells
 mediated
 through
 transforming
 growth
 factor‐ß
 (TGF‐ß;
 Salbert
 et
 al.,
 1993;
 Singh
 et
 al.,
 2007).
 
 The
 mitogen‐activated
 protein
 (MAP)
 kinase
 pathway
 can
 also
 be
 activated
 by
 RA
 in
 NB‐4
 acute
 promyelocytic
 leukemia
 and
 MCF‐7
 breast
 carcinoma
 cell
 lines
 (Alsayed
 et
 al.,
 2001).
 Also,
 In
 NB‐4
 cells,
 RA‐induced
 apoptosis
 is
 thought
 to
 be
 mediated
 by
 the
 membrane‐bound
 tumor‐selective
 death
 ligand,
 TRAIL
 (Altucci
 et
 al.,
 2001).


 
 All‐trans
 RA,
 or
 tretinoin,
 is
 approved
 to
 induce
 cytodifferentiation
 and
 decrease
 proliferation
of
acute
promyelocytic
leukemia
(APL;
Thatcher
and
Isoherranen,
2009).
RA
is
 also
utilized
in
the
treatment
of
head
and
neck
carcinoma,
and
non‐small‐cell
lung
cancer
 (Bohnsack
 and
 Hirschi,
 2004)
 and
 in
 combination
 with
 other
 drugs
 in
 the
 treatment
 of
 some
 solid
 tumors.
 However,
 clinical
 applications
 of
 RA
 in
 these
 cases
 can
 show
 limited
  
  15
  effects
 due
 to
 RA
 resistance
 (Arce
 et
 al.,
 2005;
 Hua
 et
 al.,
 2009).
 RA
 treatment
 in
 APL
 induces
complete
remission
in
about
90%
of
patients,
however
this
remission
is
short‐lived
 due
to
the
rapid
emergence
of
resistance
(Tallman
et
al.,
2002).
 
 Another
 concern
 regarding
 the
 use
 of
 RA
 as
 a
 cancer
 treatment
 is
 the
 potential
 toxicity.
 
 RA
 treatment
 is
 generally
 well
 tolerated,
 however,
 in
 about
 2‐27%
 of
 patients,
 adverse
 complications
 may
 result.
 The
 set
 of
 symptoms,
 termed
 retinoic
 acid
 syndrome
 (RAS),
 include
 fever,
 weight
 gain,
 elevated
 white
 blood
 cells,
 respiratory
 distress,
 hypotension
 and
 acute
 renal
 failure
 (Patatanian
 and
 Thompson,
 2008).
 In
 addition,
 dose‐ related
central
nervous
system
toxicity
can
develop,
especially
in
the
pediatric
population.
 This
toxicity,
termed
pseudotumor
cerebri
(PTC),
is
characterized
by
neurologic
and
ocular
 symptoms,
 as
 well
 as
 increased
 intracranial
 pressure
 (Vanier
 et
 al,
 2003).
 
 It
 has
 been
 reported
that
death
from
RAS
occurs
in
about
2%
of
patients
treated
with
RA
(Larson
and
 Tallman,
2003).
 
 Though
it
is
clear
that
RA
plays
an
important
role
in
gene
expression
and
regulation
 of
cancer
cell
growth,
very
little
is
known
about
the
downstream
pathways
of
RA‐mediated
 signaling.
 The
 mechanisms
 whereby
 RA
 regulates
 biological
 processes
 remain
 to
 be
 fully
 understood
 (Thatcher
 and
 Isoherranen,
 2009).
 Thus,
 investigations
 into
 the
 genetic
 mechanisms
 and
 signaling
 pathways
 behind
 the
 growth
 effects
 of
 RA
 in
 many
 cancer
 cell
 lines
are
subjects
of
ongoing
research
(Nakanishi
et
al.,
2008).
Information
on
direct
target
 genes
involved
in
the
anti‐proliferative
activities
of
RA
is
scarce
and
little
is
known
about
  
  16
  the
involvement
of
different
isoforms
of
the
RARs
and
RXRs
(Donato
et
al.,
2007;
Hua
et
al.,
 2009).

 
 1.4.1.
Effect
of
Retinoic
Acid
on
the
Growth
of
HepG2
 Hepatocellular
carcinomas
(liver
cancers)
are
a
relatively
common
cancer,
being
the
 fifth
most
common
but
the
third
leading
cause
of
cancer
mortality
worldwide
(Altekruse
et
 al.,
 2009;
 Alison
 et
 al.,
 2011).
 RA
 has
 been
 shown
 to
 decrease
 growth
 of
 hepatocellular
 carcinomas
through
decreasing
cell
proliferation
and
induction
of
differentiation.
Falsca
et
 al.
(1999)
observed
that
treatment
of
HepG2
(a
hepatocellular
carcinoma
cell
line)
with
5
 µM
of
RA
for
12
days
resulted
in
80%
growth
inhibition
as
well
as
progression
to
a
more
 differentiated
 phenotype.
 Alisi
 et
 al.
 (2003)
 found
 in
 HepG2
 that
 RA
 regulates
 proteins
 involved
 in
 G1/S
 transition
 in
 the
 cell
 cycle,
 particularly
 by
 altering
 the
 binding
 of
 cyclin‐ CDK
 complexes
 to
 p21
 and
 p27
 cell
 cycle
 inhibitors.
 
 Further,
 Nakanishi
 et
 al.
 (2008)
 identified
 entire
 regulatory
 cascades
 induced
 by
 RA
 involved
 in
 growth
 arrest
 in
 HepG2
 cells,
 including
 the
 MAP
 kinase
 pathway,
 which
 is
 suspected
 to
 be
 directly
 regulated
 by
 RARβ
and
RARα.
 
 Although
treatment
of
HepG2
with
RA
results
in
decreased
growth,
it
appears
to
be
 relatively
less
sensitive
than
other
cell
lines,
requiring
longer
treatment
duration
or
higher
 concentrations
of
RA.
For
example,
Nakanishi
et
al.
(2008)
observed
that
concentrations
of
 less
than
50
µM
of
RA
were
not
effective
in
decreasing
cell
number
in
HepG2
within
72
h
of
 treatment.
 
 Arce
 et
 al.
 (2005)
 found
 HepG2
 cells
 to
 be
 much
 more
 resistant
 to
 RA
  
  17
  treatment
than
Hep3B
cells,
another
hepatocarcinoma
cell
line.
They
found
a
50%
decrease
 in
 viability
 in
 Hep3B
 following
 72
 h
 of
 RA
 treatment
 at
 25
 µM
 whereas
 in
 HepG2,
 no
 reduction
in
viability
was
seen
at
this
time
point
until
treatment
reached
166
µM.

Alisi
et
 al.
(2003)
observed
that
a
treatment
of
5
µM
RA
for
2
weeks
results
in
growth
inhibition
 and
 a
 more
 differentiated
 phenotype.
 As
 a
 comparison,
 a
 reduction
 in
 growth
 was
 observed
at
RA
treatment
concentrations
as
low
as
0.001
µM
 in
MCF‐7
and
T‐47D
breast
 cancer
cell
lines
after
7
days
(Liu
et
al.,
1997).

 
 1.4.2.
Role
of
Retinoic
Acid
Receptor
β in
Cancer
 Although
 the
 exact
 genes
 involved
 in
 RA‐induced
 growth
 inhibition
 are
 unclear,
 there
 is
 much
 evidence
 that
 RARß,
 a
 member
 of
 the
 retinoic
 acid
 receptor
 family,
 is
 important
for
this
process.
In
several
in
vitro
systems,
the
retinoid
anti‐proliferative
effect
 requires
 RARß
 and
 the
 inducibility
 of
 RARß
 predicts
 responsiveness
 to
 RA
 (Altucci
 et
 al.,
 2001;
 Fabricius
 et
 al.,
 2011).
 Sun
 et
 al.
 (2000)
 demonstrated
 that
 if
 RARß
 expression
 is
 blocked,
cell
responsiveness
to
RA
is
reduced.
Because
RARß
is
RA‐inducible,
it
is
a
classical
 direct
target
of
RA
(Ross
et
al.,
2011).
Strong
evidence
suggests
that
RARß
acts
as
a
tumor
 suppressor
 as
 it
 is
 frequently
 lost
 or
 epigenetically
 silenced
 in
 various
 cancers
 and
 its
 expression
correlates
inversely
with
tumor
grade
(Ross
et
al.,
2011).

 
 1.4.3.
CYP26a1
in
Cancer
 Cellular
exposure
to
RA
is
regulated
by
controlled
synthesis
and
metabolism
by
RA
 metabolizing
enzymes,
including
CYP26a1.
CYP26a1
is
a
member
of
the
cytochrome
p450
  
  18
  superfamily
 of
 enzymes
 and
 is
 responsible
 for
 metabolizing
 RA
 to
 4‐OH
 RA,
 its
 primary
 oxidation
 product
 (Thatcher
 et
 al.,
 2010).
 It
 is
 thought
 that
 resistance
 due
 to
 rapid
 clearance
of
RA
after
continuous
oral
administration
is
due
to
the
induction
of
CYP26a1
in
 tissues
such
as
the
liver
(Thatcher
and
Isoherranen,
2009).
Ozpolat
et
al.
(2005)
observed
 that
 RA
 treatment
 induced
 mRNA
 expression
 of
 CYP26a1
 in
 human
 intestinal
 (Caco‐2),
 endothelial
(HUVEC),
APL
(NB‐4),
and
liver
(HepG2)
cell
lines
in
a
dose‐dependent
manner.
 This
theory
is
supported
by
the
observation
that
overexpression
of
CYP26a1
in
various
cell
 lines
leads
to
increased
resistance
to
RA
treatment
(Osanai
and
Petkovich,
2005).

 
 1.5.

Role
of
Zinc
in
Cancer
 
  Zinc
 appears
 to
 play
 an
 important
 role
 in
 the
 development
 and
 progression
 of
  cancer,
but
 a
 common
 relationship
 of
 zinc
 with
 cancer
 development
 and
 progression
 has
 not
yet
been
identified
(Franklin
and
Costello,
2009).
It
has
been
suggested
that
low
dietary
 zinc
 intakes
 may
 be
 a
 risk
 factor
 for
 cancer
 development.
 
 For
 example,
 in
 humans,
 zinc
 deficiency
 is
 associated
 with
 an
 increased
 risk
 of
 developing
 esophageal
 squamous
 cell
 carcinoma
(Abnet
et
al.,
2005).

The
exact
mechanisms
by
which
zinc
deficiency
increases
 the
 risk
 of
 cancer
 are
 still
 unclear
 (Song
 et
 al.,
 2009).
 
 It
 has
 been
 proposed
 that
 zinc
 deficiency
may
increase
the
risk
of
cancer
by
increasing
oxidative
stress
and
impairing
DNA
 repair
mechanisms
(Ho,
2004).
This
has
been
demonstrated
by
the
observations
that
zinc
 deficiency
 results
 in
 an
 increased
 sensitivity
 to
 oxidative
 stress
 and
 causes
 oxidative
 DNA
 damage
 (Taylor
 et
 al.,
 1988;
 Oteiza
 et
 al.,
 2000),
 and
 cells
 grown
 in
 zinc
 deficient
 media
 show
 increased
 oxidant
 production
 (Ho
 and
 Ames,
 2002).
 
 Zinc
 also
 plays
 a
 role
 in
 the
  
  19
  regulation
 of
 DNA
 replication
 and
 transcription
 through
 zinc‐finger
 proteins
 (Ho,
 2004).
 These
proteins
may
be
impaired
by
zinc
deficiency.
For
example,
p53,
a
zinc‐finger
finger
 protein
 that
 has
 been
 shown
 to
 be
 impaired
 by
 inadequate
 zinc,
 modulates
 cell
 cycle
 progression,
apoptosis,
DNA
repair,
proliferation
and
differentiation
(Ho,
2004).

 
 Further,
it
has
been
observed
that
intracellular
zinc
levels
are
altered
in
many
types
of
 cancer.
For
example,
reports
have
shown
that
zinc
levels
are
markedly
increased
in
breast
 cancer
 tissue
 as
 compared
 with
 non‐cancerous
 tissue
 (Rizk
 and
 Sky‐Peck,
 1984).
 In
 pancreatic
cancer
cells,
expression
of
the
ZIP4
zinc
importer
is
up‐regulated
in
comparison
 to
 non‐cancerous
 tissue
 resulting
 in
 an
 increased
 zinc
 levels
 in
 tumor
 cells
 and
 tumor
 growth
 (Logsdon
 et
 al.,
 2003).
 Forced
 up‐regulation
 of
 this
 transporter
 in
 cell
 culture
 studies
 results
 in
 a
 corresponding
 increase
 in
 intracellular
 zinc
 and
 increased
 cell
 proliferation
in
pancreatic
cancer
cells
(Li
et
al.,
2007).
In
contrast,
cellular
zinc
levels
are
 reduced
in
prostate
cancer
and
liver
cancer.
Normal
prostate
cells
have
high
levels
of
zinc,
 however,
malignant
cells
have
lost
the
ability
to
accumulate
zinc
due
to
a
down
regulation
 of
 hZIP1,
 a
 major
 zinc
 uptake
 transporter
 in
 prostate
 cells
 (Costello
 and
 Franklin,
 2006;
 Franklin
et
al.,
2007).

 
 1.5.1.

Zinc
in
Hepatocarcinoma
 Several
 reports
 indicate
 that
 intracellular
 zinc
 is
 significantly
 lower
 in
 hepatocellular
 cancer
 tissue
 compared
 to
 normal
 hepatic
 cells
 (Gurusami
 and
 Davidson,
 2007).
 For
 example,
 Ebara
 et
 al.
 (2000)
 observed
 55%
 lower
 zinc
 levels
 in
 hepatomas
 compared
 to
  
  20
  surrounding
 normal
 liver
 tissue
 in
 sixteen
 patients.
 
