Ucin968165255.Pdf (9.99

UNIVERSITY OF CINCINNATI

______May 18 , 20 00_____

I, Xiuqiong Wang , hereby submit this as part of the requirements for the degree of: Doctor of Philosophy in: Department of Molecular & Cellular Physiology

It is entitled: Investigation of the Role of Annexin V in Mouse Placenta: Development of Approaches to Explore the Therapeutic Potential of the Protein

Approved by: John R. Dedman, Ph.D., Chairman Robert O. Banks, Ph.D. Maria F. Czyzyk-Krzeska, M.D., Ph.D. Robert F. Highsmith, Ph.D. Gregory S. Retzinger, M.D., Ph.D. Investigation of the Role of Annexin V in Mouse Placenta: Development of Approaches to Explore the Therapeutic Potential of the Protein

A Dissertation submitted to the Division of Graduate Studies and Research of the University of Cincinnati

in partial fulfillment of the requirement for the degree of

DOCTOR OF PHILOSOPHY

in the Department of Molecular and Cellular Physiology of the College of Medicine

1999 By

XIUQIONG WANG

M.D., 1985, West China University of Medical Sciences, Chengdu, China M.S., 1988, West China University of Medical Sciences, Chengdu, China

Advisor: John R. Dedman, Ph.D., Ohio Eminent Scholar ABSTRACT

Annexin V is a promising candidate as an anticoagulant and anti-inflammatory agent due to its high affinity for anionic phospholipid surfaces. Its abundance and tissue localization in placental trophoblasts and vascular endothelial cells imply a physiological importance in these tissues. In the first section of this study, I demonstrate that annexin V is critical in maintaining murine placental integrity. I conclude that the protecting role of annexin V appears to be local because I did not detect the protein in culture media of either trophoblasts or endothelial cells. In the next portion of this study, I successfully targeted annexin V to the secretory pathway of mammalian cells and the protein was secreted into the medium. This unnatural targeting of annexin V to the lumen of the endoplasmic reticulum did not have significant effect on cellular function. Therefore, I concluded that targeting secretion of annexin V in transgenic animals should provide a valuable model for evalu- ating the therapeutic potentials of annexin V as anticoagulant and/or anti-inflammatory agent.

Acknowledgement

I would like to dedicate my thesis to my family for their endless love and support: My father Xijie Wang; My mother Xuehua Chen; My sister Xiuyun Wang; My brothers Jun Wang and Kai Wang; My nephews Jao Yang and Yifan Wang.

Very special thanks to Charles William Hall Jr, the love of my life. Table of Contents

CHAPTER I ...... 4 INTRODUCTION ...... 4 CHAPTER II...... 17 ANNEXIN V IS CRITICAL FOR THE MAINTENANCE OF MURINE PLACENTAL INTEGRITY ...... 17 INTRODUCTION ...... 17 MATERIALS AND METHODS ...... 18 RESULTS ...... 21 COMMENTS ...... 24 CHAPTER III ...... 35 DETECTION OF EXTRACELLULAR ANNEXIN V AND ATTEMPTS TO SECRET RECOMBINANT ANNEXIN V FROM CULTURED CELLS ...... 35 INTRODUCTION ...... 35 MATERIALS AND METHODS ...... 35 RESULTS AND DISCUSSION ...... 40 CHAPTER IV ...... 53 TRANSGENIC MANIPULATION TO TARGET THE SECRETION OF ANNEXIN V INTO THE CIRCULATION OF MICE ...... 53 INTRODUCTION ...... 53 MATERIALS AND METHODS ...... 55 RESULTS AND DISCUSSION ...... 58 CHAPTER V ...... 66 TARGETING ANNEXIN V TO THE SECRETORY PATHWAY OF THYROID EPITHELIAL CELLS DOES NOT SIGNIFICANTLY ALTER CELLULAR FUNCTIONS ...... 66 INTRODUCTION ...... 66 MATERIALS AND METHODS ...... 68 RESULTS ...... 73 DISCUSSION ...... 88 CHAPTER VI ...... 91 CONCLUSIONS ...... 91 BIBLIOGRAPHY...... 95

1 List of Figures

Figure 1.1. Basic structure of annexin V...... 5 Figure 1.2. The intrinsic, extrinsic and common coagulation pathways. (HMW-K=high molecular-weight kininogen; PL=phospholipid). Adapted from Highsmith, RF, in Speralakis and Banks, 1993)...... 9 Figure 2.1. Purification of recombinant epitope-tagged annexin V...... 25 Figure 2.2. Mouse placental histology (4X)...... 26 Figure 2.3. Identification of extracellular binding sites for annexin V in placenta...... 27 Figure 2.4. Immunolocalization of endogenous annexin V in placenta...... 28 Figure 2.5. Clotting assays (aPTT) with annexin V and anti-annexin V alone or in combination...... 29 Figure 2.6. Time course of anti-annexin V in the circulation after tail-vein infusion. .... 30 Figure 2.7. Effects of circulating anti-annexin V antibody on pregnancy: gross morphology...... 31 Figure 2.8. Effects of circulating anti-annexin V antibody on pregnancy: ...... histopathology...... 32 Figure 3.1. Localization of annexin V in rat placental trophoblasts and blood vessel endothelial cells...... 47 Figure 3.2. Immunofluorescent staining of annexin V in permeabilized BeWo cells and immunoblot detection of annexin V in culture media...... 48 Figure 3.3. Immunofluorescent localization of annexin V in permeabilized HUVEC and immunoblot analysis detection of annexin V in culture media...... 49 Figure 3.4. FITC-annexin V binding and immunofluorescent staining of annexin V on the surface of non-permeabilized BeWo cells...... 50 Figure 3.5. FITC-annexin V binding and localization of cell surface annexin V in human umbilical vein endothelial cells (HUVEC)...... 51 Figure 3.6. Expression and secretion of annexin V from COS-7 cells...... 52 Figure 4.1. Transgene that is under control of PEPCK promoter...... 60 Figure 4.2. PCR identification of transgenic mice...... 61 Figure 4.3. Transgene that is under control of the albumin enhancer/promoter...... 62 Figure 4.4. PCR analysis of transgenic mice...... 64 Figure 4.6. RT-PCR analysis of transgenic mice...... 65

