Department of Applied Chemistry and Microbiology, Division of Nutrition
and
Department of Biological and Environmental Sciences, Division of Genetics
University of Helsinki
Minna Pekkinen
REGULATION OF OSTEOBLAST DIFFERENTIATION AND GENE EXPRESSION BY
PHOSPHODIESTERASES AND BIOACTIVE PEPTIDES
ACADEMIC DISSERTATION
To be presented, with the permission of the Faculty of Biosciences of the University of
Helsinki, for public criticism in the large lecture hall, Bulevardi 18, Helsinki, on 28 March
2008, at 12 noon.
HELSINKI 2008
Supervisors
Acting Professor Christel Lamberg-Allardt
Department of Applied Chemistry and Microbiology
University of Helsinki
Finland
Dr. Mikael Ahlström
Department of Applied Chemistry and Microbiology
University of Helsinki
Finland
Reviewers
Acting Professor Petri Lehenkari
Department of Anatomy
University of Oulu
Finland
Adjunct Professor Anitta Mahonen
Institute of Biomedicine
Deparment of Medical Biochemistry
University of Kuopio
Finland
Opponent
Professor Kalervo Väänänen
Institute of Biomedicine
Department of Anatomy
University of Turku
Finland
© Minna Pekkinen 2008
Yliopistopaino
ISNB 978-952-92-3528-5 (paperback)
ISNB 978-952-10-4592-9 (PDF, http//:ethesis.helsinki.fi)
2
Contents
Abstract… … … … … … … … … … … … … … … … … … … … … … … … … … … … … … 6
List of original publications… … … … … … … … … … … … … … … … … … … … … … .8
Abbreviations… … … … … … … … … … … … … … … … … … … … … … … … … … … ...9
1. Introduction… … … … … … … … … … … … … … … … … … … … … … … … … .11
2. Review of the literature...................................................................................13
2.1 Bone biology… … … … … … … … … … … … … … … … … … … … … … … ..13
2.1.1 Bone tissue and bone cells.............................................................13
2.1.2 Osteoblastic differentiation............................................................13
2.1.3 Osteoblastic regulation of osteoclast differentiation......................17
2.1.4 Bone remodelling and its regulation...............................................18
2.1.5 Unbalanced bone remodelling: osteoporosis … … … … … … .........19
2.1.6 Glucocorticoids and bone.............................................................. .20
2.2 Cyclic nucleotide signalling and phosphodiesterases (PDEs).....................21
2.2.1 Cyclic adenosine monophosphate (cAMP) signalling.....................21
2.2.2 Cyclic guanosine monophosphate (cGMP) signalling....................22
2.2.3 cAMP and cGMP PDEs...................................................................23
2.2.3.1 PDE1 family............................................................................27
2.2.3.2 PDE2 family...........................................................................29
2.2.3.3 PDE3 family...........................................................................29
2.2.3.4 PDE4 family...........................................................................30
2.2.3.5 PDE5 family...........................................................................31
2.2.3.6 PDE6 family...........................................................................31
2.2.3.7 PDE7 family...........................................................................31
2.2.3.8 PDE8 family...........................................................................32
2.2.3.9 PDE9 family...........................................................................32
2.2.1.10 PDE10 family.........................................................................33
2.2.1.11 PDE11 family.........................................................................33
2.2.4 Regulation of activity and compartmentalization of PDEs.............33
2.2.5 Inhibitors of PDEs...........................................................................34
2.2.6 Bone metabolism and PDE inhibitors… … … ...… … … … ..............34
2.3 Other signal transduction pathways associated with PDEs...................... ..35
2.3.1 Exchange protein directly activated by cAMP (EPAC)
and PDEs........................................................................35
2.3.2 Mitogen-activated protein kinase (MAPK) and PDEs....................36
2.3.3 PKC and PDEs................................................................................36
3
2.4 Bioactive peptides........................................................................................36
2.4.1 Bioactive peptides in bone...............................................................38
3. Aims of the study… … … … … … … … … … … … … … … … … … … … … … ..… .39
4. Materials and methods… … … … … … … … … … … … … … … … … … … … .......40
4.1 Cell culture...................................................................................................40
4.1.1 Osteosarcoma cell lines.................................................................40
4.1.2 Human osteoblasts........................................................................40
4.1.3 Human mesenchymal stem cell (hMSC)-derived osteoblasts.......40
4.2 PDE inhibitors..............................................................................................41
4.3 Primers..........................................................................................................41
4.4 cAMP analyses.............................................................................................43
4.5 Q-sepharose chromatography.......................................................................43
4.6 Enzyme activity analyses.............................................................................44
4.6.1 Assay of PDE activity...................................................................44
4.6.2 Assay of ALP activity...................................................................44
4.7 PDE silencing...............................................................................................44
4.8 Isoleucine-proline-proline (IPP), valine-proline-proline (VPP) and leucinelysine-proline (LKP) tripeptides...................................................................45
4.9 Cell proliferation analyses............................................................................45
4.10 Mineralization analyses..............................................................................45
4.11 mRNA expression analyses........................................................................46
4.11.1 RNA isolation..............................................................................46
4.11.2 cDNA microarrays.......................................................................46
4.11.3 cDNA synthesis...........................................................................46
4.11.4 Semi-quantitative reverse transcriptase polymerase chain
reaction (RT-PCR).......................................................................47
4.11.5 Quantitative Real-time PCR (qRT-PCR).....................................47
5. Results and discussion… ..… … … … … … … … … … … … … … … … … … … … ..48
5.1 Identification of PDE families in human osteoblastic cells.............48
5.2 Identification of PDE subtypes in human osteoblastic cells............51
5.3 PDE7 and PDE8A silencing by siRNAs..........................................52
5.4 PDE7 silencing has potential for enhancing osteogenic gene
expression and differentiation in hMSC-derived osteoblasts...........53
5.5 PDE7 inhibition enhances cAMP signalling, differentiation and
mineralization in hMSC-derived osteoblasts....................................56
5.6 Down-regulation of cAMP-PDE by dexamethasone in human
osteosarcoma cells...................................................................................58
5.7 Influence of dexamethasone on PDE subtype mRNA
expression in MG-63 and SaOS-2 cells............................................58
5.8 Effects of PDEs in osteoblasts..........................................................59
4
5.9 Influences of bioactive peptides IPP, VPP and LKP on gene
expression and proliferation in hMSC-derived osteoblasts.............60
6. Conclusions… … … … … … … … … … … … … … … … … … … … … … .................63
7. Acknowledgements… … … … … … … … … … … … … … … … … … … … … … … .65
8. References… … … … … … … … … … … … … … … … … … … … … … … … … … 66
5
Abstract
Bone is a mineralized tissue that enables multiple mechanical and metabolic functions to be
carried out in the skeleton. Bone contains distinct cell types: osteoblasts (bone-forming cells),
osteocytes (mature osteoblast that embedded in mineralized bone matrix) and the osteoclasts
(bone-resorbing cells). Remodelling of bone begins early in foetal life, and once the skeleton
is fully formed in young adults, almost all of the metabolic activity is in this form. Bone is
constantly destroyed or resorbed by osteoclasts and then replaced by osteoblasts. Many bone
diseases, i.e. osteoporosis, also known as bone loss, typically reflect an imbalance in skeletal
turnover.
The cyclic adenosine monophosphate (cAMP) and the cyclic guanosine monophosphate
(cGMP) are second messengers involved in a variety of cellular responses to such
extracellular agents as hormones and neurotransmitters. In the hormonal regulation of bone
metabolism, i.e. via parathyroid hormone (PTH), parathyroid hormone-related peptide
(PTHrp) and prostaglandin E2 signal via cAMP. cAMP and cGMP are formed by adenylate
and guanylate cyclases and are degraded by phosphodiesterases (PDEs). PDEs determine the
amplitudes of cyclic nucleotide-mediated hormonal responses and modulate the duration of
the signal. The activities of the PDEs are regulated by multiple inputs from other signalling
systems and are crucial points of cross-talk between the pathways.
Food-derived bioactive peptides are reported to express a variety of functions in vivo. The
angiotensin-converting enzymes (ACEs) are involved in the regulation of the specific
maturation or degradation of a number of mammalian bioactive peptides. The bioactive
peptides offer also a nutriceutical and a nutrigenomic aspect to bone cell biology.
The aim of this study was to investigate the influence of PDEs and bioactive peptides on the
activation and the differentiation of human osteoblast cells. The profile of PDEs in human
osteoblast-like cells and the effect of glucocorticoids on the function of cAMP PDEs, were
investigated at the mRNA and enzyme levels. The effects of PDEs on bone formation and
osteoblast gene expression were determined with chemical inhibitors and siRNAs (short
interfering RNAs). The influence of bioactive peptides on osteoblast gene expression and
proliferation was studied at the mRNA and cellular levels.
This work provides information on how PDEs are involved in the function and the
differentiation of osteoblasts. The findings illustrate that gene-specific silencing with an
RNA interference (RNAi) method is useful in inhibiting, the gene expression of specific
PDEs and further, PDE7 inhibition upregulates several osteogenic genes and increases bALP
activity and mineralization in human mesenchymal stem cells-derived osteoblasts. PDEs
appear to be involved in a mechanism by which glucocorticoids affect cAMP signaling. This
may provide a potential route in the formation of glucocorticoid-induced bone loss, involving
the down-regulation of cAMP-PDE. PDEs may play an important role in the regulation of
6
osteoblastic differentiation. Isoleucine-proline-proline (IPP), a bioactive peptide, possesses
the potential to increase osteoblast proliferation, differentiation and signalling.
7
LIST OF ORIGINAL PUBLICATIONS
This thesis is based on the following original publications, which are referred to in the text by
their Roman numerals (I-IV):
I
Ahlström, M., Pekkinen M., Huttunen, M., Lamberg-Allardt C. (2005): Cyclic
nucleotide phosphodiesterases (PDEs) in human osteoblastic cells; effect of
PDE inhibition on cAMP accumulation. Cellular & Molecular Biology Letters
10(2): 305-319.
II
Pekkinen, M., Ahlström, M.E.B, Riehle, U., Huttunen, M.M, Lamberg-Allardt
C.J.E. (2007): Effects of phosphodiesterase 7 Inhibition by RNA interference on
the gene expression and differentiation of human mesenchymal stem cellderived osteoblasts. Bone (accepted).
III
Ahlström, M.*, Pekkinen, M.*, Huttunen, M., Lamberg-Allardt, C. (2005):
Dexamethasone
down-regulates cAMP-phosphodiesterase
in
human
osteosarcoma cells. Biochemical Pharmacology 69: 267-275.
IV
Huttunen, M.M.*, Pekkinen, M.*, Ahlström, M.E.B, Lamberg-Allardt C. J.E.
(2007): Effects of bioactive peptides isoleucine-proline-proline (IPP), valineproline-proline (VPP) and leucine-lysine-proline (LKP) on gene expression of
osteoblasts, differentiated from human mesenchymal stem cells. British Journal
of Nutrition 98: 780-788.
*These authors contributed equally.
These publications have been reproduced with the kind permission of their
copyright holders. In addition, some unpublished material is presented.
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ABBREVIATIONS
aa
AC
ACEs
AKAP
ATF1
ATP
bALP
BMPs
BRL 50481
BSA
BSP
CaM
cAMP
Cdc42
cGMP
CRE
CREB
CREM
Col1
Dex
dsRNA
ECM
EHNA
EPAC
ERK
FCS
FGF
FSP-1
GAPDH
GCs
hMSC
IBMX
IGF-I
Ihh
IPP
Km
bovine serum albumin
bone sialoprotein
calmodulin
cyclic adenosine monophosphate
cell division cycle 42
cyclic guanosine monophoshate
cAMP response element
cAMP response element-binding protein
cAMP response-element modulator protein
collagen type 1
dexamethasone
double-stranded RNA
extracellular matrix
erythro-9-[2-hydroxy-3-nonyl] adenine
exchange protein directly activated by cyclic AMP
extracellular signal-regulated kinase
foetal calf serum
fibroblast growth factor
fibroblast-specific protein 1
glyceraldehyde-3-phosphate dehydrogenase
glucocorticoids
human mesenchymal stem cell
3-isobutyl-1-methylxanthine
insulin-like growth factor 1
Indian hedgehog
isoleucine-proline-proline
Michaelis constant
LKP
MAPK
MC
mRNA
M-SCF
leucine-lysine-proline
mitogen-activated protein kinase
mesangial cell
messenger RNA
cytokine macrophage colony-stimulating factor
amino acid
adenylyl cyclase
angiotensin-coverting enzymes
A-kinase anchoring protein
activating transcription factor 1
adenosine triphosphate
bone-specific alkaline phosphatase
bone morphogenetic proteins
3-(N,N-dimethylsulfonamido)-4-methyl-nitrobenzene
9
8-MMX
MSC
M-SCF
MyoD
NO
OCN
OPG
OPN
Osx
PAS
PCA
PDE
PGE2
8-methoxymethyl-3-isobutyl-1-methylxanthine
bone marrow stroma mesenchymal stem cell
macrophage colony-stimulating factor
transcription factor for myogenesis
nitric oxide
osteocalcin
osteoprotegerin
osteopontin
osterix
per-arnt-sim
principal component analysis
phosphodiesterase
prostaglandin E2 prostaglandins
PGs
PKA
PKB
PKC
PKG
PKI
PLC
PPAR 2
PTH
PTHR
PTHrp
RANK
RANKL
Rap1
REC
RGD
Rho
RNAi
Runx2
siRNAs
Sox9
TGFTRANCE
Vmax
prostaglandins
protein kinase A
protein kinase B
protein kinase C
protein kinase G
protein kinase inhibitor
phospholipase C
peroxisome proliferator-activated receptor-gamma2
parathyroid hormone
PTH/PTHrp receptor
parathyroid hormone-related peptide
receptor activator for nuclear factor B
receptor activator for nuclear factor B ligand
repressor activator protein 1
response regulator receiver
arginine-glycine-aspartic acid
ras homolog gene
RNA interference
runt related transcription factor 2
short interfering RNAs
SRY-box containing gene 9
transforming growth factor
receptor activator of NF-{kappa} B ligand
maximal velocity constant
VDR
1,25-dihydroxyvitamin D3 receptor
VEGF
VPP
Wnt
vascular endothelial growth factor
valine-proline-proline
a combination of the genes (Wg) wingless and Int
10
1. Introduction
Bone has two main functions: it forms a rigid skeleton and plays a central role in calcium and
phosphate homeostasis. Bone modelling is a process associated with growth and re-shaping of
bones in childhood and adolescence. Throughout human life, bone tissue is subject to a
continuous turnover process whereby old bone is removed and replaced by new bone by the
coupled processes of bone resorption and bone formation. Bone is a very active organ and its
fixity after death is in sharp contrast to its ceaseless activity during life. In the adult skeleton,
the continuous activity largely comprises to the process of bone remodelling. Bone is
constantly destroyed or resorbed by the osteoclasts (bone-resorbing cells) and then replaced
by the osteoblasts (bone-forming cells). This process is regulated by other bone cells, namely
osteocytes, hormones, cytokines, prostaglandins and growth factors (Raisz 1999, Ducy et al.
2000, Teitelbaum 2000). A misbalance between bone resorption and formation may lead to
many bone diseases, e.g., osteoporosis and osteopetrosis. Osteoporosis, also known as bone
loss, typically reflects an imbalance in skeletal turnover such that bone resorption exceeds
bone formation. In osteopetrosis, the misbalance is caused by a defect in osteoclastogenesis or
by dysfunction of osteoclasts.
The cyclic nucleotides cAMP and cGMP are second messengers involved in a variety of
cellular responses to extracellular agents such as hormones and neurotransmitters. For
example, parathyroid hormone (PTH), an important hormone acting together with 1,25
dihydroxyvitamin D and calcitonin regulates bone metabolism, signalling via cAMP. cAMP
and cGMP are formed by adenylate and guanylate cyclases and are degraded by
phosphodiesterases (PDEs). PDEs determine the amplitudes of cyclic nucleotide-mediated
hormonal responses and modulate the duration of the signal. The activities of PDEs are
regulated by multiple inputs from other signalling systems and are crucial points of cross-talk
between the pathways (Soderling and Beavo 2000, Lugnier 2006). PDEs are an important part
of signal transduction in the cell, but very little is known about their role in bone cells.
During the last decade, fundamental studies have opened a new field of research dealing with
bioactive or biogenic substances derived from foods (Meisel 2001). The angiotensinconverting enzymes (ACEs) are involved in the regulation of the specific maturation or
degradation of a number of mammalian bioactive peptides. The antihypertensity effects of
bioactive peptides on blood pressure have been shown in several studies (e.g. Mitsui et al.
2004), and the bioactive peptides offered also provide a nutrigenomic aspect to bone cell
biology.
This work was conducted to clarify the involvement of PDEs and bioactive peptides in the
function and differentiation of human osteoblast cells and to provide more insight into the role
of PDEs in the regulation of bone metabolism. Elucidation of the role of PDEs in bone cells is
11
important because bone metabolism, especially bone formation, could be affected via PDEs.
The potential of bioactive peptides to increase osteogenic effects was also investigated.