 Tashiro
 et
 al.
 (2003)
 observed
 that
 cancerous
liver
tissue
contains
less
than
half
of
the
zinc
in
non‐cancerous
liver
tissue.
The
 mechanisms
and
functional
consequences
of
this
decrease
in
zinc
status
in
liver
cancer
are
 not
 clear.
 However,
 one
 possible
 explanation
 for
 the
 decreased
 zinc
 in
 cancerous
 liver
 tissue
is
decreased
uptake
of
zinc
due
to
the
absence
of
the
zinc
importer,
ZIP14.
Protein
 expression
of
ZIP14
is
not
detected
in
HepG2
cells
and
microarray
studies
show
that
mRNA
 expression
of
this
transporter
is
down‐regulated
(Liu
et
al.,
2007;
Franklin
et
al.,
2007).

 
 1.6.

Molecular
Effects
of
Zinc
Deficiency
 Zinc
deficiency
has
been
demonstrated
to
cause
changes
in
gene
expression.

PCR
 array
 analyses
 indicate
 that
 alterations
 in
 cellular
 zinc
 status
 affect
 hundreds
 of
 target
 genes,
 in
 a
 tissue
 and
 cell‐type
 specific
 manner
 (Cousins
 et
 al.,
 2003;
 Kindermann
 et
 al.,
 2004;
Haase
et
al.,
2007).
Low
zinc
status
has
been
shown
to
affect
gene
transcription
in
 the
liver,
including
gene
products
that
participate
in
growth
and
lipid
metabolism
(Dieck
et
 al.,
2003).

For
example,
in
rat
liver,
zinc
deficiency
results
in
the
up‐regulation
of
insulin‐ like
growth
factor
I
(IGFI)
and
fatty
acid
binding
protein
(FABP)
mRNA,
and
down‐regulation
 of
Ras‐related
protein,
among
many
others
(Dieck
et
al.,
2003).
Wong
et
al.
(2007)
found
 that
growth
of
HepG2
cells
in
medium
depleted
of
zinc
resulted
in
a
40%
decrease
in
p21
 protein
 expression
 and
 a
 70%
 decrease
 in
 p21
 mRNA.
 Reaves
 et
 al.
 (2000)
 reported
 that
 growth
 of
 HepG2
 cells
 for
 6.5
 days
 in
 medium
 depleted
 of
 zinc
 resulted
 in
 a
 two
 fold
 increase
 in
 p53
 mRNA
 compared
 to
 cells
 grown
 in
 media
 with
 adequate
 levels
 of
 zinc
 (4µM).

  
  21
  These
 observations
 demonstrate
 the
 importance
 of
 zinc
 in
 cellular
 processes
 including
 growth
 regulation;
 however,
 the
 mechanisms
 behind
 the
 effects
 of
 zinc
 deficiency
on
gene
expression
remain
elusive.

It
has
been
suggested
that
the
changes
may
 be
partially
due
to
the
role
of
zinc
as
an
essential
component
in
many
transcription
factors
 by
stabilizing
zinc‐fingers
(Dieck
et
al.,
2003).
It
is
also
thought
that
zinc
may
affect
mRNA
 stability.
For
example,
it
was
observed
that
Zinc
Transporter
5
(ZnT5)
mRNA
is
stabilized
by
 increased
zinc
availability
in
the
human
intestinal
Caco‐2
cell
line
(Jackson
et
al.,
2008).
 
 The
 importance
 of
 zinc
 for
 the
 structure
 and
 function
 of
 zinc
 finger
 proteins
 is
 demonstrated
 by
 the
 observation
 that
 zinc
 deficiency
 can
 impair
 the
 function
 of
 zinc
 fingers.
For
example,
when
wildtype
p53,
a
zinc‐finger
protein,
is
exposed
to
zinc
chelators,
 it
adopts
a
mutant
conformation
and
its
ability
to
bind
DNA
is
reduced,
whereas
addition
of
 extracellular
 zinc
 at
 concentration
 within
 the
 physiological
 range
 results
 in
 renaturation
 and
reactivation
of
the
wild‐type
protein
(Hainaut
and
Milner,
1993).

Ho
and
Ames
(2002)
 cultured
rat
gliomal
C6
cells
in
zinc
deficient
or
zinc
adequate
medium
for
five
days.
Using
 an
 electromobility
 shift
 assay
 (EMSA)
 they
 found
 that
 the
 DNA
 binding
 abilities
 of
 p53,
 nuclear
 factor
 κβ
 (NFκβ)
 and
 activator
 protein
 1
 (AP‐1),
 all
 of
 which
 are
 zinc
 finger
 proteins,
were
impaired
in
the
cells
grown
in
zinc‐deficient
media.


 
 The
 nuclear
 receptors,
 RARs
 and
 RXRs
 are
 also
 zinc‐finger
 proteins.
 These
 nuclear
 receptors
have
several
conserved
regions
in
their
amino
acid
sequence,
one
being
the
DNA
 ‐binding
domain.
This
domain
contains
two
zinc‐finger
motifs,
critical
to
the
binding
of
the
  
  22
  receptors
 to
 the
 RARE
 (Soprano
 and
 Soprano,
 2002).
 
 Currently,
 little
 research
 has
 been
 conducted
 to
 evaluate
 the
 effects
 of
 zinc
 nutritional
 status
 on
 vitamin
 A
 function,
 particularly
 its
 role
 for
 the
 vitamin
 A
 nuclear
 receptors.
 
 I
 am
 unaware
 of
 any
 published
 research
to
date
that
zinc
deficiency
affects
the
binding
efficiency
or
function
of
RARs
and
 RXRs.
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
  
  23
  
 
 
 
 
 Table
1.1:

Examples
of
genes
affected
by
zinc
status
in
the
liver
 Gene
name
(cell
 type)
  P21
(HepG2)
  Zinc
treatment
(method
 of
depletion)
  depletion

 (medium
zinc
removed
 with
Chelex‐100
resin)
 P53
(liver,
HepG2)
 depletion

 (medium
zinc
removed
 with
Chelex
100
resin)
 
 supplementation
(16
µM)
 IGF1
(rat
liver)
 depletion
(rats
fed
zinc‐ deficient
diet)
 FABP
(rat
liver)
 depletion
(rats
fed
zinc‐ deficient
diet)
 Ras‐related
 protein
 depletion
(rats
fed
zinc‐ (rat
liver)
 deficient
diet)
 
  Effect
on
 gene
 expression
 (protein
or
 mRNA)
 decrease
 (protein
and
 mRNA)
 increase
 
 
 decrease
  Reference
  Increase
 (mRNA)
 Increase
 (mRNA)
 decrease
 (mRNA)
  Dieck
et
al.,
2003
  Wong
et
al.,
2007
 Reaves
et
al.,
2000
  Dieck
et
al.,
2003
 Dieck
et
al.,
2003
  
 
 
 
 
  
  24
  1.7.

Summary
 In
summary,
vitamin
A
and
zinc
are
both
essential
nutrients
and
are
important
for
 many
cell
processes,
including
cell
growth
and
development.
Zinc
deficiency
has
effects
on
 gene
expression,
which
is
thought
to
be
partially
due
to
the
role
of
zinc
as
a
constituent
of
 many
 proteins,
 including
 enzymes
 and
 zinc‐finger
 transcription
 factors.
 Vitamin
 A,
 as
 RA,
 exerts
its
function
through
binding
to
zinc‐finger
transcription
factors
in
order
to
regulate
 gene
 expression,
 including
 genes
 involved
 in
 cell
 proliferation
 and
 growth.
 It
 has
 been
 shown
 that
 zinc
 is
 important
 for
 RA
 function
 in
 vivo,
 however,
 evidence
 of
 interaction
 between
the
two
nutrients
at
the
cellular
level
is
limited.
 
 1.8.

Hypothesis
 
  The
hypotheses
of
my
thesis
research
project
were:

 1)  Sufficient
cellular
zinc
level
is
important
for
the
effectiveness
of
RA‐induced
 growth
inhibition
in
hepatocarcinoma
HepG2
cells;
and

  2)  Influence
of
zinc
on
RA‐induced
growth
inhibition
is
through
modulating
the
 expression
 or
 function
 of
 RARs
 and
 RXRs,
 which
 in
 turn
 affects
 the
 expression
of
their
target
genes
such
as
CYP26a1
and
RARβ.
  
 1.9.

Overall
Objectives
and
Specific
Aims
 
  The
 overall
 objective
 of
 my
 thesis
 research
 project
 was
 to
 examine
 the
 effects
 of
  zinc
 on
 RA‐induced
 growth
 inhibition
 in
 hepatocarcinoma
 HepG2
 cells
 and
 the
 possible
 mechanisms
involved.

The
specific
aims
were:
  
  25
  1) To
 assess
 the
 effects
 of
 zinc
 on
 RA‐induced
 inhibition
 of
 cell
 proliferation
 in
 hepatocarcinoma
HepG2
cells.
 2) To
determine
the
effects
of
zinc
on
the
expression
of
the
nuclear
receptors
(RARα,
 RARβ,
RARγ,
RXRα,
RXRβ,
and
RXRγ).
 3) To
 determine
 the
 effects
 of
 zinc
 on
 the
 RA‐induced
 expression
 of
 CYP26a1
 and
 RARβ, downstream
targets
of
RA
signaling.
 
  
  26
  Chapter
2.

Effects
of
Zinc
on
Retinoic
Acid‐Induced
 
Growth
Inhibition
in
Human
Hepatocarcinoma
HepG2
Cells
 
 
 
 2.1.

Introduction

 
 Vitamin
 A
 has
 many
 functions
 within
 the
 human
 body
 including
 well‐understood
 roles
 in
 night
 vision,
 as
 well
 as
 regulating
 growth
 and
 development
 (Mark
 et
 al.,
 2006;
 Soprano
and
Soprano,
2007).
Research
in
more
recent
years
has
uncovered
additional
roles
 of
 vitamin
 A,
 which
 involves
 the
 action
 of
 retinoic
 acid
 (RA),
 a
 bioactive
 metabolite
 of
 vitamin
 A,
 on
 regulating
 gene
 expression
 (Schug
 et
 al.,
 2007).
 The
 circulating
 form
 of
 vitamin
 A,
 retinol,
 is
 converted
 to
 RA
 within
 the
 cell
 and
 then
 RA
 is
 transported
 into
 the
 nucleus
where
it
binds
to
nuclear
receptors
including
RARs.
RARs
form
heterodimers
with
 RXRs
and
this
complex
binds
to
the
retinoic
acid
response
element
(RARE)
within
the
target
 gene
 (Mark
 et
 al.,
 2006).
 Binding
 of
 RA
 causes
 the
 release
 of
 corepressors
 and
 the
 recruitment
of
coactivators
which
causes
a
subsequent
activation
of
transcription
(Soprano
 and
Soprano,
2002).

It
has
been
shown
that
the
expression
of
more
than
500
genes
with
 numerous
physiological
roles
is
regulated
by
RA
(Mamoon
et
al.,
2008).

 
 
  One
 function
 of
 great
 interest
 is
 the
 role
 of
 RA
 in
 the
 inhibition
 of
 cell
 growth
 in
  promyelocytic
 leukemia
 and
 many
 solid
 tumor
 cell
 lines
 including
 breast,
 pancreatic
 and
 liver
 cancers
 (Salbert
 et
 al.,
 1993;
 Liu
 et
 al.,
 1997;
 Singh
 et
 al.,
 2007;
 Thatcher
 and
 Isoherranen,
2009).
In
hepatocarcinoma
HepG2
cells,
RA
treatment
arrests
the
cell
cycle
at
  
  27
  G1/S
transition,
therefore,
slowing
cell
proliferation
and
ultimately
cell
growth
(Nakanishi
 et
al.,
2008).

 
 
  Although
 RA
 is
 successful
 in
 inducing
 remission
 in
 promyelocytic
 leukemia,
 it
 has
  numerous
side
effects
and
prolonged
treatment
results
in
resistance
(Arce
et
al.,
2005;
Hua
 et
al.,
2009).
It
is
believed
that
the
resistance
is
mediated
by
the
induction
of
CYP26a1,
a
 RA‐metabolizing
 enzyme,
 which
 is
 markedly
 induced
 by
 RA
 in
 the
 liver
 (Ozpolat
 et
 al.,
 2005).
 
 In
 addition,
 HepG2
 cells
 are
 less
 sensitive
 to
 RA
 treatment
 than
 other
 hepatocarcinoma
cells
such
as
Hep3B,
requiring
85%
higher
concentration
of
RA
to
induce
 a
decrease
in
cell
viability
(Arce
et
al.,
2005).
A
RA
treatment
of
5
X
10‐6
M
for
two
weeks
is
 required
to
reduce
the
growth
of
HepG2
cells,
whereas
only
1
x
10‐9
 M
of
RA
is
needed
to
 reduce
the
growth
of
MCF7
and
T47D
breast
cancer
cells
(Liu
et
al,
1997;
Alisi
et
al.,
2007).
 Therefore,
 finding
 means
 to
 increase
 the
 sensitivity
 of
 cancerous
 cells
 towards
 RA
 treatment
will
increase
the
effectiveness
of
RA
as
an
anticancer
agent.

 
 
  The
 regulatory
 role
 of
 RA
 in
 cell
 proliferation
 is
 mediated
 via
 gene
 expression
  through
 binding
 to
 RAR
 and
 RXR
 nuclear
 receptors
 (Mark
 et
 al.,
 2006).
 These
 nuclear
 receptors
are
zinc‐finger
proteins
and
zinc
is
an
essential
structural
component
required
for
 their
 binding
 to
 the
 RARE
 (Soprano
 and
 Soprano,
 2002).
 It
 has
 been
 shown
 that
 zinc
 deficiency
 impairs
 the
 DNA‐binding
 ability
 of
 other
 zinc‐finger
 proteins,
 such
 as
 p53
 and
 AP‐1
 (Ho
 and
 Ames,
 2002).
 
 However,
 the
 relationship
 between
 zinc
 status
 and
 the
 structure
and
function
of
RAR
and
RXR
is
presently
not
known.