2 Figure 5.1. The transgene designed to secrete annexin V via the ER/Golgi secretory pathway...... 77 Figure 5.2. Schematic description of tetracycline-inducible system (tet-on)...... 78 Figure 5.3. Comparison of the localization of ER-targeted and endogenous annexin V in PCrTTA7 cells using indirect immunofluorescent microscopy...... 79 Figure 5.4. Western blot analysis of annexin V secretion from PCrTTA7 cells...... 80 Figure 5.5. Cell growth curves evaluated by MTT assay...... 81 Figure 5.6. Equilibrium 45Ca2+ uptake and release by thapsigargin...... 82 Figure 5.7. 45Ca2+ uptake and release in saponin-permeabilized cells...... 83 Figure 5.8. The synthesis of thyroglobulin in non-induced and induced cells...... 84 Figure 5.9. The evaluation of thyroglobulin processing in non-induced and doxycycline-induced cells...... 85 Figure 5.10. Western blot analysis of thyroglobulin and chaperonin proteins in non-induced and induced cells...... 86 Figure 5.11. Pulse chase analysis to assess thyroglobulin secretion...... 87 Figure 6.1. The schematic demonstration of the role annexin V might play in the placenta. Adapted from Rand et al, New England Journal of Medicine, 1997...... 91

3 CHAPTER I

INTRODUCTION

History and Structure of Annexin V Annexin V is a member of a structurally related family of proteins that bind to phospholipid surfaces in the presence of calcium. It is a single chain polypeptide containing 319 amino acids with a molecular mass of 35 kDa (Iwasaki, 1987, Funakoshi, 1987a). The calcium phospholipid binding sites are within the four repeated domains of the protein (Figure 1.1). This core structure of 70 amino acid repeats is the most conserved element of all annexins. In contrast, the amino terminal region varies among the annexins suggesting that that region determine the specific functional role of a given annexin (Crompton, 1988, Barton, 1991).

Annexin V was first isolated from placenta and termed placental protein 4 (PP4) (Bohn, 1979). The protein was independently identified and called many other names such as 35 γ-calcimedin (Kaetzel, 1989), vascular anticoagulant protein α (VACα) (Reutelingsperger, 1985), placental anticoagulant protein I (PAP-I) (Funakoshi, 1987b), endonexin II (Kaplan, 1988), lipocortin V (Pepinsky, 1988), and anchorin II (Mollenhauer,

1984). Annexin V is the most abundant of the annexins and has the broadest expression pattern in the annexin family (Kaetzel, 1989). It is located in many tissues, with the highest concentration being in vascular endothelial cells, placental trophoblasts, exocrine, and en- docrine organs (Romisch, 1992, van Heerde, 1995).

Proposed Functions of Annexin V Physiological functions of annexin V have been of scientific research interest for about two decades. While much has been done to elucidate the protein’s pathophysiologi-

4 Conserved Repeats Calcium and Phospholipid Binding Domains

N C

Variable N-Term inal Domain

Figure 1.1. Basic structure of annexin V. Annexin V, like other annexins, has four conserved repeats that are calcium-dependent phospholipid binding sites. The N-terminus is the variable region.

5 cal significance, the precise function of annexin V in vivo remains unclear. The primary physical property of annexin V is that it binds calcium and phospholipids, especially negatively charged phospholipids such as phosphatidylserine (PS), phosphatidic acid (PA) and phosphoethanolamine (PE). The calcium binding affinity of annexins is relatively low (Kd>5x10-4M) compared with the E-F hand proteins (calmodulin, troponin C and S100 et al), but the affinity for calcium increases in the presence of phosphatidylserine (Schlaepfer, 1987). For example, annexin V binds to model membrane containing 20% PS with an affinity less than 10-10M in the presence of 3mM Ca2+ (Andree, 1990, Tait, 1989).

Many proposed functions of annexin V are based on the protein’s calcium-dependent phospholipid binding activities. Annexin V inhibits coagulation reactions in vitro by competing with coagulation factors for anionic phospholipid surfaces (Funakoshi, 1987b).

Annexin V also inhibits phospholipase A2 (PLA2) by depleting negatively-charged phospholipids, which are substrates for PLA2. Arachidonic acid release and prostaglandin formation are blocked (Ahn, 1988, Van Bilsen, 1992, Buckland, 1998, Mira, 1997). Thus, annexin V was postulated to inhibit inflammatory responses. Annexin V was also found to inhibit protein kinase C (PKC)-mediated phosphorylation of annexin I, II and other PKC substrates (Dubois, 1996). The mechanism of this inhibition is not yet clear, but is thought to involve phospholipid depletion (Raynal, 1993) and/or direct interaction with PKC (Schaepfer, 1992). Finally, annexin V (called anchorin II in bones) was proposed to be involved in cartilage calcification. The protein is localized on the outer cell surface of hypertrophic chondrocytes, chondrocyte microvilli, fibroblasts and matrix vesicles. This localization, together with the protein’s collagen binding property and a proposed intrinsic calcium channel activity, suggest that annexin V may promote matrix vesicle initiated cartilage calcification (von der Mark, 1997).