12
2. Review of the literature
2.1 Bone biology
2.1.1 Bone tissue and bone cells
Bone tissue is highly organized and consists of a mineral phase of hydroxyapatite and
amorphous calcium phosphate crystals deposited in an organic matrix. It contains the distinct
cell types of osteoblasts, osteocytes, osteoclasts and lining cells. The osteoblast is the cell
responsible for collagenand other bone proteins, forming the bone organic matrix, which
consists of approximately 90% type I collagen and 10% different non-collagenous proteins,
such as osteocalcin, matrix Gla protein, bone sialoprotein, osteopontin and fibronectin (Aubin
and Liu 1996). Osteoblasts also have an important role in the subsequent mineralization of the
matrix. Once bone is mineralized, some of the osteoblasts are retained on the bone surface in
inactive form and named as the bone-lining cells (Parfit 1994). Osteocytes are osteoblasts that
have been trapped within the bone matrix during the process of bone formation. Osteocytes
communicate with one another and with cells at the bone surface via a meshwork of cell
processes that run through canaliculi in the bone matrix (Kogiannis and Noble, 2007). Thus,
bone cells form a functional network within which cells at all stages of bone formation, from
the preosteoblast to the mature osteocyte, remain connected. Osteoclasts are multinucleated
cells formed by the fusion of the monocyte/macrophage family. They are responsible for bone
resorption, playing a central role in formation of the skeleton and regulation of its mass. They
attach on the bone surface, secreting acids and lysosomal enzymes into the space provided
between their apical surfaces, and the mineralized bone surface (Ducy et al. 2000, Teitelbaum
2000).
2.1.2 Osteoblastic differentiation
Bone marrow stromal mesenchymal stem cells (MSCs) have strong proliferative potential and
the capacity to form bone, cartilage, adipocytes and fibrous tissue. MSCs have been found in
a variety of other adult tissues and can differentiate into myoblasts, haematocytes and
possibly even neural tissues (Reyes et al. 2001, Jiang et al. 2002a, 2002b, 2003). And, in
example, osteogenic differentiation potential of human bone marrow MSCs does not decrease
by age at late adulthood (Leskelä et al. 2003). Commitment of MSCs to tissue-specific cell
types is orchestrated by morphogens, developmental signalling pathways and transcriptional
regulators that serve as key factors (Figure 1A, Table 1).
The process of osteoblast differentiation can be subdivided in to three stages: 1) proliferation,
2) extracellular matrix synthesis and maturation and 3) mineralization. Each stage is
characterized by expression of distinguishing osteoblast markers. The most frequently used
markers of osteoblast differentiation are alkaline phosphatase (ALP), collagen type 1 (Col1),
osteopontin (OPN), bone sialoprotein (BSP), osteocalcin (OCN) and PTH/PTHrp receptor
13
A
B
Figure 1. (A) Commitment of mesenchymal stem cells to tissue-specific cell types. Some of the known signalling proteins, growth factors
and transcription factors playing key regulator roles in the mesenchymal lineages are indicated. Also shown is noticeable plasticity between
osteoblasts and adipocytes. (B) Osteoblastogenesis. Selected markers of the stages of maturation are used as examples. Modified from
Lian et al. (2003).
TABLE 1. Some key factors in osteoblastogenesis and bone formation.
Signaling protein and molecules
Indian hedgehog (Ihh)
-catenin
PGE2
Transcriptional regulators and factors
Osterix
SP-3 transcription factor
Runx1
Runx2/Cbfa1
Runx3
Growth factors
BMP2,4,7
IGF-1/IGF-2
Function
control of chondrocyte proliferation and maturation and osteoblast differentiation
regulates commitment to chondrocytes (low concentration) or osteoblasts (high concentration) in Wnt signalling
increseases BMP-2 expression
bone formation and osteoblast differentiation
contributes to ossification in vivo
promotes chondrogenesis
a master regulatory gene for activating osteoblastogenesis
supports chondrocyte maturation
Endothelin
FGF
chondro-osteoprogenitor cell specification
osteoblast differentiation, a potent anti-apoptotic factor for differentiated osteoblasts and osteocytes
migration and proliferation; inhibits differentiation
stimulates MSCs to differentiate
increase bone nodules in vitro
support the development of skeletal tissues
Hormones
PTH
1,25(OH)2D3
Glucocorticoids
Sex steroids (oestrogen; androgen)
Growth Hormone
Leptin
numerous anabolic effects via regulation of many types of osteoblast genes
a potent regulator, increasing and decreasing expression of numerous osteoblast phenotypic genes
promotes osteoprogenitor cell differentiation, but increases apoptosis of mature osteoblasts
stimulates osteoprogenitor proliferation and collage synthesis; increases differentiation and mineralization
stimulates proliferation and differentiation
anabolic effects on osteoblast/inhibitor for bone formation
Platelet-derived growth factor
Epidermal growth factor
(PTHR). In general, ALP, BSP and Col1 are early markers for osteoblast differentiation,
while PTHR and OCN appears late, parallel with mineralization. Osteoprogenitor cells will
arise under the appropriate stimulus from MSCs, differentiated preosteoblasts, osteoblasts and
osteocytes (Figure 1B).
The canonical Wnt/ -catenin pathway provides early development cues that indirectly
mediate the initial cascade of gene expression for skeletal development, bone formation and
osteoblast differentiation (Church et al. 2002, Logan et al. 2004). Bone morphogenetic
proteins [BMPs, members of the transforming growth factor
(TGF- ) family] play
important roles in directing fate decisions for MCSs. TGF- can provide competence for early
stages of chondroblastic and osteoblastic differentiation, but it inhibits myogenesis,
adipogenesis and late-stage osteoblast differentiation.
BMPs also inhibit adipogenesis and myogenesis, but they strongly promote osteoblast
differentiation (Boden et al. 1997, Hughes et al. 1995, Abe et al. 2000). The necessity of the
runt homology domain factor Runx2 (Cbfa1/AML3/PEBP 1) and the zing finger protein
osterix in bone formation has been ascertained by genetic studies (Banerjee et al. 1997, Ducy
et al. 1997, Choi et al. 2000, Nakashima et al. 2002). Because Runx2 expression arises earlier
than osterix, these two factors may represent a temporal sequence of regulation crucial for the
early and final stages of osteoblast differentiation (Nakashima et al. 2002, Komori 2005,
2006). The influence of transcription factors on lineage determination and differentiation is
revealed by their capability to cause transdifferentiation of cells to other phenotypes when
these factors are overexpressed or suppressed. For example, forced overexpression of
peroxisome proliferation-activated receptor 2 (PPAR 2) in osteoblasts will modify their
phenotype to adipocytes, whereas forced expression of Runx2 in adipocytes or other nonosseous cells results in acquisition of a differentiated osteoblast phenotype (Nuttall et al.
1998, Lecka-Czernik et al. 1999, Jeon et al. 2003)(Figure 1).
The developmental expression patterns of transcription factors during osteoblast maturation
reflect their roles as key determinants of osteoblast differentiation (Figure 1B). Many steroid
and polypeptide hormones, growth factors and cytokines regulate not only the growth of
osteoprogenitors and/or their progression to mature osteoblasts but also osteoblast activity and
apoptosis (for review, see Xing and Boyce 2005). For example, PTH stimulates growth of
osteoprogenitor populations, while also inhibiting osteoblast and osteocyte apoptosis (Boyce
et al. 2002). Leptin is secreted largely by adipose tissue, but also by human mesenchymal
stem cells (hMSCs) and osteoblasts at the mineralization stage. It enhances proliferation,
collagen synthesis, matrix deposition and mineralization (Gordeladze et al. 2002) and serves
as an anti-apoptotic agent in several cell systems, including bone marrow stem cells and
osteoblasts (Shimabukuro et al. 1998, Takeda et al. 1998, Konopleva et al. 1999).
Among important concepts in regulation of osteoblast differentiation and activity is the idea
that local or endogenous production of particular growth factors and cytokines undergoes
16
changes required for osteoprogenitor differentiation to proceed and for osteoblasts to modify
their activity. For example, insulin-like growth factor 1 (IGF-I) expression is markedly
increased during early osteoblast recruitment, but declines dramatically as these cells undergo
differentiation. Data from both IGF-I or IGF-I receptor-deficient mice and mice with targeted
overexpression of IGF-I in osteoblasts show that IGF-I is involved not only in embryonic
bone development but also postnatal bone formation and remodelling (Zhao et al. 2000,
Zhang et al. 2002). Fibroblast growth factor 8 (FGF-8) has been demonstrated to regulates
different stages of MSC differentiation toward osteogenic lineage in vitro and to increase the
osteogenic capacity of bone marrow cells at the early stage of their differentiation. However,
continued exposure of osteoblastic cultures to FGF-8 leads to the inhibition of bone formation
(Valta et al. 2006).
2.1.3 Osteoblastic regulation of osteoclast differentiation
During skeletal development and throughout life, the cells from the osteoblast lineage
synthesize and secrete molecules that in turn initiate and control osteoclast differentiation
(Raisz 1999). Osteoclast differentiation calls for the binding of macrophage colonystimulating factor (M-SCF) to its receptor as well as the binding of the soluble differentiation
factor receptor activator of NF-B ligand (RANKL) to its receptor (RANK) on osteoclast
precursor cells (Teitelbaum and Ross 2003). One of the first factors cloned that regulates
osteoclast differentiation was osteoprotegerin. Osteoprotegerin, originally characterized as a
novel secreted member of the tumor necrosis factor receptor superfamily, was later found to
inhibit spontaneous or induced bone resorption and cause osteopetrosis, the opposite of
osteoporosis. Osteoprotegerin acts as a decoy receptor that binds to RANKL and prevents it
from interacting with its receptor (Figure 2). RANKL and osteoprotegerin, both of which are
produced by osteoblasts at different stages of maturity (Gori et al. 2000), account for some of
the signals in osteoblast–osteoclast communication. In conjunction with M-SCF, the RANK–
RANKL–osteoprotegerin system regulates osteoclast differentiation. Thus, mice and humans
deficient in osteoprotegerin have a high rate of bone loss (bone resorption that exceeds bone
formation; Whyte et al. 2002). As in other high bone turnover states, anti-resorptive agents
can still reduce both bone formation and resorption and compensate for osteoprotegerin
deficiency. The signal(s) that couples resorption and formation remains elusive, although
several of the anabolic ligands, such as BMPs and TGF-ß, are stored in bone matrix as bone is
formed and are released during bone resorption and can thus act on osteoblasts and precursors
in the vicinity. BMPs could also enhance osteoclast differentiation from progenitors
(Hentunen et al. 1998, Itoh et al. 2001). Signals from osteoblasts to osteoclasts can be
provided by RANKL and osteoprotegerin. In this case, preosteoblasts express a high level of
RANKL relative to osteoprotegerin, which stimulates osteoclast differentiation and function.
More mature osteoblasts, by contrast, express high levels of osteoprotegerin relative to
RANKL, which inhibits osteoclast differentiation and function (Gori et al. 2000).
17
Figure 2. Interactions among osteoblasts and osteoclasts in bone resorption and formation and
some factors that modulate their differentiation and function.
2.1.4 Bone remodelling and its regulation
Remodelling of bone begins early in foetal life, and once the skeleton is fully formed in
young adults, almost all of the metabolic activity is in this form. The process involving bone
resorption and formation is initiated by mechanical signals and is controlled by local and
systemic factors that regulate osteoblast and osteoclast differentiation and function. Bone
remodelling is not a random process and takes place in bone remodelling units comprising
osteoblasts, osteoclasts and their precursors, in which resorption and formation are coupled.
Osteocytes are also involved in the process by soluble factors or directly through cell to cell
contacts (Zhao et al. 2002, Heino et al. 2002, 2004). Bone resorption is likely the initial event
that occurs in response to local mechanical stress signals. The bone remodelling cycle
involves a complex series of sequential steps that are highly regulated (Raisz 1999). The main
phases are 1) osteoclastic resorption, 2) reversal phase, 3) preosteoblastic migration and
differentiation into osteoblasts 4) osteoblastic matrix formation and 5) mineralization. The
complete bone remodelling cycle at each microscopic site takes about 4-6 months, with the
18
resorption process lasting about two weeks and bone formation several months (Hadjidakis
and Androulakis 2006).
The metabolic functions of the skeleton are served in large part by two calcium-regulating
hormones, PTH and 1,25-dihydroxy vitamin D. A third hormone, calcitonin, which can
inhibit bone resorption, is a potent inhibitor of bone resorption and is used clinically in the
treatment of osteoporosis. The US Food and Drug Administration has approved teriparatide,
the N-terminal (1-34) fragment of recombinant human PTH, for the treatment of osteoporosis
(Mishaela et al. 2002, Hodsman et al. 2005). In many countries in Europe, teriparatide is used
after an unsuccessful course of bisphosphonate therapy or after a patient has had a previous
osteoporotic fracture (Canalis 2007). PTH regulates serum ionized calcium concentration and
its primary target cells in bone tissue are the osteoblasts (Raisz 1999). The increase in bone
formation after PTH administration in vivo has been attributed to increased production of
osteoprogenitors and differentiation of osteoblasts and decreased apoptosis of existing
osteoblasts (Jilka et al. 1999). A number of studies in animals and humans have demonstrated
that short-term, intermittent administration of low doses of PTH has a significant anabolic
effect on bone (for review, see Jilka 2007). PTH affects several typical bone formationassociated features in the osteoblast such as collagen synthesis (Dempster et al. 1993), ALP
activity (Majeska et al. 1982) and synthesis of OPN (Noda and Rodan 1989). The receptor for
PTH belongs to the superfamily of the seven membranes spanning G protein-coupled
receptors (Birnbaumer 1990). The effects of PTH are known to be mediated by cAMP and
calcium and by the activation of protein kinase C (PKC) (Sprague et al. 1996, Ahlström and
Lamberg-Allardt 1997). Plasma PTH concentration tends to increase with age, and this may
produce an increase in bone turnover and a loss of bone mass, particularly of cortical bone.
Other systemic hormones are important in regulating skeletal growth. Growth hormone can
stimulate bone formation and resorption (Rosen et al. 1998), glucocorticoids are necessary for
bone cell differentiation during development (Advani et al. 1997) and thyroid hormones can
also stimulate bone resorption and formation and are critical for maintenance of normal bone
remodelling (Kawaguchi et al. 1994). Probably the most important systemic hormone in
maintaining normal bone turnover is oestrogen. Oestrogen defiency leads to an increase in
bone remodelling in which resorption outstrips formation and bone mass decreases.
Testosterone increases osteoblast formation and affects bone formation rather than bone
resorption (Khosla et al. 2002). Many local factors, cytokines, prostaglandins (PGs) and
growth factors also regulate bone remodelling.
2.1.5 Unbalanced bone remodelling: osteoporosis
The reduced bone mass, for instance, in osteoporosis results from an imbalance between bone
resorption and formation, with the rate of resorption exceeding that of formation. Excessive
bone resorption causes changes in the microstructure of the bone matrix, which make bones
19
prone to fracture. Osteoporosis has been proposed to result from a failure of resident bone
cells to respond appropriately to mechanical load–bearing signals and to control bone mass
and architecture (Lanyon and Skerry 2001). In the majority of conditions that lead to
osteoporosis, such as age-associated gonadal hormone deficiency, remodelling rates are high.
Although bone formation and resorption are both increased, the rate of bone formation is
insufficient to keep up with resorption. RANKL has been identified as a crucial cytokine for
the formation and activation of osteoclasts. The effects of RANKL are physiologically
compensated by the decoy receptor osteoprotegerin (OPG). The RANKL/OPG system is
activated in favour of RANKL in oestrogen deficiency, inflammation, bone malignancies and
during treatment with glucocorticoids, which enhances the ratio of RANKL to OPG, thus
promoting osteoclastogenesis, accelerating bone resorption and inducing bone loss (Simonet
at al. 1997, Hsu et al. 1999). The balance between proliferation, differentiation and apoptosis
of bone cells decides the size of osteoclast or osteoblast populations. Bone cells continuously
receive signals that control their proliferation, activity and survival from neighbouring cells,
the bone matrix and hormones. Life history before and after menopause and therapeutic
interventions with drugs, such as sex hormones, glucocorticoids, PTH and bisphosphonates,
are therefore partly determined by the survival of osteoclasts, osteoblasts and osteocytes,
which contribute to the bone mass and its microarchitecture (Weinstein et al. 1998, Jilka et al.
1999, Borton et al. 2001, Calvi et al. 2001, Gordeladze et al. 2002, Iwata et al. 2006).
2.1.6 Glucocorticoids and bone
Glucocorticoids (GCs) play an important role in modifying the metabolic activity and
proliferation of bone cells. GCs are required for in vitro bone nodule formation and
mineralization, as well as for bone marrow MSC proliferation and osteogenic differentiation
(Bellows et al. 1987, Shalhoub et al. 1992, Jaiswal et al. 1997, Bellows et al. 1998). Used in
long-term therapy as immunosuppressive and anti-inflammatory drugs, glucocorticoids lead
to a reduction in bone formation, enabling the development of osteoporosis (Eastell et al.
1998, Patschan et al. 2001). GSs have two major inhibitory effect on bone formation: inhibit
cell proliferation and decrease the population of cells capable of synthesizing type I collagen,
as well as downregulate collagen gene expression in osteoblasts (Mahonen et al. 1998). GCs
decrease osteoblast and osteocyte activities and survival and have the opposite effects on
osteoclasts, i.e. increased maturation and activity as well as decreased apoptosis (Weinstein et
al. 1998, O`Brien 2004, Canalis 2005). GCs exposure enhances RANKL expression and
inhibits OPG production by osteoblasts (Hofbauer et al. 1999), and also suppresses OPG
serum levels in vivo, thus elevating the RANKL/OPG ratio (Sasaki et al. 2001). In addition,
GCs enhance the expression and activity of PPAR 2, a transcription factor that suppresses
osteoblastogenesis and supports adipogenesis (Shi et al. 2000). Most studies have not found
an association between GCs and increased PTH levels (Canalis et al. 2004), but GCs might
enhance the sensitivity to PTH by changing the number of PTH receptors and their affinity to
PTH (Urena et al. 1994).
