University of Southern California Dissertations and Theses

Protein phosphatase 2A and annexin A5: modulators of cellular functions

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Protein phosphatase 2A and annexin A5: modulators of cellular functions

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PROTEIN PHOSPHATASE 2A AND ANNEXIN A5:
MODULATORS OF CELLULAR FUNCTIONS

By

Chia-Ling Hsieh

A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(GENETIC, MOLECULAR AND CELLULAR BIOLOGY)
May 2013

Copyright 2013 Chia-Ling Hsieh
ii

Dedication

To my parents and family
This is for you.

To Han-Yu
I couldn’t have done this without you.

To Julie, Pheony, Gi Gi, Luby, and Dottie
My cutest life companies.

iii

Acknowledgements
I would like to, first and foremost, acknowledge the support of my advisor, Dr. Jeannie
Chen. None of this work would have been possible without her guidance and inspiration
through countless discussions. Dr. Chen always brought in boundless energy and brilliant
thinking to stimulate fresh and insightful views on research problems. Her enthusiasm
and dedication to science encouraged me to tackle the obstacles in research once more.
I want to show my great appreciation to my committee members, Dr. Ralf Langen and Dr.
David Hinton for their invaluable suggestions, criticism, and spiritual influence as
passionate scientists during various committee meetings. In addition, I want to thank
Robabeh Mohammadzedeh for her assistance in cardiac morphology identification and
functional analysis.
I would also like to thank all my colleagues in the Chen lab for providing me with
amiable environment and support in these years. I want to express my heartily
appreciation to Dr. Brain Soreghan and Dr. Mario Isas for their encouragement and
suggestion on my research. In particular, I want to thank Yun Yao and Helen He for their
contribution and assistance to my experiments. I also thank Dr. Wen Mao, Dr. Hormoz
Moaven, and Tian Wang for their useful discussion, feedbacks, and friendship.
Lastly, I want to thank my parents for teaching me the appreciation for education and
knowledge as well as supporting me to pursue and live all my dreams. Finally, I thank my
husband Han-Yu for always standing by me and believing in me more than I can ever do.
Without all my family’s endless support, I would never complete this achievement.
iv

Table of Contents
Dedication іі
Acknowledgements ііі
List of Figures vііі
Thesis Outline x
Abstract xі
Chapter 1: Overview of Photoreceptors and Phototransduction Cascade 1
1.1 Retina and Photoreceptor Cells 1
1.2 Photodransduction Cascade in Rod Photoreceptor 5
1.3 Phototransduction Deactivation 7
1.3.1 Dark adaptation 7
1.3.2 Retinoid cycle 8
1.4 Rhodopsin Phosphorylation and Arrestin-1 Binding 9
1.4.1 Rhodopsin phosphorylation 9
1.4.2 Arrestin-1 binding 10
1.5 Rhodopsin Dephosphorylation 13
1.5.1 Possible phosphatases for rhodopsin dephosphorylaiton 13
1.5.2 Protein phosphatase 2A (PP2A) 14

Chapter 2: The Role of Arrestin-1 in Protein Phosphatase 2 Mediated Rhodopsin
Dephosphorylation 20
2.1 Introduction 20
2.2 Materials and Methods 24
2.2.1 Isoelectric focusing (IEF) electrophoresis 24
v

2.2.2 Western blot analysis and quantification 25
2.2.3 Rod outer segment preparation 26
2.3 Results 27
2.3.1 Fostriecin can efficiently inhibit rhodopsin dephosphorylation 27
2.3.2 Arrestin-1 may play a positive role in regulating rhodopsin
dephosphorylation 30
2.3.3 Arrestin-1 has no effect in regulating the expression of PP2A subunits 32
2.3.4 Most of PP2A subunits are membrane associated 32
2.3.5 Arrestin-1 may regulate the distribution of PP2A subunits in rod outer
segments 34
2.4 Discussion 36
2.5 Summary 40

Chapter 3: Overview of Alzheimer’s Disease 41
3.1 Alzheimer’s Disease - the Most Extensively Studied Amyloid-Related
Disorders 41
3.2 Amyloidogenesis- the Generation of β-Amyloid (Aβ) Peptide 43
3.2.1 The function of amyloid precursor protein (APP) 43
3.2.2 Proteolysis process of APP 44
3.3 Amyloid Protein Aggregation 48
3.4 Aβ oligomers, the Pathological Agent in Alzheimer’s Disease 49
3.5 Neuronal Impact of Soluble Aβ Oligomers 52
3.5.1 Calcium homeostasis disruption 53
3.5.2 Generation of reactive oxygen species (ROS) 54
3.5.3 Membrane peroxidation 55
vi

3.5.4 Tau as a downstream target of Aβ oligomers 56
3.6 A Potential Protecting Factor against AD, Annexin A5 57

Chapter 4: Annexin A5 Protects Heart Function in a Mouse Model for
Alzheimer’s Disease 60
4.1 Introduction 60
4.2 Materials and methods 63
4.2.1 Transgenic animals 63
4.2.2 Trichrome staining 64
4.2.3 Sample preparation for ultrastructure examination 65
4.2.4 Electrocardiogram (ECG) recording 66
4.2.5 Western blot analysis 66
4.2.6 Cryosection and Immunocytochemistry 67
4.2.7 Mouse perfusion 67
4.2.8 Thioflavin-S staining 68
4.2.9 Western blot for β-amyloid protein 68
4.3 Results 70
4.3.1 The body weight of APP-PS1/ANXA5KO female mice is
significantly less 70
4.3.2 APP-PS1/ANXA5KO mice have low survival rate especially in female 72
4.3.3 Abnormal cardiac morphology show in young ANXA5KO and APP-
PS1/ANXA5KO mice 72
4.3.4 Diffused Z-lines and disorganized myofilaments are found in APP-
PS1/ANXA5KO mice 75
vii

4.3.5 APP-PS1/ANXA5KO mice show lower heart rate and longer
QT interval at young age 77
4.3.6 Lower heart rate and higher QT interval are found in ANXA5KO
and APP-PS1/ANXA5KO mice 79
4.3.7 The deficiency of heart in APP-PS1/ANXA5KO mice is not
caused by cardiac hypertrophy 81
4.3.8 The membrane associated αB-crystallin are increased in ANXA5KO and
APP-PS1/ANXA5KO mice 83
4.4 Discussion 85
4.5 Summary 91
4.6 Supplementary Data 92
4.6.1 APP-PS1/ANXA5KO mice do not have more β-amyloid plaques in brain 92
4.6.2 Older APP-PS1/ANXA5KO mice have more monomeric Aβ and amyloid
oligomers in brain as compared to APP-PS1 mice 94
Conclusions 97
Bibliography 100

viii

List of Figures

Figure 1-1 Retina structure and neuron cells 2
Figure 1-2 Structure of vertebrate photoreceptors 4
Figure 1-3 Schematic of the activation steps of the vertebrate phototransduction
cascade 6
Figure 1-4 Structure of protein phosphatase 2A 16
Figure 2-1 Fostriecin efficiently inhibits rhodopsin dephosphorylation 29
Figure 2-2 Rhodopsin dephosphorylation is delayed in arrestin-1
knockout mice 31
Figure 2-3 Most of PP2A subunits are expressed in arrestin-1 knock out retina 33
Figure 2-4 PP2A subunits are membrane associated under both dark and
light condition 33
Figure 2-5 The A subunit of PP2A dramatically leaves ROS under light
stimulation in arr1-/- mice 35
Figure 3-1 Proteolytic processing of amyloid precursor protein (APP) 46
Figure 4-1 The body weight of APP-PS1/ANXA5KO female mice is significantly
lower than wild type mice 71
Figure 4-2 Early sudden death of APP-PS1/ANXA5KO (APP/ANXKO) mice,
especially in the female group 73
Figure 4-3 Abnormal cardiac morphology in young ANXA5KO and APP-PS1/
ANXA5KO (APP/ANXKO) mice 74
Figure 4-4 Ultrastructure of cardiomyocytes in three months-old wildtype,
APP-PS1, ANXA5KO and APP-PS1/ANXA5KO mice 76
Figure 4-5 Lower heart rate and higher QT interval are found in
APP-PS1/ANXA5KO mice 78
ix

Figure 4-6 Both ANXA5KO and APP-PS1/ANXA5KO mice show lower
heart rate and longer QT interval in ECG recording 80
Figure 4-7 The comparison of heart weight/body weight ratio between APP-PS1/
ANXA5KO and other mice 82
Figure 4-8 Increased membrane associated αB-crystallin expression in young
ANXA5KO and APP-PS1/ANXA5KO mice 84
Figure 4-9 Both APP-PS1 and APP-PS1/ANXA5KO mice show numbers of
amyloid plaques in brain at early age 93
Figure 4-10 APP-PS1/ANXA5KO mice have more monomeric Aβ and amyloid
aggregations in brain at elder age 95

x

Thesis Outline
In this thesis, my colleagues and I concentrated on understanding how proteins
modulating cellular functions. We try to understand of the mechanism of rhodopsin
dephosphorylation and the role of annexin A5 in pathological progress of Alzheimer’s
disease by employing a combination of genetic, biochemistry, electrophysiological, and
histological techniques.
In chapter 2, the investigation was focused on a hypothesis that arrestin-1 protein
positively regulates rhodopsin dephosphorylation. PP2A is confirmed as the rhodopsin
phosphatase in this study. By using arretin-1 knockout mice, the relationship of arrestin-1
and PP2A in rhodopsin dephosphorylation was examined. The electrophoresis and
biochemistry analysis implied that arrstin-1 modulates rhodopsin dephosphorylation by
regulating the distribution of PP2A subunit in rod outer segments.
In chapter 4, APP-PS1/ANXA5KO mouse was generated to study the function of annexin
A5 in pathogenic progress of amyloid-related disease. Early death occurring in this model
was noticed first. Meanwhile, biochemical and eletrocardiogram analysis revealed severe
structural damages, functional disorders, and abnormal protein distribution in
cardiomyocytes. This study presents a novel discovery that annexin A5 is critical in
protecting cardiomyocytes from amyloid toxicity.

xi

Abstract
In the past several years, I focused on two different projects for my PhD studies. The first
project is to achieve a better understanding of the mechanism of rhodopsin
dephosphorylation. Protein phosphatase 2A (PP2A) has been recognized as the
phosphatase responsible for rhodopsin dephosphorylation for years. However, due to the
absence of in vivo evidence, the role of PP2A in regulating rhodopsin regeneration is still
questionable. This work not only clarifies the position of PP2A in modulating rhodopsin
function under physiological condition, but also presents novel findings in the detail
mechanism. Previous isoelectric focusing results revealed that arrestin-1 may play a
positive role in modulating rhodopsin dephosphorylaiton. Taking advantage of arrestin-1
knockout mice, we isolated rod outer segments to compare how arrestin-1 influences the
distribution of PP2A subunits after light stimulation. Western blot results revealed that
the movement of PP2A scaffolding subunit is regulated by arrestin-1 protein. In addition
to inactivating rhodopsin, arrestin-1 may mediate rhodopsin dephosphorylation by
modulating the cellular localization of PP2A in rod photoreceptors.
Identifying the role of annexin A5 in amyloidogenesis is the goal of my second project.
Annexin A5 is an abundant protein without clear physiological function. Earlier studies
suggested a protection effect of annexin A5 in against amyloid toxicity. We generated
APP-PS1/ANXA5KO mice by breeding APP-PS1dE9, a widely used mouse model for
Alzheimer’s disease, under annexin A5 knockout background. No overt difference in the
number of brain Aβ plaques were seen between the APP-PS1dE9 and the APP-
xii

PS1/ANXA5KO mice. Interesting, sudden death was noticed in APP-PS1/ANXA5KO
mice at early age. Morphology shown in trichrome staining and ultrastructural
examination revealed severe damages in cardiomyocytes from ANXA5KO and APP-
PS1/ANXA5KO mice. These two strain of mice also exhibited lower heart rates and
higher QT interval in electrocardiogram analysis. Furthermore, the level of αB-crystallin,
a chaperon molecule, is significantly increased in the membrane fraction of cardiac
extraction, and a specific perinuclear distribution of αB-crystallin is also observed by
immunocytochemistry in ANXA5KO and APP-PS1/ANXA5KO mice. Therefore,
annexin A5 may play an important role in regulating cardiac function in Alzheimer’s
disease model mice.

1

Chapter 1
Overview of Photoreceptors and Phototransduction Cascade

1.1 Retina and Photoreceptor Cells
Vision formation is a very complicated but interesting process. Light enters our eyes,
goes through lens, and reaches retina locating at the back of eye ball. Retina, around 0.4
millimeters thick, is a highly organized neural tissue composed of several cell layers to
convert light stimulation into electrical signals that are then transmitted to the visual
cortex for image production. These layers are comprised of six types of cells: retinal
epithelium cells, photoreceptors, bipolar cells, ganglion cells, horizontal cells, and
amacrine cells (Figure 1-1). Photoreceptors, lying at the outer region of retina, are the
places where phototransduction starts. In vertebrate, there are two kinds of photoreceptor,
rods and cones, named after their segment shapes. In particular, rods have extreme
sensitivity to response to a single photon. At brighter light levels, rods are saturated and
cones take over light detection. Due to the different light absorbing pigments, rods and
cones have different spectral sensitivity. Visible light ranges from blue (short-), green
(medium-), to red (long-wavelength) radiation. Rhodopsin, the pigment in rods, has
maximal spectra (λm) sensitivity at 500nm, whereas human blue, green and red opsins in
cones are most sensitive to short- (430nm), medium- (535nm) and long-wavelength
(560nm) light, respectively.
2

A A A
A
B
Figure 1-1 Retina structure and neuron cells.
(A) Retina can be recognized as several specialized layers. They are RPE: retinal pigment epithelium;
OS: outer segment; IS: inner segment; ONL: outer nuclear layer, fiber layer; OPL: outer plexiform
layer; INL: inner nuclear layer; IPL: inner plexiform layer; GCL: ganglion cell layer, and optic fiber
layer. (B) There are six types of cells composing these layers: retinal epithelium cells (pigmented
cells), photoreceptors (rods and cones), bipolar cells, ganglion cells, horizontal cells, and amacrine
cells.

3

Structurally, each photoreceptor has four parts: an outer segment, an inner segment, a cell
body, and a synaptic ending (Figure 1-2). Outer segment is a specialized region for
phototransduction in both rods and cones, while inner segment contains cellular
organelles such as the Golgi complex, endoplasmic reticulum, and mitochondria
responsible for molecular synthesis and maintenance of cells. Cell body containing
nucleus and synapse is the place voltage-gated ion channels regulate the release of
neurotransmitter into synaptic terminal. Most vertebrates are rod dominant, accounting
for about

97% of all photoreceptors, and cones contribute for the rest. Because of the
abundance of rods, they not only mediate phototransduction, but also play a significant
role in maintaining the overall structural integrity of retina. Rod outer segment, located
at the distal surface of retina, consists of a stack of membrane discs formed by
internalization of the plasma membrane. These membrane discs contain the light-
absorbing protein, rhodopsin; thus outer segment is the site phototransduction taking
place. Rhodopsin is a prototypical seven transmembrane protein which belongs to the
large G-protein couple receptor (GPCR) family. It is composed of two parts: 11-cis-
retinal chromophore, the light-sensitive pigment, and apoprotein opsin. The 11-cis-retinal
chromophore is covalently attached to Lys-296 (K296) (Robinson, Cohen et al. 1992) on
opsin with a protonated Schiff base linkage. Structural wise, the abundance of rhodopsin
plays an important role in structural maintaining for rod photoreceptors. Meanwhile,
rhodopsin is the initiator of phototransduction cascade; therefore it is also critical for the
function of rod photoreceptors.

4

Figure 1-2 Structure of vertebrate photoreceptor.
Each photoreceptor can be divided into four structural compartments: outer segment, inner
segment, nucleus, and synaptic terminal. Outer segments in both rods and cones are separated
from inner segments by connecting cilium. The major morphological difference between rod and
cone is from the structural shape of outer segments.
5

1.2 Phototransduction Cascade in Rod Photoreceptor
Rod photoreceptors use a G protein cascade including G protein coupled receptor
(rhodopsin), G-protein (transducin), and downstream effector (PDE), to convert and
amplify light signal into electric response (Figure 1-3). Light stimuli initiates
phototransduction by activating rhodopsin, the G-protein coupled receptor locating on the
outer segment membrane. Rhodopsin contains the photon-sensitive chromophore, 11-cis-
retinal, which isomerizes into all-trans retinal upon photon absorption. This conversion
results in a conformational change in rhodopsin and makes it transform to the active state
known as Metarhodopsin II (R*). R* then binds to downstream G protein, transducin, and
catalyzes the exchange of GTP for GDP on the transducin alpha subunit which then
dissociates from the beta/gamma subunits and rhodopsin. Next, the activated transducin
alpha subunit excites cGMP-phosphodiesterase (PDE) to hydrolyze cGMP into GMP. As
a result, a decrease concentration of intracellular cGMP closes cGMP-gated channels and
thus reduces the concentration of Na
+
and Ca
2+
inside the cell. Consequently, rod
photoreceptor becomes hyperpolarized, leading to the closure of downstream voltage-
gated Ca
2+
channels and a decrease in glutamate release in the synaptic cleft. Because this
amplified response persists as long as rhodopsin remains catalytically active (Nakatani
and Yau 1988, Chen, Makino et al. 1995, Sagoo and Lagnado 1997, Rieke and Baylor
1998), timely deactivation of rhodopsin is required for unambiguous detection of
subsequent photon absorptions.