  
  28
  Based
on
the
available
evidence,
I
hypothesized
that
sufficient
cellular
zinc
level
is
 important
 for
 the
 effectiveness
 of
 RA‐induced
 growth
 inhibition
 in
 hepatocarcinoma
 HepG2
 cells;
 and
 the
 influence
 of
 zinc
 on
 RA‐induced
 growth
 inhibition
 is
 through
 modulating
 the
 expression
 or
 function
 of
 RARs
 and
 RXRs,
 which
 in
 turn
 affects
 the
 expression
 of
 their
 target
 genes
 such
 as
 CYP26a1
 and
 RARβ.
 The
 objectives
 were:
 1)
 To
 assess
the
effects
of
zinc
on
RA‐induced
inhibition
of
cell
proliferation
in
hepatocarcinoma
 HepG2
cells,
2)
To
determine
the
effects
of
zinc
on
the
expression
of
the
nuclear
receptors
 (RARα,
RARβ,
RARγ,
RXRα,
RXRβ,
and
RXRγ),
and
3)
to
determine
the
effects
of
zinc
on
the
 RA‐induced
expression
of
Cyp26a1
and
RARβ,
two
downstream
targets
of
RA
signaling.
 
 2.2.

Materials
and
Methods
 2.2.1.

Cell
Culture
System
 HepG2,
 a
 hepatocarcinoma
 cell
 line
 (ATCC,
 Manassas,
 VA),
 was
 cultured
 in
 Minimum
 Essential
 Medium
 (MEM;
 Gibco,
 Grand
 Island,
 NY)
 containing
 10%
 fetal
 bovine
 serum
 (FBS),
 sodium
 pyruvate
 (110
 mg/L),
 sodium
 bicarbonate
 (1.5
 g/L)
 and
 penicillin/streptomycin
(5,000
U/L).

Cells
were
grown
at
37°C
in
an
atmosphere
containing
 5%
CO2.
 
 2.2.2.

Preparation
of
Chelex
100‐Treated
FBS

 In
the
culture
system
described
above,
FBS
is
the
source
of
zinc
in
the
medium.

To
 remove
zinc,
Chelex‐100
resin
(BioRad,
Hercules,
CA)
was
mixed
with
FBS
at
100
g/L
for
24
 h
 in
 an
 ice
 bath
 at
 4°C
 with
 gentle
 stirring.
 The
 Chelex‐100‐FBS
 mixture
 was
 then
  
  29
  centrifuged
at
1,000
rpm
for
15
min
at
4°C
to
remove
the
majority
of
the
resin
followed
by
 immediate
filtration
using
a
Steritop
TM
filter
(0.22
µM;
Millipore,
Billeria,
MA)
to
remove
 the
 remaining
 Chelex‐100
 resin
 and
 to
 sterilize
 the
 FBS.
 The
 Chelex‐100‐treated
 FBS
 was
 then
stored
at
‐20°C
until
needed.

 
 2.2.3.

Zinc
Manipulation
 
To
 mimic
 low,
 adequate,
 and
 high
 levels
 of
 zinc,
 cells
 were
 grown
 in
 their
 corresponding
media
supplemented
with
10%
Chelex‐100‐treated
FBS
plus
0,
5,
or
10
µM
 of
Zn2+
as
ZnSO4,
respectively.
The
concentration
of
zinc
in
normal
interstitial
fluid
is
2‐5
µM
 (Franklin
 and
 Costello,
 2009)
 and
 the
 level
 of
 zinc
 in
 normal
 medium
 supplemented
 with
 10%
FBS
is
4
µM
(Reaves
et
al.,
2000).
Zinc
concentrations
in
the
media
supplemented
with
 10%
FBS
and
Chelex‐100‐
treated
FBS
was
reported
previously
in
our
lab
to
be
4.7
µM
and
 0.15
µM,
respectively
(Tsukada,
2003).

Chelex‐100
is
also
capable
of
removing
a
number
of
 divalent
 cations.
 
 These
 divalent
 cations
 were
 added
 back
 after
 Chelex‐100
 treatment
 to
 restore
 their
 pre‐Chelex‐100
 treatment
 concentrations
 (Appendix
 1;
 table
 A.1.;
 Tsukada,
 2003).

 
 2.2.4.

Quantification
of
the
Labile
Intracellular
Pool
of
Zinc
(LIPZ)
 HepG2
cells
were
grown
in
10
cm
Petri
dishes
at
an
initial
density
of
300,000
cells
 per
dish.
To
determine
the
duration
required
for
depleting
intracellular
zinc,
the
cells
were
 grown
 in
 MEM
 medium
 supplemented
 with
 10%
 Chelex‐100‐treated
 FBS
 for
 3
 or
 6
 days
 (n=3).
Cells
in
the
control
group
were
grown
in
MEM
medium
supplemented
with
regular
  
  30
  FBS
 for
 6
 days
 (n=3).
 
 To
 determine
 the
 intracellular
 zinc
 levels
 of
 cells
 grown
 in
 low,
 adequate,
 and
 high
 zinc
 concentrations,
 cell
 were
 grown
 for
 6
 d
 in
 MEM
 medium
 supplemented
 with
 10%
 Chelex‐100‐treated
 FBS
 plus
 0,
 5,
 or
 10
 µM
 zinc,
 respectively
 (n=3).

 
 At
the
end
of
each
culture
period,
cells
were
rinsed
once
with
warm
PBS
(37°C)
and
 trypsinized
 with
 2
 ml
 of
 warm
 0.25%
 Trypsin‐EDTA.
 After
 the
 cells
 were
 detached,
 the
 trypsin
was
neutralized
with
an
equal
volume
of
the
respective
media.

The
cell
suspension
 was
 then
 transferred
 to
 a
 15
 mL
 Falcon
 tube,
 centrifuged
 at
 300
 X
 g
 for
 5
 min
 and
 the
 supernatant
aspirated.
The
cell
pellet
was
re‐suspended
in
3
mL
cold
PBS
(4°C)
using
a
26
G
 5/8
 needle
 to
 separate
 clumped
 cells,
 and
 counted
 using
 a
 particle
 counter
 (Z1
 Particle
 Counter,
 Beckman
 Coulter,
 Fullerton,
 CA)
 with
 a
 cut‐off
 point
 set
 at
 8
 µm.
 For
 each
 replicate,
 2.2
 million
 cells
 were
 transferred
 to
 a
 microcentrifuge
 tube
 and
 centrifuged
 at
 300
X
g
for
5
min.
Following
aspiration
of
the
supernatant,
cells
were
re‐suspended
in
400
 µL
Hanks
buffered
saline
solution
(HBSS;
Gibco,
Grand
Island,
NY)
and
transferred
in
180
µL
 aliquots
to
a
black
96‐well
plate
at
1
million
cells
per
well
(2
technical
replicates).
Cells
were
 then
incubated
in
the
dark
with
20
µL
of
Zinquin
working
solution
(Sigma,
St.
Louis,
MO)
to
 obtain
 a
 final
 concentration
 of
 25
 µM
 for
 30
 min
 at
 37°C
 with
 gentle
 shaking
 using
 a
 thermomixer
 (Thermomixer
 R,
 Eppendorf,
 Hauppauger,
 New
 York).
 
 A
 Zinquin
 working
 solution
was
prepared
by
diluting
the
Zinquin
stock
solution
(5
mM
Zinquin
in
DMSO)
with
 HBSS
 to
 a
 concentration
 of
 250
 µM.
 The
 abundance
 of
 the
 LIPZ
 was
 assessed
 by
 determining
 the
 intensity
 of
 the
 Zinquin‐dependent
 fluorescence
 (excitation:
 365
 nm;
  
  31
  emission:
 475
 nm;
 auto‐cutoff
 point:
 455
 nm)
 using
 a
 microplate
 reader
 (Spectra,
 Molecular
Devices,
Sunnyvale,
CA).
The
background,
which
was
the
fluorescence
intensity
 in
 the
 absence
 of
 cells
 (HBSS
 and
 Zinquin
 only),
 was
 subtracted
 from
 the
 fluorescence
 intensity
obtained
from
the
samples.

 
 2.2.5.

Quantification
of
Total
Cellular
Zinc
 HepG2
 cells
 were
 cultured,
 harvested
 and
 counted
 as
 described
 above.
 After
 re‐ suspension
 in
 cold
 PBS,
 cell
 suspension
 (1
 mL)
 was
 transferred
 to
 a
 microcentrifuge
 tube
 followed
 by
 centrifugation
 at
 300
 X
 g
 for
 5
 min
 and
 aspiration
 of
 the
 supernatant.
 The
 pellet
was
dissolved
in
100
µL
of
concentrated
nitric
acid
and
allowed
to
lyse
overnight
at
 room
 temperature.
 After
 complete
 lysis,
 double
 deionized
 water
 was
 added
 to
 a
 final
 volume
 of
 1
 ml.
 
 
 Zinc
 concentration
 was
 determined
 by
 flame
 atomic
 absorption
 spectrophotometry
as
described
by
Thompson
et
al.
(2002).

 
 2.2.6.

Assessment
of
Cell
Viability
and
Proliferation

 To
assess
cell
proliferation
in
cells
treated
with
RA
alone,
HepG2
cells
were
seeded
 into
 96‐well
 plates
 at
 an
 initial
 density
 of
 7,000
 cells
 and
 allowed
 to
 grow
 for
 2
 d.
 
 Cells
 were
then
treated
with
all‐trans
RA
(Sigma
Aldrich,
Oakville,
ON)
at
0
(DMSO
only;
Sigma
 Aldrich,
Oakville,
ON),
15,
25
or
35
µM
(n=6)
for
12
h.

RA
treatments
were
performed
in
 low
light
conditions
and
culture
dishes
containing
RA
were
protected
from
light
in
the
cell
 culture
incubator
by
aluminum
foil.


Cell
proliferation
following
RA
treatment
was
assessed
 using
 the
 5‐Bromo‐2′‐deoxy‐uridine
 (BrdU)
 colorimetric
 assay
 (Roche,
 Indianapolis,
 IN)
  
  32
  according
to
the
manufacturer’s
instructions
using
 a
microplate
reader
(SpectraMax
Plus,
 Molecular
Devices,
Sunnyvale,
CA,
USA)
at
405
nm
with
a
reference
wavelength
of
490
nm.
 BrdU
 is
 incorporated
 into
 newly
 synthesized
 DNA,
 therefore
 only
 cells
 undergoing
 active
 DNA
synthesis
were
labeled
with
BrdU
(Terry
and
White,
2006).


 
 To
assess
cell
proliferation
following
treatment
with
a
combination
of
zinc
and
RA,
 HepG2
cells
were
seeded
into
10
cm
plates
at
an
initial
density
of
300,000
cells.
Cells
were
 allowed
 to
 attach
 overnight.
 Subsequently,
 the
 cells
 were
 cultured
 in
 Chelex‐100‐treated
 medium
supplemented
with
0
(low),
5
(adequate)
or
10
µM
(high)
of
zinc
(ZnSO4;
day
0).
 On
day
4,
cells
were
transferred
into
96‐well
plates
at
a
density
of
7,000
cells
per
well,
and
 on
 day
 6,
 cells
 were
 treated
 with
 RA
 at
 0
 (DMSO
 only)
 or
 35
 mM
 (n=6)
 for
 12
 h.
 Cell
 proliferation
was
assessed
using
a
BrdU
assay
as
described
above.
 
 To
 assess
 cell
 viability
 following
 zinc
 and
 RA
 treatments,
 cells
 were
 seeded,
 cultured,
and
treated
with
zinc
and
RA
as
described
above
with
changes
as
follows.
HepG2
 cells
were
transferred
to
96‐well
plates
after
5
d,
and
on
day
6,
cells
were
treated
with
RA
 at
 0
 (DMSO
 only)
 or
 35
 µM
 (n=12)
 for
 72h.
 Cell
 viability
 was
 measured
 using
 the
 colorimetric
Thiazolyl
Blue
Tetrazolium
Bromide
(MTT)
assay
(Sigma‐Aldrich,
Oakville,
ON)
 according
 to
 the
 manufacture’s
 instruction.
 Briefly,
 at
 the
 end
 of
 the
 treatment
 period,
 medium
 was
 removed,
 and
 100
 µL
 fresh
 medium
 was
 added
 per
 well
 plus
 10
 µL
 MTT
 (5
 mg/mL
 in
 PBS)
 solution.
 The
 plates
 were
 incubated
 at
 37°C
 for
 2
 h.
 After
 removing
 the
 media,
 100
 µL
 2‐Propanol
 was
 added
 per
 well
 followed
 by
 incubation
 for
 15
 min
 at
 37°C
  
  33
  with
 gentle
 shaking
 using
 a
 thermomixer
 The
 plates
 were
 read
 in
 a
 microplate
 reader
 (SpectraMax
 Plus,
 Molecular
 Devices,
 Sunnyvale,
 California)
 at
 560
 nm
 with
 a
 reference
 wavelength
of
650
nm.


 
 2.2.7.

Cell
Cycle
Analysis
 HepG2
 cells
 were
 cultured
 into
 10
 cm
 culture
 dishes
 in
 regular
 MEM
 at
 an
 initial
 seeding
density
of
150,000
cells
and
allowed
to
attach
overnight.

Medium
was
changed
to
 MEM
 supplemented
 with
 Chelex‐100‐treated
 FBS
 plus
 0
 (low),
 5
 (adequate),
 or
 10
 µM
 (high)
of
Zn
as
ZnSO4.
The
medium
was
changed
once
more
after
3
days
and
after
6
days.

 Cells
 were
 treated
 with
 RA
 at
 35
 µM
 for
 0,
 24,
 or
 72
 h.
 
 At
 the
 end
 of
 each
 treatment
 period,
 the
 cells
 were
 analyzed
 for
 cell
 cycle
 using
 flow
 cytometry
 according
 to
 the
 established
protocol
(Terry
and
White,
2006).

The
main
steps
of
the
assay
were
as
follows.




 
 BrdU
 treatment
 and
 cell
 fixation:
 Briefly,
 dishes
 were
 treated
 with
 1
 µM
 BrdU
 (Sigma‐Aldrich,
Oakville,
ON)
for
20
min
at
37°C.
Cells
were
trypsinized
as
described
above
 and
 counted
 using
 a
 particle
 counter.
 The
 cells
 were
fixed
 by
 adding
 a
 solution
 of
 0.8
 ml
 cold
 PBS
 and
 1.2
 ml
 100%
 ethanol
 (‐20°C)
 per
 2
 X
 106
 while
 vortexing
 and
 left
 to
 fix
 overnight
at
4°C
in
the
dark.