6 Annexin V: An Anticoagulant Protein? The most studied and well-documented activity of annexin V is that of anticoagulant. There are two reasons to believe that annexin V is involved in anticoagulation. First, annexin V is abundant in cells that are constantly in contact with blood (Kaetzel, 1989, Flaherty, 1990). Vascular endothelial cells and placental trophoblasts can contain as much as 3.6 x 10-12 g/cell annexin V, or about 0.2% of their total protein (van Heerde, 1995). Second, as a calcium-dependent phospholipid binding protein, annexin V has a 1000-fold higher affinity for anionic phospholipids than coagulation factors (Tait, 1989, Andree, 1992, Sun, 1993a). These factors, such as factor VII, V and X, require calcium and negatively- charged phospholipids for activation (Figure 1.2). Therefore, annexin V can deplete anionic phospholipids and inhibit the activation of coagulation factors. In vitro studies have shown that annexin V significantly prolongs the activated partial thromboplastin time (aPTT) and the dilute Russel’s viper venom time (dRVVT). The aPTT tests the integrity of the intrinsic coagulation pathway and the dRVVT tests the assembly of the prothrombinase complex that catalyzes the formation of thrombin (Andree, 1992, Van Heerde, 1994a, Reutelingsperger, 1988, Almus, 1993, Hauptmann, 1989, Campos, 1998).

The ability of annexin V to interfere with the tissue factor-factor VII complex is controversial. Some studies show that annexin V inhibits the formation of this complex

(Van Heerde, 1994a, Kondo, 1987), while others show that annexin V fails to do so (Rao, 1992, Almus, 1993) or that this inhibition happens only when very high concentrations of annexin V are applied (Ravanat, 1992).

The coagulation inhibitor protein C is also activated on the surface of endothelial cells. However, studies show that activation of protein C is only slightly inhibited by annexin V (Ravanat, 1992,) whereas the activation of protein S is significantly inhibited by annexin V on the surface of activated platelets (Sun, 1993a). Annexin V strongly inhibits the inac-

7 tivation of factor Va by activated protein C/protein S complex (Sun, 1993b). Collectively, these studies suggest that activation of protein C is probably independent of anionic phospholipids while the formation of activated protein C/protein S complex requires the avail- ability of negatively charged phospholipid surfaces. The fact that annexin V can be involved in both anticoagulation and inactivation of activated factors by protein C/protein S pathway suggests multi-dimensional roles of annexin V in vivo.

In vivo and ex vivo experiments have been performed to test the anticoagulant activities of annexin V. So far, all these studies have shown that annexin V is an inhibitor of thrombin formation and fibrin deposition under arterial and venous blood flow. In a laser and a photochemical-induced thrombus model in rat mesenteric arterioles and venules, infusion of annexin V significantly inhibited thrombus formation at a dose level of 0.3 mg/kg (Romisch, 1991). In an ex vivo model where blood from the anticubital vein of healthy volunteers passes directly over extracellular matrix of endothelial cells stimulated by tumor necrosis factor (TNF-ECMs), pre-incubation of the endothelial cells with annexin V significantly inhibited fibrin deposition and platelet-matrix adhesion (van Heerde, 1994).

The potency of annexin V as an anticoagulant in vitro has also been compared to heparin in the in vivo models. Chollet, et al., demonstrated that annexin V inhibited in-

traocular postoperative fibrin formation, and it is as potent as heparin (Chollet, 1992). In another experiment, Van Ryn-McKenna, et al., showed that on the injured jugular vein of a rabbit, annexin V, but not prophylactic doses of heparin (20U/kg), inhibited fibrin deposition on the surface of de-endothelialized vessel wall. This finding indicates that annexin V is a more potent anticoagulant than heparin (Van Ryn-McKenna, 1993). A more recent study of chemically induced thrombus model on rabbit carotid artery provided further evidence to support annexin V as a vascular anticoagulant (Thiagarajan, 1997). Further studies of annexin V in the whole animal are needed to evaluate the true role annexin V plays in hemostasis.

8 Figure 1.2. The intrinsic, extrinsic and common coagulation pathways. (HMW-K=high molecular-weight kininogen; PL=phospholipid). Annexin V inhibits the tenase complex and the prothrombinase complex thereby prolonging in vitro coagulation tests (activated partial throboplastin test and dilute Russel’s viper venom test). Protein C is activated on lipid surfaces of endothelial cells and platelets. Activated protein C, together with its cofac- tor, protein S, cleaves factors Va and VIIIa. Anionic phospholipids are required for these activities. Adapted from Highsmith, RF, in Speralakis and Banks, 1993).

9 Activated platelets are important sources of negatively-charged phospholipids involved in the activation of coagulation factors. When blood vessels are injured, platelets adhere to the exposed sub-endothelium and are activated by collagen and small amounts of thrombin. The activated platelets undergo a series of changes including the reorganization of surface membrane phospholipids, and the exposure of phosphatidylserine for the activation of coagulation factors. The increase of annexin V binding sites from 5000 on a non- stimulated platelet to 200,000 on thrombin-stimulated and other agonists-stimulated platelet, suggest that phosphatidylserine is externalized (Sun, 1993a, Thiagarajan, 1990). The binding of annexin V does not have an effect on collagen and thrombin-induced platelet aggregation, and does not cause platelet aggregation by itself (Thiagarajan, 1990). How- ever, as mentioned above, the binding of protein S to the platelet surface is significantly inhibited (Sun, 1993a).