6

Figure 1-3 Schematic of the activation steps of the vertebrate phototransduction cascade.
Upon light stimulation (hv), rhodopsin becomes activated and binds to downstream G protein,
transducin. This interaction catalyzes the exchange of GTP for GDP and induces the dissociation of
transducin α subunit. The activated transducin α then turns on the catalytic activity of
phosphodiesterase (PDE) to hydrolyze cGMP into GMP. This hydrolysis decreases the intracellular
concentration of cGMP, resulting in the close of cyclic nucleotide channels. An influx of Na
+
and Ca
2+

is terminated as a consequence of channel close which causes the hyperpolarization of the membrane
potential. (Figure adapted from Lamb and Pugh 2000)
7

1.3 Phototransduction Deactivation
1.3.1 Dark adaptation
The termination of phototransduction cascade and the restoration of activated proteins
involved are essential for the recovery of photoreceptor back into a state ready to absorb
next photon; this process is known as “dark adaptation”. Deactivating rhodopsin and
regenerating 11-cis-retinal are two important events in dark adaptation. The light-
activated rhodopsin is deactivated by incorporation of multiple phosphates on a cluster of
serine and threonine residues at its C terminus (Wilden and Kuhn 1982, Thompson and
Findlay 1984, Sibley, Strasser et al. 1986, Palczewski and Benovic 1991) by rhodopsin
kinase (GRK1) and the binding of arrestin-1 protein. In mice, the calcium-bound protein,
recoverin, is bound to GRK1 under dark condition to inhibit its enzyme activity on the
light-triggered rhodopsin. As the Ca
2+
concentration decreases due to the closure of
cGMP-gated channel under light stimulation, recoverin is freed of calcium and
dissociates from GRK1. This allows GRK1 to phosphorylate rhodopsin. The process of
phosphorylation alone not only decreases the catalytic activity of light-activated
rhodopsin by at least 50% (Chen, Makino et al. 1995, Sagoo and Lagnado 1997, Xu,
Dodd et al. 1997, Chen, Burns et al. 1999), but also prepares rhodopsin for high-affinity
arrestin-1 binding (Gurevich and Gurevich 2004). The following arrestin-1 binding is
required for complete turnoff (Wilden, Hall et al. 1986, Xu, Dodd et al. 1997) and
prevention of further interaction with transducin. The arrestin-1 binding then causes
subsequent release of all-trans retinal and induce rhodopsin dephosphorylation. This two-
8

step process makes rhodopsin in its chromophore-free state, opsin, and awaits binding of
a new chromophore to get ready for further light absorption termed rhodopsin
regeneration.
When rhodopsin is regenerated to response next photon, available transducin and PDE
are required to carry out this cascade again. In mice, the deactivation of activated
transducin-PDE complex involves a few steps. The activity of transducin alpha subunit is
terminated when bound GTP is hydrolyzed into GDP. Although transducin has intrinsic
slow GTPase activity, a GAP complex made up of RGS9, Gβ5, and R9AP speeds up
GTP hydrolysis. The hydrolysis of GTP deactivates transducin alpha and dissociates it
from PDE, which terminates cGMP hydrolysis.

1.3.2 Retinoid cycle
As previously discussed, rhodopsin regeneration is the counterpart to rhodopsin
deactivation as the essential steps of dark adaptation. After deactivation of rhodopsin, it
must be reconstituted with a new 11-cis-retinal chromophore to restore photosensitivity.
The regeneration process is known as retinoid cycle, or visual cycle, in which the all-
trans-retinal on deactivated rhodopsin is replaced by new 11-cis-retinal. The retinoid
cycle has four steps: photochemistry, removal of retionoid, reconversion of retinoid, and
delivery of retinoid (Lamb and Pugh 2004). Upon photon absorption, 11-cis-retinal is
photoisomerized to all-trans retinal, and activates rhodopsin. After rhodopsin is
deactivated by C-terminal phosphorylation and arrestin-1 binding, all-trans retinal is
9

reduced into all-trans retinol by all-trans retinol dehydrogenase (RDH) either in rod outer
segment cytoplasm or when still non-covalently bound to opsin. All-trans retinol is then
guided to RPE by inter-photoreceptor retinoid binding protein (IRBP). Within RPE,
cellular retinol binding protein (CRBP) leads all-trans retinol to lecithin retinol acyl
transferase (LRAT) for esterification. RPE65 then guides now all-trans retinyl ester to
become isomerized into 11-cis retinol by retinyl ester isomerohydrolase. 11-cis retinol is
further oxidized to 11-cis retinal by 11-cis RDH with the guidance of cellular
retinaldehyde binding protein (CRALBP). Finally, IRBP directs 11-cis retinal to diffuse
back into rod outer segment disc membrane and covalently binds to opsin to regenerate
photosensitive rhodopsin.

1.4 Rhodopsin Phosphorylation and Arrestin-1 Binding
1.4.1 Rhodopsin phosphorylation
The first step of phototransduction termination is the deactivation of light-stimulated
rhodopsin by GRK1 phosphorylation. Most mammals have seven GRK subtypes, of
which GRK1 and GRK7 are specific for photoreceptors. GRK1 expresses in both rods
and cones, and GRK7 presents exclusively in cones (Lorenz, Inglese et al. 1991) (Weller,
Virmaux et al. 1975, Shichi and Somers 1978, Weiss, Raman et al. 1998). GRK1 was
shown to be crucial for timely signal shutoff in both types of photoreceptors (Cideciyan,
10

Zhao et al. 1998, Chen, Burns et al. 1999). There are three serine and three threonine
residues located at the C-terminus of rhodopsin in mice. The importance of these residues
on rhodopsin phosphorylation has been revealed by electrophysiological recordings with
rhodopsin mutated transgenic mice carrying different numbers of available
phosphorylation sites at C-terminus (Chen, Makino et al. 1995, Mendez, Burns et al.
2000). The minimal number of phosphorylations required to inactivate the photoresponse
in a timely manner is three and all six residues are needed for following recovery.
Moreover, the serine residues located on the C-terminus of rhodopsin are phosphorylated
in a highly ordered pattern (Kennedy, Lee et al. 2001). Phosphorylation starts with
Ser343, the residue most distant from the plane of membrane, then Ser338 and finally
the most proximal Ser334. Studies showed that the rate of phosphorylation of each
residue correlates with its mobility and the frequency of collisions with kinase (Langen,
Cai et al. 1999, Kennedy, Lee et al. 2001). Thus, residues located at the C-terminal tip are
more likely to become phosphorylated than residues at the C-terminal base. With the
same rule, rhodopsin dephosphorylation follows the same pattern as phosphorylation
(Ser343 → Ser338 → Ser334).

1.4.2 Arrestin-1 binding
Next, the residual activity of rhodopsin is completely terminated by arrestin-1 binding.
Arrestin-1, the first member of the arrestin family, was discovered twice: first as S-
antigen causing uveitis (Wacker, Donoso et al. 1977), then as a 48kDa protein that binds
11

light-activated rhodopsin (Kuhn, Hall et al. 1984). In mammals, there are four arrestin
proteins, arrestin-1 or visual arrestin, arrestin-4 or cone arrestin, arrestin-2 and arrestin-3.
Arrestin-1 expressed at very high levels in both rods (Strissel, Sokolov et al. 2006, Song,
Vishnivetskiy et al. 2011) and cones (Nikonov, Brown et al. 2008), whereas the cone-
specific arrestin-4 (Murakami, Yajima et al. 1993, Craft, Whitmore et al. 1994)
constitutes only about 2% of the total arrestin complement in cone photoreceptors
(Nikonov, Brown et al. 2008). Arrestins are a small family of proteins that regulate G
protein-coupled receptors. Arrestins specifically bind to phosphorylated receptors,
terminating G protein coupling, targeting receptors to endocytotic vesicles, and initiating
G protein-independent signaling. Receptor-bound arrestins not only serve as an adaptor
for internalization through binding to clathrin and AP2 but also function as a scaffold
protein to promote the formation of multiprotein signaling complexes (Ferguson,
Downey et al. 1996, Goodman, Krupnick et al. 1998, Laporte, Oakley et al. 1999, Perry
and Lefkowitz 2002). In contrast to arrestin-2 and -3, arrestin-1 does not have a clathrin-
binding site localized in the C-terminus (Goodman, Krupnick et al. 1996), and therefore
does not effectively support rhodopsin internalization.
Studies showed that arrestin-1 specifically binds to activated phosphorylated rhodopsin
(p-R*) with 10-12 times higher affinity as compared to non-activated phosphorylated (p-
R) and activated unphosphorylated rhodopsin (R*) (Gurevich and Benovic 1993). A
model of sequential multiple interactions between arrestin-1 and rhodopsin was proposed
to explain the binding selectivity
12, 36, 37
. Two regions of arrestin-1 are involved, a
phosphorylation sensor (residues 49-90, β-strands V and VI and adjacent loops) in the N-
12

domain and an activation sensor (residues 237-268, β-strands XV and XVI) in C-domain.
Simultaneous engagement of both sensors triggers a global conformation change of
arrestin-1 and transits it from inactive state into the high-affinity receptor-binding state.
Therefore, when rhodopsin is both activated and phosphorylated, arrestin-1 can recognize
and interact with rhodopsin to terminate rhodopsin’s activity. On the other hand, the
mechanism of arrestin activation by receptor-attached phosphates suggests that arrestin
cannot be particularly sensitive to the sequence context of phosphorylated residues.
Indeed, both under in vitro (Vishnivetskiy, Raman et al. 2007) and in vivo (Mendez,
Burns et al. 2000) conditions, arrestin-1 requires three phosphates on rhodopsin for high-
affinity binding, but not particular on which residues out of serines and threonines. The
nonphosphorylated residues can function as additional targets for rhodopsin kinase
(GRK1) to increase the binding efficiency with activated rhodopsin, thus accelerating
rhodopsin turnoff (Caruso, Bisegna et al. 2010).

13

1.5 Rhodopsin Dephosphorylation
1.5.1 Potential phosphatases for rhodopsin dephosphorylaiton
Posttranslational modification of proteins by kinases and phosphatases plays an important
role in the regulation of cellular signaling especially in neurochemistry. This also applies
to vertebrate photoreceptors where phosphorylation of rhodopsin causes uncoupling from
the signal transduction cascade. The activity of rhodopsin is regained after substitution of
the bleached photopigment 11-cis-retinal and by dephosphorylation of the opsin moiety.
For other G protein coupled receptors, the binding of arrestin recruits AP2 and clathrin to
form a complex which initiates endocytosis and the removal of G protein coupled
receptor from cell membrane. In endosomes, the receptor molecules are
dephosphorylated and recycled. Finally, the G protein coupled receptor is sent back to
cell membrane and ready for the next signal transduction (Sorkin and Von Zastrow 2002).
However, unlike other G protein coupled receptors, rhodopsin is not endocytosed under
normal condition in vertebrate (Young and Bok 1969, Basinger, Bok et al. 1976). That
means the dephosphorylation of rhodopsin has to be carried out at the disk membrane.
Since dephosphorylation has been known as a key step in recycling of rhodopsin, many
studies have focused on searching the corresponding phosphatase. Studies in Drosophila
provided a possible candidate, RDGC (Steele and O'Tousa 1990, Steele, Washburn et al.
1992), but the elimination of rdgc homologs in mice has no effect in preventing
rhodopsin dephosphorylation (Ramulu, Kennedy et al. 2001). On the other hand,
Palczewski et al. (Palczewski, Hargrave et al. 1989) found that the extracts from rod
14

outer segments have the ability to dephosphorylate phospho-opsin in a concentration-
dependent manner, and this phosphatase activity was later characterized originating from
PP2A by DEAE-cellulose chromatography and HPLC gel permeation chromatography.
More in vitro data all suggested the possibility of PP2A being as the rhodopsin
phosphatase (Fowles, Akhtar et al. 1989, Palczewski, Hargrave et al. 1989, King,
Andjelkovic et al. 1994). Therefore, people generally believe that rhodopsin
dephosphorylation is carried by PP2A function.

1.5.2 Protein phosphatase 2A (PP2A)
Reversible protein phosphorylation is an essential regulatory mechanism in many
biological processes. There are three structurally distinct families of serine/threonine
protein phosphatase that have been classified, and protein phosphatase 2A (PP2A), which
belongs to the PPP family, is the major serine/threonine phosphatase involved in many
cellular functions such as cell cycle regulation, cell growth control, development,
cytoskeleton dynamics, and cell mobility (Janssens and Goris 2001), as well as a tumor-
suppressor protein (Janssens, Goris et al. 2005). PP2A is a heterotrimeric protein which
consists of a 65 kDa scaffolding A (PR65) subunit, a 36 kDa catalytic C subunit, and a
variable regulatory B subunit (Figure 1-4). A core enzyme consisting of only A and C
subunits has been reported to contain physiological functions and are abundantly existing
in cells. However, the prevalent PP2A enzymes are ABC heterotrimers in vivo. It is
believed that individual subunits do not exist as stable monomers in vivo. The A and C
15

subunits are each encoded by two highly related and widely expressed genes. The C
subunit of PP2A is among the most conserved subunit in species ranging from yeast to
mammal. To gain full activity of protein phosphatase, one of the variable regulatory
subunits needs to join the PP2A core enzyme to form a heterotrimeric holoenzyme. The
regulatory subunit is composed by four different families: B (B55 or PR55), B’ (B56 or
PR61), B’’ (PR48/PR72/PR130), and B’’’ (PR93/PR110), and there are at least 16
members in these families (Virshup 2000, Janssens and Goris 2001, Lechward,
Awotunde et al. 2001, Xu, Xing et al. 2006).
16

Figure 1-4 Structure of protein phosphatase 2A.
PP2A is a holoenzyme containing three major components: a scaffolding subunit (A subunit), a
regulatory subunit (B subunit), and a catalytic subunit (C subunit). In mammalian, A and C subunits
are both encoded by two genes (α and β). The regulatory B subunit can be classified into four different
families: B, B’, B”, and B’’’. The B/PR55 subunits are encoded by four related genes (α, β, γ, and δ).
The B’/PR61 family contains five variant genes (α, β, γ, δ, and ε). The B” family has three related
genes, encoding PR72, PR130, and PR59 or PR48. PR93 (SG2NA) and PR110 (striatin) are the two
components of the B’’’ family. (Figure adapted from Janssens and Goris 2001)
17

Except the B’’’ subunits, direct interactions between the PP2A core enzyme and the
regulatory subunits have been demonstrated. There is no detectable sequence homology
among the four families of regulatory subunits, and the expression levels of various
regulatory subunits are highly diverse depending on cell types and tissues. For example,
the regulatory subunits which belong to B” subfamily are specifically expressed in heart
and muscle. The B”’ subfamily contains neuron-specific proteins and the ones from B or
B’ subfamilies are widely expressed. In this regard, the B subunits of PP2A appear to
have a number of functions such as regulating PP2A activity, determining substrate
specificity of PP2A, and targeting PP2A to distinct intracellular locations. The B subunits
show different binding affinities to the core enzyme, and their interaction modulates the
catalytic activity of PP2A. Despite the fact that PP2A is primary a serine/threonine
phosphatase, it can function as a phosphotyrosyl phosphatase (PTPase) under certain
conditions. PTPA, an activator of the PTPase activity of PP2A, has been cloned and
characterized (Cayla, Van Hoof et al. 1994). B subunits may differentially inhibit PTPA-
induced PP2A catalytic activity (Cayla, Van Hoof et al. 1994). In addition to modulate
PP2A activity, the variety of B subunit also greatly influences substrate specificity. A
unique feature of the B subunit is their ability to compete and replace each other in the
PP2A holoenzyme. Therefore, changing the ratio between different B subunit carried
holoenzymes has a great influence on substrate selection (Garcia, Cereghini et al. 2000).
For example, a PP2A complex containing a 72kDa (PR72) regulatory subunit
dephosphorylates casein kinase I sites on SV40 large T antigen, and PP2A with a 55kDa
regulatory B subunit dephosphorylates cyclin-dependent kinase sites (Cegielska, Moarefi
18

et al. 1994). Moreover, numbers of studies indicate that distinct B subunits are
differentially expressed and distributed in tissue and cells, therefore, their unique
distribution certainly affects the targeting of PP2A (Sontag 2001). For instance, B56γ1
directs PP2A to paxillin at focal adhesions (Ito, Kataoka et al. 2000), and an isoform of
the 55kDa B subunits, Bα, is responsible for the microtubule targeting of PP2A (Sontag,
Nunbhakdi-Craig et al. 1995, Sontag, Nunbhakdi-Craig et al. 1999). Lastly, the
regulatory subunits of PP2A may function as receptors of second messengers (Csortos,
Zolnierowicz et al. 1996, McCright, Rivers et al. 1996, Zolnierowicz, Van Hoof et al.
1996, Strack, Chang et al. 1999, Xing, Xu et al. 2006). Several studies have suggested
that the lipid ceramide activates PP2A through the B subunit.
Besides the regulation carried by the B subunit mentioned above, the catalytic activity
and substrate specificity of PP2A complex are also modulated by the post-translational
modifications of the C subunit. There are two phosphorylation sites (Thr304, Tyr307) and
a methylation site (Leu309) locating at the C-terminal of the catalytic subunit (Sontag
2001). The phosphorylation of both threonine and tyrosine residues is associated with
PP2A inactivation and the modification is reversed by its unique ability to catalyze
intramolecular autodephosphorylation (Guo, Reddy et al. 1993). Leu309 is methylated by
methyltransferase (De Baere, Derua et al. 1999) and demethylated by a methylesterase,
PME-1 (Ogris, Du et al. 1999). Although the methylation on the C subunit increases
PP2A activity (Favre, Zolnierowicz et al. 1994), these effects are indirect (De Baere,
Derua et al. 1999). On the other hand, these modifications of C subunit significantly
affect the binding of B subunit, and modulate the functional specificity of PP2A. It has
19

been reported that deletion or mutations of the Leu309, Thr304, and Tyr307 residues
dramatically reduce the binding of Bα to AC in vitro and in cells, and alter substrate
specificity (Ogris, Gibson et al. 1997, Bryant, Westphal et al. 1999).
Although PP2A has been recognized as the phosphatase responsible for rhodopsin
dephosphorylation with lots of supports from in vitro evidence, the certainty is not
secured yet. One of the concerns is that the opsin phosphatase activity might come from
other compartment of rod cell or other cells in retina because of the contamination during
rod outer segment preparation. On the other hand, the detail mechanism of rhodopsin
dephosphorylation is still not clear today. Therefore, experiments done with physiological
conditions are required to verify the role of PP2A in rhodopsin dephosphorylation, and
the dephosphorylation process needs to be uncovered as well.