 
 Staining
 with
 PI
 and
 BrdU
 antibody
 incubation:
 Briefly,
 fixed
 cells
 were
 incubated
 with
 1.5
 ml
 2N
 HCl/
 2
 X
 106
 cells
 at
 37°C
 for
 20
 min.
 Subsequently,
 the
 cells
 were
 rinsed
 with
6
ml
of
0.1
M
sodium
borate
while
vortexing,
and
then
centrifuged
at
350
X
g
for
4
  
  34
  min
at
20°C.
The
supernatant
was
aspirated
and
6
ml
of
cold
PBTB
(4°C;
PBS
+
0.5%
Tween‐ 20
+0.5%
BSA)
was
added
while
vortexing
and
the
solution
was
centrifuged
at
350
X
g
for
4
 min
 and
 20°C.
 Following
 aspiration
 of
 the
 supernatant,
 the
 cells
 were
 incubated
 with
 a
 1:100
 dilution
 of
 anti‐BrdU
 monoclonal
 antibody
 (Santa
 Cruz
 Biotechnology,
 Santa
 Cruz,
 CA)
in
PBT
(PBS
+
0.5%
Tween‐20)
for
60
min
at
room
temperature,
in
the
dark,
then
rinsed
 with
PBTB.

The
cells
were
then
incubated
with
FITC‐conjugated
goat
anti‐mouse
antibody
 (1:100;
Santa
Cruz
Biotechnology,
Santa
Cruz,
CA)
in
PBTG
(PBTB
+
1.0%
goat
serum)
for
45
 min
 at
 room
 temperature
 in
 the
 dark,
 followed
 by
 rinse
 with
 PBTB.
 
 The
 cells
 were
 then
 stained
 with
 10
 µg/mL
 propidium
 iodine
 (PI;
 Sigma‐Aldrich,
 Oakville,
 ON)
 in
 PBTB
 overnight.
 Thirty
 minutes
 prior
 to
 analysis,
 RNAse
 A
 (1mg/ml
 in
 water;
 Sigma‐Aldrich,
 Oakville,
ON)
was
added
to
each
tube.

 
 FACS
 analysis:
 The
 samples
 (10,000
 cells/treatment
 group)
 were
 analyzed
 using

 flow
 cytometry
 (CellQuest,
 Becton‐Dickinson,
 Franklin
 Lakes,
 NJ)
 at
 excitation
 of
 488
 nm.
 BrdU
(FITC,
green
fluorescence)
was
measured
using
a
logarithmic
amplifier
with
a
530
nm
 short‐pass
filter
and
linear
DNA
content
(PI,
red
fluorescence)
was
measured
using
a
610
 nm
long‐pass
filter.
The
percentage
of
cells
in
the
G1,
S
and
G2/M
phases
were
determined
 using
FlowJo
8.7
analysis
software.

 
 2.2.8.

RNA
Extraction
and
Real
Time
Quantitative
PCR
 
 HepG2
 cells
 were
 cultured
 in
 6
 cm
 culture
 dishes
 at
 a
 seeding
 density
 of
 150,000
 cells
in
regular
MEM
and
allowed
to
attach
overnight.
The
culture
medium
was
changed
to
  
  35
  low‐,
adequate‐
or
high‐zinc
medium.
Medium
was
changed
once
more
after
3
days.
After
 6
days,
dishes
were
treated
with
RA
at
0
(DMSO
only)
or
10
µM
(n=4)
for
6
h
for
assessing
 the
expression
of
RARβ,
and
at
35
µM
for
48
h
for
assessing
the
expression
of
Cyp26a1.

 
 Total
 cellular
 RNA
 was
 isolated
 using
 TriZol
 (Invitrogen,
 Burlington,
 ON)
 in
 accordance
 with
 the
 manufacturer's
 directions,
 with
 the
 following
 modifications.
 Briefly,
 1mL
 of
 TriZol
 was
 added
 per
 plate,
 and
 scraped
 using
 a
 cell
 scraper.
 At
 the
 precipitation
 step,
samples
were
incubated
in
equal
volumes
of
2‐propanol
and
high
salt
solution
(0.8
M
 sodium
 citrate,
 1.2
 M
 sodium
 chloride)
 for
 10
 min
 at
 ‐80oC.
 The
 final
 RNA
 product
 was
 dissolved
in
30
µL
of
DEPC‐treated
ddH2O
and
stored
at
‐80oC
until
analysis.
The
integrity
of
 RNA
 was
 assessed
 by
 electrophoretic
 separation
 of
 total
 RNA
 on
 a
 0.8%
 agarose
 gel
 containing
GelRed
Nucleic
Acid
Stain
(Biotium,
Hayward,
CA,
USA)
at
80
V
for
45
minutes
 followed
 by
 visualization
 under
 UV
 light.
 
 RNA
 purity
 was
 assessed
 by
 determining
 the
 OD260/OD280
ratio
using
a
spectrophotometer
(Nanodrop
ND‐1000,
Wilmington,
DE,
USA).
 Samples
with
an
OD260/OD280
or
≥1.7
were
used
for
the
synthesis
of
cDNA.

 
 Total
 RNA
 (5
 µg)
 was
 used
 to
 generate
 cDNA
 with
 the
 SuperScript
 III
 First‐Strand
 Synthesis
 System
(Invitrogen,
 Burlington,
 ON)
 according
 to
 the
 manufacturers’
 directions.
 Briefly,
5
µg
RNA
dissolved
in
DEPC‐treated
water
was
mixed
with
1
µL
of
50
µM
Oligo‐DT
 and
1
µL
of
100
mM
dNTP
mix

and
DEPC‐treated
water
to
a
total
reaction
volume
of
10
µL
 and
incubated
at
65°C
for
5
min.
Then
10
µL
of
master
mix
was
added
to
each
reaction
(1X
 RT
buffer,
5
mM
MgCl2,
10
mM
DTT,
1
µL
RNase
OUT
and
1
µL
Superscript
III
RT)
and
the
  
  36
  reaction
was
carried
out
at
50°C
for
50
min
and
then
85°C
for
20
min.
The
product
(50
ng)
 was
amplified
using
RT2
Real‐TimeTM
SYBR
Green/ROX
PCR
2X
master
mix
(SABiosciences,
 Frederick,
MD,
USA)
with
500
nM
each
reverse
and
forward
primers
(Table
A.2)
to
a
final
 volume
of
20
µL
with
DEPC‐treated
water
in
a
thermocycler
(7500
Real‐Time
PCR
System,
 Applied
Biosytems,
Foster
City,
CA,
USA).
The
abundance
of
the
mRNA
of
the
target
gene
 was
normalized
on
the
fluorescence
of
the
corresponding
cyclophilin
control.

 
 2.2.9.

Protein
Extraction
and
Western
Blot
Analysis
 
 HepG2
cells
were
cultured
into
10
cm
culture
dishes
at
an
initial
seeding
density
of
 250,000
cells
in
MEM
plus
10%
FBS
and
allowed
to
attach
overnight.
Medium
was
changed
 to
MEM
plus
10%
Chelex
100‐treated
FBS
plus
0
µM
zinc
(low),
5
µM
zinc
(adequate)
or
10
 µM
zinc
(high;
n=4).
Medium
was
changed
once
more
after
3
days.

After
6
days,
the
cells
 were
harvested
and
counted,
as
described
above,
except
that
the
cells
were
re‐suspended
 in
1.5
mL
of
cold
PBS
twice
following
centrifugation
at
1000
rpm
for
5
m
at
4°C
to
rinse
the
 cells.

The
cell
suspension
was
then
transferred
to
a
microcentrifuge
tube
and
centrifuged
 14
 X
 g
 for
 5
 min
 at
 40C.
 After
 aspiration
 of
 the
 supernatant,
 whole
 cell
 radioimmunoprecipitation
assay
buffer
(RIPA:
50
mM
Tris
HCl,
pH
7.4,
150
mM
NaCl,
1
mM
 EDTA,
0.1%
SDS,
1%
sodium
deoxycholate,
1%
triton
X‐100,
1
mM
phenylmethylsulphonyl‐ fluoride
 (PMSF),
 10%
 Protease
 Inhibitor
 Cocktail)
 was
 added
 at
 30
 µL/106
 cells.
 Samples
 were
 incubated
 on
 ice
 for
 30
 min
 and
 then
 centrifuged
 for
 10
 min
 at
 14
 X
 g.
 The
 total
 protein
 was
 quantified
 using
 the
 BioRad
 DC
 method
 (BioRad,
 Hercules,
 CA,
 USA)
 and
 the
 samples
were
stored
at
‐80°C.

  
  37
  Protein
 samples
 (35
 µg/lane
 for
 RXRα
 and
 50
 µg/lane
 for
 RARα)
 were
 size‐ fractioned
 by
 10%
 SDS‐PAGE
 at
 175
 V
 for
 60
 min
 and
 transferred
 to
 a
 PVDF
 membrane
 (Millipore,
 Billerica,
 MA)
 for
 3
 h
 at
 a
 constant
 current
 of
 200
 mA.
 The
 membrane
 was
 subsequently
 blocked
 with
 2.5%
 skim
 milk
 powder
 in
 TTBS
 buffer
 (50
 mM
 Tris,
 150
 mM
 NaCl,
0.05%
Tween
20)
for
1
h,
incubated
with
an
appropriate
primary
antibody
as
listed
in
 Appendix
 4
 (Santa
 Cruz
 Biotechnology,
 Santa
 Cruz,
 CA),
 washed
 with
 TTBS
 for
 10
 min
 3
 times.
 
 Finally,
 the
 membrane
 was
 incubated
 with
 horseradish
 peroxidase‐conjugated
 secondary
antibody
(Santa
Cruz
Biotechnology,
Santa
Cruz,
CA)
in
1%
skim
milk
powder
in
 TTBS
for
45
min
at
dilutions
listed
in
Table
A.3.
Membranes
were
stripped
and
re‐probed
 with
a
β‐actin
antibody
(Santa
Cruz
Biotechnology,
Santa
Cruz,
CA)
to
confirm
equal
loading
 of
protein.
Briefly,
the
membrane
to
be
stripped
was
incubated
at
room
temperature
for
5
 min
twice
in
stripping
buffer
(pH
2.2;
0.2M
glycine,
3.5mM
SDS,
1%
Tween‐20),
and
then
 rinsed
for
10
min
twice
in
PBS
and
5
min
twice
in
TTBS.
Presence
of
the
target
protein
was
 visualized
using
an
enhanced
chemiluminescence
kit
(Thermo
Scientific,
Rockford,
IL,
USA)
 and
chemiluminescence
film
(Kodak
BioMax
Light
film,
Sigma‐Aldrich,
St.
Louis,
MO,
USA).
 Images
 of
 the
 bands
 were
 captured
 by
 a
 digital
 camera
 and
 the
 band
 intensity
 was
 measured
using
a
software
(Kodak
ID
v.
3.6.5,
Scientific
Imaging
Systems,
New
Haven,
CT,
 USA).
 
 
 
 
  
  38
  2.2.10.

Statistical
Analyses
 The
results
were
analyzed
using
one‐way
ANOVA
followed
by
Tukey’s
Honesty
Test
 (p
 <
 0.05).
 Statistical
 analyses
 were
 performed
 using
 GraphPad
 Prism
 (5.0
 for
 Mac
 OS
 X;
 GraphPad
Software,
San
Diego,
CA,
USA).

 
 2.3.

Results
 2.3.1.

RA‐Mediated
Reduction
in
Cell
Proliferation
 In
the
absence
of
zinc
manipulation,
no
reduction
in
cell
proliferation
was
observed
 in
 HepG2
 cells
 treated
 with
 0,
 15,
 and
 25
 µM
 RA
 for
 12
 h
 (Figure
 2.1).
 
 At
 35
 µM
 RA
 treatment,
a
44%
reduction
in
cell
proliferation
was
observed.

 
 2.3.2.
 Treatment
 Duration‐Dependent
 Reduction
 in
 Total
 Cellular
 Zinc
 Level
 and
 the
 Abundance
of
LIPZ
 In
 HepG2
 cells,
 the
 abundance
 of
 LIPZ
 was
 reduced
 by
 74
 and
 86%
 after
 being
 cultured
 in
 the
 low
 zinc
 medium
 for
 3
 and
 6
 d,
 respectively,
 compared
 to
 the
 control
 (Figure
2.2A).
Total
cellular
zinc
level
was
significantly
decreased
by
29%
after
6
d
of
culture
 (Figure
2.2B).
These
results
suggested
that
treating
the
HepG2
cells
with
low
zinc
for
a
total
 of
6
d
was
sufficient
to
reduce
the
abundance
of
LIPZ
as
well
as
the
total
cellular
zinc
levels
 in
the
cells.
 
 
 
  
  39
  2.3.3.

Dose‐Dependent
Reduction
in
Total
Cellular
Zinc
Level
and
the
Abundance
of
LIPZ
 
  
LIPZ
 abundance
 doubled
 in
 HepG2
 cells
 cultured
 for
 6
 d
 in
 high‐zinc
 medium
  compared
 to
 low‐
 and
 adequate‐zinc
 media
 (Figure
 2.3A);
 however,
 there
 was
 no
 significant
 difference
 in
 LIPZ
 abundance
 between
 the
 low‐
 and
 adequate‐zinc
 groups
 (Figure
2.3A).
Total
cellular
zinc
concentration
was
doubled
in
the
adequate‐
and
high‐zinc
 groups
 compared
 to
 the
 low‐zinc
 group,
 but
 the
 levels
 were
 the
 same
 between
 the
 adequate‐
and
high‐zinc
groups
(Figure
2.3B).

These
results
suggested
that
the
abundance
 of
LIPZ
and
total
cellular
zinc
level,
responded
to
zinc
treatment
in
HepG2.
 
