ROLE OF THE P2Y2 NUCLEOTIDE RECEPTOR IN INFLAMMATION AND INFLAMMATORY DISEASES
A Dissertation presented to
the Faculty of the Graduate School University of Missouri-Columbia
In Partial Fulfillment
of the Requirements for the Degree Doctor of Philosophy
by
CHEN CAO
Dissertation Advisors: Drs. Gary A. Weisman and Laurie Erb
MAY 2015
The undersigned, appointed by the dean of the Graduate School, have examined the dissertation entitled
ROLE OF THE P2Y2 NUCLEOTIDE RECEPTOR IN
INFLAMMATION AND INFLAMMATORY DISEASES presented by Chen Cao, a candidate for the degree of doctor of philosophy, and hereby certify that, in their opinion, it is worthy of acceptance.
______Dr. Gary A. Weisman
______Dr. Laurie Erb
______Dr. Michael J. Petris
______Dr. Dennis B. Lubahn
______Dr. Virginia H. Huxley
ACKNOWLEDGEMENTS
First of all, I would like to sincerely thank my advisor Dr. Gary Weisman, who has always been patient, supportive, and encouraging to me during his guidance of my Ph.D. study. His passion for science, insightful scientific views, and perseverance through challenges during research have shown me an excellent model of a scientist. I would like to deeply thank Dr. Laurie Erb as well, who has been constantly providing me inspirations and advice for experimental design and data interpretation over the past years.
I am also thankful to other members of my doctoral committee: Dr. Michael Petris,
Dr. Dennis Lubahn, and Dr. Virginia Huxley, who have generously given their precious time, thoughtful comments, and inspiring suggestions.
I am grateful for having worked with the current and former members of Dr.
Weisman’s and Dr. Petris’ groups, who are all wonderful colleagues teaching me experiment techniques, and helping me whenever I am in need: Jean Camden, Deepa
Ajit, Lucas Woods, Mahmoud Khalafalla, Farid El-Sayed, Olga Baker, Jennifer
Hamilton, Qixing Liang, Zhongji Liao, Troy Peterson, Christina Thebeau, Yanfang
Wang, Victoria Hodgkinson, Erik Ladomersky, Karen Nickelson, Vinit Shanbhag, Ying
Wan, and Sha Zhu.
Finally, I would like to thank my family. My husband, Meng Sun, and my parents,
Jingfu Cao and Zhaoli Zhou, who have been unconditionally supporting me so that I can pursue my goal. I especially owe to my grandfather, Songqin Zhou, who I was not able to visit before him passing away during my last semester of Ph.D. study. In addition, I am in ii gratitude to my brothers and sisters in Columbia Chinese Christian Church for their encouragement, help, prayers, and love, and I thank our Father in heaven for His blessings to me, and for His beautiful and wonderful creatures.
iii TABLE OF CONTENT
Acknowledgments ...... ii
List of Figures ...... vi
List of Tables ...... viii
List of Abbreviations ...... ix
Abstract ...... xiii
Chapter
I. Introduction ...... 1
A. Inflammation ...... 1
B. Leukocytes and Vascular Endothelial Cells in Inflammation ...... 2
1. Leukocyte Chemotaxis and Chemokinesis ...... 3
2. Vascular Barrier and Leukocyte Transendothelial Migration ...... 5
C. Hematopoiesis and Leukocyte Emigration from the Bone Marrow ...... 9
1. Hematopoiesis ...... 9
2. Emigration of Hematopoietic Stem/Progenitor Cells and Leukocyte from the Bone Marrow to Blood ...... 13
3. Homing of Hematopoietic Stem/Progenitor Cells and Leukocytes to the Bone Marrow ...... 19
D. Alzheimer’s Disease ...... 21
E. Purinergic Signaling in Inflammation ...... 25
1. Nucleotides and Purinergic Receptors ...... 25
2. Purinergic Signaling in Inflammation ...... 28
3. Purinergic Signaling in Alzheimer’s Disease ...... 33 iv II. The P2Y2R Interacts with VE-Cadherin and VEGF Receptor-2 to Regulate Rac1 Activity in Endothelial Cells ...... 47
Abstract ...... 47
Introduction ...... 48
Results ...... 51
Discussion ...... 58
Materials and Methods ...... 64
III. P2Y2 Receptor Deficiency in an Alzheimer’s Disease Mouse Model Decrease CXCL12, SCF and VCAM-1 Expression in the Bone Marrow and Reduces Levels of Circulating Monocytes and Neutrophils ...... 98
Abstract ...... 98
Introduction ...... 99
Results ...... 103
Discussion ...... 109
Materials and Methods ...... 116
IV. Summary and Future Directions ...... 140
A. Summary ...... 140
B. Future Directions ...... 141
References ...... 147
Vita ...... 195
v LIST OF FIGURES
Figure I-1. Leukocyte chemotaxis, recruitment, and transendothelial migration ...... 36
Figure I-2. Hematopoiesis...... 38
Figure I-3. HSC mobilization from and homing to the bone marrow ...... 40
Figure I-4. P2Y receptor function in Alzheimer’s disease ...... 44
Figure II-1. Translocation of eGFP-P2Y2R to peripheral membranes of endothelial cells in response to UTP ...... 70
Figure II-2. UTP causes transient interaction between the HA-P2Y2R, VE-cadherin and VEGFR-2 ...... 72
Figure II-3. VE-cadherin inhibits UTP-induced internalization of the HA-hP2Y2R in 1321N1 astrocytoma cells ...... 74
Figure II-4. Activation of the P2Y2R by UTP causes sustained tyrosine phosphorylation of VE-cadherin ...... 76
Figure II-5. Src and VEGFR-2 kinase activity are necessary for P2Y2R-mediated tyrosine phosphorylation of VE-cadherin and Rac1 activation ...... 78
Figure II-6. The Src-binding domain of the P2Y2R is required for the P2Y2R to interact with and phosphorylate VE-cadherin ...... 80
Figure II-7. UTP stimulation does not affect the pool of β- or γ-catenin or the binding of β- or γ-catenin to VE-cadherin ...... 82
Figure II-8. VE-cadherin expression is required for P2Y2R-mediated activation of Rac1, but not ERK1/2 ...... 84
Figure II-9. Effect of down-regulation of VE-cadherin on UTP-induced activation of Rho in HCAECs ...... 86
Figure II-10. P2Y2R activation causes tyrosine phosphorylation of p120 catenin that is dependent upon Src and VEGFR-2 kinase activities and expression of VE-cadherin ...... 88
Figure II-11. Expression of p120 catenin is required for P2Y2R-mediated activation of Rac1 ...... 90
vi Figure II-12. P2Y2R activation causes interaction between p120 catenin and vav2 ...... 92
−/− Figure II-13. In vivo permeability in spleen, colon, and kidney is reduced in P2Y2R mice ...... 94
Figure II-14. A schematic of the signaling pathways activated by the P2Y2R in endothelial cells that are thought to regulate vascular permeability ...... 96
Figure III-1. Cellularity in the peripheral blood ...... 124
Figure III-2. Cellularity in the bone marrow ...... 126
Figure III-3. Marginated pools of neutrophils and monocytes in the spleen ...... 129
Figure III-4. Expression levels of CXCL12, SCF, and VCAM-1 in the bone marrow ...131
Figure III-5. CXCL12 levels in the peripheral blood, cortex and spleen ...... 134
Figure III-6. Expression levels of M-CSF in the bone marrow and cortex ...... 136
Figure III-7. Expression level of CCL2 in the cortex ...... 138
vii LIST OF TABLES
Table I-1. P2Y receptor agonist, signaling pathways, and functions ...... 42
Table III-1. Cellularity in the peripheral blood ...... 122
viii LIST OF ABBREVIATIONS
Aβ amyloid-β
ACKR atypical chemokine receptor
AD Alzheimer’s disease
ADAM a disintegrin and metalloproteinase
ADP adenosine 5’-diphophate
APP amyloid precursor protein
ATP adenosine 5’-triphophate
BBB blood-brain barrier
BM bone marrow
CAA cerebral amyloid angiopathy
CAR cells CXCL12 abundant reticular cells
CBC Complete Blood Count
CCL C-C chemokine ligand
CCR C-C chemokine receptor
CKR chemokine receptor
CSF colony-stimulating factor
CXCL C-X-C chemokine ligand
CXCR C-X-C chemokine receptor eGFP enhanced green fluorescent protein
ERK extracellular signal-regulated kinases
ix FPR formyl peptide receptor
GAPDH glyceraldehyde 3-phosphate dehydrogenase
G-CSF granulocyte colony-stimulating factor
GEF guanine nucleotide-exchange factor
GM-CSF granulocyte/macrophage colony-stimulating factor
GPCR G protein-coupled receptor
GTPase guanosine 5’-triphosphatase
HA hemagglutinin
HCAEC human coronary artery endothelial cell
HPC hematopoietic progenitor cell
HSC hematopoietic stem cell
HSPC hematopoietic stem/progenitor cell
IB/WB immunoblotting/Western blotting
ICAM-1 intercellular adhesion molecule-1
IF interferon
IL interleukin
IP immunoprecipitation
JAM junctional adhesion molecule
LPS lipopolysaccharide
LSK Lin–Sca-1+c-Kit+
LT-HSC long-term hematopoietic stem cell
M-CSF macrophage colony-stimulating factor
x MAPK mitogen-activated protein kinase
MPP multipotent progenitor
N-cadherin neural cadherin
NF-κB nuclear factor κ-light-chain-enhancer of activated B cells
NO nitric oxide
P2Y2R P2Y2 receptor
PECAM-1 platelet-endothelial cell adhesion molecule-1
PLC phospholipase C
PRR pattern recognition receptor
RBC red blood cell
RGD arginine-glycine-aspartate
RGE arginine-glycine-glutamate
Sca-1 stem cell antigen 1
SCF stem cell factor
ST-HSC short-term hematopoietic stem cell
TEM transendothelial migration
TLR Toll-like receptor
TNF tumor necrosis factor
UDP uridine 5’-diphosphate
UTP uridine 5’-triphosphate
VE-cadherin vascular endothelial cadherin
VCAM-1 vascular cell adhesion molecule-1
xi VEGF vascular endothelial growth factor
VEGFR vascular endothelial growth factor receptor
WBC white blood cell
WT wild type
xii Role of the P2Y2 Nucleotide Receptor in Inflammation and
Inflammatory Diseases
Abstract
Nucleotides (ATP, ADP, UTP and UDP) are released by apoptotic and leaky cells, or from activated/stimulated neurons, platelets, vascular endothelial and smooth muscle cells, leukocytes, and bone stromal cells in non-lytic ways. By activating receptors
(P2X1-7 ion channel receptors and P2Y1,2,4,6,11-14 G protein-coupled receptors) on the cell surface, extracellular nucleotides trigger purinergic signaling that regulates inflammation, a critical process in response to traumatic, toxic, infectious, ischemic and autoimmune injuries. During inflammation, leukocytes are recruited from the bone marrow (BM) and to the bloodstream and cross the vasculature at sites of inflammation. However, chronic or exaggerated inflammation is detrimental to tissue healing. Therefore, it is important to modulate inflammatory responses in numerous diseases, including multiple sclerosis, arteriosclerosis, rheumatoid arthritis, asthma, juvenile diabetes and Alzheimer’s disease
(AD). Accumulating evidence indicates that the P2Y2 nucleotide receptor (P2Y2R) is an important regulator of many inflammatory events. Here, the role of P2Y2R in leukocyte migration, recruitment, and vascular inflammation are explored by using both in vitro experiments and in vivo studies with an animal model of AD.
Leukocyte infiltration is a crucial step during inflammation and P2Y2R has been shown to regulate leukocyte chemotaxis, chemokinesis and recruitment to sites of injury or infection. A regulatory role of P2Y2R in endothelial cells is also recognized, including upregulation of adhesion molecules that facilitate leukocyte attachment to blood vessels.
xiii We hypothesized that the endothelial P2Y2R also contributes to the transmigration of leukocytes and macromolecules through blood vessels by regulating interendothelial adhesion junctions that control the permeability of the vascular barrier. Vascular endothelial cadherin (VE-cadherin) is the major component of adhesion junctions between adjacent endothelial cells whose adhesion function is controlled by Rho
GTPases. In Chapter II, we show that P2Y2R when activated by UTP formed a transient complex between VE-cadherin and VEGF receptor-2 (VEGFR-2) and triggered RhoA and Rac1 activation in human coronary artery endothelial cells (HCAECs). Knockdown of VE-cadherin expression with siRNA did not affect UTP-induced activation of extracellular signal-regulated kinases 1/2 (ERK1/2) but led to a loss of UTP-induced
Rac1 activation and tyrosine phosphorylation of p120 catenin, a protein that interacts with the cytoplasmic tail of VE-cadherin. Activation of the P2Y2R by UTP also caused a prolonged interaction between p120 catenin and vav2 (a guanine nucleotide exchange factor for Rac1) that correlated with the kinetics of UTP-induced tyrosine phosphorylation of p120 catenin and VE-cadherin. Inhibitor of the kinase activity of
VEGFR-2 (SU1498) or Src (PP2) significantly diminished UTP-induced Rac1 activation, tyrosine phosphorylation of p120 catenin and VE-cadherin, and association of the P2Y2R with VE-cadherin and p120 catenin with vav2. These findings suggest that the P2Y2R uses Src and VEGFR-2 to mediate its association with VE-cadherin in endothelial adherens junctions to activate Rac1. Blood flow-caused shear stress is known to cause
ATP release from red blood cells and endothelium as well as association of VE-cadherin and VEGFR-2 in endothelial cells. We used an in vivo permeability assay to show
−/− reduced permeability in splenic, colonic and renal vessels in the P2Y2R mice compared
xiv to age-matched wild type controls, which suggests that increase in basal vascular permeability properties in response to shear stress in spleen, colon and kidney are likely physiological effects of the P2Y2R-mediated VE-cadherin signaling including Rac1 activation. Whether signaling through the P2Y2R regulates vascular permeability and leukocyte extravasation under inflammatory condition needs to be determined in future studies.
Alzheimer’s disease (AD), featuring chronic inflammation and neurodegeneration, is the most prevalent form of dementia in Western society. Immune responses involving microglia-mediated clearance of amyloid-β (Aβ) plaques in the AD brain have been intensively studied, while it is unclear how systemic immune responses contribute to disease progression. Recently, circulating monocytes and neutrophils derived from the
BM have been shown to accumulate around Aβ plaques in the brain and are thought to contribute to Aβ clearance in AD mouse models. We hypothesized that the protective role for the P2Y2R in AD is not only limited within the brain but also in regulation of systemic immunity. In Chapter III, we report mild neutrophilia in peripheral blood samples from the TgCRND8 (Tg+) mouse model of AD. Neutrophilia occurs in male Tg+ mice at 10 weeks of age but is abrogated in Tg+ mice with heterozygous knockout of the
+ +/− + P2Y2R (Tg P2Y2R ). Similarly, the number of circulating monocytes in Tg mice was modestly increased compared to wild type mice, whereas the circulating monocyte level
+ +/− in Tg P2Y2R mice was abnormally low. We previously reported that suppression of
+ + P2Y2R expression in Tg mice accelerates AD-like pathology and reduces CD11b microglia/leukocyte accumulation around Aβ plaques in the brain. The deficiency of
+ +/− circulating leukocytes in Tg P2Y2R mice likely contributes to the reduced
xv accumulation of leukocytes around Aβ plaques as well as the accelerated progression of
AD symptoms observed in these mice. Assessment of gene expression in BM samples by
+ +/− quantitative real time polymerase chain reaction (RT-PCR) indicated that Tg P2Y2R mice have a striking reduction in the expression of C-X-C chemokine ligand 12
(CXCL12, also known as stromal cell-derived factor 1), stem cell factor (SCF, also known as Kit ligand) and vascular cell adhesion protein 1 (VCAM-1) compared to Tg+ mice. Loss of these molecules in the BM is generally considered to facilitate leukocytes and hematopoietic stem/progenitor cells emigration out of BM to the peripheral blood.
Therefore, the inability of leukocytes to emigrate out of the BM in Tg+ mice lacking sufficient P2Y2R expression is likely due to the essential role of the P2Y2R in sensing and amplifying chemoattractant signals. This work has revealed a novel function of the
P2Y2R in AD in the regulation of leukocyte release into the bloodstream and the modulation of BM homeostasis.
Taken together, these data expand our understanding of P2Y2R-mediated signaling in inflammatory diseases that regulates leukocyte mobilization, recruitment and transendothelial migration, i.e., the P2Y2R regulates the BM homeostasis and leukocyte chemotaxis/chemokinesis to modulate leukocyte emigration out of the BM into the circulation, and nucleotide release from stressed tissues activates the P2Y2R in endothelial cells to promote VCAM-1-dependent adherence of circulating leukocytes to endothelium, followed by P2Y2R association with VE-cadherin and VEGFR-2 at the adhesion junctions to promote nucleotide-induced phosphorylation of VE-cadherin and
Rac1 activation that facilitate leukocyte infiltration into inflamed tissue.
xvi CHAPTER I
Introduction
A. Inflammation
Inflammation is a response recognized long ago conventionally featuring redness
(rubor), swelling (tumor), heat (calor) and pain (dolor) in response to traumatic, toxic, infectious, ischemic or autoimmune injury [1]. Nowadays, many diseases have been found to be inflammatory, including Alzheimer’s disease, multiple sclerosis, arteriosclerosis, rheumatoid arthritis, asthma, juvenile diabetes and others [1]. While the role and mechanism of inflammation in each condition are still under intensive investigation, inflammation is generally considered as a beneficial response initially but can be detrimental if prolonged or exaggerated [1, 2]. Therefore, appropriate immune homeostasis must be maintained by inflammatory mediators, such as cytokines, which have effects on many types of cells [2].
Cytokines are small molecules (mostly proteins or glycoproteins) secreted by a variety of cells to properly regulate immune responses, and include interleukins (ILs), interferons (IFNs) and growth factors. Based on the effects in terms of causing or preventing inflammation, cytokines can be called proinflammatory or anti-inflammatory, respectively, yet some cytokines have both potentials depending on the cell type and state in response to the cytokine. As Borish and Steinke summarized, “cytokines are involved in virtually every facet of immunity and inflammation, including innate immunity, antigen presentation, bone marrow differentiation, cellular recruitment and activation,
1 and adhesion molecule expression” [3]. Chemokines are a subset of cytokines featuring chemotactic properties and share conserved cysteine residues in the N-terminus.
Depending on the sequence, chemokines are grouped into four families, the C-X-C ligands (CXCLs), C-C ligands (CCLs), XC ligands (XCLs, also referred to as C ligands), and C-X3-C ligands (CX3CLs), and their receptors are named as CXCRs, CCRs, XCRs, and CXC3Rs, respectively. These chemokine receptors (CKRs) are all G protein-coupled receptors (GPCRs) but a few atypical CKRs (ACKRs) only resemble conventional CKRs structurally without similar signaling functions; instead, ACKRs negatively regulate chemotaxis by ligand scavenging or gradient shaping [4]. Cytokines and chemokines can be up- or down-regulated in inflammatory diseases yet must be finely tuned to modulate cellular activities in innate and adaptive immunity so as to react but not overreact to inflammatory stimulation. In some cases, stimuli such as molecules of microbes in infection and amyloid-β in AD, are chemoattractant for immune cells.
B. Leukocytes and Vascular Endothelial Cells in Inflammation
Typically when an inflammatory condition occurs, leukocytes in the circulation sense the inflammatory signals (e.g., bacterial lipopolysaccharides and chemokines) from the sites of infection or injury, and are recruited to these sites to react to the stimuli. As the semipermeable barrier between the circulation and peripheral tissues, the monolayer of vascular endothelial cells formed throughout the vascular system is a critical component of inflammatory responses by actively controlling the traffic of leukocytes as well as cytokines, chemokines and other molecules.
2 1. Leukocyte Chemotaxis and Chemokinesis
Both directional and random motility, termed chemotaxis and chemokinesis, respectively, are important in leukocyte activities. Leukocytes are highly dynamic cells capable of rearranging their cytoskeleton to reshape and polarize the cell according to the environment. In a migrating leukocyte, lamellipodia are formed at the leading edge while uropods appear at the trailing edge. During the transformation and migration repeating polymerization and depolymerization of F-actin occurs, along with membrane protrusions and contractions, and modulation of adhesive properties. The conventional CKRs when activated by their chemokine ligands, release associated Gα and Gβγ subunits. The Gβγ subunit are generally thought to activate the phospholipase C (PLC) pathway that hydrolyzes phosphatidylinostitol-4,5-bisphophate (PIP2) into inositol-1,4,5-trisphosphate
(IP3) and diacylglycerol (DAG), leading to mobilization of intracellular calcium and downstream events [5, 6]. Many small guanosine 5’-triphosphatases (GTPases) have been shown to regulate leukocyte motility, such as Rap in integrin activation and induction of polarity, Rac in lamellipodia formation, Rho in integrin activation and uropod contraction, and Cdc42 in stabilization of a leading edge and directed motility [6]. By modulating their kinases and guanine nucleotide-exchange factors (GEFs), activated CKRs rearrange cytoskeleton to form lamellipodia and uropods via small GTPase signaling [6]. For example, as a consequence of CXCR4 activation GalDAG-GEFI, a Rap1 GEF is activated to render integrin activation through Rap1 signaling in lymphocytes, neutrophils, and platelets [7]. Blockage of the Rho kinase ROCK inhibited T lymphocyte chemotaxis toward the CCR7 ligands, CCL19 and CCL21 [8], while loss of Rac GEF
DOCK2 in lymphocytes (but not monocytes) resulted in impaired F-actin formation, cell
3 polarity and migration [9, 10]. Recently, vesicle fusion during leukocyte chemotaxis was shown to be mediated by members of the synaptotagmin family of calcium-dependent regulators using RNA-interference to identify the mediators as SYT2, SYT7 and a related protein SYTL5. In both T lymphocytes and monocytes, knockdown of SYT2 enhanced and knockdown of SYTL5 diminished their migration towards CXCL12 and CCL2, respectively; loss of the lysosomal membrane protein SYT7, a positive regulator of lysosome exocytosis, inhibited T lymphocyte chemotaxis towards CXCL12 with impaired uropod release [11]. Meanwhile, in the Rab family of GTPases involved in lysosome trafficking to the membrane, Rab27a and Rab3a were determined to play a role in leukocyte chemotaxis [11].
Besides chemokines, the innate immune system senses microorganism-associated molecular patterns (MAMPs, also referred to as pathogen-associated molecule patterns, or PAMPs) such as bacterial lipopolysaccharide (LPS), di- or triacylated lipopeptides, peptidoglycan, flagellin, nucleic acids and formylated peptides, which bind to pattern recognition receptors (PRRs) and trigger immediate responses by the innate immune system including chemotaxis, or initiates further responses by the adaptive immune system [12]. The toll-like receptor (TLR) family of type I transmembrane proteins with an ectodomain is essential among the PRRs for their capability of detecting many of the
MAMPs [13]. After the binding of specific MAMPs, TLRs recruit Toll-IL-1 receptor
(TIR) domain-containing adaptor proteins, e.g., MyD88, TIRAP (also known as Mal),
TRIF, or TRAM, and activate of MAPK (mitogen-activated protein kinase)- and NF-κB
(nuclear factor κ-light-chain-enhancer of activated B cells)-dependent pathways giving rise to production of inflammatory cytokines [13]. TLR4 may be the most complex
4 member in the family, since it utilizes both MyD88- and TRIF-dependent pathways.
When activated by LPS in presence of the accessory molecule MD2 [14], the MyD88- dependent pathway leads to early activation of MAPK and NF-κB which upregulates production of proinflammatory cytokines such as IL-8; then endocytosed TLR4 also triggers the TRIF-dependent pathway, causing late activation of NF-κB and expression of type I IFN [13]. Another type of intensively studied PRRs is the formyl peptide receptor
(FPR) family. Although the function of FPR3 has not been well understood, FPR1 is known to sense bacterial formyl peptides while FPR2 responds to phenol-soluble modulins and some other MAMPs [12]. A prototypical agonist of FPR1 is N-formyl-Met-
Leu-Phe (fMLF) from Escherichia coli, which is well known to induce calcium flux and chemotaxis in neutrophils [15]. An interesting recent finding about the FPR1 and FPR2 in mouse neutrophils is that when these FRPs were activated by the common non-peptidyl agonist compound 43, the expression of CXCR2 was diminished and LPS-induced recruitment was impaired [15]. In addition, GPCR41 and GPCR43 (also known as free fatty acid receptor 3 and 2, respectively) have been found to respond to short-chain fatty acids including acetate, propionate and butyrate, and mediate chemotaxis through intracellular calcium mobilization [16, 17].
2. Vascular Barrier and Leukocyte Transendothelial Migration
When resident immune cells are unable to resolve the inflammatory condition, more leukocytes are recruited to the site from the blood across the vascular barrier. This process is termed diapedesis or transendothelial migration (TEM) (see Figure I-1). The barrier function of endothelium is largely dependent on interendothelial junctions,
5 including adherens junctions, tight junctions and gap junctions, which are stabilized by the binding of integrins to extracellular matrix [18]. Adherens junctions are mainly comprised of cadherins, and endothelial cells express their exclusive vascular endothelial cadherin (VE-cadherin) as well as neural cadherin (N-cadherin), which share similar structures and catenins bound in the cytoplasmic domain [19]. Only VE-cadherin mediates homotypic interendothelial adhesion, but N-cadherin locates on the abluminal surface and mediates heterotypic interaction between endothelial cells and vascular smooth muscle cells or pericytes [19]. While the N-terminal extracellular domain of VE- cadherin form Ca2+-dependent homophilic adhesion between adjacent endothelial cells, the C-terminal cytoplasmic tail contains binding sites for other junctional proteins, such as β- or γ-catenin (also known as plakoglobin) that forms a dynamic interaction with α- catenin, which is linked to the actin cytoskeleton [20]. Another VE-cadherin-bound protein, p120 catenin regulates the activity of small Rho GTPases, thereby modulating actin cytoskeletal organization and cell motility [21-23].
Tight junctions are dense net-like ultrastructural organizations involving the transmembrane proteins occludin, junctional adhesion molecules (JAMs), and the claudin family as well as their intracellular adaptors zonula (or zona) occludins, ZO-1, -2 and -3
[24-26]. Besides the tight junction transmembrane proteins (i.e., claudins, occludin and
JAMs), ZO-1 can form independent complexes with ZO-2 and ZO-3 [27], and has domains for binding of actin, α-catenin, and several signaling proteins [24]. Despite contradictory results from epithelial cells, the regulation of the actomyosin cytoskeleton by ZO-1 in endothelial cells was demonstrated recently. In endothelium, ZO-1 is required for junctional localization of active myosin II and maintenance of VE-cadherin-mediated
6 cell-cell tension, as well as junctional recruitment of claudin-5 and JAM-A (also named
JAM-1), which are critical for the barrier function [28]. ZO-1 was also found to associate with p120 catenin, which is mediated by Afadin and induced by oxidized phospholipids with coincident endothelial barrier reinforcement [29].
In addition, gap junctions formed by connexons (also called hemichannels, consisting of connexins) mediate cell-to-cell communication by allowing the passage of electrical signals and small molecules [30]. Moreover, other molecules at the junctional region may also contribute to regulation of leukocyte TEM, such as platelet-endothelial cell adhesion molecule-1 (PECAM-1, also known as CD31). This transmembrane protein is expressed on endothelial cells, platelets, and subsets of leukocytes, and mediates cell-cell adhesion between adjacent endothelial cells and serves as a scaffold for recruiting and docking of other junctional proteins, including β- and γ-catenins, as well as phosphatases and kinases that regulate the junctional function [25].
Disruption of interendothelial junctions can be triggered by inflammatory mediators such as vascular endothelial growth factor (VEGF), thrombin, tumor necrosis factor α
(TNF-α) and histamine, whereas the contractility of endothelial cells provided by the actomyosin cytoskeleton under the control of Rho GTPases also contributes to the passage of macromolecules and leukocytes [18, 31]. In fact, endothelial cells need to be activated by proinflammatory mediators to express cytokines, growth factors, chemokines (e.g., IL-1, IL-6, IL-8, granulocyte-macrophage colony-stimulating factor,
CCL2, CCL5), and cell adhesion molecules that mediate the chemoattraction, tethering, rolling, adhesion and TEM of leukocytes [1]. Leukocytes initially attach to the endothelial cells via the affinity of L-selectin, PSGL-1, CD44, CD43, and E-selectin
7 ligand-1 (ESL-1) for the endothelial E- and P-selectins. The interaction between rolling leukocytes and endothelial cells is reversible until reinforced by the binding of α4β1 integrin (also called very late antigen-4 or VLA4) and two members of the β2 integrin family, αLβ2 (function-associated antigen-1 or LFA-1, or CD11a/CD18) and αMβ2
(macrophage antigen-1 or Mac-1, or CD11b/CD18) to their cognate ligands, vascular cell adhesion molecule-1 (VCAM-1) and intercellular adhesion molecule-1 (ICAM-1), respectively, i.e., stable adhesion [32]. Specifically, neutrophil adhesion was found to be mediated by the αLβ2 integrin (LFA-1) while crawling was mediated by the αMβ2 integrin
(Mac-1) [33].
Nonetheless, just as the transportation of plasma proteins, solutes, and liquid occurs via paracellular and transcellular pathways, TEM of leukocytes also has a “transcellular” route besides the relatively well-understood “paracellular” route, but the mechanism underlying the transcellular pathway remains elusive [25]. Subcellular membranous structures in endothelial cells involved in the transcellular passage of fluid and plasma proteins, such as caveolae, fenestrae, and vesiculo-vacuolar organelles, may also affect transcellular TEM of leukocytes [25]. ICAM-1-αLβ2 integrin-mediated adhesion seems to favor transcellular over paracellular TEM of neutrophils, and the cytoplasmic tail of
ICAM-1 is critical [34]. The preference for the transcellular route may rely on the strength of intercellular junctions within the endothelial monolayer under the influence of local cytokine exposure. For example, IL-1β reduces transcellular TEM of monocytes by disruption of the adherens junctions [35]. In transcellular TEM, the endothelial cells were found to form a dome to seal the migrating neutrophil and prevent leakage [36].
8 C. Hematopoiesis and Leukocyte Emigration from the Bone Marrow
Before leukocytes become available in the circulation to be recruited in response to inflammatory stimuli, they need to be mobilized primarily from the bone marrow (BM), where they are produced and preserved in adult mammals. The primitive hematopoietic stem cells (HSCs) give rise to all varieties of blood cells, i.e., a process termed hematopoiesis, in complex microenvironments (termed “niches”) in the BM surrounded by stromal cells including mesenchymal stem cells and their progeny, chondrocytes, osteoblasts, fibroblasts, adipocytes, endothelial cells, and myocytes [37].
1. Hematopoiesis
The hierarchy of HSC differentiation into mature blood cells is well established, and subsets of hematopoietic cells present specific surface proteins that can serve as markers for cell type identification (Figure I-2, recently reviewed in [38]). While lineage (Lin)- committed hematopoietic cell markers are not found on HSCs, CD34 has been successfully used as a specific marker of human HSCs, but not murine HSCs which express low levels of or no CD34 [39]. Instead, stem cell antigen 1 (Sca-1, also known as
Ly6A/E) and c-Kit (also known as CD117, or stem cell factor receptor or SCFR) are considered as markers for murine HSCs, so the Lin–Sca-1+c-Kit+ (LSK) subpopulation is often taken as HSCs. In fact, the majority of LSK cells are CD34+ short-term HSCs (ST-
HSCs with limited self-renewing capacity) and committed multipotent progenitors
(MPPs), while the rest are long-term HSCs (LT-HSCs with extensive self-renewing potential) without expression of CD34 or Flt3 (Fms-like tyrosine kinase 3, also known as
CD135, or fetal liver kinase-2 or Flk-2) [39-42]. Therefore, LSK cells should be referred
9 to as hematopoietic stem/progenitor cells (HSPCs), among which LSKCD34+Flt3– ST-
HSCs produce less-primitive LSKCD34+Flt3+ MPPs [42]. Alternatively, labeling of members of the signaling lymphocytic activation molecule (SLAM) family can be used, i.e., CD150+CD244 − CD48 − for HSCs, CD244+CD150 − CD48 − for MPPs, and
CD48+CD244+CD150− for most restricted progenitors [43, 44]. The MPPs give rise to common lymphoid progenitors, common myeloid progenitors, and mast cell-committed progenitors, each of which differentiate into specific linages [38, 45] (see Figure I-2).
Hematopoiesis is largely dependent on the intimate contact of HSPCs with the stromal cells in the BM (see Figure I-3). While the committed progenitors stay more in the central regions of the BM, primitive HSPCs are mostly localized to the endosteum
(i.e., endosteal niche) of the trabecular portions of long bones, which are lined primarily by osteoblasts [46]. In particular, the spindle-shaped N-cadherin+CD45– osteoblastic
(SNO) cells were identified to contact HSCs [47]. Osteoblasts have been shown to directly affect hematopoiesis in vivo. Increase in osteoblasts enlarged the population of
HSCs via Notch activation, although the numbers of progenitors were not affected [47-
49], while inversely, ablation of osteoblasts resulted in reduced cell numbers in the BM, including the LSK population [50, 51]. Besides the structural support for HSPCs, osteoblasts secrete a variety of hematopoietic growth factors, such as granulocyte colony- stimulating factor (G-CSF) [52], hepatocyte growth factor [53], angiopoietin [54], osteopontin [55], and CXCL12, also known as stromal cell-derived factor 1 (SDF-1), or pre-B cell growth-stimulating factor (PBGF) [56]. Osteoblasts also express Sca-1, IL-7, and adhesion molecules including VCAM-1, ICAM-1 and β3 integrin (CD61) [57].