Binding of annexin V to the activated platelet surface also significantly inhibits factor IXa-catalyzed factor X activation. The mechanism involves depletion of negatively- charged phospholipids (London, 1996) and inhibiting the binding of factor Xa to thrombin- stimulated platelets (Thiagarajan, 1990). Exposure of phosphatidylserine on the platelet surface provides a tool for targeting platelet-containing thrombi with annexin V. This possibility was evaluated by Tait et al, who employed iodinated annexin V to target arterial thrombi and found that platelet-containing thrombi selectively take up annexin V (Tait,

1994). These studies were continued using 99mTc-labeled human annexin V to detect acute left atrial thrombi in vivo in swine (Stratton, 1995). The specific interaction between annexin V and activated platelets prompted the notion that annexin V could be used to target throm- bolytic agents to arterial thrombi. The feasibility of this proposal was explored by fusing annexin V with pro-urokinase. The chimeric protein possesses urokinase activity as well as specific ability to bind to membranes containing phosphatidylserine (Tait, 1995).

10 Annexin V is also present in platelets, but it is not located on the secretory granules nor can it be released upon stimulation by thrombin or other agonists (Murphy, 1992, Flaherty, 1990, Trotter, 1994). However, thrombin stimulation causes intracellular relocation of annexin V in platelets (Trotter, 1994). Upon stimulation, platelet annexin V associates tightly with plasma membrane and is EGTA-resistant. Triton X-100 treatment can extract EGTA-resistant annexin V (Trotter, 1994, Trotter, 1993). The significance of this relocation is not clear. The involvement of annexin V in membrane processing, such as phospholipid reorganization and platelet secretion, was suggested by Trotter, et al (Trotter, 1994).

The placental trophoblast is another site where annexin V may be actively involved in the inhibition of blood coagulation. The syncytiotrophoblast is the outer most layer of fetal cells in the placenta. It is in contact with maternal blood, and is a site of nutritional exchange between mother and fetus. Because of cell fusion, syncytiotrophoblasts also provide an important barrier between maternal and fetal circulations. In order to efficiently supply the fetus with nutrients from maternal blood, the maternal blood pools in intravillous spaces (as in human placenta) or sinuses in fetal tissue labyrinthine (as in mouse placenta) and flows slowly across the surface of syncytiotrophoblasts. The purpose is to maximize contact area and time between maternal blood and syncytiotrophoblasts for the adequate exchange of metabolites.

Syncytialization of the trophoblasts is accompanied by externalization of anionic phospholipids as observed in the fusion of myoblasts (Sessions, 1983). Externalization was confirmed using various approaches (Rote, 1995, Lyden, 1992, Lyden, 1993, Katsuragawa, 1995). The slowly moving blood and the possible externalization of anionic phospholipids on the surface of syncytiotrophoblasts create an ideal environment for blood clot formation. In addition, pregnancy is usually accompanied by an increase in plasma concentration of several coagulants and no correspondent increase of coagulation inhibitors (Delmore, 1992).

11 Thrombin generation and fibrin deposition in the placenta are increased and there must be a local regulatory mechanism to maintain the hemostatic balance (Delorme, 1992). There- fore, a sophisticated anticoagulation system is required to maintain blood circulation in the placenta.

So far, both in vitro and in vivo experiments have suggested that annexin V is involved in anticoagulation in the placenta. When cell surface annexin V is neutralized by anti-annexin V antibodies, the coagulation time performed on the cell surface is significantly shorter than when annexin V is present (Rand, 1997a). Findings in the whole animal support this observation: infusion of anti-annexin V antibodies into Balb/c mice caused fetal reabsorption, placental thrombosis and inflammation (Wang, 1999). Antiphospholipid syndrome is a hypercoagulable condition characterized by arterial and venous thrombosis and recurrent fetal loss with circulating autoantibodies against anionic phospholipids. Ex- actly how antiphospholipid autoantibodies cause thrombotic events remains unknown (Triplett, 1993). Antiphospholipid antibodies, however, have been found to remove trophoblast surface annexin V and to facilitate thrombin binding (Vogt, 1997).

Studies performed using the BeWo trophoblast cell line and cultured primary placental villi by Rand, et al., suggested that annexin V plays a protective role in the placenta by “shielding” anionic phospholipids. This shielding prevents the catalysis of coagulation reactions by phospholipid surfaces, and antiphospholipid antibodies cause placenta thrombosis by removing annexin V from trophoblasts (Rand, 1997a, Rand, 1997b, Rand, 1994). Furthermore, the physiological role of annexin V was implied when it was found to be deficient in placentas from patients with antiphospholipid syndrome, characterized by hypercoagulablity with thrombotic events in the placenta and other organs (Rand, 1994, Devine, 1996). Similar results were found with cultured endothelial cells after treating the cells with antiphospholipid antibodies isolated from patients (Rand, 1997a). A new class of

12 diseases supposedly caused by abnormality of annexins was proposed recently, and defi- ciency of annexin V was thought to be the mechanism of thrombosis seen in placenta from patients with antiphospholipid syndrome (Rand, 1999).

The above results were challenged by the work of Lakasing, et al., who demonstrated that there was no change in the concentration of annexin V or tissue factor and thrombomodulin between placentas from antiphospholipid syndrome and those from normal controls (Lakasing, 1999). The exact mechanism by which annexin V and antiphospholipid antibodies interact in the placenta is speculative. The cellular mechanism of providing cell surface annexin V is not understood (Rote, 1997, Cheng, 1997).