 2.3.4.
 Zinc
 Appeared
 to
 Sensitize
 HepG2
 Cells
 to
 RA‐Induced
 Reduction
 in
 Cell
 Proliferation
 In
HepG2,
in
the
absence
of
RA
treatment,
cells
grown
in
high‐zinc
media
exhibited
 lower
cell
proliferation
as
measured
by
BrdU
incorporation
compared
to
cells
grown
in
low‐
 or
 adequate‐zinc
 media.
 RA
 suppressed
 cell
 proliferation
 at
 all
 levels
 of
 zinc
 treatment
 compared
 to
 their
 corresponding
 RA
 controls.
 The
 greatest
 suppression
 was
 in
 the
 adequate
 zinc
 group
 (50%),
 compared
 to
 the
 low
 zinc
 group
 (35%)
 in
 response
 to
 RA
 treatment.

In
the
high‐zinc
group,
suppression
of
cell
proliferation
was
only
30%
compared
 to
the
RA
control.
However
overall,
treatment
with
high‐zinc
and
35
µM
RA
resulted
in
the
 lowest
cell
proliferation
(Figure
2.4).

 
 Cell
 cycle
 analysis
 was
 performed
 on
 HepG2
 cells
 following
 6
 d
 of
 growth
 in
 low‐,
 adequate‐,
and
high‐zinc
media
and
0,
24,
and
72
h
of
treatment
with
35
µM
RA.
No
clear
  
  40
  trend
 in
 the
 proportion
 of
 cells
 in
 the
 G2/M
 phase
 was
 observed
 following
 zinc
 and
 RA
 treatment.
RA
increased
the
proportion
of
cells
in
the
G1
phase
after
24
and
72
h
following
 growth
in
adequate‐
and
high‐zinc
media
(Table
2.1;
Figure
2.5).
In
contrast,
cells
grown
in
 low‐zinc
responded
to
RA
treatment
with
a
decrease
in
the
percentage
of
cells
in
the
G1
 phase
 after
 24
 h
 and
 no
 change
 after
 72
 h.
 In
 cells
 grown
 in
 the
 low‐zinc
 medium,
 the
 percentage
 of
 cells
 in
 G1
 phase
 was
 67,
 63
 and
 68%
 in
 0,
 24,
 and
 72
 h
 RA
 treatments,
 respectively.
When
cells
were
grown
in
the
adequate‐zinc
medium,
the
proportion
of
cells
 in
the
G1
phase
was
57,
69
and
68%.
In
cells
grown
in
the
high‐zinc
medium,
the
proportion
 of
cells
in
G1
was
58,
68,
and
70%.

 
 In
the
absence
of
RA,
zinc
appeared
to
slightly
increase
the
proportion
of
cells
in
the
 S‐phase
with
19,
20
and
22%
of
cells
in
the
S‐phase
in
the
low‐,
adequate‐,
and
high‐zinc
 groups,
respectively
(Figure
2.5A).
In
contrast,
when
RA
was
present,
the
proportion
of
cells
 in
 the
 S‐phase
 was
 decreased
 with
 increasing
 zinc
 concentration.
 After
 treating
 the
 cells
 with
35
µM
of
RA
for
24
h
the
proportion
of
cells
in
the
S‐phase
was
reduced
from
16%
in
 cells
grown
in
the
low‐zinc
media
to
13%
in
the
adequate‐zinc
media
to
9%
in
the
high‐zinc
 media
 (Figure
 2.5B).
 At
 72
 h
 of
 treatment,
 the
 proportion
 of
 cells
 in
 the
 S‐phase
 was
 reduced
 from
 9
 to
 8
 to
 6%
 (Figure
 2.5C).
 These
 results
 suggested
 that
 RA
 reduced
 the
 proportion
 of
 cells
 in
 the
 S‐phase
 at
 24
 and
 72
 h
 of
 treatment
 and
 this
 effect
 was
 dependent
on
the
level
of
media
zinc.
 
  
  
  
  41
  2.3.5.

Cell
Viability
Was
Unaffected
by
RA
and
Zinc
 The
cell
viability
was
affected
by
neither
zinc
nor
RA
treatment
(Figure
2.6).
 
 2.3.6.

Growth
of
HepG2
in
the
Low‐Zinc
Medium
Increased
mRNA
Abundance,
But
Not
 Protein
Levels
of
the
RXRα
Nuclear
Receptor
 
  Abundance
 of
 mRNA
 of
 RARα,
 RARβ,
 RARγ,
 RXRβ
 and
 RXRγ
 was
 generally
  unaffected
 by
 zinc
 treatment
 in
 HepG2
 cells.
 However,
 abundance
 of
 RXRα mRNA
 was
 increased
 by
 2‐fold
 in
 cells
 grown
 in
 the
 low‐zinc
 medium
 compared
 to
 those
 grown
 in
 adequate‐zinc
 medium
 (Figure
 2.7).
 Western
 blot
 analysis
 revealed
 no
 changes
 in
 abundance
of
RXRα or
RARα
protein
levels
among
zinc
treatment
groups

(Figure
2.8).
 
 2.3.7.
 
 Zinc
 Treatment
 Did
 Not
 Affect
 the
 RA‐Induced
 Increase
 in
 mRNA
 Abundance
 of
 Downstream
Targets,
RARβ
and
CYP26a1
 In
 HepG2
 cells,
 treatment
 of
 RA
 for
 6
 h
 at
 10
 µM
 caused
 3
 fold
 increase
 in
 RARβ
 mRNA
expression
in
all
zinc
treatment
groups
compared
to
their
RA
control
(DMSO
only;
 data
 not
 shown).
 There
 was
 no
 difference
 in
 induction
 of
 RARβ
 by
 RA
 among
 the
 zinc
 treatment
groups
(Figure
2.9)
 
 Treatment
of
RA
for
24
h
at
35
µM
following
zinc
treatment
caused
an
approximate
 6‐fold
increase
in
CYP26a1
mRNA
in
all
zinc
treatment
groups
compared
to
the
RA
control
 (DMSO
only;
data
not
shown).
However,
there
was
no
difference
in
induction
of
CYP26a1
  
  42
  among
the
zinc
treatment
groups
(Figure
2.10).
This
data
suggests
that
zinc
status
did
not
 affect
RA‐induced
expression
of
the
downstream
targets:
RARβ
and
CYP26a1.
 
 2.4.

Discussion
 
 2.4.1.

Growth
in
Medium
Depleted
of
Zinc
was
Effective
in
Reducing
Cellular
Zinc
in
 HepG2
 My
 hypothesis
 was
 that
 zinc
 status
 is
 important
 for
 RA
 function,
 particularly
 the
 growth
 suppressive
 effects
 in
 HepG2
 hepatocarcinoma
 cells.
 In
 order
 to
 test
 this,
 it
 was
 first
 necessary
 to
 manipulate
 cellular
 zinc.
 Since
 a
 change
 in
 medium
 zinc
 does
 not
 necessarily
 induce
 a
 change
 in
 cellular
 zinc,
 to
 confirm
 the
 effects
 of
 zinc
 treatment
 on
 cellular
zinc
levels,
a
time
course
was
performed
to
determine
the
length
of
time
necessary
 to
 deplete
 cellular
 zinc
 and
 a
 dosage
 test
 to
 confirm
 a
 difference
 among
 the
 zinc
 treatments.
 
 Cellular
zinc
status
was
assessed
by
measuring
both
total
cellular
zinc
level
and
the
 abundance
of
the
LIPZ.
Total
cellular
zinc
is
a
measure
of
the
amount
of
zinc
present
in
the
 entire
cell,
much
of
which
is
tightly
bound
to
proteins.
LIPZ
is
a
dynamic
pool
of
zinc
loosely
 bound
to
low
molecular
weight
ligands
and
a
small
amount
of
free
ionic
zinc
(in
the
fM‐pM
 range),
with
the
entire
pool
making
up
less
than
10%
of
the
total
cellular
zinc
(Franklin
and
 Costello,
 2009).
 
 This
 labile
 pool
 provides
 a
 supply
 of
 zinc
 for
 cellular
 signaling,
 second
 messenger
 metabolism,
 and
 the
 function
 of
 enzymes
 (Kambe
 et
 al.,
 2004).
 Total
 cellular
  
  43
  zinc
therefore
represents
mostly
zinc
incorporated
into
proteins,
and
LIPZ
represents
zinc
 readily
available
to
the
cell.

 
 It
 was
 observed
 that
 growth
 of
 HepG2
 cells
 in
 the
 low‐zinc
 medium
 significantly
 reduced
 the
 level
 of
 total
 cellular
 zinc
 and
 the
 abundance
 of
 the
 LIPZ
 after
 6
 d
 and
 that
 cellular
 zinc
 is
 significantly
 different
 between
 low‐,
 adequate‐,
 and
 high‐zinc
 treatments.
 Similarly,
one
other
study
by
Reaves
et
al.
(2000)
found
that
growth
of
HepG2
in
a
low‐zinc
 media
 (0.4
 µM)
 also
 significantly
 depleted
 total
 cellular
 zinc
 levels
 after
 6.5
 d.
 
 These
 observations
 confirm
 that
 growth
 in
 media
 depleted
 of
 zinc
 was
 a
 successful
 method
 of
 depleting
cellular
zinc
in
HepG2
cells
and
that
cellular
zinc
levels
were
influenced
by
media
 zinc
levels
in
HepG2
hepatocarcinoma
cells.

 
 An
alternate
method
of
zinc
depletion
involves
incubation
with
N,N,N',N'‐tetrakis(2‐ pyridylmethyl)ethylenediamine
(TPEN),
a
membrane
permeable
zinc
chelator,
(Kolenko
et
 al.,
2001).
However,
this
method
was
not
chosen
in
my
research
as
TPEN
exhibits
high
zinc‐ binding
 affinity,
which
could
 remove
the
entire
labile
 pool
 of
zinc
as
well
as
a
fraction
of
 immobile
 macromolecular‐bound
 zinc.
 This
 would
 not
 be
 achievable
 under
 normal
 pathological
 conditions
 and
 could
 ultimately
 result
 in
 cell
 death
 (Franklin
 and
 Costello,
 2009).
In
addition,
excess
TPEN
is
not
zinc‐specific
and
could
remove
the
cellular
pools
of
 Mn
and
Fe
(Franklin
and
Costello,
2009).
Reaves
et
al.
(2000)
concluded
that
culturing
cells
 in
low‐zinc
medium
to
deplete
cellular
zinc
more
closely
resembles
physiologic
conditions
 than
exposing
the
cells
directly
to
a
metal
chelator.

  
  44
  2.4.2.

Zinc

Appeared
to
Sensitize
HepG2
Cells
to
RA‐Induced
Reduction
in
Cell
 Proliferation
 I
hypothesized
that
zinc
status
would
effect
the
growth
suppressant
function
of
RA
 in
HepG2
cells.
To
determine
if
the
altered
zinc
status
in
HepG2
cells
had
an
effect
on
RA‐ induced
inhibition
of
cell
proliferation,
cells
were
treated
with
RA
following
growth
in
low‐,
 adequate‐,
 and
 high‐zinc
 media.
 The
 level
 of
 reduction
 of
 cell
 proliferation
 following
 RA
 treatment,
 as
 measured
 by
 the
 BrdU
 assay,
 appeared
 to
 be
 dependent
 on
 the
 level
 of
 media
zinc
in
HepG2
cells.
Cells
grown
in
high‐zinc
media
exhibited
lower
cell
proliferation
 than
those
grown
in
low‐
or
adequate‐zinc
medium.
 
 Further
 investigation
 into
 the
 effect
 of
 RA
 on
 cell
 cycle
 progression
 following
 a
 similar
 treatment
 regime
 revealed
 a
 positive
 correlation
 between
 media
 zinc
 and
 the
 percentage
of
cells
in
the
S‐phase
of
the
cell
cycle
following
 RA
treatment.
RA
treatment
 also
 caused
 an
 increase
 in
 the
 percentage
 of
 cells
 in
 the
 G1‐phase
 compared
 to
 control
 following
 growth
 in
 adequate‐
 and
 high‐zinc
 media,
 but
 not
 in
 low‐zinc
 media.
 These
 observations
suggest
that
zinc
may
be
important
to
the
growth‐suppressant
function
of
RA
 in
HepG2
cells,
particularly
the
role
of
RA
in
the
inhibition
of
cell
cycle
progression
from
G1‐
 to
S‐phase.
 
 In
another
study,
a
correlation
between
media
zinc
and
RA
function
was
observed
 when
testing
the
effects
of
zinc
deficiency
on
RA‐induced
differentiation
of
neuronal
cells
 (Gower‐Winters,
2008).
However,
to
my
knowledge,
this
is
the
only
other
study
attempting
  
  45
  to
assess
the
effect
of
zinc
status
on
RA
function.
The
authors
did
not
uncover
a
mechanism
 but
 hypothesized
 that
 the
 impaired
 RA
 function
 during
 growth
 in
 media
 containing
 low
 levels
of
zinc
is
due
to
impaired
RAR/RXR
nuclear
receptor
function.

 
 An
 inconsistency
 between
 the
 BrdU
 assay
 and
 cell
 cycle
 analysis
 was
 observed
 in
 cells
grown
in
high‐zinc
medium
and
treated
with
0
µM
RA.
In
this
group,
the
BrdU
assay
 showed
 a
 decrease
 in
 cell
 proliferation
 compared
 to
 low‐
 and
 adequate‐zinc
 treatments,
 however
cell
cycle
analysis
showed
a
slight
increase
in
BrdU
incorporation
in
this
treatment
 group.
 
 One
 possible
 explanation
 for
 this
 inconsistency
 is
 that
 high‐zinc
 conditions
 may
 induce
apoptosis,
which
would
affect
the
BrdU
assay
results
by
decreasing
the
cell
number
 per
 well.
 This
 would
 not
 be
 detected
 in
 cell
 cycle
 analysis
 with
 flow
 cytometry,
 as
 this
 method
 measures
 the
 percentage
 of
 live
 cells
 stained
 with
 BrdU
 (dead
 cells
 are
 not
 included
 in
 the
 analysis).
 However,
 I
 am
 unaware
 of
 any
 published
 data
 showing
 observations
of
increased
apoptosis
in
HepG2
cells
due
to
zinc
supplementation.
 