20

Chapter 2
The Role of Arrestin-1 in Protein Phosphatase 2A Mediated Rhodopsin
Dephosphorylation

2.1 Introduction
Rod photoreceptors use a G protein cascade to convert the signal of single absorbed
photon into electric response. Absorption of a photon induces a conformational change
and activates the G protein coupled receptor rhodopsin, which catalyzes the exchange of
GTP for GDP on the G protein transducin. Transducin-GTP in turn, activates cGMP
phosphodiesterase (PDE) to hydrolyze cGMP and reduces the intracellular cGMP
concentration. This makes cGMP-gated channels to close and hyperpolarize the cell.
Because this amplified response persists as long as rhodopsin remains catalytically active
(Nakatani and Yau 1988, Chen, Makino et al. 1995, Sagoo and Lagnado 1997, Rieke and
Baylor 1998), timely deactivation of rhodopsin is required for unambiguous detection of
subsequent photon absorptions.
Two critical events, phosphorylation and arrestin-1 binding, participate in the
deactivation of these G protein coupled receptors. In rod photoreceptors, the
phosphorylation of light-activated rhodopsin by rhodopsin kinase (GRK1) initiates
deactivation and limits the response amplitude (Chen, Makino et al. 1995, Sagoo and
Lagnado 1997, Chen, Burns et al. 1999). Like other G protein coupled receptors, there is
21

a cluster of serine and threonine residues which can undergo stimulus-dependent
phosphorylation (Wilden and Kuhn 1982, Thompson and Findlay 1984, Sibley, Strasser
et al. 1986, Palczewski and Benovic 1991) at the carboxyl terminus of rhodopsin. The
process of phosphorylation alone not only decreases the catalytic activity of light-
activated rhodopsin by at least 50% (Xu, Dodd et al. 1997), but also prepares rhodopsin
for high-affinity arrestin-1 binding (Gurevich and Gurevich 2004). Arrestin-1, or visual
arrestin, is the first member identified in the arrestin family. It was discovered twice: first
as S-antigen causing uveitis (Wacker, Donoso et al. 1977), then as a 48kDa protein that
binds light-activated rhodopsin (Kuhn, Hall et al. 1984). Shortly thereafter, Pfister et al
(Pfister, Chabre et al. 1985) established that both are one and the same protein. The
following arrestin-1 binding is required for the complete shutoff of light-activated
rhodopsin (Wilden, Hall et al. 1986, Xu, Dodd et al. 1997).

Endocytosis is a mechanism used to recycle G protein coupled receptors. After the C
terminal phosphorylation, arrestin binds to G protein coupled receptors and recruits AP2
and clathrin to form a complex which can initiate endocytosis (Sorkin and Von Zastrow
2002). In endosomes, phosphatase removes phosphate molecules and finally G protein
coupled receptors are regenerated and sent back to cell membrane. However, this
restoration mechanism is not applied on rhodopsin. In contract to arrestin-2 and -3,
arrestin-1 does not have a clathrin-binding site which can support rhodopsin
internalization at the C-terminus (Goodman, Krupnick et al. 1996). Instead of occurring
22

in endosomes, the rhodopsin dephosphorylation has to take place in the outer segment
membrane which is a unique mechanism as compared to the regular endocytosis.
The phosphatase working for rhodopsin dephosphorylation is still unclear. The
suggestion has been made that the vertebrate rhodopsin phosphatase may be related to
RDGC, a Ca
2+
-dependent phosphatase that dephosphorylates Drosophila rhodopsin
(Steele, Washburn et al. 1992, Vinos, Jalink et al. 1997). However, the mammalian
homologue of RDGC was identified but it appears to be localized primary to rod inner
segments (Sherman, Sun et al. 1997). Ramulu et al. (Ramulu, Kennedy et al. 2001)
deleted the two rdgC homologs, PPEF-1 and PPEF-2, in mice but found that the
elimination of PPEF function has no effect in preventing rhodopsin dephosphorylation.
On the other hand, the activity that dephosphorylates rhodopsin has characteristics of
protein phosphatase 2A (PP2A). Both PP1 and PP2A have been detected in rod outer
segment preparations but only PP2A recognizes and dephosphorylates rhodopsin (Fowles,
Akhtar et al. 1989, Palczewski, Hargrave et al. 1989). PP2A holoenzyme has been
purified from photoreceptors as a catalytic subunit and two larger polypeptides (King,
Andjelkovic et al. 1994).
Protein phosphatase 2A (PP2A) is predominantly found as a heterotrimeric holoenzyme
consisting of a catalytic subunit (C), a scaffolding subunit (A), and one member of 4
families of regulatory subunits (B). PP2A-catalyzed dephosphorylation of target substrate
protein is widespread and critical for cellular functions. PP2A is very diverse in
vertebrates, and this is achieved by expression of two C subunits, two A subunits, and
23

approximately sixteen B subunits. The B subunits are derived from four diverse gene
families (B, B’, B’’, and B’’’) that have little sequence similarity between families but
maintain high sequence similarity within families (Slupe, Merrill et al. 2011). It is
believed that the variation of the B subunit is responsible for substrate targeting and
binding specificity of PP2A.
The detail mechanism of rhodopsin dephosphorylation is still unclear to date. Although
PP2A has been identified as the possible enzyme for rhodopsin dephosphorylation, the
conclusion is still questionable because of the results are from in vitro experiments. Here,
the whole intact retina was used to test the function of PP2A in rhodopsin
dephosphorylation with the treatment of PP2A inhibitors. On the other hand, arrestin-1
has been suggested to inhibit rhodopsin dephosphorylation due to the occupation of
phosphorylated residues (Palczewski, McDowell et al. 1989). In order to test this
hypothesis, we used arrestin-1 knock mice to study the role of arrestin-1 in regulating
rhodopsin dephosphorylation.

24

2.2 Materials and Methods
2.2.1 Isoelectric Focusing (IEF) electrophoresis
Mice were dark adapted for overnight and whole retinas were dissected out under
infrared light. Retinas were incubated with 5 μM fostriecin in Ame’s bicarbonate buffer
(8.9g/L Ame’s medium, Sigma; 1.9g/L sodium bicarbonate) at 37 ℃ for 1 hour in dark.
Retinas were exposed to intensive light for 10 minutes and transferred back to dark again
for recovery. Then retinas were homogenized in 400 µl of homogenization buffer
[ 25mM Hepes (pH7.5), 100mM EDTA, 50mM NaF, 5 mM adenosine, 1mM PMSF, and
0.5mg/ml protease inhibitor cocktail (Roche Inc)]. Samples were centrifuged at 19,000 x
g for 15 minutes to separate membrane fraction. Membrane fraction was rinsed with
10mM Hepes (pH7.5) and resuspended in 1ml of regeneration buffer [ 10mM Hepes
(pH7.5), 0.1mM EDTA, 1mM MgCl
2
, 50mM NaF, 5mM adenosine, 1mM PMSF,
2%BSA, 100µM 9-cis retinal, and 0.5mg.ml protease inhibitor cocktail (Roche Inc)] for
regeneration at 4 ℃ overnight. After regeneration with 9-cis retinal, membrane fractions
were collected by centrifugation at 19,000 x g for 20 minutes and solubilized in 100µl of
solubilization buffer [ 20mM Hepes (pH7.5), 1mM MgCl
2
, 10mM NaCl, 0.1mM EDTA,
50mM NaF, 5 mM adenosine, 1mM PMSF, 1% dodecyl-maltoside (Calbiochem), 1mM
DTT, and 0.5mg/ml protease inhibitor cocktail (Roche Inc)] at 4℃ for overnight. The
solubilized samples were centrifuged again at 19,000 x g for 20 minutes. The supernatant
was collected and 5µl of each sample was loaded on an IEF gel after mixing 1:1 with
glycerol. Then isoelectric focusing electrophoresis was performed on pH gradient of 3-8
25

gel (with 0.04M glutamic acid as the anode solution and 1.0M NaOH as the cathode
solution) as described before(Adamus, Arendt et al. 1993). The gel was run at a constant
23W on a Pharmacia Flat Bed Apparatus FBE300 at 10 ℃ for 2 hours. The proteins were
then transferred to nitrocellulose membrane and phosphorylated rhodopsin species were
detected by Western blot using monoclonal antibody 4D2. For the experiment of
rhodopsin dephosphorylation in arrestin-1 knockout mice, indicated mice were dark
adapted overnight and exposure to diffuse white light (5000lux intensity at cage level) for
1 hour after pupils were dilated with 0.5% tropicamide and 2.5% phenylephrine
hydrochloride. Then mice were transferred to dark room for recovery. Retinas were taken
at different time point and prepared for IEF as described above.

2.2.2 Western blot analysis and quantification
Indicated mice were dark adapted overnight and exposed to 5000 luxs light with pupil
dilation for 1 hour. Retinas were taken from these mice and retinal proteins were
extracted with non-detergent [80mM Tris (pH8.0), 4mM MgCl
2
, and 0.5mg/ml protease
inhibitor cocktail (Roche Inc)] or 1% Triton X-100 containing buffer [80mM Tris
(pH8.0), 4mM MgCl
2
, 1% Triton X-100, and 0.5mg/ml protease inhibitor cocktail (Roche
Inc)]. One dimensional PAGE was carried out and proteins were transferred onto
nitrocellulose membranes. The blots were saturated in TBS-T buffer [20mM Tris (pH7.5),
136.8mM NaCl, and 0.1% Tween 20] containing 5% non-fat milk. The following primary
antibodies were used for Western blot: rabbit polyclonal anti-PP2A A subunit (1:500,
26

Cell signaling), mouse monoclonal anti-PP2A B subunit (1:1000, Cell signaling), mouse
monoclonal anti-PP2A C subunit (1:2000, Cell signaling), rabbit polyclonal anti-PP2A
B’-beta subunit (1:1000, courtesy of Dr. David Virshup), rabbit polyclonal anti-PP2A B’-
delta subunit (1:1000, courtesy of Dr. David Virshup), rabbit polyclonal anti-PP2A B’-
epsilon subunit (1:1000, courtesy of Dr. Cheryl Craft), and rabbit polyclonal anti-Gβ5
(1:2000, Caltech). The secondary antibodies of IRDye 680 goat anti-mouse and IRDye
800 goat anti-rabbit antibodies (LI-COR Biosciences, Lincoln NE) were applied
accordingly. The proteins were visualized and quantified using Odyssey Infrared Imaging
System (LI-COR, Biosciences).

2.2.3 Rod outer segment preparation
The detail process has been described before(Nickell, Park et al. 2007). 10 retinas were
collected from indicated mice for each sample. Retinas were vortexed in 120 µl of 8%
opti-prep/Ringers buffer (OptiPrep density gradient medium, Sigma; Ringers: 130mM
NaCl, 3.6mM KCl, 2.4mM MgCl
2
, 1.2mM CaCl
2
, 10mM Hepes, 0.02mM EDTA, pH7.4,
osmolarity at 313 mosM) at full speed for 2 minutes. After a quick spin at 510 x g for 1
minute, 100 µl of supernatant was taken out and another 100µl of 8% opti-prep buffer
was added. Pellets were resuspended again, and repeat this process for five times to
collect 500 µl of supernatant. Then the supernatant was loaded to the top of an opti-prep
gradient which is made by mixing 1.4 ml of 10% opti-prep/Ringers buffer with 1.5 ml of
18% opti-prep/Ringers buffer. After centrifuging at 70,000 rmp (Beckman TLA100) at
27

4℃ for 1 hour, carefully take out the outer segments and add 3X volume of chilled
Ringer’s buffer. Samples were centrifuged again at 50,000 rmp (Beckman TLA100) at
4℃ for another 20 minutes. Pellets were washed with ice cold Ringer’s buffer and kept at
-80℃ for later use.

2.3 Results
2.3.1 Fostriecin can efficiently inhibit rhodopsin dephosphorylation.
There are six serine/threonine residues functioning as phosphorylation sites on the C
terminus of mouse rhodopsin, and the different phosphorylated rhodopsin species can be
easily separated by isoelectric focusing (IEF) electrophoresis because of the difference of
PI value. Retinas from C57 mice were incubated with fostriecin first and used to monitor
rhodopsin dephosphorylation. Dark sample shows no rhodopsin phosphorylation as
expected. Light stimuli triggers phototransduction cascade and rhodopsin molecules are
partially inactivated by GRK1 catalyzed C-terminal phosphorylation. The
dephosphorylation of rhodopsin starts within 30 minutes of dark recovery and rhodopsin
are fully dephosphorylated after 1 hour in control group. However, this
dephosphorylation process is significantly inhibited with the treatment of fostriecin. As
compared to control group, even after 30minute of dark recovery, the phosphorylation
status of rhodopsin is still maintained (Fig. 2-1). Fostriecin is known as a specific
28

inhibitor of PP2A (Walsh, Cheng et al. 1997, Lewy, Gauss et al. 2002). Therefore, we
further confirm that PP2A may be the phosphatase responsible for rhodopsin
dephosphorylation.

29

Figure 2-1 Fostriecin efficiently inhibits rhodopsin dephosphorylation.
Retinas were first incubated with or without fostriecin to see the subsequent dark recovery after light
exposure. The different phosphorylated level of rhodopsin is shown by isoelectric focusing (IEF).
Rhodopsin is fully phosphorylated after light stimulation and the dephosphorylation process starts
within 30 minute in the control group. Rhodopsin is completely dephosphorylated after 1 hour dark
recovery. The rhodopsin phosphorylation also shows in the retina with the treatment of fostriecin.
However, in the same group, the dephosphorylation of rhodopsin is significantly inhibited even after
1 hour dark recovery.

30

2.3.2 Arrestin-1 may play a positive role in regulating rhodopsin dephosphorylation
Since PP2A may play a role in rhodopsin dephosphorylation, the next question is how it
participates in this process. Previously, some studies showed that arrestin-1 may inhibit
rhodopsin dephosphorylation due to its binding to phosphorylation sites on the C-
terminus of rhodopsin (Palczewski, McDowell et al. 1989). In order to test this
hypothesis, we used arrestin-1 knock mice as model to study the dephosphorylation of
rhodopsin under the absence of arrestin-1. In the IEF results, rhodopsin in both C57 and
arrestin-1 knockout mice are phosphorylated after light exposure. The inactivated
rhodopsin is fully dephosphorylated after 2 hour dark recovery in the control group.
However, in arrestin-1 knockout mice, it takes almost 4 hours to reach the same status
(Fig 2-2). Here, we found that the absence of arrestin-1 causes the delay of rhodopsin
dephosphorylation. This clearly points out that arrestin-1 may play a positive role in
regulating rhodopsin dephosphorylation.

31

Figure 2-2 Rhodopsin dephosphorylation is delayed in arrestin-1 knockout mice.
Retinas from WT (W) and arrestin-1 knockout mice (A) were stimulated with light and recovered
under dark for different time points. The level of rhodopsin phosphorylation is also analyzed and
shown by IEF. Rhodopsin is fully phosphorylated in arr1-/- mice as compared to WT under light
exposure. After 2 hour dark adaptation, rhodopsin is completely dephosphorylated in WT mice.
However, most of the rhodopsin molecules are still phosphorylated in arr1-/- at the same time point.
After 4 hours, rhodopsin is almost dephosphorylated in arr1-/- mice.

32

2.3.3 Arrestin-1 has no effect in regulating the expression of PP2A subunits
We have demonstrated the positive effect of arrestin-1 and the function of PP2A in
rhodopsin dephosphorylation. Our next question is whether arrestin-1 works as a
regulator for PP2A in rod photoreceptors. Therefore, we first compared the retinal
expression of PP2A subunits in arrestin-1 knock and C57 mice. From the Western blot
results, most of the PP2A subunits except the B’-beta subunit are equally expressed in
both arrestin-1 knock out and C57 mice (Fig. 2-3). This indicates that arrestin-1 has no
effect in regulating the expression of PP2A subunits in retina.

2.3.4 Most of PP2A subunits are membrane associated
Under normal condition, rhodopsin is not endocytosed. This suggests that protein
phosphatases have to go to membrane discs for rhodopsin dephosphorylation. In order to
test this hypothesis, the retinal proteins are separated into cytoplasmic and membrane
associated fractions according to the detergent solubility. Abundant protein levels of
PP2A subunits are found in the membrane fraction under both dark and light stimulated
condition (Fig. 2-4). Meanwhile, this membrane localization of PP2A is not changed in
arrestin-1 knockout mice. Here, we found that most of the PP2A subunits are membrane
associated. Neither the stimulation of light nor the absence of arrestin-1 in retina has an
effect on changing the membrane localization of PP2A subunits.
33

Figure 2-3 Most of PP2A subunits are expressed in arrestin-1 knock out retina.
Retinal proteins were extracted from arrestin-1 knock out (A) and control mice (W), and equal
amount of each sample was loaded. Actin is used as the loading control here. Most subunits of PP2A
we checked here are abundantly expressed in retina except B’-beta subunit. The absence of arrestin-1
in retina has no effect on the protein expression and there is no expression difference between arr1-/-
and control mice as well.

Figure 2-4 PP2A subunits are membrane associated under both dark and light condition.
Retinas were taken from arr1-/- and wild type mice and proteins were separated into cytoplasmic (C)
and membrane fraction (M) base on the detergent solubility. Equal amount of proteins were loaded in
each sample. All the PP2A subunits we checked here are located in the membrane fraction, and neither
the absence of arrestin-1 nor the stimulation of light has effect on determination of the membrane
associated distribution.