Extracellular Annexin V To function as an anticoagulant that works by competing for anionic phospholipids with coagulation factors, the protein needs to be extracellular. The annexins do not possess a traditional secretory signal sequence (Funakoshi, 1987a). It is generally accepted that the annexins are intracellular proteins. There are mixed and controversial reports of extracellular annexin V. Annexin V was detected in conditioned medium of cultured endothelial cells, in amniotic fluid and in platelet-rich plasma after the stimulation of platelets with arachidonic acid (Flaherty, 1990). Low amounts of annexin V were also found in the serum and plasma of healthy volunteers. The reported levels range from 0 to 50 ng/ml. The minimum concentration of annexin V that inhibits blood coagulation is about 50 ng/ml (Shirotake, 1986, Goulding, 1990, Uemura, 1992, Tait, 1988). Studies have also shown that annexin V mRNA levels increase when cells are treated with phorbol myristate acetate (PMA) and dexamethasone, suggesting that annexin V may be involved in anti-inflammatory processes (Solito, 1991).

13 Some investigators have argued that if annexin V is detectable in the blood, it might be released from damaged annexin V-abundant cells rather than actively secreted (Raynal, 1994). However, the reported results of extracellular annexin V are all based on the study of cells exceptionally rich in annexin V. As previously mentioned, annexin V in vascular endothelial cells makes up as much as 0.2% of total cellular protein (van Heerde, 1995). In support of this theory, elevated plasma annexin V was detected in patients suffering from ischemic damage (Relton, 1991, Romish, 1992, Kaneko, 1996). Annexin V was also proposed to be one of the new markers for early detection of myocardial damage (Mair, 1997). The study of non-stimulated and various agonist-stimulated platelets failed to show release of annexin V from cells, even though annexin V was found to be relocated and tightly associated with the platelet membrane (Murphy, 1992, Flaherty, 1990, Trotter, 1994). How- ever, the inability to detect lactate dehydrogenase (LDH) level in the medium excluded substantial cell lysis (Flaherty, 1990).

Several studies have shown annexin V to be present on cell surfaces of cultured trophoblasts and placental villi (Rand, 1994, Vogt, 1997). Annexin V was also found on hypertrophic chondrocytes as well as the surfaces of microvilli using immunofluorescence, immunoelectronmicroscopy and cell surface iodination (von der Mark, 1997). It has been proposed that the intrinsic channel activity of annexin V mediates Ca2+ influx into chondrocytes. Annexin V has been localized to the perimatrix of hypertrophic cartilage

where it is thought to be the receptor for certain types of collagen and mediates Ca2+ flux into matrix vesicles, promoting the initiation of cartilage matrix mineralization (Mollenhauer, 1999, Pfaffle, 1988). The above findings are consistent with the notion that annexin V is on the surface of apoptotic cells, where it prevents phospholipid-initiated coagulation and inflammation (Boersma, 1996, Van Engeland, 1996). Experiments have also suggested that the human liver cell membrane possess annexin V, and on that cell the protein is a very specific hepatitis B virus (HBV) receptor protein (Hertogs, 1994).

14 Most of the findings described above suggest that annexin V is extracellular, either in culture media or on cell surfaces. While lack of a hydrophobic signal sequence makes it unlikely the protein is processed and secreted by the endoplasmic reticulum (ER) and Golgi, alternative secretory routes have been suggested. Many proteins are readily secreted into extracellular spaces in spite of lacking a hydrophobic signal peptide. Annexin I and annexin II, other members of annexin family without signal peptides, have been detected on cell surfaces, both in culture medium and in prostate fluid (annexin I) (Christmas, 1991, Serres, 1994, Siever, 1997). Proteins other than annexins, e.g., endothelial cell growth factors (Jaye, 1986), interleukin-1 (Andrei, 1999, Corradi, 1995) and fibroblast growth factor 2 (Albuquerque, 1998), are also secreted in the absence of a signal peptide. The proposed route of secretion of interleukin-1 is exocytosis of preterminal endocytic vesicles (Andrei, 1999).

Hypotheses of this Thesis Project Annexin V is important to the maintenance of placental integrity. To test this hypothesis, I established the possible roles of annexin V in the placenta by neutralizing the protein with anti-annexin V antibodies followed by the evaluation of the fetal viability and placental integrity.

Annexin V is secreted or released from mammalian cells. To test this hypothesis, I set out to detect extracellular annexin V in culture media and on cell surfaces of trophoblasts and endothelial cells, two cell types rich in endogenous annexin V.

Annexin V can be secreted to the circulation of transgenic animals. To test this hypothesis, I targeted the annexin V gene to hepatocytes of transgenic mice and expression and secretion of the transgene product were analyzed.

15 Targeting annexin V to the secretory pathway of cells affects cellular functions. To test this hypothesis, I stably transfected the annexin V gene into thyroid epithelial cells and ER functions of those cells were analyzed in terms of calcium homeostasis and protein synthesis, processing and secretion.

16 CHAPTER II

ANNEXIN V IS CRITICAL FOR THE MAINTENANCE OF MURINE PLACENTAL INTEGRITY

INTRODUCTION

Patients with some autoimmune diseases often suffer from a syndrome of early fetal loss, thrombosis and thrombocytopenia. This syndrome, thought to be caused by antibodies against anionic phospholipids such as phosphatidylserine and cardiolipin, is termed the antiphospholipid syndrome (APS) (Hughes, 1983). However, some patients with APS test negative for antiphospholipid antibodies but produce immunoglobulins that are pathogenic to murine pregnancy (Silver, 1997). Thus, other autoantibodies may be involved in causing APS.