Osteoblast depletion leads to defective B lymphopoiesis, which is dependent on VCAM-1, 10 CXCL12, and IL-7 signaling, while the HSPC numbers are less affected [57]. In a mouse arthritis model, which displays osteoporosis, the long-term repopulating potential was not impaired [58]. In addition to the osteoblastic niche, the vascular niche around the sinusoidal endothelial cells in the BM is a second type of niche for HSPCs. Regeneration of sinusoidal endothelial cells was found to be necessary for the engraftment and reconstitution of HSPCs after myeloablation [59]. Intraperitoneal injection of endothelial progenitor cells accelerated hematopoietic recovery in addition to restoring BM sinusoidal vessels in mice after total body irradiation [60]. Similarly, transplantation of highly purified CD31+ microvascular endothelial cells from brain and lung consistently restored hematopoiesis in mice which received bone marrow lethal doses of irradiation for BM [61]. In BM sections, many CD150+CD48−CD41−Lin− HSCs were found localized next to the sinusoidal endothelium [44]. Around the sinusoidal endothelium or near the endosteum, most HSCs were found in contact with a subset of reticular cells that express high levels of CXCL12, called CXCL12 abundant reticular cells (CAR cells), a proposed key player in the HSC niches [62], which also have constitutive expression of VCAM-1 and is also a major resource of SCF secretion in the BM [63]. CAR cells have adipogenic and osteogenic differentiation potential, and short-term ablation of CAR cells in vivo caused a severe reduction in the number of HSCs and lymphoid and erythroid progenitors, presumably due to the lack of production of the cytokines CXCL12 and stem cell factor
(SCF, also known as Kit ligand) by CAR cells [63], both of which are important to HSPC survival as well as migration (discussed in the following section). Especially for the maintenance of LT-HSCs, genetically modified mouse models underscored the critical role of CXCL12 and SCF signaling. Mice with a mutant gene for c-Kit (KitW41/W41), the 11 receptor for SCF, had reduced numbers of quiescent LT-HSCs at steady state [64]. In conditional Cxcl12 homozygous knockout (CXCL12-cKO) mice, the BM and blood exhibited increased LSK HSPCs with reduced quiescent LT-HSCs in the BM [65]. Of note, SCF is decreased in the BM of the CXCL12-cKO mice at both the mRNA and secreted protein levels [65]. A connection between CXCL12 and SCF levels is the metalloprotease-9, which releases soluble SCF from a transmembrane precursor and was found to be upregulated in the BM in response to elevation of CXCL12 in plasma, leading to increased plasma levels of SCF [66]. Other signaling pathways involved in the stem cell-niche interactions that regulate hematopoiesis include bone morphogenic protein, Wnt, Notch, hedgehog, Ca2+-sensing receptor, angiopoietin-1/Tie-2 and c-
Mpl/thrombopoietin [37, 67].
Three colony-stimulating factors (CSFs), granulocyte/macrophage CSF (GM-CSF; also known as CSF2), macrophage CSF (M-CSF; also known as CSF1) and granulocyte
CSF (G-CSF; also known as CSF3) were identified for their abilities to stimulate precursor cells to generate in vitro colonies of mature myeloid cells, as their name indicates [68]. Later, these CSFs were found to also play roles in immune responses, and a link has been identified between the expression and action of CSFs and the proinflammatory cytokines [69]. Constitutive expression M-CSF in the steady state was found in vitro in fibroblasts, endothelial cells, stromal cells, macrophages, smooth muscle cells and osteoblasts, whereas GM-CSF and G-CSF are only upregulated under certain pro-inflammatory conditions [69]. Loss of M-CSF production caused toothlessness, low body weight, severe abnormalities in skeletal structure and deficiency in macrophages in mice, which was considered to be the result of impaired osteoclast formation and
12 subsequently abnormal bone structure [70]. Despite the requirement in development and maturation of macrophages in other tissues, M-CSF was reported to be dispensable for development of Langerhans cells, BM monocytes, and macrophages in lymph nodes and thymus [71]. Administration of M-CSF in mice enhanced the circulating monocyte numbers from 3% to 30% and increased macrophages in the periphery [72]. In contrast, a study on 6 patients with acute leukemia who received cytotoxic chemotherapy showed that M-CSF injection did not have an effect on mobilization of HSPCs [73]. Instead, M-
CSF mobilizes endothelial progenitor cells to sites of vascular injury and accelerates neovascularization, which can be also triggered by G-CSF [74]. As for GM-CSF and G-
CSF, systemic or intraperitoneal injection of GM-CSF into mice augments numbers of both circulating neutrophils and cycling peritoneal macrophages [75], while G-CSF is clinically used to treat or prevent neutropenia [76]. In addition, both GM-CSF and G-CSF are clinical HSPC mobilizers, although G-CSF appears predominant [77]. How hematopoietic cells are released from the BM in response to G-CSF, GM-CSF and other mobilizing stimuli is discussed in the next section.
2. Emigration of Hematopoietic Stem/Progenitor Cells and Leukocytes from the
Bone Marrow to Blood
Recent studies suggest that HSPCs are able to respond directly and immediately to infections and inflammatory signals, migrate to the inflammatory sites, and differentiate into effector cells to provide rapid immunosurveillance [78, 79]. HSPCs also emigrate from the BM to the blood at a low rate during steady state [80]. The mechanism of regulation of hematopoietic cell migration from the BM to the circulation remains elusive,
13 although several signaling axes have been identified. Besides the role of supporting hematopoiesis in the niches, CXCL12 is known for its chemokine property to potently attract HSPCs, pro-B and pre-B cells, megakaryocyte progenitors, T lymphocytes and monocytes [81]. The CXCL12 expressed by niche cells, consisting of osteoblasts, endothelium, and CAR cells, is thought to interact with its primary receptor CXCR4 on the hematopoietic cells to retain the cells in their niches [65]. For example, G-CSF depresses CXCL12 levels in the BM, primarily through induction of osteoblast apoptosis and inhibition of osteoblast differentiation [82-84]. Administration of AMD3100, a specific CXCR4 antagonist, or selective deletion of CXCR4 in the myeloid lineage was found to facilitate mature neutrophil emigration from the BM to the circulation, [85, 86].
In CXCR4−/− bone marrow chimeras of mice, pro- and pre-B cells, and granulocytic cells were more abundant in the blood but less so in the BM, and HSPCs in the blood and spleen were also increased [87, 88]. N-terminal processing of CXCR4 and/or CXCL12 by several proteases from HSPCs and leukocytes, often as a consequence of G-CSF stimulation, was correlated to their mobilization [89-92]. In contrast, CTCE-0214, an analogue peptide of CXCL12 and an agonist of CXCR4, synergize with G-CSF [93] and
AMD3100 [94-97] to mobilize HSPCs and neutrophils. Elevation of CXCL12 levels in the blood by introducing an adenoviral vector expressing CXCL12 also promoted HPSC mobilization [98, 99]. Inversely, injection of CXCL12 and CXCR4 neutralizing antibodies into healthy mice prevented G-CSF-induced mobilization of HSPCs and
CD45+ leukocytes [92]. Similarly, impaired neutrophil mobilization in a cecal ligation and puncture model of sepsis was found in mice pretreated with CXCL12 antisera, and septic mice had significantly decreased expression of CXCL12 in the BM but reciprocal
14 increase in the spleen [100]. RNA in situ hybridization determined that the G-CSF- triggered loss of CXCL12 expression was limited to osteoblasts and not other CXCL12- expressing cells in the BM, including endothelial cells and CAR cells [88]. Osteoblasts undergo apoptosis to attenuate retention factors including CXCL12, SCF and VCAM-1 after G-CSF treatment [101], which primarily activates monocytic G-CSF receptor [102].
Hence, one may hypothesize that CXCL12 expression on osteoblasts, which retains
CXCR4-expressing HSPCs and leukocytes in the endosteal niche, is disrupted during mobilization and the sequential ascending gradient of CXCL12 towards sinusoidal endothelium and surrounding CAR cells guide them to the vasculature [92]. There, a second gradient of CXCL12, as a result of local degradation by protease-mediated cleavage or the plasma level elevation derived from peripheral tissues, might facilitate the transendothelial migration of the hematopoietic cells from the BM to the peripheral blood
[100]. Yet, the pivotal role of the CXCL12-CXCR4 axis in HSPC migration under steady state is challenged by Sasaki et al. [103], who reported that the CXCR4 expression in freshly isolated LSK cells was very low to undetectable.
While SCF seems to play a role in mobilization of HSPCs, research has generated inconsistent conclusions. One study observed chemotaxis and chemokinesis on cultured
HSPCs in respond to SCF [104], whereas other studies suggested that SCF is not chemotactic or chemokinetic by itself, but SCF increased G-CSF-induced mobilization of
HSPCs in vivo [105], and enhanced CXCL12-induced chemotaxis of CD34+ HSPCs via downstream signaling pathways shared with CXCL12 [106]. Recently, an in vivo mouse study showed administration of SCF, similar to G-CSF, increased numbers of circulating
15 and splenic HSPCs, and reduced CXCL12 expression in the osteoblasts and the osteoblast counts in the BM [88].
The retention of leukocytes in the BM also has been liked to the interaction between
α4β1 integrin and VCAM-1. VCAM-1 is constitutively expressed by BM stromal cells like CAR cells [63] and sinusoidal endothelium [107], while in endothelium of other tissues it is only upregulated upon inflammation [108]. The immunostaining of VCAM-1 was found to be greatly reduced in the BM during HPC mobilization in response to G-
CSF and SCF [109]. Two serine proteases, neutrophil elastase and cathepsin G, were found to cleave VCAM-1 and facilitate CD34+ HPC detachment in vitro, which, however, was dispensable for G-CSF-induced HSPC mobilization [109, 110]. In vitro experiments showed that treatment with CXCL12 enhanced adhesion of α4β1 integrin- expressing CD34high BM cells, hematopoietic cell lines and neutrophils to VCAM-1 or
VCAM-1-expressing stromal cells [81, 111]. The interaction between α4β1 integrin and
VCAM-1 retains sinusoidal B cells inside BM [112] while the expression of α4β1 integrin on neutrophils was found to decrease as they mature, and injection of α4 blocking antibody caused efflux of BM neutrophils [111]. However, Burdon et al. [113] reported that the expression of α4 integrin was elevated in neutrophils mobilized by rat CXCL2
(also known as macrophage inflammatory protein-2). Moreover, in transgenic mice that lack VCAM-1 expression selectively in endothelial and hematopoietic cells, peripheral blood showed mild leukocytosis with only a statistically significant increase in the numbers of monocytes and B220+IgDlowIgMhigh immature B cells [114]. Additionally in the BM, the numbers of IgDlowIgMhigh (immature), IgD+IgMhigh and IgD+IgM+ B cells as well as CD8+ T cells were decreased compared to controls, whereas B220+CD43+ pro-B
16 cell levels remained the same [114]. Obviously, more research is required to better understand the impact of VCAM-1 and α4β1 integrin on the mobilization of various hematopoietic cell types in the BM. Of note, administration of anti-α4β1 integrin antibodies induced mobilization of HSPCs, which was not affected by the lack of functional receptors for G-CSF, IL-7 and IL-3α but was dependent on cooperative signaling through SCF and its receptor c-Kit [115]. The α4β1 integrin and other adhesion molecules including αLβ2 integrin, lymphocyte function-associated antigen 3 (LFA-3, also known as CD58), and ICAM-1 are expressed at lower levels in CD34+ cells mobilized in human subjects by GM-CSF, as compared to BM CD34+ cells, suggesting a role for ICAM-1-αLβ2 integrin adhesion interaction in GM-CSF-induced mobilization of
HSPCs [116]. Like VCAM-1, ICAM-1 is expressed in BM stromal cells, and the interaction with αLβ2 integrin supports HSPC attachment to stromal cells [117]. Yet, in contrast to the enhancing effect of anti-α4β1 integrin antibody on HSPC mobilization
[115], antibody blocking of αLβ2 integrin ablated IL-8 induced mobilization of HSPCs in mice [118].
Mobilization of lymphocytes, including B and NK cells through disruption of
CXCL12-CXCR4-mediated retention in the BM has been documented [119-121], and
CXCR4 antagonism also promotes mobilization of T cells that mature in the thymus
[122]. The chemoattractant sphingosine-1-phosphate (S1P) has emerged as a central mediator of lymphocyte trafficking especially for T and NK cells [123]. In the case of
NK cells, a deficiency in the S1P receptor S1PR5 leads to their accumulation in the BM and lymph node [124, 125]. While the CXCR antagonist AMD3100 mobilizes all types of NK cells, the CCR1/5 agonist CCL3 (also known as macrophage inflammatory protein
17 1α or MIP1α) selectively mobilizes mature NK cells but not immature or precursor NK cells, probably through a decreased presence of CXCR4 on mature NK cells implied by impaired chemotaxis to CXCL12, which is not observed in CCL3-treated immature NK cells [120]. Neutrophil release from the BM is believed to be regulated primarily through the CXCL12-CXCR4 axis, and secondarily through the interaction of CXCR2 with its ligands, the ELR+ chemokines CXCL1 and CXCL2 that are constitutively expressed in
BM endothelial cells and osteoblast [126]. CXCR2-null neutrophils preferentially retained in the BM and are not mobilized by transient disruption of CXCR4, whereas neutrophils lacking both CXCR2 and CXCR4 showed neutrophilia similar to CXCR4- deficient neutrophils [126]. In contrast, the most important chemokine signaling pathway for the mobilization of classical monocytes (Ly6Chigh) to inflammatory sites is CCR2 and its ligands CCL2 and CCL7 (also known as monocyte chemoattractant protein-1 and -3, or MCP-1 and -3, respectively). In contrast, nonclassical monocytes (Ly6Clow) regularly patrolling along the luminal surface of endothelial cells of microvessels do not highly express CCR2 like classical monocytes and their emigration mechanism remains elusive
[127, 128]. CCR2 –/– mice display severe loss of monocytes in blood and spleen while monocytes are increasingly retained in the BM, which was also observed in mice lacking
CCL2 or CCL7 [129, 130]. LPS, a TLR4 ligand, induced rapid CCL2 expression in BM mesenchymal stem cells and their progeny including CAR cells in proximity to BM vascular sinuses leading to monocytosis [131]. Loss of CCL2 expression impairs mobilization of monocytes/macrophages out of the BM [131, 132]. In addition, CCR2 ligands (predominantly CCL2) were found to be upregulated in the BM and blood in response to murine cytomegalovirus infection in mice, and were produced mainly by
18 F4/80+ cells in the BM [132]. Nonetheless, the CXCL12-CXCR4 axis is critical in monocyte mobilization, as demonstrated by inhibition of CCR2 antagonist-induced monocytopenia by the CXCR antagonist AMD3100 [133]. Of note, CCR2 was found to be expressed on subsets of HSPCs and mediated their recruitment to inflammatory sites in mice through the chemotaxis to CCL2 and CCL7 [134]. In addition, CCR1 mediates mobilization of myeloid progenitors from the BM triggered by CCL3 [135].
3. Homing of Hematopoietic Stem/Progenitor Cells and Leukocytes to the Bone
Marrow
Homing of HSPCs and leukocytes from the circulation to the BM is also critical to their homeostasis while the process is asymmetric to the egress from the BM to the bloodstream. In contrast, HSPC transendothelial migration during homing to the BM employs adhesion molecules such as VCAM-1, ICAM-1 and VE-cadherin, similarly to leukocyte extravasation at inflammatory sites in peripheral tissues [136]. CXCL12-
CXCR4 binding mediates HSPC adhesion and transendothelial migration during homing of HSPCs to the BM, which is dependent on α4β1 and α5β1 integrins [137]. Using
CXCL12-cKO mice, impaired homing of HSPCs was observed in CXCL12-cKO stroma culture and CXCL12-cKO recipients in a BM cell transplantation experiment [65]. Long- term plasma cells (antibody-secreting memory B cells) also require CXCR4 expression to home to the BM [138]. They exhibit high expression of adhesion molecules including
+ α4β1 and αLβ2 integrins, and greater adhesion to VCAM-1 than IgM cells [139]. Selectins may also contribute to this process in the initial steps [139-141], which is controversial
19 based on a study with selectin-deficient mice showing unimpaired mobilization of HSPCs and leukocytes in response to sulfated polysaccharides [141].
Circulating senescent neutrophils need to be cleared in the liver and spleen as well as bone marrow, where the CXCL12-CXCR4 interaction is also important for homing of circulating neutrophils before these cells undergo apoptosis and are phagocytosed by macrophages in the BM [142, 143]. Progressive reduction of in CXCR2 expression and increases in CXCR4 expression were observed as the neutrophils become senescent [86,
144]. Sequestration of intravenously injected CXCR4high neutrophils in the femoral BM was more efficient than for CXCR4low neutrophils [86, 142]. Moreover, the CXCR4 intracellular signaling pathway seems essential for the neutrophil homing to the BM, which was suppressed by pretreatment with pertussis toxin, a Gi protein-coupled receptor inhibitor that blocks CXCR4 intracellular signaling [145]. Interestingly, the BM macrophages that had phagocytosed apoptotic neutrophils had a substantial increase of
G-CSF production, which is likely a homeostatic link between the clearance and the production/release of neutrophils [145].
Homing of Ly6Chigh classical monocytes to BM in the absence of inflammation has been demonstrated, and unlike senescent neutrophils returning to the BM which become apoptotic and are cleared by macrophages, returning classical monocytes are speculated to differentiate into nonclassical monocytes that can reenter the circulation [146]. The mechanism of monocyte homing to the BM is not clear but, in contrast to the crucial role in classical monocyte emigration from the BM to peripheral blood, CCR2 is not required for monocyte homing to the BM or inflamed peripheral tissues [130, 147]. Since the expression of CXCR4 is not detectable in classical monocytes in the blood of naïve mice,
20 the CXCL12-CXCR4 signal is unlikely to be employed in the circulating monocyte homing to the BM [133].
D. Alzheimer’s Disease
Alzheimer disease (AD) is the most prevalent form of dementia characterized by progressive cognitive decline [148]. The neuropathological lesions manifested in AD include senile amyloid plaques and neurofibrillary tangles, which are accompanied by reactive microgliosis, dystrophic neurites and loss of neurons and synapses [149]. A consensual hypothesis for the pathogenesis of AD is the amyloid cascade hypothesis that deposition of amyloid-β (Aβ) peptides, the products of the cleavage of amyloid precursor protein (APP) by β- and γ-secretases [150, 151] and the main components of senile plaques, trigger neuronal dysfunction and death in the brain associated with disordered intracellular accumulation of the microtubule-associated τ protein, which gives rise to neurofibrillary tangles [152, 153]. Although some hypotheses have been proposed, the question of how Aβ affects τ protein in neurons is still unresolved [148]. Research in the past two decades in animal models of AD has revealed a crucial role for neuroinflammation in AD pathology, and efforts have been made to target inflammation regulators for clinical benefit. For example, conventional nonsteroidal anti-inflammatory drugs have been shown to be preventive and therapeutic in AD [154].
Microglia, the residential immune cells of the myeloid lineage in the central nervous system (CNS), have been reported to produce nitric oxide (NO), reactive oxygen species
(ROS), proinflammatory cytokines, e.g., tumor necrosis factor (TNF)-α, IL-1β, IL-6 and
IL-18, and prostaglandins (e.g., PGE2) in response to Aβ, a process referred to as
21 classical M1 inflammatory activation that contributes to neuronal death [155]. However, microglia can alternatively undergo M2 activation characterized by expression of anti- inflammatory cytokines that promote tissue repair, which was observed in AD patient subjects and an AD mouse model [156]. It has been experimentally demonstrated that microglia-mediated clearance of soluble Aβ is via macropinocytosis, both in vitro and in vivo [157]. The clearance of insoluble fibrillar Aβ (fAβ) and fAβ-containing plaques by microglia is carried out by phagocytosis, during which fAβ interacts with a cell surface receptor complex formed by the B-class scavenger receptor CD36, the α6β1 integrin and
CD47/integrin associated protein (IAP) [158, 159], followed by fAβ phagocytosis and degradation [160]. The importance of microglia in reducing Aβ levels has been confirmed not only in AD mouse models, but also in Aβ-immunized AD patients who showed decreased plaque load and increased microglial cell accumulation [161-163].
Furthermore, studies have also shown that microglia as well as other cells involved in the
CNS immune response, such as dendritic cells and astrocytes, are able to detect Aβ through TLR signaling [164].
Recently findings support roles in AD of immune cells other than resident microglia in the brain. Blood-borne monocytes have been shown to infiltrate the brain in AD mouse models, and restrict plaque burden like microglial cells [165-167]. Fiala and colleges found that Aβ1-42-treated human peripheral monocytes secreted proinflammatory cytokines, including TNF-α, IL-1β, and IL-12, as well as chemokines CCL2, CCL3,
CCL4, and CXCL8 (IL-8), which was speculated to help monocytes migrate across the blood-brain barrier (BBB) based on data with an in vitro model comprised of brain endothelial and astroglial cells [168]. Macrophages from AD patients appear apoptotic 22 and bind but fail to phagocytose Aβ compared to cells from age-matched controls [169,
170]. AD brain sections were shown to have oligomeric Aβ in neurons and apoptotic macrophages, which accumulated around microvessels with fibrillar Aβ and contribute to cerebral amyloid angiopathy (CAA) [170]. Loss of CCR2 or treatment of CCL2 did not change phagocytosis of Aβ by adult microglia and bone marrow-derived macrophages; however, CCR2 deficiency in AD mice caused significant reductions in numbers of perivascular macrophages (PVMs) around the Aβ+ vessels, Aβ phagocytosis by microglia and BM-derived macrophages and survival rate, and exacerbated vascular Aβ deposition in spite of normal parenchymal Aβ deposition in the brain [171]. Therefore, CCR2- dependent recruitment of PVMs to Aβ+ vessels seems crucial for Aβ clearance. Yet, while some believe that macrophages emigrate to the blood to export Aβ out of the brain, others have questioned this speculation since Aβ-engorged macrophages, either from AD patients or control subjects, adhered to the endothelium and accumulated in the perivascular regions due to inhibited emigration [170]. This controversy was addressed by a recent study using live intravital two-photon microscopy, which showed that circulating Ly6Clow monocytes were attracted to and crawled selectively onto Aβ+ veins, where they engulfed Aβ and returned it back to the bloodstream, thus contributing to the clearance of Aβ deposition from the brain [172].
Furthermore, Shad et al. reported that AD patients, regardless of their age or sex, had significantly higher blood counts of monocytes and/or neutrophils compared to healthy controls [173], whereas Jaremo et al. showed that circulating granulocytes
(neutrophils and eosinophils) in the blood of AD patients in the light fractions were less than in control subjects, which was hypothesized to be a sign of impaired granulocyte
23 turnover and cell damage [174]. A third study, which involved the largest sampling sizes, detected an increase in the blood neutrophil-lymphocyte ratio (NLR) in AD patients compared to patients with normal cognitive function [175]. Such elevated NLR has been suggested to be a marker of systemic inflammation in many diseases/conditions, including cancer [176], hypertension [177], diabetes mellitus [177], coronary artery bypass grafting [178], familial Mediterranean fever [179], ankylosing spondylitis [180], placental inflammatory response [181], schizophrenia [182] and cerebral infarction [183].
An interesting finding about peripheral blood neutrophils in AD patients is that their engulfment of microbes remained the same as normal neutrophils, while their digestive activity decreased at the early stage of AD, and the percentage of phagocytizing neutrophils decreased at the late stage [184]. Although the effects of changes in neutrophils in AD is still enigmatic, two-photon in vivo imaging demonstrated that Gr-1
(Ly6G/C) antibody-labeled neutrophils transmigrated into the brain parenchyma and towards amyloid plaques in 5XFAD mice (an AD model), a phenomenon not observed in wild type mice [185]. Yet, the chemotactic activity of neutrophils in 5XFAD mice was likely indirectly induced by Aβ, probably via release of chemokines from resident brain cells [185]. It has been reported that microglia are able to directly engulf invading both apoptotic and, surprisingly, viable and motile neutrophils [186]. Hence, it is possible that activated neutrophils engulf Aβ and then are cleared by microglia. In a typical acute inflammatory response, neutrophils accumulate at the inflammatory site first and help recruit monocytes [187] and, therefore, a similar pattern may occur in the AD brain.
Neutrophils are usually considered to be detrimental to cells since they can secrete proteases, as well as oxidative and tissue-degrading enzymes at the inflammatory sites,
24 although a beneficial role for neutrophils in proper tissue regeneration has emerged [188].
For example, antibody-depletion of Ly6G/Gr-1+ neutrophils in mice subjected to spinal cord lesion unexpectedly eliminated ROS [189], but inhibited evolution of astrocyte reactivity (a wound healing response), altered cytokine and chemokine production levels, and worsened the neurological outcome [190]. Neutrophil-depleted mice had significantly higher levels of G-CSF, CCL2, CCL9 (also known as macrophage inflammatory protein 1-γ) and CXCL1 (also known as keratinocyte-derived chemokine) as a compensatory attempt to recruit more inflammatory cells [190]. Moreover, circulating HSPCs have also emerged as a factor in AD pathology. Patients with early
+ AD have reduced numbers of circulating CD34 HSCs, which correlates with age, Aβ1-42 levels in the cerebrospinal fluid, and the ratio of Aβ1-42/Aβ1-40 [191].
In addition to a neuroinflammatory phenotype, AD is often preceded by malfunctions in the cerebrovascular system, including CAA and decreased cerebral blood flow, which lead to hypoxia, breakdown of the BBB and ultimately to brain atrophy and death [192-
196]. Studies in mouse models and human subjects of AD indicate that Aβ peptides cause forceful constriction of cerebral blood vessels [197, 198], suggesting that vascular factors play an early pathogenic role in AD.
E. Purinergic Signaling in Inflammation
1. Nucleotides and Purinergic Receptors
In addition to their roles in intracellular metabolism, nucleotides released into the extracellular space can activate P2 purinergic receptors on the cell surface and evoke many important signaling events. Adenosine 5’-triphosphate (ATP) was identified as a
25 neurotransmitter in 1970 [199] and, therefore, the receptors and signaling pathways of extracellular nucleotides were termed “purinergic”. In contrast to intracellular concentrations of ATP in the millimolar range [200] and an ATP concentration within nerve vesicles that reaches 1M [201], extracellular concentrations of ATP are maintained at much lower levels, for example, at submicromolar level in human plasma [200]. Other than vesicular secretion from neuronal cells as a transmitter [199, 202, 203], ATP and its metabolites can be released from non-excitable cells such as activated/stimulated platelets
[204, 205], leukocytes [200, 206], red blood cells (RBCs) [207, 208], vascular endothelial and smooth muscle cells [209, 210], bone stromal cells [211] and cancer cells [212, 213].
Obviously, apoptotic or damaged cells also release large amounts of nucleotides that were compartmentalized within the cells. Release of uridine 5’-triphosphate (UTP) without cell lysis has been less studied than ATP, but UTP release was observed in shear force-stressed bovine endothelial cells, leukocytes, airway epithelial cells, astrocytes and several cell lines, as well as activated human platelets [214, 215], thereby elevating the extracellular UTP concentrations up to 20-fold above the baseline nanomolar range [214].
On the other hand, purinergic receptors are widely expressed in excitable and nonexcitable cells of vertebrates. Seven ATP-gated P2X receptors (P2X1-7 receptors), and eight P2Y receptors with varying subtype specificity for ATP, adenosine 5’- diphosphate (ADP), UTP, uridine 5’-diphosphate (UDP) and UDP-glucose (P2Y1,2,4,6,11-14 receptors) as well as four P1 adenosine receptors (A1, A2A, A2B, and A3 receptors) have been identified to date, and several ectonucleotidases are known to catalyze hydrolysis of extracellular ATP to ADP, and UTP to UDP, as well as their further breakdown [216].
P2Y receptors (P2YRs) and P1 receptors (P1Rs) are GPCRs (also known as seven-
26 transmembrane domain receptors), whereas P2X receptors (P2XRs) are ligand-gated non- selective cation channels that contain two transmembrane domains, a large extracellular loop and intracellular N- and C-termini [217], and were found to form stable homo- or hetero-trimers [218-221]. Adenosine is generally considered to be involved in immunosuppression (mainly acting at A2A receptors) and cell growth regulation (mainly through A3 receptors), while ATP has more complex signaling leading to proinflammatory, immunosuppressive, and growth-promoting effects [212].
Among the metabolic P2YRs (see Table I-1 for summary of agonists, signaling pathways and function), the P2Y2 receptor (P2Y2R) subtype responds to ATP and UTP with equipotency [222]. The P2Y2R contains an extracellular Arg-Gly-Asp (RGD) domain that binds the fibronectin-binding integrins, αvβ3 and αvβ5 [223], thereby enabling the P2Y2R to activate small Rho GTPases, Rac1 and RhoA [224, 225]. The P2Y2R also has SH3-binding motifs in its intracellular domain for interaction with Src and promotes the Src-dependent activation of several growth factor receptors, including VEGF receptor
2 (VEGFR-2) [226], epithelial growth factor receptor (EGFR), and platelet-derived growth factor receptor (PDGFR) [227]. The P2Y2R is able to activate two members of the ADAM (a disintegrin and metalloproteinase) family, ADAM10 and ADAM17 (also known as TNF-α converting enzyme or TACE) [228-230], which shed the membrane- bound precursor and release the active form of TNF-α [231], as well as other cytokines, chemokines, growth factors, and their receptors [232]. The P2Y2R is one of the most widely expressed purinergic receptors in different cell types and tissues [216, 233] and has dominant roles in purinergic signaling under many inflammatory conditions, as discussed below.
27 2. Purinergic Signaling in Inflammation
Purinergic signaling is involved in the regulation of functions in all types of immune cells [200]. Both autocrine and paracrine purinergic signaling is suggested to be crucial for immune cell responses to inflammatory stimulations. An example of an autocrine mechanism is T cell receptor stimulation at the immune synapse between T cells and antigen-presenting cells that rapidly releases ATP, which evokes calcium influx, activation of nuclear factor of activated T cells (NFAT), and upregulation of IL-2 and the
P2X7 receptor (P2X7R) in T cells [234, 235], followed by additional ATP release through pannexin hemichannels that open in response to elevated cytosolic calcium levels
[235]. Interestingly, ATP induces translocation of pannexin-1 hemichannels as well as
P2X1 and P2X4 receptors (P2X1R and P2X4R, respectively) but not P2X7R to the immune synapse, and P2X1R and P2X4R but not P2X7R contribute to NFAT activation and IL-2 expression, while P2X7R, P2X1R, and P2X4R all contribute to calcium entry
[236]. Purinergic signaling through P2XRs in cooperation with pannexin-1 is also crucial for the activation of inflammasomes and the consequential release of the pro- inflammatory cytokine IL-1β [206]. At the leading edge of migrating neutrophils, ATP is released and activates the P2Y2R to amplify chemotactic signals and orientates the cells towards the bacterial chemoattractant formyl-Met-Leu-Phe in vitro [237]. Autocrine purinergic signaling through the P2Y2R is also known to mediate chemokinesis of macrophages and their chemotaxis towards the chemoattractant Complement C5a [238,
239]. Besides the autocrine feedback loop, ATP released by stressed cells has been suggested to be a “find-me” signal to guide phagocytes to the inflammatory sites to promote cell clearance by the phagocytes [240].
28 −/− While under standard diet the P2Y2R knockout (P2Y2R ) mice do not exhibit significantly different body weight, plasma osmolality and some other blood parameters
−/− but increased arterial blood pressure comparing to wild type mice [241], the P2Y2R mice have impaired migration of neutrophils to sites of Staphylococcus bacterial infection
[237] and to smoke-injured lungs [242], migration of eosinophils and monocyte-derived
DCs to allergen-infected lungs [243], and migration of monocytes and macrophages to apoptotic cells in an air-pouch model [240]. The accumulating in vivo evidence has manifested the importance of the P2Y2R in chemotaxis and infiltration of leukocytes under inflammatory conditions. As mentioned earlier, leukocyte recruitment requires contributions from both leukocytes and endothelial cells. While leukocytes have high levels of P2Y2R expression, the P2Y2R, but not P2Y4 receptor (P2Y4R), is upregulated in stressed or injured vascular endothelium and smooth muscle [244]. In collared rabbit carotid arteries, an in vivo model of vascular stress, perivascular infusion of ATP or UTP
(a selective agonist for P2Y2R and P2Y4R), but not UDP (an agonist for P2Y6R) induced intimal hyperplasia and promoted monocyte/macrophage infiltration into intima, which are responses characteristic of the development of atherosclerotic plaques [244]. Such findings highlighted the regulatory role of the P2Y2R in endothelial cells and smooth muscle cells during leukocyte recruitment. In endothelial cells, the UTP-stimulated
P2Y2R transactivated VEGFR-2 in a Src-dependent manner, which upregulates the expression of VCAM-1, facilitating the binding of leukocytes to endothelial cells [226,
227]. Another pivotal adhesion molecule for leukocyte-endothelium interaction, ICAM-1 was upregulated in UTP-stimulated smooth muscle cells in consequence of P2Y2R- mediated release of lymphotoxin-α (LTA) [245]. LTA also has been shown to also induce
29 ICAM-1 expression in endothelial cells [246], therefore, the endothelial P2Y2R might contribute to leukocyte attachment to endothelium before TEM by upregulating ICAM-1 in addition to VCAM-1.
In hematopoiesis that gives rise to leukocytes and other blood cells, purinergic signaling also plays a role. In the colony-forming unit (CFU) assay, ATP and, to a higher extent, UTP potently enhanced the clonogenic growth of human CD34+ HSCs that were confirmed to functionally express all seven known subtypes of P2XRs and the P2Y1R and P2Y2R (but not P2Y4R, P2Y6R, or P2Y11R) [247]. UTP also synergizes with cytokines to induce colony-forming activities of human CD34+ HSCs and Lin−CD34−
HPCs in vitro [247]. In another study, BM suspensions treated with 1mM ATP for 4 h had a decreased percentage of HSCs, common myeloid progenitors and granulocyte- macrophage progenitors but not megakaryocytic-erythroid progenitors, and in vivo administration of ATP caused increases in the mature myeloid cell population (i.e., granulocytes and monocytes) at the expense of the granulocyte–macrophage progenitor population, suggesting that ATP boosts cell differentiation [248]. The ATP-induced acceleration of proliferation and differentiation towards the myeloid lineage was prevented by the cytokines GM-CSF, SCF and IL-3, as well as by suramin, a P2 receptor antagonist, showing the involvement of P2 receptors [248]. For the traffic of HSCs, UTP was found to significantly enhance CXCL12-stimulated human CD34+ HSC chemotaxis in vitro, inhibit downregulation of CXCR4 on the cell surface, and increase cell adhesion to fibronectin [249]. UTP-pretreated human CD34+ cells homed to the BM of immunodeficient mice more efficiently than untreated cells [249]. The UTP-induced
+ motility of CD34 cells likely occurs though activation of P2Y2R, since the other UTP-
30 + activating receptors, P2Y4R and P2Y6R are not expressed in human CD34 HSCs [247].