20
2.2 Cyclic nucleotide signalling and phosphodiesterases (PDEs)
2.2.1 Cyclic adenosine monophosphate (cAMP) signalling
The cAMP is a second messengers involved in a variety of cellular responses to such
extracellular agents such as hormones, growth factors and neurotransmitters. cAMP is a part
of the intracellular signalling pathway of several hormones involved in the regulation of bone
metabolism such as PTH, parathyroid hormone-related peptide (PTHrp) and prostaglandin E2
(Beavo 1995, Soderling and Beavo 2000). The level of intracellular cAMP is regulated by the
balance between the activity of two types of enzymes: adenylyl cyclase (AC) and the cyclic
nucleotide PDE. Both enzymes are encoded by a large number of genes that differ in their
expression patterns and mechanisms of regulation. The cAMP-dependent signal transduction
cascades from cell-surface hormone receptors to AC activation through the “coupling
proteins”that we know as heterotrimeric G proteins (Ross and Gilman 1977, Benovic et al.
1986, Lohse et al. 1990). Nine different ACs exist in mammals. Their activities are stimulated
by interaction with the subunit of Gs proteins. Gs subunits are coupled with different
types of membrane receptors in heterotrimeric complexes together with the ß and subunits,
from which they dissociate after the binding of specific ligands (Taussig et al. 1995).
Degradation of cAMP is regulated by a large family of PDEs (Beavo 1995). The activities of
ACs and PDEs are regulated positively and negatively by other signalling systems, such as
calcium signalling [through calmodulin (CaM), CaM-dependent protein kinase II and IV, and
calcineurin], subunits of other G proteins (e.g. Gi, Go and Gq proteins), inositol lipids (by
PKC) and receptor tyrosine kinases [through ERK and protein kinase B (PKB)] (Gilman
1990, Daub et al. 1996, Dessauer and Gilman 1997, Luttrell et al. 1999, Fisher et al. 2003).
Three major targets of cAMP have been identified: protein kinase A (PKA), the GTPexchange protein for the small GTPase or the exchange protein directly activated by cAMP
(EPAC) and the cyclic nucleotide-gated ion channels (Kopperud et al. 2003). cAMP could
also directly regulate the mitogen-activated protein kinase (MAPK) pathway by binding to
and activating EPAC and repressor activator protein 1 (Rap1) (Ehses et al. 2003). Although
the role of Rap1 in the MAPK pathway is incompletely understood, it seems to act by binding
to and activating mitogen-activated protein kinase kinase kinase B-Raf and/or inhibiting the
Ras-Raf (Ras as a G protein and Raf as a mitogen-activated protein kinase kinase) pathway
(Zhong 1995, Stork and Schmitt 2002).
PKA is composed of a complex of two regulatory (R) subunits and two catalytic (C) subunits.
Four genes (RI , RIß, RII and RIIß) encode the R subunits, and three (C , Cß, C and
PrKX) encode the C subunits (Skalhegg and Tasken 2000). PKA is activated by the binding of
cAMP to the R subunits, which induces their dissociation from the C subunit (for review, see
Taylor et al. 2005) Apart from the R subunits, PKA activity is down-regulated by protein
21
kinase inhibitors (PKIs). Three known isoforms of PKI , ß and ) are encoded by different
genes, each of which has a specific expression pattern. Interestingly, PKI could act as a
chaperone for nuclear export of the C subunit, thereby interfering with the extent of PKA
activity in the nucleus (Taylor et al. 2005, Dalton and Dewey 2006). An important step
forward in our understanding of the action of PKA has been the identification of PKAanchoring proteins (AKAPs). These proteins allow specificity in cAMP signal transduction by
placing PKA close to specific effectors and substrates (Huang et al. 1997).
A large number of proteins have been identified as substrates for PKA. PKA modulates the
cAMP response by phosphorylating other components of the cAMP pathway such as
receptors, ACs and PDEs. Depending on the cellular system involved, phosphorylation of
these proteins results in an enhancement of the cAMP signal (e.g. repressing the activity of
PDE1) (Giorgi et al. 2002, Goraya et al. 2004) or in a rapid down-regulation of the pathway to
obtain a transient effect (e.g. by repressing the activity of AC V and AC VI, or the 2
adrenergic receptor) (for review, see Beazly and Watts 2006). Regulation of transcription by
PKA is mainly achieved by direct phosphorylation of transcription factors. cAMP response
element-binding protein (CREB), cAMP response-element modulator protein (CREM) and
activating transcription factor 1 (ATF1), originally identified as the activators that respond to
cAMP, are phosphorylated by PKA in their activation domains. Several lines of evidence
support the notion that CREB, CREM and ATF1 can be phosphorylated by many different
kinases (Ziff 1990, Foulkes et al. 1991, Rehfuss et al. 1991, Lalli and Sassone-Corsi 1994,
Sassone-Corsi 1995).
2.2.2 Cyclic guanosine monophosphate (cGMP) signalling
Natriuretic peptides and nitric oxide (NO) activate the cGMP/cGMP-dependent protein kinase
G (PKG) signalling pathway. The role of cGMP as a second messenger has long been
realized, but the mechanism of action has only been addressed in the last couple of decades
(Francis and Corbin 1994, Forte and Currie 1995). Most of cGMP's downstream actions
(Eigenthaler et al. 1999), and possibly some of cAMP's actions (White et al. 2000), involve
activation of PKGs. Unlike cAMP, however, cGMP has been found to regulate other protein
classes such as ion channels (Mery et al. 1991, Delay et al. 1997, Hagen et al. 1998, Anthony
et al. 2000) and PDEs (Francis et al. 1994). The two forms of mammalian PKG have distinct
and overlapping tissue distributions. Type I isoforms exist as alpha and beta splice variants,
are cytosolic and have been recognized in the brain (Tsou et al. 1993), smooth muscle
(Komalavilas and Lincoln 1996), platelets, cardiac myocytes (Kwak et al. 1995) and kidney
(Gambaryan et al. 1996). The type II isoform is myristylated and membrane-bound and is
present in intestinal mucosa (French et al. 1995), kidney (Gambaryan et al. 1996), brain (elHusseini et al. 1995) and bone (Pfeifer et al. 1996). Both PKGs contain amino-terminal
leucine zipper motifs, and, in contrast to PKA, exist as homodimers in native tissues (Francis
and Corbin 1994).
22
Little is known about PKG regulation of gene expression, and gene expression profiling is
just beginning to contribute to the growing list of cGMP-regulated genes (Pilz and Broderik
2005). Expression of the immediate early genes c-fos and junB, but not of c-jun (a component
of the transcription factor activator protein 1) or proto-oncogene junD, is increased upon
activation of the cGMP pathway. Furthermore, this induction can be inhibited with the
selective PKG inhibitor KT5823 (Haby et al. 1994). cAMP response element (CRE)
dependent gene transcription is induced by transfected PKG, but weakly, as compared with
PKA (Collins and Uhler 1999). Both nitric oxide and atrial natriuretic peptide activate the cfos promoter through a guanylyl cyclase- and PKG-dependent mechanism (Gudi et al. 1996,
Idriss et al. 1999, Lincoln et al. 2006). Many other genes probably are targeted by
cGMP/PKG, including vascular endothelium growth factor (VEGF) and the VEGF receptor
(Tuder et al. 1995, Dulak et al. 2000), smooth muscle actin and calponin (Boerth et al. 1997,
Lincoln et al. 1998), but whether these genes are activated directly by PKG or indirectly
through cross-talk with other signalling systems is obscure. Re-expression of PKG in cultures
of mammalian cells that produce low levels of endogenous protein has been helpful in testing
the requirement for PKG in various signal transduction systems. Gudi et al. (1997) observed
that a specific cGMP-dependent nuclear localization signal controls PKG translocation to the
nucleus and transactivation of the fos promoter. These data suggest that the cGMP/PKG
system plays a potentially important and direct role in transcriptional regulation. However,
Collins and Uhler (1999) observed exclusively cytoplasmic localization of both PKG
isoforms and could only demonstrate nuclear translocation and transcriptional activation when
the protein was mutated to a chimeric form containing the C subunit of PKG and the PKA
catalytic domain. These findings indicate that PKGs, in contrast to PKA, may, in fact, utilize
additional cytoplasmic factors to regulate gene expression.
2.2.3. cAMP and cGMP PDEs
The second messengers, cAMP and cGMP, are inactivated by the cyclic nucleotide PDEs
(Figure 3). Mammalian PDEs have an HD domain (a superfamily of metal-dependent
phosphohydrolases; Aravind and Koonin 1998), the C-terminal half and show high affinity
for cAMP and/or cGMP. Protein domains involved in regulation of PDE enzymatic activity
and subcellular localization are mainly present in the N-terminal half. Some PDEs have
phosphorylation sites targeted by protein kinases and lipid modification sites. Approximately
270 aa in the C-terminal catalytic domain are conserved, with a sequence identity of 35%50% among different PDE families (Figure 4). Some PDE families are composed of 2- 4
subfamily genes showing sequence identity of more than 70% and having identical protein
domain organization. Twenty-one PDE genes have been identified in humans, rats, and mice.
They are categorized into 11 structurally, biochemically and pharmacologically distinct PDE
families, PDE1-PDE11 (Manganiello et al. 1995, Soderling et al. 1998a, 1998b, Fawcett et al.
23
Figure 3. Signalling using 3`5` cAMP and cGMP. A. Natriuretic peptide receptor (NPR)
activates guanylate cyclase (GC) upon receptor binding, followed by cGMP production from
precursor GTP. Soluble GC, activated by NO also catalyses the production of cGMP. Three
known cGMP target molecules, protein kinase G (PKG), cyclic nucleotide phosphodiesterase
(PDE) and cyclic nucleotide gated channels (CNG channel), may be activated, leading to
initiation of cellular responses. B. Receptors (GPCRs) bind specific receptor proteins.
Receptors then interact with G proteins found at the cytosol face of the membrane, causing
them to bind GTP, change conformation and subsequently activate the enzyme adenylyl
cyclase (AC). This enzyme produces cyclic AMP from adenosine triphosphate (ATP) within
the cell interior. Cyclic AMP then exerts its action in the cell by binding to and activating
forms of protein kinase A (PKA). This family of enzymes elicits the phosphorylation of
specific targets proteins, causing functional changes within the cell. Modified from Omari et
al. (2007).
24
Figure 4. General structure of phosphodiesterases. The different domains of the PDEs are
drawn as boxes connected by wires. PDEs share a common structure, with a conserved
catalytic domain (light blue) flanked by regulator domains (grey and white) the N-terminus
and C-terminus. The white box corresponds to regulator domain 1, where several
phosphorylation sites or allosteric binding sites have been mapped. Modified from Mehats et
al. (2002).
2000, Conti and Beavo 2007), some of which specifically hydrolyse cAMP (PDE3, PDE4,
PDE7 and PDE8) and others cGMP (PDE5, PDE6 and PDE9). The PDEs belonging to the
PDE1, PDE2, PDE10 and PDE11 families are dual-substrate enzymes, hydrolysing both
cAMP and cGMP (Soderling et al. 1998a, 1998b, Fawcett et al. 2000). Most PDE families
contain multiple genes (subtypes), and most of these genes can produce multiple transcripts or
variants (Conti and Jin 2000). The unique characteristics of each PDE gene family are defined
by protein domains located in the N-terminal to the catalytic unit. As shown in Figure 5,
approximately half of the PDE gene families (PDE2, PDE5, PDE6, PDE10 and PDE11) have
a protein domain termed GAF in tandem and are therefore designated a GAF-PDE
25
Figure 5. Domain and motif organization of each PDE family. Modified from Soderling and
Beavo (2000).
26
subfamily.The known functions of GAF domains are cGMP binding-mediated allosteric
regulation and dimerization of GAF-PDEs. Some GAF domains have also been reported to
bind cAMP. Other PDEs (PDE1, PDE3, PDE4 and PDE7–9) have no GAF domain and belong
to the non–GAF-PDE subfamily. PDE1 contains a Ca2+/CaM-binding site, PDE3 has a
transmembrane domain, PDE4 has upstream conserved regions (UCRs) and PDE8 has a
response regulator receiver (REC) domain and a per–arnt–sim (PAS) domain. PDE7 and
PDE9 have no specific protein domain in addition to the PDE catalytic domain.
Each PDE family is distinguished functionally by its unique combination of enzymatic
characteristics and pharmacological inhibitory profiles (Table 2). Different families, subtypes
and variants are varied in different tissues and cell types. One of the main roles of the PDEs is
to determine the amplitude of cyclic nucleotide-mediated hormonal responses and to
modulate the duration of the signal. Therefore, the PDEs can function as modulators of the
cyclic nucleotide-mediated signal transduction pathways (Manganiello et al. 1995, Mehats et
al. 2002, Housley et al. 2003, Conti et al. 2003). The activities of PDE are regulated by
multiple inputs from other signalling systems and PDEs are crucial in cross-talk between
pathways.
In the nomenclature used for each PDE isozyme (e.g. HsPDE1A1), the first two letters
indicate the animal species and the first Arabic number after PDE designates the PDE gene
family. This number is followed by a single capital letter indicating a distinct subfamily gene.
The last Arabic number indicates a specific splice variant or a specific transcript generated
from a unique transcription initiation site (http://depts.washington.edu/pde/pde.html).
2.2.3.1 PDE1 family
Three subfamily genes, PDE1A–C with various splice variants encode Ca 2+/CaM-dependent
cAMP- and cGMP-hydrolysing PDEs. PDE1 is mainly present in the cytosolic fraction;
however PDE1 has also been found in the fibres of several neurons from the dorsal root
ganglion (Giorgio et al. 2002). In humans, PDE1A shows high affinity for cGMP (Loughney
et al. 1996). PDE1B hydrolyses cGMP with a Km value lower and a Vmax value higher than
those for cAMP (Bender et al. 2005). High affinity for both cAMP and cGMP is observed
with PDE1C (Loughney et al. 1996). In cells expressing PDE1, hormones that raise cytosolic
Ca2+ would active PDE1, in this way diminishing the cyclic AMP response to hormones
stimulating cAMP or cGMP synthesis. Sustained Ca 2+ entry into the cells is required to
activate PDE1A in the astroma cell line, and PDE1A reportedly cannot discriminate between
the different sources of Ca2+ entry (Goraya et al. 2004). By contrast, in vivo phosphorylation
of PDE1 would likely result in the potention of cAMP or cGMP accumulation, involving the
elevation of cytosolic Ca2+ and activation of guanylyl or adenylyl cyclases, and would play a
major role in the amplification and prolongation of cyclic nucleotide effects. Different
alterations in tissue and cell PDE1 isozymes may occur by short-term (phosphorylation,
proteolysis) or long-term regulation (at the mRNA level) in response to hormonal stimulation
27
TABLE 2. Superfamily of phosphodiesterases.
Family
Substrate
Property
Number
of genes
Selective inhibitors
PDE1
cAMP,
cGMP
Ca2+-CaMstimulated
3
PDE2
cAMP,
cGMP
cGMP-stimulated
1
PDE3
cGMP-inhibited
2
PDE4
cAMP,
cGMP
cAMP
4
PDE5
cGMP
PDE6
cGMP
PDE7
cAMP
2
+
PDE8
cAMP
2
Unknown
-
PDE9
cGMP
1
Unknown
-
PDE10
cAMP,
cGMP
cAMP,
cGMP
cGMPinsensitive
PKA/PKGphosphorylated
Transducinactivated
Rolipraminsensitive
Rolipraminsensitive
cGMP high
affinity
Unknown
8-methoxy-IBMX,
phenothiazines,
nimodipine
erythro-9-[2hydroxy-3-nonyl]
adenine (EHNA)
Cilostamide,
milrinone
Rolipram,
roflumilast, Ariflo TM
Zaprinast, sildenafil
(Viagra TM)
Zaprinast, DMPPO,
E4021, Sildenafil
BRL 50481, ICI242
1
Papaverine
+
Unknown
1
Unknown
+
PDE11
1
4
IBMX
Sensitivity
+
+
+
+
+
+
and probably play a role in spatial-temporal regulation of PDE1 activity (Spence et al. 1997,
Michibata et al. 2001, Yan et al. 2001). In humans, PDE1A5 and PDE1A6 expression is
brain-specific, whereas PDE1A1 and PDE1A4 expression is omnipresent, but particularly
high in the kidney, liver, pancreas and thyroid gland (Michibata et al. 2001, Fidock et al.
2002). PDE1A10 expression is testis-specific (Michibata et al. 2001).
28
2.2.3.2 PDE2 family
PDE2A hydrolyses both cGMP and cAMP with similar maximal rates and relatively high K m
values. PDE2A is allosterically stimulated by cGMP binding to its GAF domain (Martinez et
al. 2002), enabling the simultaneous regulation of both cAMP and cGMP signalling. PDE2
was shown to play a major feedback role by restoring the basal level in cyclic nucleotides in
response to hormonal stimulation of the adrenal gland (MacFarland et al. 1991). Three
variants of a single gene were cloned for PDE2: PDE2A1, PDE2A2 and PDE2A3
(Sonnenburg et al. 1991, Yang et al. 1994, Rosman et al. 1997). PDE2 activity is up-regulated
in vivo at the post-transcriptional level by PKC under 4 beta-phorhol 12-myristate 13 acetate
(Geoffroy et al. 1999). In endothelial cells, PDE2 is up-regulated during phenotype changes
(Keravis et al. 2000) as well as under stimulation by VEGF (Favot et al. 2004), indicating
PDE2 participation in endothelial cell proliferation. PDE2 protein is mainly present in the
adrenal medulla, heart, rat ventricle (Yanaka et al. 2003), brown adipose tissue (Coudray et al.
1999), liver and brain.