34

2.3.5 Arrestin-1 may regulate the distribution of PP2A subunits in rod outer segments
Arrestin-1 has been reported to move from rod inner segment to outer segment upon light
stimuli (Broekhuyse, Tolhuizen et al. 1985, Broekhuyse, Janssen et al. 1987, Philp,
Chang et al. 1987, Mangini and Pepperberg 1988, Mendez, Lem et al. 2003). In order to
see whether arrestin-1 modulates the localization of PP2A subunits in rod outer segment
(ROS), we isolated dark adapted and light exposed rod outer segments from arrestin-1
knock out and C57 mice for Western blot. We evaluated the purity of ROS by checking
Western blot image of Gβ5. The G protein β subunit, Gβ5, is predominantly expressed in
the central nervous system. A splicing variant of Gβ5, Gβ5 long form (Gβ5L), was
identified in retina that contains an N-terminal 42 amino acid extension. Furthermore, it
was found that retinal Gβ5L, but not Gβ5, was present in the ROS membranes (Watson,
Aragay et al. 1996). In the same approach, we found out that our purified ROS contains
the Gβ5L only, while both the short and long forms were detected in the whole retinal
homogenates (data not shown). This indicated that there is no contamination from other
cell compartments. Light stimulation makes most of PP2A subunits except the A subunit
leave ROS in both arrestin-1 knock and C57 mice (Fig. 2-5A). Averagely, about 60-70%
of the B, C, B’-delta, and B’-epsilon subunit are left in light exposed ROS as compared to
dark condition. However, the A subunit moves out ROS in arrestin-1 knockout mice
under light condition and there is only about 40% of the A subunit left, rather than the
90% in C57 mice (Fig. 2-5B). Therefore, arrestin-1 may specifically regulate the
localization of PP2A A subunit in ROS.
35

Figure 2-5 The A subunit of PP2A dramatically leaves ROS under light stimulation in arr1-/-
mice.
(A) Rod outer segments (ROS) were isolated from light-stimulated (L) or control (dark, D) retinas
in indicated mice. Gβ5 long form (Gβ5L) is used as the indicator of ROS purity and the loading
control as well. Light stimulation makes most of the PP2A subunits leaving ROS except the A
subunit in wild type mice. The amount of the A subunit is even less in the arr1-/- ROS as
compared to wild type under light condition. (B) Quantification data (n=4) indicate that 60-80% of
the B, C, B’-delta, and B’-epsilon subunits are left in light-stimulated ROS in both arr1-/- and wild
type mice. Most of the A subunit leaves ROS in the absence of arrestin-1 protein. There is only
40% of the A subunit left in light exposed ROS in arr1-/- mice, rather than the 95% in wild type
mice.

36

2.4 Discussion
Reversible phosphorylation plays an important role in modulating many cellular
processes. Many protein kinases have been identified and their regulation mechanisms
are characterized as well. However, the kinase antagonist, phosphatase, is less understood.
Protein phosphatase 2A (PP2A) is a heterotrimeric holoenzyme which consists of three
different subunits. Among other enzymes, PP2A accounts for as much as 1% of total
cellular proteins and for the major portion of serine/threonine phosphatase activity in
most tissues and cells (Cohen 1997). PP2A has been identified as the possible
phosphatase for rhodopsin dephosphorylation in vitro (Fowles, Akhtar et al. 1989,
Palczewski, Hargrave et al. 1989, King, Andjelkovic et al. 1994), and here we further
confirm the role of PP2A in rhodopsin dephosphorylation by using fostriecin, a known
PP2A specific inhibitor, and intact retina to closely reflect the real physiological
condition.
Arrestin-1 is the protein that binds to the phosphorylated rhodopsin and completely turns
off the activity of light-activated rhodopsin. Besides deactivating rhodopsin, the
suggestion has been made that arrestin-1 may inhibit rhodopsin dephosphorylation
(Palczewski, McDowell et al. 1989). However, the process of rhodopsin
dephosphorylation is significantly slowed down in arrestin-1 knockout mice (Fig. 2-2).
Therefore, we hypothesize that arrestin-1 may play a positive role in regulating rhodopsin
dephosphorylation. Due to the absence of clathrin binding motif, arrestin-1 and AP2
along is not enough to support endocytosis, a mechanism used to recycle inactivated G
37

protein coupled receptors through dephosphorylation, for rhodopsin. Therefore,
rhodopsin phosphatase should be localized in or close to cell membrane. Although
arrestin-1 does not affect the retinal expression of PP2A, while comparing the
distribution of PP2A subunits between cytoplasm and membrane, we show that most of
the PP2A subunits are membrane associated under dark and light condition (Fig. 2-4). It
does conform to the requirement of membrane localization for rhodopsin phosphatase.
Photoreceptors are neurons with a unique design: they have a specialized signaling
compartment, the outer segment, connected to the rest of the cells via a narrow cilium.
This provides an opportunity to modulate the gain of signaling by changing the effective
concentrations of proteins such as arrestin-1 and transducin, through moving them in and
out of OS, rather than by changing their expression like most other cells do (Gurevich,
Hanson et al. 2011). In figure 2-5, we showed that most of PP2A subunits (B, C, and B’)
except A subunit leave ROS under light stimulation in C57 mice. Recently, a signal-
mediated, transient cytoplasm-membrane translocation and subsequent activation of a yet
unidentified PP2A holoenzyme have just been reported (Ludowyke, Holst et al. 2000).
Brown et al (Brown, Carlson et al. 2002) also proposed a model of PP2A
dephosphorylating phosducin and rhodopsin by translocating subunits between cytoplasm
and membrane. All these support a hypothesis that rod cells may also regulate the effect
of PP2A by changing subunits’ concentration in ROS. On the other hand, in arrestin-1
knock mice, there is only about 40% of A subunit left in ROS as compared to C57. It
seems like that the ability to keep the ROS localization of A subunit is lost in the absence
of arrestin-1 protein. The A subunit functions as the scaffolding part to provide the
38

interacting base for regulatory and catalytic subunits. We hypothesize that keeping the A
subunit in ROS by arrestin-1 provide an available base which can interact with the B and
C subunits to form a functional PP2A holoenzyme for rhodopsin dephosphorylation in
short time. Without the presence of A subunit in ROS, the regeneration of a PP2A
holoenzyme in ROS will take more time. This also explains the reason that the rhodopsin
dephosphorylation is much slower in arrestin-1 knockout mice. Moreover, the substrate
targeting specificity is insured as well through the proper localization of the A subunit in
ROS.
The modulation of PP2A subcellular localization has been widely reported and some
studies show that the protein-protein interactions between PP2A and other intracellular
components contribute to the specificity of PP2A signaling (Sontag 2001). For instance,
the interaction with paxillin, vimentin, tubulin, and CG-NAP directs selective PP2A
holoenzyme to discrete cellular domains, i.e. focal adhesions (Ito, Kataoka et al. 2000),
intermediate filaments (Turowski, Myles et al. 1999), microtubules (Sontag, Nunbhakdi-
Craig et al. 1995, Sontag, Nunbhakdi-Craig et al. 1999), and the Golgi apparatus
(Takahashi, Shibata et al. 1999), respectively. In addition, PP2A interacts with proteins
that target the phosphatase to specific cellular regulatory functions, such as the PP2A
complex comprising PR48 and Cdc6 regulates initiation of DNA replication(Yan,
Fedorov et al. 2000). Thirdly, the selective association of PP2A with scaffolding proteins
directs the phosphatase to specific signaling modules. For example, PP2A regulates Wnt
signaling through associating with a multi-protein complex containing axin, APC, β-
catenin, and GSK-3β (Kikuchi 1999). All these studies strongly support the idea that
39

PP2A can be targeted to restricted intracellular microenvironments through protein-
protein interactions (Sontag 2001).
Non-visual arrestins were reported to interact with hundreds of non-receptor signaling
proteins (Xiao, McClatchy et al. 2007), ranging from components of the internalization
machinery, such as clathrin (Goodman, Krupnick et al. 1996), AP2 (Laporte, Oakley et al.
1999), and N-ethylmaleimide-sensitive factor (NSF) (McDonald, Cote et al. 1999) to
MAP kinases activating JNK3 (McDonald, Chow et al. 2000), ERK1/2 (Luttrell,
Roudabush et al. 2001), and p38 (Bruchas, Macey et al. 2006) and several E3 ubiquitin
ligases (Bhandari, Trejo et al. 2007, Ahmed, Zhan et al. 2011). As compared to other
arrestin, arrestin-1 has been reported to exclusively interact with rhodopsin. However,
polymerized tubulin, was discovered as non-receptor binding protein for arrestin-1 in
2004 (Nair, Hanson et al. 2004). Later, more proteins such as MAP kinases JNK3 (Song,
Raman et al. 2006), ERK2 (Coffa, Breitman et al. 2011), Ca
2+
-liganded calmodulin (Wu,
Hanson et al. 2006), E3 ubiquitin ligases Mdm2 (Song, Raman et al. 2006, Hanson,
Cleghorn et al. 2007), parkin (Ahmed, Zhan et al. 2011), and NSF (Huang, Brown et al.
2010) as well as enolase (Smith, Bolch et al. 2011) were identified as interacting partners
with arrestin-1. Therefore, arrestin-1 may modulate the localization of PP2A by protein-
protein interaction and the future studies will focus on elucidating the molecular basis of
this mechanism.

40

2.5 Summary
The activity of rhodopsin, a G protein coupled receptor on rod outer segment, is turnoff
by C-termial phosphorylation and the binding of arrestin-1 protein. Protein phosphatase
2A (PP2A) has been reported as the rhodopsin phosphatase from the results of many in
vitro experiments. Arrestin-1, a critical protein in the terminating of light response in
vertebrate rod photoreceptor, has been suggested to negatively regulate rhodopsin
dephosphorylation. However, the detail mechanism of rhodopsin dephosphorylation is
still not fully understood today. In this study, we show that PP2A is the phosphatase
responsible for rhodopsin dephosphorylation and arrestin-1 function as a positive
regulator in this dephosphorylation process. Moreover, most of PP2A subunits leave ROS
upon light stimulation except the A subunit, and the localization of PP2A scaffolding
subunit (A subunit) in rod outer segment is modulated by arrestin-1. These results
establish a hypothesis that the movement of PP2A subunits into ROS is a mechanism
used to regulate rhodopsin dephosphorylation by efficiently changing the functional
concentration of this phosphatase. Arrestin-1 keeps the A subunit in ROS to provide a
base for immediately binding of the B and C subunits at proper timing. This mechanism
not only regenerates the functional herterotrimeric holoenzyme in short time but also
insures the targeting specificity of PP2A in ROS.

41

Chapter 3
Overview of Alzheimer’s Disease

3.1 Alzheimer’s Disease - the Most Extensively Studied Amyloid-Related Disorders
Alzheimer’s disease (AD) is the most common age-associated dementia that leads to
cognitive, memory and behavioral impairments in elder population. Current estimates
suggest that it may affect more than 12 million individuals worldwide (Ferri, Prince et al.
2005). The pathological characterists of AD patients include the extracellular
accumulation of 38 to 43 amino-acid β-amyloid (Aβ) peptides in senile plaques, the
presence of intracellular hyperphosphorylated tau in neurofibrillary tangles, synaptic loss
and neuronal degeneration. Plaques and tangles are present mainly in brain regions
involved in learning, memory and emotional behaviors such as the entorhinal cortex,
hippocampus, basal forebrain and amygdale. These affected regions that typically exhibit
reduced numbers of synapses and neuritis are usually damaged, hence it suggests that
synapses and neuritis are the targets of Aβ (Mattson 2004). Due to the high prevalence of
AD, it is one of the most extensively studied amyloid-related disorders. However, the
lack of effective treatments and the uncertain mechanisms of pathogenesis are still the
nightmare of AD patients.
Medical studies estimate that only about 3 percent of all Alzheimer cases, which
translating into 200,000 people in the U.S., are early-onset familial Alzheimer’s disease
42

(FAD). Symptoms of FAD can start before a person’s sixties. FAD is caused by
autosomal dominant mutations in amyloid precursor protein (APP), presenilin 1 (PSEN1),
and presenilin 2 (PSEN2) genes locating on chromosomes 21, 14, and 1, respectively.
Mutations found in APP either increase Aβ production (Haass, Lemere et al. 1995) or
generate highly fibrillogenic Aβ variants (Vetrivel and Thinakaran 2006). PSEN1 and
PSEN2 encode for homologous polytopic membrane proteins that regulate the production
of Aβ (Sisodia, Kim et al. 1999). Together, mutations in APP and presenilin 1 or
presenlilin 2 result in the increased production of pathologic Aβ (Cai, Golde et al. 1993,
Scheuner, Eckman et al. 1996, Borchelt, Ratovitski et al. 1997, Oyama, Sawamura et al.
1998), which rapidly oilgomerizes, aggregates into fibrils, and finally deposits as the
main component of neurotic plaques (Selkoe 2001, Wirths, Weis et al. 2006). In addition
to early-onset FAD mutations, it was discovered that the presence of ε4 allele of APOE
gene is also a risk factor for AD. ApoE plays vital role in the metabolism and clearance
of Aβ along with α2M and low-density lipoprotein receptor (LRP). Overexpression of
APOE ε4 allele leads to the increased Aβ aggregation and decreased clearance (Vetrivel
and Thinakaran 2006). Besides inherent issue, the majority of AD cases are the sporadic,
late-onset form of the disease, indicating that environmental factors may involve in
disease development.

43

3.2 The Generation of Β-Amyloid (Aβ) Peptide
3.2.1 The function of amyloid precursor protein (APP)
The human amyloid precursor protein (APP) gene, located on chromosome 21, was first
identified in 1987 by several laboratories independently. APP is an integral type I
membrane glycoprotein that is trafficked through the constitutive secretory pathway. It
has a large N-terminal extracellular domain (ectodomain) and a short cytoplasmic tail.
APP is expressed as three alternative spliced isoforms: APP695 (neuronal form), and
APP770/751 isoforms (peripheral and glial isoforms). Since the discovery of APP, its
actual functions remain unclear because the removal of APP gene does not lead to any
phenotype in gene-targeting model mice (Zheng, Jiang et al. 1996). Recent studies reveal
few possible physiological functions of APP in neurons. First, APP may functions as an
axonal transport receptor. APP binds to the light chain subunit of kinesin 1, a microtubule
motor protein (Kamal, Stokin et al. 2000) and directs kinesin-mediated axonal transport
(Kamal, Almenar-Queralt et al. 2001). Secondly, APP was found to associate with the G
protein G
o
in brain (Nishimoto, Okamoto et al. 1993), and missense mutations near the γ-
secretase site in APP lead to the constitutive activation of G
o
-inked receptors (Okamoto,
Takeda et al. 1996). A signal transduction pathway might also link APP to apoptosis
through the activation of a G-protein dependent pathway (Mbebi, See et al. 2002).
Moreover, the cytoplasmic C-terminal domain of APP, AICD, can be transported into
nucleus to from a transcriptionally active complex which modulates calcium signaling
(Cao and Sudhof 2001).
44

3.2.2 Proteolytic process of amyloid precursor protein (APP)
There are two pathways responsible for APP metabolism. Amyloidogenesis is the favored
metabolic pathway in neurons, and nonamyloidogenesis is the one predominant in all
other types of cells (Figure 3-1). Commitment of APP to these pathways can be
modulated by the activation of cell-surface receptors, which respond to transduction
pathways including calcium, arachidonic acid metabolites, and PKC (Checler 1995,
Racchi, Solano et al. 1999). In nonamyloidogenic proteolytic pathway, APP is first cut by
α-secretase, a membrane associated metalloproteinase, within the Aβ domain (between
residues 16 and 17). This cleavage results in the release of the large soluble extracellular
N-terminal portion of APP (APPsα) which precludes the liberation of an amyloidogenic
species, and a C-terminal fragment consisting of 83 residues (C83). C83 then undergo
further proteolysis by γ-secretase to liberate the P3 peptide, which has no amyloid toxic,
but exists in diffused plaques (Lalowski, Golabek et al. 1996, Tekirian, Saido et al. 1998)
(Figure 3-1B). Aβ, the main component of amyloid deposits in AD, is generated by a
different proteolytic process. To release the Aβ peptide through amyloidgenic pathway,
APP must undergo two sequential endoproteolytic steps that are mediated by distinct
enzymes known as β- and γ-secretase. Β-secretase, the enzyme also known as BACE (the
β-amyloid precursor cleaving enzyme) (Sinha, Anderson et al. 1999, Vassar, Bennett et al.
1999)), cleaves APP at the N-terminal region of Aβ sequence. Cleavage by β-secretase
generates a shorter amino terminus (APPsβ) and the Aβ containing fragment (C99)
(Selkoe 2001, Sisodia and St George-Hyslop 2002). Gamma-secretase is an aspartyl
protease complex consisting of at least four components: nicastrin, APH-1, PEN-2, and
45

presenilin (Wolfe 2006), and presenilin contains the proteolytic activity that carries out
the further cleavage of APP (Wolfe, Xia et al. 1999). Cleavage of C99 by γ-secretase
occurs at the ε-site within the transmembrane domain, releasing the C terminus of APP,
known as the APP intracellular carboxyl-terminal domain (AICD). This Aβ-containing
fragment is then cut at the γ-secretase site to release Aβ (LaFerla 2002) (Figure 3-1C).
APPsα, the large soluble extracellular portion generated by α-secretase in the
nonamyloidogenesis has been reported to have neurotrophic properties. In contrast, Aβ
peptides exert cytotoxicity to neuronal survival (Vetrivel and Thinakaran 2006). Hence,
the nonamyloidgenic pathway appears to be neuroprotective compared to the
neurodegenerative effect generated in amyloidogenic pathway.