Moreover, UTP promoted CXCL12-dependent HSC migration by activating Rac2 and the downstream effectors Rho GTPase-activated kinases 1 and 2 (ROCK1/2) [249].
Consistently, ATP synergizes with CXCL12 to induce chemotaxis of BM nucleated cells in vitro [250]. Among these cells, HSPCs respond well to CXCL12 or CXCL12 + ATP, although ATP alone did not trigger significant chemotaxis [250]. In contrast, the migration of CD14+ cells (mainly monocytes) or CXCR4+ cells was not enhanced in the presence of CXCL12 and/or ATP gradient compared to their spontaneous basal migration, albeit the basal migration of HSPCs was lower than CD14+ or CXCR4+ cells [250]. These findings indicate a role for purinergic signaling in the directed migration of HSPCs.
As for the niches supporting hematopoiesis and involved in release of hematopoietic cell from the BM, bone homeostasis relies on the balance between formation by osteoblasts and resorption by osteoclasts. Osteoblasts are under the influence of mechanical stimulation, and in vitro studies showed that human BM stromal cells released ATP through vesicular exocytosis in response to oscillatory fluid flow, increasing the mean extracellular ATP concentration from 0.3 nM to 6.8 nM [211]. The
P2Y2R was found to sense fluid sheer stress and mediate the formation of actin stress fiber via RhoA activation in a mouse osteoblast cell line, where ATP- and UTP-induced formation of actin stress fibers maximized at 100 µM comparable to concentrations
2 generated by fluid shear stress at a velocity of 12 dyn/cm [251]. The P2Y1 receptor
(P2Y1R) responds to ~0.1 to 10 µM ADP or ATP to stimulate both the formation and resorption of murine osteoclasts [252, 253], while increased expression of the UDP- responsive P2Y6R correlates with the differentiation of osteoclasts to mediate bone
31 resorption [254]. The requirement of purinergic signaling to maintain bone homeostasis was also implied by abnormalities in bones of knockout (P2X7R, P2Y1R, P2Y2R, P2Y6R, and P2Y13 receptor) mouse models although conflicting results were observed in two
−/− distinct P2X7R knockout mouse models [255]. A recent study showed that P2Y2R mice exhibit a significant reduction in bone formation [256]. In vitro experiments using
−/− primary osteoblasts from P2Y2R mice further confirmed the important role of the
P2Y2R in bone construction through its positive correlation to activation of MAP kinases induced by fluid flow or ATP (5 and 20 µM) [256]. These findings appear contradictory to an earlier report that 10-100 µM ATP or 1-100 µM UTP inhibited bone module formation by rat osteoblasts in vitro [257]. Considering that the P2Y4R expression was undetectable in osteoblasts, ATP/UTP-induced inhibition of bone formation was attributed to the activation of the P2Y2R [257]. This conclusion was challenged by a following study from the same group, in which male P2Y2R−/− mice did show increased limb bone mineral content compared to wild type mice [258]. However, osteoblasts express ectonucleotide phosphodiesterase/pyrophosphatase-1 (ENPP-1), an ectonucleotidase that hydrolyzes nucleotide triphosphates to generate pyrophosphate
(PPi), which, in turn, is further hydrolyzed by alkaline phosphatase to produce orthophosphate (Pi) ions; PPi potently inhibits mineralization through antagonism of Pi- dependent mineralization [258]. The application of ATP or UTP to osteoblasts inhibited the expression and activity of alkaline phosphatase, whereas the inhibition of mineralization was also achieved without affecting alkaline phosphatase activity by addition of guanosine 5’-triphosphate (GTP) or cytosine 5’-triphosphate (CTP), which are not P2 receptor agonists, but can be hydrolyze to produce PPi like ATP and UTP
32 [258]. Despite the complexity of the mechanisms involved, the importance of purinergic signaling in bone homeostasis is supported by many other studies, including the
® demonstration that the selective antagonist of P2Y12 receptor Clopidogrel (Plavix ) can inhibit osteoblast proliferation, differentiation, function and survival [259], although further studies are required to resolve the relation between purinergic signaling and phosphate metabolism in the bone.
3. Purinergic Signaling in Alzheimer’s Disease
Purinergic signaling through activation of P1 and P2 receptors is implicated in neuroinflammation, vascular regulation and the progression of many neurodegenerative diseases, including AD [260-265] (see Figure I-4 for P2Y receptor-mediated signaling pathways in AD). Aβ-induced ATP release has been observed in rat primary cortical astrocytes [266], mouse primary microglial cells [267] and microglial N13 cells [268].
Aβ-induced release of ATP from microglial cells, as well as microglial cell activation, are dependent on the expression of the ionotropic P2X7R, which is thought to release ATP through the formation of a membrane pore [268]. In addition, Aβ treatment increases expression of the P2Y2R in microglia [267] and neurons [228, 269], which is likely triggered by P2X7R-mediated IL-1β release [268, 270-276]. In the vasculature, hypoxia causes the release of ATP from erythrocytes into the blood stream [207], and a 24 h exposure of human erythrocytes to Aβ inhibited ATP release from deoxygenated cells
[277]. P2Y receptors expressed in human vascular endothelium, including P2Y1,2,4,6 receptors, trigger vasodilation through release of NO, endothelium-derived hyperpolarizing factor (EDHF) and prostacyclin (PGI2) [264, 278, 279]; thus activation
33 of these receptors may help to counteract the vasoconstrictive effects of Aβ, relieve the hypoxia in the AD brain and ensure vascular perfusion [198, 280].
ATP and UTP have been shown to increase the MAPK-dependent production and secretion of APP in primary rat cortical astrocytes [281], presumably through activation of the P2Y2R and P2Y4R. However, P2Y2R activation in primary rat cortical neurons and human astrocytoma cells activates two members of the ADAM (a disintegrin and metalloproteinase) family, ADAM10 and ADAM17, to enhance α-secretase-mediated
APP processing and produce a non-amyloidogenic fragment [228-230], thereby inhibiting the β-secretase pathway and decreasing the generation of neurotoxic Aβ1-40 and Aβ1-42 through a competitive mechanism [282-285].
As mentioned before, activated microglia are the primary immune effector cells that are recruited to clear Aβ. Although direct evidence is still lacking, P2Y6R and P2Y12R may contribute to the pathophysiology of AD by modulating microglial responses.
P2Y12R was identified to mediate ADP-induced activation and chemokinesis of microglia both in vitro and in vivo [286-288], while UDP-activating P2Y6R has been thought to mediate phagocytosis and chemotaxis/migration in microglia and monocyte infiltration into the injured brain by inducing expression of the chemokines CCL2 (MCP-1) and
CCL3 (MIP-1α) [289, 290]. Although the P2Y2/4R agonist UTP was reported to be insufficient to induce ruffle formation in rat primary microglia [287, 291], it enhanced migration of mouse primary microglia in vitro [267]. Soluble Aβ clearance by pinocytosis (uptake of liquids) in primary rat microglia stimulated with UTP, ATP or
ATPγS, a non-hydrolyzable ATP homologue, was found to occur through activation of
P2Y4R [288]. In contrast, the phagocytosis and degradation of insoluble fibrillar and
34 oligomeric Aβ1–42 aggregates by primary microglia stimulated with UTP or ATP is mediated by the P2Y2R [267]. In the TgCRND8 mouse model of AD, hippocampal expression of CD11b, a marker for activated microglia/macrophage, was reduced in mice
+ +/− with heterozygous knockout of the P2Y2R (TgCRND8 P2Y2R ) [292]. These
+ +/− TgCRND8 P2Y2R mice had accelerated AD pathogenesis, including significantly increased Aβ deposition and neurological deficits when compared to age-matched
+ +/+ + +/− TgCRND8 P2Y2R mice [292]. All TgCRND8 P2Y2R mice exhibited premature
+ +/+ death (by 10-12 weeks) in contrast to a 100% survival rate in TgCRND8 P2Y2R mice
+ −/− at 90 days, whereas TgCRND8 P2Y2R mice had an even shorter lifespan (mean = 25.5 days) [292]. It has been shown that neuropathology in AD (e.g., synapse loss) correlated with a decrease in P2Y2R expression in the parietal cortex of human postmortem AD brain samples as compared to controls [293], which is similar to the enhanced
+ +/− neuroinflammatory and neurodegenerative phenotype seen in the TgCRND8 P2Y2R mice [292]. The neuronal P2Y2R may also contribute to the prevention of these deficits by promoting neurite extension [269, 294]. Interestingly, while previous studies with post-mortem human AD brain tissue show decreased expression of P2Y2Rs as compared to non-AD controls [295], TgCRND8 mice initially showed P2Y2R upregulation in the brain during early stage disease pathology (10-25 weeks of age), however, the expression of the P2Y2R decreased during late stage with disease progression (by 25-48 weeks of age) [292]. Thus, loss of protective functions regulated by the P2Y2R in brain neurons and glial cells in humans and mice appears to enhance the progression of the AD phenotype. In other words, activation of P2Y2Rs during the early stage of AD may serve to delay neurodegeneration caused by inflammation in the AD brain.
35 Figure I-1. Leukocyte chemotaxis, recruitment, and transendothelial migration.
36 Blood)Flow) Erythrocyte)
Leukocyte)
Rolling) Tethering)) Adhesion)) Transendothelial)) FlaEening) migra9on)
Interendothelial)) Selec9ns) VCAM/ICAM) Chemotac9c)gradient) junc9ons)) Endothelial)cell) Basement) membrane) Inflammatory)s9muli)
37 Figure I-2. Hematopoiesis. LT-HSC, long-term hematopoietic stem cell; ST-HSC, short-term hematopoietic stem cell; MPP, multipotent progenitor; LMPP, lymphoid- primed; CLP, common lymphoid progenitor; MCP, mast cell-committed progenitor;
CMP, common myeloid progenitor; MEP, megakaryocyte–erythrocyte progenitor; GMP, granulocyte–macrophage progenitor; NK, natural killer cell; EP, erythrocyte progenitor;
MkP, megakaryocyte progenitor; MDP, monocyte–dendritic cell progenitor; GP, granulocyte progenitor; CDP, common dendritic progenitor; DC, dendritic cell.
38 Primi%ve(stem/progenitor(cells( Commi3ed(( Lineage(Commi3ed(cells( progenitors( B'Lymphocyte' Pro'B' T'Lymphocyte' Pro'T' CLP' NK'cell' Pro'NK' Mast'cell' LMPP' '' Erythrocyte' EP' LT#HSC'' ST#HSC'' MPP' MCP' Platelets' MEP' MkP' Megakaryocyte' CMP' '' CDP' DC' GMP' MDP'
Monocyte' GP' Neutrophil' Macrophage' Basophil' Eosinophil'
39 Figure I-3. HSC mobilization from and homing to the bone marrow. HSC, hematopoietic stem cell; SNO cell, Spindle-shaped N-cadherin+CD45− osteoblastic cell;
CAR cell, CXCL12-abundant reticular cell. Small shapes represent CXCL12, SCF, and
VCAM-1 as indicated, and their binding partners.
40 Bone(
HSC$ SNO$cell$ Osteoblast$ Emigra9ng$HSC$ Progenitor$ Leukocyte$
Homing$HSC$ Bone( Endothelial$ cell$
SCF$ Vascular( Emigra9ng$$ (sinus( leukocyte$ CXCL12$
VCAMA1$
Vascular( (sinus( CAR$cell$ Chemoa:ractant$
41 Table I-1. P2Y receptor agonist, signaling pathways, and functions. Modified from
Erb and Weisman [296]. AC, adenylate cyclase; HDL, high-density lipoprotein.
42 Receptor Agonist G protein−Main Function Effector(s)
P2Y1R ADP > ATP Gq−PLCβ, Rac, Rho Platelet shape change/aggregation activation Bone resorption Leptin secretion from adipocytes Mechanical and thermal nociception Atherosclerosis Angiogenesis Synaptic plasticity
P2Y2R ATP = UTP Gq−PLCβ activation Bone formation
Go−PLCβ, Rac Immune cell recruitment and phagocytosis activation Vascular tone, inflammation, thrombosis
G12−Rho activation Blood pressure regulation Mechanical and thermal nociception HIV infection of T cells Epithelial K+/Cl− secretion Pancreatic and renal functions Liver regeneration Wound healing
P2Y4R UTP (human) Gq−PLCβ activation Visual and auditory transmission + − UTP ≥ ATP (rodent) Go−PLCβ activation Intestinal K /Cl secretion
P2Y6R UDP Gq−PLCβ activation Bone resorption
G12/13−Rho activation Vascular tone and inflammation Cardiac fibrosis Macrophage cytokine/chemokine release Microglial phagocytosis Epithelial Cl− secretion
P2Y11R ATP Gq−PLCβ activation Inhibition of neutrophil apoptosis
Gs−AC activation Negative regulation of TLR signaling Pancreatic Cl− secretion Cell cycle arrest in endothelium
P2Y12R ADP Gi/o−AC inhibition Platelet aggregation PLCβ, RhoA Microglial activation/migration activation Dendritic cell activation/macropinocytosis Peripheral anti-nociception, central nociception
P2Y13R ADP Gi/o−AC inhibition Bone formation PLCβ, RhoA Liver uptake of HDL activation Inhibition of ATP release from RBCs Peripheral anti-nociception
P2Y14R UDP-glucose Gi/o−AC inhibition Immune function/IL-8 release in epithelium PLCβ activation Gastric function/stomach contractility Peripheral anti-nociception 43 Figure I-4. P2Y receptor function in Alzheimer’s disease. (A) Overview of cell types involved in neuroinflammation and the neurovascular unit. Areas b–e in panel (A) are magnified in panels (B–E). Although intimately involved, neuron-associated astrocytes and oligodendrocytes and microvessel-associated pericytes are not shown for simplicity.
Aβ alters cellular release of nucleotides, including increased ATP release from microglia and decreased ATP release from hypoxic erythrocytes. In perivascular neurons, ATP is released from synaptic vesicles and causes vasoconstriction by activating P2X1 and
P2Y2,4,6 receptors on vascular smooth muscle cells. P2Y1,2,4,6 receptors in vascular endothelial cells promote vasodilation by responding to ATP in the blood stream. The endothelial P2Y2R facilitates monocyte adhesion to the vascular wall, through VEGFR-2- dependent upregulation of vascular cell adhesion molecule-1 (VCAM-1) as well as monocyte extravasation. Activation of neuronal P2Y1,2 receptors promotes neurite extension and stabilization, whereas P2Y13R activation inhibits neurite extension.
Neuronal P2Y2R facilitates non-amyloidogenic processing of APP through ADAM10/17- dependent production of soluble APPα (sAPPα). P2Y1,12,13 receptors enhance glycine transport in the synaptic cleft and P2Y11R activation in glutamatergic neurons delays apoptosis. P2YRs increase dopamine and glutamate release, but can also inhibit neurotransmitter release in the cerebral cortex and hippocampus. In microglia,
P2Y1,2,6,12,13 receptor activation increases microglial cell migration and P2Y2, 6 receptors promote Aβ uptake and degradation. In astrocytes, P2Y1,6 receptors stimulate the production of proinflammatory cytokines and chemokines, P2Y2,4 receptors increase the production and secretion of APP and P2Y14R increases expression of MMP9, which degrades Aβ.
44
45
46 Chapter II
The P2Y2 Receptor Interacts with VE-Cadherin and VEGF Receptor-2
to Regulate Rac1 Activity in Endothelial Cells
Abstract
Vascular endothelial cadherin (VE-cadherin) mediates homotypic adhesion between endothelial cells and is an important regulator of angiogenesis, blood vessel permeability and leukocyte trafficking. Rac1, a member of the Rho family of GTPases, controls VE- cadherin adhesion by acting downstream of several growth factors, including angiopoietin-1 and vascular endothelial growth factor (VEGF). Here we show that UTP- induced activation of the Gq protein-coupled P2Y2 nucleotide receptor (P2Y2R) in human coronary artery endothelial cells (HCAECs) activated Rac1 and caused a transient complex to form between P2Y2R, VE-cadherin and VEGF receptor-2 (VEGFR-2).
Knockdown of VE-cadherin expression with siRNA did not affect UTP-induced activation of extracellular signal-regulated kinases 1/2 (ERK1/2) but led to a loss of UTP- induced Rac1 activation and tyrosine phosphorylation of p120 catenin, a cytoplasmic protein that interacts with VE-cadherin. Activation of the P2Y2R by UTP also caused a prolonged interaction between p120 catenin and vav2 (a guanine nucleotide exchange factor for Rac) that correlated with the kinetics of UTP-induced tyrosine phosphorylation of p120 catenin and VE-cadherin. Inhibitors of VEGFR-2 (SU1498) or Src (PP2) significantly diminished UTP-induced Rac1 activation, tyrosine phosphorylation of p120 catenin and VE-cadherin, and association of the P2Y2R with VE-cadherin and p120
47 catenin with vav2. These findings suggest that the P2Y2R uses Src and VEGFR-2 to mediate association of the P2Y2R with VE-cadherin complexes in endothelial adherens junctions to activate Rac1. An in vivo permeability assay revealed reduced permeability
−/− of splenic, colonic and renal vessels to albumin in the P2Y2R mice compared to age- matched wild type controls during steady state, suggesting that a possible physiological effect of the P2Y2R-mediated VE-cadherin signaling and Rac1 activation is to maintain vascular permeability in response to shear stress in spleen, colon and kidney.
Introduction
Vascular endothelium functions as a semipermeable barrier between the bloodstream and peripheral tissues. Dysfunction of the vascular endothelium often occurs during inflammatory and allergic reactions, accompanied by edema and infiltration of leukocyte in the tissues. The formation and disruption of vascular endothelial cell intercellular adhesion structures, including adherens junctions, tight junctions and gap junctions, as well as the contractility provided by the actomyosin cytoskeleton are key to transendothelial migration (TEM) of immune cells and blood vessel permeability to macromolecules [18]. Many signaling molecules involved in inflammation such as vascular endothelial growth factor (VEGF), thrombin, tumor necrosis factor α (TNFα) and histamine also can increase endothelial permeability through modulation of protein distribution in adherens junctions and the activities of Rho GTPases [31]. The Rho family
GTPases (RhoA, Rac1 and Cdc42) regulate actin cytoskeletal organization and stability of intercellular junctions [297]. However, it has become apparent recently that Rho
GTPases can either stabilize or destabilize the endothelial barrier depending on the
48 scenario and also through complex interactions with regulators of G protein signaling
(guanine nucleotide exchange factors, GEFs; guanine dissociation inhibitors, GDIs; and
GTPase accelerating proteins, GAPs) [31, 298].
Recently, the P2Y2 nucleotide receptor (P2Y2R), a Gq protein-coupled receptor activated by ATP and UTP with equipotency, has emerged as an important regulator of blood vessel permeability and immune cell recruitment. In vivo studies indicated that activation of the P2Y2R transiently increases microvascular leakage to macromolecules
[299] and promotes extravasation of leukocytes and lymphocytes in many inflammatory conditions, including atherosclerosis, asthmatic airway inflammation, Alzheimer’s disease, autoimmune diseases, bacterial infection and chronic obstructive pulmonary disease (COPD) [237, 244, 292, 300-302]. In addition, accumulating evidence recently demonstrated a role for the P2Y2R in metastasis of multiple types of cancer cells. Besides the regulation by P2Y2R within the cancer cells [304], ATP is released from cancer cells
[213] and platelets [303] to activate endothelial P2Y2R to promote transendothelial migration and metastasis of tumor cells.
By virtue of an arginine-glycine-aspartate (RGD) integrin-binding motif in its extracellular domain, the P2Y2R mediates the activation of small Rho GTPases, Rac1 and
RhoA [224, 225]. The P2Y2R also has SH3-binding motifs in its intracellular domain for interaction with Src and promotes the Src-dependent activation of several growth factor receptors including VEGF receptor 2 (VEGFR-2), which upregulates the expression of vascular cell adhesion molecule-1 (VCAM-1), an immune cell binding protein in endothelial cells [226, 227]. Considering the roles of the P2Y2R in the regulation of vascular barrier integrity, leukocyte adhesion and TEM, and Rho GTPase activities [224-
49 227, 237, 244, 292, 299-302, 305], we speculated that the P2Y2R may modulate the permeability of endothelium by affecting the stability of adherens junctions and Rho
GTPase signaling.
Vascular endothelial cadherin (VE-cadherin) is an endothelial-exclusive adhesion molecule [306] that has been well recognized for its crucial role in regulating vascular permeability as well as leukocyte recruitment and extravasation under inflammatory conditions [307-310]. Deletion of VE-cadherin in mice causes severe defects in vascular development and embryonic death [311, 312]. Down-regulation of VE-cadherin has been associated with vascular tumor growth [313], and neutralization of VE-cadherin with a specific antibody increased VEGF-induced VEGFR-2 activity in endothelial cells [314].
VE-cadherin-null endothelial cells have thinner actin stress fibers, decreased vinculin- positive focal contacts, and attenuated activity of Rac1 [315]. The N-terminal extracellular domain of VE-cadherin mediates Ca2+-dependent homophilic adhesion between adjacent cells while the cytoplasmic domain interacts with various intracellular binding partners, including α-, β-, γ- and p120 catenins, which provide a linkage to actin cytoskeleton and trigger downstream signaling events [20]. For example, p120 catenin regulates actin cytoskeletal organization and cell motility by activation of Rho GTPase
[21-23]. In addition, interactions between VE-cadherin and VEGFR-2, intracellular signaling molecules, such as Shc and Csk [316, 317], and vascular endothelial protein tyrosine phosphatase (VE-PTP) [318], are suggested to regulate cell-cell contacts, cell adhesion, and growth factor signaling [306].
Previous studies performed in our laboratory demonstrated that activation of the
P2Y2R promotes monocyte adhesion and extravasation into rabbit carotid arteries and
50 increases the development of atherosclerotic plaques [244]. Furthermore, mechanistic studies in human coronary artery endothelial cells (HCAECs) revealed that the activated
P2Y2R associates transiently with VEGFR-2 in a Src-dependent manner and that
VEGFR-2 activity is necessary for VCAM-1 upregulation induced by UTP [226]. In this work, we investigated how activation of the P2Y2R affects its distribution and association with VE-cadherin in HCAECs. Our findings demonstrate that activation of the endothelial P2Y2R causes translocation of this receptor to intercellular junctions and transient association with VE- cadherin. We also show that Rac1 activation by the UTP- stimulated P2Y2R in HCAECs requires tyrosine phosphorylation of VE-cadherin and p120 catenin, and association of p120 catenin with vav2, a GEF for Rac1.
Results
UTP causes clustering of the P2Y2R in endothelial intercellular junctions
To visualize the distribution of P2Y2 receptors in endothelial cells we transfected
HCAECs with cDNA encoding the human P2Y2R (hP2Y2R) tagged with enhanced green fluorescent protein (eGFP). Eighty-four hours after transfection, the GFP-hP2Y2R appeared uniformly distributed in the plasma membrane (Figure II-1). In ~50% of the transfected cells, stimulation with UTP for 5 min caused clustering of eGFP-hP2Y2R in the junctional area between adjacent cells indicated by immunostaining of VE-cadherin, an adherens junction protein involved in endothelial cell-cell adhesion (Figure II-1).
UTP induces transient association of VE-cadherin with P2Y2R and VEGFR-2
51 Since the P2Y2R translocated to endothelial intercellular junctions upon activation
(Figure II-1), we examined whether the activated P2Y2R interacts with the adherens junction protein VE-cadherin. For this experiment, HCAECs were transfected with cDNA encoding the HA-tagged hP2Y2R and immunoprecipitation (IP) was performed with anti-HA-conjugated agarose beads. An interaction between the HA-hP2Y2R and
VE-cadherin in HCAECs was observed by co-IP within 5 min of exposure to 100 µM
UTP (Figure II-2A). Similarly, we also found that UTP treatment caused a transient association (maximum after 5 min) between VE-cadherin and VEGFR-2 in HCAECs
(Figure II-2B). Such rapid and transient association between VEGFR-2 and HA-P2Y2R after UTP stimulation was detected by co-IP in a previous study performed in our laboratory [226]; therefore, a transient complex of P2Y2R, VEGFR-2 and VE-cadherin seem to form upon P2Y2R activation by UTP.
Other previous studies by our laboratory also found that the activated P2Y2R interacts with Src kinase and induces the Src-dependent transactivation of growth factor receptors, including VEGFR-2 [226], the epithelial growth factor receptor (EGFR), and platelet-derived growth factor receptor (PDGFR) [227]. In particular, activation of the
P2Y2R caused a transient (peaking at 5 min) association between P2Y2R and VEGFR-2 and tyrosine phosphorylation of VEGFR-2, which was required for the UTP-induced expression of VCAM-1 [226]. Pretreatment with the VEGFR-2 tyrosine kinase inhibitor
SU1498 diminished both the UTP-induced VEGFR-2 phosphorylation and VCAM-1 expression, while cells pretreated with Src kinase inhibitor PP2 or expressing a mutant
P2Y2R that lacks the C-terminal SH3-binding domains (Del-hP2Y2R) were deficit in
VEGFR-2 phosphorylation in response to UTP stimulation [226]. Hence, Src activity was
52 suspected as being a required signaling mediator for association between VE-cadherin and the P2Y2R, and so was VEGFR-2 tyrosine kinase activity. Accordingly, we pretreated HCAECs with 1 µM of the Src inhibitor PP2 or with 10 µM of the VEGFR-2 tyrosine kinase activity inhibitor SU1498, and found inhibition of the UTP-induce association between VE-cadherin and P2Y2R by either inhibitor (Figure II-2C). This suggested that both Src activity and VEGFR-2 tyrosine kinase activity are required for recruitment of the P2Y2R to endothelial adherens junctions.
Considering interaction between VE-cadherin and VEGFR-2 inhibits the internalization of VEGFR-2 in response to VEGF and, thus, VEGF-induced downstream signaling [319, 320], we overexpressed VE-cadherin in 1321N1 astrocytoma cells lacking endogenous P2 receptors but expressing exogenous HA-hP2Y2R. Similarly, these cells exhibited inhibited UTP-induced internalization of the HA-hP2Y2R (Figure II-3).
The P2Y2R mediates tyrosine phosphorylation of VE-cadherin
It has been shown that VEGF treatment causes interaction between VE-cadherin and
VEGFR-2 in endothelial cells [316] and that VEGF stimulates the tyrosine phosphorylation of adherens junction proteins, including VE-cadherin and its cytoplasmic binding partners, β-catenin, γ-catenin (plakoglobin), and p120 catenin [321]. Likewise, we found that the P2Y2R agonist UTP induced tyrosine phosphorylation of VE-cadherin in HCAECs within 5 min of treatment (Figure II-4A). Unlike the transient association between P2Y2R, VE-cadherin and VEGFR-2, the phosphorylation of VE-cadherin was sustained for more than 30 min (data not shown). To confirm that UTP-induced tyrosine phosphorylation is mediated by the P2Y2R, we knocked down the expression of the
53 endogenous P2Y2R in HCAECs with specific siRNA duplexes and observed inhibited
UTP-induced tyrosine phosphorylation of VE-cadherin in the siRNA-pretreated HCAECs
(Figures II-4B-C). In addition, expression of HA-hP2Y2R and VE-cadherin in P2 receptor-deficient 1321N1 astrocytoma cells enabled UTP to induce tyrosine phosphorylation of VE-cadherin (data not shown), further confirming that the P2Y2R responses to UTP and mediate VE-cadherin phosphorylation.
Src and VEGFR-2 activity are required for P2Y2R-mediated tyrosine phosphorylation of
VE-cadherin and Rac1 activation
We have proved that the UTP-induced interaction between P2Y2R and VE-cadherin depends on functional kinase activity of Src and VEGFR-2 by pretreatment of HCAECs with PP2 or SU1498, which inhibited the HA-hP2Y2R and VE-cadherin interaction
(Figure II-2C). Likewise, UTP-induced tyrosine phosphorylation of VE-cadherin was found to be dependent of the kinase activity of Src and VEGFR-2 (Figures II-5A-B). The role of Src in mediating signal transduction between the P2Y2R and VE-cadherin was further demonstrated by using the Del-hP2Y2R mutant that functions identically to the wild type (WT) receptor in terms of calcium and ERK signaling. This deletion mutant was previously used to show that Src binds to the activated P2Y2R via C-terminal SH3- binding domains [227] and that these domains are required for Src-dependent transactivation of VEGFR-2 and upregulation of VCAM-1 [226]. In this study, we found that UTP did not cause association between VE-cadherin and Del-hP2Y2R nor did UTP induce tyrosine phosphorylation of VE-cadherin in 1321N1 cells expressing Del-hP2Y2R
(Figure II-6). Together, these experiments suggest that binding of Src to SH3-binding
54 domains in the P2Y2R is required for VEGFR-2 transactivation and the stimulation of
VE-cadherin phosphorylation.
Furthermore, the presence of VE-cadherin was associated with activation of Rac1 as well as a Rac1 GEF, vav2 [315, 322], both of which were activated by UTP treatment in transfected P2Y2R-expressing 1321N1 astrocytoma cells accompanied by enhanced cell migration [225]. Here we report that UTP induced Rac1 activation in HCAECs in a Src- and VEGFR-2-dependent manner, demonstrated by inhibition with PP2 and SU1498, respectively (Figure II-5C), suggesting roles for Src and VEGFR-2 in the P2Y2R- mediated regulation of the vascular barrier.
VE-cadherin and VEGFR-2 are required for UTP-induced Rac1 activation but not
MAPK signaling
Previous studies have revealed involvement of VE-cadherin-containing adherens junctions in regulating cell proliferation [319, 320] as well as actin cytoskeletal organization [315]. To investigate the role of VE-cadherin in P2Y2R-mediated signal transduction in endothelial cells, VE-cadherin-specific siRNA was used to downregulate the expression of VE-cadherin in HCAECs. It was reported that in VEGF-stimulated confluent VE-cadherin-null endothelial cell cultures, numbers of proliferating cells and the phosphorylation levels of extracellular signal-regulated kinases 1/2 (ERK1/2; i.e., p44/42 mitogen-activated protein kinase or MAPK) were reduced compared to the confluent VE-cadherin-positive cells [320]. In other words, the presence of VE-cadherin restrained the phosphorylation of VEGFR-2 in response to VEGF and contributes to contact inhibition of cell proliferation, which was found to be dependent on β-catenin and
55 its binding to VE-cadherin [320]. However, while VE-cadherin-binding or total levels of
β-catenin or its paralogue γ-catenin (plakoglobin) were not affected by UTP stimulation
(Figure II-7), we also found that P2Y2R-mediated MAPK activation in HCAECs was not affected by VE-cadherin knockdown with specific siRNA (Figure II-8A). In contrast, pretreatment with VE-cadherin siRNA resulted in an inhibition of UTP-induced activation of Rac1 (Figure II-8B). Consistent with a previous report [315], we found that knockdown of VE-cadherin expression in HCAECs caused an increase in basal Rho activity, which was not further stimulated by UTP (Figure II-9). Since both UTP-induced phosphorylation of VE-cadherin and activation of Rac1 are regulated by VEGFR-2, we conclude that VEGFR-2 tyrosine kinase activity is required for VE-cadherin-dependent activation of Rac1 mediated by the P2Y2R.
The P2Y2R regulates p120 catenin phosphorylation via VEGFR-2 and VE-cadherin
Catenins that interact with the cytoplasmic domain of VE-cadherin provide a linkage with actin cytoskeleton as well as intracellular signaling pathways [20]. Among these adherens junction proteins, β- or γ-catenin provides a linkage to the actin cytoskeleton through a dynamic interaction with α-catenin [20], whereas p120 catenin regulates the activity of small Rho GTPases, thereby modulating actin cytoskeletal organization and cell motility [21-23]. Tyrosine phosphorylation of p120 catenin is dependent upon an interaction between p120 and VE-cadherin, thereby regulating endothelial barrier function [323]. We found that in HCAECs, UTP caused tyrosine phosphorylation of p120 catenin, which was mediated by the P2Y2R as demonstrated by the abrogation by P2Y2R- specific siRNA transfection (Figure II-10A). Similar inhibition was found in HCAECs
56 pretreated with PP2 (Figure II-10B) or SU1498 (Figure II-10C) prior to UTP stimulation, indicating a role for Src-dependent VEGFR-2 transactivation in UTP-induced p120 catenin phosphorylation. Furthermore, knockdown of VE-cadherin expression with specific siRNA also inhibited P2Y2R-mediated tyrosine phosphorylation of p120 catenin
(Figure II-10D), suggesting that VE-cadherin is involved in the modulation of p120 catenin phosphorylation in response to P2Y2R activation.
P2Y2R-mediated activation of Rac1 is regulated by p120 catenin
To examine the role of p120 catenin in P2Y2R-mediated VE-cadherin-dependent
Rac1 activation, HCAECs were transfected with p120 catenin-specific siRNA to downregulate expression of p120 catenin. UTP-induced activation of Rac1 was inhibited by p120 catenin-specific siRNA transfection (Figure II-11), further supporting the idea that adherens junction proteins are necessary for the P2Y2R to modulate Rho GTPase activity in endothelial cells. In addition, similar to VE-cadherin-specific siRNA (Figure
II-9), knockdown of p120 catenin expression by siRNA also increased the basal activity of Rho in HCAECs although no further activation by UTP was seen (data not shown).
Previously, we found that the Rac1 GEF vav2 regulated P2Y2R-mediated Rac1 activation, since the expression of dominant negative vav2 inhibits Rac1 activity in UTP- treated 1321N1 cells expressing the P2Y2R [225]. It has been postulated that vav2 binding to p120 is required for P2Y2R-mediated Rac1 activation [21], which is demonstrated here by the UTP-induced increase in the interaction between p120 catenin and vav2 in HCAECs (Figure II-12A). Furthermore, the UTP-induced binding of vav2
57 and p120 catenin was inhibited by SU1498 or PP2 (Figure II-12B), consistent with a role for Src-dependent VEGFR-2 transactivation in P2Y2R-mediated Rac1 activity.