2.2.3.3 PDE3 family
PDE3A and PDE3B, subfamily genes of PDE3, show a high affinity for both cAMP and
cGMP. A low Vmax value for cGMP compared with that for cAMP allows cGMP to function
as a competitive inhibitor for cAMP hydrolysis (Degerman et al. 1997). Therefore, PDE3s are
termed cGMP-inhibited cAMP PDEs. The presence of a 44-aa insert in the catalytic domain is
a unique characteristic of the PDE3 family. Another special feature is the presence of Nterminal hydrophobic membrane association domains (Wechsler et al. 2002). Short-term
activation of PDE3 was firstly demonstrated in rat fat cells in response to insulin and
isoprenaline stimulations, which induce PDE3 phosphorylation (Degerman et al. 1990). A
single phosphorylated serine site (Ser 302) was identified (Rahn et al. 1996). This site could
be phosphorylated either by PKA or by PKB, but the major site of PKB phosphorylation is
Ser 273 (Kitamura et al. 1999). Possible activation of PDE3B by phosphoinositide 3-kinase
phosphorylation (Rondinone et al. 2000) was recently implicated in the hypothalamic action
of leptin on feeding (Zhao et al. 2002) as well as in -cell insulin secretion (Zhao et al. 1998).
Furthermore, hormonal stimulation (insulin, glucagon) was shown to activate in vivo PDE3
associated with the Golgi-endosomal fraction (Geoffroy et al. 2001). PDE 3A is mainly
present in the heart muscle cells, platelets, vascular smooth muscle cells and oocytes
(Reinhardt et al. 1995, Degerman et al.1997, Wechsler et al. 2002), whereas PDE3B is mainly
associated with adipocytes, hepatocytes and spermatocytes (Miki et al. 1996).
29
2.2.3.4 PDE4 family
The PDE4 family exclusively hydrolyses cAMP (Km=2-4 µM). Currently, the PDE4 family
represents the largest and most studied PDE family, comprising four highly similar subfamily
genes, PDE4A-D, that encode rolipram-sensitive PDEs. The PDE4 family includes various
alternative splice variant groups encoding long PDE4 and short PDE4 isozymes, with a
minimum of 35 different PDE4 proteins (Houslay 2001) based on the presence or absence of
N-terminal upstream conserved region (UCR) domains (Bolger et al. 1993). The long-form
variants have UCR1, linker region (LR) 1, UCR2, LR2, and a catalytic domain. The shortform variants contain LR1-UCR2-LR2 and UCR2 (truncated)-LR2 in the N-terminal region.
UCR1, which includes one PKA phosphorylation site, is connected to UCR2 by LR1. UCR2
has a hydrophilic N-terminal region, which intramolecularly interacts with the hydrophobic Cterminal portion of UCR1 (Beard et al. 2000). UCR1 and UCR2 are involved in PDE4
enzymatic regulation (Sette and Conti 1996, Hoffman et al. 1999, Grange et al. 2000) through
UCR2 interaction with the catalytic domain (Lim et al. 1999) and have also been reported to
participate in PDE4 dimerization (Richter and Conti 2002). Although the PDE4 catalytic site
binds to the competitive inhibitor rolipram, the affinity of this site for rolipram can change
markedly depending on the conformation of the enzyme. PDE4 activity is controlled by
phosphorylation, a relationship with a protein or endogenous mediator and proteolysis. The
presence of an acceptor site in UCR1 for PKA-mediated phosphorylation allows a fast change
in PDE4 activity. In vivo, this augmentation of PDE4 activity was shown as a result of a
extended increase of cAMP resulting from hormonal stimulation; this was demonstrated to be
a short-term feedback mechanism allowing cAMP levels to return to the basal cellular state
(Sette et al. 1994, Oki et al. 2000). An extended accretion of cAMP induces a long-term
regulation of PDE4 induction. Follicle-stimulating hormone and dibutyryl cAMP treatment
was shown to start the specific induction of short-form PDE4D1 and PDE4D2 in rat Sertolli
cells, whereas the expression of the long-form PDE4D3 was unchanged (Swinnen et al.
1991). Conti et al. (1995) suggested that the PDE4D gene had two distinct transcriptional
units, the unit controlling the short-form expression being up-regulated by cAMP.
Correspondingly, cAMP-dependent up-regulation was shown for PDE4A and PDE4B in
U937 (Torphy et al. 1995), for PDE4B2 in human monocytes (Manning et al. 1996) and,
more recently, in human myometrial cells (Oger et al. 2002). Furthermore, the downregulation of PDE4A and PDE4B could be connected to PDE4D up-regulation, as shown in
endothelial cells during phenotypic variance (Keravis et al. 2000), whereas PDE4A was upregulated and PDE4B and PDE4D were down-regulated in the brain during fluoxetine
treatment (Miro et al. 2002). In humans, PDE4A-PDE4D expression is fundamentally
ubiquitous (Engels et al. 1994, 1995, Bolger et al. 1997, Wang et al. 1999, Wang et al. 2003),
with a variant-specific tissue distribution pattern, but PDE4A has been reported to be
relatively high in the brain (Bolger et al. 1993) and PDE4B low in the lung and absent in
blood (Engels et al. 1995).
30
2.2.3.5 PDE5 family
To date, only one gene product has been reported for this family, PDE5A, which has two
GAF domains (GAF A and GAF B) in the N-terminal half and specifically hydrolyses cGMP.
The GAF A domain of PDE5A has been reported to be responsible for this enzyme`s
allosteric binding to cGMP (Liu et al. 2002), and therefore PDE5A is termed a cGMP-binding
cGMP-specific PDE. One PKG- and PKA-dependent phosphorylation site in the N-terminal
region is related to activation of the PDE5A enzyme (Corbin et al. 2000). cGMP binding to
the PDE5A GAF A domain promotes phosphorylation, which not only activates the catalytic
function but also increases cGMP-binding affinity (Corbin et al. 2000, Francis et al. 2002,
Zoraghi et al. 2005). In humans, PDE5A transcripts are abundantly found in various tissues,
especially in smooth muscle tissues, and are also detected in platelets (Yanaga et al. 1998,
Kotera et al. 1999). PDE5A1 and PDE5A2 transcripts are widely distributed (Kotera et al.
1999, Lin et al. 2000). In contrast, specific expression of PDE5A3 in smooth and/or cardiac
muscle has been suggested (Lin et al. 2000).
2.2.3.6 PDE6 family
In retinal rod and cone cells, the level of cGMP, a second messenger in visual signal
transduction, is tightly controlled through regulated cGMP hydrolysis by PDE6A-C subfamily
genes (photoreceptor PDEs) (Cote 2004). Light-activated transducin stimulates PDE6 activity
by removing the inhibitory subunit . Membrane hyperpolarization is caused by cGMP
elimination, leading to an electrical cellular response. PDE6 and PDE6ß subunits encoded by
the PDE6A and PDE6B genes, respectively, form a holoenzyme, rod PDE, with two copies of
the smaller inhibitory subunit (PDE6 ) encoded by PDE6G. The GAF A domain is related
to PDE6 ß heterodimerization (Muradov et al. 2003). In cone cells, a homodimer of 2 '
subunits (PDE6 ') encoded by PDE6C comprises cone PDE with cone-specific inhibitory
subunits encoded by PDE6H. Thus, regulation of PDE6 activity by small inhibitory subunits is
a unique aspect of this PDE family. Several different genetic diseases caused by defects in
PDE6 function or expression have been characterized; in humans, for instance, autosomal
dominant congenital stationary night blindness and autosomal recessive retinitis pigmentosa
have been attributed to PDE6B mutations (Gal et al. 1994).
2.2.3.7 PDE7 family
This family includes two genes, PDE7A and PDE7B, that encode rolipram-insensitive highaffinity cAMP-specific PDEs (Km value approximately 0.2 µmol/L) (Michaeli et al. 1993).
The PDE7 family contains neither GAF domains nor regulatory domains (Beavo 1995).
PDE7A and PDE7B subtypes show approximately 70% homology (Gardner et al. 2000,
Sasaki et al. 2000). A PKA pseudosubstrate site is present in the N-terminus of the PDE7A
subfamily. Alternative splicing for PDE7A and tissue-specific expression of PDE7 splice
31
variants have been identified (Bloom and Beavo 1996, Han et al. 1997). PDE7A1 mRNA and
protein are distributed ubiquitously across human proinflammatory and immune cells (Smith
et al. 2003). On the other hand, PDE7A2, which is generated by 5'-splicing and differs from
PDE7A1 at its N-terminus (Bloom and Beavo 1996, Han et al. 1997), has never been detected
at the protein level, despite unequivocal identification of its mRNA (Smith et al. 2003). The
distribution of PDE7A3 is largely unknown, but in human it has been found in T-lymphocytes
(Glavas et al. 2001) and may also be present in many PDE7A1-expressing cells, as both
transcripts are probably regulated by the same promoter (Torras-Llort and Azorin, 2003). In
rats, three N-terminal splice variants, PDE7B1-3, exist. Only PDE7B1 has been identified in
humans and mice and it is abundant in the brain, liver, heart, thyroid glands and skeletal
muscles (Gardner et al. 2000).
2.2.3.8 PDE8 family
PDE8A and PDE8B are the subfamily genes of PDE8. PDE8s are high-affinity cAMPspecific PDEs insensitive to rolipram and 3-isobutyl-1-methylxanthine (IBMX) (Gamanuma
et al. 2003), and contain response regulator receiver (REC) and PAS domains in the Nterminal portion. The REC domain functions as a receiver of signals from the sensor
component in a 2-component signal transduction system in lower organisms. The PAS domain
is involved in the binding of small ligands and protein–protein interactions (Huang et al.
1993). However, regulation of PDE8s via REC or PAS domain is unknown, and obvious
PDE8 activity has not yet been demonstrated in either tissue or cell extracts. PDE8A
transcripts are expressed in various tissues and are abundant in the testis, ovary, small
intestine and colon (Fisher et al. 1998). The expression of PDE8B seems to be confined to the
thyroid gland and the brain (Kobayashi et al. 2003).
2.2.3.9 PDE9 family
PDE9A is the only reported subfamily gene of PDE9, although more than 20 splice variants
have been observed in humans, indicating that the PDE9A gene appears to have a complex
regulation of expression (Rentero et al. 2003). PDE9A specifically hydrolyses cGMP with
high affinity (Fisher et al. 1998, Guipponi et al. 1998). However, no reports are available on
the regulation of PDE9A activity or on the presence of endogenous PDE9A activity in either
tissue or cell extracts. IBMX-insensitive cGMP PDE activity, which has been shown to date,
is likely attributable to PDE9. PDE9 mRNA is highly conserved between species and is
widely distributed throughout the rodent brain (Van Staveren et al. 2002). PDE9A mRNA is
expressed in the spleen, small intestine, brain, colon, prostate, kidney and placenta (Fisher et
al. 1998, Guipponi et al. 1998). In humans, the PDE9A5 splice variant is highly expressed in
immune tissues (Wang et al. 2003).
32
2.2.3.10 PDE10 family
PDE10A contains two N-terminal GAF domains and hydrolyses both cAMP and cGMP. Two
major variants, PDE10A1 and PDE10A2, and several minor variants have been identified in
humans (Fujishige et al. 1999a, 2000). High affinity for cAMP inhibits cGMP hydrolysis,
making this enzyme a cAMP-inhibited dual-substrate PDE. Among newly discovered PDEs,
the enzymatic activity of PDE10A is clearly demonstrated in tissue extracts (Fujishige et al.
1999b). The enzymatic activity of a chimeric construct of the PDE10A GAF domain and
cyanobacterial adenyl cyclase is stimulated by cAMP, suggesting a possible allosteric
modulation of PDE10A activity by cAMP (Gross-Langenhoff et al. 2006). PDE10 transcripts
are particularly abundant in the brain, thyroid and testis. The PDE10 family was recently
shown to be associated with a progressive neurodegenerative disease, Huntington`s disease
(Hebb et al. 2004, Hu et al. 2004).
2.2.3.11 PDE11 family
The most recently identified family, PDE11, contains four N-terminal variants, PDE11A1PDE11A4. A full-length form, PDE11A4, contains two GAF domains and a catalytic domain.
PDE11A hydrolyses both cAMP and cGMP with similar Km values (Fawcett et al. 2000,
Hetman et al. 2000, Yuasa et al. 2000). Although a cyanobacterial adenyl cyclase fused with
PDE11A4 GAF domains is activated by cGMP (Gross-Langenhoff et al. 2006), no reports
exist on the allosteric regulation of the PDE11A enzyme. PDE11A activity has not yet been
clearly demonstrated in tissue or cell extracts. In humans, PDE11A expression is strong in the
prostate and moderate in the testis and several other tissues (Fawcett et al. 2000, Yuasa et al.
2000).
2.2.4 Regulation of activity and compartmentalization of PDEs
PDEs exist universally, from prokaryotes to eukaryotes. In all organisms, multiple PDEs
appear to be involved in tight feedback regulation of cAMP and cGMP. Observations also
imply that different combinations of PDEs can regulate the same general function in different
species. Divergent PDE expression patterns have been detected in the hearts of the rodents
and humans. Two general types of PDE regulation occur. Short-term regulation involves the
activation of second messenger pathways and consequent allosteric modulation of enzyme
activity, and long-term regulation occurs through increased enzyme synthesis. Several PDE
families are subject to one or both forms of regulation. Functional evidence indicates that
cyclic nucleotide signals are confined in compartments and that different PDEs, as well as the
presence of PDE/PKA or PDE/PKG complexes, may contribute to establishment or
stabilization of these compartments. However, the exact mechanism restricting cyclic
33
nucleotide diffusion remains unclear, and convincing evidence for PDE-generated cyclic
nucleotide gradients is still lacking (for review, see Conti and Beavo 2007).
2.2.5 Inhibitors of PDEs
The PDEs are targets for a number of synthetic PDE isozyme inhibitors. Theophylline was the
first chemical inhibitor described for PDEs (Butcher and Sutherland 1962). One decade later,
a new xanthine analogue, IBMX, was shown to be 100-fold more potent than theophylline
(for review, see Chasin and Harris 1976). These two compounds are both non-specific PDE
inhibitors. The last decade, numerous more or less selective inhibitors have been designed and
synthesized for scientific and pharmacological purposes. Some of them inhibit PDE activity
very potently in the nanomolar and subnanomolar ranges and are specific to certain PDE
families. Thus, in the span of several decades, the potency of PDE inhibitors has increased by
108, and the specificity of some of them by 104. Recently, several selective inhibitors for
individual genes within the PDE family have been discovered and synthesized, e.g. OPC33540 inhibits PDE3A (IC50= 0.32 nM) (Sudo et al. 2000) and IC242 inhibits PDE7A
selectively, with an IC50 of 0.37 µM (Lee et al. 2002a, 2002b). Efforts are now focused on
developing third-generation inhibitors that are subtype-specific. PDE inhibitors are
experimental tools singularly useful in studies of the role of PDEs in cyclic nucleotide
signalling and also represent a novel class of pharmacological agents suited for “signal
transduction pharmacotherapy” (Levitzki 1997). Currently, PDE1, PDE2, PDE3, PDE4,
PDE5, PDE6, PDE7 and PDE10 have more or less selective inhibitors, but no selective
chemical inhibitors have yet been described for PDE8, PDE9 or PDE11 (Table 2). Because of
their great market potential and therapeutic relevance, PDE inhibitors became recognized as
important therapeutical agents for a variety of diseases, such as heart failure, depression,
asthma, inflammation and erectile dysfunction (Conti et al. 1995, Torphy 1998, Mehats et al.
2002, Rotella 2002, Halena and Siegel 2007). Several PDE inhibitors are currently on the
market, e.g. sildenfil (Viagra®), tadalafil (Cialis®) and vardenafil (Levitra®); all of these are
inhibitors of PDE5.
2.2.6 Bone metabolism and PDE inhibitors
Several studies have shown that selective chemical PDE inhibitors promote osteoblastic
differentiation and increase bone formation both in vivo and in vitro (Miyamoto et al. 1997,
2003, Kinoshita et al. 2000, Wakabayashi et al. 2002, Scutt et al. 2004). Most of these effects
are likely to be mediated by elevated cAMP levels, although cAMP-independent effects have
also been reported (Rawadi et al. 2001). In osteoblasts, for example, cAMP produced in
response to PTH or PGs regulates osteoblastic differentiation (Kumegawa et al. 1984,
Farndale et al. 1988, Partridge et al. 1994, Ishizuya et al. 1997). There are corresponding data
that show that administration of PTH or PGs also leads to increases in cancellous bone
34
volume in animal models (Jee et al. 1985, 1987, High 1987, Finkelstein et al.1994, Whitfield
and Morley 1995, Reeve 1996). In rat bone marrow culture, XT-44 (a PDE4 inhibitor) and
rolipram stimulated mineral-nodule formation, whereas it inhibited osteoclast-like cell
formation in mouse bone marrow culture (Waki et al. 1999). Therefore, PDE inhibitors are
considered to be promising candidates for anti-osteoporosis drug therapies. A PDE4 inhibitor,
XT-611, has been shown to inhibit osteoclast multinucleation or maturation by acting
synergistically on osteoclast progenitors with prostaglandin E2 (PGE 2) secreted from stromal
cells, but not by influencing the cell-to-cell interaction between stromal cells and osteoclast
progenitors (Yamagami et al. 2003). Rolipram has augmented the inhibitory effects of PGE2
on osteoclast progenitor cell viability; thus combined treatment with PGE 2 and rolipram
suppresses osteoclast formation by directly reducing osteoclast progenitor cell viability (Park
and Yim 2007). By contrast, PDE inhibitors have been shown to stimulate osteoclast
formation by inducing receptor activator of NF-{kappa} B ligand (TRANCE) expression in
mice calvarial osteoblasts and UAMS-32 cells (Takami et al. 2005). Pentoxifylline and
rolipram have reportedly been to increase bone mass in animals given these drugs in daily
injections (Kinoshita et al. 2000). PTH as well as PDE inhibitors induced osteoclast formation
in a mouse co-culture system. Injections of PTH to animals once per day produced a net
anabolic effect in vivo (Tam et al. 1982, Fu et al. 2002). Continuous exposure to PTH leads to
an increase in both bone formation and bone resorption, with a net loss of bone mass (Tam et
al. 1982). PDE inhibitors may induce bone loss if administered continuously (Takami et al.