Anti-annexin V antibody has been detected in patients with diseases such as systemic lupus erythematosus (SLE), rheumatic arthritis (RA), preeclampsia and habitual fetal loss (Kaburaki, 1997, Rodriguez-Garcia, 1996, Matsuda, 1994a, Matsuda, 1994b). These findings suggest that anti-annexin V autoantibodies may be the cause of thrombotic events in patients, possibly by interfering with an anticoagulant function of annexin V. However, it has been reported that anti-annexin V antibodies in vitro prolonged the activated partial thromboplastin time (aPTT), similar to antiphospholipid antibody and lupus anticoagulant (Nakamura, 1995). The high concentration of annexin V in placental trophoblasts has been proposed to maintain blood flow on the constantly regenerating surface of trophoblasts so that maternal and fetal nutrition exchange and fetal viability are unimpaired (Krikrun, 1994). In fact, it has been reported that the level of annexin V is reduced on placental villi of women with recurrent spontaneous pregnancy loss, especially those with APS (Rand, 1994,

17 Rand, 1997a). Antiphospholipid antibodies also were shown to decrease the level of annexin V on the surfaces of cultured trophoblasts and placental villi (Rand, 1997, Vogt, 1997).

With these known interactions, the role of annexin V and anti-annexin V in the placenta were explored. I demonstrate the presence of binding sites for annexin V on the surface of placental trophoblasts and investigate the consequence of passive infusion of anti-annexin V antibodies on the pregnancy of BALB/c mice. Anti-annexin V autoantibodies may be pathogenic and cause clinical presentations consistent with APS.

MATERIALS AND METHODS

Preparation of immunoglobulins. Immune sera from rabbits immunized with annexin V were collected, and anti-annexin V (IgG) was purified by affinity chromatography using a Sepharose 4B column covalently linked to annexin V (Dedman, 1978). Anti- annexin V IgG was stored in phosphate-buffered saline (PBS) at 4oC. Nonimmune rabbit IgG was isolated from normal rabbit serum.

Clotting assay. The activated partial thromboplastin time (aPTT) was used to evaluate if anti-annexin V antibody blocks the effects of annexin V in vitro. aPTT was measured using citrated (13 mM) pooled normal human plasma, and Dade actin FSL according to a modification of Fritsma (Fristma GA, 1989). The prolongation of aPTT by annexin V was measured as described elsewhere (Campos, B, 1998). Annexin V (10 µM) and anti-annexin V (100 µM) were prepared in PBS and mixed with the aPTT reagent, which was diluted 2.5 times in order to increase the sensitivity of the assay. PBS was used as a negative control.

Preparation of FLAG-tagged annexin V. In order to distinguish infused annexin V from endogenous annexin V, an epitope-tagged annexin V was constructed. FLAG is an

18 eight amino acid peptide epitope (Eastman Kodak). Annexin V cDNA was cloned into the bacterial expression system pFLAG-CTC (Eastman Kodak) and the fusion protein was expressed in E. coli. Two hours after induction with isopropyl-β-D-thiogalactopyranoside (IPTG), the bacteria were pelleted and lysed to release the fusion protein. FLAG-tagged annexin V from the supernatant of the lysed bacteria was affinity purified as described elsewhere (Kaetzel, 1989). The bacterial extracts were applied to a column of phenyl- Sepharose that had been treated with brain phospholipids (Sigma). After many washes with buffers containing salt concentrations 0.5 M and 1.0 M in the presence of 1 mM CaCl2, FLAG-annexin V fusion protein was eluted with (ethylene glycol-bis (β-aminoethyl ether) N,N,N’,N’-tetraacetic acid, EGTA). The protein eluted from the affinity column was subjected to sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE), and then either stained with Coomassie Blue or transferred to nitrocellulose. The membrane was probed with either mouse anti-FLAG monoclonal antibodies (Eastman Kodak) or affinity purified rabbit anti-annexin V antibodies, and peroxidase-conjugated secondary antibodies sequentially. Before infused into mice, FLAG-annexin V was dialyzed against normal saline.

Determination of the serum level of anti-annexin V antibodies after infusion. Three-month old mice were tail vein-infused with 10 µg anti-annexin V antibody. Blood samples were collected using retro-orbital bleeding at 4, 17, 24, 48, 72, 96, 120 and 144 hours after infusion. Two mice were used at each time point. Sera were obtained and stored at -20oC. Serum levels of anti-annexin V were detected using Enzyme-Linked Immunosorbent Assay (ELISA). The values were determined using a standard curve gener- ated with rabbit anti-annexin V antibody prepared in normal mouse serum. ELISA plates were coated with annexin V overnight at room temperature. After blocking (0.25% BSA and 0.05% Tween-20 in PBS), samples were added to the plate, incubated at room temperature for an hour, and then exposed to peroxidase-conjugated goat anti-rabbit secondary an-

19 tibodies for another hour. The peroxidase activity was visualized with H2O2 and O-phe- nylenediamine. The optical density was determined using a kinetic microtiter-plate reader set to a wavelength of 450 nm.

Evaluation of fetal and placental development. Antigen affinity-purified rabbit anti-annexin V antibodies and normal rabbit IgG were prepared in normal saline to a concentration of 0.1 µg/µl immediately prior to infusion. Ten micrograms of either anti-annexin V or preimmune IgG was infused into the tail veins of 3 month-old BALB/c mice (Jackson Laboratory) using a 27-gauge needle. Female mice infused with affinity-purified anti-annexin V antibodies, control females infused with normal rabbit IgG and saline-treated females were mated with three-month old males immediately after the infusion, in the late after- noon. Coitus was confirmed by observation of vaginal plugs the following morning. Ten out of 12 anti-annexin V-infused, 4 out of 5 pre-immune IgG-infused and 4 out of 6 saline- treated mice had vaginal plugs 16 hours after mating. Only those mice with vaginal plugs were chosen for the experiments.