The P2Y2R regulates vascular permeability in vivo
To evaluate whether the P2Y2R regulates the vascular barrier function in vivo, we
−/− performed a permeability assay in P2Y2R and age-matched wild type (WT) mice as previously described [324]. In brief, mice were subjected to intravenous injection of the dye Evans blue that has very high affinity for albumin. After 1 h, the mice were sacrificed, perfused with saline to deplete Evans blue-labeled albumin from the vasculature, and the organs were harvested and incubated in formamide to extract Evans blue-labeled albumin that had leaked into the tissues. Statistical analysis indicated significantly lower permeability to Evans blue-labeled albumin in the spleen, colon and
−/− kidney of the P2Y2R than WT mice (P < 0.001) (Figure II-13), indicating a role for
P2Y2R in the regulation of vasculature permeability in vivo.
Discussion
The endothelium plays an active role in vascular barrier function [325-329] and in vivo evidence indicates that the P2Y2R is an important regulator of microvascular permeability
[299] and transendothelial migration of leukocytes into injured or infected tissue [237,
244, 292, 300-302]. Here, we examined the distribution of eGFP-tagged P2Y2Rs in quiescent and UTP-treated endothelial cells. In contrast to findings in migrating cells, where the activated P2Y2R is found evenly distributed on the cell surface [224, 237], we found that activation of the P2Y2R in confluent endothelial cell monolayers by UTP
58 caused a rapid and transient translocation of eGFP-tagged P2Y2Rs to the cell-cell junctional zones, indicated by VE-cadherin (Figure II-1), a crucial adhesion molecule in adherens junctions of endothelial cells that is critical for maintaining the vascular barrier
[330]. Further experiments utilizing co-IP demonstrated a transient complex formed between the P2Y2R and VE-cadherin, in which VEGFR-2 was also engaged (Figure II-2A and B). Based on these findings, we speculated that the endothelial P2Y2R affects vascular barrier function by relocating to endothelial cell junctions and interacting with VE- cadherin.
The endothelial-exclusive adhesion molecule, VE-cadherin is a well-known regulator of vascular barrier function [330, 331]. In non-angiogenic endothelium, VE-cadherin molecules of neighboring cells form homophilic adhesion through the extracellular domain, while the cytoplasmic tail is directly linked to other adherens junction proteins, such as p120, and β- and γ-catenins, as well as indirectly to α-catenin through an interaction with β- or γ-catenin [20]. Several mechanisms have been found to regulate VE- cadherin signaling so as to control the vascular barrier function, including serine and tyrosine phosphorylation, clathrin-dependent internalization and metalloprotease
ADAM10-dependent shedding of VE-cadherin [18, 330, 332-335]. In the present work, we report that the UTP-activated P2Y2R mediates sustained overall phosphorylation of
VE-cadherin (Figure II-4) even after the transient complex of P2Y2R/VE- cadherin/VEGFR-2 has disassembled (Figure II-2), although which residues are phosphorylated awaits further investigation. In addition, we observed UTP-induced tyrosine phosphorylation of p120 catenin, a VE-cadherin-associated protein (Figure II-10), and coincident activation of Rac1 (Figures II-8B-C) and interaction between the Rac1
59 GEF vav2 and p120 catenin (Figure II-12), which seems to mimic the previously reported signaling pathway (i.e., Src-vav2-Rac1-PAK1 or p21 activated kinase 1) for VEGF- induced serine phosphorylation and β-arrestin-dependent endocytosis of VE-cadherin
[336]. However, we did not observe noticeable translocation of VE-cadherin in HCAECs by immunostaining after a 5 min or 30 min treatment with UTP, whereas VEGF was reported to induce rapid internalization of VE-cadherin within minutes [333], suggesting that the P2Y2R may not regulate endothelial barrier integrity via VE-cadherin endocytosis.
Other potential mechanisms for the P2Y2R to regulate vascular integrity via VE-cadherin exist, but were not explored in this study. For example, P2Y2R activation has been reported to increase cytoplasmic calcium levels, activate ADAM10 metalloprotease [337], phosphorylate myosin light chain (MLC) [338] and focal adhesion kinase (FAK) [339] and upregulate VCAM-1 and intercellular adhesion molecule-1 (ICAM-1) expression in vascular cells [226, 245]. As mentioned above, VE-cadherin can be cleaved by
ADAM10 in the ectodomain in response to calcium influx, and ADAM10 activity is necessary for permeability of fluorescently labeled dextran and T cell TEM [335].
Phosphorylation of MLC is critical for monocyte-induced tyrosine phosphorylation of VE- cadherin and monocyte TEM [340], while FAK activity is required for the VEGF-induced increase in vascular permeability and for β-catenin phosphorylation and dissociation from
VE-cadherin [341]. Moreover, induction of VE-cadherin tyrosine phosphorylation has been found in endothelial cells treated with cross-linking antibodies against VCAM-1 and
ICAM-1 to mimic the clustering of these adhesion molecules that occurs during leukocyte
TEM [340, 342].
60 On the other hand, the role of VE-cadherin in controlling cell proliferation is well recognized and requires interactions between VE-cadherin and Src kinases, catenins and the cell cytoskeleton [317, 343, 344]. In the case of VEGF signaling in confluent endothelial cell culture, the presence of VE-cadherin restrains cell proliferation by preventing internalization of VEGFR-2 that promotes the activation of the mitogen- activated protein kinases ERK1/2 [319]. We found that overexpression of VE-cadherin in cells stably expressing the P2Y2R tagged with hemagglutinin (HA) inhibits UTP-induced internalization of the P2Y2R (Figure II-3). Unlike Rac1 activation, however, UTP- induced ERK1/2 activation was unaffected by either downregulation or overexpression of
VE-cadherin (Figure II-8). This is not surprising and could be attributed to different signaling pathways for internalized VEGFR-2 and P2Y2R. While signaling by the P2 family of G protein-coupled receptors is shut off by agonist-induced desensitization and internalization [345], internalized VEGFR-2 remains phosphorylated at the binding site for phospholipase C-γ (PLC-γ) and continues to activate PLC-γ and downstream cascades, including MAPK pathway [319].
The tyrosine kinase activities of Src and VEGFR-2 are indispensable to the association and crosstalk between P2Y2R and VE-cadherin in response to UTP, demonstrated by the inhibition of the binding of P2Y2R and VE-cadherin, phosphorylation of VE-cadherin and downstream signaling events by pretreatment of endothelial cells with PP2 or SU1498, compounds that inhibit Src and VEGFR-2 kinase activity, respectively (Figures II-2C, II-5, II-10B-C, and II-12B). In addition to inducing
VE-cadherin/P2Y2R interaction, UTP also caused a rapid and transient association between VEGFR-2 and VE-cadherin (Figure II-2B), suggesting that P2Y2R activation
61 recruits VEGFR-2 to the VE-cadherin/P2Y2R complex. These findings are consistent with previous reports that both Src and VEGFR-2 can associate with the activated P2Y2R
[226, 227]. VEGFR-2 is known to associate with VE-cadherin in response to VEGF stimulation as well as shear stress in vascular endothelial cells [346]. Since nucleotides are known to be released in response to mechanical stress [204], this implies that the
P2Y2R acts as a shear stress sensor in the vasculature to control VE-cadherin complexes and associated vascular barrier function.
Besides Src and VEGFR-2, integrins may also play a role in regulating P2Y2R/VE- cadherin interactions. It has been shown that fibronectin binding to integrins disrupts VE- cadherin-containing adherens junctions in a Src-dependent manner [347]. The P2Y2R contains an extracellular RGD domain that renders affinity for the fibronectin-binding integrins, αvβ3 and αvβ5 [223]. When the RGD sequence was mutated to arginine-glycine- glutamate (RGE) that is devoid of binding affinity for the αvβ3/β5 integrins, the RGE-
P2Y2R mutant did not exhibit UTP-induced phosphorylation of VE-cadherin (data not shown), suggesting that P2Y2R/integrin interactions also play a role in signaling to VE- cadherin although further studies are required.
In fact, intracellular signaling transduced by VE-cadherin is complex and varies depending on whether the vasculature is growing (angiogenic) or established [331]. In confluent endothelial cultures, VE-cadherin activity more closely resembles an established, resting vascular bed, with VE-cadherin clustered in adherens junctions at cell-to-cell borders and small GTPase activity favoring inhibition of RhoA activity, which decreases actomyosin contractility. We found that activation of the P2Y2R with
UTP in confluent HCAECs cultures activates both RhoA and Rac1 and in vivo studies
62 show that the Rho-associated protein kinase inhibitor Y27632 blocks the increase in microvascular permeability induced by UTP [299]. Since Rac1 is known to regulate the integrity of adherens junctions and participate in cytoskeletal rearrangements important for endothelial permeability induced by thrombin and histamine [348] and transendothelial leukocyte migration [349], we evaluated vascular permeability in adult mice during steady state, and found that the vasculature of spleen, colon and kidney in the P2Y2R-null mice are significantly less permeable to Evans blue-labeled albumin than age-matched WT mice (Figure II-13). It has been shown that blood flow-induced shear stress causes ATP release from red blood cells [208] and endothelial cells [210]. Such an observation is consistent with previously reported UTP-induced increases in cremaster
−/− venular permeability to albumin in WT but not P2Y2R mice [299], and might reflects a role of the P2Y2R in splenic, colonic and renal endothelial cells in response to the nucleotides released under mechanical stress [204]. Yet, the tissue specificity of the response, and the relevance of P2Y2R-mediated Rac1 activity and VE-cadherin signaling and P2Y2R-mediated vasodilation [264, 278] have not been explored. Interestingly, inhibition or activation of Rac1 has been shown to increase the permeability of endothelial cells [348], suggesting that precise control of Rac1 activity is needed to maintain the integrity of the vascular barrier. Extensional investigation based on the present work would be beneficial to elucidate the molecular mechanisms of P2Y2R- regulated permeability and leukocyte extravasation.
To summarize, we present here that in endothelial cells, activation of the G protein- coupled P2Y2R induces the Src- and VEGFR-2-dependent activation of VE-cadherin in adherens junctions, which leads to the p120 catenin- and vav2-dependent activation of
63 Rac1 (Figure II-14). Since the P2Y2R can associate with VE-cadherin as well as VEGFR-
2 [226] and αvβ3/β5 integrins [223], these results suggest that the P2Y2R participates in a multi-receptor complex to regulate the function of adherens junctions. This study elucidates a novel mechanism whereby nucleotides act as signaling cues to modulate adherens junctions and activate Rac1 signaling in endothelium, an important process for regulating vascular permeability and inflammation.
Materials and Methods
Materials
−/− tm1Bhk P2Y2R mice (B6.129P2-P2ry2 /J, stock #009132) were purchased from
Jackson Laboratories (Bar Harbor, ME), and bred at the Christopher S. Bond Life
Sciences Center Animal Facility of the University of Missouri, Columbia, MO. Goat anti-human VE-cadherin polyclonal antibody targeting the C-terminus of VE-cadherin, rabbit anti-human Flk-1 (VEGFR-2) polyclonal antibody, and rabbit anti-vav2 polyclonal antibody were purchased from Santa Cruz Biotechnology (Dallas, TX). The mouse anti- phosphotyrosine antibody, mouse anti-β catenin antibody, and mouse anti-γ catenin
(plakoglobin) antibody were purchased from BD Bioscience (San Jose, CA). Mouse anti-
HA antibody-conjugated agarose beads and rabbit anti-HA antibody were purchased from
Covance (Berkeley, CA). Anti-p120 catenin antibodies were purchased from Santa Cruz
Biotechnology and BD Bioscience. Rabbit polyclonal anti-phospho-ERK1/2 antibody was purchased from Cell Signaling (Beverly, MA). Specific inhibitors for VEGFR-2 tyrosine phosphorylation (SU1498) and Src (PP2) were obtained from Calbiochem
(Indianapolis, IN). ON-TARGETplus SMARTpool siRNA duplexes targeting the human
64 P2Y2 receptor, VE-cadherin and p120 were purchased from Dharmacon (Chicago, IL).
VE-cadherin cDNA was a kind gift from Dr. Elisabetta Dejana (IFOM-IEO, Milan,
Italy). All other reagents including nucleotides were obtained from Sigma-Aldrich (St.
Louis, MO), unless otherwise specified.
Cell Culture and Transfection
HCAECs were cultured in endothelial growth medium-2 (EGM-2; Clonetics,
Walkerville, MD) supplemented with 100 U/mL penicillin and 100 µg/mL streptomycin
at 37°C in a humidified atmosphere of 5% CO2 and 95% air. HCAECs between the fourth and eighth passages were used. For transient transfections, siRNA or plasmid constructs were delivered using Targefect F-2 plus Virofect or Targefect-HUVEC from Targeting
Systems (Santee, CA), respectively, according to the manufacturer’s instructions. In both cases, transfection efficiency of at least 60% was achieved. Human P2Y2R (hP2Y2R) cDNA encoding a hemagglutinin (HA) tag at the N-terminus in pcDNA3.1(–) [224] or cDNA encoding the hP2Y2R cDNA with an eGFP tag at the C-terminus in pEGFP-N1 (a kind gift from Dr. Fernando A. González, Department of Chemistry, University of Puerto
Rico) were transiently expressed in HCAECs. Endogenously P2 receptor-null human
1321N1 astrocytoma cells were cultured in Dulbecco's modified Eagle's medium
(DMEM; Life Technologies,) containing 5% (v/v) fetal bovine serum (FBS; Life
Technologies), 100 U/mL penicillin and 100 µg/mL streptomycin (P/S; Life
Technologies) and maintained at 37°C in a humidified atmosphere of 5% CO2 and 95%
air. Cells were stably transfected with cDNA encoding either the wild type P2Y2R or a mutant P2Y2R in which the C-terminal proline-rich SH3-binding domains where deleted
65 (Del), as previously described [227]. These receptor constructs contained sequence
encoding a HA tag at the N-terminus of the P2Y2R, as previously described [227]. In addition, the cells were transiently transfected with either pcDNA3 vector or pcDNA-VE- cadherin using the Lipofectamine 2000 reagent (Life Technologies).
Confocal Laser Scanning Microscopy Visualization
HCAECs plated on glass coverslips were cultured to ~90% confluence before being transfected with hP2Y2R-eGFP cDNA. The cell transfectants were maintained in EGM-2 for 72 h and transferred to serum-free endothelial basal medium-2 (EBM-2; Clonetics) for 12 h. Then, cells were treated with or without 100 µM UTP for 5 min at 37°C, washed in ice-cold PBS, fixed for 10 min in 4% (w/v) paraformaldehyde, treated with 0.1% (v/v)
Triton X-100 for 5 min, and rinsed in PBS. Fixed cells were incubated with mouse anti- human VE-cadherin antibody (1:100 dilution, BD Bioscience, San Jose, CA) for 1 h, washed and stained with Alexa Fluor 594 goat anti-mouse IgG (1:200 dilution; Life
Technologies) for 1 h, followed by incubation with Hoechst (Life Technologies).
Coverslips were mounted on glass slides in ProLong antifade reagent (Life Technologies) and examined using a Zeiss inverted LSM 510 META confocal laser scanning microscope (CLSM) equipped with a C Apochromat 40× objective. Images were acquired, processed and analyzed with a Zeiss LSM Image Examiner.
RNA Extraction and RT-PCR
Isolation of RNA, cDNA synthesis, and RT-PCR were performed as previously described [350]. Amplification of P2Y2R cDNA was performed by RT-PCR using the
66 following oligonucleotide primers: sense 5’-CTTCAACGAGGACTTCAAGTACGTGC-
3’, and antisense 5’-CATGTTGATGGCGTTGAGGGTGTGG-3’. Primers for amplification of human glyceraldehyde 3-phosphate dehydrogenase (GAPDH) cDNA were: sense 5'-TGAAGGTCGGAGTCAACGGATTTGGT-3', and antisense 5'-
CATGTGGGCCATGAGGTCCACCAC-3'. Thirty-five amplification cycles were used, with annealing temperatures of 60°C as previously described [350]. PCR products were resolved by 2% (w/v) agarose gel electrophoresis.
Rac1 and RhoA Activity Assay
A Rac1 activation assay kit (EMD Millipore, Billerica, MA) was used to assess Rac1 activity according to the manufacturer's instructions. Briefly, cells were cultured in 6-well tissue culture dishes in EGM-2 and then transferred to serum-free EBM-2 for 12 h before incubation with or without UTP for 5 min at 37°C. Then, cells were washed three times with ice-cold PBS, suspended in Lysis Buffer containing 125 mM HEPES, pH 7.5, 750
mM NaCl, 5% (v/v) Igepal CA-630, 50 mM MgCl2, 5 mM EDTA and 10% (v/v) glycerol, and the lysates were transferred to 1.5 mL tubes. Thirty microliters of agarose- conjugated p21 binding domain of PAK1 that only recognizes GTP-bound Rac1 were added to 500 µl of lysate for 1 h at 4°C. The beads were collected by centrifugation and washed three times with Lysis Buffer. Finally, the beads were resuspended in 40 µL of 2×
Laemmli sample buffer (120 mM Tris-HCl, pH 6.8, 2% (w/v) SDS, 10% (w/v) sucrose, 1 mM EDTA, 50 mM dithiothreitol and 0.003% (w/v) Bromophenol Blue) and Western blotting or immunoblotting (IB) analysis (see below) was performed with a 1:1000 dilution of mouse anti-human Rac1 antibody (Upstate Biotechnology, NY). Rho activity
67 was determined similarly except that p21-PAK-1-agarose and anti-Rac1 antibody were replaced with Rhotekin Rho binding domain (RBD)-agarose and mouse anti-human Rho antibody (EMD Millipore, Billerica, MA), respectively.
Immunoprecipitation and Immunoblotting
Immunoprecipitation (IP) and immunoblotting (IB) were performed, as previously described [224]. Cells were transferred to serum-free medium for 12 h before treatment with the indicated inhibitors and/or UTP at 37°C and cell lysates were used for IP with the indicated antibodies. The immune complexes were precipitated with protein A- or protein G-conjugated beads and analyzed by IB with antibodies against the proteins of interest. After IB, the membranes were stripped and reprobed with the same antibody used for IP to verify the consistency of protein precipitation between samples. IP also was performed with lysates from cells expressing the HA-tagged hP2Y2R using anti-HA- conjugated agarose beads.
Internalization of HA-hP2Y2R
Internalization of the HA-hP2Y2R was examined indirectly by determining the uptake of HA antibodies added to 1321N1 astrocytoma cells stably expressing the wild type HA- hP2Y2R, as described [351]. Briefly, the cells were transfected with either pcDNA3 or pcDNA3-VE-cadherin. Then, cell transfectants were incubated at 37°C in serum-free medium supplemented with 5 µg/mL anti-HA antibodies in the absence or presence of 1 mM UTP for 5 min to allow endocytosis of anti-HA antibody bound to the HA-hP2Y2R.
Cells were then placed on ice to prevent further receptor internalization, washed with ice-
68 cold PBS, and surface-bound antibodies were removed by three washes with ice-cold acidic buffer (100 mM glycine, 20 mM magnesium acetate, 50 mM potassium chloride, pH 2.2). After an additional wash with ice-cold PBS, the cells were lysed with 2×
Laemmli sample buffer, and lysates were analyzed for anti-HA antibodies by immunoblotting. Anti-HA antibodies were detected by chemiluminescence using horseradish peroxidase-conjugated antibodies (1:1000 dilution).
In Vivo Permeability Assay
Mice were handled using protocols approved by the Institutional Animal Care and
Use Committee (IACUC) of the University of Missouri (protocol #7793). As previously
−/− described [324], P2Y2R and age-matched WT mice were intravenously injected with
100 µL 4% (w/v) Evans blue solution in 0.9% (w/v) sodium chloride saline. One hour after the injection, the mice were sacrificed and perfused with saline. The brain, lung, liver, spleen, colon and kidney of each mouse were collected, weighed, and incubated in formamide for 48 h at 55°C. Then the supernatant was transferred to an optical cullet, and the absorbance at 620 nm was measured and the absorbance of the blank (formamide only) was subtracted. The corrected absorbance was normalized to the organ weight and
−/− plotted. The statistical difference between the P2Y2R and WT groups was determined by two-tailed t test where P < 0.05 indicated a significant difference between experimental groups.
69 Figure II-1. Translocation of eGFP-P2Y2R to peripheral membranes of endothelial cells in response to UTP. A, HCAECs expressing eGFP-tagged human P2Y2R (shown in green) were stimulated with or without 100 µM UTP for 5 min, as indicated. Cells were washed, fixed, permeabilized and labeled with mouse anti-VE-cadherin antibody. Alexa
Fluor 594-conjugated anti-mouse IgG was then used to stain VE-cadherin (shown in red).
B, Graphs of the distribution of VE-cadherin (red line) and eGFP-P2Y2R (green line) across the eGFP-P2Y2R-labeled cells. The yellow lines indicated by the arrows in (A), mark the sections where the distribution graphs were generated. For each treatment, ~30 cell transfectants from 4 independent experiments were examined.
70 A' Control ' ' ''' ' '+'UTP'(5'min)'
B' Intensity' Intensity'
Distance'(µm)' Distance'(µm)'
71 Figure II-2. UTP causes transient interaction between the HA-P2Y2R, VE-cadherin and VEGFR-2. (A) HCAECs expressing HA-tagged human P2Y2R were treated with
100 µM UTP for the indicated time. Cell lysates were prepared and subjected to IP with mouse anti-HA antibody conjugated to agarose beads. IB analysis of the IP samples was then performed with anti-VE-cadherin antibody to detect co-precipitation of VE-cadherin or with rabbit anti-HA antibody to detect relative levels of HA-tagged P2Y2R precipitated in each sample. (B) HCAECs were treated with 100 µM UTP for the indicated time and cell lysates were subjected to IP with anti-VEGFR-2 antibody and IB with anti-VE- cadherin antibody. The membrane was stripped and re-blotted with anti-VEGFR-2 antibody. (C) HCAECs expressing HA-tagged human P2Y2R were treated with the indicated concentration of the Src kinase inhibitor, PP2, or the VEGFR-2 kinase inhibitor, SU1498, for 30 min followed by incubation with or without 100 µM UTP for 5 min. Cell lysates were prepared and subjected to IP with anti-HA matrix beads and IB with anti-VE-cadherin or anti-HA antibody. Cell lysates also were subjected to IB with anti-VE-cadherin antibody to detect total VE-cadherin. Blots representative of 4 experiments are shown.
72
73 Figure II-3. VE-cadherin inhibits UTP-induced internalization of the HA-hP2Y2R in
1321N1 astrocytoma cells. Human 1321N1 astrocytoma cells stably expressing the wild type HA-hP2Y2R were transiently transfected with pcDNA3 or pcDNA3-VE-cadherin.
Anti-HA antibodies were added to the medium and the cells were incubated with or without 1mM UTP for 5 min to allow endocytosis to proceed. Surface-bound antibodies were removed from cells by acidic buffer wash and internalized anti-HA antibodies were detected by IB, as described in “Materials and Methods”. Total cell lysates also were subjected to IB with anti-VE-cadherin or anti-actin antibodies. Blots representative of 3 experiments are shown.
74
75 Figure II-4. Activation of the P2Y2R by UTP causes sustained tyrosine phosphorylation of VE-cadherin. (A) HCAECs were treated with100 µM UTP for the indicated time. Cell lysates were subjected to IP with anti-phosphotyrosine antibody followed by IB with anti-VE-cadherin antibody to detect tyrosine phosphorylated VE- cadherin. Cell lysates also were subjected to IB with anti-VE-cadherin antibody to detect total VE-cadherin in each sample. (B and C) HCAECs were transfected with either scrambled siRNA or P2Y2R-specific siRNA for 36 h. B, Total RNA was extracted from cell lysates and RT-PCR was performed with either P2Y2R primers or GAPDH primers.
(C) Transfected cells were incubated with or without 100 µM UTP for 5 min. Cell lysates were prepared and subjected to IP with anti-phosphotyrosine antibody and IB with anti-
VE-cadherin antibody. The data shown are representative of results from 4 experiments.
76
77 Figure II-5. Src and VEGFR-2 kinase activity are necessary for P2Y2R-mediated tyrosine phosphorylation of VE-cadherin and Rac1 activation. (A and B) HCAECs expressing HA-tagged human P2Y2R were treated with the indicated concentration of the
Src kinase inhibitor, PP2, or the VEGFR-2 kinase inhibitor, SU1498, for 30 min and then with 100 µM UTP for 5 min. Cell lysates were prepared and subjected to IP with anti- phosphotyrosine antibody and IB with anti-VE-cadherin antibody. Cell lysates also were subjected to IB with anti-VE-cadherin antibody to detect total VE-cadherin. Blots representative of 3 experiments are shown. (C) HCAECs were treated as indicated with
PP2 or SU1498 for 30 min, and then with 100 µM UTP for 5 min prior to measuring
Rac1 activity. Cell lysates also were subjected to IB with anti-Rac1 antibody to detect total Rac1. Blots representative of 3 experiments are shown.
78
79 Figure II-6. The Src-binding domain of the P2Y2R is required for the P2Y2R to interact with and phosphorylate VE-cadherin. Human 1321N1 cells expressing the
HA-tagged WT (wild type) or Del (deletion of prolines in the Src-binding domains) mutant hP2Y2R were transfected with VE-cadherin cDNA in pcDNA3 and incubated with or without 100 µM UTP for 5 min. Cell lysates were prepared and subjected to IP with A, anti-HA matrix or B, anti-phosphotyrosine antibody, followed by IB with anti-
VE-cadherin antibody. Cell lysates also were prepared and analyzed by IB with anti-VE- cadherin antibody. Blots representative of 3 experiments are shown.
80
81 Figure II-7. UTP stimulation does not affect the pool of β- or γ-catenin or the binding of β- or γ-catenin to VE-cadherin. HCAECs were incubated with 100 µM UTP for the indicated times, and cell lysates were prepared and subjected to IB for β-catenin
(A) or γ-catenin (C), or IP of VE-cadherin prior to IB for β-catenin (B) or γ-catenin (D).
Blots representative of 3 experiments are shown.
82
83 Figure II-8. VE-cadherin expression is required for P2Y2R-mediated activation of
Rac1, but not ERK1/2. (A and B) HCAECs were transfected with either scrambled siRNA or VE-cadherin-specific siRNA for 36 h. (A) Transfected cells were incubated with 100 µM UTP for the indicated time and analyzed by IB with either anti-VE-cadherin antibody or anti-phospho-p42/44 (ERK1/2) antibody. (B) Transfected cells were subjected to a Rac1 activity assay, as described in the “Materials and Methods”.
84
85 Figure II-9. Effect of downregulation of VE-cadherin on UTP-induced activation of
Rho in HCAECs. HCAECs were transfected with either scrambled or VE-cadherin- specific siRNA for 36 h, then incubated with the indicated concentration of UTP for 5 min prior to performing a Rho activity assay, as described in “Materials and Methods”.
Cell lysates also were prepared and analyzed by IB with anti-Rho antibody to detect total
Rho in the samples. Blots representative of 3 experiments are shown.
86
87 Figure II-10. P2Y2R activation causes tyrosine phosphorylation of p120 catenin that is dependent upon Src and VEGFR-2 kinase activities and expression of VE- cadherin. (A and D) HCAECs were transfected with the indicated siRNA for 36 h and then incubated with or without 100 µM UTP for 5 min. (B and C) HCAECs were treated with the indicated concentration of PP2 or SU1498 for 30 min, and then incubated with or without 100 µM UTP for 5 min. (A-D) Cell lysates were prepared and subjected to IP with anti-phosphotyrosine antibody and IB with anti-p120 catenin antibody. Cell lysates also were subjected to IB with anti-p120 catenin antibody to detect total p120 catenin.
Blots representative of 3 experiments are shown.
88
89 Figure II-11. Expression of p120 catenin is required for P2Y2R-mediated activation of Rac1. HCAECs were transfected with either scrambled siRNA or p120 catenin- specific siRNA for 36 h and then incubated with or without 100 µM UTP for 5 min prior to measuring Rac1 activity. Cell lysates were prepared and subjected to IB with anti-Rac1 antibody or anti-p120 catenin antibody to detect total Rac1 and p120 catenin, respectively. Blots representative of 3 experiments are shown.
90
91 Figure II-12. P2Y2R activation causes interaction between p120 catenin and vav2.
(A) HCAECs were incubated with 100 µM UTP for the indicated times. (B) HCAECs were treated with the indicated concentration of PP2 or SU1498 for 30 min and then with100 µM UTP for 5 min. (A and B) Cell lysates were prepared and subjected to IP with anti-vav2 antibody and IB with anti-p120 catenin or anti-vav2 antibody. Blots representative of 3 experiments are shown.
92
93 Figure II-13. In vivo permeability in spleen, colon, and kidney is reduced in
−/− −/− P2Y2R mice. Eight-week old P2Y2R (N = 13; open bars) and age-matched wild type mice (N = 15; solid bars) were subjected to the in vivo permeability assay as described in the “Methods and Materials”. Permeabilized albumin-binding dye in major organs as indicated was quantified and normalized to the organ weight. The results were analyzed by unpaired two-tailed t test, and the groups with statistically significant differences (P <
0.05) were labeled with asterisks (***, P < 0.001).
94
95 Figure II-14. A schematic of the signaling pathways activated by the P2Y2R in endothelial cells that are thought to regulate vascular permeability. After activation by ATP or UTP, P2Y2R translocates to the cell-cell adherens junctions and transiently associates with VEGFR-2 and VE-cadherin, which are phosphorylated via a Src- dependent pathway. The kinase Src also phosphorylates VE-cadherin-bound p120 catenin, and possibly β- and γ-catenins as well, to further disrupt the adherens junctions.
Then, p120 catenin associates with the GEF vav2, activating Rac1, which is likely to induce cytoskeletal rearrangements to further facilitate the passage of macromolecules or leukocytes through the endothelial intercellular space. Grey ovals represent the phosphorylation sites. Catenins are represented by light blue ovals, and pink circles represent heterotrimeric G protein subunits.
96
97 Chapter III
P2Y2 Receptor Deficiency in an Alzheimer’s Disease Mouse Model
Decreases CXCL12, SCF and VCAM-1 Expression in the Bone Marrow
and Reduces Levels of Circulating Monocytes and Neutrophils
Abstract
Immune responses in the brain occur in Alzheimer’s disease (AD), yet it is unclear how systemic immune responses contribute to disease progression. Recently, circulating monocytes and neutrophils derived from the bone marrow (BM) have been shown to accumulate around amyloid-β (Aβ) plaques in the brain and are thought to contribute to
Aβ clearance in AD mouse models. Here we report that male TgCRND8 (Tg+) mice, a model of human AD that expresses the Swedish and Indiana mutations of amyloid precursor protein (APP), exhibit neutrophilia at 10 weeks of age that is abrogated by heterozygous
+ + +/− deletion of the P2Y2 nucleotide receptor gene in Tg mice (Tg P2Y2R ). Similarly, the number of circulating monocytes in male Tg+ mice was modestly increased compared to
+ +/− wild type mice, whereas the circulating monocyte level in Tg P2Y2R mice was
+ abnormally low. We previously reported that suppression of P2Y2R expression in Tg mice accelerates AD-like pathology and reduces CD11b+ microglia/leukocyte accumulation around Aβ plaques in the brain. The deficiency in circulating leukocytes in
+ +/− Tg P2Y2R mice described in the current study likely contributes to the reduced accumulation of leukocytes around Aβ plaques in the brain and the accelerated rate of progression of AD symptoms observed in these mice. Assessment of gene expression in
98 BM samples by quantitative real time polymerase chain reaction (RT-PCR) indicated that
+ +/− Tg P2Y2R mice show a striking reduction in the expression of C-X-C chemokine ligand 12 (CXCL12, also known as stromal cell-derived factor 1), stem cell factor (SCF, also known as Kit ligand) and vascular cell adhesion molecule 1 (VCAM-1) as compared to Tg+ mice. Loss of these molecules in the BM is generally considered to facilitate leukocytes and hematopoietic stem/progenitor cells emigrate out of BM into the peripheral blood. Therefore, the inability of leukocytes to emigrate out of the BM in Tg+ mice lacking sufficient P2Y2R expression is likely due to the essential role of the leukocytic P2Y2R in sensing and amplifying chemoattractant signals. This work has revealed a novel function of the P2Y2R in AD in the regulation of the leukocyte release into the bloodstream and modulation of BM homeostasis.
Introduction
Alzheimer’s disease (AD) is a chronic neuroinflammatory disorder that is thought to be initiated by alterations in the processing of amyloid precursor protein (APP) and production of amyloid-β peptides (Aβ) that aggregate and form senile plaques [352, 353].
APP when cleaved by β- and γ-secretases, gives rise to Aβ [353], which induces microglial cells, the resident macrophages in the central nervous system, to produce nitric oxide, reactive oxygen species, and proinflammatory cytokines, leading to neuroinflammation and neuronal death [354]. On the other hand, microglial cells carry out neuroprotective functions in AD by migrating towards Aβ plaques and clearing neurotoxic Aβ by endocytosis and sequential degradation [160, 355-358]. AD patients are often found to also have cerebral amyloid angiopathy (CAA), where apoptotic
99 macrophages and fibrillar Aβ accumulate around vessels [170]. While most studies of
AD have focused on the pathology of the brain, increasing evidence suggests that disorders in the systemic immune system are also involved. In particular, blood-borne monocytes have been shown to infiltrate into the brain in AD mouse models, and assist resident microglial cells to clear Aβ plagues (Reviewed in [167]). Fiala and colleagues found that human AD patient-derived macrophages were defective in terms of phagocytosis; they had exhibited only surface up-take of Aβ and appeared to be apoptotic instead of carrying out intracellular phagocytosis of Aβ, as in cells from age-matched controls [169, 170]. AD mice lacking C-C chemokine receptor 2 (CCR2, also known as monocyte chemotactic protein 1) were found to have significantly reduced perivascular macrophage (PVM) numbers around the Aβ+ vessels and exhibited diminished Aβ phagocytosis by microglia and BM-derived macrophages, lower survival rates, and elevated vascular Aβ deposition, but normal parenchymal Aβ deposition in the brain as compared to AD mice with CCR2 [171]. More recently, live intravital two-photon microscopy was used to demonstrate that circulating Ly6Clow monocytes were attracted selectively to Aβ-positive veins in the brain and enhanced the Aβ clearance through phagocytosis [172]. Furthermore, Shad et al. [173] noticed that AD patients had significantly higher monocyte numbers in the blood regardless of their age and sex; otherwise, significantly higher neutrophils were found in those AD patients with normal monocyte levels.