2005).
2.3 Other signal transduction pathways associated with PDEs
PDE isoenzymes are targets of intricate regulatory mechanisms that integrate different
signalling pathways, function in feedback to modulate signalling pathways and serve as
effectors of signalling pathways. An additional property that only recently is being explored is
the subcellular distribution of PDEs and their involvement in establishing microdomains and
channelling of signals. Future pathway studies are anticipated to shed light on the role of
PDEs in cyclic nucleotide signal transduction and connections between other signal pathways.
2.3.1 Exchange protein directly activated by cAMP (EPAC) and PDEs
In the heart, signalling events such as the onset of cardiac hypertrophy are influenced by
muscle-specific mAKAP signalling complexes that target PKA, the EPAC and PDE4 (DodgeKafka et al. 2005). Mediation of signalling events by AKAPs might also have a role in
lipolysis in adipocytes. cAMP either stimulates or inhibits PKB phosphorylation, acting
through different effectors, in rat adipocytes. Hence, a cAMP pool that activates EPAC and is
regulated by PDE3B and PDE4 appears to mediate a decrease in PKB phosphorylation. PKB
35
phosphorylation, and therefore activation, in adipocytes is regulated by an orchestrated crosstalk between insulin and cAMP pathways, where PDE3B seems to play an important role
(Zmuda-Trzebiatowska et al. 2007).
2.3.2 Mitogen-activated protein kinase (MAPK) and PDEs
PDEs have been shown to be involved in MAPK signalling; e.g. pentoxifylline, a nonselective PDE inhibitor, induces osteoblastic differentiation in pluripotent mesenchymal cell
lines C3H10T1/2 and C2C12. Pentoxifylline treatment enhances osteoblast differentiation,
which occurs via signalling by MAPK cascades and is independent of PKA activation
(Rawadi et al. 2001). Rat mesangial cells (MCs) possess functionally compartmentalized
intracellular pools of cAMP that are differentially regulated by cAMP PDEs. Inhibition of
PDE3, but not PDE4, has been demonstrated to suppress MC mitogenesis (Tsuboi et al.
1996). MC mitogenesis is regulated through "negative cross-talk" between cAMP PKA and
extracellular signal-regulated kinase (ERK) signalling. PDE3 inhibitors act by suppressing
ERK activation. Other targets of cAMP signalling directed by PDE3 include cyclins D, E and
A, and the cell cycle inhibitor p21 (Cheng et al. 2004).
2.3.3 PKC and PDEs
PDEs have been reported to be involved in the non-canonical Wnt signalling pathway. Wnt
binds to a Frizzled receptor and possibly also the co-receptor, Knypek or Ror2. This can lead
to activation of a pathway involving calcium/CaM-dependent kinase II and protein kinase C.
Frizzled may activate heterotrimeric GTP-binding proteins, leading to activation of
phospholipase C (PLC) and PDE. Frizzled can also recruit Dishevelled, leading to activation
of small GTP-binding proteins, such as Rho and Cdc42 (Veeman et al. 2003). PKC, cAMP,
PLC, Ca2+ and PDEs have been shown to be involved oocyte maturation in amphibians
(Toranzo et al. 2007). The roles of PKC and PDE have been demonstrated in -opioid
agonist-induced reductions in cAMP levels in the isolated iris-ciliary bodies of the rabbit
(Dortch-Carnes and Potter 2002). In addition, -opioid receptor stimulation in rat heart has
shown PDE4 activation by PKC (Bian et al. 2000).
2.4 Bioactive peptides
During the last decade, fundamental studies have opened a new field of research concerning
bioactive or biogenic substances derived from foods (Meisel 2001). Proteins and peptides
from food have been found to be physiologically active or bioactive either in a direct manner
through their presence in the undisturbed food itself or after their release from the respective
host proteins by hydrolysis in vivo or in vitro, e.g. in cheese ripening (Rizzello et al. 2005a),
36
food fermentation (Fitzgerald and Murray 2006, Korhonen and Pihlanto 2006) or enzymatic
reactions in the gut after the ingestion of foods containing precursor proteins (Meisel et al.
2003, Hernandez-Ledesma et al. 2004). Bioactive peptides have been found, among others, in
meat, fish, egg, soy and grain products, but thus far most have been isolated from milk-based
products. Functionally bioactive peptides are usually unknown sequences 2-5 aa in length and
have a proline residue in the C-terminus (Curtis et al. 2002). Two human di-/tri-peptide
transporters have been identified; hPepT1 is expressed both in the small intestine and in the
proximal tubule, whereas hPepT2 is expressed only in the proximal tubule (see Nielsen and
Brodin 2003, Steffansen et al. 2004). The cellular mechanisms and signalling of bioactive
peptides remain obscure.
Different health effects have been attributed to food-derived peptides. Antimicrobial peptides
have been recognized in many protein hydrolysates, particularly in milk. The most studied are
the lactoferricins, which are derived from bovine and human lactoferrin (Kitts and Weiler
2003). In addition, antibacterial peptides have also been recognized in s1-casein and s2casein (Rizzello et al. 2005b, McCann et al. 2006). Antimicrobial peptides operate against
different Gram-positive and Gram-negative bacteria, yeast and filamentous fungi. The ACEs
are involved in the regulation of the specific maturation or degradation of a number of
mammalian bioactive peptides. Since the first detection of exogenous ACE inhibitors in snake
venom (Ondetti et al. 1977), a large number of ACE peptides have been discovered in the
digestion of different food proteins, being established particularly in milk but also in fish and
meat. ACE inhibitory peptides are generally short-chain peptides, often carrying polar amino
acid residues like proline (Fuglsang et al. 2003) The hypertensive and immunomodulator
peptides Val-Pro-Pro and Ile-Pro-Pro, for example, can be released from precursor proteins
-casein and -casein) by enzymes from Lactobacillus helveticus (Hata et al. 1996,
Korhonen and Pihlanto 2003). The antihypertensity activities of bioactive peptides on blood
pressure have been shown in several studies (e.g. Mitsui et al. 2004, Miguel and Aleixandre
2006). Hypocholesterolemic effects have been reported for casein and whey-derived peptides,
in addition to peptides from soy protein (Hori et al. 2001). Antithrombotic and antioxidant
activities of bioactive peptides (Kitts and Weiler 2003) have been shown, as has enhancement
of mineral absorption and/or bioavailability in rats (Erba et al. 2002) and in the human
intestine (Chabance et al. 1998, Meisel et al. 2003). Immunomodulatory peptides can improve
immune cell functions, measured as lymphocyte proliferation, natural killer cell activity,
antibody synthesis and cytokine regulation (Horiguchi et al. 2005, Fitzgerald and Murray
2006). Furthermore, immunomodulatory peptides might reduce allergic reactions in atopic
humans and augment mucosal immunity in the gastrointestinal tract (Korhonen and Pihlanto
2003). Cytochemical studies have given increasing evidence that bioactive peptides modulate
the proliferation, differentiation and apoptosis of various cell types. Some milk-derived
peptides, for instance, have been shown to trigger apoptosis, mainly in malignant cells,
whereas normal cells seem to be less susceptible (Meisel and Fitzgerald 2003). Peptides have
also been observed to have opioid activities; for example, opioid receptor ligands with
agonistic activity originate from different milk proteins and exert naloxone-inhibitable opioid
37
activities in both receptor studies and bioassays (Kayser and Meisel 1996, Mullally et al.
1996).
2.4.1 Bioactive peptides in bone
Effects of bioactive peptides have been studied in dental/orthopaedic biomaterials as well as
in basic bone biology. Proactive or new-generation biomaterials achieve specific, timely and
desirable responses from surrounding cells and tissues. Osteoblast adhesion is a crucial
process at bone implant interfaces before multiple osteoblast functions (such as proliferation
and mineralization), which occur after cell adhesion and are necessary for the success of an
implant (Dee 1996a). In the past two decades, many cell adhesion molecules have been
discovered, and their functions in cell morphology, locomotion, mitosis, cytokinesis,
phagocytosis and maintenance of cell polarity have been investigated (Pavalko and Otey
1994, Haass et al. 2005, Turowski et al. 2005). A key finding remains that the amino acid
sequence arginine-glycine-aspartic acid (RGD) in fibronectin serves as a primary cell
attachment cue. The RGD sequence is expressed in many extracellular matrix (ECM)
molecules, and the responsiveness of the RGD sequence for all attachments has been shown
numerous times. The small synthetic peptides (often hundreds of daltons) that contain the
amino acid sequence RGD can apparently mediate cell attachment similarly to their
considerably larger parental molecules (100 000 Daltons). According to Craig et al. (1995),
the potential exists to develop RGD-based therapeutics that function either as agonist, to
promote the interaction of cells and tissues with artificial matrices or as antagonists to control
the nature of cell-cell and cell-ECM interactions. The coating of biomaterials with RGD
peptides offers an attractive strategy for controlling the cell-material interface and achieving a
bioactive implant (Kantlehner et al. 1999, Verrier et al. 2002). Peptides have been shown to
support and enhance osteoblast adhesion (Dee et al. 1998, Schuler et al. 2006), proliferation
(Huag et al. 2003, Benoit and Anseth, 2005) and mineralization (Dee et al. 1996b). Based on
the RGD structure, synthetic RGD-mimetic compounds have been designed. These mimetic
compounds are effective inhibitors of bone resorption in vitro and in vivo (James et al. 1996,
Engleman et al. 1997).
In addition, the effect of other small peptides (without RGD sequence), such as isoleucineproline-proline (IPP) and valine-proline-proline (VPP), have been studied in bone cells. IPP
and VPP, fermented from milk by Lactobacillus helveticus, increase activation of bone
formation in osteoblast culture, but do not have an impact on osteoclast activation and bone
resorption in vitro (Narva et al. 2004). L. helveticus-fermented milk has been demonstrated to
prevent bone loss in ovariectomized rats, but this phenomenon could not be seen following
synthetic VPP treatment (Narva et al. 2007).
38
3. Aims of the study
This study focused on the areas of bone cell biology and signalling transduction. PDEs are an
important part of signal transduction in the cell, but very little is known about them in bone
cells. The inhibition of some PDEs could have therapeutic potential in the treatment of
osteoporosis. Short bioactive peptides have been shown to have modulatory effects on bone
biology.
The aim of this study was to identify PDE families in human osteoblasts and osteosarcoma
cells as well as to investigate the role of PDEs and bioactive peptides on the activation and the
differentiation of human osteoblast cells.
Specific aims were as follows:
1) To identify the expression of PDE families in human osteoblasts.
2) To investigate the effects of PDE inhibition on osteoblast differentiation and function
with RNA interference (RNAi) and pharmacological inhibitors.
3) To analyse the expression and regulation of different PDEs during osteoblast
differentiation at the mRNA and enzyme levels.
4) To examine the effects of bioactive peptides on gene expression in the osteoblast.
39
4. Materials and methods
4.1 Cell culture
4.1.1 Osteosarcoma cell lines (I, III, IV)
Human and rat osteosarcoma cell lines, widely used as models for osteoblasts, were used in
studies I, III and IV. The human osteosarcoma MG-63 (III), which is considered to represent a
mature osteoblastic phenotype was the primarily used cell line, whereas SaOS-2 cells (I, III)
were mainly used to confirm results obtained with MG-63 cells (III) or NHOst (I). Both cell
lines were purchased from the American Type Culture Collection. MG-63 cells were cultured
in α-MEM supplemented with 10% foetal calf serum and SaOS-2 cells in McCoy`s 5A,
supplemented with 12.5% foetal calf serum, with antibiotics (50 IU/ml penicillin and 50
µg/ml streptomycin) in a humidified atmosphere with 5% CO2 at 37°C. UMR-106 (IV) rat
osteosarcoma cells (American Type Culture Collection) were grown in Dulbecco’s modified
Eagle’s medium (DMEM), supplemented with 10% foetal calf serum (FCS), 50 IU penicillin
and 50 g/ml streptomycin at 37°C in a humified atmosphere with 5% CO2 on 96–well plates.
The cells were seeded at 1 x 104 cells/cm².
4.1.2 Human osteoblasts (I)
Normal human osteoblasts (NHOst) and all the cell culture reagents for the NHOst cells were
purchased from Cambrex, BioWhittaker. Two strains of NHOst cells isolated from bone chips
from female donors were used. The cells were seeded at a density of 5000 cells/cm2 and
cultured in osteoblast growth medium supplemented with 10% FCS, 50 µg/ml ascorbic acid, 5
mM ß-glycerophosphate, 200 nM hydrocortisone-21-hemisuccinate and 0.1%
gentamin/amphotericin-B solution, at 37ºC in a 5% CO2/95% air atmosphere on plastic Petri
dishes.
4.1.3 Human mesenchymal stem cells (hMSC)-derived osteoblasts (II, IV)
Human bone marrow-derived mesenchymal stem cells (hMSC, Cambrex) were cultured in
mesenchymal stem cell growth medium with mesenchymal cell growth supplements
(Cambrex) at 37°C in a humified atmosphere with 5% CO2 on 100-mm plastic dishes at a
seeding density of 3100 cells/cm2. The differentiation into osteoblasts was performed
according to the instructions of the supplier after the cultures became 50% confluent. The
differentiation medium consisted of -MEM supplemented with 10% foetal bovine serum,
10-8 M dexamethasone (Dex), 50 g/ml L-ascorbic acid, 10 mM –glycerophosphate, 100
U/ml penicillin and 100 U/ml streptomycin and was changed every 3-4 days.
40
4.2 PDE inhibitors
The PDE chemical inhibitors used in the studies are listed in Table 3.
TABLE 3. PDE inhibitors used in Studies I-III.
Family
Inhibitor
Study
PDE1
8-MMX (8-methoxymethyl-3-isobutyl-1-methylxanthine)
I, III
PDE2
EHNA
I, III
PDE3
Cilostamide (N-cyclohexyl-N-methyl-4-[1,2-dihydro-2oxo-6-quinolyloxy] butyramide)
III
PDE3
Milrinone (6-methyl-2-oxo-5-pyridin-4-yl-1H-pyridine-3carbonitrile)
I
PDE4
Rolipram (4-[3-{cyclopentyloxy}-4-methoxyphenyl]-2pyrrolidinone)
I, II, III
PDE7
BRL 50481 (3-(N,N-dimethylsulfonamido)-4-methylnitrobenzene)
II
PDE3
cGMP
I, III
non-specific PDE
inhibition (PDE1-7,
10-11)
IBMX
I, III
4.3 Primers (I, II, III, IV)
The primers were used as presented in Table 4.
TABLE 4. Primers used in Studies I-IV. Forward primer (F), Reverse Primer (R).
cDNA
Sequence
Study
PDE1A
F: AAAATGGGGATGACAAAAAAGAA
R: GATATTCATTTCTTCTTCTTGCAT
F: ATGCAGGATGATGAGATGAA
R: CGGAAGAATTCCTCCAT
F: AGACAGGGTGACAGAGAAGC
R: TGGCCTTCTCCTCTTTGGGT
F: TCACCTCTCCAAGGGACTCCT
R: CAGCATGTAAAACATCAGTGGC
I, III
PDE1B
PDE1C
PDE3A
41
I, III
I, III
I, III
PDE4A
PDE4B
PDE4B
PDE4C
PDE4C
PDE4D
PDE7A
PDE7B
PDE8A
PDE8A
PDE10A
PDE11A
GAPDH
Beta-actin
Beta-actin
Osteocalcin
Caspase-8
CREB-5
Beta-catenin
Vitamin D
receptor
BMP-1
BMP-2
BMP-5
PTHrp
T7-promotor
+ PDE7
F: CTCGCACAAGTTCAAAAGGATG
R: GCCTCCAGCGTAATCCGACA
F: GACATTGCAACAGAAGACAAG TCCCC
R: ACTCAAGTAACTGAAGGCCAGGTGG
F: AGCTCATGACCCAGATAAGTG
R: ATAACCATCTTCCTGAGTGTC
F: ACCTCAGCGCCAAGCAGC
R: TGGGTGGGAAAGTGAAGCAGG
F: ACACTGAACTCCTGTCCCCTGAAG
R: GATGTGACTCAAGAGTGACCACTGG
F: CCCTGGACTGTTATCATGCA
R: TGATTGGACACACCAGGATG
F: GGACGTGGGAATTAAGCAAGC
R: TCCTCACTGCTCGACTGTTCT
F: TGCACAGGACAGGCACTTTA
R: CTTCTGTGCTGCCTGGGCAA
F: GCC TGT TTC CTG GACAAA C
R: GCA TTA CGG ACA ACT CTT CTC
F: CAGCCAGAGACGACACTCTTCCAT
R: ATGATCTTAGCGTTGACTCGGAGC
F: GTGTGGTGCAGATGGTCAAC
R: GGGTGAATTGCATGAGACCT
F: ATCTGCCTCAGTATCCCCCT
R: CCACCAGGAAAAGAGAGCAG
F: AAGGCTGGGGCTCATTTG
R: CCACCAGGAAAAGAGAGCAG
F: CATCGTCACCAACTGGGACGAC
R: CGTGGCCATCTCTTGCTCGAAG
F: AGGCCAACCGCGAGAAGATGACC
R: GAAGTCCAGGGCGACGTAGCAC
F: ATGAGAGCCCTCACACTCCTCG
R: GTCAGCCAACTGGTCACAGTCC
F: AGGAGGAGATGGAAAGGGAACTT
R: ACCTCAATTCTGATCTGCTCACTTCT
F: GCTTTGGTGCTTTTCTCCAG
R: GGTGACACCACAGCACAAAC
F: TTCTGGTGCCACTACCACAGC
R: TGCATGCCCTCATCTAATGTC
F: CCAGTTCGTGTGAATGATGG
R: CCTTTTGGATGCTGTAACTG
F: AGGCCCACTTCTTCTCAGAAAAG
R: GAAGAGAGCACCAGTTCTGTCCG
F: GGAGAAGGAGGAGGCAAG
R: GACACGTCCATTGAAAGAGC
F: AAGAAGACAAGAAGGACTAAAAATAT
R: GTAGAGATCCAGCATAAAGAGAGGT
F. GTCTCAGCCGCCGCCTCAA
R: GGAAGAATCGTCGCCGTAAA
F: TAATACGACTCACTATAGGTTCAAGAAGATTAC
R: TAATACGACTCACTATAGGTTTCACTCCACT
42
I, III
II
I, III
I
III
I, III
I, II, III
I, II, III
II
III
III
III
III
I
II, IV
II, IV
II, IV
II, IV
II
II
II
IV
II, IV
IV
II
T7promotor +
PDE8A
F: TAATACGACTCACTATAGGAAGAAGGTAGCAG
R: TAATACGACTCACTATAGGTTCATTGTACGAG
II
4.4 cAMP analyses (I, II, III)
Measurement of cAMP accumulation was determined with a luminometric cAMP assay
(cAMP-Glo TM Assay, Promega) two days after transfections with PDE7 siRNAs (II) or using
the method of Frandsen and Krishna (1976), as described earlier (Ahlström et al. 2000).