46

Figure 3-1 Proteolytic processing of amyloid precursor protein (APP).
(A) Schematic structure of transmembrane APP is shown in yellow and the Aβ domain is enlarged
and marked in red. The amino acid residues are numbered from the N-terminus of Aβ and the major
sites of cleavage by α-, β-, and γ-secretases are indicated by arrows. (Figure adapted from Vetrivel
and Thinakaran 2006)
47

Figure 3-1 (Continued) Proteolytic processing of amyloid precursor protein (APP).
(B) Nonamyloidogenic process of APP refers to a proteolytic process carried by membrane bound α-,
and γ-secretases. The cleavage site of α-secretase locates within the Aβ domain, thus preventing the
generation of intact Aβ peptide. The following cleavage made by γ-secretases generates the N-
terminally truncated Aβ (p3) and APP intracellular domain (AICD). (C) In amyloidogenic process,
APP is first cleaved by β-secretase to generate the Aβ containing peptide. With the function of γ-
secretases, Aβ peptide is released. (Figure adapted from Vetrivel and Thinakaran 2006)
48

3.3 Amyloid Protein Aggregation
Alzheimer’s disease belongs to protein misfolding diseases (PMD) that the dysregulation
of protein folding is the primary pathological cause of diseases. Several reasons can lead
to protein misfolding and the most common one is self-aggregation. Aβ, the 4 kDa
peptide generated from a series of proteolytic cleavages of APP, is the basic structural
unit of mature β-amyloid fibril. The two most abundant forms of Aβ are the 40 and 42
residue peptides, Aβ40 and Aβ42 respectively. Aβ is an amphipathic surface-active
peptide, and oligomerization can starts in a concentration dependent manner under proper
hydrophobic environment (Soreghan, Kosmoski et al. 1994, Tjernberg, Pramanik et al.
1999, LeVine 2002). Monomeric Aβ is not neurotoxic, and aggregation is required for
toxic gain-of-function (Lorenzo and Yankner 1994). First, Aβ peptides assemble into a
60-100Å diameter β-sheet rich secondary structure which can accommodate almost
unlimited numbers of polypeptide chains (Kelly 1996, Walsh, Hartley et al. 1999, Nelson,
Sawaya et al. 2005). Then due to the intermolecular interactions, the β-sheet structure is
stabilized, and oligomerization is initiated. Studies using synthetic, cell-derived and
purified recombinant proteins revealed a number of soluble intermediates, including
spherical oligomers, Aβ-derived diffusible ligands (ADDLs), protofibrils, and annular
protofibrils in the aggregation process (Harper, Wong et al. 1997, Lashuel, Hartley et al.
2003). Spherical oligomers are small assembles of misfolded proteins sizing from dimers
to 24-mers (Glabe 2006, Walsh and Selkoe 2007). Further association of these oligomers
results in higher molecular weight assemblies called ADDLs and protofibrils which
eventually exceed solubility limits and finally deposit as amyloid fibrils (Ferreira, Vieira
49

et al. 2007). Aβ-derived diffusible ligands (ADDLs), which only forms from Aβ42,
ranges in molecular weight between 17,000 and 42,000 Da. Protofibrils are larger
aggregates showing as curvilinear structures of 4-11 nm diameter and < 200nm long
under electron microscopy (Walsh, Hartley et al. 1999, Caughey and Lansbury 2003).
Protofibrils are capable in elongation by growing on their ends in a concentration
dependent manner (Harper, Wong et al. 1999). Annular protofibrils are ring-like pore
structures that form on cell membrane and may contribute to cell death (Lashuel, Hartley
et al. 2002). Amyloid fibrils are long, straight and unbranched assembles with
approximately 10 nm diameter and several µm lengths. A unique feature of amyloid
fibrils is the binding with dyes Congo red and thioflavin, hence this property is usually
used to distinguish amyloid fibrils from oligomers (Nelson, Sawaya et al. 2005). Once the
amyloid fibril has been developed, it can continuously grow by the addition of monomer
onto the ends of the fibrils (Collins, Douglass et al. 2004).

3.4 Aβ Oligomers, the Pathological Agent in Alzheimer’s Disease
The pathogenesis of Alzheimer’s disease is complex, and involves in many molecular,
cellular, and physiological mechanisms. Over the years, evidence from biochemical,
genetic, and pathologic investigations overwhelmingly support the idea that
amyloidogenesis is a critical and early event in AD pathogenesis. Early pathological
studies identified the 6-10 nm amyloid fibrillar protein aggregates in intracellular
50

inclusions or in extracellular deposits, and named as amyloid plaques (Terry, Gonatas et
al. 1964, Maloy, Longnecker et al. 1981). Amyloid plaques were found in variant organs
in different amyloid diseases, such as brain in Alzheimer’s diseases. Regardless of the
difference of amino acid sequence, amyloid proteins share many common physical and
tinctorial properties (Dobson 2003). Because of the common characteristics and the
ubiquitous presence in affected tissues, amyloid fibrils were highly speculated as the
pathological cause of amyloid diseases (Castano, Ghiso et al. 1986, Westermark,
Engstrom et al. 1990, Gustavsson, Engstrom et al. 1991). Studies using transgenic animal
models of amyloid diseases demonstrated that fibril containing amyloid plaques
abundantly present in affected tissues (Quon, Wang et al. 1991, Wirak, Bayney et al.
1991). Moreover, synthetic fibrils were also proved as toxic to a variety of primary cells
in culture (Lorenzo and Yankner 1994, Lopes, Colin et al. 2004). These all supported the
idea that amyloid fibrils played crucial roles in the pathogenesis of amyloid diseases.
Therefore, amyloid cascade hypothesis suggested that the accumulation of Aβ driven by
increased generation of fibrillogenic Aβ over its clearance is the initiating molecular
event for the subsequent formation of neurofibrillary tangles, senile plaques, neuronal
dysfunction, neuronal death, microglial cell activation in AD pathogenesis (Hardy and
Selkoe 2002).
However, despite of these strong supports, the “fibril hypothesis” is questionable due to
the failure in explaining crucial clinical and pathological issues of several degenerative
diseases. In Alzheimer’s disease, it has been pointed out that there is poor or absence of
correlation between plaques and dementia (Terry, Masliah et al. 1991, Giannakopoulos,
51

Herrmann et al. 2003). Two interesting studies showed that passive immunization using
anti-Aβ antibodies leads to the reversal of memory loss without reduction in the number
of amyloid plaques in different transgenic mouse models of AD (Dodart, Bales et al.
2002, Kotilinek, Bacskai et al. 2002). Therefore, the fibril hypothesis does not fully
account for pathogenesis in patients and in animal models of amyloid diseases, hence
amyloid fibrils is no longer thought as the real “killer” in amyloid diseases.
Indeed, recent studies have shown that soluble oligomers of the corresponding
proteins/peptides are associated with pathology in many amyloid diseases. The increasing
number of soluble Aβ oligomers in the brains of affected individuals has been recognized
as an additional neuropathological hallmark of AD (Klein, Krafft et al. 2001). Early
evidence indicated the elevated levels of soluble Aβ and presence of pre-fibrillar Aβ
aggregates showing in AD-affected brains (Tabaton, Nunzi et al. 1994, Kuo, Emmerling
et al. 1996). This unusual increase of soluble Aβ oligomers have also been found in
different transgenic mouse models of AD, further supporting the idea that Aβ oligomers
play crucial roles in AD pathogenesis (Lesne, Koh et al. 2006, Oddo, Caccamo et al.
2006). Lambert et al. showed that soluble Aβ oligomers formed spontaneously in vitro
under conditions in which fibril formation was inhibited. Moreover, they also found that
such oligomers, named as ADDLs (Aβ-Derived Diffusible Ligands), were neurotoxins
that killed neurons in organotypic cultures (Lambert, Barlow et al. 1998). Both
protofibrils and annular protofibrils have also been reported neurotoxic in different
studies (Hartley, Walsh et al. 1999, Lashuel, Hartley et al. 2002). On the other hand,
other groups have also demonstrated that synthetic Aβ monomers aggregate in vitro in a
52

time-dependent fashion to form oligomers, which eventually may form fibrils (Huang,
Yang et al. 2000, Ward, Jennings et al. 2000, Bitan, Lomakin et al. 2001, Kayed, Head et
al. 2003). These observations all suggested that soluble oligomeric Aβ, accumulating as
the precursors of amyloid deposits, is the primary pathological causes of AD (Ferreira,
Vieira et al. 2007).

3.5 Neuronal Impact of Soluble Aβ Oligomers
Oligomeric Aβ is thought to be the neurotoxicity which impairs learning, memory and
synaptic plasticity (Walsh and Selkoe 2004, Haass and Selkoe 2007). Recently, studies
indicate Aβ oligomers with the ability to initiate a series of events including membrane
peroxidation, oxidative stress induction, calcium homeostasis disruption and tau
hyperphosphorylation at the early stage in AD pathogenesis (Kelly and Ferreira 2006, De
Felice, Velasco et al. 2007, De Felice, Wu et al. 2008). All these events are considered to
play important roles in the progressive damage and loss of neurons. Collectively, these
observations imply Aβ oligomerization as an upstream event leading to neuronal
dysfunction and, eventually, to dementia in AD.

53

3.5.1 Calcium homeostasis
Aβ oligomers have been shown to destabilize neuronal calcium homeostasis which
generally leads to an increase of cytosolic calcium (Mattson, Cheng et al. 1992, Mattson,
Tomaselli et al. 1993). A major mechanism by which Aβ oligomers are believed to alter
cellular Ca
2+
homeostasis involves disruption of membrane Ca
2+
permeability. The three
major proposed mechanisms of Aβ interaction with cell membrane, involving interactions
with endogenous Ca
2+
-permeable channels, disruption of membrane lipid integrity, and
formation of Ca
2+
-permeable channels by Aβ (Demuro, Parker et al. 2010). Various Ca
2+
-
permeable channels including voltage-gated Ca
2+
channels, nicotinic acetylcholine
channels, glutamate receptors (AMPA and NMDA), and dopamine receptors (Rovira,
Arbez et al. 2002, Verdurand, Dalton et al. 2010), have been reported as targets of Aβ,
and the binding of Aβ to these channels directly induce Ca
2+
influx into cells. In addition,
Aβ peptides are known to interact with membrane lipids such as phosphoinositides
phosphatidylglycerol (Terzi, Holzemann et al. 1995), phosphatidylcholine (Avdulov,
Chochina et al. 1997), and gangliosides (McLaurin, Franklin et al. 1998). A direct
interaction of Aβ with cell membranes was initially proposed by Cotman and co-workers
(Cribbs, Pike et al. 1997). Fluorescence spectroscopy measurements indicate that Aβ
interaction with the synaptic plasma membrane causes substantial changes in the
membrane fluidity and results in increasing membrane permeability to Ca
2+
, Na
+
, and K
+

ions (Muller, Koch et al. 1995, McLaurin and Chakrabartty 1996). Another different
mechanism of action posits that Aβ peptides form nonselective high conductance cation
pores by directly incorporating into cell membrane (Arispe, Rojas et al. 1993, Lin, Bhatia
54

et al. 2001). This pore-forming mechanism has been further supported by studies using
atomic force microscopy (Lin, Bhatia et al. 2001), electron microscopy (Lashuel, Hartley
et al. 2002), and theoretical modeling (Durell, Guy et al. 1994). The Aβ channels are
capable for conducting huge amount of Na
+
, K
+
, and Ca
2+
which can rapidly disrupt
cellular homeostasis and cause severe Aβ toxicity (Arispe, Rojas et al. 1993).
These studies all indicate that calcium dysregulation is an early molecular defect in the
pathogenesis of AD. However, because calcium destabilization can also induce the
hallmark features of the disease (calcium hypothesis) such as the hyperphosphorylation
of tau, the accumulation of amyloid β, and neuronal death (LaFerla 2002), there may be a
potential reciprocal mechanism between Ca
2+
pathways and amyloid pathology. These
two factors continuously potentiate each other, generate a feed-forward cycle, and finally
lead to further neurodegeneration (LaFerla 2002).

3.5.2 Generation of reactive oxygen species (ROS)
Sustained increases in cytosolic Aβ oligomer levels trigger several intracellular events
and one of them is the production of reactive oxygen species (ROS) (Chinopoulos and
Adam-Vizi 2006). Two mechanisms have been described involving in the elevation of
ROS, metal interaction and mitochondria dysfunction. During peptide oligomer
formation, Aβ generate hydrogen peroxide through a process enhanced by iron (Fe
2+
) and
copper (Cu
2+
) (Hensley, Carney et al. 1994, Huang, Atwood et al. 1999). Secondly, the
increase of free radicals is resulted from the dysfunction of mitochondria caused by Aβ-
55

mediated Ca
2+
homeostasis disruption. Increased Ca
2+
levels in mitochondria leads the
generation of superoxide radicals by altering mitochondrial oxidative phosphorylation
and activating oxygenases (Mattson 1995, Mattson, Barger et al. 1995, Bezprozvanny
and Mattson 2008). The interaction of Aβ with mitochondria also causes the impairment
of the electron transport chain, leading to the decrease of ATP production, cytochrome c
release, and caspase activation (Mattson 2004). On the other hand, Aβ may disrupt
calcium homeostasis by triggering oxidative stress which leads to membrane calcium
pumps impair and enhances calcium influx through voltage-dependent channels and
glutamate receptors (Mattson 2004).

3.5.3 Membrane peroxidation
Aβ oligomers can initiate lipid peroxidation during AD progression (Schubert, Behl et al.
1995, Lambert, Barlow et al. 1998, Selkoe 2001, Butterfield, Castegna et al. 2002,
Butterfield and Lauderback 2002, Butterfield, Reed et al. 2007) by membrane integration
and/or surface binding (Hertel, Terzi et al. 1997, Kayed, Sokolov et al. 2004). When Aβ
aggregation occurs at the cell membrane, oxidative stress is induced resulting in lipid
peroxidation (Hensley, Carney et al. 1994) and the consequent generation of 4-
hydroxynonenal (4HNE) (Mattson 1997), which impairs the function of ion-motive
ATPase and other transporters, leading to the elevation of basal intracellular calcium
levels (Mark, Hensley et al. 1995). The proteins modified by this Aβ-induced oxidative
stress include membrane transporters (ion-motive ATPase, glucose transporters, and
56

glutamate transporters), receptors, GTP-binding proteins and ion channels (VDCC:
voltage-dependent chloride channel; NMDAR: N-methyl-D-aspartate receptor)
(Butterfield, Drake et al. 2001).

3.5.4 Tau as a downstream target of Aβ oligomers
Neurofibrillary tangles (NFTs) are intracellular fibrillar aggregates consisting of
hyperphosphorylated tau protein. Under physiological conditions, tau plays important
roles in microtubule stabilization and dynamics. Previously, in vitro and in vivo studies
had shown that Aβ deposits are associated with neuronal tau hyperphosphorylation (Gotz,
Chen et al. 2001, Oddo, Caccamo et al. 2003, Oddo, Billings et al. 2004), and increased
brain levels of soluble Aβ were found correlatively with NFT density both in AD patients
and transgenic mouse models (McLean, Cherny et al. 1999). The level of phosphorylated
tau was selectively increased in a subpopulation of neurons that specifically exhibited Aβ
oligomer binding in immunocytochemical analysis, and an anti-Aβ oligomer antibody
could significantly block tau hyperphosphorylation (De Felice, Wu et al. 2008). Hoshi et
al. then demonstrated that Aβ oligomers activate glycogen synthase kinase-3β (Hoshi,
Sato et al. 2003), a kinase involving in the pathological hyperphosphorylation of tau. The
misregulation of CK2 kinase has also been reported participating in the
hyperphosphorylation process (Pigino, Morfini et al. 2003, Lee and Trojanowski 2006,
LaPointe, Morfini et al. 2009, Pigino, Morfini et al. 2009). Another recent study
demonstrated that tau mediates the destabilization of neuronal microtubules induced by
57

Aβ oligomers, which results in neuronal dysfunction triggered by cytoskeletal changes
(King, Kan et al. 2006). These observations suggest that Aβ oligomers trigger the
activation of intracellular signaling pathways leading to tau hyperphosphorylation, and
the hyperphosphorylation of tau is a crucial step in neurodegeneration in AD.

3.6 A Potential Protecting Factor against AD, Annexin A5
Alzheimer’s disease is one of the most common dementia in elder population today, and
effective treatments against this disease are still in search. Many factors involving in
calcium regulation or antioxidation have been reported as potential targets for medical
therapy. Currently, annexin A5, a Ca
2+
-dependent membrane-binding protein, has been
proposed to directly interact with amyloidogenic proteins and reduces their toxicity by
competitive interaction at membrane phosphatidylserine (Lee, Pollard et al. 2002)
,
(Bedrood, Jayasinghe et al. 2009). Therefore, we speculate that annexin A5 may play a
protecting role in the progress of Alzheimer’s disease.
Annexins are one class of Ca
2+
regulated proteins. In vertebrate, 12 annexin subfamilies
(A1-A11 and A13) with different splice variants have been identified. They are
abundantly expressed in a wide range of tissues (Reutelingsperger 2001, Rescher and
Gerke 2004) and found in intra- and extracellular locations (Flaherty, West et al. 1990,
Romisch, Schuler et al. 1992). Annexins are characterized by the unique architecture of
58

their Ca
2+
binding site, which enables them to peripherally dock onto negatively charged
membrane surfaces in their Ca
2+
bound conformation. Each annexin is composed of two
principal domains: the various N-terminal region, and the conserved C-terminal core
domain representing the major part of annexins. The N-terminal domain is diverse in
sequence and length, involving in the regulation of protein interaction and membrane
association. The C-terminal domain is consisted by four 70 amino residue repeats called
annexin repeats, and only annexin A6 has eight. An important α-helical domain carrying
a Ca
2+
binding site is presented in each repeat, and this is responsible for the different
binding affinity to phospholipids (Raynal and Pollard 1994, Camors, Monceau et al.
2005). This highly α-helical core domain forms a compact structure not only harboring
the Ca
2+
- and membrane-binding sites but also being available for other types of
interaction/regulation (Raynal and Pollard 1994, Gerke and Moss 2002). Depending on
Ca
2+
concentration, annexins have been reported to participate in a variety of membrane-
related events such as exocytosis, endocytosis, apoptosis, binding to cytoskeletal proteins,
and organization of membrane domains (Reutelingsperger 2001, Gerke and Moss 2002,
Rescher and Gerke 2004, Gerke, Creutz et al. 2005). They have also been reported to
regulate protein activities.
Even after decades of study, the physiological function of annexin A5 is still not clear.
Annexin A5 is the only annexin that has extracellular presence in addition to intracellular
localization. The most well-known feature of annexin A5 is its high affinity to
membranes carrying phosphatidylserine (PS). This binding of annexin A5 to PS has been
accounted for its anticoagulant (Thiagarajan and Tait 1990, van Heerde, Sakariassen et al.
59