Recently, elevation of the blood neutrophil-lymphocyte ratio (NLR) has emerged as a marker of systemic inflammation in may diseases/conditions, including cancers [176], hypertension [177], diabetes mellitus [177], coronary artery bypass grafting [178],
100 familial Mediterranean fever [179], ankylosing spondylitis [180], placental inflammatory response [181], schizophrenia [182], and cerebral infarction [183]. Consistent with this,
AD patients were found to have a significantly higher NLR compared to patients with normal cognitive function [175]. In another report, circulating granulocytes (neutrophils and eosinophils) from AD patients as well as controls were separated into fractions according to weight, and AD patients had fewer granulocytes in the light fractions as compared with controls, which was hypothesized to be a sign of impaired granulocyte turnover and cell damage [174]. More recently, two-photon in vivo imaging showed migration of Gr-1 (Ly6G/C) antibody-labeled neutrophils into the brain parenchyma and towards amyloid plaques in 5XFAD mice (an AD model) but not in wild type mice [185].
Yet, the chemotactic activity of neutrophils in 5XFAD mice was likely indirectly induced by Aβ via release of chemokine from resident brain cells [185]. These findings have suggested a role for neutrophils in AD pathology. Moreover, circulating hematopoietic stem/progenitor cells (HSPCs) have also emerged as a factor in AD pathology. Patients with early AD have reduced levels of circulating CD34+ hematopoietic stem cells
(HSCs), which significantly correlates with age, Aβ1-42 levels in the cerebrospinal fluid, and the ratio of Aβ1-42/Aβ1-40 [191].
Nucleotides (ATP, ADP, UTP and UDP) and their cell surface receptors (i.e., the ion channel P2X1-7 and the G protein-coupled P2Y1,2,4,6,11-14 receptors) have emerged as key players in immune responses [216, 359]. In injured tissue, millimolar quantities of ATP and, to a lesser extent, other nucleotides are released from damaged cells, where they act as danger signals by activating P2X and P2Y receptors, leading to a variety of responses, including platelet aggregation, blood vessel dilation, upregulation of cell adhesion
101 proteins, leukocyte recruitment, cytokine release and the perception of pain [296].
Extracellular nucleotides are then hydrolyzed by ectonucleotidases to form nucleosides that induce a variety of pro- and anti-inflammatory responses by activating G protein- coupled adenosine receptors [296].
In vivo studies focusing on the P2Y2 receptor subtype (P2Y2R) indicate that this receptor is important for leukocyte recruitment to stressed, injured or infected tissue. The
P2Y2R, which is activated equally well by ATP or UTP [222], is expressed at high levels in leukocytes and is upregulated in various tissues, including vascular endothelium, smooth muscle and epithelium in response to injury or stress [244]. In rabbit carotid arteries undergoing surgical stress, application of UTP promoted monocyte infiltration into stressed arteries and increased the development of atherosclerotic plaques [244].
Mice lacking the P2Y2R display defects in the migration of neutrophils to sites of
Staphylococcus bacterial infection [237] and to smoke-injured lungs [242], eosinophils and monocyte-derived dendritic cells to allergen-affected lungs [243], and monocytes and macrophages to apoptotic cells in an air-pouch model [240]. In vitro experiments have shown that the release of ATP at the leading edge of migrating neutrophils, through activation of the P2Y2R, direct the orientation and enhanced chemotaxis of neutrophils towards bacterial chemoattractant [237]. Few studies, however, have investigated whether the P2Y2R regulates leukocyte production and emigration from the bone marrow
(BM) to the circulation under inflammatory conditions.
Our laboratory previously reported that in the TgCRND8 mouse model of AD (Tg+), which expresses human APP containing the Swedish (K670N/M671L) and Indiana
(V717F) mutations, loss of the P2Y2R resulted in neurological deficits and striking
102 + −/− + +/− premature death (Tg P2Y2R mice by 4 weeks and Tg P2Y2R by 12 weeks) [292].
+ + +/− Compared to age-matched Tg mice, the Tg P2Y2R mice at 10 weeks of age had increased Aβ plaque load and soluble Aβ1–42 levels, while expression of the leukocyte/microglia marker CD11b in the brain was significantly reduced [292].
Considering P2Y2R’s role in chemotaxis and infiltration of leukocytes in other models of inflammation, we hypothesized that recruitment of circulating leukocytes in addition to
+ +/− the residential microglial population in the brain were impaired in Tg P2Y2R mice. In the present study, we evaluated the leukocytes reservoirs in the peripheral blood and BM
+ of Tg mice with or without heterozygous knockout of the P2Y2R. The results suggest a novel role for the P2Y2R in AD mice in the promotion of monocyte and neutrophil emigration from BM, which likely involves the regulation of leukocyte retention signals in the BM and orientation of leukocytes towards gradients of CXCL12 (C-X-C chemokine ligand 12).
Results
Elevated Levels of Neutrophils and Monocytes in Peripheral Blood of Tg+ Mice is
Prevented by Suppression of P2Y2R Expression
Peripheral blood samples were collected by cardiac puncture from 10-week old male
− +/+ − − +/− + +/+ mice of four genotypes, Tg P2Y2R (i.e., wild type or Tg ), Tg P2Y2R , Tg P2Y2R
+ + +/− (Tg ) and Tg P2Y2R , and the samples were subjected a Complete Blood Count (CBC) test. Although we did not find appreciable differences in the number of total white blood cells (WBCs; i.e., leukocytes) in peripheral blood between the genotypes (Table III-1 and
Figure III-1A), variations were observed in leukocyte subtypes. Consistent with previous
103 reports in human AD patients [174, 175], a significant increase of neutrophil numbers
(Table III-1 and Figure III-1B ) as well as the percentages of neutrophils in total WBCs
(Table III-1) were found in Tg+ mice compared to the Tg− littermate controls, whereas the
+ +/− neutrophil numbers were not elevated in Tg P2Y2R mice (Table III-1 and Figure III-
1B). Similarly, monocyte numbers (Table III-1 and Figure III-1C) and percentages (Table
+ +/− + − +/− III-1) in Tg P2Y2R mice were significantly lower than in Tg and Tg P2Y2R littermate controls. Furthermore, the percentages of lymphocytes in Tg+ mice were
− + +/− significantly lower than in Tg and Tg P2Y2R mice (Table III-1), but the absolute numbers were not significantly different (Table III-1). This suggests that the differences in lymphocyte percentages were due to the changes in the numbers of neutrophils and monocytes instead of changes in the lymphocytes. The numbers of red blood cells (RBCs) and platelets were not significantly different in the examined genotypes (Table III-1).
+ Suppression of P2Y2R Expression in Tg Mice Inhibits Leukocyte Emigration from the
Bone Marrow
To determine whether the changes in the numbers of circulating neutrophils and
+ + +/− monocytes in Tg versus Tg P2Y2R mice arose from alterations in their production in or the emigration from the BM to the peripheral blood, the cellular composition of BM in these mice was assessed. Marrow cells in the femur were collected from 10-week old
− − +/− + + +/− male Tg , Tg P2Y2R , Tg , and Tg P2Y2R mice, and flow cytometry was performed
(Figure III-2) as previously described [360]. In the BM cell population (excluding red blood cells), the percentage of CD45+Ly6G+CD11b+ neutrophils was similar in all four genotypes (Figure III-2B), suggesting that the reduction in circulating neutrophils in
104 + +/− + Tg P2Y2R mice vs. Tg (Figure III-1B) was due to differences in the efficiency of emigration out of the BM. The percentage of CD45+Ly6G−CD115+ monocytes in BM
+ +/− + − +/− − significantly increased in Tg P2Y2R vs. Tg and Tg P2Y2R vs. Tg mice (Figure III-
2C), which also suggests that the reduced number of circulating monocytes in
+ +/− + Tg P2Y2R vs. Tg mice (Figure III-1C) was not due to decreased production of monocytes in the BM but due to their impaired emigration out of the BM. Hence, we hypothesize that the P2Y2R mediates the release of neutrophils and monocytes from BM, especially under inflammatory conditions such as AD.
Other sources of circulating neutrophils and monocytes are the marginated pools. For neutrophils, these include the liver, spleen, and lung, although the reason and mechanism for these organs to retain large populations of neutrophils is not yet well understood [143,
361]. Similarly, the spleen was identified as a storage site for monocytes secondarily to the BM [362]. Therefore, we also investigated the pools of neutrophils and monocytes in the spleen, the common organ of marginated neutrophils and monocytes. It was first
− − +/− + determined that the spleens of 10-week old male Tg , Tg P2Y2R , Tg , and
+ +/− Tg P2Y2R mice were of similar sizes (Figure III-3A). This was a rough indication that no dramatic changes occurred in the spleen, since the weight of the spleen was found to correlate with the splenic neutrophil pool [363] and inflammatory monocyte pool [364].
We used RT-PCR analysis to evaluate the expression levels of neutrophil elastase (Elane) and allograft inflammatory factor 1 (AIF-1, also known as ionized calcium-binding adapter molecule 1, IBA-1), as a measure of splenic neutrophil and monocyte/macrophage pools, respectively. No significant differences were found in neutrophil elastase levels in the spleen of all four genotypes examined (Figure III-3B).
105 + + +/− Although Tg and Tg P2Y2R mice had significantly lower level of AIF-1 in the spleen
− − +/− than their littermate controls, Tg and Tg P2Y2R , respectively, the AIF-1 levels in
+ +/− + Tg P2Y2R mice did not differ significantly from those of Tg mice (Figure III-3C).
These results confirm that the deficiency in neutrophils and monocytes in the peripheral
+ +/− + blood of Tg P2Y2R mice compared to Tg mice was likely due to impaired efflux from the BM rather than a shift from circulating pools to marginated pools.
+ Suppression of P2Y2R Expression in Tg Mice Decreases Bone Marrow Expression of
CXCL12, SCF and VCAM-1
A variety of cytokines and chemokines are known to regulate leukocyte mobilization from the BM. To examine whether the P2Y2R regulates cytokine/chemokine expression in the BM, we extracted total RNA from BM cells of 10-week old male Tg−,
− +/− + + +/− Tg P2Y2R , Tg , and Tg P2Y2R mice, and performed real time polymerase chain reaction (RT-PCR) with selective primers (Figures III-A-C). We assessed the BM expression of CXCL12, which is a C-X-C chemokine that potently attracts HSPCs, pro-B and pre-B cells, megakaryocyte progenitors, T lymphocytes, neutrophils and monocytes
[81]. CXCL12 is expressed in niche-constituting cells in the BM, namely osteoblasts, endothelium, and CXCL12-abundant reticular (CAR) cells [65]. The interaction of
CXCL12 and its primary physiologic receptor, C-X-C chemokine receptor 4 (CXCR4), is not only important for stem cell migration and survival, but also regulates leukocyte mobilization and recruitment [65]. CXCL12-CXCR4 interaction in the BM is generally considered as a retention signal for cells [85, 86], whereas a low-to-high gradient of
CXCL12 from osteoblasts to sinusoidal endothelium and then to the bloodstream is
106 critical for hematopoietic cell emigration [92, 100]. Surprisingly, we found that the
+ +/− mRNA levels of CXCL12 in total BM cells of male Tg P2Y2R mice were much lower
− − +/− + than in the other genotypes, i.e., Tg , Tg P2Y2R and Tg (Figure III-4A).
Immunohistochemistry (IHC) staining of mouse femur using an anti-CXCL12 antibody
+ +/− confirmed that CXCL12 protein levels were reduced in the BM of Tg P2Y2R mice
(Figure III-4D). A possible explanation for the impaired emigration of leukocyte with
+ +/− decreased CXCL12 levels in Tg P2Y2R mice is that the plasma CXCL12 levels in
+ +/− Tg P2Y2R mice are also diminished. However, we found that the CXCL12α level in
+ +/− + plasma of Tg P2Y2R were not significantly reduced compared to Tg mice (Figure III-
5A), although levels of the other isoform CXCL12β were not tested. Also, no significant differences in CXCL12 mRNA expression in the cortex or the spleen were observed
+ + +/− between Tg and Tg P2Y2R mice (Figures III-5B-C), suggesting that regulation of
CXCL12 expression by the P2Y2R is specific to BM.
The expression of stem cell factor (SCF, or Kit ligand) and vascular cell adhesion
+ +/− molecule-1 (VCAM-1) mRNAs were also much lower in BM from Tg P2Y2R mice compared to Tg+ mice (Figures III-4B-C). SCF expression is suppressed when CXCL12 is deleted in adult mice [65], therefore, lower expression levels of CXCL12 in
+ +/− + Tg P2Y2R mice compared to Tg mice may account for the reduction in SCF. Also, activation of the P2Y2R is known to upregulate VCAM-1 expression in endothelial cells
+ +/− [226, 365, 366], so it was not surprising to find that Tg P2Y2R BM cells having lower
VCAM-1 expression levels than Tg+ BM cells. Although the interaction between VCAM-
1 on the stromal cells and its binding partner α4β1 integrin on the leukocytes is generally
107 considered to be a retention signal for leukocytes in the BM cells, a reduction in VCAM-
1 levels is not pivotal to the mobilization of hematopoietic progenitor cells [110].
We also examined the expression levels of three colony-stimulating factors (CSFs) that are cytokines known for monocytic and granulocytic lineage differentiation, granulocyte-CSF (G-CSF, also known as CSF3) and granulocyte-macrophage CSF (GM-
CSF, also known as CSF2) and macrophage CSF (M-CSF, also known as CSF1) [69].
Administration of these CSFs has been shown to increase levels of circulating monocytes or neutrophils [72, 75, 76]. Patients with AD were reported to have increased plasma levels of M-CSF compared to patients with mild cognitive impairment and healthy controls [367] while the plasma levels of G-CSF in early AD patients were lower than healthy controls [368]. Particularly, G-CSF is known to cause a decrease in CXCL12 expression in osteoblasts [88]. Our results show that the mRNA levels of GM-CSF and
G-CSF in BM and cortex are too low to be assessed by RT-PCR (data not shown), while the M-CSF expression is detectable in both types of tissues but not significantly different between the examined genotypes (Figures III-6A-B). Finally, we tested the BM expression of C-C chemokine ligand 2 (CCL2, also known as monocyte chemotactic protein 1), the key chemokine for mobilizing inflammatory monocytes, which has been found to be increased in AD brain and suggested to be involved in Aβ clearance [375-
377]. It has been reported that CCL2 is rapidly upregulated in BM mesenchymal stem cells, including CAR cells, in response to circulating LPS or bacteria, leading to monocyte mobilization [131]. Although CCL2 levels were marginally increased in cortex
+ + +/− − of 10-week old male Tg and Tg P2Y2R mice relative to Tg controls (Figure III-7),
108 the levels of CCL2 in BM samples from all four genotypes were too low to be accurately quantified (data not shown).
Discussion
The most prevalent form of dementia in Western society is AD, which features chronic inflammation and neurodegeneration. The pathophysiology of AD has been intensively studied and recently associated with extracellular nucleotides and purinergic signaling [359], and immune cells other than the microglial cells resident in the CNS, such as bone marrow-derived monocytes [167, 169, 170, 172], myeloid cells [171], neutrophils [174, 175, 185] and HSPCs [191]. Similar to observations in human AD patients [174, 175], we demonstrate here that circulating levels of leukocytes are increased in the TgCRND8 (Tg+) mouse model of AD (Figure III-1). Furthermore, we
+ + +/− show that suppression of P2Y2R expression in Tg mice (Tg P2Y2R ) reduces the number of circulating leukocytes, which likely leads to the reduced brain accumulation of microglia/leukocytes, the inadequate clearing of Aβ plaques, the accelerated neurological deficits and the shorter lifespans that we previously observed in these mice as compared to Tg+ mice [292]. It also was determined that deficiencies in circulating leukocytes in
+ +/− Tg P2Y2R mice are not due to changes in the marginated pools of leukocytes, but is most likely due to impaired emigration from the bone marrow (Figures III-2-3).
+ Suppression of P2Y2R expression in Tg mice also reduced expression of CXCL12, SCF and VCAM-1 in BM (Figure III-4), although further research is needed to reveal the underlying mechanisms. Our preliminary results showed that UTP-induced activation of the P2Y2R in primary BM stromal cell culture increases expression of CXCL12, SCF and
109 VCAM-1, whereas treatment with Aβ decreases CXCL12 and SCF expression but increases VCAM-1 expression (data not shown). These results suggest a novel role for the P2Y2R in regulation of monocyte and neutrophil emigration from BM under inflammatory conditions, which likely involves a yin-yang effect, i.e., counteraction of mobilizing reagent-induced reduction in retention signals in the BM, and promotion of leukocytes chemotaxis towards chemokine gradients.
The chemokine CXCL12 potently attracts HSPCs, pro-B and pre-B cells, megakaryocyte progenitors, T lymphocytes, neutrophils, and monocytes [81], and it is important for leukocyte recruitment and the regulation of stem cell migration and survival
[65]. In BM, CXCL12 is expressed in osteoblasts, endothelium and CXCL12-abundant reticular (CAR) cells [65]. The interaction between CXCL12 and its receptor CXCR4 is generally considered to be a retention signal for HSPCs and leukocytes in BM, since many studies conclude that disruption of this interaction induces cell mobilization. For example, G-CSF, a well-known mobilizing agent for mature neutrophil emigration from the BM to the circulation, acts by reducing CXCL12 levels in BM [85, 86]. Similarly, administration of a CXCR4 antagonist or selective deletion of CXCR4 in the myeloid lineage was found to facilitate neutrophil mobilization from the BM [85, 86]. Several proteases from hematopoietic cells have been found to process the N-terminus of CXCR4 and/or CXCL12 to facilitate their mobilization [89-92]. RNA in situ hybridization determined that the G-CSF-induced decreases in CXCL12 expression occurred specifically in osteoblasts, but not other CXCL12-expressing cells in the BM, including endothelial and CAR cells [88]. Based on this finding and results from other studies, it is hypothesized that emigration of HSPCs and leukocytes from BM to blood requires
110 CXCL12 gradient both within the BM and also across the BM sinusoidal vessels.
CXCL12 expression in osteoblasts that retains CXCR4+ HSPCs and leukocytes in osteoblastic niches is reduced during cell mobilization, and whereupon CXCR4+ cells migrate towards relatively higher concentrations of CXCL12 produced by sinusoidal endothelial cells and surrounding CAR cells in BM. There, a second concentration gradient of CXCL12, as a result from local degradation by protease-mediated cleavage or the enhanced level in circulation delivered from peripheral tissues, or a gradient of other chemokines that have increased levels in the blood enables hematopoietic cells from the
BM side to the circulation side [92, 93, 98-100]. Here, we report a remarkable decrease
+ +/− of CXCL12 expression in the BM of Tg P2Y2R mice (Figures III-4A and D), yet also observe reduced mobilization of leukocytes from the BM in these mice (Figures III-1-2).
+ +/− This inability of leukocytes to emigrate out of the BM in Tg P2Y2R mice is likely due to an essential role of the P2Y2R in sensing and amplifying chemoattractant signals. For example, neutrophils have been shown to rely on the P2Y2R to respond to ATP that is released at the leading edge of cells migrating towards the bacterial chemoattractant formyl-Met-Leu-Phe, which orients the cells and promotes their migration towards the chemoattractant [237]. Autocrine signaling through the P2Y2R is also known to mediate chemokinesis of macrophages and promote their chemotaxis towards the chemoattractant
Complement C5a [238, 239], and P2Y2R-mediated chemotaxis and/or chemokinesis in other cell types have been demonstrated [225, 378]. Also, ATP and UTP have been reported to synergize with CXCL12 to induce chemotaxis of HSPCs in vitro, and UTP- pretreated HSCs homed to the BM in vivo more efficiently than untreated HSCs, which is very likely mediated by the P2Y2R, the only UTP receptor expressed in HSCs [249, 250].
111 Hence, with inadequate P2Y2R expression leukocytes and HSPCs are unlikely to be able to respond to CXCL12 gradients and thereby efficiently emigrate out of the BM to the circulation during inflammatory diseases, e.g., AD.
Besides regulating cell release from BM, CXCL12 was well known for its role in lymphopoiesis, but the recently constructed conditional Cxcl12 homozygous knockout
(CXCL12-cKO) mice only had significantly higher percentages of neutrophils in blood than wild type mice, whereas lymphocyte and monocyte percentage levels were not significantly affected [65]. In contrast, levels of the Lineage-Sca1+c-Kit+ (LSK) cell population comprising long- and short-term HSCs and multipotent progenitors in the BM and blood increased, while quiescent long-term HSCs in the BM were reduced in
CXCL12-cKO mice, suggesting a crucial role for CXCL12 in the quiescence of long- term HSCs [65]. Consistent with this report, we did not find significant differences in
+ +/− + lymphocyte numbers between 10-week old male Tg P2Y2R and Tg mice and their littermate controls (Table III-1), despite of the significant reduction in CXCL12
+ +/− + expression observed in the BM of male Tg P2Y2R vs. Tg mice (Figure III-4A). Our
+ +/− preliminary data also suggest an augment of LSK cell population in BM of Tg P2Y2R mice as compared to Tg+ mice (data not shown). It remains to be determined whether
+ +/− increases of LSK cell levels in Tg P2Y2R mice are due to impaired egress like leukocytes, or disturbed quiescence of long-term HSCs in the BM. The circulating HSCs have been suggested to contribute to regeneration of brain tissues through transdifferentiation into neuronal and glial cells, and early AD patients were found to have decreased circulating CD34+ HSCs [191]. Of note, an AD mouse model (3xTg-AD) receiving neural stem cells injection in the hippocampal regions had improved cognition
112 despite of the persisting Aβ plaques and neurofibrillary tangles [379], which underscores the potential of stem cells. How HSC cellularity in the BM and peripheral blood is
+ changed upon P2Y2R deletion in Tg mice in relation to AD pathogenesis is currently being investigated. In addition, P2Y2R is also important for bone homeostasis through its role in the proliferation and maturation in addition to emigration of hematopoietic cells.
−/− Compared to wild type mice, P2Y2R mice exhibit a significant reduction in bone volume, thickness and stiffness at 17 weeks, and loss of trabecular bone mass and thickness at 8 weeks of age [256]. In vitro studies have showed that BM stromal cells released ATP through vesicular exocytosis in response to shear stress [211], which can be sensed by osteoblastic P2Y2R and induce stress actin formation [251]. The CXCL12-
CXCR4 axis is involved in estrogenic differentiation of mesenchymal stem cells [380,
381] and the regulation of osteoclasts [382, 383], and our finding that normal P2Y2R expression is necessary for CXCL12 expression in the BM (Figures III-4A and D) supports a role for P2Y2R in bone formation.
+ +/− + The BM levels of SCF are reduced in male Tg P2Y2R vs. Tg mice (Figure III-4B), similar to decreases in SCF levels in the BM of CXCL12-cKO mice [65], which is consistent with the conclusion that P2Y2R deletion reduces CXCL12-dependent SCF production. Activation of the c-Kit receptor by SCF is a principal factor in the maintenance of HSPC self-renewal and survival. For example, mice with a mutant c-Kit gene (KitW41/W41) exhibit reductions in long-term quiescent HSCs at steady state [64]. In addition, SCF can induce chemotaxis and chemokinesis of cultured HSPCs [104], and addition of SCF enhances G-CSF induced mobilization of HSPCs in vivo [105], and
CXCL12-induced chemotaxis of CD34+ HSCs by activating the same downstream
113 signaling pathways [106]. Moreover, BM ablation by 5-fluorouracil treatment was found to elevate plasma levels of both CXCL12 and SCF; the latter was thought to result from upregulation of metalloprotease-9 by CXCL12 [66]. Administration of SCF by subcutaneous injection increased circulating and splenic HSPCs, when CXCL12 production by osteoblasts was reduced in the BM of mice [88], a necessary step to initiate
HSPC mobilization in the BM as discussed above. Here we report the lower expression
+ +/− of SCF in the total BM cells of male Tg P2Y2R mice, yet how this abnormality affect the animals’ HSPC functions, including mobilization, is to be determined.
VCAM-1 is constitutively expressed in BM stromal cells like CAR cells [63] and sinusoidal endothelium [107] while in other tissues its expression level is upregulated only in response to inflammation [108]. Studies have shown that the interaction between
α4β1 integrin and VCAM-1 inhibits hematopoietic cell release from the BM. The immunostaining of VCAM-1 was greatly reduced in the BM during HSPC mobilization in responsive to G-CSF and SCF [109]. Two serine proteases, neutrophil elastase and cathepsin G were determined to be responsible for most cleavage of VCAM-1, which, however, was not indispensible for G-CSF-induced HSPC mobilization [109, 110]. Of note, the mobilization of HSPCs induced by anti-α4β1 integrin antibodies appears to require cooperative signaling by SCF and its receptor c-Kit [115]. On the other hand, adhesion molecules such as VCAM-1, ICAM-1 and VE-cadherin have been demonstrated to regulate HSPC transendothelial migration during homing from the peripheral blood to the BM [136]. It is also established that VCAM-1 in inflamed peripheral tissue is upregulated in the vascular endothelium to enable α4β1 integrin- expressing leukocytes to adhere to the endothelium, which precedes leukocyte
114 extravasation and migration to sites of inflammation [386]. In this scenario, it has been demonstrated, both in vivo and in vitro, that the P2Y2R upregulates VCAM-1 expression in vascular endothelial cells through a mechanism involving transactivation of VEGFR-2
[226, 365, 366]. P2Y2R activation not only upregulates the membrane-bound form of
VCAM-1, but also increases soluble VCAM-1 levels, as consequence of cleavage of membrane-bound VCAM-1 [366]. It has been postulated that soluble VCAM-1 acts as competitive feedback inhibitor of α4β1 integrin and membrane-bound VCAM-1 binding that regulates endothelial-leukocytes interaction [387-389], but recent evidence suggests that soluble VCAM-1 is also as a chemoattractant for monocytes, eosinophils, and some
T-cells [390-392]. It has been reported that shedding of VCAM-1 stimulated by phorbol
12-myristate 13-acetate (PMA) is mediated by ADAM17 (also known as tumor necrosis factor-α-converting enzyme, TACE) [393], a metalloprotease coupled to P2Y2R activation in cortical neurons, astrocytoma cells and epithelial cells [228, 230, 394].
Thus, P2Y2R likely regulates soluble VCAM-1 levels through activation of ADAM17 in addition to VCAM-1 mRNA levels. The reduction in BM expression of VCAM-1 in 10-
+ +/− + week old male Tg P2Y2R vs. Tg mice reported in the present study (Figure III-4B) indicates a role for P2Y2R in regulation of VCAM-1 in BM stromal cells, although further studies are needed to evaluate the participation of P2Y2R-mediated VCAM-1 signaling pathway in trafficking of HSPC and leukocytes in vivo.
When inflammation is initiated, mature leukocytes are mobilized from the BM and recruited in response to chemokines produced at the inflammatory site [395].
Accordingly, many studies have demonstrated the importance of the P2Y2R in leukocyte recruitment to damaged or infected tissues [237, 240, 242-244], although little attention
115 has been paid to the role of the P2Y2R in leukocyte emigration from the BM. Here, we demonstrate that suppression of P2Y2R expression in an AD mouse model prevents monocyte and neutrophil emigration from the BM to the circulation. To our knowledge, leukocyte counts in the peripheral blood have only been documented in one other study concerning the P2Y2R. Inoue et al. reported that in a sepsis model (cercal ligation and puncture, CLP), wild type mice maintained stable levels of leukocytes in the peripheral
−/− blood after CLP, whereas leukocyte levels in P2Y2R mice were elevated 2 and 4 h after
CLP, which were attributed to a decrease in circulating leukocyte recruitment to
−/− inflammatory sites (i.e., peritoneum and lungs) after CLP in P2Y2R mice [396].
Therefore, it is likely that the P2Y2R plays different roles in leukocyte mobilization in response to acute bacterial infection and chronic inflammation occurring in AD. Findings in this study suggest that the P2Y2R plays two important roles in bone marrow homeostasis during the development of AD: 1) control of leukocyte retention molecule
(CXCL12, SCF and VCAM-1) expression in bone marrow stromal cells and 2) control of leukocyte emigration from the BM by assisting in chemotaxis.
Materials and Methods
Reagents
The following reagents were purchased from Biolegend, San Diego, CA: RBC lysis buffer (10×), cell staining buffer, purified anti-mouse CD16/CD32 antibody, Alexa
Fluor® 488-labeled anti-mouse CD45 antibody, PE/Cy7-labeled anti-mouse/human
CD11b antibody, APC-labeled anti-mouse CD115 antibody and PE-labeled anti-mouse
Ly6G antibody. Rabbit anti-mouse CXCL12 polyclonal antibody was purchased from
116 Santa Cruz Biotechnology, Dallas, TX. All other reagents were purchased from Sigma-
Aldrich (St. Louis, MO) unless otherwise stated.
Mice
Animals were handled using protocols approved by the Institutional Animal Care and
Use Committee (IACUC) of the University of Missouri. Heterozygous TgCRND8 mice
(Tg+) that express human APP with Swedish and Indiana mutations [397] were obtained from Dr. David West away (University of Toronto) [397] and were maintained on a mixed C3H/C57BL/6 strain background. Non-transgenic littermate (Tg−) mice on the mixed C3H/C57BL/6 strain background that do not express human APP mutations were
−/− tm1Bhk used as negative controls for experiments. P2Y2R mice (B6.129P2-P2ry2 /J, stock
#009132) were purchased from Jackson Laboratories (Bar Harbor, ME) and were
+ −/− maintained on a C57BL/6 strain background. Tg mice were bred with P2Y2R mice to
+ +/− − +/− generate Tg P2Y2R heterozygotes and littermate Tg P2Y2R mice. All mice were bred at the Christopher S. Bond Life Sciences Center Animal Facility of the University of
Missouri, Columbia, MO. Animals were housed in vented cages with 12 h light/dark cycles and received food and water ad libitum. All mice were genotyped by PCR using appropriate primers (Integrated DNA Technologies, Coralville, IA), as described previously [397, 398]. Age-matched male mice were utilized for all experiments.
Peripheral Blood Collection and Plasma Preparation
Mice were euthanized by overdose of isoflurane inhalation, and peripheral blood was immediately collected by cardiac puncture with 25G needles and 1-mL syringes.
117 Collected blood was immediately transferred to Microvette® K3EDTA-coated tubes
(Sarstedt, North Rhine-Westphalia, Nümbrecht, Germany) for complete blood counts
(CBCs) or BD Microtainer® K2EDTA-coated tubes (BD Biosciences, Franklin Lakes,
NJ) for plasma preparation. To prepare plasma, the blood samples were centrifuged at
10,000 × g for 15 min at 4°C to completely remove platelets and the plasma supernatant was collected and frozen at -80 °C until use.
Complete Blood Count (CBC) Test
Complete blood counts (CBCs) were generated on an automated hematology instrument (Hemavet 950FS, Drew Scientific, Dallas, TX) validated for use in mice. For each blood sample, a smear was made, stained and a 100 white blood cell differential was performed. An inspection of samples for red and white cell morphological changes and platelet clumping was performed, and samples that were found with platelet clumps were excluded.
Tissue/Cell Collection
Spleen and cortex were dissected from mice and flash frozen immediately in liquid nitrogen and stored at -80°C until use. Cells in the BM were isolated from mouse femurs by truncating both ends of the bones and flushing the marrow cavity with cold PBS in syringes with 22G needles. For mRNA extraction, BM cells were pelleted by centrifugation at 400 × g for 7 min at 4°C. For flow cytometry, single-cell suspensions were made by repeated pipetting with a 1 mL pipet tip and passing through a 100 µm-
118 filter. Then the cells were then repelleted by centrifugation and cell pellets were flash frozen immediately in liquid nitrogen and stored at -80°C until use.
Flow Cytometry
Isolated BM cells were treated with 1× RBC lysis buffer on ice for 5 min, followed by washing twice with cell staining buffer. Cell counting was performed with a
Countess® automated cell counter (Life Technologies, Carlsbad, CA). Accordingly, cell suspension of 1×107 cells/mL was made with cell staining buffer, and 1×106 cells in 100
µL were incubated with 1.0 µg of purified anti-mouse CD16/CD32 antibody for 5-10 min on ice to block non-specific binding of immunoglobulin to Fc receptors prior to staining with fluorescently labeled antibodies for 15 min on ice. Finally, stained cells were washed twice with cell staining buffer by centrifugation at 350 × g for 5 min at 4°C, and resuspended in 0.5 mL of cell staining buffer, and analyzed by a LSRFortessa™ X-20 cell analyzer (BD Biosciences, Franklin Lakes, NJ).
Quantitative Real Time Polymerase Chain Reaction (RT-PCR)
To assess gene expression levels, total RNA extraction from tissues and RT-PCR analysis were performed as previously described [292]. Briefly, tissues or cell pellets were homogenized in TRI reagent using an IKA® T10 basic homogenizer (Wilmington,
NC), and mixed with formamide at a 5:1 ratio (by volume) by vortexing. After 5-15 min incubation at room temperature, the samples were centrifuged at 12,000 × g for 15 min at
4°C, followed by RNA extraction using a RNA RNeasy Mini Plus Kit (Qiagen, Hilden,
Germany) according to the manufacture’s instruction. cDNA was synthesized from 5 µg
119 of total RNA using RNA to cDNA EcoDry™ Premix (Clontech, Mountain View, CA) according to the manufacturer’s instructions. TaqMan probes for mouse P2Y2R,
CXCL12, SCF, VCAM-1, M-CSF, G-CSF, GM-CSF, VEGF, CCL2 and 18S rRNA endogenous control were obtained from Applied Biosystems® (Life Technologies,
Carlsbad, CA). The samples were prepared in duplicate and subjected to RT-PCR analysis on an Applied Biosystems® 7500 Real-Time PCR machine (Life Technologies).
Quantification of each target gene expression was normalized to 18S rRNA for each sample, and expressed as relative quantities (RQs) normalized to wild type (Tg−) samples.
Immunohistochemistry
Intact femurs were isolated, fixed in 4% (v/v) paraformaldehyde in PBS, decalcified in
Formic Acid-Sodium Citrate solution (1:1 mixture of 50% formic acid and 20% sodium citrate), and paraffin embedded for immunohistochemistry. Specific immunohistochemical detection of CXCL12 was performed with a rabbit anti-mouse
CXCL12 polyclonal antibody, visualized by the indirect peroxidase-antibody conjugate technique using a goat anti-rabbit IgG antibody. The sections were also counterstained with Mayer's hematoxylin. Images were obtained by Leica DM5500B microscope
(Wetzlar, Germany).