Briefly, the cells were grown in 24-well plates without treatment (I) or treated with either
vehicle (ethanol) or Dex (1-100 nM) for 48 h (III). The cells were then induced with forskolin
for 7 min at 37°C and rinsed twice with ice-cold phosphate-buffered saline (PBS) (III) or
washed with Hanks Balanced Salt Solution (HBSS) (I). For determination of basal cAMP
levels, cells were incubated at 37°C for 30 min before washing with ice-cold PBS (III) or
HBSS (I). cAMP was extracted from the cells with 96% ethanol at -18°C for 3 h. The extract
was evaporated with N2 gas and dissolved in assay buffer (0.05 M sodium acetate, pH 6.2). If
necessary, the sample was further diluted with assay buffer, and the cAMP concentration was
determined by radioimmunoassay with iodinated cAMP as tracer.
4.5 Q-sepharose chromatography (I, III)
Five confluent cultures of either NHOst, SaOS-2 or MG-63 cells, grown on 100-mm plastic
dishes were washed twice with PBS buffer, harvested with a cell scraper into 6 ml of ice-cold
homogenization buffer (buffer A), containing 20 mM Bis-tris, pH 6.5, 5 mM
mercaptoethanol, 0.1% of a protease inhibitor cocktail (PIC), consisting of 0.5 mg/ml each of
bestain, pepstatin A, aprotinin, leupeptin, E-64 and 12.5 mg/ml AEBSF. The harvested cells
were then homogenized on ice by ten passages with a Teflon/glass homogenizer. The
homogenization was repeated three times. The homogenate was then centrifuged for 20 min
at 20 000 g. Five millilit of the supernatant was diluted with 20 ml of buffer B, containing 20
mM Bis-tris, pH 6.5, 0.1 M sodium acetate, 0.02% PIC (v/v), 0.1 mM EDTA, 1 mM
benzamidine and 1 mM mercaptoethanol, filtered through a 0.22-µm syringe filter and
applied to a column (5-ml bed volume) of Q-sepharose High Performance (Amersham
Pharmacia Biotech) previously equilibrated with buffer B. After washing the column with 10
bed volumes, the PDE activities were eluted with a 0.1-1.0 M linear sodium acetate gradient
in buffer B, at a flow rate of 2.5 ml/min. Fractions of 2 ml were collected into tubes
containing 50 µl of 5% bovine serum albumin (BSA) and assayed for either cAMP-PDE or
cGMP-PDE activity, as described below.
43
4.6 Enzyme activity analyses
4.6.1 Assay of PDE activity (I, II, III)
PDE activity was assayed according to the method described by Thompson and Appleman
(1971). Samples from either Q-sepharose elutions or whole-cell homogenates were added to
the incubation buffer (40 mM Tris-HCl, pH 8.0, 0.1% PIC, 0.05% BSA, 5 mM MgCl2, 1 mM
EGTA, 0.5 µM cAMP and 3H –cAMP (100000 cpm/tube) to give a final reaction volume of
200 µl. For the cGMP-PDE assay, cAMP was replaced with 0.25 µM cGMP and 3H –cGMP.
The samples were incubated at 34°C for 30-60 min, followed by boiling for 1 min. After
cooling on ice, 50-100 µl of Crotalus atrox snake venom nucleotidase (1 mg/ml, Sigma) was
added and incubated at 34°C for 10 min. Then 500 µl of a 1:3 slurry of AGI-X8 resin
(BioRad) was added, and the tubes were mixed and centrifuged for 5 min at 8000 rpm. The
radioactivity of the supernatant was determined with a β-counter. Selective inhibitors used for
different PDE types are summarized in Table 3.
4.6.2 Assay of ALP activity (II)
Bone-specific alkaline phosphatase (bALP) activity was measured with a immunoenzymetric
assay (OCTEIATM Ostase® BAP, Immunodiagnostic Systems). Prior to measurements, the
cells were washed with PBS and after addition of 300 l of Tris –Triton (0.1 M Tris–base,
0.2% Triton X-1000), the cells were frozen at -70 C. After thawing, the cells were centrifuged
(12000 g, 5 min). The supernatant was used for analysis. The ALP activity was corrected with
total protein content.
4.7 PDE silencing (II)
PDE silencing was performed with siRNAs, and the PDE7 selective inhibitor BRL 50481
(Table 3) was mainly used to confirm results obtain with PDE7 siRNAs. The short doublestranded RNA (dsRNA) mixtures for PDE7 and PDE8 inhibition were generated from PDE7
and PDE8 PCR products. Primers with appended T7 promoters sequenced for the PDE7 PCR
products were designed to amplify both PDE7A and PDE7B. PDE8 PCR products were
designed to amplify only PDE8A. The BLAST database program (NBCI) was used to confirm
the specificity of the target sequences. The large dsRNAs were first synthesized by in vitro
transcription from the DNA templates. The large (150-1500 bp) dsRNAs were processed by
ShortCut RNase III digestion in the presence of manganese, which produced a heterogeneous
population of short (18-25 bp) siRNAs. For generation of siRNAs, we used a ShortCut RNAi
kit (New England BioLabs). After 12-14 days of differentiation, the osteoblast phenotype was
44
confirmed by staining cultures and control cultures with a histological alkaline phosphatase
kit (Sigma). At this stage, the medium was changed to a low serum (2.5%), antibiotic-free
medium, and 24 h later, the siRNA mixtures were transfected into the cells by Lipofectamin
2000 (Invitrogen). Control cells were simultaneously treated with lipofectamin without
siRNAs. The transfection time varied between 6 and 24 h, and the cell culture was then
changed to a new differentiation medium. The transfection efficiency was determined with
BLOCK-iT™ Fluorescent Oligos (Invitrogen) according to the manufacturer`s instructions.
4.8 Isoleucine-proline-proline (IPP), valine-proline-proline (VPP) and leucine-lysine-proline
(LKP) tripeptides (IV)
The peptides were synthesized by 9-fluorenylmethyloxycarbonyl (Fmoc) chemistry and
purified using HPLC reverse phase columns. Peptides were synthesized in the Core Facility
for Synthetic Peptides, Division of Biochemistry, Department of Biological and
Environmental Sciences, University of Helsinki, Finland. Peptides were dissolved in sterile
water prior to administration.
4.9 Cell proliferation analyses (II, IV)
The proliferation rate was assayed with a luminometric ATP detection assay (ATPlite, Perkin
Elmer) 48 h after the siRNA transfections (II), with multilabel counter (Victor 3, Perkin
Elmer) or measured the activity for DNA synthesis (IV) with a Cell Proliferation ELISA kit
(Roche Diagnostics). The 5-bromo-2’-deoxyuridine (BrdU) binds to the newly synthesized
cellular DNA. BrdU incorporation was measured with a spectroscopic plate reader at 450 nm
(Multiskan Ex, Thermo Labsystems).
4.10 Mineralization analyses (II)
Mineralization was assessed by Alizarin Red S staining in cells grown on 96-well plates
using the method described by Stanford et al. (1995). After staining, the dye was eluated with
10% cetylpyridinium chloride in 10 mM sodium phosphate (pH 7.0, 15 min, RT). Absorbance
was measured with a spectroscopic plate reader at 590 nm.
45
4.11 mRNA expression analyses
4.11.1 RNA isolation (I-IV)
RNA of the osteoblastic cells was extracted using the QuickPrep Micro mRNA purification
kit (Amersham Pharmacia Biotech) or the RNeasy Protect mini kit (Qiagen GmbH). The
RNA concentration was measured at 260 nm with a spectrophotometer. The integrity of the
RNA was confirmed by 1.2% formaldehyde agarose gel electrophoresis.
4.11.2 cDNA microarrays (II, IV)
The cDNA microarray hybridizations were performed at the Finnish DNA Microarray Centre
of the Turku Centre for Biotechnology. The expression profiles of osteoblasts differentiated
from hMSC were compared after a 48-h treatment with 35 nM PDE7 or PDE8A siRNA
mixtures (III), or after a 24 h treatment with 50 M IPP, VPP or LKP (IV), with Hum16-K
protocol. The array contains 16 000 human gene probes (accession number: A-MEXP-557,
http://www.ebi.ac.uk/ arrayexpress/). Samples were labelled with FluoroLink Cy-3-dUTP and
Cy-5-dUTP (Amersham Biosciences) using 20 g of total RNA for direct labelling and
hybridized using an augmented reference design (Kerr and Churchill 2001). Discrete images
for Cy-3 and Cy-5 dyes were obtained using a Scan Express laser-scanning microscope
(Packard BioSciences) and gene transcript-levels were determined from the fluorescence
intensities of scanned data image files with the QuantArray Microarray Analysis software
(Packard BioSciences). T-test and gene-expression profiles were analysed by Kensington
Discovery Edition 2.0 software. At least a 1.6 fold difference in expression was used in
control/treatment comparison. The cDNA microarray experiment is available at accession
number E-MEXP-1021 (II) and E-MEXP-885 (IV) (http://www.ebi.ac.uk/arrayexpress/). The
gene expression data for IPP- treated osteoblasts were analysed by principal component
analysis (PCA) to determine the main components affected. Results were considered
significant at a 95% significance level (p < 0.05) (IV).
4.11.3 cDNA synthesis (I, II, III, IV)
cDNA was prepared from 1-2 µg of total RNA or mRNA by reverse transcription with the
following buffer and conditions: RT buffer (Promega) containing 50 mM Tris-HCl (pH 8.3)
75 mM KCl, 3 mM MgCl2 and 10 mM DTT, 13 U RNAguard, Rnase inhibitor (Amersham
Pharmacia Biotech), 0.9 mM each of dATP, dCTP, dGTP and dTTP, 15 pmol Oligo(dT) 15
primer (Promega) and 100 U M-MLV-reverse transcriptase primer (Promega) in a final
volume of 25 µl, with incubation for 1 h at 37°C.
46
4.11.4 Semi-quantitative reverse transcriptase polymerase chain reaction (RT-PCR) (I, III)
PCR amplifications were conducted in 30-µl aliquots containing 2-4 µl of cDNA, 1x buffer
(Finnzymes), comprising 10 mM Tris-HCl, pH 8.8 at 25°C, 1.5 mM MgCl2, 50 mM KCl and
0.1% Triton X-100, 0.2 mM each of dATP, dCTP, dGTP and dTTP, 0.35 µM of each primer
and 1 U of DyNAzyme DNA polymerase (Finnzymes). Primers are listed in Table 4. The
amplification protocol was run on a PTC-100 thermocycler (MJ Research). After denaturation
at 94°C for 4 min, the reactions underwent cycles of denaturing for 1 min at 95°C, annealing
for 90 s at 58-63°C and extension for 2 min at 72°C. This was followed by a single step at
72°C for 10 min. To ensure that the PCR was in the exponential phase, the conditions of the
semi-quantitative RT-PCR were optimised for the number of PCR cycles and the amount of
total RNA applied to the reactions for the PDE subtypes/isoforms and for glyceraldehyde-3phosphate dehydrogenase (GAPDH) or -actin, which was used as an external standard.
Human brain RNA and a cocktail of RNA from rat spleen, brain, kidney and heart was used
to prepare cDNAs used as positive controls. Negative controls included samples in which
cDNA synthesis was performed in the absence of reverse transcriptase. Reactions in which no
cDNA was added were also performed to check for possible non-specific products. Volumes
of 10 µl of the PCR-amplified products were analysed on 1.5% agarose gels containing
ethidium bromide. Gel results were quantified by AlphaDigiDoc™ System 1000 (Alpha
Innotech Corporation). The results were analysed as a ratio of integrated optical density of the
PDE subtypes/isoforms to the external standard.
4.11.5 Quantitative Real-time PCR (qRT-PCR) (II, IV)
PCR amplification was performed in a real-time quantitative PCR engine Mx3000P
(Stratagene). The PCR reactions contained cDNA templates and Brilliant SYBR Green QPCR
Master mix kit (Stratagene) in a 25-µl reaction volume containing 200 nM of each genespecific primer (Table 4). Fluorescence data were collected during the annealing step and
analysed with Mx3000P software. Amplification was obtained by denaturing at 95°C for 10
min, followed by 40 cycles of denaturing at 95°C for 30 s, annealing at 58°C for 1 min, and
extension at 72°C for 30 s. The amplification cycles were followed by for 1 min at 95°C and
for 30 s at 55°C, and data for dissociation plots were collected as the temperature was
returned to 95°C. cDNA from human brain total RNA (BD Biosciences) was used as a
calibration standard, and the data were normalized with amplification data of ß-actin. All
reactions were run in triplicate with three dilution concentrations, and the mean value was
used to calculate the ratio of target gene/ß-actin expression in each sample. Using the ratio in
the untreated sample as a standard (100), the relative ratio of treated sample was presented as
the relative expression level of the target gene.
47
5. Results and discussion
5.1 Identification of PDE families in human osteoblastic cells (I, III)
PDEs has been extensively studied in kidney, brain, heart, muscle and immune system cells of
humans, rats and mice (Lugnier 2006), but very little is known about them in bone cells.
There are no reports of PDEs in human osteoblasts. In Studies I and III, we therefore
examined the profile of PDEs in different types of osteoblastic cells, comprising SaOS-2 and
MG-63 osteosarcoma cell lines and normal human osteoblast (NHOst) cells (Table 5). MG-63
and SaOS-2 are osteoblast-like cells widely used as osteoblast models in in vitro studies
(Murray et al. 1987, Rodan et al. 1987, Lajeunesse et al. 1990).
The cAMP PDE activity of NHOst, SaOS-2 and MG-63 cells separated by Q-sepharose ionexchange chromatography was reflected in three peaks, A1-A3 in Figure 6 and, 1-3 in Figure
7. The first peak of NHOst, SaOS-2 (I) and MG-63 (III) cells consisted of CaM-stimulated
PDE1 activity, which is sensitive to 8-MMX. This peak was also sensitive to cGMP, which is
explained by the dual substrate nature of PDE1. IBMX, but not EHNA, cilostamide,
milrinone or rolipram, also affected this peak. The second peak activity of NHOst and SaOS-2
cells was insensitive to the addition of high levels of several tested inhibitors. IBMX,
however, inhibited the activity of the second peak. PDE7, the cAMP-specific, rolipraminsensitive PDE family, has been shown to be insensitive to most known PDE inhibitors, but
relatively sensitive to IBMX (Hetman et al. 2000), suggesting that the most likely candidate
for peak A2 of both NHOst and SaOS-2 cell is PDE7.
TABLE 5. Summary of PDE families and subtypes expressed in human osteoblasts and
osteoblast-like cells (+ identified, - not identified, / not determined).
PDE1
PDE3
PDE4
_______
1A 1C
_______
3A 3B
__________________
4A
4B 4C 4D
NHost
+
+
+
-
+
+
-
-
+
+
/
/
/
SaOS-2
-
+
+
-
+
+
+
+
+
+
/
/
/
MG-63
+
+
+
-
+
+
+
+
+
+
+
+
+
48
PDE7
PDE8 PDE10 PDE11
_______ ____
7A 7B
8A
_____
10A
___
11A
Figure 6. Elution profile of cAMP-PDE activity of NHOst and SaOS-2 cells: Cell extracts
were applied to a Q-sepharose column and eluted with a linear sodium acetate gradient
(straight line). The eluted fractions of NHOst (A) and SaOS-2 (B) cells were assayed for
cAMP-PDE activity in the absence (o) and presence (g) of 6 µM cGMP, and in the
presence of 20 U/tube CaM/10 µM CaCl2 (n). The eluted cGMP-PDE activities of NHOst
(C) and SaOS-2 cells (D) were assayed in the absence (o) and presence (n) of 20 U/tube
CaM/10 µM CaCl2.
The identity of the third peak was determined by assaying sensitivity to cGMP, and PDE
inhibitors indicated a different identity of A3 peaks of NHOst and SaOS-2 cells (Table 6). In
NHOst cells, A3 was shown to be sensitive to milrinone, a PDE3 selective inhibitor, but
insensitive to rolipram, a PDE4 selective inhibitor. Peak A3 of SaOS-2 cells was sensitive to
rolipram, but relatively insensitive to milrinone. The high sensitivity of the third NHOst peak
to cGMP further suggested that the NHOst peak A3 activity is produced by a member of the
PDE3 family. The only detected cGMP PDE activity in both NHOst and SaOS-2 consisted of
two closely eluting CaM-stimulated peaks. The first of these peaks mostly hydrolysed cGMP,
but the second peak, corresponding to the fractions of cAMP-hydrolysing peak A1,
hydrolysed both cAMP and cGMP. The two activities of MG-63 cells (peaks 2 and 3)
contained rolipram-sensitive PDE4 activity (Figure 7). EHNA, cilostamide and cGMP did not
reduce the activity of these two peaks.