1994), antiapoptotic (Gidon-Jeangirard, Hugel et al. 1999), and anti-inflammatory
(Reutelingsperger and van Heerde 1997) effects. On the other hand, annexin A5 has been
widely used as a molecular imaging agent to visualize PS-expressing apoptotic cells in
vitro and in vivo in animal models and patients over the past decade (Boersma, Kietselaer
et al. 2005). Recently, some studies proposed that annexin A5 may form Ca
2+
channels to
mediate Ca
2+
influx (Rojas, Arispe et al. 1992, Arispe, Rojas et al. 1996)
,
(Kirsch,
Harrison et al. 2000)
,
(Wang, Xu et al. 2003), but more evidence are required to support
this hypothesis.
Another unique property of annexin A5 and several other annexins is to self-assemble
into two-dimensional ordered arrays on membrane binding (Pigault, Follenius-Wund et al.
1994, Voges, Berendes et al. 1994, Reviakine, Bergsma-Schutter et al. 1998, Gerke,
Creutz et al. 2005). In the presence of mM Ca
2+
concentration, annexin A5 molecules
bind to PS-containing membranes and rapidly self-assemble into trimmers (Concha, Head
et al. 1992, Langen, Isas et al. 1998, Oling, Bergsma-Schutter et al. 2001), and then the
trimers are further align into two-dimensional ordered arrays (Mosser, Ravanat et al.
1991, Voges, Berendes et al. 1994, Reviakine, Bergsma-Schutter et al. 1998, Oling,
Bergsma-Schutter et al. 2001). This property of annexin A5 is used to form an
anticoagulant shield covering the surface of placental syncytiotrophoblasts. Furthermore,
annexin arrays have been proposed to affect membrane properties such as fluidity,
rigidity, and lipid segregation, therefore participating in the regulation and/or stabilization
of membrane domains (Brisson, Mosser et al. 1991, Pigault, Follenius-Wund et al. 1994,
Reviakine, Bergsma-Schutter et al. 1998, Reviakine, Bergsma-Schutter et al. 2000).
60

Chapter 4
Annexin A5 protects heart function in a mouse model for Alzheimer’s
disease

4.1 Introduction
Alzheimer’s disease (AD) is the most common form of age-associated dementia which
leads to cognitive, memory, and behavioral impairment. The characteristic and invariant
lesions in brains of afflicted individuals include the accumulation of 38- to 43-amino-acid
β-amyloid peptides (Aβ) in plaques, and the appearance of hyperphosphorylated tau in
neurofibrillary tangles. The pathogenesis of Alzheimer’s disease is very complicated, and
involves in many molecular, cellular, and physiological mechanisms. Amyloid cascade
hypothesis, which was proposed over a decade ago, suggests that the accumulation of Aβ
peptides driven by increased generation of Aβ over its clearance is the initiating
molecular event that triggers neurodegeneration in Alzheimer’s disease (Hardy and
Selkoe 2002).
Currently, annexin A5 has been proposed to offer protection from the cytotoxicity of
Alzheimer’s β-protein (Aβ) through competitive interaction at membrane
phosphatidylserine (Lee, Pollard et al. 2002). On the other hand, Bedrood et al. further
indicated that annexin A5 directly interacts with other amyloidogenic proteins and
61

reduces their toxicity (Bedrood, Jayasinghe et al. 2009). Therefore, we speculated that
annexin A5 may play a role in preventing the Aβ cytotoxicity in Alzheimer’s disease.
In order to study the function of annexin A5 in the pathogenesis of Alzheimer’s disease,
we generate APP-PS1/ANXA5KO mice by breeding APP-PS1dE9 mice, a widely used
mouse model Alzheimer’s disease, with ANXA5KO mice. No overt difference in the
number of brain Aβ plaques were seen between the APP-PS1dE9 and the APP-
PS1/ANXA5KO mice. Interestingly, APP-PS1/ANXA5KO mice develop severe heart
deficiency and die from sudden death at an early age. The association between AD and
cardiac failure has been noted in the past. In addition to brain, presenilin 1 and 2 genes
are also expressed in heart (Levy-Lahad, Poorkaj et al. 1996, Hebert, Serneels et al.
2004), and gene targeting experiments in murine models have shown that they are critical
to cardiac development (Donoviel, Hadjantonakis et al. 1999, Nakajima, Moriizumi et al.
2004). Recently, sequence variation analysis of PSEN1 and PSEN2 was performed in
dilated cardiomyopathy (DCM) patients, and the results indicate that PSEN1 and PSEN2
mutations are associated with heart failure and implicate novel mechanism of myocardial
disease (Li, Parks et al. 2006, Gianni, Li et al. 2010). Likewise, amyloid oligomers have
been found in the failing heart of DCM patients (Gianni, Li et al. 2010), and in mice
model of desmin-related cardiomyopathy (Sanbe, Osinska et al. 2005). Since the genes
for presenilins (Levy-Lahad, Poorkaj et al. 1996, Hebert, Serneels et al. 2004) and APP
(Sandbrink, Masters et al. 1994) are expressed in hearts, as is γ-secretase activity (Hebert,
Serneels et al. 2004), the association between cardiac dysfunction and AD should be an
important area of investigation.
62

Αlpha-crystallin is a member of the small heat-shock protein (sHsp) family which
contains two main types: αA-crystallin and αB-crystallin. αB-crystallin is heat- and
stress-inducible, whereas αA-crystallin is not heat-inducible. Both αA- and αB-crystallin
form homo- as well as hetero-multimers of various sizes (van den Oetelaar, van Someren
et al. 1990), and exhibit molecular-chaperone-like activity in preventing aggregation and
denature of unfolded proteins (Horwitz 1992, Raman and Rao 1994, Sun, Das et al. 1997,
Datta and Rao 1999), with αB-crystallin being more efficient than αA-crystallin. αB-
crystallin was originally discovered and classified as a lens protein (Piatigorsky 1984).
Later, αB-crystallin is also found in nonlenticular tissues and abundantly expressed in
cardiac and skeletal muscle (Iwaki, Kume-Iwaki et al. 1989, Longoni, Lattonen et al.
1990). αB-crystallin binds both desmins and cytoplasmic actins in vitro (Bennardini,
Wrzosek et al. 1992, Nicholl and Quinlan 1994, Wang and Spector 1996). The
upregulation of gene and subsequent accumulation of αB-crystallin occurs in a number
of degenerative neural pathologies such as Alexander and Alzheimer’s disease (Iwaki,
Kume-Iwaki et al. 1989, Shinohara, Inaguma et al. 1993), as well as cardiac disorders
including familial hypertrophic cardiomyopathy and desminopathy (Hwang, Dempsey et
al. 1997, Arbustini, Morbini et al. 1998, Xiao and Benjamin 1999), but the functional
consequences are unknown.

63

4.2 Materials and Methods
4.2.1 Transgenic animals
Male double transgenic mice APP-PS1dE9 bearing chimeric human/mouse amyloid
precursor protein (Mo/HuAPP695swe) gene combined with a mutant human presenilin 1
(PS1-dE9) gene were purchased from Jackson Laboratory, Bar, Harbor, ME, USA
[B6C3-Tg(APPswe,PSEN1dE9)85Dbo/J]. The mice were originally on a B6C3
background and backcrossed for at least 6 generation to a C57BL/6J background. Female
annexin A5 mice (courtesy of Dr. Ralf Langen) bred with APP-PS1dE9 to generate APP-
PS1/ANXA5KO mice. Mice were genotyped by PCR amplification. Genomic DNA was
obtained from mouse-tail biopsy samples. The human/mouse amyloid precursor protein
transgene was detected with primers IMR1597 (5’-GAC TGA CCA CTC GAC CAG
GTT CTG-3’) and IMR1598 (5’-CTT GTA AGT TGG ATT CTC ATA TCC G-3’),
while mutated human presenilin 1 gene was detected with primers IMR1644 (5’-AAT
AGA GAA CGG CAG GAG CA-3’) and IMR1645 (5’-GCC ATG AGG GCA CTA
ATC AT-3). Internal control of presenilin 1 gene was amplified with primers IMR0944
(5’-CCT CTT TGT GAC TAT GTG GAC TGA TGT CGG-3’) and IMR1588 (5’-GTG
GAT AAC CCC TCC CCC AGC CTA GAC C-3’). Transgenic mice bred to annexin A5
-/- mice were genotyped for the annexin A5 locus. PCR was performed to detect the
presence of the annexin A5 null allele by using primers 56990H (5’-CGA GAG GCA
CTG TGA CTG ACT TCC CTG GAT-3’) and 56990I (5’-GCC AGT TTG AGG GGA
CGA CGA CAG-3’), while the annexin A5 allele was detected with primers 56990F (5’-
64

GAA GCA ATG CTC AGC GCC AGG A-3’) and 56990G (5’-CTG TAC TCT ATC
ACT ATC ACT GAC TGT TTA ATC-3’).

4.2.2 Gomori’s modified trichrome staining
3 months old APP-PS1/ANXA5KO and other mice were euthanized and hearts were
taken out immediately and put in 50mM KCl/PBS until not beating. Hearts were carefully
cleaned and fixed with 4% paraformaldehyde/PBS for overnight. Then heart samples
were paraffin embedded, sectioned, and stained with modified Gomori’s trichrome
staining. Sections were deparaffinzed and rehydrated with zylene and different
concentration of ethanol first. Then place sections in Bouin’s fluid (75 ml of picric acid,
saturated aqueous; 25 ml of formaldehyde, 37-40%; 5 ml of acetic acid) at 56 ℃for 1 hour.
Rinse sections in tap water for 3-5 minutes until yellow color is removed. Place sections
in Working Weigert’s Iron Hematoxyin (1:1 mix of solution A [2g of hematoxylin
crystals, 100 ml of 90% ethanol] and solution B [4 ml of ferric chloride, 62% aqueous; 1
ml of hydrochloric acid, concentrated; 95 ml of distilled water]) for 10 minutes and rinse
sections in tap water for 5-10 minutes. Then stain sections in Beibrich Scarlet-Acid
Fuchsin solution (0.9% Biebrich scarlet and 0.1% in 1% acetic acid) for 5-10 minutes to
achieve the desired intensity. After rinsing in distilled water for 30 seconds, place
sections in Phosphotungstric-Phosphomolybdic acid solution (2.5% phosphotungstric
acid, 2.5% phosphomolybdic acid) for 5 minutes. Next, stain sections in Anline Blue
Stain Solution (2.5 g Anline Blue dissolve in 2 ml of acetic acid and 100 ml of distilled
65

water) for 5-10 minutes followed by one minute incubation in 1% acetic acid. After
washing with distilled water, sections were dehydrated, cleaned, and mounted for
observation.

4.2.3 Sample preparation for ultrastructural examination
Hearts from 3 month old APP-PS1/ANXA5KO and other mice were taken out and put in
50mM KCl/PBS until fully relaxed. The papillary muscles were carefully dissected out
and fixed overnight at 4 ℃ with ½ Karnovsky buffer (2% paraformaldehyde, 2.5%
glutaraldehyde, 0.1M cacodylate, pH7.2). The fixed papillary muscles were washed for 3
x 10 minutes with 0.1 M cacodylate buffer, pH7.2 and further fixed with 1% oxmium
tetraoxide in 0.1M cacodylate buffer for 1.5-2.0 hours at room temperature. Papillary
muscles were washed 2 X 10 minutes with 0.1M cacodylate buffer, pH7.2 and
dehydrated in the following manner: 15 minute each in 50%, 70%, 85%, and 95% ethanol;
3 x 10 minutes 100% ethanol; and 2 x 10 minutes 100% propylene oxide. Infiltration with
epoxy resin (28.7% w/w Epon 812, 14.4% w/w Epon 826, 4.8% w/w Epon 871, 46.4%
w/w DDSA, 5.7% w/w NMA, and 3% EBMA) were performed as follows: overnight
incubation with 1:1 epon : proprylene oxide; overnight incubation with 2:1 epon :
proprylene oxide; and 6 hours to overnight incubation with 100% epon. The papillary
muscles were placed in molds filled with epon with proper orientation. The samples were
baked for 3-4 days at 55 ℃. Once hardened, the epon-embedded papillary muscles were
cut for ultrastructural examination.
66

4.2.4 Electrocardiogram (ECG) recording
Electrocardiogram was performed on female APP-PS1/ANXA5KO and other mice every
two week from 2.5 months old. Mice were weighted and anesthetized for recording based
on standard procedure. This part of analysis was done by Robabeh Mohammadzadeh.
After ECG, hearts were dissected out, weighted, and prepared for paraffin section.

4.2.5 Western blot analysis
Hearts were taken from indicated mice at 3 months old and homogenized in TBS buffer
(15mM Tris, pH7.4; 140mM NaCl, 3mM KCl, and 0.5mg/ml protease inhibitor cocktail
[Roche Inc]). After centrifugation, the supernatant was collected as the cytoplasm
fraction and the pellets were homogenized again with TBST buffer (15mM Tris, pH7.4;
140mM NaCl, 3mM KCl, 1% triton X-100, and 0.5mg/ml protease inhibitor cocktail
[Roche Inc]) to isolate membrane associated proteins. One dimensional PAGE was
carried out and proteins were transferred onto nitrocellulose membranes. The blots were
saturated in TBS-T buffer [20mM Tris (pH7.5), 136.8mM NaCl, and 0.1% Tween 20]
containing 5% non-fat milk. Mouse monoclonal anti-Aβ peptide (6E10, Signet
Laboratories), rabbit polyclonal anti-α-B crystalline, and mouse monoclonal anti-actin
antibodies were used as primary antibodies. The secondary antibodies of IRDye 680 goat
anti-mouse and IRDye 800 goat anti-rabbit antibodies (LI-COR Biosciences, Lincoln NE)
were applied accordingly. The proteins were visualized and quantified using Odyssey
Infrared Imaging System (LI-COR, Biosciences).
67

4.2.6 Cryosection and immunocytochemistry
Ventricular muscles were dissected from 3 months old mice and immersed in 30%
sucrose/PBS for overnight followed by embedding into O.C.T. medium (Tissue Tek).
Cross-sectional and longitudinal sections were collected at 10µm thickness on glass
slides. Sections were dried and fixed with cold 100% methanol for 5 minutes. After
washing with PBS twice, samples were permeabilized with 0.1% Triton X-100 in PBS
for 30 minutes. Wash slides with PBS, 1% BSA/PBS, and 3% BSA/PBS for 5 minute
each. Samples were incubated with primary antibody (α-B-crystallin, 1:1000) in 1%
BSA/PBS at 4 ℃ overnight followed by 4 x 5 minute of wash with PBS, 1% BSA/PBS,
2% BSA/PBS, and 3% BSA/PBS accordingly. Sections then were incubated for 1hour
with 1:400 dilutions of FITC-conjugated goat anti-rabbit IgG (Vector Laboratories, Inc.)
and washed with PBS, 1% BSA/PBS, 2% BSA/PBS, 3% BSA/PBS again. A tiny drop of
Vectashield (Vector Laboratories, Inc.) was placed on the sections which were then cover
slipped. Images were acquired on an Axioplan2microscope (Zeiss Co.) All images for
each section were taken at the same detection gain.

4.2.7 Mouse perfusion
Mice at 3 month age were anesthetized with an intraperitoneal injection of 100 mg
ketamine and 10 mg xylazine per kg body weight. Quickly open chest, cut right atrium to
release blood, and inject a needle connecting with PBS filled syringe into left ventricle.
Pump PBS into heart at 5 ml/min speed for circulation. Wash out blood until liver turns
68

white. Then keep the needle in ventricle and carefully switch it to another syringe
containing 4% paraformaldehyde (PFA)/PBS. Pump 4% PFA into heart at 5 ml/min for
circulation until mouse tail is stark and stiff. Open head and chest quickly to take our
brain and heart for further processing.

4.2.8 Thioflavin-S staining
Age matched APP-PS1/ANXA5KO and other mice were anesthetized and perfused with
4% paraformaldehyde. Brains were taken out and embedded in 3% agarose gel. Samples
were then prepared for vibratome section at 100 µm thickness. Make 1% thioflavin-S
solution freshly and filter to remove undissolved particles. Drop thioflavin-S on sections
and incubate for 10 minute at room temperature. Cover sections with foil to protect
solution from light. Decant thioflavin-S and wash sections with 80% ethanol for 1 minute.
Change to new ethanol for another 2 minute incubation. Remove 80% ethanol and wash
in 95% ethanol for 1 minute. Wash sections with three exchanges of distilled water. Put
on coverslip with aqueous mounting media.

4.2.9 Western blot for β-amyloid protein
Brain were taken from indicated mice at 1 month or 1 year old and homogenized in TBS
buffer (15mM Tris, pH7.4; 140mM NaCl, 3mM KCl, and 0.5mg/ml protease inhibitor
cocktail [Roche Inc]). After centrifugation, the supernatant was collected as the
69

cytoplasm fraction and the pellets were homogenized again with TBST buffer (15mM
Tris, pH7.4; 140mM NaCl, 3mM KCl, 1% triton X-100, and 0.5mg/ml protease inhibitor
cocktail [Roche Inc]) to isolate membrane associated proteins. After second
centrifugation, 80% of formic acid was added to the pellets and samples were nutated for
overnight at 4 ℃. Remove undissolved tissue by centrifugation and neutralize formic acid
extracted fraction with 2M Tris-base, pH11 before loading to gels. One dimensional
PAGE was carried out and proteins were transferred onto nitrocellulose membranes. The
blots were saturated in TBS-T buffer [20mM Tris (pH7.5), 136.8mM NaCl, and 0.1%
Tween 20] containing 5% non-fat milk. Mouse monoclonal anti-Aβ peptide (6E10,
Signet Laboratories), rabbit polyclonal OC antibody (courtesy of Dr. Chaeles Glabe)
were used as primary antibodies. The secondary antibodies of IRDye 680 goat anti-
mouse and IRDye 800 goat anti-rabbit antibodies (LI-COR Biosciences, Lincoln NE)
were applied accordingly. The proteins were visualized and quantified using Odyssey
Infrared Imaging System (LI-COR, Biosciences).