 Although
 the
 BrdU
 assay
 and
 cell
 cycle
 analysis
 appeared
 to
 show
 a
 sensitizing
 effect
 of
 zinc
 on
 RA‐induced
 inhibition
 of
 cell
 proliferation,
 it
 is
 unclear
 whether
 this
 relationship
 is
 significant
 to
 produce
 a
 clinically
 relevant
 change.
 RA
 treatment
 following
 growth
in
low‐,
adequate‐,
or
high‐zinc,
resulted
in
no
change
in
cell
viability
between
RA‐ treated
 and
 un‐treated
 cells
 in
 all
 levels
 of
 media
 zinc
 after
 72
 h,
 indicating
 that
 the
 changes
observed
in
cell
proliferation
may
not
be
significant
enough
to
induce
a
change
in
 viability
within
this
time
frame.

  
  46
  2.4.3.

Growth
of
HepG2
in
Low
Zinc
Media
Resulted
in
Increased
mRNA
Abundance,
But
 Not
Protein
Levels
of
the
RXRα
Nuclear
Receptor

 The
 effects
 of
 zinc
 status
 on
 RAR
 and
 RXR
 nuclear
 receptor
 expression
 was
 investigated
 as
 a
 possible
 mechanism
 for
 the
 change
 in
 sensitivity
 of
 HepG2
 cells
 to
 RA
 following
 zinc
 manipulation.
 RAR/RXR
 nuclear
 receptors
 are
 critical
 for
 RA
 function
 (Soprano
and
Soprano,
2002).
Zinc
deficiency
has
been
shown
to
have
wide‐ranging
effects
 on
 gene
 expression
 (Dieck
 et
 al.,
 2003),
 but
 it
 was
 not
 known
 if
 zinc
 status
 affects
 the
 expression
of
RAR/RXR
receptors
in
HepG2
cells.

 
 The
only
receptor
observed
to
be
affected
by
zinc
status
was
RXRα,
which
exhibited
 an
 increase
 in
 mRNA
 abundance,
 but
 not
 protein
 level,
 following
 growth
 in
 low‐zinc
 medium
 compared
 to
 adequate‐zinc.
 It
 is
 not
 known
 if
 this
 receptor
 is
 involved
 in
 the
 growth
regulation
pathway.
However,
if
this
receptor
were
involved,
the
increase
in
mRNA
 abundance
 might
 indicate
 a
 compensatory
 mechanism
 for
 decreased
 function,
 similar
 to
 the
response
of
p53,
another
zinc‐finger,
to
zinc
deficiency.
Low
zinc
results
in
an
increased
 mRNA
expression
of
p53,
whereas
its
DNA
binding
ability
is
reduced
(Ho
and
Ames,
2002).
 However,
 it
 is
 thought
 that
 the
 increase
 in
 p53
 expression
 may
 be
 a
 response
 to
 DNA
 damage
resulting
from
low
intracellular
zinc
(Ho
et
al.,
2004).

The
level
of
nuclear
receptor
 mRNA
is
not
necessarily
reflective
of
the
function
of
this
protein.
Therefore,
it
is
possible
 that
 the
 DNA‐binding
 ability
 of
 the
 nuclear
 receptors,
 RARs
 and
 RXRs,
 is
 impaired
 by
 low
 zinc.
 
  
  47
  An
inconsistency
between
mRNA
and
protein
level
in
RXRα could
possibly
be
due
to
 improper
 protein
 folding.
 Zinc‐fingers
 are
 essential
 for
 proper
 protein
 folding
 (Klug
 and
 Schwabe,
1995).

An
effect
of
zinc
depletion
on
improper
folding
of
zinc‐finger
proteins
is
 demonstrated
 by
 p53,
 which
 adopts
 a
 mutant
 conformation
 following
 induction
 of
 zinc‐ deficiency
(Hainaut
and
Milner,
1993).

Misfolded
proteins
are
degraded
via
the
ubiquitin‐ proteasome
system
(Eisele
and
Wolf,
2008).
 
 2.4.4.
 
 Zinc
 Treatment
 Did
 Not
 Affect
 the
 RA‐Induced
 Increase
 in
 mRNA
 Abundance
 of
 Downstream
Targets,
RARβ
and
CYP26a1
 To
 determine
 if
 the
 function
 of
 RXR/RAR
 nuclear
 receptors
 was
 affected
 by
 zinc
 status,
 the
 mRNA
 expression
 of
 two
 well
 established
 RA‐inducible
 genes,
 CYP26a1
 and
 RARβ
(Ozpolat
et
al.,
2005;
Altucci
et
al.,
2007),
was
measured.
It
has
been
observed
that
 zinc
 deficiency
 causes
 impaired
 DNA
 binding
 of
 some
 zinc‐finger
 proteins,
 including
 p53,
 NFkB,
 and
 AP‐1
 in
 rat
 gliomal
 cells
 grown
 in
 medium
 supplemented
 with
 Chelex‐100
 treated
 FBS
 (Ho
 and
 Ames,
 2002).
 Since
 RAR
 and
 RXR
 nuclear
 receptors
 contain
 zinc‐ fingers,
then
theoretically,
low
zinc
could
lead
to
impaired
DNA‐binding
of
these
receptors
 and
ultimately,
impaired
expression
of
downstream
genes.

 
 A
 difference
 among
 zinc
 groups
 in
 RA‐induced
 expression
 of
 CYP26a1
 or
 RARβ
 following
 growth
 in
 low‐,
 adequate‐,
 and
 high‐zinc
 media
 was
 not
 observed.
 This
 implied
 that
the
function
of
the
RAR/RXR
receptors
was
not
affected
by
zinc.


 
  
  48
  It
is
likely
that
some
other
mechanism
is
responsible
for
the
observation
that
zinc
 sensitized
 HepG2
 cells
 to
 RA‐induced
 reduction
 in
 cell
 proliferation.
 Firstly,
 since
 zinc
 has
 wide‐ranging
 effects
 on
 gene
 expression,
 low
 zinc
 could
 alter
 the
 expression
 of
 genes
 involved
 in
 the
 RA‐induced
 inhibition
 of
 cell
 proliferation.
 For
 example,
 p21
 is
 a
 key
 regulator
 of
 the
 cell
 cycle.
 
 This
 protein
 binds
 to
 and
 inhibits
 the
 activity
 of
 cyclin‐CDK2
 complexes,
therefore
inhibiting
cell
cycle
progression
and
slowing
growth
(Maddika
et
al.,
 2007).
 Wong
 et
 al.
 (2007)
 found
 that
 growth
 of
 HepG2
 cells
 in
 medium
 depleted
 of
 zinc
 resulted
 in
 a
 40%
 decrease
 in
 p21
 protein
 expression
 and
 a
 70%
 decrease
 in
 p21
 mRNA.

 Therefore,
 if
 low
 zinc
 decreases
 the
 expression
 of
 p21,
 less
 of
 this
 protein
 would
 be
 available
to
participate
in
RA‐induced
inhibition
of
cell
proliferation.

 
 
Secondly,
 if
 zinc
 deficiency
 impairs
 zinc‐finger
 binding,
 this
 may
 also
 apply
 to
 protein‐protein
interactions,
as
zinc‐fingers
are
thought
to
mediate
protein‐protein
binding
 (Gamsjaeger
 et
 al.,
 2006).
 Cell
 cycle
 regulation
 involves
 numerous
 protein‐protein
 interactions.
For
example,
p21
is
a
zinc‐finger
protein,
and
the
region
required
for
binding
 to
cyclins
is
within
the
zinc‐finger
domain
of
the
protein
(The
UniProt
Consortium,
2011).
 Impaired
protein‐protein
interactions
among
cell
cycle
regulating
proteins
could
potentially
 result
in
altered
cell
cycle
progression.
 
 Third,
 the
 expression
 of
 p21
 is
 controlled
 by
 the
 zinc‐finger
 protein,
 p53
 (Hawkes
 and
Alkan,
2011).
Zinc
deficiency
has
been
shown
to
impair
DNA
binding
ability
of
p53
(Ho
 and
 Ames,
 2002).
 Recently
 it
 was
 discovered
 that
 RA
 induces
 hypomethylation
 in
 the
  
  49
  promoter
region
of
cell
cycle
regulatory
genes,
including
p21,
via
down‐regulation
of
DNA
 methyltransferases.
 This
 hypomethylation
 facilitates
 the
 binding
 of
 p53
 to
 the
 p21
 promoter,
resulting
in
up‐regulation
of
p21
and
subsequent
blockage
in
the
G1‐phase
and
 cell
 cycle
 arrest
 (Lim
 et
 al.,
 2011).
 
 Therefore,
 it
 is
 possible
 that
 impaired
 DNA‐binding
 ability
 of
 p53
 could
 lead
 to
 impaired
 up‐regulation
 of
 p21
 and
 interference
 with
 the
 inhibition
of
cell
proliferation.

 
 2.4.5.

Summary
 
  In
 summary,
 my
 thesis
 research
 revealed
 that
 growth
 in
 zinc‐deficient
 medium
 is
  sufficient
 to
 reduce
 intracellular
 zinc
 levels
 in
 HepG2
 cells.
 
 When
 HepG2
 cells
 grown
 in
 low‐,
adequate‐,
or
high‐zinc
medium
were
subsequently
treated
with
RA,
it
appeared
that
 zinc
 sensitized
 HepG2
 cells
 to
 RA‐induced
 inhibition
 of
 cell
 proliferation.
 However,
 the
 mechanism
behind
this
interaction
remains
unclear.
Zinc
deficiency
increased
expression
of
 one
nuclear
receptor,
RXRα,
however
it
is
unknown
if
this
protein
is
involved
in
RA‐induced
 inhibition
of
cell
proliferation.
Further,
the
function
of
the
nuclear
receptors,
measured
as
 RA‐induced
expression
of
two
downstream
targets,
RARβ
and
CYP26a1,
did
not
appear
to
 be
affected
by
zinc
status.


 
 
 
 
 
  
  50
  
 
 
 
 Table
 2.1:
 The
 effect
 of
 retinoic
 acid
 on
 proportion
 of
 cells
 in
 the
 G1‐
 and
 S‐phases
 following
zinc
treatment.*
 
 Retinoic
Acid
 Proportion
of
cells
in
the
S,
G1,
and
G2/M
phases
of
the
cell
cycle
 Treatment
Duration
 (%)
 (h)
















 Low
 Adequate
 High
 0

  24
  72
  
  *  
 
 
 
 
 
 
 
 
 
  S=
19
  S=
20
  S=
22
  G1=67
  G1=57
  G1=58
  G2/M=12
  G2/M=21
  G2/M=17
  S=
16
  S=
13
  S=
9
  G1=63
  G1=69
  G1=68
  G2/M=16
  G2/M=14
  G2/M=18
  S=
9
  S=
8
  S=
6
  G1=68
  G1=71
  G1=70
  G2/M=20
  G2/M=17
  G2/M=20
  Data
from
the
cell
cycle
analysis
shown
in
Figure
2.6
was
used
in
this
analysis.

  51
  
 
 
 
 
 
 
 
 
 
  
 
 
  
 
 
 
 
 
 
 
 
  
 
 
 
 
 
 
 Figure
2.1:
Cell
proliferation
of
HepG2
cells
treated
with
retinoic
acid.
HepG2
cells
were
 cultured
in
medium
supplemented
with
0,
15,
25
and
35
µM
retinoic
acid
(RA)
for
12
h.
Cell
 proliferation
 was
 quantified
 using
 the
 BrdU
 incorporation
 assay
 with
 absorbance
 as
 a
 measure
 of
 BrdU
 incorporation.
 Values
 represent
 mean
 ±
 SEM
 (n=6;
 the
 experiment
 was
 repeated
 once
 independently).
 Means
 with
 different
 letters
 are
 significantly
 different
 (p<0.05).
 
  
  52
  
 A.
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 B.
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 Figure
 2.2:
 
 Time‐dependent
 reduction
 in
 total
 cellular
 zinc
 level
 and
 the
 abundance
 of
 the
labile
intracellular
pool
of
zinc.
HepG2
cells
were
grown
in
media
supplemented
with
 Chelex‐100
 treated
 FBS
 for
 0,
 3
 and
 6
 days.
 (A)
 Labile
 intracellular
 pools
 of
 zinc
 (LIPZ)
 as
 measured
 by
 Zinquin
 fluorescent
 assay.
 (B)
 Total
 cellular
 zinc
 as
 measured
 by
 atomic
 absorption
 spectrophotometry.
 Values
 represent
 mean
 ±
 SEM
 (n=3;
 the
 experiment
 was
 repeated
 once
 independently).
 Means
 with
 different
 letters
 are
 significantly
 different
 (p<0.05).
 
 
 
  53
  
 A.
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 B.
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 Figure
2.3:
Dose‐dependent
reduction
in
total
cellular
zinc
level
and
the
abundance
of
the
 labile
intracellular
pool
of
zinc.
HepG2
cells
were
grown
in
media
containing
Chelex‐100‐ treated
FBS
and
0
(Low),
5
(Adequate),
and
10
(High)
µM
of
Zn
(ZnSO4)
for
6
days.

(A)
Total
 cellular
zinc
as
measured
by
atomic
absorption
spectrophotometry.
(B)
Labile
intracellular
 pools
of
zinc
(LIPZ)
as
measured
by
the
Zinquin
fluorescent
assay.
Values
represent
mean
±
 SEM
 (n=3;
 the
 experiment
 was
 repeated
 once).
 Means
 with
 different
 letters
 are
 significantly
different
(p<0.05).
 