ELISA
Plasma prepared from mouse peripheral blood was subjected to ELISA using a mouse
CXCL12α ELISA kit from R&D systems (Minneapolis, MN) following the
120 manufacturer’s instruction. Experimental samples and standard samples with known concentrations of recombinant mouse CXCL12α were assayed in duplicates and data obtained represent the absorbance at 450 nm minus by the absorbance at 570 nm (for correction). The concentration of CXCL12α in experimental samples was calculated according to the standard curve.
Statistical Analysis
All experiments were independently performed at least 3 times with similar results. Data were processed in GraphPad Prism 5 software (GraphPad Software, La Jolla, CA) and expressed as mean ± SEM in each bar graph. Two-tail t test was used and statistical significance was reported when the P value was less than 0.05.
121 Table III-1. Cellularity in the peripheral blood. Peripheral blood of from 10-week old
− − +/− + + +/− Tg , Tg P2Y2R , Tg , and Tg P2Y2R mice was subjected to CBC test. The results are presented as mean ± SEM with the sample size (N) noted. Statistical analysis using t test was performed and significantly different P values between indicated groups are shown
(*P < 0.05; **P < 0.01; ***P < 0.001). WBC, white blood cell number (103/µL); LY#, lymphocyte number (103/µL); NE#, neutrophil number (103/µL); MO#, monocyte number (103/µL); EO, eosinophil number (103/µL); LY%, lymphocyte percentages (%);
NE%, neutrophil percentages (%); MO%, monocyte percentages (%); EO%, eosinophil percentages (%); RBC, red blood cell number (106/µL); PLT, platelet number (103/µL).
122 − − + + Tg Tg Tg Tg PAB PCD PAC PBD PAD +/− +/− (A) P2Y2R P2Y2R (B) (C) (D)
WBC 3.626 ± 2.736 ± 3.578 ± 3.273 ± 0.1066 0.6057 0.9357 0.3233 0.5229 0.3876 0.3688 0.4432 0.3862 N=16 N=16 N=13 N=16 LY# 3.105 ± 2.162 ± 2.557 ± 2.724 ± 0.0381 0.7188 0.2586 0.1906 0.4103 0.3329 0.2800 0.3305 0.3128 * N=16 N=16 N=13 N=16 NE# 0.4367 ± 0.4654 ± 0.9285 ± 0.5142 ± 0.8200 0.0401 0.0057 0.7539 0.5533 0.06550 0.1065 0.1635 0.1114 * ** N=16 N=16 N=13 N=16 MO# 0.0264 ± 0.03813 ± 0.05077 ± 0.01063 ± 0.4178 0.0262 0.2212 0.0237 0.1540 0.009667 0.01050 0.01806 0.004784 * * N=16 N=16 N=13 N=16 EO# 0.05803 ± 0.07188 ± 0.03489 ± 0.02438 ± 0.6348 0.5716 0.3566 0.0649 0.1430 0.01896 0.02176 0.01421 0.01187 N=16 N=16 N=13 N=16 LY% 85.50 ± 79.81 ± 71.46 ± 84.56 ± 0.0534 0.0030 0.0001 0.1977 0.7512 1.339 2.492 3.108 2.606 ** *** N=16 N=16 N=13 N=16 NE% 12.50 ± 16.63 ± 26.08 ± 14.50 ± 0.1307 0.0082 0.0003 0.5368 0.4841 1.313 2.307 3.292 2.498 ** *** N=16 N=16 N=13 N=16 MO% 0.6250 ± 1.250 ± 1.385 ± 0.3125 ± 0.1109 0.0172 0.1179 0.0083 0.2238 0.2213 0.3096 0.4463 0.1197 * ** N=16 N=16 N=13 N=16 EO% 1.375 ± 2.313 ± 1.077 ± 0.6250 ± 0.1934 0.2950 0.5794 0.0135 0.1022 0.3750 0.5966 0.3662 0.2394 * N=16 N=16 N=13 N=16 RBC 9.769 ± 9.897 ± 9.422 ± 9.632 ± 0.4903 0.3929 0.1240 0.2059 0.5311 0.1380 0.1195 0.1724 0.1689 N=15 N=14 N=13 N=13 PLT 836.2 ± 802.9 ± 871.0 ± 822.5 ± 0.5083 0.5571 0.6111 0.7608 0.8347 36.36 33.64 59.37 55.66 N=15 N=14 N=13 N=13
123 Figure III-1. Cellularity in the peripheral blood. Complete blood counts (CBCs) were
− − +/− + performed on peripheral blood collected from 10-week old male Tg , Tg P2Y2R , Tg ,
+ +/− and Tg P2Y2R mice. The number of total white blood cells (WBCs) (A), neutrophils
(B) and monocytes (C) are presented as means ± SEM. Statistical analysis was performed using the t test, where P values < 0.05 indicate significant differences (*P < 0.05; **P <
0.01; N = 13-16 per group).
124 A
B
C
125 Figure III-2. Cellularity in the bone marrow. BM cells were isolated from 10-week old
− − +/− + + +/− Tg , Tg P2Y2R , Tg , and Tg P2Y2R mice. After lysis of the RBCs, the BM cells were stained with the indicated fluorescently labeled antibodies against specific cell markers and subjected to flow cytometry. (A) Representative figures of the gating of
CD45+ leukocytes, CD45+Ly6G+CD11b+ mature neutrophils, CD45+Ly6G−CD115+ monocytes (FSC, forward scatter; SCC, side scatter). (B-C) Statistical bar graphs of the percentages of (B) CD45+Ly6G+CD11b+ mature neutrophils and (C)
CD45+Ly6G−CD115+ monocytes in the total BM cells are presented as mean ± SEM.
Statistical analysis was performed using the t test, where P values < 0.05 indicate significant differences between groups (*P < 0.05; **P < 0.01; N = 8-14 per group).
126 A
SSC SSC
FSC CD45-AF488
PE
- PE - Ly6G Ly6G CD11b-PE/Cy7 CD115-APC B
127 C
128 Figure III-3. Marginated pools of neutrophils and monocytes in the spleen. Spleens
− − +/− + + +/− were isolated from 10-week old Tg , Tg P2Y2R , Tg , and Tg P2Y2R mice and weighed. (A) Bar graph of the wet weight of spleens over the total body weight is presented as mean ± SEM (N = 9-18 per group). (B and C) Spleen cells subjected to RT-
PCR analysis for (B) neutrophil elastase and (C) AIF-1 levels, as indicators of the splenic pools of neutrophils and monocytes, respectively. Relative quantities (RQ) are presented as mean ± SEM. Statistical analysis was performed using the t test, where P values < 0.05 indicate significant differences between groups (*P < 0.05; ** P < 0.01; N = 10-12 per group).
129 A
B
C
130 Figure III-4. Expression levels of CXCL12, SCF, and VCAM-1 in the bone marrow.
− − +/− + (A-C) BM cells were isolated from 10-week old Tg , Tg P2Y2R , Tg , and
+ +/− Tg P2Y2R mice, followed by total RNA extraction and RT-PCR analysis for (A)
CXCL12, (B) SCF and (C) VCAM-1 expression. Bar graphs of the relative quantities
(RQ) of CXCL12, SCF and vCAM-1 mRNAs are presented as mean ± SEM. Statistical analysis was performed using the t test, where P values < 0.05 indicate significant differences (*P < 0.05, **P < 0.01; ***P < 0.001; N = 10 to 12 per group). (D) Femurs
+ + +/− of 10-week old Tg and Tg P2Y2R mice were subjected to IHC staining for CXCL12
(brown) with hematoxylin counter staining (purple; N = 3 per group). A representative
+ + +/− view of a femur of a Tg (left image) and Tg P2Y2R mouse (right image) is presented.
Scale bar in the left image is 100 µm.
131 A
B
C
132 D
133 Figure III-5. CXCL12 levels in the peripheral blood, cortex and spleen. (A) Plasma
− − +/− + was prepared from the peripheral blood of 10-week old Tg , Tg P2Y2R , Tg , and
+ +/− Tg P2Y2R mice and CXCL12α levels in the peripheral blood were measured by
ELISA. Bar graph of the calculated CXCL12α levels is presented as mean ± SEM (N = 5 to 6 per group). (B and C) The (B) cerebral cortex or (C) spleen was isolated from 10-
− − +/− + + +/− week old Tg , Tg P2Y2R , Tg , and Tg P2Y2R mice, followed by total RNA extraction and RT-PCR analysis for CXCL12 expression. Bar graphs of the relative quantities (RQ) of CXCL12 are presented as mean ± SEM. Statistical analysis was performed using the t test, where P values < 0.05 indicate significant differences between groups (*P < 0.05; ns = non-significant; N = 5-10 per group).
134 A
B
C
135 Figure III-6. Expression levels of M-CSF in the bone marrow and cortex. (A) BM
− − +/− + cells or (B) cerebral cortex was isolated from 10-week old Tg , Tg P2Y2R , Tg , and
+ +/− Tg P2Y2R mice, followed by total RNA extraction and RT-PCR analysis for M-CSF.
Bar graphs of the relative quantities (RQ) of M-CSF mRNA are presented as mean ±
SEM. Statistical analysis was performed using the t test, where no P values between groups were below 0.05 (N = 5-11 per group).
136 A
B
137 Figure III-7. Expression level of CCL2 in the cortex. Cortex was isolated from 10-
− − +/− + + +/− week old Tg , Tg P2Y2R , Tg , and Tg P2Y2R mice, followed by total RNA extraction and RT-PCR analysis for CCL2 mRNA. Bar graphs of the relative quantities
(RQ) of CCL2 mRNA are presented as mean ± SEM. Statistical analysis was performed using the t test, where no P values between groups were below 0.05 but marginal differences were noted with the P values (ns = non-significant; N = 6 per group).
138
139 Chapter IV
Summary and Future Directions
A. Summary
Inflammation is prevalent in mammalian diseases and fine-tuned inflammatory responses are crucial for a beneficial outcome. In a typical inflammatory response, when a microbial or aseptic inflammatory stimulus is present, resident immune cells sense and migrate towards the chemoattractants (i.e. chemotaxis) and release cytokines and chemokines to recruit circulating leukocytes to the inflammatory site if the initial inflammatory response is not resolved. For this process, leukocytes must transmigrate across the vascular barrier where endothelial cells are activated to enhance leukocyte attachment by upregulation of adhesion molecules, such as VCAM-1 and ICAM-1. In addition, interendothelial adhesion junctions are weakened, through phosphorylation of junctional proteins and rearrangement of the actin cytoskeleton to form a “leaky” state.
To increase numbers of circulating leukocytes, stromal cells and leukocytes in the BM respond to inflammatory mediators to promote leukocyte production and emigration from the BM to the peripheral blood. The importance of the P2Y2R in leukocyte infiltration of peripheral tissues has been highlighted in several in vivo studies using different models of inflammation [237, 240, 242, 243]. Some of the mechanisms underlying P2Y2R-mediated inflammatory responses have been determined, including autocrine activation of the
P2Y2R in migrating leukocytes to enhance chemotaxis towards chemoattractants [237-
239], P2Y2R-mediated release of vasodilators from endothelial cells to induce
140 vasodilation [264, 278] and upregulation of VCAM-1 expression to facilitates binding of leukocytes to endothelial cells [226, 227]. In addition, the P2Y2R in HSPCs is likely the mediator of ATP/UTP induced promotion of hematopoiesis in synergy with hematopoietic cytokines [247]. The present work reveals a novel role of endothelial
P2Y2R to interact with VE-cadherin and activate Rac1, a pathway that modulates vascular barrier function to increase permeability of macromolecules and leukocytes. The work completed also demonstrates the role of the P2Y2R in leukocyte emigration from the BM to peripheral blood likely by regulating the stromal cell expression of CXCL12,
SCF, and VCAM-1, and leukocyte chemotaxis.
B. Future Directions
1. Identify Novel P2Y2R-mediated Signaling in Endothelial Cells
We show here that in endothelial cells the P2Y2R interacts directly with VE-cadherin and VEGFR-2 to induce VE-cadherin phosphorylation and Rac1 activation, responses known to loosen adherens junctions and rearrange cytoskeleton in response to thrombin and histamine [348], which facilitates transendothelial leukocyte migration [349]. A recent in vivo study showed that phosphorylation of different tyrosine residues of VE- cadherin control leukocyte extravasation (Tyr731) and vascular permeability (Tyr685), and dephosphorylation of VE-cadherin at Tyr731 in cultured endothelial cells is triggered by leukocytes via the activation of the tyrosine phosphatase SHP-2, while the tyrosine phosphatase VE-PTP selectively dephosphorylates of Tyr685 [399]. Likewise, P2Y2R- induced phosphorylation of VE-cadherin should be further investigated to determine which residues of VE-cadherin are phosphorylated using antibodies that bind specific
141 phosphotyrosine residues in VE-cadherin, and whether the phosphatases SHP-2 and VE-
PTP are involved in VE-cadherin dephosphorylation in endothelial cells using specific siRNA’s. Moreover, future experiments are required to which of the currently known
P2Y2R signaling pathways regulate leukocyte extravasation and vascular permeability.
Our preliminary data utilized an in vitro transmigration assay with a modified Boyden chamber [400] and a monocytic cell line and human endothelial cells monolayer formed between the two chambers. Results obtained indicate that UTP promotes monocyte transendothelial migration (TEM), which is significantly reduced when the expression of
P2Y2R is inhibited by siRNA in the endothelial cells (data not shown). In another in vitro assay, the transendothelial resistance of an endothelial monolayer, a measure of barrier function, was reduced by UTP treatment (data not shown). These experiments should be repeated with endothelial cells transfected with dominant negative Rac1 (T17NRac1) to determine the role of P2Y2R-induced Rac activation in these physiological events.
2. Delineate the Role of Endothelial P2Y2R In Vivo
We also have evaluated the vascular permeability of adult mice during steady state,
−/− and found the vascular permeability in spleen, colon and kidney in the P2Y2R mice was significantly lower than in age-matched wild type (WT) mice, indicating a role for the P2Y2R in the maintenance of baseline permeability of the vasculature in these organs, probably triggered by ATP release from red blood cells and endothelial cells in response to blood flow-induced shear stress [208, 210]. The investigation on the regulation of vascular permeability by the P2Y2R should be extended to other inflammatory
142 −/− conditions, i.e., to assess vascular permeability in P2Y2R vs. WT mice under different inflammatory conditions, such as chemically-induced intestinal inflammation [401].
Although many in vivo studies have shown that the P2Y2R is critical for leukocyte recruitment under inflammatory conditions, none of those studies clearly distinguished the roles of leukocytic P2Y2R and endothelial P2Y2Rs. We hypothesize that the endothelial P2Y2R is required for the regulation of vascular permeability and leukocyte infiltration during inflammation. To evaluate this hypothesis, mice with conditional (c) knockout of P2Y2R in endothelium (e) (eP2Y2R-cKO) could be generated by crossing
flox/flox P2Y2R mice that we possess with Tie2-Cre mice in which expression of Cre is under the control of the endothelium-specific Tie2 promoter. These eP2Y2R-cKO mice will be a powerful tool for in vivo studies of the endothelial P2Y2R. For example, a mild
−/− dose of LPS could be injected intraperitoneally into the eP2Y2R-cKO, P2Y2R and wild type (WT) mice, followed by collection and numeration of the peritoneal leukocytes. If the hypothesis is true, the number of peritoneal leukocytes in eP2Y2R-cKO mice will be
−/− less than WT mice, and the comparison between eP2Y2R-cKO and P2Y2R will indicate the extent to which the leukocytic P2Y2R contributes to leukocyte infiltration. Of note, we found in the TgCRND8 mouse model of AD that neutrophil and monocyte mobilization was impaired when P2Y2R expression was decreased. Therefore, it will be worth comparing whether the leukocyte populations in the BM and the circulation are
−/− altered in eP2Y2R-cKO and P2Y2R mice to further establish the role of this receptor in the regulation of vascular function.
143 3. Confirming the Role of the P2Y2R in Promoting Leukocyte Emigration from the
BM
The role of the P2Y2R in promoting leukocyte emigration from the BM to the circulation has never been addressed, and there are only a few in vitro studies that demonstrating that ATP/UTP stimulates proliferation and differentiation [247, 248], and synergizes with CXCL12 to induce chemotaxis of HSPCs [249, 250]. Here, we have provided in vivo evidence for the role of the P2Y2R in neutrophil and monocyte mobilization from BM using TgCRND8 (Tg+) mice, an AD model. The expression levels of CXCL12, SCF, and VCAM-1, which have been generally considered to be leukocyte retention signals, were determined to be significantly reduced in the BM cells from
+ +/− + Tg P2Y2R mice compared with Tg mice. We also found that UTP treatment increased expression of CXCL12, SCF, and VCAM-1 in isolated primary BM stromal cells from a
WT mouse, while Aβ1-42 decreased CXCL12 and SCF but increased VCAM-1 expression
(preliminary data not shown). These experiments should be repeated with cells from
−/− P2Y2R mice to determine the role of P2Y2R in modulating the expression of CXCL12,
+ + +/− SCF and VCAM-1 in BM stromal cells. Femur sections from Tg and Tg P2Y2R could also be subjected to immunostaining for CXCL12, SCF, and VCAM-1 and specific cell markers for different cell types in the BM to illustrate the type and distribution of the cells that are regulated by the P2Y2R. The Boyden chamber assay also could be
−/− performed using neutrophils and monocytes isolated from WT and P2Y2R mice to evaluate their chemotaxis/chemokinesis. Our preliminary data show that in the IL-14α overexpressing mouse model of autoimmunity characterized by increased serum levels of immunoglobulins, infiltration of the salivary glands with lymphocytes, and the
144 development of lymphoma [402], loss of P2Y2R expression significantly reduces infiltration of lymphocytes into salivary glands as well as the level of circulating lymphocytes (data not shown). Therefore, the P2Y2R is also likely to regulate lymphocyte mobilization from the BM to circulation in these mice, and similar experiments including assessment of the lymphocyte population in the BM and their mobility in a chemotactic gradient could be performed to prove this speculation.
4. Further Investigate the Role of the P2Y2R in AD
Circulating HSCs have been suggested to contribute to regeneration of brain tissues through transdifferentiation, and early AD patients were found to have decreased circulating CD34+ HSCs compared to controls [191]. Surprisingly, neural stem cell injection into the hippocampal regions of an AD mouse model improved cognition despite the persisting Aβ plaques and neurofibrillary tangles [379], which underscored the great potential of stem cells in the treatment of disease. Our preliminary data indicated an increase in the numbers of LSK (Lineage−Sca-1+c-Kit+) HSPCs in the BM of
+ +/− Tg P2Y2R vs. WT mice (data not shown), which possibly results from their impaired emigration like neutrophils and monocytes due to loss of the P2Y2R. The peripheral
+ + +/− blood of Tg and Tg P2Y2R mice should be collected and analyzed by flow cytometry to enumerate HSPCs in circulation. Furthermore, the endothelial P2Y2R could mediate vasodilation to counter Aβ-induced vasoconstriction [198, 264, 278, 280]. We also
+ + +/− noticed that compared to Tg mice, Tg P2Y2R mice have higher cortical and hippocampal expression of VEGF (data not shown), a typical response to hypoxia since
VEGF dilates microvessels by inducing production of nitric oxide (NO) and prostacyclin
145 (PGI2) [403, 404]. Besides, VEGF was reported to inhibit leukocyte rolling and adherence on and transmigrate cross the endothelium [405], whereas VEGF in the plasma is also able to mobilize HSCs as well as vasculogenic stem cells by activating VEGFR-2
+ +/− [374]. Hence, how vascular dysfunction in the brain of Tg P2Y2R mice contributes to the accelerated AD progress is an interesting question to be answered in future studies.
− − +/− + CBC analysis of the blood of 10-week old male Tg , Tg P2Y2R , Tg , and
+ +/− Tg P2Y2R mice showed that all genotypes have similar platelet numbers, however,
+ +/− − Tg P2Y2R mice had significantly larger mean platelet volume (MPV) than Tg and
− +/− + Tg P2Y2R mice, and were marginally larger than Tg mice (data not shown). The elevation of MPV has been found in cardio- and cerebrovascular disorders and low-grade inflammatory conditions, and a correlation between increased MPV and the risk of thrombosis has been suggested [406]. Larger platelets contain more granules and produce greater amounts of vasoactive and prothrombotic factors including ATP, aggregate more rapidly and express a greater number of adhesion molecules [407]. It has been recently noticed that AD patients also have increased MPV compared to controls [408]. A major component of blood clots, fibrinogen, has been found to interact with Aβ and form abnormal clots that are resistant to degradation, which contributes to cerebral amyloid atrophy (CAA); depletion of fibrinogen or an inhibitor of Aβ-fibrinogen interaction reduced CAA and cognitive decline [409, 410]. CXCL12 has been shown to be a critical regulator of thrombopoiesis and mobilization [411, 412]. Therefore, it seems worth investigating the contribution of platelet-related factors to the accelerated AD pathology
+ +/− and premature death of Tg P2Y2R mice.
146 REFERENCES
1. Winsauer, G. and R. de Martin, Resolution of inflammation: intracellular
feedback loops in the endothelium. Thromb Haemost, 2007. 97(3): p. 364-9.
2. Pawelec, G., D. Goldeck, and E. Derhovanessian, Inflammation, ageing and
chronic disease. Curr Opin Immunol, 2014. 29: p. 23-8.
3. Borish, L.C. and J.W. Steinke, 2. Cytokines and chemokines. J Allergy Clin
Immunol, 2003. 111(2 Suppl): p. S460-75.
4. Griffith, J.W., C.L. Sokol, and A.D. Luster, Chemokines and chemokine
receptors: positioning cells for host defense and immunity. Annu Rev Immunol,
2014. 32: p. 659-702.
5. Wu, D., G.J. LaRosa, and M.I. Simon, G protein-coupled signal transduction
pathways for interleukin-8. Science, 1993. 261(5117): p. 101-3.
6. Thelen, M. and J.V. Stein, How chemokines invite leukocytes to dance. Nat
Immunol, 2008. 9(9): p. 953-9.
7. Pasvolsky, R., et al., A LAD-III syndrome is associated with defective expression
of the Rap-1 activator CalDAG-GEFI in lymphocytes, neutrophils, and platelets. J
Exp Med, 2007. 204(7): p. 1571-82.
8. Bardi, G., V. Niggli, and P. Loetscher, Rho kinase is required for CCR7-mediated
polarization and chemotaxis of T lymphocytes. FEBS Lett, 2003. 542(1-3): p. 79-
83.
9. Fukui, Y., et al., Haematopoietic cell-specific CDM family protein DOCK2 is
essential for lymphocyte migration. Nature, 2001. 412(6849): p. 826-31. 147 10. Nombela-Arrieta, C., et al., A central role for DOCK2 during interstitial
lymphocyte motility and sphingosine-1-phosphate-mediated egress. J Exp Med,
2007. 204(3): p. 497-510.
11. Colvin, R.A., et al., Synaptotagmin-mediated vesicle fusion regulates cell
migration. Nat Immunol, 2010. 11(6): p. 495-502.
12. Bloes, D.A., D. Kretschmer, and A. Peschel, Enemy attraction: bacterial agonists
for leukocyte chemotaxis receptors. Nat Rev Microbiol, 2015. 13(2): p. 95-104.
13. Kawai, T. and S. Akira, Toll-like receptors and their crosstalk with other innate
receptors in infection and immunity. Immunity, 2011. 34(5): p. 637-50.
14. Schromm, A.B., et al., Molecular genetic analysis of an endotoxin nonresponder
mutant cell line: a point mutation in a conserved region of MD-2 abolishes
endotoxin-induced signaling. J Exp Med, 2001. 194(1): p. 79-88.
15. Sogawa, Y., et al., Inhibition of neutrophil migration in mice by mouse formyl
peptide receptors 1 and 2 dual agonist: indication of cross-desensitization in vivo.
Immunology, 2011. 132(3): p. 441-50.
16. Brown, A.J., et al., The Orphan G protein-coupled receptors GPR41 and GPR43
are activated by propionate and other short chain carboxylic acids. J Biol Chem,
2003. 278(13): p. 11312-9.
17. Le Poul, E., et al., Functional characterization of human receptors for short chain
fatty acids and their role in polymorphonuclear cell activation. J Biol Chem,
2003. 278(28): p. 25481-9.
18. Mehta, D. and A.B. Malik, Signaling mechanisms regulating endothelial
permeability. Physiol Rev, 2006. 86(1): p. 279-367.
148 19. Liebner, S., U. Cavallaro, and E. Dejana, The multiple languages of endothelial
cell-to-cell communication. Arterioscler Thromb Vasc Biol, 2006. 26(7): p. 1431-
8.
20. Vincent, P.A., et al., VE-cadherin: adhesion at arm's length. Am J Physiol Cell
Physiol, 2004. 286(5): p. C987-97.
21. Noren, N.K., et al., p120 catenin regulates the actin cytoskeleton via Rho family
GTPases. J Cell Biol, 2000. 150(3): p. 567-80.
22. Anastasiadis, P.Z., et al., Inhibition of RhoA by p120 catenin. Nat Cell Biol, 2000.
2(9): p. 637-44.
23. Grosheva, I., et al., p120 catenin affects cell motility via modulation of activity of
Rho-family GTPases: a link between cell-cell contact formation and regulation of
cell locomotion. J Cell Sci, 2001. 114(Pt 4): p. 695-707.
24. Balda, M.S. and K. Matter, Tight junctions at a glance. J Cell Sci, 2008. 121(Pt
22): p. 3677-82.
25. Wittchen, E.S., Endothelial signaling in paracellular and transcellular leukocyte
transmigration. Front Biosci (Landmark Ed), 2009. 14: p. 2522-45.
26. Luscinskas, F.W., et al., Leukocyte transendothelial migration: a junctional
affair. Semin Immunol, 2002. 14(2): p. 105-13.
27. Wittchen, E.S., J. Haskins, and B.R. Stevenson, Protein interactions at the tight
junction. Actin has multiple binding partners, and ZO-1 forms independent
complexes with ZO-2 and ZO-3. J Biol Chem, 1999. 274(49): p. 35179-85.
28. Tornavaca, O., et al., ZO-1 controls endothelial adherens junctions, cell-cell
tension, angiogenesis, and barrier formation. J Cell Biol, 2015. 208(6): p. 821-38.
149 29. Birukova, A.A., et al., Afadin controls p120-catenin-ZO-1 interactions leading to
endothelial barrier enhancement by oxidized phospholipids. J Cell Physiol, 2012.
227(5): p. 1883-90.
30. Simon, A.M. and D.A. Goodenough, Diverse functions of vertebrate gap
junctions. Trends Cell Biol, 1998. 8(12): p. 477-83.
31. Spindler, V., N. Schlegel, and J. Waschke, Role of GTPases in control of
microvascular permeability. Cardiovasc Res, 2010. 87(2): p. 243-53.
32. Leick, M., et al., Leukocyte recruitment in inflammation: basic concepts and new
mechanistic insights based on new models and microscopic imaging technologies.
Cell Tissue Res, 2014. 355(3): p. 647-56.
33. Phillipson, M., et al., Intraluminal crawling of neutrophils to emigration sites: a
molecularly distinct process from adhesion in the recruitment cascade. J Exp
Med, 2006. 203(12): p. 2569-75.
34. Yang, L., et al., ICAM-1 regulates neutrophil adhesion and transcellular
migration of TNF-α-activated vascular endothelium under flow. Blood, 2005.
106(2): p. 584-92.
35. Ferreira, A.M., et al., Interleukin-1β reduces transcellular monocyte diapedesis
and compromises endothelial adherens junction integrity. Microcirculation, 2005.
12(7): p. 563-79.
36. Phillipson, M., et al., Endothelial domes encapsulate adherent neutrophils and
minimize increases in vascular permeability in paracellular and transcellular
emigration. PLoS One, 2008. 3(2): p. e1649.
150 37. Yin, T. and L. Li, The stem cell niches in bone. J Clin Invest, 2006. 116(5): p.
1195-201.
38. Rieger, M.A. and T. Schroeder, Hematopoiesis. Cold Spring Harb Perspect Biol,
2012. 4(12).
39. Osawa, M., et al., Long-term lymphohematopoietic reconstitution by a single
CD34-low/negative hematopoietic stem cell. Science, 1996. 273(5272): p. 242-5.
40. Adolfsson, J., et al., Upregulation of Flt3 expression within the bone marrow
Lin−Sca1+c-kit+ stem cell compartment is accompanied by loss of self-renewal
capacity. Immunity, 2001. 15(4): p. 659-69.
41. Christensen, J.L. and I.L. Weissman, Flk-2 is a marker in hematopoietic stem cell
differentiation: a simple method to isolate long-term stem cells. Proc Natl Acad
Sci U S A, 2001. 98(25): p. 14541-6.
42. Yang, L., et al., Identification of Lin−Sca1+kit+CD34+Flt3− short-term
hematopoietic stem cells capable of rapidly reconstituting and rescuing
myeloablated transplant recipients. Blood, 2005. 105(7): p. 2717-23.
43. Morita, Y., H. Ema, and H. Nakauchi, Heterogeneity and hierarchy within the
most primitive hematopoietic stem cell compartment. J Exp Med, 2010. 207(6): p.
1173-82.
44. Kiel, M.J., et al., SLAM family receptors distinguish hematopoietic stem and
progenitor cells and reveal endothelial niches for stem cells. Cell, 2005. 121(7):
p. 1109-21.
45. Kitamura, Y. and A. Ito, Mast cell-committed progenitors. Proc Natl Acad Sci U
S A, 2005. 102(32): p. 11129-30.
151 46. Wilson, A. and A. Trumpp, Bone-marrow haematopoietic-stem-cell niches. Nat
Rev Immunol, 2006. 6(2): p. 93-106.
47. Zhang, J., et al., Identification of the haematopoietic stem cell niche and control
of the niche size. Nature, 2003. 425(6960): p. 836-41.
48. Calvi, L.M., et al., Osteoblastic cells regulate the haematopoietic stem cell niche.
Nature, 2003. 425(6960): p. 841-6.
49. Chitteti, B.R., et al., Impact of interactions of cellular components of the bone
marrow microenvironment on hematopoietic stem and progenitor cell function.
Blood, 2010. 115(16): p. 3239-48.
50. Visnjic, D., et al., Hematopoiesis is severely altered in mice with an induced
osteoblast deficiency. Blood, 2004. 103(9): p. 3258-64.
51. Visnjic, D., et al., Conditional ablation of the osteoblast lineage in Col2.3Δtk
transgenic mice. J Bone Miner Res, 2001. 16(12): p. 2222-31.
52. Taichman, R.S. and S.G. Emerson, Human osteoblasts support hematopoiesis
through the production of granulocyte colony-stimulating factor. J Exp Med,
1994. 179(5): p. 1677-82.
53. Taichman, R., et al., Hepatocyte growth factor is secreted by osteoblasts and
cooperatively permits the survival of haematopoietic progenitors. Br J Haematol,
2001. 112(2): p. 438-48.
54. Arai, F., et al., Tie2/angiopoietin-1 signaling regulates hematopoietic stem cell
quiescence in the bone marrow niche. Cell, 2004. 118(2): p. 149-61.
55. Stier, S., et al., Osteopontin is a hematopoietic stem cell niche component that
negatively regulates stem cell pool size. J Exp Med, 2005. 201(11): p. 1781-91.
152 56. Taichman, R.S., Blood and bone: two tissues whose fates are intertwined to
create the hematopoietic stem-cell niche. Blood, 2005. 105(7): p. 2631-9.
57. Zhu, J., et al., Osteoblasts support B-lymphocyte commitment and differentiation
from hematopoietic stem cells. Blood, 2007. 109(9): p. 3706-12.
58. Ma, Y.D., et al., Defects in osteoblast function but no changes in long-term
repopulating potential of hematopoietic stem cells in a mouse chronic
inflammatory arthritis model. Blood, 2009. 114(20): p. 4402-10.
59. Hooper, A.T., et al., Engraftment and reconstitution of hematopoiesis is
dependent on VEGFR2-mediated regeneration of sinusoidal endothelial cells.
Cell Stem Cell, 2009. 4(3): p. 263-74.
60. Salter, A.B., et al., Endothelial progenitor cell infusion induces hematopoietic
stem cell reconstitution in vivo. Blood, 2009. 113(9): p. 2104-7.
61. Li, B., et al., Endothelial cells mediate the regeneration of hematopoietic stem
cells. Stem Cell Res, 2010. 4(1): p. 17-24.
62. Sugiyama, T., et al., Maintenance of the hematopoietic stem cell pool by
CXCL12-CXCR4 chemokine signaling in bone marrow stromal cell niches.
Immunity, 2006. 25(6): p. 977-88.
63. Omatsu, Y., et al., The essential functions of adipo-osteogenic progenitors as the
hematopoietic stem and progenitor cell niche. Immunity, 2010. 33(3): p. 387-99.
64. Thoren, L.A., et al., Kit regulates maintenance of quiescent hematopoietic stem
cells. J Immunol, 2008. 180(4): p. 2045-53.
65. Tzeng, Y.S., et al., Loss of Cxcl12/Sdf-1 in adult mice decreases the quiescent
state of hematopoietic stem/progenitor cells and alters the pattern of
153 hematopoietic regeneration after myelosuppression. Blood, 2011. 117(2): p. 429-
39.
66. Heissig, B., et al., Recruitment of stem and progenitor cells from the bone marrow
niche requires MMP-9 mediated release of kit-ligand. Cell, 2002. 109(5): p. 625-
37.
67. Chotinantakul, K. and W. Leeanansaksiri, Hematopoietic stem cell development,
niches, and signaling pathways. Bone Marrow Res, 2012. 2012: p. 270425.
68. Burgess, A.W. and D. Metcalf, The nature and action of granulocyte-macrophage
colony stimulating factors. Blood, 1980. 56(6): p. 947-58.
69. Hamilton, J.A., Colony-stimulating factors in inflammation and autoimmunity.
Nat Rev Immunol, 2008. 8(7): p. 533-44.
70. Ryan, G.R., et al., Rescue of the colony-stimulating factor 1 (CSF-1)-nullizygous
mouse (Csf1 op/Csf1op) phenotype with a CSF-1 transgene and identification of
sites of local CSF-1 synthesis. Blood, 2001. 98(1): p. 74-84.