49
Figure 7. Q-sepharose elution profile of cAMP-PDE activity present in MG-63 cells. Cell
extracts were applied to a Q-sepharose column and eluted with a linear sodium acetate
gradient (0.05–1.2 M). Fractions were collected and assayed for cAMP-PDE activity in the
presence (ο) or absence (•) of PDE inhibitors and cGMP. The concentrations of the inhibitors
were as follows: 8-MMX, 25 M; EHNA, 10 M; cilostamide, 5 M; rolipram, 10 M;
IBMX, 50 M. The concentration of cGMP was 5 M.
50
TABLE 6. Sensitivity of the eluted peaks towards selective PDE inhibitors. The results are
given as µM inhibiting half of the maximal PDE activity (IC50).
peak A1
peak A2
peak A3
inhibitor
NHOst
SaOS-2
NHOst
SaOS-2
NHOst
SaOS-2
8-MMX
Milrinone
Rolipram
IBMX
cGMP
11
>100
>100
25
8
>100
>100
17
>100
>100
>100
7
100
>100
>100
>100
6
100
>100
0.13
>100
0.01
>100
10
0.19
100
Peaks 2 and 3 were also sensitive to IBMX. 8-MMX had a mild inhibitory effect on the
activity of peaks 2 and 3, possibly due to non-specific inhibition. IBMX inhibited the activity
of all fractions. The relative PDE1 and PDE4 activities of MG-63 and SaOS-2 cell
homogenates were also assayed (III). MG-63 cells contained higher levels of PDE4 activity
than SaOS-2 cells.
According to biochemical identification, PDE1 is one of the major PDEs in NHOst and
SaOS-2 cells (I) and PDE4 is one of the major PDEs in MG-63 cells (III). PDE expression
may depend on the biological state and environment of the cell. For example, the transcript of
PDE4B2 is more abundant in the myometria of pregnant women than in non-pregnant women
(Leroy et al. 1999). PDE3 activity was detected in NHOst and MG-63 cells, but SaOS-2 cells
lacked this activity. MG-63 and SaOS-2 cells are malignant cell lines that are in different
stages of osteoblastic differentiation, which could in part explain their dissimilar PDE
profiles.
5.2 Identification of PDE subtypes in human osteoblastic cells (I, III)
The presence of mRNA transcripts of all known PDE1, PDE3, PDE4 and PDE7 subtypes was
examined by RT-PCR (Table 5). PDE1A and PDE1C transcripts were expressed in NHOst
and MG-63 cells, whereas SaOS-2 cells only expressed the PDE1C subtype. Of the two PDE3
subtypes, PDE3A and PDE3B, mRNA of the former was detected in all three cell types. The
existence of mRNA transcripts of all known PDE4 subtypes (PDE4A-4D) (Bolger et al. 1993)
was examined; PDE4A and PDE4B subtypes were found in NHOst cells, whereas all PDE4
subtypes were found in MG-63 and SaOS-2 cells. HOSM-1, an osteosarcoma cell line
established from a human mandible, has demonstrated the expression of PDE4A, PDE4B and
PDE4C, and rolipram has been shown to inhibit the proliferation of these cells (Narita et al.
2003). Lower PDE4 activity in NHOst than in SaOS-2 and MG-63 cells may occur because
51
cells expressed only PDE4A and PDE4B subtypes. The expression of PDE4C and PDE4D
could be involved in the proliferation and the malignant phenotype of osteosarcoma cells.
The rolipram-insensitive/IBMX-sensitive cAMP-specific PDE activity detected in NHOst,
SaOS-2 and MG-63 cells indicated the presence of PDE7. PDE7 has two subtypes in
mammals; both mRNA transcripts (PDE7A and PDE7B) were detected in all studied cells. In
addition, the presence of mRNA transcripts of the remaining cAMP PDEs (PDE8A, PDE10A
and PDE11A) was identified in MG-63 cells.
Profiles of PDE subtypes in different osteoblastic cells have previously been studied in rat
UMR-106 osteoblast-like cells (Ahlström and Lamberg-Allardt 1999, 2000), mouse
osteoblastic cell line MC3T3-E1 and bone marrow stromal cell line ST-2 (Wakabayashi et al.
2002). In mouse cell lines, PDE1, PDE2, PDE3, PDE4, PDE7, PDE8 and PDE9 mRNA
transcripts have been identified. In rat UMR-106 osteoblast-like cells, the main PDE fractions
are PDE1 and PDE4, but PDE2, PDE3, PDE7 and PDE8 expression has also been detected.
These findings are line with our results from human osteoblasts and osteosarcoma cells MG63 and SaOS-2. PDE1, PDE2, PDE3, PDE4, PDE7 and PDE8 may be the major PDEs in
osteoblasts, but profiles and subtypes could vary during the differentiation and with
environmental changes.
5.3 PDE7 and PDE8A silencing by siRNAs (II)
In in vivo and in vitro studies, the inhibition of PDE4 has been demonstrated to increase
osteoblastic differentiation and bone formation in rats and mice (Miyamoto et al. 1997, 2003,
Kinoshita et al. 2000, Wakabayashi et al. 2002, Scutt et al. 2004), but the effect of PDE7
inhibition on bone biology has not previously been investigated. At the beginning of our
study, no chemical inhibitors for PDE7 or PDE8 were available. siRNAs have provided the
field of molecular biology with a new and efficient tool to efficiently knock-down specific
genes (Mousses et al. 2003, Bantounas et al. 2004, Semizarov et al. 2004). We investigated
the effect of specific silencing of phosphodiesterase subtypes PDE7A, PDE7B and PDE8A
on gene expression in hMSC-derived osteoblasts. For the silencing of specific PDEs, siRNAs
generated from PDE7 and PDE8 were utilized. Quantitative real-time PCR was used to
confirm the specific PDE silencing. PDE7 and PDE8 inhibition by RNAi significantly
decreased the gene expression of PDE7A, PDE7B and PDE8A. At the level of mRNA, PDE7
and PDE8 inhibition by RNAi decreased the gene expression of PDE7A by 60-70%, PDE7B
by 40-50% and PDE8A by 30%.
Since the discovery of the RNAi method various genes involved in the cell cycle, apoptosis,
differentiation and signalling have successfully been inhibited in different mammalian cell
types, in vitro and in vivo, to elucidate gene function, identify drug targets and treat a variety
of diseases. β-catenin expression has been silencing, leading to the inhibition of growth of
52
colon cancer cells (Verma et al. 2003). The reduction of cyclin B1 levels in HeLa cells by
siRNAs caused inhibition of proliferation by arresting cells in the G2 phase and inducing
apoptosis (Yuan et al. 2006). RNAi targeting TGF-β1 significantly ameliorated the
progression of matrix expansion in experimental glomerulonephritis (Takabatake et al. 2005).
In addition, the gene expression of PDE4B and PDE4D has been reduced with synthetic
siRNAs in human embryonic kidney (HEK293B2) cells (Lynch et al. 2005). Silencing
efficiency of siRNAs is dependent on the sequence of siRNAs, the number of siRNA
molecules and the cell doubling rate. In slowly proliferating or non-dividing cells, siRNAs
seem to have a longer gene silencing effect than in rapidly dividing cells (Pai et al. 2006). For
example, siRNA activity lasts for 3-7 days in proliferating cells, yet persists for three weeks
or more in terminally differentiated cells such as neurons (Omi et al. 2004). We used hMSCderived osteoblasts in our study, when cells differentiated the rate of proliferation has become
limited (Amedee et al. 1994). Our results show that specific gene silencing with the RNAi
method is useful for inhibiting the gene expression of PDE7 and PDE8 subtypes.
5.4 PDE7 silencing has potential for enhancing osteogenic gene expression and
differentiation in hMSC-cells derived osteoblasts (II)
Gene expression profiling was performed using a Hum-16K cDNA microarray. The array
contains 16 000 human genes probes and is designed for genome-wide screening. The array
therefore contains a limited number of known key factors for osteoblastogenesis and bone
formation; Runx 2, Rankl and Lrp5 genes are missing. Treatment for 48 h with PDE7 siRNAs
up-regulated 435 and down-regulated 60 loci ( 1.6-fold difference). The respective numbers
for siRNA treatment of PDE8A were 38 and 96. Selected microarray results are displayed in
Table 7. Quantitative real-time PCR results confirmed up-/down-regulation of a set of genes
from the microarray results (Table 8). The qRT-PCR results were in line with the microarray
analysis. PDE7 silencing up-regulated the gene encoding OCN, a non-collagenous bonespecific matrix protein secreted by normal maturing osteoblasts (Price et al. 1976). OCN
expression is highly up-regulated by 1,25(OH)2-D3 in MG-63 osteosarcoma cells (Mahonen et
al. 1990, Lajeunesse et al. 1991). In addition, the activation of the cAMP/PKA signalling
pathway increases OCN mRNA and the secretion of OCN in rat osteoblast-like cells (Noda et
al. 1988, Theofan et al. 1989). In human osteosarcoma cells, the results of cAMP/PKA
activation on OCN expression are conflicting; OCN expression has been reported to be either
up- or down-regulated by compounds that elevate cAMP (Lajeunesse et al. 1991, Drissi et al.
1997, Boguslawski et al. 2000).
53
TABLE 7. Effects of PDE7 and PDE8A RNAi silencing on gene expression in hMSC-derived human
osteoblasts. A set of interesting genes were selected from cDNA microarray results.
_________________________________________________________________________________________
Accession number
Gene/protein
Fold-change
___________________________________________________________________________
Inhibition of PDE7
Up-regulated
Enzymes and metabolism
H87471
Differentiation
AA779457
Apoptosis
N94588
Kynureninase (L-kynurenine hydrolase)
2.1
Bone morphogenetic protein 5
Catenin beta-like 1
Calcium-sensing receptor
Glutamate receptor metabotropic 3
Tumour necrosis factor (ligand)
superfamily member 10
3.5
2.0
2.0
3.1
CASP8 and FADD-like apoptosis regulator
2.1
3.2
Down-regulated
Hormone and growth factor receptors
Vitamin D (1,25 dihydroxyvitamin D3 receptor) -1.7
Fibroblast growth factor receptor
-5.4
Apoptosis
T95823
cyclin L2
-1.8
Inhibition of PDE8A
Up-regulated
N55459
Metallothionein 1F (functional)
1.6
R01682
Metalloprotease-related protein 1
1.6
Down-regulated
Apoptosis
N94588
CASP8 and FADD-like apoptosis regulator
-1.9
Differentiation
AA045327
Osteoglycin (osteoinductive factor, mimecan)
-1.5
R56774
Bone morphogenetic protein 1
-1.5
___________________________________________________________________________
Fold-change describes the relative expression of the given factor in osteoblast cells treated for
48 h with 35 nM PDE7 or PDE8A siRNAs.
54
TABLE 8. Summary of qPCR results. Effects of PDE7 and PDE8A RNAi silencing on gene
expression in hMSC-derived human osteoblasts.
Gene
PDE7 silencing
PDE8A silencing
OCN
β-catenin
Caspase 8
CREB-5
VDR
BMP-1
BMP-5
PDE7A
PDE7B
PDE8A
PDE4A
PDE4B
Ostoglycin
↑
↑
↑
↑
↓
↔
↑
↓
↓
↔
↔
↔
↔
↔
↔
↓
↔
↔
↓
↔
↔
↔
↓
↔
↔
↓
The effects of gene expression are indicated as follows: up-regulation ↑, down-regulation ↓,
no effect ↔.
The BMPs are known to be important in the process of osteoblast differentiation,
proliferation, apoptosis and morphogenesis (Canalis et al. 2003). The BMP family can be
divided into several subgroups based on BMP gene sequence identity. BMP-1 is unrelated to
the other BMPs and does not regulate the growth and differentiation of skeletal cells (Uzel et
al. 2001). BMP-5, BMP-6, BMP-7 and BMP-8 belong to the same subgroup and all have
osteoinductive properties (Wozney 2002). In our study, PDE7 inhibition caused a threefold
up-regulation of BMP-5 gene expression. The BMPs have been shown to interact closely with
Wnt peptides. The Wnt proteins are secreted by osteoblasts and are required for BMPmediated stimulation of osteoblast proliferation and function (Nakashima et al. 2004, Nelson
and Nusse 2004). Inhibition of PDE7 up-regulated -catenin, a protein that plays a central role
in the Wnt signalling pathway (Rawadi and Roman-Roman 2005), suggesting that PDE7
could participate in Wnt-mediated effects on osteoblast differentiation. In parallel with
differentiation, osteoblasts show increased apoptosis, which is established to be associated
with the activation of caspases (Canalis et al. 2005). Evidence indicates that caspase-mediated
activation of apoptosis is necessary for osteoblastic differentiation (Mogi and Togari 2003).
The effect of PDE7 silencing was to significantly up-regulate caspase-8. In contrast to
caspase-8, vitamin D receptor (VDR) mRNA was reduced in PDE7-silenced cells. VDR
knock-out mice show increased bone volume and density compared with wild-type mice
(Tanaka and Seino 2004), and osteoblasts lacking VDR have enhanced differentiation,
manifested as an increase in both ALP activity and mineralized matrix formation (Sooy et al.
2005). The increased expression of caspase-8 and the decreased expression of VDR seen in
PDE7-silenced cells therefore support PDE7 inhibition promoting an osteoblastic geneexpression pattern. The transcription factors of the CREB family stimulate basal transcription
55
of CRE-containing genes and mediate induction of transcription upon phosphorylation by
protein kinase A. CREB has been demonstrated to have a major role in the expression of
skeleton-specific genes (Sakamoto et al. 1998). In our study, silencing of PDE7 up-regulated
CREB-5, which further suggests that PDE7 is involved in gene expression during osteoblastic
differentiation.
In human T cells, PDE7 has the potential to regulate cell function, including cytokine
production, proliferation and expression of activation markers (Nakata et al. 2002). PDE8A
silencing was also performed, resulting in decreased BMP-1, osteoglycin and collagen 1 gene
expression. As the expression of these genes is known to be associated with the osteoblastic
phenotype, PDE8A inhibition seems unlikely to enhance the differentiation of osteoblasts. In
fact, PDE8 silencing appears to have a negative effect on osteoblastic differentiation. The
microarray findings of PDE7 silencing illustrate up-regulation of several osteogenic gene
expressions, which could reflect augmented osteoblastic differentiation.
5.5 PDE7 inhibition enhances cAMP signalling, differentiation and mineralization in hMSCderived osteoblasts (II)
The effects of PDE7 silencing on cAMP accumulation were studied to evaluate the role of
PDE7 on cAMP signalling. The cells were cultured in differentiation media for two weeks.
PDE7 was silenced with siRNAs for 48 h, and cAMP synthesis was then stimulated with 10
µM forskolin for 20 min. PDE7 silencing increased the forskolin-stimulated cAMP response
more than threefold. As the PDE7 silencing increased both cAMP accumulation and
osteocalcin expression, we investigated the possible involvement of PKA in PDE7 siRNAstimulated osteocalcin expression. Treatment with 5 µM PKA inhibitor H-89 after siRNA
transfections completely abolished the effects of PDE7 siRNAs on OCN expression.
To evaluate bALP activity in hMSC-derived osteoblasts, the transfections were performed
after 13 days of differentiation and bALP activity was measured two weeks after
transfections. The treatment with PDE7 siRNAs significantly increased the bALP levels
compared with control cells. Silencing PDE8 did not significantly change bALP. In
undifferentiated hMSC cells, bALP levels were low (Figure 8A). Mineralization was assessed
by Alizarin red S staining. Cells were treated with siRNAs or with 20 µM of the PDE7
selective inhibitor BRL 50481 (3-(N,N-dimethylsulfonamido)-4-methyl-nitrobenzene). After
21-22 days of differentiation, the mineralization of PDE7 siRNA-treated cells was increased
threefold compared with controls (Figure 8B). Treatment with PDE8A siRNAs had no effect
on mineralization (Figure 8C). Treatment with 20 µM BRL-50481 also increased
mineralization (Figure 8D).
56
(A)
(B)
**
4000
µM/mg protein
100
***
***
0
**
**
**
3000
2000
1000
(C)
(D)
750
1000
nM
nM
nM
10
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35
nM
15
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nM
C
on
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ro
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si
R
H
-8
N
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9
PD
E7
+
H
-8
9
PD
E7
si
R
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A
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on
tr
ol
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5
% Change
200
µM/mg protein
500
250
750
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250
C
on
tr
ol
on
tr
ol
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nM
10
nM
15
nM
35
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20
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R
L
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µM
µM/mg protein
***
Figure 8. Effect of PDE7 and PDE8A silencing with siRNAs on ALP activity and
mineralization in osteoblasts differentiated from hMSC cells. (A) To evaluate the bALP
activity, the cells were grown on 96-well plates for two weeks in differentiation media. After
a two-week treatment with PDE7- or PDE8A-specific siRNAs, bone-specific ALP activity
(bALP) was assayed by immunoassay. To determinate the effect of PDE7 and PDE8A on
mineralization, the siRNAs transfections were performed after 12-14 days of differentiation,
and mineralization was assayed by Alizarin red S staining after 21-22 days of differentiation.
Treatment with PDE7 siRNAs dose-dependently increased mineralization (B), whereas
PDE8 siRNA treatment did not (C). Treating the cultures with the PDE7 selective inhibitor
BRL-50481 (20 µM) for 21 days also increased mineralization compared with controls (D).