In order to evaluate the progression of the effects of anti-annexin V antibody on pregnancy, the uteri, fetuses and placentas were surgically harvested and photographed at day 11 or 15 of confirmed pregnancy. The number of intact embryos was counted in both experimental and controls groups. It is impossible to accurately count the number of re- absorptions because in most cases, there are increased empty spaces between intact embryos, especially after day 15 of pregnancy. The intact uteri with fetuses and placentas were fixed in 10% PBS-buffered formalin for 20 hours and then embedded in paraffin. Four- micron thick sections were stained with haematoxylin-eosin (H & E). Paraffin embedded sections of the same thickness were also prepared for immunolocalization studies. In a separate experiment, 150 µg of FLAG-annexin V in normal saline was infused into two 15- day pregnant mice. Another two 15-day pregnant mice were infused with normal saline.

20 Mice infused with FLAG-annexin V and normal saline were sacrificed after 3 or 20 hours. Uteri were collected and fixed, and paraffin embedded sections were prepared as described above.

Immunohistology. Localization of annexin V in mouse placenta was determined by immunoperoxidase-staining using the VECTASTAIN Elite ABC peroxidase system (Vec- tor Laboratories, USA). Paraffin-embedded tissue sections were deparaffinized, hydrated, and blocked with 0.3% hydrogen peroxide to quench endogenous peroxidase activity. The sections were then incubated with affinity-purified sheep anti-annexin V antibodies followed by peroxidase-conjugated goat anti-sheep secondary antibodies. The tissue sections incubated only with secondary antibodies served as controls. The same immunoperoxidase staining system was employed to detect the FLAG-tagged annexin V in placental tissues except that the first antibody was rabbit anti-FLAG (Zymed). The tissue sections from normal saline-infused mice were used as controls.

Statistical analysis. Unpaired Student’s t-test was used to compare the number of intact embryos between groups. p < 0.05 was considered significant.

All procedures involving mice were approved by the Institutional Animal Care and

Use Committee.

RESULTS

Expression and purification of FLAG-tagged annexin V. In order to distinguish infused annexin V from endogenous annexin V, the protein was tagged with an eight amino acid epitope, FLAG. Coomassie Blue staining showed that a single 36 kDa protein was present in the elution fractions from phenyl Sepharose column treated with brain phospho-

21 lipids. The molecular weight of wild type recombinant annexin V without the epitope tag is 35 kDa (Figure 2.1). The fusion protein from the elution was confirmed to be FLAG- annexin V by immunoblotting with both affinity purified anti-annexin V antibodies and anti-FLAG monoclonal antibodies (data not shown).

Murine placental tissue contains annexin V binding surfaces. Similar to the human placenta, mouse placenta is hemochorial, i.e., fetal trophoblasts are directly bathed in maternal blood (Rossant, J, 1985). However, mouse placenta is histologically distinct from human placenta in maternal and fetal tissue interdigitation. Instead of being villous as in humans, mouse placenta is labyrinthine. Murine fetal tissue forms sinuses that are filled with maternal blood (Figure 2.2). Mice, 15 days pregnant, were infused with FLAG-tagged annexin V. The tagged annexin V was localized on the surfaces of trophoblasts which form the sinuses containing maternal blood (Figure 2.3A). There was no reaction to anti-FLAG antibodies in placental tissues from saline-infused mice (Figure 2.3B). These results demonstrate annexin V binding sites on the surfaces of trophoblasts.

Immunohistological localization of endogenous annexin V in mouse placenta. The placental tissue from mice at day 15 of pregnancy was stained for endogenous annexin V. Annexin V was concentrated along the surfaces of sinuses formed by fetal tissue. The

localization of endogenous annexin V in the placental tissues had a similar pattern to that of FLAG-tagged annexin V. There was no detectable annexin V stained in decidual tissue (Figure 2.4A). The control, stained with secondary antibody only, did not show any positive staining (Figure 2.4B).

Anti-annexin V antibodies block the anticoagulant properties of annexin V in vitro. When added alone, annexin V prolonged the aPTT from 34.5 ± 1.2 to 54.2 ± 3.5 seconds (n=3), this prolongation was blocked when anti-annexin V antibody was added

22 with annexin V (aPTT 37.7 ± 5.1 seconds, n=3). Antibody alone does not change the aPTT (35.0 ± 0.9 seconds, n=3) (Figure 2.5).

Infusion of anti-annexin V antibodies into pregnant BALB/c mice causes fetal loss. Anti-annexin V antibody can be detected in mouse circulation up to 5 days after infusion. Serum antibody levels decreased rapidly from 16 ng/µl to 10 ng/µl during the first 24 hours after infusion and then plateau at 8 ng/µl for the next 2 days. The second rapid drop in the level of serum anti-annexin V occurred 72 hours after infusion. Five days after infusion, anti-annexin V can not be detected by ELISA (Figure 2.6).

Fetal reabsorption was observed at day 11 or day 15 of pregnancy in all mice infused with affinity-purified anti-annexin V IgG. At day 11, most of the affected embryos were only partially reabsorbed and some of the remnant embryonic and placental tissues were recognizable (Figure 2.7D). At day 15, the embryos and placentas affected were almost completely reabsorbed and only bluish spots of the necrotic tissue mixed with blood remained (Figure 2.7B). No fetal loss was observed in the four saline infused and four normal rabbit IgG infused mice at day 11 or day 15 of pregnancy (Figure 2.7A & C).

Mice infused with normal saline carried 12 ± 3 embryos (n=4); mice infused with normal rabbit IgG also bore 12 ± 3 embryos (n=4). However, mice infused with anti-annexin V only have 6 ± 2 embryos left (n=8), which is significantly lower than either of the control groups (p<0.01).