71. Cecchini, M.G., et al., Role of colony stimulating factor-1 in the establishment
and regulation of tissue macrophages during postnatal development of the mouse.
Development, 1994. 120(6): p. 1357-72.
72. Stanley, E.R., et al., Biology and action of colony--stimulating factor-1. Mol
Reprod Dev, 1997. 46(1): p. 4-10.
73. Eto, T., et al., Effects of macrophage colony-stimulating factor (M-CSF) on the
mobilization of peripheral blood stem cells. Bone Marrow Transplant, 1994.
13(2): p. 125-9.
154 74. Minamino, K., et al., Macrophage colony-stimulating factor (M-CSF), as well as
granulocyte colony-stimulating factor (G-CSF), accelerates neovascularization.
Stem Cells, 2005. 23(3): p. 347-54.
75. Hamilton, J.A. and G.P. Anderson, GM-CSF Biology. Growth Factors, 2004.
22(4): p. 225-31.
76. Semerad, C.L., et al., G-CSF is an essential regulator of neutrophil trafficking
from the bone marrow to the blood. Immunity, 2002. 17(4): p. 413-23.
77. Pelus, L.M., et al., Peripheral blood stem cell mobilization. A role for CXC
chemokines. Crit Rev Oncol Hematol, 2002. 43(3): p. 257-75.
78. Jaiswal, S. and I.L. Weissman, Hematopoietic stem and progenitor cells and the
inflammatory response. Ann N Y Acad Sci, 2009. 1174: p. 118-21.
79. Baldridge, M.T., K.Y. King, and M.A. Goodell, Inflammatory signals regulate
hematopoietic stem cells. Trends Immunol, 2011. 32(2): p. 57-65.
80. Wright, D.E., et al., Physiological migration of hematopoietic stem and
progenitor cells. Science, 2001. 294(5548): p. 1933-6.
81. Hidalgo, A., et al., Chemokine stromal cell-derived factor-1α modulates VLA-4
integrin-dependent adhesion to fibronectin and VCAM-1 on bone marrow
hematopoietic progenitor cells. Exp Hematol, 2001. 29(3): p. 345-55.
82. Semerad, C.L., et al., G-CSF potently inhibits osteoblast activity and CXCL12
mRNA expression in the bone marrow. Blood, 2005. 106(9): p. 3020-7.
83. Katayama, Y., et al., Signals from the sympathetic nervous system regulate
hematopoietic stem cell egress from bone marrow. Cell, 2006. 124(2): p. 407-21.
155 84. Christopher, M.J. and D.C. Link, Granulocyte colony-stimulating factor induces
osteoblast apoptosis and inhibits osteoblast differentiation. J Bone Miner Res,
2008. 23(11): p. 1765-74.
85. Eash, K.J., et al., CXCR4 is a key regulator of neutrophil release from the bone
marrow under basal and stress granulopoiesis conditions. Blood, 2009. 113(19):
p. 4711-9.
86. Martin, C., et al., Chemokines acting via CXCR2 and CXCR4 control the release
of neutrophils from the bone marrow and their return following senescence.
Immunity, 2003. 19(4): p. 583-93.
87. Ma, Q., D. Jones, and T.A. Springer, The chemokine receptor CXCR4 is required
for the retention of B lineage and granulocytic precursors within the bone
marrow microenvironment. Immunity, 1999. 10(4): p. 463-71.
88. Christopher, M.J., et al., Suppression of CXCL12 production by bone marrow
osteoblasts is a common and critical pathway for cytokine-induced mobilization.
Blood, 2009. 114(7): p. 1331-9.
89. Levesque, J.P., et al., Disruption of the CXCR4/CXCL12 chemotactic interaction
during hematopoietic stem cell mobilization induced by GCSF or
cyclophosphamide. J Clin Invest, 2003. 111(2): p. 187-96.
90. McQuibban, G.A., et al., Matrix metalloproteinase activity inactivates the CXC
chemokine stromal cell-derived factor-1. J Biol Chem, 2001. 276(47): p. 43503-8.
91. Christopherson, K.W., 2nd, S. Cooper, and H.E. Broxmeyer, Cell surface
peptidase CD26/DPPIV mediates G-CSF mobilization of mouse progenitor cells.
Blood, 2003. 101(12): p. 4680-6.
156 92. Petit, I., et al., G-CSF induces stem cell mobilization by decreasing bone marrow
SDF-1 and up-regulating CXCR4. Nat Immunol, 2002. 3(7): p. 687-94.
93. Pelus, L.M., et al., The CXCR4 agonist peptide, CTCE-0021, rapidly mobilizes
polymorphonuclear neutrophils and hematopoietic progenitor cells into
peripheral blood and synergizes with granulocyte colony-stimulating factor. Exp
Hematol, 2005. 33(3): p. 295-307.
94. Broxmeyer, H.E., et al., Rapid mobilization of murine and human hematopoietic
stem and progenitor cells with AMD3100, a CXCR4 antagonist. J Exp Med, 2005.
201(8): p. 1307-18.
95. Liles, W.C., et al., Mobilization of hematopoietic progenitor cells in healthy
volunteers by AMD3100, a CXCR4 antagonist. Blood, 2003. 102(8): p. 2728-30.
96. Devine, S.M., et al., Rapid mobilization of CD34+ cells following administration
of the CXCR4 antagonist AMD3100 to patients with multiple myeloma and non-
Hodgkin's lymphoma. J Clin Oncol, 2004. 22(6): p. 1095-102.
97. Flomenberg, N., et al., The use of AMD3100 plus G-CSF for autologous
hematopoietic progenitor cell mobilization is superior to G-CSF alone. Blood,
2005. 106(5): p. 1867-74.
98. Moore, M.A., et al., Mobilization of endothelial and hematopoietic stem and
progenitor cells by adenovector-mediated elevation of serum levels of SDF-1,
VEGF, and angiopoietin-1. Ann N Y Acad Sci, 2001. 938: p. 36-45; discussion
45-7.
157 99. Hattori, K., et al., Plasma elevation of stromal cell-derived factor-1 induces
mobilization of mature and immature hematopoietic progenitor and stem cells.
Blood, 2001. 97(11): p. 3354-60.
100. Delano, M.J., et al., Neutrophil mobilization from the bone marrow during
polymicrobial sepsis is dependent on CXCL12 signaling. J Immunol, 2011.
187(2): p. 911-8.
101. Singh, P., et al., Expansion of bone marrow neutrophils following G-CSF
administration in mice results in osteolineage cell apoptosis and mobilization of
hematopoietic stem and progenitor cells. Leukemia, 2012. 26(11): p. 2375-83.
102. Christopher, M.J., et al., Expression of the G-CSF receptor in monocytic cells is
sufficient to mediate hematopoietic progenitor mobilization by G-CSF in mice. J
Exp Med, 2011. 208(2): p. 251-60.
103. Sasaki, Y., et al., Marginal expression of CXCR4 on c-kit+Sca-1+Lineage−
hematopoietic stem/progenitor cells. Int J Hematol, 2009. 90(5): p. 553-60.
104. Okumura, N., et al., Chemotactic and chemokinetic activities of stem cell factor
on murine hematopoietic progenitor cells. Blood, 1996. 87(10): p. 4100-8.
105. Yan, X.Q., et al., Mobilization of long-term hematopoietic reconstituting cells in
mice by the combination of stem cell factor plus granulocyte colony-stimulating
factor. Blood, 1994. 84(3): p. 795-9.
106. Dutt, P., J.F. Wang, and J.E. Groopman, Stromal cell-derived factor-1α and stem
cell factor/kit ligand share signaling pathways in hemopoietic progenitors: a
potential mechanism for cooperative induction of chemotaxis. J Immunol, 1998.
161(7): p. 3652-8.
158 107. Schweitzer, K.M., et al., Constitutive expression of E-selectin and vascular cell
adhesion molecule-1 on endothelial cells of hematopoietic tissues. Am J Pathol,
1996. 148(1): p. 165-75.
108. Furze, R.C. and S.M. Rankin, Neutrophil mobilization and clearance in the bone
marrow. Immunology, 2008. 125(3): p. 281-8.
109. Levesque, J.P., et al., Vascular cell adhesion molecule-1 (CD106) is cleaved by
neutrophil proteases in the bone marrow following hematopoietic progenitor cell
mobilization by granulocyte colony-stimulating factor. Blood, 2001. 98(5): p.
1289-97.
110. Levesque, J.P., et al., Characterization of hematopoietic progenitor mobilization
in protease-deficient mice. Blood, 2004. 104(1): p. 65-72.
111. Petty, J.M., et al., Crosstalk between CXCR4/stromal derived factor-1 and VLA-
4/VCAM-1 pathways regulates neutrophil retention in the bone marrow. J
Immunol, 2009. 182(1): p. 604-12.
112. Pereira, J.P., et al., Cannabinoid receptor 2 mediates the retention of immature B
cells in bone marrow sinusoids. Nat Immunol, 2009. 10(4): p. 403-11.
113. Burdon, P.C., C. Martin, and S.M. Rankin, The CXC chemokine MIP-2 stimulates
neutrophil mobilization from the rat bone marrow in a CD49d-dependent manner.
Blood, 2005. 105(6): p. 2543-8.
114. Koni, P.A., et al., Conditional vascular cell adhesion molecule 1 deletion in mice:
impaired lymphocyte migration to bone marrow. J Exp Med, 2001. 193(6): p.
741-54.
159 115. Papayannopoulou, T., G.V. Priestley, and B. Nakamoto, Anti-VLA4/VCAM-1-
induced mobilization requires cooperative signaling through the kit/mkit ligand
pathway. Blood, 1998. 91(7): p. 2231-9.
116. Watanabe, T., et al., GM-CSF-mobilized peripheral blood CD34+ cells differ from
steady-state bone marrow CD34+ cells in adhesion molecule expression. Bone
Marrow Transplant, 1997. 19(12): p. 1175-81.
hi 117. Teixido, J., et al., Role of β1 and β2 integrins in the adhesion of human CD34
stem cells to bone marrow stroma. J Clin Invest, 1992. 90(2): p. 358-67.
118. Pruijt, J.F., et al., Anti-LFA-1 blocking antibodies prevent mobilization of
hematopoietic progenitor cells induced by interleukin-8. Blood, 1998. 91(11): p.
4099-105.
119. Ueda, Y., et al., Inflammation controls B lymphopoiesis by regulating chemokine
CXCL12 expression. J Exp Med, 2004. 199(1): p. 47-58.
120. Bernardini, G., et al., CCL3 and CXCL12 regulate trafficking of mouse bone
marrow NK cell subsets. Blood, 2008. 111(7): p. 3626-34.
121. Sawada, S., et al., Disturbed CD4+ T cell homeostasis and in vitro HIV-1
susceptibility in transgenic mice expressing T cell line-tropic HIV-1 receptors. J
Exp Med, 1998. 187(9): p. 1439-49.
122. Kean, L.S., et al., Significant mobilization of both conventional and regulatory T
cells with AMD3100. Blood, 2011. 118(25): p. 6580-90.
123. Cyster, J.G. and S.R. Schwab, Sphingosine-1-phosphate and lymphocyte egress
from lymphoid organs. Annu Rev Immunol, 2012. 30: p. 69-94.
160 124. Jenne, C.N., et al., T-bet-dependent S1P5 expression in NK cells promotes egress
from lymph nodes and bone marrow. J Exp Med, 2009. 206(11): p. 2469-81.
125. Walzer, T., et al., Natural killer cell trafficking in vivo requires a dedicated
sphingosine 1-phosphate receptor. Nat Immunol, 2007. 8(12): p. 1337-44.
126. Eash, K.J., et al., CXCR2 and CXCR4 antagonistically regulate neutrophil
trafficking from murine bone marrow. J Clin Invest, 2010. 120(7): p. 2423-31.
127. Gautier, E.L., C. Jakubzick, and G.J. Randolph, Regulation of the migration and
survival of monocyte subsets by chemokine receptors and its relevance to
atherosclerosis. Arterioscler Thromb Vasc Biol, 2009. 29(10): p. 1412-8.
128. Ingersoll, M.A., et al., Monocyte trafficking in acute and chronic inflammation.
Trends Immunol, 2011. 32(10): p. 470-7.
129. Tsou, C.L., et al., Critical roles for CCR2 and MCP-3 in monocyte mobilization
from bone marrow and recruitment to inflammatory sites. J Clin Invest, 2007.
117(4): p. 902-9.
130. Serbina, N.V. and E.G. Pamer, Monocyte emigration from bone marrow during
bacterial infection requires signals mediated by chemokine receptor CCR2. Nat
Immunol, 2006. 7(3): p. 311-7.
131. Shi, C., et al., Bone marrow mesenchymal stem and progenitor cells induce
monocyte emigration in response to circulating toll-like receptor ligands.
Immunity, 2011. 34(4): p. 590-601.
132. Crane, M.J., K.L. Hokeness-Antonelli, and T.P. Salazar-Mather, Regulation of
inflammatory monocyte/macrophage recruitment from the bone marrow during
161 murine cytomegalovirus infection: role for type I interferons in localized
induction of CCR2 ligands. J Immunol, 2009. 183(4): p. 2810-7.
133. Wang, Y., et al., CCR2 and CXCR4 regulate peripheral blood monocyte
pharmacodynamics and link to efficacy in experimental autoimmune
encephalomyelitis. J Inflamm (Lond), 2009. 6: p. 32.
134. Si, Y., et al., CCR2 mediates hematopoietic stem and progenitor cell trafficking to
sites of inflammation in mice. J Clin Invest, 2010. 120(4): p. 1192-203.
135. Broxmeyer, H.E., et al., Dominant myelopoietic effector functions mediated by
chemokine receptor CCR1. J Exp Med, 1999. 189(12): p. 1987-92.
136. van Buul, J.D., et al., Migration of human hematopoietic progenitor cells across
bone marrow endothelium is regulated by vascular endothelial cadherin. J
Immunol, 2002. 168(2): p. 588-96.
137. Peled, A., et al., The chemokine SDF-1 activates the integrins LFA-1, VLA-4, and
VLA-5 on immature human CD34+ cells: role in transendothelial/stromal
migration and engraftment of NOD/SCID mice. Blood, 2000. 95(11): p. 3289-96.
138. Hargreaves, D.C., et al., A coordinated change in chemokine responsiveness
guides plasma cell movements. J Exp Med, 2001. 194(1): p. 45-56.
139. Harada, H., et al., Phenotypic difference of normal plasma cells from mature
myeloma cells. Blood, 1993. 81(10): p. 2658-63.
140. Frenette, P.S., et al., Endothelial selectins and vascular cell adhesion molecule-1
promote hematopoietic progenitor homing to bone marrow. Proc Natl Acad Sci U
S A, 1998. 95(24): p. 14423-8.
162 141. Frenette, P.S. and L. Weiss, Sulfated glycans induce rapid hematopoietic
progenitor cell mobilization: evidence for selectin-dependent and independent
mechanisms. Blood, 2000. 96(7): p. 2460-8.
142. Rankin, S.M., The bone marrow: a site of neutrophil clearance. J Leukoc Biol,
2010. 88(2): p. 241-51.
143. Summers, C., et al., Neutrophil kinetics in health and disease. Trends Immunol,
2010. 31(8): p. 318-24.
144. Nagase, H., et al., Cytokine-mediated regulation of CXCR4 expression in human
neutrophils. J Leukoc Biol, 2002. 71(4): p. 711-7.
145. Furze, R.C. and S.M. Rankin, The role of the bone marrow in neutrophil
clearance under homeostatic conditions in the mouse. FASEB J, 2008. 22(9): p.
3111-9.
146. Varol, C., et al., Monocytes give rise to mucosal, but not splenic, conventional
dendritic cells. J Exp Med, 2007. 204(1): p. 171-80.
147. Lim, J.K., et al., Chemokine receptor Ccr2 is critical for monocyte accumulation
and survival in West Nile virus encephalitis. J Immunol, 2011. 186(1): p. 471-8.
148. Ballard, C., et al., Alzheimer's disease. Lancet, 2011. 377(9770): p. 1019-31.
149. Serrano-Pozo A1, F.M., Masliah E, Hyman BT, Neuropathological alterations in
Alzheimer disease. Cold Spring Harb Perspect Med. , 2011.
150. Mayeux, R. and Y. Stern, Epidemiology of Alzheimer disease. Cold Spring Harbor
perspectives in medicine, 2012. 2(8).
163 151. Haass, C. and D.J. Selkoe, Soluble protein oligomers in neurodegeneration:
lessons from the Alzheimer's amyloid β-peptide. Nature reviews. Molecular cell
biology, 2007. 8(2): p. 101-12.
152. Selkoe, D.J., Alzheimer's disease: genes, proteins, and therapy. Physiological
reviews, 2001. 81(2): p. 741-66.
153. Liu, F., et al., Dephosphorylation of tau by protein phosphatase 5: impairment in
Alzheimer's disease. The Journal of biological chemistry, 2005. 280(3): p. 1790-6.
154. McGeer, P.L., M. Schulzer, and E.G. McGeer, Arthritis and anti-inflammatory
agents as possible protective factors for Alzheimer's disease: a review of 17
epidemiologic studies. Neurology, 1996. 47(2): p. 425-32.
155. Akiyama, H., et al., Inflammation and Alzheimer's disease. Neurobiology of
aging, 2000. 21(3): p. 383-421.
156. Colton, C.A., et al., Expression profiles for macrophage alternative activation
genes in AD and in mouse models of AD. J Neuroinflammation, 2006. 3: p. 27.
157. Mandrekar, S., et al., Microglia mediate the clearance of soluble Aβ through fluid
phase macropinocytosis. J Neurosci, 2009. 29(13): p. 4252-62.
158. Bamberger, M.E., et al., A cell surface receptor complex for fibrillar β-amyloid
mediates microglial activation. J Neurosci, 2003. 23(7): p. 2665-74.
159. Koenigsknecht, J. and G. Landreth, Microglial phagocytosis of fibrillar β-amyloid
through a β1 integrin-dependent mechanism. J Neurosci, 2004. 24(44): p. 9838-
46.
160. Jiang, Q., et al., ApoE promotes the proteolytic degradation of Aβ. Neuron, 2008.
58(5): p. 681-93.
164 161. Nicoll, J.A., et al., Neuropathology of human Alzheimer disease after
immunization with amyloid-β peptide: a case report. Nature medicine, 2003. 9(4):
p. 448-52.
162. Boche, D., et al., Neuropathology after active Aβ42 immunotherapy: implications
for Alzheimer's disease pathogenesis. Acta neuropathologica, 2010. 120(3): p.
369-84.
163. Schenk, D., et al., Immunization with amyloid-β attenuates Alzheimer-disease-like
pathology in the PDAPP mouse. Nature, 1999. 400(6740): p. 173-7.
164. Liu, S., et al., TLR2 is a primary receptor for Alzheimer's amyloid β peptide to
trigger neuroinflammatory activation. J Immunol, 2012. 188(3): p. 1098-107.
165. Simard, A.R., et al., Bone marrow-derived microglia play a critical role in
restricting senile plaque formation in Alzheimer's disease. Neuron, 2006. 49(4): p.
489-502.
166. Hao, W., et al., Myeloid differentiation factor 88-deficient bone marrow cells
improve Alzheimer's disease-related symptoms and pathology. Brain : a journal of
neurology, 2011. 134(Pt 1): p. 278-92.
167. Feng, Y., L. Li, and X.H. Sun, Monocytes and Alzheimer's disease. Neurosci Bull,
2011. 27(2): p. 115-22.
168. Fiala, M., et al., Amyloid-β induces chemokine secretion and monocyte migration
across a human blood--brain barrier model. Mol Med, 1998. 4(7): p. 480-9.
169. Fiala, M., et al., Ineffective phagocytosis of amyloid-β by macrophages of
Alzheimer's disease patients. J Alzheimers Dis, 2005. 7(3): p. 221-32; discussion
255-62.
165 170. Zaghi, J., et al., Alzheimer disease macrophages shuttle amyloid-beta from
neurons to vessels, contributing to amyloid angiopathy. Acta Neuropathol, 2009.
117(2): p. 111-24.
171. Mildner, A., et al., Distinct and non-redundant roles of microglia and myeloid
subsets in mouse models of Alzheimer's disease. J Neurosci, 2011. 31(31): p.
11159-71.
172. Michaud, J.P., et al., Real-time in vivo imaging reveals the ability of monocytes to
clear vascular amyloid beta. Cell Rep, 2013. 5(3): p. 646-53.
173. Shad, K.F., et al., Peripheral markers of Alzheimer's disease: surveillance of
white blood cells. Synapse, 2013. 67(8): p. 541-3.
174. Jaremo, P., et al., Alzheimer's disease and granulocyte density diversity. Eur J
Clin Invest, 2013. 43(6): p. 545-8.
175. Kuyumcu, M.E., et al., The evaluation of neutrophil-lymphocyte ratio in
Alzheimer's disease. Dement Geriatr Cogn Disord, 2012. 34(2): p. 69-74.
176. Guthrie, G.J., et al., The systemic inflammation-based neutrophil-lymphocyte
ratio: experience in patients with cancer. Crit Rev Oncol Hematol, 2013. 88(1): p.
218-30.
177. Imtiaz, F., et al., Neutrophil lymphocyte ratio as a measure of systemic
inflammation in prevalent chronic diseases in Asian population. Int Arch Med,
2012. 5(1): p. 2.
178. Gibson, P.H., et al., Preoperative neutrophil-lymphocyte ratio and outcome from
coronary artery bypass grafting. Am Heart J, 2007. 154(5): p. 995-1002.
166 179. Uslu, A.U., et al., Is neutrophil/lymphocyte ratio associated with subclinical
inflammation and amyloidosis in patients with familial Mediterranean fever?
Biomed Res Int, 2013. 2013: p. 185317.
180. Gokmen, F., et al., Neutrophil-Lymphocyte Ratio Connected to Treatment Options
and Inflammation Markers of Ankylosing Spondylitis. J Clin Lab Anal, 2014.
181. Kim, M.A., Y.S. Lee, and K. Seo, Assessment of predictive markers for placental
inflammatory response in preterm births. PLoS One, 2014. 9(10): p. e107880.
182. Semiz, M., et al., Elevated neutrophil/lymphocyte ratio in patients with
schizophrenia. Psychiatr Danub, 2014. 26(3): p. 220-5.
183. Lee, J.H., et al., Characteristics of platelet indices, neutrophil-to-lymphocyte ratio
and erythrocyte sedimentation rate compared with C reactive protein in patients
with cerebral infarction: a retrospective analysis of comparing haematological
parameters and C reactive protein. BMJ Open, 2014. 4(11): p. e006275.
184. Davydova, T.V., et al., Phagocytic activity and state of bactericidal systems in
polymorphonuclear leukocytes from patients with Alzheimer's disease. Bull Exp
Biol Med, 2003. 136(4): p. 355-7.
185. Baik, S.H., et al., Migration of neutrophils targeting amyloid plaques in
Alzheimer's disease mouse model. Neurobiol Aging, 2014. 35(6): p. 1286-92.
186. Neumann, J., et al., Microglia cells protect neurons by direct engulfment of
invading neutrophil granulocytes: a new mechanism of CNS immune privilege. J
Neurosci, 2008. 28(23): p. 5965-75.
187. Soehnlein, O., L. Lindbom, and C. Weber, Mechanisms underlying neutrophil-
mediated monocyte recruitment. Blood, 2009. 114(21): p. 4613-23.
167 188. Neirinckx, V., et al., Neutrophil contribution to spinal cord injury and repair. J
Neuroinflammation, 2014. 11: p. 150.
189. de Castro, R., Jr., et al., Evidence that infiltrating neutrophils do not release
reactive oxygen species in the site of spinal cord injury. Exp Neurol, 2004.
190(2): p. 414-24.
190. Stirling, D.P., et al., Depletion of Ly6G/Gr-1 leukocytes after spinal cord injury in
mice alters wound healing and worsens neurological outcome. J Neurosci, 2009.
29(3): p. 753-64.
191. Maler, J.M., et al., Decreased circulating CD34+ stem cells in early Alzheimer's
disease: Evidence for a deficient hematopoietic brain support? Mol Psychiatry,
2006. 11(12): p. 1113-5.
192. Iadecola, C., Neurovascular regulation in the normal brain and in Alzheimer's
disease. Nat Rev Neurosci, 2004. 5(5): p. 347-60.
193. Kelleher, R.J. and R.L. Soiza, Evidence of endothelial dysfunction in the
development of Alzheimer's disease: Is Alzheimer's a vascular disorder? Am J
Cardiovasc Dis, 2013. 3(4): p. 197-226.
194. Bell, R.D., et al., SRF and myocardin regulate LRP-mediated amyloid-β
clearance in brain vascular cells. Nat Cell Biol, 2009. 11(2): p. 143-53.
195. Sagare, A.P., R.D. Bell, and B.V. Zlokovic, Neurovascular defects and faulty
amyloid-β vascular clearance in Alzheimer's disease. J Alzheimers Dis, 2013. 33
Suppl 1: p. S87-100.
196. Zlokovic, B.V., The blood-brain barrier in health and chronic neurodegenerative
disorders. Neuron, 2008. 57(2): p. 178-201.
168 197. Paris, D., et al., Vasoactive effects of Aβ in isolated human cerebrovessels and in
a transgenic mouse model of Alzheimer's disease: role of inflammation. Neurol
Res, 2003. 25(6): p. 642-51.
198. Niwa, K., et al., Aβ-peptides enhance vasoconstriction in cerebral circulation.
Am J Physiol Heart Circ Physiol, 2001. 281(6): p. H2417-24.
199. Burnstock, G., et al., Evidence that adenosine triphosphate or a related nucleotide
is the transmitter substance released by non-adrenergic inhibitory nerves in the
gut. Br J Pharmacol, 1970. 40(4): p. 668-88.
200. Bours, M.J., et al., Adenosine 5'-triphosphate and adenosine as endogenous
signaling molecules in immunity and inflammation. Pharmacol Ther, 2006.
112(2): p. 358-404.
201. Burnstock, G., Historical review: ATP as a neurotransmitter. Trends Pharmacol
Sci, 2006. 27(3): p. 166-76.
202. Burnstock, G., et al., Direct evidence for ATP release from non-adrenergic, non-
cholinergic ("purinergic") nerves in the guinea-pig taenia coli and bladder. Eur J
Pharmacol, 1978. 49(2): p. 145-9.
203. Lew, M.J. and T.D. White, Release of endogenous ATP during sympathetic nerve
stimulation. Br J Pharmacol, 1987. 92(2): p. 349-55.
204. Born, G.V. and M.A. Kratzer, Source and concentration of extracellular
adenosine triphosphate during haemostasis in rats, rabbits and man. J Physiol,
1984. 354: p. 419-29.
169 205. Schumacher, D., et al., Platelet-derived nucleotides promote tumor-cell
transendothelial migration and metastasis via P2Y2 receptor. Cancer Cell, 2013.
24(1): p. 130-7.
206. Di Virgilio, F., Liaisons dangereuses: P2X7 and the inflammasome. Trends
Pharmacol Sci, 2007. 28(9): p. 465-72.
207. Bergfeld, G.R. and T. Forrester, Release of ATP from human erythrocytes in
response to a brief period of hypoxia and hypercapnia. Cardiovasc Res, 1992.
26(1): p. 40-7.
208. Wan, J., W.D. Ristenpart, and H.A. Stone, Dynamics of shear-induced ATP
release from red blood cells. Proc Natl Acad Sci U S A, 2008. 105(43): p. 16432-
7.
209. Pearson, J.D. and J.L. Gordon, Vascular endothelial and smooth muscle cells in
culture selectively release adenine nucleotides. Nature, 1979. 281(5730): p. 384-
6.
210. Yamamoto, K., et al., Visualization of flow-induced ATP release and triggering of
Ca2+ waves at caveolae in vascular endothelial cells. J Cell Sci, 2011. 124(Pt 20):
p. 3477-83.
211. Riddle, R.C., et al., ATP release mediates fluid flow-induced proliferation of
human bone marrow stromal cells. J Bone Miner Res, 2007. 22(4): p. 589-600.
212. Di Virgilio, F., Purines, purinergic receptors, and cancer. Cancer Res, 2012.
72(21): p. 5441-7.
170 213. Jin, H., et al., P2Y2 receptor activation by nucleotides released from highly
metastatic breast cancer cells increases tumor growth and invasion via crosstalk
with endothelial cells. Breast Cancer Res, 2014. 16(4): p. R77.
214. Lazarowski, E.R. and T.K. Harden, Quantitation of extracellular UTP using a
sensitive enzymatic assay. Br J Pharmacol, 1999. 127(5): p. 1272-8.
215. Lazarowski, E.R. and R.C. Boucher, UTP as an extracellular signaling molecule.
News Physiol Sci, 2001. 16: p. 1-5.
216. Burnstock, G., Purinergic signalling: Its unpopular beginning, its acceptance and
its exciting future. Bioessays, 2012. 34(3): p. 218-25.
217. Valera, S., et al., A new class of ligand-gated ion channel defined by P2X receptor
for extracellular ATP. Nature, 1994. 371(6497): p. 516-9.
218. Nicke, A., et al., P2X1 and P2X3 receptors form stable trimers: a novel structural
motif of ligand-gated ion channels. EMBO J, 1998. 17(11): p. 3016-28.
219. Barrera, N.P., et al., Atomic force microscopy imaging demonstrates that P2X2
receptors are trimers but that P2X6 receptor subunits do not oligomerize. J Biol
Chem, 2005. 280(11): p. 10759-65.
220. Antonio, L.S., et al., Identification of P2X2/P2X4/P2X6 heterotrimeric receptors
using atomic force microscopy (AFM) imaging. FEBS Lett, 2014. 588(12): p.
2125-8.
221. Compan, V., et al., P2X2 and P2X5 subunits define a new heteromeric receptor
with P2X7-like properties. J Neurosci, 2012. 32(12): p. 4284-96.
171 222. Velazquez, B., et al., Differential agonist-induced desensitization of P2Y2
nucleotide receptors by ATP and UTP. Mol Cell Biochem, 2000. 206(1-2): p. 75-
89.
223. Erb, L., et al., An RGD sequence in the P2Y2 receptor interacts with αVβ3
integrins and is required for Go-mediated signal transduction. J Cell Biol, 2001.
153(3): p. 491-501.
224. Liao, Z., et al., The P2Y2 nucleotide receptor requires interaction with αV
integrins to access and activate G12. J Cell Sci, 2007. 120(Pt 9): p. 1654-62.
225. Bagchi, S., et al., The P2Y2 nucleotide receptor interacts with αV integrins to
activate Go and induce cell migration. J Biol Chem, 2005. 280(47): p. 39050-7.
226. Seye, C.I., et al., The P2Y2 nucleotide receptor mediates vascular cell adhesion
molecule-1 expression through interaction with VEGF receptor-2 (KDR/Flk-1). J
Biol Chem, 2004. 279(34): p. 35679-86.
227. Liu, J., et al., Src homology 3 binding sites in the P2Y2 nucleotide receptor
interact with Src and regulate activities of Src, proline-rich tyrosine kinase 2, and
growth factor receptors. J Biol Chem, 2004. 279(9): p. 8212-8.
228. Kong, Q., et al., Interleukin-1β enhances nucleotide-induced and α-secretase-
dependent amyloid precursor protein processing in rat primary cortical neurons
via up-regulation of the P2Y(2) receptor. J Neurochem, 2009. 109(5): p. 1300-10.
229. Leon-Otegui, M., et al., Opposite effects of P2X7 and P2Y2 nucleotide receptors
on α-secretase-dependent APP processing in Neuro-2a cells. FEBS Lett, 2011.
585(14): p. 2255-62.
172 230. Camden, J.M., et al., P2Y2 nucleotide receptors enhance α-secretase-dependent
amyloid precursor protein processing. J Biol Chem, 2005. 280(19): p. 18696-702.
231. Mezyk-Kopec, R., et al., Identification of ADAM10 as a major TNF sheddase in
ADAM17-deficient fibroblasts. Cytokine, 2009. 46(3): p. 309-15.
232. Huovila, A.P., et al., Shedding light on ADAM metalloproteinases. Trends
Biochem Sci, 2005. 30(7): p. 413-22.
233. Moore, D.J., et al., Expression pattern of human P2Y receptor subtypes: a
quantitative reverse transcription-polymerase chain reaction study. Biochim
Biophys Acta, 2001. 1521(1-3): p. 107-19.
234. Yip, L., et al., Autocrine regulation of T-cell activation by ATP release and P2X7
receptors. FASEB J, 2009. 23(6): p. 1685-93.
235. Schenk, U., et al., Purinergic control of T cell activation by ATP released through
pannexin-1 hemichannels. Sci Signal, 2008. 1(39): p. ra6.
236. Woehrle, T., et al., Pannexin-1 hemichannel-mediated ATP release together with
P2X1 and P2X4 receptors regulate T-cell activation at the immune synapse.
Blood, 2010. 116(18): p. 3475-84.
237. Chen, Y., et al., ATP release guides neutrophil chemotaxis via P2Y2 and A3
receptors. Science, 2006. 314(5806): p. 1792-5.
238. Kronlage, M., et al., Autocrine purinergic receptor signaling is essential for
macrophage chemotaxis. Sci Signal, 2010. 3(132): p. ra55.
239. Isfort, K., et al., Real-time imaging reveals that P2Y2 and P2Y12 receptor agonists
are not chemoattractants and macrophage chemotaxis to complement C5a is
173 phosphatidylinositol 3-kinase (PI3K)- and p38 mitogen-activated protein kinase
(MAPK)-independent. J Biol Chem, 2011. 286(52): p. 44776-87.
240. Elliott, M.R., et al., Nucleotides released by apoptotic cells act as a find-me
signal to promote phagocytic clearance. Nature, 2009. 461(7261): p. 282-6.
241. Rieg, T., et al., Mice lacking P2Y2 receptors have salt-resistant hypertension and
facilitated renal Na+ and water reabsorption. FASEB J, 2007. 21(13): p. 3717-
26.
242. Cicko, S., et al., Purinergic receptor inhibition prevents the development of
smoke-induced lung injury and emphysema. J Immunol, 2010. 185(1): p. 688-97.
243. Muller, T., et al., The purinergic receptor P2Y2 receptor mediates chemotaxis of
dendritic cells and eosinophils in allergic lung inflammation. Allergy, 2010.
65(12): p. 1545-53.