 
  54
  
 
 
 
 
 
 
 
  
  
 
 
  
  
  
 
 
 
 
 
  
 
 
 
 
 
 
 
 Figure
2.4:
The
effect
of
retinoic
acid
on
the
proliferation
of
HepG2
cells
following
growth
 in
 low‐,
 adequate‐,
 and
 high‐zinc
 medium.
 HepG2
 cells
 were
 grown
 in
 media
 containing
 Chelex
100‐treated
FBS
plus
0
(Low),
5
(Adequate),
and
10
(High)
µM
Zn
(ZnSO4)
for
6
days
 followed
by
0
or
35
µM
retinoic
acid
(RA)
for
12
h.
Cell
proliferation
was
quantified
using
 the
BrdU
colorimetric
assay
with
absorbance
as
a
measure
of
BrdU
incorporation.
Values
 represent
 mean
 ±
 SEM
 (n=6).
 Means
 with
 different
 letters
 are
 significantly
 different.
 Asterisks
represent
a
significant
difference
between
the
RA
group
and
its
respective
control
 (p<0.05).

  
  55
  





 
 
Low
 
  
  













Adequate
  
  










High
  















  
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
  
 
 
  
 
  
 
  
 
 
 
 
 
 
 Figure
2.5:
The
effect
of
RA
on
the
percentage
of
cells
in
the
S‐Phase
following
growth
in
 low‐,
 adequate‐,
 and
 high‐zinc
 medium.
 HepG2
 cells
 were
 grown
 in
 media
 containing
 Chelex‐100‐treated
FBS
plus
0
(Low),
5
(Adequate),
and
10
(High)
µM
of
Zn
(ZnSO4)
for
6
d
 followed
by
35
µM
of
RA
for
0
(A),
24
(B)
and
72
(C)
h.
Cell
cycle
analysis
was
performed
by
 BrdU
 incorporation
 for
 determination
 cells
 within
 S‐phase
 and
 PI
 staining
 for
 total
 DNA
 content
 using
 flow
 cytometry.
 Cells
 in
 the
 S‐phase,
 G1/G0
 and
 G2/M
 phases
 are
 located
 within
the
small
box,
lower
left
quadrant,
and
lower
right
quadrant,
respectively.

 
 
 
  
  56
  
 
 
 
 
 
 
 
 
  
 
 
 
 
 
 
 
 
  
 
 
 
 
 
 
 
 
 
 Figure
2.6:
The
effect
of
retinoic
acid
on
viability
of
HepG2
cells
following
growth
in
low‐,
 adequate‐,
 and
 high‐zinc
 medium.
 HepG2
 cells
 were
 grown
 in
 media
 containing
 Chelex‐ 100‐treated
 FBS
 plus
 0
 (Low),
 5
 (Adequate),
 and
 10
 (High)
 µM
 of
 Zn
 (ZnSO4)
 for
 6
 d
 followed
by
0
or
35
µM
of
retinoic
acid
(RA)
for
72
h.
Cell
viability
was
quantified
using
the
 MTT
 colorimetric
 assay
 with
 absorbance
 as
 a
 measure
 of
 cell
 viability.
 Values
 represent
 mean
±
SEM
(n=12).
No
significant
difference
among
the
treatment
groups
was
identified
 (p<
0.05)


  
  57
  
 
 
 
 
 
  
 
 
 
  
 
 
 
 
 
  
 
 
 
 
 
 Figure
2.7:
The
relative
expression
of
retinoic
acid
receptor
(RAR)
and
retinoid
X
receptor
 (RXR)
 nuclear
 receptor
 mRNA
 in
 HepG2
 following
 growth
 in
 low‐
 and
 high‐
 zinc
 media.
 HepG2
 cells
 were
 grown
 in
 media
 containing
 Chelex‐100‐treated
 FBS
 plus
 0
 (Low;
 open
 bars),
10
(High;
black
bars)
and
5
(Adequate;
control;
grey
bars)
µM
of
Zn
(ZnSO4)
for
6
d.
 Values
represent
mean
fold
difference
in
comparison
to
adequate‐zinc
group
±
SEM
(n=4;
 the
 experiment
 was
 repeated
 once
 independently).
 The
 asterisk
 indicates
 a
 statistically
 significant
difference
in
∆∆CT
value
from
adequate
zinc
treatment.

 
 
 
 
 
 
  
  58
  
 
 
 
 
 
 
 
 
 
  
  
  
 
  
 
  
  
 
  
 
 
  
  
 
  
 
  
 
 Figure
 2.8:
 Western
 blot
 of
 retinoid
 X
 receptor‐α (RXRα)
 and
 retinoic
 acid
 receptor‐ α (RARα)
following
growth
in
low‐,
adequate‐,
and
high‐zinc
medium.

HepG2
cells
were
 grown
in
media
supplemented
with
Chelex
100‐treated
FBS
plus
0
(Low),
5
(Adequate)
or
 10
 (High)
 µM
 of
 Zn
 (ZnSO4)
 for
 6
 d.
 Protein
 levels
 were
 assessed
 using
 Western
 blot
 analysis.
Following
probing
with
anti‐RARα
antibody,
the
membrane
was
stripped
and
re‐ probed
 with β‐actin
 antibody
 to
 ensure
 equal
 protein
 loading.
 The
 same
 protein
 samples
 were
used
for
RARα
and
RXRα
Western
blot
analysis.
 
 
 
 
 
 
 
 
 
  59
  
 
 
 
 
 
 
 
 
  
  
 
 
 
 
 
 
 
 
 
 
  
 
 
 
 
 
 
 
 Figure
 2.9:
 The
 relative
 expression
 of
 retinoic
 acid
 receptor‐β (RARβ)
 mRNA
 in
 retinoic
 acid‐treated
 HepG2
 cells
 following
 growth
 in
 low‐,
 adequate‐,
 and
 high‐zinc
 medium.
 HepG2
 cells
 were
 grown
 in
 media
 containing
 Chelex
 100‐treated
 FBS
 plus
 0
 (Low),
 5
 (Adequate)
or
10
(High)
µM
of
Zn
(ZnSO4)
for
6
d
and
subsequently
treated
with
0
or
10
µM
 retinoic
acid
(RA)
for
6
h.
Values
represent
mean
fold
difference
in
comparison
to
control
 (adequate
zinc
treatment
+
0
µM
RA)
±
SEM
(n=4;
the
experiment
was
repeated
once).
No
 statistically
significant
difference
in
∆∆CT
value
was
observed
when
compared
to
control.



 
 
 
 
 
 
 
  
  60
  
 
 
 
 
 
 
 
 
 
  
  
  
  
  
 
 
 
 
 
 
 
 
  
 
 
 
 
 
 Figure
 2.10:
 The
 relative
 expression
 of
 CYP26a1
 mRNA
 in
 retinoic
 acid‐treated
 HepG2
 cells
following
growth
in
low‐,
adequate‐,
and
high‐zinc
medium.
HepG2
cells
were
grown
 in
media
containing
Chelex
100‐treated
FBS
plus
0
(Low),
5
(Adequate)
or
10
(High)
µM
of
 Zn
(ZnSO4)
for
6
d
and
subsequently
treated
with
0
or
35
µM
of
retinoic
acid
(RA)
for
24
 hours.
 Values
 represent
 mean
 fold
 difference
 in
 comparison
 to
 control
 (adequate
 zinc
 treatment
 +
 0
 µM
 RA)
 ±
 SEM
 (n=4;
 the
 experiment
 was
 repeated
 once).
 No
 statistically
 significant
difference
in
∆∆CT
value
was
observed
when
compared
to
control.

 
 
 
 
 
  
  61
  Chapter
3.
General
Discussion,
Limitations,
and
Future
Directions

 
 3.1.

General
Discussion
 
 Initial
experiments
were
performed
in
AML12,
a
murine
liver
cell
line,
in
order
to
 compare
zinc
levels
between
HepG2
and
a
non‐cancerous
liver
cells
and
whether
a
 difference
in
zinc
level
is
correlated
with
sensitivity
to
RA
treatment.

During
these
initial
 experiments
I
noted
that
increased
cellular
zinc
was
correlated
with
an
increased
sensitivity
 to
RA
treatment.
For
example,
the
total
cellular
zinc
levels
in
HepG2
were
much
lower
than
 those
observed
in
AML12
(about
75%
less
zinc;
Figure
A.1).

Interestingly,
lower
 concentrations
of
RA
were
required
to
inhibit
growth
in
AML12
than
HepG2.
A
reduction
in
 cell
proliferation
was
seen
at
as
low
as
15
µM
of
RA
treatment
in
AML12,
whereas
 reduction
in
cell
proliferation
was
not
observed
until
35µM
of
treatment
in
HepG2
cells
 (Figure
A.2;
Figure
2.1).
In
addition,
cell
viability
was
reduced
in
AML12
after
72
h
of
RA
 treatment,
whereas
no
reduction
was
seen
in
HepG2
(Figure
A.3,
Figure
2.6).
This
 relationship
between
zinc
level
and
sensitivity
to
RA
treatment
was
further
supported
by
 the
observation
that
growth
in
high‐zinc
medium
sensitized
HepG2
hepatocarcinoma
cells
 to
RA‐induced
reduction
in
cell
proliferation.
 



 One
possibility
for
an
interaction
between
zinc
and
vitamin
A
at
the
cellular
level
is
 that
zinc
deficiency
impairs
the
DNA‐binding
ability
of
RAR/RXR
receptors,
as
an
effect
of
 zinc
on
the
binding
ability
of
other
zinc‐finger
proteins,
including
p53
and
AP‐1,
has
been
  
  62
  observed
(Ho
and
Ames,
2002).
However,
my
observation
that
zinc
level
did
not
affect
the
 RA‐induced
expression
of
the
downstream
targets,
RARβ
and
CYP26a1,
fails
to
support
this
 hypothesis.
 This
 suggests
 that
 other
 mechanisms
 are
 likely
 present.
 
 These
 mechanisms
 could
include
the
influence
of
zinc
on
the
expression
or
function
of
other
proteins
involved
 in
regulation
of
the
cell
cycle,
such
as
p21,
a
cell
cycle
regulator.

 
 The
results
of
my
thesis
point
to
the
possibility
 that
zinc
may
be
important
 in
the
 growth‐suppressant
function
of
RA
in
HepG2.
This
observation
brings
to
light
the
possibility
 that
a
similar
phenomenon
could
be
present
in
other
tumor
and
cell
types.

In
one
study,
 zinc
deficiency
impaired
the
RA‐induced
differentiation
of
neuronal
cells
(Gower‐Winters,
 2008).
 If
 it
 is
 the
 case
 that
 zinc
 deficiency
 impairs
 the
 function
 of
 RA
 in
 other
 cancer
 cell
 types,
then
it
would
be
important
to
ensure
that
the
patients’
zinc
levels
are
adequate
for
 optimum
RA
function.

 
 However,
 in
 order
 for
 zinc
 manipulation
 to
 be
 a
 viable
 method
 of
 increasing
 the
 sensitivity
of
cancer
cells
to
RA
treatment,
it
would
be
necessary
for
the
sensitivity
of
non‐ cancerous
cells
to
be
unaffected,
or
decreased,
by
zinc.

In
my
comparisons
between
the
 effects
of
zinc
treatment
on
RA‐induced
reduction
in
cell
proliferation
between
HepG2
and
 AML12
 cells,
 I
 observed
 that,
 similar
 to
 HepG2,
 high
 zinc
 treatment
 resulted
 in
 a
 greater
 reduction
 in
 cell
 proliferation
 following
 RA
 treatment
 in
 AML12
 cells
 (figure
 A.4).
 This
 observation
suggests
that
zinc
treatment
may
not
be
an
appropriate
method
of
increasing
 the
 sensitivity
 of
 liver
 cancer
 to
 RA
 treatment,
 however,
 because
 AML12
 is
 a
 murine
 cell
  
  63
  line,
 confirmation
 of
 a
 similar
 response
 in
 a
 non‐cancerous
 human
 liver
 cell
 line
 is
 necessary.

 
 Currently,
 RA
 is
 used
 clinically
 in
 the
 treatment
 of
 acute
 promyelocytic
 leukemia
 (APL),
 head
 and
 neck
 carcinoma,
 and
 non‐small‐cell
 lung
 cancer
 (Bohnsack
 and
 Hirschi,
 2004).
 In
 APL,
 treatment
 induces
 differentiation
 and
 often
 results
 in
 complete
 remission
 (McNamara
 et
 al.,
 2010).
 However,
 these
 results
 are
 not
 long
 lasting,
 as
 prolonged
 treatment
leads
to
the
emergence
of
RA
resistance
(Tallman
et
al.,
2002).

In
addition,
in
 about
2‐27%
of
patients,
RA
treatment
can
have
severe
side
effects,
including
acute
renal
 failure
and
central
nervous
system
toxicity,
with
about
2%
of
treatments
resulting
in
death
 (Vanier
et
al,
2003;
Patatanian
and
Thompson,
2008).
In
the
case
of
APL,
it
is
understood
 that
 in
 33%
 of
 cases,
 resistance
 is
 brought
 on
 by
 an
 acquired
 mutation,
 however,
 in
 the
 remaining
67%,
the
mechanisms
of
resistance
are
unknown
(McNamara
et
al.,
2010).
 
  It
has
been
proposed
that
a
major
contributing
factor
to
acquired
RA
resistance
is
  the
 induction
 of
 CYP26a1
 (a
 P450
 enzyme)
 in
 tissues
 such
 as
 the
 liver
 (Thatcher
 and
 Isoherranen,
2009).
In
cases
of
resistance,
instead
of
increasing
RA
dosage,
P450
inhibitors
 are
co‐administered
to
prevent
the
metabolism
and
clearance
of
RA
(Sun
and
Lotan,
2002).
 These
inhibitors
are
referred
to
as
RA
metabolism
blocking
agents
(RAMBAs;
Verfaille
et
al.,
 2008).

In
one
case
study,
co‐administration
of
fluconazole
(a
P450
inhibitor)
led
to
a
70%
 reduction
in
RA
dosage
(Vanier
et
al.,
2003).

 
  
  64
  An
 attractive
 alternative
 to
 administering
 additional
 drugs
 would
 be
 manipulation
 of
 nutrient
 intake,
 such
 as
 zinc.
 
 According
 to
 my
 results,
 zinc
 status
 did
 not
 affect
 RA‐ induced
 CYP26a1
 expression
 in
 HepG2
 liver
 cells.
 
 Therefore,
 zinc
 manipulation
 does
 not
 appear
to
be
a
viable
option
for
controlling
RA
metabolism
through
CYP26a1
expression.