Results represent means ±SEM of triplicate determinations (**P<0.01,**P<0.001).
57
Silencing of PDE7 increased both bALP activity and mineralization in hMSC-derived
osteoblasts, implying that PDE7 also has an impact on osteoblast cell functions. Our study
shows that inhibition by PDE7-specific siRNAs and by the chemical PDE7 inhibitor BRL50481 can control osteoblastic differentiation and mineralization.
5.6 Down-regulation of cAMP-PDE by dexamethasone in human osteosarcoma cells (III)
The effect of glucocorticoids on the function of PDEs was studied at the mRNA and enzyme
levels in vitro. Dex, a synthetic glucocorticoid, has been shown to increase cell osteogenic
differentiation and proliferation in bone marrow mesenchymal stem cell (Bellows et al. 1987,
Shalhoub et al. 1992, Jaiswal et al. 1997, Bellows et al. 1998), but impair the proliferative and
metabolic activity of osteoblasts (Patschan et al. 2001). Treatment of MG-63 cells with Dex
significantly decreased the cAMP-PDE activity in a dose- and time-dependent manner. The
effect of Dex on cAMP-PDE activity was reduced by up to 50%, with the half-maximal
inhibition of the cAMP-PDE activity being 5 nM. Treatment with 100 nm Dex for 48 h did
not affect the cGMP-PDE activity in MG-63 cells, indicating that Dex specifically affects
cAMP-PDE activity. The effect of Dex on the cAMP-PDE activity of SaOS-2 cells was less
pronounced, probably due to the lower PDE4 activity in these cells. The sensitivity of
homogenates towards PDE inhibitors EHNA and cilostamide was not affected by Dex
treatment. However, homogenates of the Dex-treated cells were, as expected, less sensitive to
rolipram. The Dex treatment also resulted in homogenates that were slightly less sensitive to
8-MMX. Stimulation with forskolin or rolipram separately only increased the cAMP levels in
the cells moderately. Simultaneous treatment with rolipram and forskolin resulted in 20-fold
and 60-fold increases in cAMP levels in MG-63 and SaOS-2 cells, respectively. Treatment
with Dex did not increase cAMP levels in either cell type. However, the Dex treatment
significantly reduced the potency of rolipram to increase forskolin-stimulated cAMP
response. We detected a substantial decrease in the cAMP-specific PDE activity after
treatment with Dex. The Dex treatment alone did not affect the intracellular cAMP levels, but
taking into account that PDE4 is abundant PDE type in both MG-63 and SaOS-2 cells, the
reduction of the cAMP response in the Dex-treated cell might better describe the effect of an
altered PDE profile of the cells. A previous study with SaOS-2 cells has presented similar
results, showing that Dex alone does not affect cAMP levels (Mori et al. 1999).
5.7 Influence of dexamethasone on PDE subtype mRNA expression in MG-63 and SaOS-2
cells (III)
Expression of the following cAMP-PDE subtypes was detected by reverse transcriptase PCR:
PDE1A, PDE1C, PDE2A, PDE3A, PDE4A, PDE4B, PDE4C, PDE4D, PDE7A, PDE7B,
PDE8A, PDE10A and PDE11A (Table 5). Of these subtypes, Dex reduced the mRNA of
PDE4A and PDE4B subtypes in MG-63 and SaOS-2 cells. The decrease in PDE4 mRNA was
58
dose- and time-dependent, with the half-maximal inhibition being 5–10 nM Dex, with
significant effects seen within 4 h of treatment. Further analysis revealed a reduction in the
mRNA of the PDE4A4 and PDE4B1 isoforms. Transcription of PDE4A1, 4A7, 4A10, 4B2
and 4B3 was also evident, but we could not detect any reduction with Dex treatment in these
isoforms. The observed effects on cAMP-PDE activity and cAMP accumulation were
accompanied by a decrease in the mRNA levels of PDE4A4 and PDE4B1 isoforms. These
presented results thereby support the hypothesis that PDEs play a role in Dex-mediated
effects on cAMP signalling in osteoblasts. In rat cultured osteoblasts, two glucocorticoid
actions have been shown: the stimulation of adenylate cyclase and the inhibition of PDE
(Chen and Feldman 1979). In addition, treatment with Dex increases PTHR mRNA
expression in ROS 17/2.8 cells, but not in opossum kidney cells. Glucocorticoids might
enhance sensitivity to PTH by changing the number of PTH receptors and their affinity to
PTH; this suggests the involvement of a post-transcriptional mechanism (Urena et al. 1994).
5.8 Effects of PDEs in osteoblasts
In summary, Studies I-III revealed that the profiles of PDEs in human osteoblasts are similar
to these in rat and mouse osteoblasts. The main PDEs are PDE1, PDE2, PDE3, PDE4, PDE7
and PDE8, but families and subtypes can vary depending on differentiation stage and
circumstances around the cells. In Study II, we investigated the effects of silencing of PDE7
and PDE8 on gene expression and differentiation in hMSC-derived osteoblasts. Inhibition of
PDE7 enhances differentiation and mineralization. The inhibitors of PDE4 have earlier been
shown to increase osteoblastic differentiation and bone formation. At the sequence level, the
PDE7 family is closely related to the PDE4 family (Omori et al. 2007). This could be one
reason why effects of PDE inhibition are osteogenic in hMSC-derived osteoblasts. A
synthetic glucocorticoid, Dex, could alter the profile of PDE4 subtypes in osteoblastic cells.
Dex did not have a direct effect on intracellular cAMP levels, but did cause a reduction in
cAMP response. Our results suggest an additional mechanism by which glucocorticoids could
be involved in bone loss.
Only a small number of studies exist on the effects of PDEs in bone or bone cells, and most of
these have investigated the effect of PDE4 inhibitors. In rats inoculated with Walker 256/S
carcinoma to induce an artificial osteoporotic state, bone loss was reduced by over 50% after
a two-week treatment with the PDE4 selective inhibitor denbufylline (Miyamoto et al. 1997).
PDE4 inhibitors rolipram, XT-44 and XT-611 have been shown to increase mineralized
nodule formation (Waki et al. 1999) and PGE2-mediated anabolic effects (Miyamoto et al.
2003) in rat bone marrow culture. XT-44 also increases the bone mineral density of
ovariectomized rats. Rolipram and pentoxifylline, a non-selective inhibitor, have been
demonstrated to increase both cortical and cancellous bone mass in mice (Kinoshita et al.
2000). Daily injections of pentoxifylline have been reported to enhance bone formation in
BMP-impregnated collagen disks implanted into the back muscles of mice (Horiuchi et al.
59
2001). Pentoxifylline also promotes the osteoblastic differentiation of ST2 mouse bone
marrow-derived stromal cells, although a recent report suggests that the effects of
pentoxifylline are protein kinase A-independent (Rawadi et al. 2001). More information about
the role of other PDEs in bone formation and how these enzymes are involved in the
regulation of bone metabolism is needed.
Future studies should clarify the profile of PDEs in various differentiation stages, from stem
cells to osteoblasts, osteocytes and osteoclast. The role of PDE7 in bone metabolism could be
investigated in vivo with, for instance, knock-out mice. T-cell functional studies have
revealed that PDE7A knock-out mice have approximately 10% lower body fat than PDE7A /+ and PDE7A -/- mice (Yang et al. 2003). The lower body fat in PDE7A knock-out mice
might also indicate changes in bone tissue.
5.9 Influences of bioactive peptides IPP, VPP and LKP on gene expression and proliferation
in hMSC-derived osteoblasts (IV)
The Hum-16K cDNA microarray data analysis revealed that the bioactive peptide IPP
regulated more genes in hMSC-derived osteoblasts than the bioactive peptides VPP and LKP.
Treatments with 20 µM tripeptides showed that IPP up-regulated 270 genes and downregulated 100 genes. The respective numbers for VPP were 25 and 10, and for LKP 16 and
14. IPP not only regulated a larger number of genes than VPP or LKP, but also was the only
tripeptide to up-regulate osteogenic differentiation factors.
The PCA included 245/270 up-regulated genes and 47/100 down-regulated genes in the
different components. The PCA revealed 45% of the variation in up-regulated genes from the
IPP-treated osteoblasts to be attributed to the first principal component (up-regulated cell
differentiation). The second principal component explained 25.3% (up-regulated cell growth)
and the third 22.6% (up-regulated cell transcription) of the variation (Table 9). The first
component (115 genes) had heavy positive accumulations of genes related to differentiation,
receptors, mitochondria, apoptosis, enzymes, signal transduction, membrane architecture,
membrane function and membrane transport. The second component (47 genes) had heavy
positive accumulations of genes related to cell adhesion, osteogenic differentiation and cell
growth and proliferation, while and third component (83 genes) had heavy positive
associations with genes related to transcription, splicing, translation, EST and ‘unknown
functions’. The up-regulation of these genes indicates that IPP enhances osteoblast
proliferation and differentiation.
Our results also show that IPP increases UMR-106 and hMSC proliferation, but not
proliferation of mature osteoblasts. When the state of differentiation increases, the
60
TABLE 9. Summary of the microarray analysis: number of up-regulated (Up) and down-regulated (Down) genes by IPP, VPP and LKP in
osteoblasts differentiated from hMSCs and the average fold change.
EST, expressed sequence tags.
61
proliferation of the mature osteoblast become limited (Amedee et al. 1994). These results are
in line with ours. Biomaterial studies conducted with an Arg-Gly-Asp-containing peptide
sequence also reveal increased primary calvarial osteoblast cell proliferation in response to
the peptide (Huag et al. 2003, Benoit and Anseth 2005). A set of IPP-regulated genes was
further analysed by qRT-PCR (Figure 9). Microarray analysis had revealed IPP-treated
osteoblasts to express more differentiation genes (BMP-5), transcription factor genes (CREB5) and hormone-related genes (PTHrP) than control cells. Apoptosis-related caspase-8 and
vitamin D receptor gene expression, were decreased due to IPP treatment. In this study, 24-h
treatment with IPP upregulated PTHrP and CREB-5 genes. PTH exerts anabolic and catabolic
effects in vivo on bone (Dempster et al. 1993). In vitro studies on osteoblasts have shown
continuous treatment with very low concentration of PTH to stimulate UMR-106-01 and
Figure 9. The gene expression of IPP treated osteoblasts analysed by a cDNA microarray
method ( ) and quantitative real-time PCR (qRT-PCR; ). At least 1.8-fold difference was
used in control v. treatment comparison in microarray data, and the ratio in the untreated
sample was used as a control (100). qRT-PCR results were normalized by amplification of the
result of -actin. Values are means, with their standard errors depicted by vertical bars, of
triplicate determinations, where all amplified genes have their own controls. Mean values
were significantly different from those of the controls (unpaired Student's t test): ***P<0.001,
**P<0.01,*P<0.05.
62
primary osteoblast proliferation (Dempster et al. 1993, Swarthout et al. 2001). PTHrp is a
polypeptide hormone sharing a common sequence with PTH (Walsh et al. 1997), and both can
function with the PTHR (Walsh et al. 1997). PTHrp produced by osteoblasts is thought to
function locally. Within the skeletal microenvironment, PTHrp propels pluripotential bone
marrow stromal cells towards the osteogenic lineage and also exerts antiapoptotic effects
(Karaplis and Goltzman 2000). The ability of PTH to drive changes in gene expression is
dependent upon activation of such transcription factors as the CREB (Swarthout et al. 2002).
In response to PTH, CREB is phosphorylated and in turn activates transcription. The
increased proliferation in IPP-treated hMSC could be due to increased PTHrp and CREB.
Because cell proliferation and maturation are sequential processes, increased numbers of
pluripotent cells and their augmented maturation into osteoblasts could increase the total
number of osteoblasts. The ability of IPP to induce BMP-5 production can increase osteoblast
differentiation and augment fracture repair, a feature associated with a protein-rich diet
(Bastow et al. 1983, Delmi et al. 1990, Rizzoli and Bonjour 2004).
In summary, IPP, one of the three investigated tripeptides, increase the gene expression of
osteogenic factors in hMSC-derived osteoblasts. IPP has also shown to increase matrix
production, mineralization and decrease RANKL/OPG ration in the same cells (Huttunen et
al. 2007). Tripeptides could transport via hPepT1 transporter both in the small intestine and in
the proximal tubule or hPepT2 in the proximal tubule (see Nielsen and Brodin 2003,
Steffansen et al. 2004), but the transporter and signaling mechanisms in bone cells is still
unclear. Forthcoming studies could focus on the the receptor interactions and cellular
mechanisms of bioactive tripeptides in bone cells.
6. Conclusions
We identified PDE families and their subtypes present in normal human osteoblasts and in
human osteosarcoma (SaOS-2, MG-63) cell lines. The main difference in PDE profiles was
the obvious lack of PDE3 activity in osteosarcoma cells, while PDE3 contributed largely to
the total PDE activity of normal human osteoblasts. The major PDE subtype of MG-63 cells
was PDE4, and MG-63 cells also contained higher levels of PDE4 activity than SaOS-2 cells.
The reason for this difference in PDE profiles is unknown. The osteosarcoma cell lines might
represent a different differentiation stage that is associated with high levels of PDE4
expression, or PDE profiles might have altered during cell transformation from the normal to
the malignant phenotype.
Our results showed that inhibition by PDE7-specific siRNAs and by the chemical PDE7
inhibitor BRL-50481 can control osteoblastic differentiation and mineralization. The findings
also indicated that gene-specific silencing with the RNAi method is useful for inhibiting the
gene expression of specific PDEs. The silencing of PDE7 up-regulates several osteogenic
63
genes at the mRNA level, and PDE7 may therefore play an important role in the regulation of
osteoblastic differentiation.
At the mRNA and enzyme levels, Dex, a synthetic glucocorticoid, decreased gene expression
of PDE4 subtypes 4A and 4B, but did not affect other cAMP-PDEs in MG-63 and SaOS-2
cells. The observed effects on cAMP-PDE activity and cAMP accumulation were
accompanied by a decrease in the mRNA levels of PDE4A4 and PDE4B1 isoforms. The
results support the hypothesis that PDEs play a role in Dex-mediated effects in cAMP
signalling in osteoblasts. PDEs appears to be involved in a mechanism by which
glucocorticoids affect cAMP signalling, demonstrating a new possible route in the formation
of glucocorticoid-induced bone loss involving the down-regulation of cAMP-PDE.
Based on our results, tripeptide IPP increases cell proliferation at the cellular and mRNA
levels. Genes associated with differentiation were up-regulated and apoptosis inducers were
down-regulated. These findings indicate that IPP enhances gene expression in such a way that
increases the differentiation of hMSC into osteoblasts and prolongs their viability.
This work has shown that PDEs possess mechanisms that enhance osteoblast differentiation
and affect cAMP signalling by glucocorticoids. Bioactive peptide IPP augmented the potential
to up-regulate osteogenic and cell viability genes in osteoblasts. Bioactive peptides, such as
IPP, might well contribute to the positive effects that dietary protein has on bone mineral
mass. Understanding the role of PDEs in bone cells is important since bone metabolism,
especially bone formation, could be affected by PDEs. These enzymes are potentially
significant targets for the development of new signal transduction therapies for the treatment
of metabolic bone diseases, for example osteoporosis. However, additional research is needed
to elucidate the mechanisms of PDEs and bioactive peptides on osteoblast differentiation and
bone formation.
64
7. Acknowledgements
This work was carried out at the Calcium Research Unit, Division of Nutrition, Department of
Applied Chemistry and Microbiology, University of Helsinki. I am grateful to Professors
Antti Ahlström and Marja Mutanen for providing excellent working facilities.
I also wish to thank Professor Tapio Palva, Head of the Division of Genetics, Department of
Environmental and Biosciences, for interest in my work and for guiding me through the
bureaucracy.
My warmest gratitude is due to my supervisors, Dr. Christel Lamberg-Allardt for the
opportunity to work with her group and Dr. Mikael Ahlström for introducing me to the world
of bone cells and phosphodiesterases. Both (of you) gave me a great deal of scientific
freedom, guidance when needed and were always ready to listen to my ideas.
I am grateful to the reviewers, Dr. Petri Lehenkari and Dr. Anitta Mahonen, their advice,
constractive criticism and insightful comments that substantially improved this thesis.
I am grateful to my author-editor, Carol Ann Pelli, for revising the English language of the
manuscript.
I thank present and past colleagues at the Calcium Research Unit; Heini Karp, Merja
Kärkkäinen, Virpi Kemi, Marika Laaksonen, Zahirul Islam, Heli Viljakainen and particularly
Minna Huttunen for stimulating teamwork, help and enthusiasm, and Ulrike Riehle for her
contribution and for being such a nice person. It has been a pleasure to share the ups and
downs of academic life with all of you.
My heartfelt thanks are due to my friends; providing for relaxing company and all kinds of
fun over the years. I greatly appreciate the unwavering support of my mother Kerttu, my
brother Harri, my mother-in-law Elise and her husband Juhani. Finally, my deepest love and
gratitude are owed to my husband Pasi and dauhgter Iisa. I am so lucky to have you in my
life.
This work has been financially supported by the Finnish Graduate School in Musculo-Skeletal
Diseases, the National Technology Agency of Finland (TEKES), the Oskar Öflund
Foundation, the Finnish Konkordia Fund, the Finnish Cultural Foundation, the Ella ja Georg
Ehrnrooth Foundation and the Betty Väänänen Foundation.
Sipoo, March 2008
Minna Pekkinen
65
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Erratum sheet
Page 17- line 34, Hentunen et al. 1995 not 1998
Page 20- line 28, GCs not GSs
Page 57- in Figure 8 (A) and publication II (manuscript), page 25- in Figure 6 (A)
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