244. Seye, C.I., et al., Functional P2Y2 nucleotide receptors mediate uridine 5'-
triphosphate-induced intimal hyperplasia in collared rabbit carotid arteries.
Circulation, 2002. 106(21): p. 2720-6.
245. Seye, C.I., et al., P2Y2 receptor-mediated lymphotoxin-α secretion regulates
intercellular cell adhesion molecule-1 expression in vascular smooth muscle cells.
J Biol Chem, 2012. 287(13): p. 10535-43.
246. Madge, L.A., et al., Lymphotoxin-α1β2 and LIGHT induce classical and
noncanonical NF-κB-dependent proinflammatory gene expression in vascular
endothelial cells. J Immunol, 2008. 180(5): p. 3467-77.
247. Lemoli, R.M., et al., Extracellular nucleotides are potent stimulators of human
hematopoietic stem cells in vitro and in vivo. Blood, 2004. 104(6): p. 1662-70.
174 248. Barbosa, C.M., et al., Differentiation of hematopoietic stem cell and myeloid
populations by ATP is modulated by cytokines. Cell Death Dis, 2011. 2: p. e165.
249. Rossi, L., et al., The extracellular nucleotide UTP is a potent inducer of
hematopoietic stem cell migration. Blood, 2007. 109(2): p. 533-42.
250. Laupheimer, M., et al., Selective Migration of Subpopulations of Bone Marrow
Cells along an SDF-1α and ATP Gradient. Bone Marrow Res, 2014. 2014: p.
182645.
251. Gardinier, J., et al., P2Y2 receptors regulate osteoblast mechanosensitivity during
fluid flow. Am J Physiol Cell Physiol, 2014. 306(11): p. C1058-67.
252. Morrison, M.S., et al., ATP is a potent stimulator of the activation and formation
of rodent osteoclasts. J Physiol, 1998. 511 ( Pt 2): p. 495-500.
253. Hoebertz, A., et al., Extracellular ADP is a powerful osteolytic agent: evidence
for signaling through the P2Y1 receptor on bone cells. FASEB J, 2001. 15(7): p.
1139-48.
254. Orriss, I.R., et al., The P2Y6 receptor stimulates bone resorption by osteoclasts.
Endocrinology, 2011. 152(10): p. 3706-16.
255. Orriss, I., et al., Bone phenotypes of P2 receptor knockout mice. Front Biosci
(Schol Ed), 2011. 3: p. 1038-46.
256. Xing, Y., et al., The roles of P2Y2 purinergic receptors in osteoblasts and
mechanotransduction. PLoS One, 2014. 9(9): p. e108417.
257. Hoebertz, A., et al., ATP and UTP at low concentrations strongly inhibit bone
formation by osteoblasts: a novel role for the P2Y2 receptor in bone remodeling. J
Cell Biochem, 2002. 86(3): p. 413-9.
175 258. Orriss, I.R., et al., Extracellular nucleotides block bone mineralization in vitro:
evidence for dual inhibitory mechanisms involving both P2Y2 receptors and
pyrophosphate. Endocrinology, 2007. 148(9): p. 4208-16.
259. Syberg, S., et al., Clopidogrel (Plavix), a P2Y12 receptor antagonist, inhibits bone
cell function in vitro and decreases trabecular bone in vivo. J Bone Miner Res,
2012. 27(11): p. 2373-86.
260. Burnstock, G., Purinergic signalling and disorders of the central nervous system.
Nat Rev Drug Discov, 2008. 7(7): p. 575-90.
261. Takenouchi, T., et al., P2X7 receptor signaling pathway as a therapeutic target
for neurodegenerative diseases. Arch Immunol Ther Exp (Warsz), 2010. 58(2): p.
91-6.
262. Rahman, A., The role of adenosine in Alzheimer's disease. Curr Neuropharmacol,
2009. 7(3): p. 207-16.
263. Weisman, G.A., et al., P2 receptors for extracellular nucleotides in the central
nervous system: role of P2X7 and P2Y2 receptor interactions in
neuroinflammation. Mol Neurobiol, 2012. 46(1): p. 96-113.
264. Burnstock, G. and V. Ralevic, Purinergic signaling and blood vessels in health
and disease. Pharmacol Rev, 2014. 66(1): p. 102-92.
265. Erlinge, D. and G. Burnstock, P2 receptors in cardiovascular regulation and
disease. Purinergic Signal, 2008. 4(1): p. 1-20.
266. Peterson, T.S., et al., P2Y2 nucleotide receptor-mediated responses in brain cells.
Mol Neurobiol, 2010. 41(2-3): p. 356-66.
176 267. Kim, H.J., et al., Nucleotides released from Aβ1-42-treated microglial cells
increase cell migration and Aβ1-42 uptake through P2Y2 receptor activation.
Journal of neurochemistry, 2012. 121(2): p. 228-238.
268. Sanz, J.M., et al., Activation of microglia by amyloid β requires P2X7 receptor
expression. J Immunol, 2009. 182(7): p. 4378-85.
269. Peterson, T.S., et al., Up-regulation and activation of the P2Y2 nucleotide
receptor mediate neurite extension in IL-1β-treated mouse primary cortical
neurons. J Neurochem, 2013. 125(6): p. 885-96.
270. Chakfe, Y., et al., ADP and AMP induce interleukin-1β release from microglial
cells through activation of ATP-primed P2X7 receptor channels. J Neurosci,
2002. 22(8): p. 3061-9.
271. Suzuki, T., et al., Production and release of neuroprotective tumor necrosis factor
by P2X7 receptor-activated microglia. J Neurosci, 2004. 24(1): p. 1-7.
272. Choi, H.B., et al., Modulation of the purinergic P2X7 receptor attenuates
lipopolysaccharide-mediated microglial activation and neuronal damage in
inflamed brain. J Neurosci, 2007. 27(18): p. 4957-68.
273. Mingam, R., et al., In vitro and in vivo evidence for a role of the P2X7 receptor in
the release of IL-1β in the murine brain. Brain Behav Immun, 2008. 22(2): p.
234-44.
274. Takenouchi, T., et al., Modulation of the ATP-lnduced release and processing of
IL-1β in microglial cells. Crit Rev Immunol, 2009. 29(4): p. 335-45.
177 275. D'Alimonte, I., et al., Activation of P2X7 receptors stimulates the expression of
P2Y2 receptor mRNA in astrocytes cultured from rat brain. Int J Immunopathol
Pharmacol, 2007. 20(2): p. 301-16.
276. Takenouchi, T., et al., The activation of P2X7 receptor induces cathepsin D-
dependent production of a 20-kDa form of IL-1β under acidic extracellular pH in
LPS-primed microglial cells. J Neurochem, 2011. 117(4): p. 712-23.
277. Misiti, F., et al., Amyloid peptide inhibits ATP release from human erythrocytes.
Biochem Cell Biol, 2008. 86(6): p. 501-8.
278. Guns, P.J., et al., Endothelium-dependent relaxation evoked by ATP and UTP in
the aorta of P2Y2-deficient mice. Br J Pharmacol, 2006. 147(5): p. 569-74.
279. Wangensteen, R., et al., Contribution of endothelium-derived relaxing factors to
P2Y-purinoceptor-induced vasodilation in the isolated rat kidney. Gen
Pharmacol, 2000. 35(3): p. 129-33.
280. Doganay, G., et al., Pharmacological manipulation of the vasoconstrictive effects
of amyloid-β peptides by donepezil and rivastigmine. Curr Alzheimer Res, 2006.
3(2): p. 137-45.
281. Tran, M.D., P2 receptor stimulation induces amyloid precursor protein
production and secretion in rat cortical astrocytes. Neurosci Lett, 2011. 492(3):
p. 155-9.
282. O'Brien, R.J. and P.C. Wong, Amyloid precursor protein processing and
Alzheimer's disease. Annual review of neuroscience, 2011. 34: p. 185-204.
178 283. Skovronsky, D.M., et al., Protein kinase C-dependent α-secretase competes with
β-secretase for cleavage of amyloid-β precursor protein in the trans-golgi
network. The Journal of biological chemistry, 2000. 275(4): p. 2568-75.
284. Lichtenthaler, S.F., Alpha-secretase cleavage of the amyloid precursor protein:
proteolysis regulated by signaling pathways and protein trafficking. Current
Alzheimer research, 2012. 9(2): p. 165-77.
285. Deuss, M., K. Reiss, and D. Hartmann, Part-time α-secretases: the functional
biology of ADAM 9, 10 and 17. Curr Alzheimer Res, 2008. 5(2): p. 187-201.
286. Nasu-Tada, K., S. Koizumi, and K. Inoue, Involvement of β1 integrin in microglial
chemotaxis and proliferation on fibronectin: different regulations by ADP
through PKA. Glia, 2005. 52(2): p. 98-107.
287. Haynes, S.E., et al., The P2Y12 receptor regulates microglial activation by
extracellular nucleotides. Nat Neurosci, 2006. 9(12): p. 1512-9.
288. Li, H.Q., et al., P2Y4 receptor-mediated pinocytosis contributes to amyloid beta-
induced self-uptake by microglia. Mol Cell Biol, 2013. 33(21): p. 4282-93.
289. Koizumi, S., et al., UDP acting at P2Y6 receptors is a mediator of microglial
phagocytosis. Nature, 2007. 446(7139): p. 1091-5.
290. Kim, B., et al., Uridine 5'-diphosphate induces chemokine expression in microglia
and astrocytes through activation of the P2Y6 receptor. J Immunol, 2011. 186(6):
p. 3701-9.
291. Honda, S., et al., Extracellular ATP or ADP induce chemotaxis of cultured
microglia through Gi/o-coupled P2Y receptors. J Neurosci, 2001. 21(6): p. 1975-
82.
179 292. Ajit, D., et al., Loss of P2Y2 nucleotide receptors enhances early pathology in the
TgCRND8 mouse model of Alzheimer's disease. Mol Neurobiol, 2014. 49(2): p.
1031-42.
293. Lai, M.K., et al., Selective loss of P2Y2 nucleotide receptor immunoreactivity is
associated with Alzheimer's disease neuropathology. J Neural Transm, 2008.
115(8): p. 1165-72.
294. Arthur, D.B., K. Akassoglou, and P.A. Insel, P2Y2 receptor activates nerve
growth factor/TrkA signaling to enhance neuronal differentiation. Proc Natl Acad
Sci U S A, 2005. 102(52): p. 19138-43.
295. Lai, M.K., et al., Selective loss of P2Y2 nucleotide receptor immunoreactivity is
associated with Alzheimer's disease neuropathology. Journal of neural
transmission, 2008. 115(8): p. 1165-72.
296. Erb, L. and G.A. Weisman, Coupling of P2Y receptors to G proteins and other
signaling pathways. Wiley Interdiscip Rev Membr Transp Signal, 2012. 1(6): p.
789-803.
297. Wojciak-Stothard, B. and A.J. Ridley, Rho GTPases and the regulation of
endothelial permeability. Vascul Pharmacol, 2002. 39(4-5): p. 187-99.
298. Beckers, C.M., V.W. van Hinsbergh, and G.P. van Nieuw Amerongen, Driving
Rho GTPase activity in endothelial cells regulates barrier integrity. Thromb
Haemost, 2010. 103(1): p. 40-55.
299. Harvey, J., Erb, L., Huxley, V., Weisman, G.A., Garrad, R. and Wang, J., P2Y2
Receptor Dependent Modulation of Microvascular Barrier Function. FASEB J,
2012. 26(1): p. 855.4.
180 300. Muller, T., et al., The purinergic receptor P2Y2 receptor mediates chemotaxis of
dendritic cells and eosinophils in allergic lung inflammation. Allergy. 65(12): p.
1545-53.
301. Cicko, S., et al., Purinergic receptor inhibition prevents the development of
smoke-induced lung injury and emphysema. J Immunol. 185(1): p. 688-97.
302. Agca, C., et al., Development of a novel transgenic rat overexpressing the P2Y(2)
nucleotide receptor using a lentiviral vector. J Vasc Res, 2009. 46(5): p. 447-58.
303. Schumacher, D., et al., Platelet-Derived Nucleotides Promote Tumor-Cell
Transendothelial Migration and Metastasis via P2Y Receptor. Cancer Cell, 2013.
304. Li, W.H., et al., P2Y2 receptor promotes cell invasion and metastasis in prostate
cancer cells. Br J Cancer, 2013. 109(6): p. 1666-75.
305. Wheelock, M.J. and K.R. Johnson, Cadherin-mediated cellular signaling. Curr
Opin Cell Biol, 2003. 15(5): p. 509-14.
306. Dejana, E., G. Bazzoni, and M.G. Lampugnani, Vascular endothelial (VE)-
cadherin: only an intercellular glue? Exp Cell Res, 1999. 252(1): p. 13-9.
307. Breviario, F., et al., Functional properties of human vascular endothelial
cadherin (7B4/cadherin-5), an endothelium-specific cadherin. Arterioscler
Thromb Vasc Biol, 1995. 15(8): p. 1229-39.
308. Corada, M., et al., Vascular endothelial-cadherin is an important determinant of
microvascular integrity in vivo. Proc Natl Acad Sci U S A, 1999. 96(17): p. 9815-
20.
309. Matsuyoshi, N., et al., In vivo evidence of the critical role of cadherin-5 in murine
vascular integrity. Proc Assoc Am Physicians, 1997. 109(4): p. 362-71.
181 310. Gotsch, U., et al., VE-cadherin antibody accelerates neutrophil recruitment in
vivo. J Cell Sci, 1997. 110 ( Pt 5): p. 583-8.
311. Carmeliet, P., et al., Targeted deficiency or cytosolic truncation of the VE-
cadherin gene in mice impairs VEGF-mediated endothelial survival and
angiogenesis. Cell, 1999. 98(2): p. 147-57.
312. Gory-Faure, S., et al., Role of vascular endothelial-cadherin in vascular
morphogenesis. Development, 1999. 126(10): p. 2093-102.
313. Zanetta, L., et al., Downregulation of vascular endothelial-cadherin expression is
associated with an increase in vascular tumor growth and hemorrhagic
complications. Thromb Haemost, 2005. 93(6): p. 1041-6.
314. Rahimi, N. and A. Kazlauskas, A role for cadherin-5 in regulation of vascular
endothelial growth factor receptor 2 activity in endothelial cells. Mol Biol Cell,
1999. 10(10): p. 3401-7.
315. Lampugnani, M.G., et al., VE-cadherin regulates endothelial actin activating Rac
and increasing membrane association of Tiam. Mol Biol Cell, 2002. 13(4): p.
1175-89.
316. Zanetti, A., et al., Vascular endothelial growth factor induces SHC association
with vascular endothelial cadherin: a potential feedback mechanism to control
vascular endothelial growth factor receptor-2 signaling. Arterioscler Thromb
Vasc Biol, 2002. 22(4): p. 617-22.
317. Baumeister, U., et al., Association of Csk to VE-cadherin and inhibition of cell
proliferation. Embo J, 2005. 24(9): p. 1686-95.
182 318. Nawroth, R., et al., VE-PTP and VE-cadherin ectodomains interact to facilitate
regulation of phosphorylation and cell contacts. Embo J, 2002. 21(18): p. 4885-
95.
319. Lampugnani, M.G., et al., Vascular endothelial cadherin controls VEGFR-2
internalization and signaling from intracellular compartments. J Cell Biol, 2006.
174(4): p. 593-604.
320. Grazia Lampugnani, M., et al., Contact inhibition of VEGF-induced proliferation
requires vascular endothelial cadherin, β-catenin, and the phosphatase DEP-
1/CD148. J Cell Biol, 2003. 161(4): p. 793-804.
321. Esser, S., et al., Vascular endothelial growth factor induces VE-cadherin tyrosine
phosphorylation in endothelial cells. J Cell Sci, 1998. 111 ( Pt 13): p. 1853-65.
322. Birukova, A.A., et al., VE-cadherin trans-interactions modulate Rac activation
and enhancement of lung endothelial barrier by iloprost. J Cell Physiol, 2012.
227(10): p. 3405-16.
323. Iyer, S., et al., VE-cadherin-p120 interaction is required for maintenance of
endothelial barrier function. Am J Physiol Lung Cell Mol Physiol, 2004. 286(6):
p. L1143-53.
324. Radu, M. and J. Chernoff, An in vivo assay to test blood vessel permeability. J Vis
Exp, 2013(73): p. e50062.
325. Kukulski, F., et al., Endothelial P2Y2 receptor regulates LPS-induced neutrophil
transendothelial migration in vitro. Mol Immunol, 2010. 47(5): p. 991-9.
326. van Buul, J.D. and P.L. Hordijk, Signaling in leukocyte transendothelial
migration. Arterioscler Thromb Vasc Biol, 2004. 24(5): p. 824-33.
183 327. Huang, A.J., et al., Endothelial cell cytosolic free calcium regulates neutrophil
migration across monolayers of endothelial cells. J Cell Biol, 1993. 120(6): p.
1371-80.
328. Vestweber, D., Adhesion and signaling molecules controlling the transmigration
of leukocytes through endothelium. Immunol Rev, 2007. 218: p. 178-96.
329. Muller, W.A., Mechanisms of transendothelial migration of leukocytes. Circ Res,
2009. 105(3): p. 223-30.
330. Dejana, E., F. Orsenigo, and M.G. Lampugnani, The role of adherens junctions
and VE-cadherin in the control of vascular permeability. J Cell Sci, 2008. 121(Pt
13): p. 2115-22.
331. Giannotta, M., M. Trani, and E. Dejana, VE-cadherin and endothelial adherens
junctions: active guardians of vascular integrity. Dev Cell, 2013. 26(5): p. 441-
54.
332. Fernandez-Borja, M., J.D. van Buul, and P.L. Hordijk, The regulation of
leucocyte transendothelial migration by endothelial signalling events. Cardiovasc
Res. 86(2): p. 202-10.
333. Gavard, J. and J.S. Gutkind, VEGF controls endothelial-cell permeability by
promoting the β-arrestin-dependent endocytosis of VE-cadherin. Nat Cell Biol,
2006. 8(11): p. 1223-34.
334. Mukherjee, S., M. Tessema, and A. Wandinger-Ness, Vesicular trafficking of
tyrosine kinase receptors and associated proteins in the regulation of signaling
and vascular function. Circ Res, 2006. 98(6): p. 743-56.
184 335. Schulz, B., et al., ADAM10 regulates endothelial permeability and T-Cell
transmigration by proteolysis of vascular endothelial cadherin. Circ Res, 2008.
102(10): p. 1192-201.
336. Hebda, J.K., et al., The C-terminus region of β-arrestin1 modulates VE-cadherin
expression and endothelial cell permeability. Cell Commun Signal, 2013. 11(1):
p. 37.
337. Ratchford, A.M., et al., P2Y2 nucleotide receptors mediate metalloprotease-
dependent phosphorylation of epidermal growth factor receptor and ErbB3 in
human salivary gland cells. J Biol Chem. 285(10): p. 7545-55.
338. Korczynski, J., et al., Is MLC phosphorylation essential for the recovery from
ROCK inhibition in glioma C6 cells? Acta Biochim Pol, 2011. 58(1): p. 125-30.
339. Kaczmarek, E., et al., Modulation of endothelial cell migration by extracellular
nucleotides: involvement of focal adhesion kinase and phosphatidylinositol 3-
kinase-mediated pathways. Thromb Haemost, 2005. 93(4): p. 735-42.
340. Haidari, M., et al., Myosin light chain phosphorylation facilitates monocyte
transendothelial migration by dissociating endothelial adherens junctions.
Cardiovasc Res, 2011. 92(3): p. 456-65.
341. Chen, X.L., et al., VEGF-induced vascular permeability is mediated by FAK. Dev
Cell, 2012. 22(1): p. 146-57.
342. Allingham, M.J., J.D. van Buul, and K. Burridge, ICAM-1-mediated, Src- and
Pyk2-dependent vascular endothelial cadherin tyrosine phosphorylation is
required for leukocyte transendothelial migration. J Immunol, 2007. 179(6): p.
4053-64.
185 343. Nelson, C.M. and C.S. Chen, VE-cadherin simultaneously stimulates and inhibits
cell proliferation by altering cytoskeletal structure and tension. J Cell Sci, 2003.
116(Pt 17): p. 3571-81.
344. Ferber, A., et al., An octapeptide in the juxtamembrane domain of VE-cadherin is
important for p120ctn binding and cell proliferation. Exp Cell Res, 2002. 274(1):
p. 35-44.
345. Hoffmann, C., et al., Agonist-selective, receptor-specific interaction of human
P2Y receptors with β-arrestin-1 and -2. J Biol Chem, 2008. 283(45): p. 30933-41.
346. Shay-Salit, A., et al., VEGF receptor 2 and the adherens junction as a mechanical
transducer in vascular endothelial cells. Proc Natl Acad Sci U S A, 2002. 99(14):
p. 9462-7.
347. Wang, Y., et al., Integrins regulate VE-cadherin and catenins: dependence of this
regulation on Src, but not on Ras. Proc Natl Acad Sci U S A, 2006. 103(6): p.
1774-9.
348. Wojciak-Stothard, B., et al., Rho and Rac but not Cdc42 regulate endothelial cell
permeability. J Cell Sci, 2001. 114(Pt 7): p. 1343-55.
349. van Wetering, S., et al., VCAM-1-mediated Rac signaling controls endothelial
cell-cell contacts and leukocyte transmigration. Am J Physiol Cell Physiol, 2003.
285(2): p. C343-52.
350. Schrader, A.M., J.M. Camden, and G.A. Weisman, P2Y2 nucleotide receptor up-
regulation in submandibular gland cells from the NOD.B10 mouse model of
Sjogren's syndrome. Arch Oral Biol, 2005. 50(6): p. 533-40.
186 351. Mao, X., et al., A histidine-rich cluster mediates the ubiquitination and
degradation of the human zinc transporter, hZIP4, and protects against zinc
cytotoxicity. J Biol Chem, 2007. 282(10): p. 6992-7000.
352. Mayeux, R. and Y. Stern, Epidemiology of Alzheimer disease. Cold Spring Harb
Perspect Med, 2012. 2(8).
353. Haass, C. and D.J. Selkoe, Soluble protein oligomers in neurodegeneration:
lessons from the Alzheimer's amyloid β-peptide. Nat Rev Mol Cell Biol, 2007.
8(2): p. 101-12.
354. Akiyama, H., et al., Inflammation and Alzheimer's disease. Neurobiol Aging,
2000. 21(3): p. 383-421.
355. Bolmont, T., et al., Dynamics of the microglial/amyloid interaction indicate a role
in plaque maintenance. J Neurosci, 2008. 28(16): p. 4283-92.
356. Frautschy, S.A., et al., Microglial response to amyloid plaques in APPsw
transgenic mice. Am J Pathol, 1998. 152(1): p. 307-17.
357. Kim, H.J., et al., Nucleotides released from Aβ1-42-treated microglial cells
increase cell migration and Aβ1-42 uptake through P2Y(2) receptor activation. J
Neurochem, 2012. 121(2): p. 228-38.
358. Meyer-Luehmann, M., et al., Rapid appearance and local toxicity of amyloid-β
plaques in a mouse model of Alzheimer's disease. Nature, 2008. 451(7179): p.
720-4.
359. Erb, L., et al., P2Y receptors in Alzheimer's disease. Biol Cell, 2015. 107(1): p. 1-
21.
187 360. Benarafa, C., et al., SerpinB1 protects the mature neutrophil reserve in the bone
marrow. J Leukoc Biol, 2011. 90(1): p. 21-9.
361. Kolaczkowska, E. and P. Kubes, Neutrophil recruitment and function in health
and inflammation. Nat Rev Immunol, 2013. 13(3): p. 159-75.
362. Swirski, F.K., et al., Identification of splenic reservoir monocytes and their
deployment to inflammatory sites. Science, 2009. 325(5940): p. 612-6.
363. Rohrer, C., U. Arni, and K.A. Deubelbeiss, Influence of the spleen on the
distribution of blood neutrophils. Quantitative studies in the rat. Scand J
Haematol, 1983. 30(2): p. 103-9.
364. Valdes-Ferrer, S.I., et al., HMGB1 mediates splenomegaly and expansion of
splenic CD11b+Ly-6Chigh inflammatory monocytes in murine sepsis survivors. J
Intern Med, 2013. 274(4): p. 381-90.
365. Seye, C.I., et al., The P2Y2 nucleotide receptor mediates UTP-induced vascular
cell adhesion molecule-1 expression in coronary artery endothelial cells. J Biol
Chem, 2003. 278(27): p. 24960-5.
366. Vanderstocken, G., et al., P2Y2 receptor regulates VCAM-1 membrane and
soluble forms and eosinophil accumulation during lung inflammation. J Immunol,
2010. 185(6): p. 3702-7.
367. Laske, C., et al., Macrophage colony-stimulating factor (M-CSF) in plasma and
CSF of patients with mild cognitive impairment and Alzheimer's disease. Curr
Alzheimer Res, 2010. 7(5): p. 409-14.
188 368. Laske, C., et al., Decreased plasma levels of granulocyte-colony stimulating
factor (G-CSF) in patients with early Alzheimer's disease. J Alzheimers Dis,
2009. 17(1): p. 115-23.
369. Grunewald, M., et al., VEGF-induced adult neovascularization: recruitment,
retention, and role of accessory cells. Cell, 2006. 124(1): p. 175-89.
370. Salvucci, O., et al., Regulation of endothelial cell branching morphogenesis by
endogenous chemokine stromal-derived factor-1. Blood, 2002. 99(8): p. 2703-11.
371. Unoki, N., et al., SDF-1/CXCR4 contributes to the activation of tip cells and
microglia in retinal angiogenesis. Invest Ophthalmol Vis Sci, 2010. 51(7): p.
3362-71.
372. Majka, M., et al., Stromal-derived factor 1 and thrombopoietin regulate distinct
aspects of human megakaryopoiesis. Blood, 2000. 96(13): p. 4142-51.
373. Kijowski, J., et al., The SDF-1-CXCR4 axis stimulates VEGF secretion and
activates integrins but does not affect proliferation and survival in
lymphohematopoietic cells. Stem Cells, 2001. 19(5): p. 453-66.
374. Hattori, K., et al., Vascular endothelial growth factor and angiopoietin-1
stimulate postnatal hematopoiesis by recruitment of vasculogenic and
hematopoietic stem cells. J Exp Med, 2001. 193(9): p. 1005-14.
375. Sokolova, A., et al., Monocyte chemoattractant protein-1 plays a dominant role in
the chronic inflammation observed in Alzheimer's disease. Brain Pathol, 2009.
19(3): p. 392-8.
376. Westin, K., et al., CCL2 is associated with a faster rate of cognitive decline
during early stages of Alzheimer's disease. PLoS One, 2012. 7(1): p. e30525.
189 377. Cho, H., et al., Microfluidic chemotaxis platform for differentiating the roles of
soluble and bound amyloid-β on microglial accumulation. Sci Rep, 2013. 3: p.
1823.
378. Weisman, G.A., et al., Molecular determinants of P2Y2 nucleotide receptor
function: implications for proliferative and inflammatory pathways in astrocytes.
Mol Neurobiol, 2005. 31(1-3): p. 169-83.
379. Blurton-Jones, M., et al., Neural stem cells improve cognition via BDNF in a
transgenic model of Alzheimer disease. Proc Natl Acad Sci U S A, 2009. 106(32):
p. 13594-9.
380. Liu, C., et al., CXCL12/CXCR4 signal axis plays an important role in mediating
bone morphogenetic protein 9-induced osteogenic differentiation of mesenchymal
stem cells. Int J Med Sci, 2013. 10(9): p. 1181-92.
381. Zhu, W., et al., A novel regulatory role for stromal-derived factor-1 signaling in
bone morphogenic protein-2 osteogenic differentiation of mesenchymal C2C12
cells. J Biol Chem, 2007. 282(26): p. 18676-85.
382. Shahnazari, M., et al., CXCL12/CXCR4 signaling in the osteoblast regulates the
mesenchymal stem cell and osteoclast lineage populations. FASEB J, 2013.
27(9): p. 3505-13.
383. Wright, L.M., et al., Stromal cell-derived factor-1 binding to its chemokine
receptor CXCR4 on precursor cells promotes the chemotactic recruitment,
development and survival of human osteoclasts. Bone, 2005. 36(5): p. 840-53.
190 384. Kawakami, Y., et al., A small interfering RNA targeting Lnk accelerates bone
fracture healing with early neovascularization. Lab Invest, 2013. 93(9): p. 1036-
53.
385. Cao, J., et al., A comparison of stromal cell-derived factor-1 expression during
distraction osteogenesis and bone fracture in the mandible. J Craniofac Surg,
2013. 24(3): p. 805-8.
386. Alon, R., et al., The integrin VLA-4 supports tethering and rolling in flow on
VCAM-1. J Cell Biol, 1995. 128(6): p. 1243-53.
387. Rose, D.M., et al., Soluble VCAM-1 binding to α4 integrins is cell-type specific
and activation dependent and is disrupted during apoptosis in T cells. Blood,
2000. 95(2): p. 602-9.
388. Matsuda, Y., et al., Serum levels of soluble adhesion molecules in stem cell
transplantation-related complications. Bone Marrow Transplant, 2001. 27(9): p.
977-82.
389. Miwa, K., A. Igawa, and H. Inoue, Soluble E-selectin, ICAM-1 and VCAM-1
levels in systemic and coronary circulation in patients with variant angina.
Cardiovasc Res, 1997. 36(1): p. 37-44.
390. Tokuhira, M., et al., Soluble vascular cell adhesion molecule 1 mediation of
monocyte chemotaxis in rheumatoid arthritis. Arthritis Rheum, 2000. 43(5): p.
1122-33.
391. Ueki, S., et al., Soluble vascular cell adhesion molecule-1 induces human
eosinophil migration. Allergy, 2009. 64(5): p. 718-24.
191 392. Kitani, A., et al., Soluble VCAM-1 induces chemotaxis of Jurkat and synovial
fluid T cells bearing high affinity very late antigen-4. J Immunol, 1998. 161(9): p.
4931-8.
393. Garton, K.J., et al., Stimulated shedding of vascular cell adhesion molecule 1
(VCAM-1) is mediated by tumor necrosis factor-α-converting enzyme (ADAM 17).
J Biol Chem, 2003. 278(39): p. 37459-64.
394. El-Sayed, F.G., et al., P2Y2 nucleotide receptor activation enhances the
aggregation and self-organization of dispersed salivary epithelial cells. Am J
Physiol Cell Physiol, 2014. 307(1): p. C83-96.
395. Zhao, E., et al., Bone marrow and the control of immunity. Cell Mol Immunol,
2012. 9(1): p. 11-9.
396. Inoue, Y., et al., A3 and P2Y2 receptors control the recruitment of neutrophils to
the lungs in a mouse model of sepsis. Shock, 2008. 30(2): p. 173-7.
397. Chishti, M.A., et al., Early-onset amyloid deposition and cognitive deficits in
transgenic mice expressing a double mutant form of amyloid precursor protein
695. J Biol Chem, 2001. 276(24): p. 21562-70.
398. Wang, M., et al., P2Y nucleotide receptor interaction with αV integrin mediates
astrocyte migration. J Neurochem, 2005. 95(3): p. 630-40.
399. Wessel, F., et al., Leukocyte extravasation and vascular permeability are each
controlled in vivo by different tyrosine residues of VE-cadherin. Nat Immunol,
2014. 15(3): p. 223-30.
400. Chen, H.C., Boyden chamber assay. Methods Mol Biol, 2005. 294: p. 15-22.
192 401. Wirtz, S., et al., Chemically induced mouse models of intestinal inflammation. Nat
Protoc, 2007. 2(3): p. 541-6.
402. Shen, L., et al., Development of autoimmunity in IL-14α-transgenic mice. J
Immunol, 2006. 177(8): p. 5676-86.
403. Laham, R.J., et al., Spatial heterogeneity in VEGF-induced vasodilation: VEGF
dilates microvessels but not epicardial and systemic arteries and veins. Ann Vasc
Surg, 2003. 17(3): p. 245-52.
404. Murohara, T., et al., Vascular endothelial growth factor/vascular permeability
factor enhances vascular permeability via nitric oxide and prostacyclin.
Circulation, 1998. 97(1): p. 99-107.
405. Scalia, R., G. Booth, and D.J. Lefer, Vascular endothelial growth factor
attenuates leukocyte-endothelium interaction during acute endothelial
dysfunction: essential role of endothelium-derived nitric oxide. FASEB J, 1999.
13(9): p. 1039-46.
406. Gasparyan, A.Y., et al., Mean platelet volume: a link between thrombosis and
inflammation? Curr Pharm Des, 2011. 17(1): p. 47-58.
407. Bath, P.M. and R.J. Butterworth, Platelet size: measurement, physiology and
vascular disease. Blood Coagul Fibrinolysis, 1996. 7(2): p. 157-61.
408. Yesil, Y., et al., Increased mean platelet volume (MPV) indicating the vascular
risk in Alzheimer's disease (AD). Arch Gerontol Geriatr, 2012. 55(2): p. 257-60.
409. Ahn, H.J., et al., A novel Aβ-fibrinogen interaction inhibitor rescues altered
thrombosis and cognitive decline in Alzheimer's disease mice. J Exp Med, 2014.
211(6): p. 1049-62.
193 410. Cortes-Canteli, M., et al., Fibrinogen and β-amyloid association alters thrombosis
and fibrinolysis: a possible contributing factor to Alzheimer's disease. Neuron,
2010. 66(5): p. 695-709.
411. Perez, L.E., et al., Increased plasma levels of stromal-derived factor-1 (SDF-
1/CXCL12) enhance human thrombopoiesis and mobilize human colony-forming
cells (CFC) in NOD/SCID mice. Exp Hematol, 2004. 32(3): p. 300-7.
412. Niswander, L.M., et al., SDF-1 dynamically mediates megakaryocyte niche
occupancy and thrombopoiesis at steady state and following radiation injury.
Blood, 2014. 124(2): p. 277-86.
194 VITA
Chen (Cheery) Cao was born on May 19, 1986, in Jiangshan, Zhejiang Province,
China. After attending the Jiangshan High School in her hometown, she studied in the
Department of Biological Sciences, Life Sciences College at Zhejiang University in
Hangzhou, China, and received a B.S. degree in 2008. Since 2008, she has been a Ph.D. student in the Department of Biochemistry at the University of Missouri-Columbia, USA, under the supervision of Dr. Gary A. Weisman and Dr. Laurie Erb.
195