70

4.3 Results
4.3.1 The body weight of APP-PS1/ANXA5KO female mice is significantly lower
When we got the APP-PS1/ANXA5KO mice, we noticed that these mice seem to be
smaller than wild type mice. Therefore, we weighted the APP-PS1/ANXA5KO and other
mice once every week from 1 month old to sequentially check the change of body weight.
From the statistic results, female APP-PS1/ANXA5KO mice show significantly lower
body weight as compared to other female mice (Fig. 4-1A). ANOVA analysis suggests
significant weight differences between the groups. The table shows the p values from
one-tail t-test for the lighter weight of APP-PS1/ANXA5KO female as compared to the
other three groups. Cells marked in red represent the p value smaller than 0.05. However,
the lighter body weight of APP-PS1/ANXA5KO mice does not show in male group (Fig.
4-1B).

71

Figure 4-1 The body weight of APP-PS1/ANXA5KO female mice is significantly lower than wild
type mice.
(A) Female mice from different groups, WT, blue line (n=5) ; APP-PS1, green line (n=6) ;
ANXA5KO, pink line (n=4) ; APP-PS1/ANXA5KO(APP/ANXKO), red line (n=6) are weighed
once every week from 1 month of age. The body weight of the APP-PS1/ANXA5KO group is
significantly less than the other groups. ANOVA analysis suggests significant weight differences
between the groups with p-value: 7.02E-10 at week 4, 8.89E-07 at week 5, 6.24E-05 at week 6,
3.99E-05 at week 7, and 1.91E-4 at week 8. The lower table shows p-values from one-tail t-test for
the lower weight of APP-PS1/ANXA5KO female as compared to the other three groups. Cells of p-
value < 0.05 are marked in red. (B) Male mice from different groups, WT, blue line (n=3) ; APP-
PS1, green line (n=5) ; ANXA5KO, pink line (n=5) ; APP-PS1/ANXA5KO(APP/ANXKO), red line
(n=6) are weighed from 1 month of age. Male APP-PS1/ANXA5KO mice do not show significant
weight difference from WT mice.

72

4.3.2 APP-PS1/ANXA5KO mice have low survival rate especially in the female group
The early death occurring in the APP-PS1/ANXA5KO mice also attracted our attention.
The statistic data monitoring total 101 (male: 46; female: 55) APP-PS1/ANXA5KO and
other mice from 1 to 6 months old shows that the survival rate of male APP-
PS1/ANXA5KO mice is only 47% at 6 months old and even lower to 34% in female
group (Figure 4-2).

4.3.3 Abnormal cardiac morphology show in young ANXA5KO and APP-
PS1/ANXA5KO mice
Previous studies have shown that presenillin 1 and 2 are both expressed in heart and the
mutations of these two genes have been identified in many cardiomyophathies (Levy-
Lahad, Poorkaj et al. 1996, Hebert, Serneels et al. 2004, Li, Parks et al. 2006, Gianni, Li
et al. 2010). In order to figure out whether cardiac deficiency may cause the early death
of APP-PS1/ANXA5KO mice, hearts were collected, prepared, sectioned, and stained
with Gomori’s modified Trichrome staining for morphological examination (Figure 4-3).
Enlarged right ventricles are found in ANXA5KO and APP-PS1/ANXA5KO mice (Fig.
4-3C, 4-3D). Under higher magnification, the cardiomyocytes are properly aligned and
well organized in wild type and APP-PS1 mice (Fig. 4-3E, 4-3F). In contrast, the muscle
bundles in ANXA5KO and APP-PS1/ANXA5KO mice are loosely packed (Fig. 4-3G, 4-
3H).
73

Figure 4-2 Early sudden death of APP-PS1/ANXA5KO (APP/ANXKO) mice, especially in the
female group.
The viability of male (n=46) and female (n=55) APP-PS1/ANXA5KO mice are monitored from 1 to
6 months. The survival rate is 46.67% in male and 34.83% in female at 6 months while APP-PS1
and ANXA5KO mice are both 100%.

74

Figure 4-3 Abnormal cardiac morphology in young ANXA5KO and APP-PS1/ANXA5KO
(APP/ANXKO) mice.
Three months old mice were anesthetized and perfused with 4% paraformaldehyde in PBS. Hearts
were embedded into paraffin, sectioned, and stained with Gomori’s modified trichrome stain. Both
ANXA5KO and APP-PS1/ANXA5KO shows enlarged right ventricle under low magnification. At
higher magnification, both ANXA5KO and APP-PS1/ANXA5KO cardiac muscles show loose
cardiomyocyte bundles when compared to the wildtype and APP-PS1 mice.

75

4.3.4 Diffused Z-lines and disorganized myofilaments are found in APP-
PS1/ANXA5KO mice
In order to check the detail morphology of cardiomyocytes in APP-PS1/ANXA5KO mice,
papillary muscles were taken from ventricles and prepared for the ultrastructural
examination. Normal Z-lines, intercalated disks (ID), and mitochondria are shown in wild
type. APP-PS1 mice have clear Z-lines, intercalated disks, and organized mitochondria as
well. Diffused Z-lines are found in ANXA5KO and APP-PS1/ANXA5KO mice.
Additionally, sarcomeres in APP-PS1/ANXA5KO mice show disorganized and
disintegrated myofilaments and increased number of mitochondria (Fig. 4-4).

76

Figure 4-4 Ultrastructure of cardiomyocytes in three months-old wildtype, APP-PS1, ANXA5KO
and APP-PS1/ANXA5KO mice.
Papillary muscles were fixed and prepared for electron microscopy examination. Normal Z-lines,
intercalated disks (ID), and mitochondria (M) are shown in wild type mice. APP-PS1 mice have
clear Z-lines and intercalated disks (ID) as wild type. Sarcomeres in ANXA5KO and APP-
PS1/ANXA5KO (APP/ANXKO) mice show diffuse Z-lines. Additionally, cardiomyocytes of APP-
PS1/ANXA5KO mice showed disorganized and disintegrated myofilaments (asterisks) and
increased number of mitochondria.

77

4.3.5 APP-PS1/ANXA5KO mice show lower heart rate and longer QT interval at
young age
In order to see whether these morphological deficiencies affect normal heart function, a
series of electrocardiogram (ECG) was used to analyze the performance of heart in age
matched APP-PS1/ANXA5KO and wild type mice from 10 weeks old. ECG is usually
used to measure the rate and regularity of heartbeats, as well as the size and position of
the chambers, and the presence of any damage to the heart. Heart rate and the QT interval
were collected and used to present the physiological condition of heart. The lower heart
rate has been noticed in APP-PS1/ANXA5KO mice at the beginning, while the QT
interval shows no difference from the one in wild type mice (Fig. 4-5A). At 15 weeks old,
the average heart rate of wild type mice is 468±35.0 bmp (beats per minute), but it goes
down to 398±21.5 bmp which is a 15% decrease as compared to wild type in APP-
PS1/ANXA5KO mice (Fig. 4-5C). Meanwhile, there is a 1.14 fold increase of the QT
interval in APP-PS1/ANXA5KO mice (QT= 51.4±1.2 ms, milliseconds) as compared to
wild type (QT=44.9±1.0 ms) (Fig. 4-5C). Here, it already shows a significant difference
in heart rate and the QT interval between APP-PS1/ANXA5KO and wild type mice. To
16 weeks old, APP-PS1/ANXA5KO mice still have lower heart rate (422±47.3 bmp), and
the QT interval is even longer (52.5±6.3 ms), a 1.16% increase as compared to wild type
(45.2±1.9 ms) (Fig. 4-5D).

78

Figure 4-5 Lower heart rate and higher QT interval are found in APP-PS1/ANXA5KO mice.
(A) Electrocardiogram (ECG) recordings were performed on age matched wild type (WT) and APP-
PS1/ANXA5KO (KO) female mice at different ages. WT, HR=510±54.9, QT=43.7±2.5 (n=5); KO,
R=454±21, QT=44±2 (n=5). (B) ECG recording at 12 weeks. WT, HR=471±78.5, QT=47.0±3.6
(n=5); KO, HR=436±31,QT=49.2±2.3 (n=5). (C) ECG recording at 15 weeks. WT, HR=468±35.0,
QT=44.9±1.0 (n=4); KO, HT=398±21.5, QT=51.4±1.2 (n=4). (D) ECG recording at 16 weeks. WT,
HR=519±26.2, QT=45.2±1.9 (n=4); KO, HR=422±47.3, QT=52.5±6.3 (n=4).

79

4.3.6 Lower heart rate and higher QT interval are found in ANXA5KO and APP-
PS1/ANXA5KO mice
Next, ECG recording was performed on APP-PS1/ANXA5KO and other control mice at
3 months old. Like APP-PS1/ANXA5KO mice, the lower heart rate (HR=437±21.2 bmp)
and higher QT interval (QT=54.5±3.9 ms) were found in ANXA5KO mice which show
abnormalities in cardiac morphology as well. However, APP-PS1/ANXA5KO mice still
have the lowest heart rate (HR=417±50.9 bmp) and the highest QT interval
(QT=56.9±1.2 ms) among these mice. Although APP-PS1 mice have higher QT interval
than wild type mice, the heart rate in these two mice are quite similar (Fig. 4-6).

80

Figure 4-6 Both ANXA5KO and APP-PS1/ANXA5KO mice show lower heart rate and longer QT
interval in ECG recording.
Age matched wild type (WT), APP-PS1 (APP), ANXA5KO (ANX), and APP-PS1/ANXA5KO (KO)
female mice were performed ECG recording at 4 months. WT, HR=558±63.2, QT=44.1±2.9 (n=5);
APP-PS1, HR=563±31.8, QT=51.5±3.4 (n=3); ANXA5KO, HR=437±21.2, QT=54.5±3.9 (n=3),
APP-PS1/ANXA5KO, HR=417±50.9, QT=56.9±1.2 (n=5).

81

4.3.7 The deficiency of heart in APP-PS1/ANXA5KO mice is not caused by cardiac
hypertrophy
The QT interval represents the electric depolarization and repolarization of right and left
ventricles, thus it is usually used as an indicator for ventricular function. It is known that
a prolonged QT interval indicates ventricular deficiency and is a risk factor for sudden
death. However, QT interval is just a general indicator, and many cardiomyopathies such
as hypertrophy cardiomyopathy which is caused by the unusual thickening of heart
muscles, show prolonged QT interval. In order to further identify the pathological
mechanism which causes the heart deficiency in APP-PS1/ANXA5KO mice, we
calculated the ratio of the heart weight versus the body weight in these mice. The heart
weight (HW) /body weight (BW) ratio is about 9mg/g in wild type mice, and APP-PS1
mice is about 8mg/g. ANXA5KO mice have higher heart weight/body weight ratio, but
the ratio in APP-PS1/ANXA5KO mice is also close to 8mg/g (Fig. 4-7). Since APP-
PS1/ANXA5KO mice did not show larger HW/BW ratio, and the unusual thickness of
ventricle muscles was not found in Figure 4-3 either. Therefore, the dysfunction of heart
in APP-PS1/ANXA5KO mice is not caused by hypertrophy.
82

0.00
2.00
4.00
6.00
8.00
10.00
12.00
14.00
WT APP ANX A5 DO
HW/BW ratio (mg/g)
Figure 4-7 The comparison of heart weight/body weight ratio between APP-PS1/ANXA5KO and
other mice.
4 months old APP-PS1/ANXA5KO and other mice were anesthetized and weighted for body weight.
After ECG recording, hearts were taken out from euthanized mice and weighted. The heart
weight/body weight ratio of APP/ANXA5KO (DO) mice is not significantly different from other
mice. Wild type (WT, n=5); APP-PS1 (APP, n=3); ANXA5KO (ANXA5, n=3); APP/ANXA5KO
(DO, n=5).

83

4.3.8 The membrane associated αB-crystallin are increased in ANXA5KO and APP-
PS1/ANXA5KO mice
αB-crystallin is a small heat shock protein which functions as a molecular chaperone. It is
initially identified as a major structural protein in lens(Piatigorsky 1984), and later found
abundantly in cardiac and skeletal muscles(Iwaki, Kume-Iwaki et al. 1989, Longoni,
Lattonen et al. 1990). Recently, the upregulation of αB-crystallin was observed in many
cardiac disorders including familial hypertrophic cardiomyopathy and desmin-related
cardiomyopathy (Hwang, Dempsey et al. 1997, Arbustini, Morbini et al. 1998, Xiao and
Benjamin 1999). Therefore, we want to see whether this upregulation also happens in
APP-PS1/ANXA5KO mice. Cardiac proteins were extracted and separated into
cytoplasmic and membrane fractions. Cardiac expression of transgenic human APP gene
was detected in both APP-PS1 and APP-PS1/ANXA5KO mice as expected (Fig. 4-8A).
αB-crystallins were abundantly shown in the cytoplasm fraction and the amount is similar
in all four groups. However, the membrane associated αB-crystallins were significantly
increased in ANXA5KO and APP-PS1/ANXA5KO as compared to APP-PS1 and wild
type mice (Fig. 4-8B, 4-8C). In addition, the perinuclear localization of αB-crystallin was
shown in APP-PS1 mice with the immunocytochemistry, and this is even more evident in
the cardiomyopathies from ANXA5KO and APP-PS1/ANXA5KO mice as compared to
the diffused basal signal in wild type mice (Fig. 4-8D).

84

Figure 4-8 Increased membrane associated αB-crystallin expression in young ANXA5KO and APP-
PS1/ANXA5KO mice.

Heart samples were collected from 1 month-old mice. Proteins were extracted with Tris-HCl buffer
containing 1% Triton X-100 and supernatants were collected for Western blot analysis.
(A) The human amyloid precursor protein (APP) transgene is expressed in both APP-PS1 and APP-
PS1/ANXA5KO(APP/ANXKO) mice. (B) Membrane associated αB crystallin is significantly
increased in 1 month-old ANXA5KO and APP-PS1/ANXA5KO(APP/ANXKO) mice. APP-PS1 mice
have similar amount of αB crystallin protein as wildtype. (C) Quantification of Western blot from
membrane fraction results (n=4 for each group) shows significantly increased of αB crystallin in
ANXA5KO and APP-PS1/ANXA5KO mice. Band intensities are normalized against actin as a
loading control. Ratio is expressed as fold-change over mean WT value ± S.E. (D) Ventricle muscles
were collected for immunostaining. Wildtype tissue showed diffuse basal staining. In contrast, many
cells in the APP-PS1 section showed perinuclear localization of αB-crystallin. This is even more
evident in the tissue from ANXA5KO and APP-PS1/ANXA5KO(APP/ANXKO) mice.

85

4.4 Discussion
The human APP gene, located on chromosome 21, was first identified in 1987 by several
laboratories independently. Since the discovery of APP, a number of physiological roles
have been attributed to the molecule, but its actual functions remain unclear. Aβ is the
proteolytic product of APP through an enzymatic process of β- and γ-secretase, and the
increased Aβ deposition in brain is the primary cause of Alzheimer’s disease. Previously,
annexin A5, an abundantly expressed Ca
2+
-dependent membrane binding protein has
been reported to offer protection from the cytotoxicity of Aβ in vitro. In order to elucidate
this function of annexin A5, APP-PS1/ANXA5KO mice were generated and we found
that the number of brain Aβ plaques in APP-PS1/ANXA5KO mice is similar to APP-
PS1dE9 mice at young age. However, the early death was noticed in APP-
PS1/ANXA5KO mice. At 6 months of age, the survival rate of APP-PS1/ANXA5KO
mice is 46.67% in male and only 34.83% in female as compared to APP-PS1dE9 and
ANXA5KO mice (Fig. 4-2). Further studies indicated that the sudden death is caused by
a severe cardiac deficiency. Diffused Z-lines, disorganized myofilaments, and increased
number of mitochondria were shown in the cardiomyocytes in APP-PS1/ANXA5KO
mice (Fig. 4-4). Lower heart rates and higher QT interval were reported from functional
analysis (Fig. 4-5, 4-6). APP-PS1/ANXA5KO mice have the similar ratio of heart
weight/body weight as wild type, thus the cardiac hypertrophy is excluded as the cause of
this heart disorder (Fig. 4-7). αB-crystallin, a small heat shock protein functioning as
molecular chaperon, was increased in the membrane fraction of cardiac protein extract
86

(Fig. 4-8). Therefore, we speculate that the presence of annexin A5 may protect
cardiomyocytes from the damages caused by mechanical stress and Aβ cytotoxicity.
Annexins A1, A2, A4, A5, A6, and A7 have been detected in cardiomyocytes, and
annexin A5 and A6 are the most abundant among them. They can be co-expressed in the
certain cells but with their own localization. Annexin A5 is associated to sacrolemma
(SL), intercalated disks, and T-tubules connected to the terminal cisternae of
sarcoplasmic reticulum (SR) where excitation-contraction coupling takes place (Doubell,
Lazure et al. 1993, Luckcuck, Trotter et al. 1997, Jans, de Jong et al. 1998, Trouve, Legot
et al. 1999, Benevolensky, Belikova et al. 2000, Matteo and Moravec 2000). It has also
been described as associated to the Z lines (Wang, Rahman et al. 1995). Diffused Z-lines
showing in ANXA5KO and APP-PS1/ANXA5KO mice (Fig. 4-4) may result from the
absence of annexin A5. Annexins are Ca
2+
-binding proteins which may be involved in
regulating Ca
2+
signaling. In cardiomyocytes, Ca
2+
-handling proteins contribute to the
rise and fall of the intracellular free Ca
2+
that induces contraction and relaxation. Annexin
A5 may be involved in this modulation by functioning as a Ca
2+
channel or as an
essential signaling intermediate in a Ca
2+
influx pathway (Kirsch, Nah et al. 1997,
Kubista, Hawkins et al. 1999). Therefore, the functional deficiency which represented as
lower heart rate and elongated QT interval in ANXA5KO and APP-PS1/ANXA5KO
mice (Fig. 4-6) may be caused by the dysregulation of Ca
2+
homeostasis.
Another distinctive property of annexin A5 and several other annexins is to self-assemble
into two-dimensional ordered arrays on membrane binding (Pigault, Follenius-Wund et al.
87