 
 Because
 inadequate
 zinc
 is
 so
 widely
 prevalent
 in
 the
 developed
 and
 under‐ developed
 world
 (Prasad,
 2003;
 Song
 et
 al.,
 2009),
 it
 is
 important
 to
 understand
 the
 implications
 of
 zinc
 on
 the
 function
 of
 other
 nutrients.
 Since
 vitamin
 A
 has
 many
 crucial
 functions
 within
 the
 body
 and
 RA
 is
 used
 as
 a
 cancer
 treatment
 (Bohnsack
 and
 Hirschi,
 2004;
Mark
et
al.,
2006),
an
understanding
of
the
implications
of
zinc
status
on
vitamin
A
 function
is
important,
especially
if
zinc
manipulation
can
alter
the
sensitivity
of
cancer
cells
 to
RA
treatment.
 
 3.2.

Limitations
 There
are
several
limitations
apparent
in
my
thesis
research.
The
first
limitation
is
 the
use
of
an
in
vitro
system.


Cell
systems
do
not
replicate
the
complexity
of
whole
body
 systems,
therefore
it
is
difficult
to
extrapolate
the
results
of
this
project.
 
 The
 second
 limitation
 is
 the
 method
 of
 zinc
 depletion.
 In
 order
 to
 mimic
 low‐zinc
 conditions,
zinc
was
removed
by
treating
the
FBS
with
Chelex‐100
resin.
The
filtration
steps
 required
during
preparation
of
the
low
zinc
FBS
may
have
removed
small
peptides
which
 could
potentially
be
important
in
cellular
processes.
Also,
it
is
possible
that
zinc
within
the
  
  65
  FBS
 bound
 to
 low
 molecular
 weight
 proteins
 was
 removed
 by
 the
 Chelex‐100
 resin.
 This
 zinc
was
subsequently
replaced
by
free
zinc.
Free
zinc
and
zinc‐bound
to
proteins
may
have
 different
availability
to
the
cell.

 
 Thirdly,
the
treatment
durations
were
relatively
short‐term,
with
a
maximum
of
72
 h.
This
is
likely
not
reflective
of
a
clinical
treatment
situation,
where
treatment
durations
 are
weeks
to
months
(Huang
et
al.,
1988).



 
 Finally,
the
cell
cycle
analysis
experiment
was
performed
only
once.
Although
each
 treatment
 consists
 of
 the
 analysis
 of
 10,000
 individual
 cells,
 a
 repeat
 of
 the
 experiment
 would
strengthen
the
observation.

 
 3.3.

Future
Directions
 I
examined
the
gene
expression
of
the
RAR
and
RXR
receptors
in
response
to
zinc
 manipulation.
However,
the
level
of
these
receptors
does
not
necessarily
indicate
a
change
 in
function.
Therefore,
it
would
be
valuable
to
investigate
the
effects
of
zinc
status
on
the
 DNA‐binding
ability
of
these
zinc‐finger
proteins
using
protein‐DNA
binding
assays
such
as
 the
 electromobility
 shift
 assay
 (EMSA),
 which
 is
 used
 to
 detect
 protein
 complexes
 with
 nucleic
 acids
 (Hellman
 and
 Fried,
 2007).
 This
 process
 was
 described
 by
 Ho
 and
 Ames
 (2002).
 I
 had
 attempted
 to
 perform
 an
 EMSA,
 however
 could
 not
 obtain
 useable
 results
 due
to
technical
challenges.
Alternatively,
a
new
method
of
assessing
protein‐DNA
binding,
 negative‐ion
electrosparay
ionization
mass
spectrometry,
is
described
by
Park
et
al.
(2011).

  
  66
  Since
 zinc
 has
 wide
 ranging
 effects
 on
 gene
 expression,
 it
 is
 plausible
 that
 other
 proteins
involved
in
the
metabolism,
transport
and
function
of
vitamin
A
may
be
affected
 by
zinc
status.
Therefore
it
would
be
interesting
to
investigate
the
effect
of
zinc
status
on
 other
 vitamin
 A
 related
 proteins,
 including
 transport
 proteins
 such
 as
 CRABPII,
 which
 is
 necessary
for
the
transport
of
RA
into
the
nucleus
(Donato
et
al.,
2007).

 
 RA
 as
 a
 cancer
 treatment
 has
 toxic
 side
 effects
 (Fabricius
 et
 al.,
 2011).
 It
 may
 be
 beneficial
 to
 research
 the
 effects
 of
 zinc
 on
 RA‐induced
 growth
 suppression
 in
 other
 cancerous
and
non‐cancerous
cell
types
including
human
hepatic
and
extrahepatic
cells,
as
 well
 as
 whole
 body
 animal
 models.
 Increasing
 the
 sensitivity
 of
 cancerous
 cells
 or
 decreasing
 the
 sensitivity
 of
 non‐cancerous
 cells
 to
 the
 effects
 of
 RA
 during
 cancer
 treatment
could
be
an
attractive
method
of
attenuating
the
toxic
effects
of
this
treatment
 regime
or
could
perhaps
postpone
the
onset
of
resistance.
 
 
 
 
 
 
 
 
 
  
  67
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  78
  
 
 
 
 
 
 
 
 
  Appendices
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
  
 
 
  79
  Appendix
1
  
 
 
 
 Table
A.1:

Mineral
supplements
added
to
Chelex‐100
treated
medium.

 
  
 
 
 
  
  
  Mineral
 
  Weight
(mg/L
medium)
  Ca(CH3COO)2.H20
 
  50.06

  KH2PO4
  3.845
  MgSO4
  10.94
  MnCO3
  0.0027
  MoO3
  0.0009
  CuCO3.H20
 
  0.0275
  FeC6H5O7
  0.0175
  
 
 
 
 
 
  
  80
  Appendix
2
 
 
 
 
 
 Table
A.2:

Primer
Sequences
for
real
time
quantitative
PCR

 Gene

  Primer
  Reference
  RARα
  Forward:
5’‐CTG
TGA
GAA
ACG
ACC
GAA
AC‐3’
 Reverse:
5’‐TTG
TCC
CCA
GAG
GTC
AAT
GTC‐3’
  Wei
et
al.,
2007
  RARβ
  Forward:
5’GGA
ACG
CAT
TCG
GAA
GGC
T‐3’
 Reverse:
5’‐TTC
CCA
GCC
CCG
AAT
CAT‐3’
  Wei
et
al.,
2007
  RARγ
  Forward:
5’‐ACC
AGG
AAT
CGC
TGC
CAG
TA‐3’
 Reverse:
5’TGG
GCT
TTG
CTG
ACC
TTG
G‐3’
  Wei
et
al.,
2007
  RXRα
  Forward:
5’‐GAC
AAG
CGG
CAG
CGG
AAC‐3’
 Reverse:
5’‐CTC
TCC
ACC
GGC
ATG
TCC
T‐3’
  Wei
et
al.,
2007
  RXRβ
  Forward:
5’‐GGG
GTG
CGA
AAA
GAA
ATG‐3’
 Reverse:
5’‐CGG
GGT
TTG
TTG
TTC
TCC‐3’
  Wei
et
al.,
2007
  RXRγ
  Forward:
5’‐GAG
GAC
GAT
AAG
GAA
GGA
C‐3’
 Reverse:
5’AAG
CGA
CTT
CTG
ATA
GCG‐3’
  Wei
et
al.,
2007
  CYP26a1
  Forward:
5’‐TGG
TAC
TGC
AGC
GGA
GGA‐3’
 Reverse:5’‐GCG
TCT
TGT
AGA
TGA
AGC
CGT
A‐3’
  Deng
et
al.,
2003
  Cyclophilin
  Forward:
5’‐ACG
GCG
AGC
CCT
TGG
 Reverse:
5’‐TTT
CTG
CTG
TCT
TTG
GGA
CCT‐3’
  Deng
et
al.,
2003
  
 
 
 
  
 
 
  81
  Appendix
3
 
 
 
 
 
 
 Table
A.3:

Western
Blot
antibody
dilutions
and
incubation
times
 Protein
 RARα
  Primary
Ab

 (dilution)
 Rabbit
 polyclonal
 (1:400)
  RXRα
  Rabbit
 polyclonal
 (1:400)
  Actin
  Mouse
 monoclonal
 (1:400)
  Primary
Ab
 incubation

 Overnight
at



 4
°C
in
2.5%
 skim
milk
 powder
in
TTBS
 1
hour
at
room
 temperature
in
 2.5%
skim
milk
 powder
in
TTBS
 Overnight
at



 4
°C
in
2.5%
 skim
milk
 powder
in
TTBS
  Secondary
Ab
 (dilution)
 HRP
conjugated
 goat
anti‐rabbit
 (1:10,000)
  Secondary
Ab
 incubation
 45
min
at
room
 temperature
in
 1%
skim
milk
in
 TTBS
 HRP
conjugated
 45
min
at
room
 goat
anti‐rabbit
 temperature
in
 (1:10,000)
 1%
skim
milk
in
 TTBS
 HRP
conjugated
 45
min
at
room
 goat
anti‐ temperature
in
 mouse
 1%
skim
milk
in
 (1:10,000)
 TTBS
  
 
 
 
 
 
 
 
  
  82
  Appendix
4
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 Figure
 A.1:
 Total
 cellular
 zinc
 levels
 in
 HepG2
 and
 AML12
 cells.
 HepG2
 and
 AML12
 cells
 were
 cultured
 into
 10
 cm
 plates
 at
 initial
 seeding
 densities
 of
 300,000
 and
 200,000
 cells
 respectively,
 for
 6
 d.
 HepG2
 were
 cultured
 in
 MEM
 and
 AML12
 were
 cultured
 in
 F12/DMEM
1:1
(Gibco,
Grand
Island,
NY)
containing
10%
FBS,
sodium
pyruvate
(110
mg/L),
 sodium
bicarbonate
(1.2
g/L),
insulin‐transferin‐selenium
(Invitrogen;
1%),
Hepes
buffer
(15
 mM),
penicillin/streptomycin
(5,000
U/L),
and
dexamethasone
(40
ng/ml)
At
the
end
of
the
 culture
 period,
 cells
 were
 harvested
 and
 total
 cellular
 zinc
 was
 measured
 by
 atomic
 absorption
 spectrophotometry.
 Values
 represent
 mean
 ±
 SEM
 (n=3;
 the
 experiment
 was
 repeated
once).
Asterisk
indicates
a
significant
difference
between
AML12
and
HepG2
cells
 (p<0.05).

 
 
  
  83
  Appendix
5
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 Figure
A.2:
Cell
proliferation
of
AML12
cells
treated
with
retinoic
acid.
AML12
cells
were
 cultured
 into
 96‐well
 plates
 at
 an
 initial
 seeding
 density
 of
 5000
 cells/well
 in
 F12/DMEM
 1:1
medium
and
allowed
to
grow
for
2
d.
Cells
were
then
treated
with
all‐trans
retinoic
acid
 (RA)
at
0
(DMSO
only),
15,
25
and
35
µM
for
12
h.
Cell
proliferation
was
quantified
using
 the
BrdU
colorimetric
assay
with
absorbance
as
a
measure
of
BrdU
incorporation.
Values
 represent
 mean
 ±
 SEM
 (n=6;the
 experiment
 was
 repeated
 once).
 Means
 with
 different
 letters
are
significantly
different
(p<0.05).
 
 
 
 
 
  84
  Appendix
6
 
 
 
  
 
 
 
 
 
 
 
 Figure
A.3:
The
effect
of
retinoic
acid
on
viability
of
AML12
cells
following
growth
in
low‐,
 adequate‐,
and
high‐zinc
medium.
AML12
cells
were
cultured
into
10
cm
plates
at
an
initial
 density
of
200,000
cells
in
F12/DMEM
1:1
medium.
Cells
were
allowed
to
attach
overnight.

 Subsequently,
the
cells
were
cultured
in
medium
containing
Chelex
100‐treated
FBS
plus
0
 (Low),
5
(Adequate),
and
10
(High)
µM
zinc.
On
day
5,
cells
were
transferred
into
96‐well
 plates
at
a
density
of
5,000
cells
per
well,
and
on
day
6,
cells
were
treated
with
all‐trans
 retinoic
acid
(RA)
at
0
(DMSO
only)
or
35
µM
for
72
h.
Cell
viability
was
quantified
using
the
 MTT
 colorimetric
 assay
 with
 absorbance
 as
 a
 measure
 of
 cell
 viability.
 Values
 represent
 mean
 ±
 SEM
 (n=12).
 Means
 with
 different
 letters
 are
 significantly
 different.
 Asterisks
 represent
 a
 significant
 difference
 between
 RA‐treatment
 and
 its
 respective
 control.
 (p<0.05).

 
 
 
  
  85
  Appendix
7
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 Figure
A.4:
The
effect
of
RA
on
the
proliferation
of
AML12
cells
following
growth
in
low‐,
 adequate‐,
and
high‐zinc
medium.
AML12
cells
were
cultured
into
10
cm
plates
at
an
initial
 density
of
200,000
cells
in
F12/DMEM
1:1
medium.
Cells
were
allowed
to
attach
overnight.

 Subsequently,
the
cells
were
cultured
in
medium
containing
Chelex
100‐treated
FBS
plus
0
 (Low),
5
(Adequate),
and
10
(High)
µM
zinc.
On
day
4,
cells
were
transferred
into
96‐well
 plates
at
a
density
of
5,000
cells
per
well,
and
on
day
6,
cells
were
treated
with
all‐trans
 retinoic
 acid
 (RA)
 at
 0
 (DMSO
 only)
 or
 35
 µM
 for
 12
 h.
 Cell
 proliferation
 was
 quantified
 using
 the
 BrdU
 colorimetric
 assay
 with
 absorbance
 as
 a
 measure
 of
 BrdU
 incorporation.
 Values
represent
mean
±
SEM
(n=6).
Means
with
different
letters
are
significantly
different.
 Asterisks
 represent
 a
 significant
 difference
 between
 RA‐treatment
 and
 its
 respective
 control.
(p<0.05).

  
  86
  

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