1994, Voges, Berendes et al. 1994, Reviakine, Bergsma-Schutter et al. 1998, Gerke,
Creutz et al. 2005). This is shown in cryo-electron and atomic-force microscopy images
of annexin A5 bound to phosphatidylserine-containing bilayers (Pigault, Follenius-Wund
et al. 1994, Voges, Berendes et al. 1994, Reviakine, Bergsma-Schutter et al. 1998, Oling,
Santos et al. 2000). In such images, the protein forms two-dimensional (2D) crystals on
the bilayers, with annexin A5 trimers representing the principal building block. In vitro
studies have indicated that annexin A5 2D arrays rigidify lipid monolayers and reduce the
lateral diffusion of phospholipids (Saurel, Cezanne et al. 1998, Venien-Bryan, Lenne et al.
1998). Bouter et al. proposed that the formation of annexin A5 2D array at a ruptured
membrane strengthens the membrane and prevents the expansion of the tear by
counteracting membrane tension due to cytoskeleton attachment (Bouter, Gounou et al.
2011). Given the abundance of annexin A5 in some cells and its localization to the
plasma membrane, 2D assemblies of the protein might function in promoting membrane
repair, stabilizing certain plasma-membrane structures, and/or in changing membrane
curvature and therefore cell shape (Gerke, Creutz et al. 2005).
The neurotoxicity of Aβ involves the disruption of cellular calcium homeostasis and the
generation of reactive oxygen species. Aβ may disrupt calcium regulation by triggering
oxidative stress which damages membrane calcium pumps and enhances calcium influx
through ionotropic glutamate receptors and voltage-dependent channels (Mattson 2004).
On the other hand, toxic Aβ (1-42) oligomers can initiate lipid peroxidation during AD
progression (Schubert, Behl et al. 1995, Lambert, Barlow et al. 1998, Selkoe 2001,
Butterfield, Castegna et al. 2002, Butterfield and Lauderback 2002, Butterfield, Reed et
88

al. 2007) by membrane integration and/or surface binding (Hertel, Terzi et al. 1997,
Kayed, Sokolov et al. 2004). Oligomer-induced peroxidation has the potential to disrupt
phosphatidylinositol-4,5-bisphosphate metabolism (Berman, Dall'Armi et al. 2008),
increase bilayer conductance through alternation of dielectric structure (Strissel, Sokolov
et al. 2006) and increase bilayer permeability (Arispe and Doh 2002, Kayed, Sokolov et
al. 2004, Demuro, Mina et al. 2005). Given the potential membrane repair function of
annexin A5 and the importance of membrane in maintaining Ca
2+
homeostasis, we
hypothesize that the severe cardiac defect seen in APP/ANXA5KO mice is resulted from
the membrane damage caused by Aβ cytotoxicity. Although cardiomyocytes in APP-
PS1dE9 mice suffer from the Aβ cytotoxicity, no membrane damages are made because
of the repairing function of annexin A5. However, the absence of annexin A5 in
APP/ANXA5KO mice cannot provide a proper repairing mechanism for the Aβ damaged
membrane which could disrupt Ca
2+
homeostasis, thus develop severe cardiac
dysfunction. In order to prove our hypothesis, pSIVA, an annexin-based biosensor (Kim,
Chen et al. 2010), was injected into ventricles to monitor the cardiac membrane structure
in APP/ANXA5KO mice. Illuminated fluorescence was generated from the membrane
binding of p-SIVA which indicates severe membrane damage in hearts from APP-
PS1/ANXA5KO (data not shown). This supports our hypothesis that cell membranes are
severely damaged in APP-PS1/ANXA5KO mice and the membrane repairing mechanism
of annexin A5 is critical for cardiac function in APP-PS1dE9 mice. Cardiomyocytes are
exposed to constant mechanical stress which often causes the tears in their plasma
membranes (McNeil and Steinhardt 2003). ANXA5KO mice show abnormal cardiac
89

phenotype and function as well, but other annexins may be able to compensate the
function of annexin A5 in repairing the mechanical damaged membrane and the
unbalance of Ca
2+
homeostasis. Therefore, the cardiac disorder does not affect the
survival of ANXA5KO mice.
When Aβ aggregation occurs at the cell membrane, Aβ-induced oxidative stress leads to
lipid peroxidation and the consequent generation of 4-hydroxynonenal (4HNE), a
neurotoxic aldehyde that covalently modifies proteins on cysteine, lysine, and histidine
residues (Mattson 1997). Those proteins modified by this Aβ-induced oxidative stress
include membrane transporters (ion-motive ATPase, a glucose transporter and a
glutamate transporter), receptors, GTP-binding proteins, and ion channels (VDCC:
voltage-dependent chloride channel; NMDAR: N-methyl-D-aspartate receptor)
(Butterfield, Drake et al. 2001). Aβ can also cause mitochondrial oxidative stress and
dysregulation of calcium homeostasis, resulting in impairment of the electron transport
chain, elevated production of ROS, and decreased production of ATP (Mattson 2004). In
order to maintain the energy level in cardiomyocytes without a proper membrane
repairing mechanism, elevating the number of mitochondria may be used as a strategy to
overcome the decreased production of ATP caused by Aβ at the early stage. Thus, an
increased number of mitochondria were found in APP-PS1/ANXA5KO mice (Fig 4-5).
αB-crystallin is a small heat shock protein highly expressed in cardiac and skeletal
muscles. It is induced by cellular stress and functions as a molecular chaperone. The
upregulation of αB-crystallin was found in the membrane fraction of the cardiac protein
90

extract from ANXA5KO and APP-PS1/ANXA5KO mice (Fig. 4-8B). The increased
number of αB-crystallin in ANXA5KO mice may be caused by the cellular stress. On the
other hand, the significant elevation of αB-crystallin in APP-PS1/ANXA5KO mice
results from the accumulation of Aβ and the impairment of cell membrane. αB-crystallin
is present in brain tissues as well, and previously study has shown that its expression is
elevated in Alzheimer’s disease (Shinohara, Inaguma et al. 1993). The increase in levels
of αB-crystallin found in AD reveals an important role of the small heat-shock proteins in
AD. Here, we found that APP-PS1dE9 mice have similar level of cardiac αB-crystallin as
wild type. The presence of annexin A5 provides a functional membrane repairing
mechanism which is effective in against amyloid neurotoxicity. Therefore, APP-PS1dE9
mice do not develop cardiac deficiency, and this also further support the hypothesis that
annexin A5 can protect cells from amyloid cytotoxicity. In summary, the cardiac function
in the APP-PS1dE9 AD model is modulated by annexin A5.

91

4.5 Summary
Alzheimer’s disease (AD) is an age-associated neurodegenerative disorder characterized
by the presence of neurofibrillary tangels, neuritic plaques, synapse damage, and
neuronal cell loss. Although the pathology of AD is complex, the amyloid cascade
hypothesis suggests that the accumulation of Aβ driven by its increased generation over
its clearance is the primary cause of the subsequent formation of those clinical dementias
in AD pathogenesis. Previously, annexin A5, an abundantly expressed Ca
2+
-dependent
membrane binding protein, has been reported to offer protection from the cytotoxicity of
Aβ in vitro. Therefore, we speculate that annexin A5 may play a role in preventing the
Aβ cytotoxicity in AD. We generated APP-PS1/ANXA5KO mice by breeding APP-
PS1dE9 mice, a widely used mouse model for Alzheimer’s disease, with ANXA5KO
mice. No overt difference in the number of brain plaques were seen between the APP-
PS1dE9 and the APP-PS1/ANXA5KO mice. Interestingly, APP-PS1/ANXA5KO mice
develop severe heart deficiency and die from sudden death at an early age. Both
ANXA5KO and APP-PS1/ANXA5KO mice have significantly lower heart rate and
higher QT interval than wild type mice, as well as the upregulation of αB-crystallin in
cardiac tissues. These all suggest that the presence of annexin A5 may protect
cardiomyocytes from the damages caused by mechanical stress and Aβ. The heart defect
seen in the ANXA5KO and APP-PS1/ANXA5KO mice may be related to the recent
discovered role of annexin A5 in promoting membrane repair following mechanical
disruption to the plasma membrane. Together, our results show that cardiac function in
the APP-PS1dE9 AD model is modulated by annexin A5.
92

4.6 Supplementary Data
In this project, we demonstrated that annexin A5 has the function to protect
cardiomyocytes from Aβ toxicity with APP-PS1/ANXA5KO mouse model. Since brain
is the organ presenting most significant features in the pathology of Alzheimer’s disease,
it is reasonable to speculate that whether the loss of annexin A5 causes more severe brain
damage in the disease pathogenesis. Hence, in the other part of this project, we also spent
some time in analyzing brain samples from APP-PS1/ANXA5KO and other mice.

4.6.1 APP-PS1/ANXA5KO mice do not have more β-amyloid plaques in brain
3 months old APP-PS1/ANXA5KO and age matched control mice were anesthetized and
prepared for perfusion. Blood was washed out with PBS first and organs were fixed with
4% paraformaldehyde. Brains were then taken out carefully and prepared for vibrato-
section. Whole brain was cut and parts of sections selected from the anterior, middle and
posterior were then stained with thioflavin-S, a dye known to bind mature amyloid fibrils,
to detect the plaques in brain samples. Many plaques were shown by thioflavin-S in both
APP-PS1 and APP-PS1/ANXA5KO mice at this young age (Figure 4-9A). However, if
we look at whole brain, no matter from the anterior, middle, or posterior part, there is no
significant difference in the number of plaques in these two mice (Figure 4-9B).

93

APP/ANXKO ANXA5KO
APP-PS1 WT
A
B
Figure 4-9 Both APP-PS1 and APP-PS1/ANXA5KO mice show numbers of amyloid plaques in brain
at early age.
(A) Brain vibratome section collected from APP-PS1 and APP-PS1/ANXA5KO mice both show
numbers of thioflavin-S positive stains indicating the presence of amyloid plaques. (B) The view of
whole section selected from the middle part of brain further reveal the high density of plaques in APP-
PS1 and APP-PS1/ANXA5KO mice. However, APP-PS1/ANXA5KO mice do not develop more
plaques as compared to APP-PS1 mice.
94

4.6.2 Older APP-PS1/ANXA5KO mice have more monomeric Aβ and amyloid
oligomers in brain as compared to APP-PS1 mice
Next, brain samples collected from old and young mice were homogenized with
detergent containing buffer or 80% formic acid, and proteins were extracted according to
their solubility. The level of monomeric Aβ and amyloid aggregation were analyzed by
Western blot. The human APP gene is expressed in both APP-PS1 and APP-
PS1/ANXA5KO mice as expected. In the formic acid extraction, more monomeric Aβ
were showing in the older APP-PS1/ANXA5KO mice as compared to APP-PS1 mice (4-
10A). On the other hand, in the results shown with OC antibody, a structure-specific
antibody used to detect amyloid fibrils; APP-PS1/ANXA5KO mice apparently have more
amyloid aggregations in brain rather than APP-PS1 mice at elder age (4-10B, C).

95

APP
amyloid oligomers
actin
A
B
96

0
0.5
1
1.5
APP APP/ANXKO
oligomeric A β level
C
Figure 4-10 APP-PS1/ANXA5KO mice have more monomeric Aβ and amyloid aggregations in
brain at elder age.
(A) Transgenic human APP gene was expressed in brains from both APP-PS1 and APP-
PS1/ANXA5KO mice. The level of monomeric Aβ, extracted with 80% formic acid, is higher in
elder APP-PS1/ANXA5KO mice. (B) On the other hand, APP-PS1/ANXA5KO mice also have more
amyloid oligomers in brain at old age as compared to APP-PS1 mice. (C) Quantification results from
three individual experiments show the level of amyloid oligomers in APP-PS1/ANXA5KO mice is
about 1.25 fold higher than age matched APP-PS1 mice.

97

Conclusions
Exploring the magic of life and the physiological mechanisms composing creatures are
always my dreams since childhood. In years of my Ph. D. career, I focused on how
proteins modulating cellular functions. In my two totally different projects, I successfully
identify new function of arrstin-1 and annexin A5. Furthermore, possible mechanisms are
also established to explain how these two proteins involve in regulating cellular functions.

In the first study, we demonstrated PP2A as rhodopsin phosphatase by employing intact
retina and PP2A inhibitor, fostriecin. This part of results confirms the findings shown in
previous studies and excludes the potential phosphatase activity contamination which has
been questionable for years. More interesting, we discovered a new role of arrestin-1 in
positively regulating rhodopsin dephosphorylation by modulating the distribution of
PP2A subunits in rod outer segment. Arrestin-1 was recognized as an inhibitor in
rhodopsin dephosphorylation process. However, by using arrestin-1 knockout mice, our
findings reverse arrestin-1 from an inhibitor to a positive regulator, and additional
experiments further demonstrate that the distribution of PP2A subunits, especially A
subunit, may be controlled by arrestin-1. Here, we not only reported a novel function of
arrestin-1 in addition to terminating rhodopsin activity, but also established a possible
mechanism which was not shown before. Although these results are very exciting, some
experiments may need to be done for more support. For example, using intact retinal as
material is better than purified opsin; however, it is still not able to reflect the real
98

physiological situation inside eyes. Currently, PP2A conditional knockout mice are
available, and if the fact of PP2A as rhodopsin phosphatase could be demonstrated in this
mouse model, that will be a very strong support to our findings. On the other hand,
revealing the regulation mechanism in controlling PP2A subunit distribution will be the
major focus in the future. Identifying protein interactions between arrestin-1 and PP2A A
subunit by biochemistry strategies will be the beginning and with more information
available we will gradually establish the complete mechanism of rhodopsin
dephosphorylation.

Alzheimer’s disease (AD) is the most common age-associated dementia that leads to
cognitive, memory and behavioral impairments in elder population. Due to the lack of
effective treatments and the uncertain mechanisms of pathogenesis, AD is one of the
most extensively studied amyloid-related disorders. In the model mice APP-
PS1/ANXA5KO we generated, the absence of annexin A5 which has been reported as a
protector against β-amyloid toxicity does not accelerate the accumulation of Aβ
aggregations in brain at early age. However, short life time is remarkably noticed in these
mice. Cardiomyocytes in APP-PS1/ANXA5KO and annexin A5 knockout mice show
severe damages. Electrocardiogram also reported cardiac dysfunction in these two strains
of mice. Previous study in the development of annexin A5 knockout mice showed no
significant alternation in viability, fertility, metabolism, and skeletal functions. Here, we
first reported cardiac deficiencies in annexin A5 knockout mice, and these results
99

provides a novel and strong evidence for the role of annexin A5 in cardiac development.
Given carrying cardiac disorders, annexin A5 knockout mice have normal viability, but
APP-PS1/ANXA5KO mice do not. Therefore, this study not only demonstrated the
function of annexin A5 in resisting β-amyloid toxicity in an animal model, but also
revealed the importance of annexin A5 in maintaining cardiac function. Hearts have been
shown as a possible target for β-amyloid toxicity, and one of the damages caused by Aβ
aggregations is membrane impairment. Currently, annexin A5 has been reported
involving in the process of membrane repairing. Taking together, we hypothesize that the
way provided by annexin A5 to support cardiomyocytes against the damage caused by
Aβ aggregations is a strong membrane repairing mechanism. In order to test this
hypothesis, the presence of membrane damage in APP-PS1/ANXA5KO cardiac tissues is
the first question needed to be answered. Recently, an annexin-derived biomarker, p-
SIVA has been developed with the ability to detect membrane impairment. With the help
of this molecule, we will be able to provide more evidence to support our hypothesis.

100

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Abstract (if available)

Abstract In the past several years, I focused on two different projects for my PhD studies. The first project is to achieve a better understanding of the mechanism of rhodopsin dephosphorylation. Protein phosphatase 2A (PP2A) has been recognized as the phosphatase responsible for rhodopsin dephosphorylation for years. However, due to the absence of in vivo evidence, the role of PP2A in regulating rhodopsin regeneration is still questionable. This work not only clarifies the position of PP2A in modulating rhodopsin function under physiological condition, but also presents novel findings in the detail mechanism. Previous isoelectric focusing results revealed that arrestin-1 may play a positive role in modulating rhodopsin dephosphorylaiton. Taking advantage of arrestin-1 knockout mice, we isolated rod outer segments to compare how arrestin-1 influences the distribution of PP2A subunits after light stimulation. Western blot results revealed that the movement of PP2A scaffolding subunit is regulated by arrestin-1 protein. In addition to inactivating rhodopsin, arrestin-1 may mediate rhodopsin dephosphorylation by modulating the cellular localization of PP2A in rod photoreceptors. ❧ Identifying the role of annexin A5 in amyloidogenesis is the goal of my second project. Annexin A5 is an abundant protein without clear physiological function. Earlier studies suggested a protection effect of annexin A5 in against amyloid toxicity. We generated APP-PS1/ANXA5KO mice by breeding APP-PS1dE9, a widely used mouse model for Alzheimer’s disease, under annexin A5 knockout background. No overt difference in the number of brain Aβ plaques were seen between the APP-PS1dE9 and the APP-PS1/ANXA5KO mice. Interesting, sudden death was noticed in APP-PS1/ANXA5KO mice at early age. Morphology shown in trichrome staining and ultrastructural examination revealed severe damages in cardiomyocytes from ANXA5KO and APP-PS1/ANXA5KO mice. These two strain of mice also exhibited lower heart rates and higher QT interval in electrocardiogram analysis. Furthermore, the level of αB-crystallin, a chaperon molecule, is significantly increased in the membrane fraction of cardiac extraction, and a specific perinuclear distribution of αB-crystallin is also observed by immunocytochemistry in ANXA5KO and APP-PS1/ANXA5KO mice. Therefore, annexin A5 may play an important role in regulating cardiac function in Alzheimer’s disease model mice.

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Asset Metadata

Creator Hsieh, Chia-Ling (author)

Core Title Protein phosphatase 2A and annexin A5: modulators of cellular functions

School Keck School of Medicine

Degree Doctor of Philosophy

Degree Program Genetic, Molecular and Cellular Biology

Publication Date 04/30/2015

Defense Date 03/21/2013

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag Alzheimer's disease,annexin A5,OAI-PMH Harvest,protein phosphatase 2A,rhodopsin

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Language English

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Advisor Chen, Jeannie (committee chair), Hinton, David R. (committee member), Langen, Ralf (committee member)

Creator Email chialing@usc.edu,irene.hs@yahoo.com.tw

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c3-248384

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