Pathological changes in the affected placentas. At day 11, the atretic embryos in mice infused with anti-annexin V showed nearly intact placenta and embryo debris. Histo- logically, there were many small thrombi located at the junction of decidual and fetal tissues. The fetal tissue displayed large-scale necrosis and fibrosis with islands of normal

23 tissues. The decidua basalis demonstrated an inflammatory response, with infiltration of lymphocytes. There was no necrosis or thrombus formation in the blood vessels (Figure 2.8A & B). At day 15, there is no tissue left in absorbed embryos for pathological studies (Figure 2.7B). In one uterus, some of the embryos were smaller than others (4x3 mm and 9x6 mm)(Figure 2.8C). Placentas from the smaller embryos had no visible changes in fetal tissues. The decidual tissues, however, displayed focal necrosis and fibrosis (Figure 2.8D). No pathological changes were found in the placental tissues from saline and normal IgG- infused mice (Figure 2.8E & F).

COMMENTS

An autoantibody is defined as being pathogenic if it causes clinical manifestation when passively transferred to a normal animal (Bona, 1991). In the present study, we demonstrated that passive transfer of anti-annexin V antibodies into pregnant mice caused multiple fetal losses, possibly due to placental thrombosis and necrosis. These findings strongly argue that annexin V plays a critical role in sustaining placental viability. Results obtained from our defined murine model correlate well with clinical findings.

I speculate that, in the placenta, annexin V is released from syncytiotrophoblasts while these cells undergo rapid growth and turnover. The released annexin V then binds to the surfaces of the apoptotic cells and by so doing blocks the exposed anionic phospholipids, mainly phosphatidylserine. The presence of the annexin V binding sites was confirmed by the infusion of FLAG-tagged annexin V into pregnant mice (Figure. 2.3). Thus, a “shielding” effect of annexin V may prevent the activation of the blood coagulation system and, consequently, keep the slow moving blood from clotting on the placental surfaces. The shielding of anionic phospholipids also prevents the initiation of inflammatory responses.

24 h g V u n h h rd i ro h s x s a s n a e th a a d n w w w n n w t d tio ta s n rd lu s rA flo 1 2 3 e

35 Kd 31 Kd

Figure 2.1. Purification of recombinant epitope-tagged annexin V. Recombinant FLAG- annexin V was purified using phospholipid affinity chromatography. Fractions from flow- through, 3 wash with buffers of different salt concentrations and elution were subjected to SDS-PAGE and stained with Coomassie Blue. The molecular mass of wild-type recombinant annexin V (rAnnexin V) is 35 kDa, the eight amino acid FLAG epitope tag increases the size of the FLAG-annexin V to 36 kDa (elution).

25 B

F D

Figure 2.2. Mouse placental histology (4X). Mouse placenta contains maternal tissue decidua (D) and fetal tissue (F). Fetal trophoblasts form labyrinthine and endodermal sinuses that are bathed directly in maternal blood supplied by blood vessels (B) from decidua.

26 Figure 2.3. Identification of extracellular binding sites for annexin V in placenta. A, Immunoperoxidase staining with polyclonal rabbit anti-FLAG antibody shows that the infused annexin V binds to the surface of the sinuses containing maternal blood. The “wire loop” pattern of distribution corresponds to the trophoblast surfaces in mouse placenta (40X). B, Control. A placental tissue section from a control mouse stained with the same anti- FLAG antibodies does not show positive staining for FLAG-tagged annexin V (20X). D: Decidua; F: Fetal tissue.

27 D F

Figure 2.4. Immunolocalization of endogenous annexin V in placenta. A, immunoperoxidase staining with rabbit anti-annexin V antibodies demonstrates that annexin V is located mainly in the fetal tissue (F). It is abundant on the surface of sinuses containing maternal blood. The decidua (D) does not stain (40X).

28 60

)

s 50

T T 40

30 PBS Anx V A n x V + Anti- Anti-anx V anx V

Figure 2.5. Clotting assays (aPTT) with annexin V and anti-annexin V alone or in combination. Annexin V prolongs the aPTT from 34.5 ± 1.2 seconds (PBS control) to 54.2 ± 3.5 seconds (annexin V). Anti-annexin V antibody prevents the prolongation of aPTT by annexin V from 54.2 ± 3.5s (annexin V) to 37.7 ± 5.1s (annexin V + anti-annexin V). Anti-annexin V alone does not affect aPTT (35.0 ± 0.9s).

29 18

l) 12

(n 10

exin

n 8

ti-an

n 6

0 0 20 40 60 80 100 120 140 160

Hours After Infusion

Figure 2.6. Time course of anti-annexin V in the circulation after tail-vein infusion. Ten µg anti-annexin V was tail-vein infused to mouse circulation. Anti-annexin V level dropped rapidly from 16 ng/µl to 10 ng/µl within the first 24 hour after infusion, and then remained at ~8 ng/µl for the next 48 hours. Five days after the infusion, serum anti-annexin V level can not be detected by ELISA.

30 AB

Figure 2.7. Effects of circulating anti-annexin V antibody on pregnancy: gross morphology. A, a uterus from a 15-day pregnant mouse infused with nonimmune rabbit IgG. All of the embryos are intact and of normal size. B, a uterus from a 15-day pregnant mouse infused with rabbit anti-annexin V antibodies immediately before mating. There are about 5 embryos that were absorbed, leaving necrotic tissues at the point of implantation (arrows). The remaining embryos are normal in size and gross appearance. C, a uterus from a 11-day pregnant mouse infused with nonimmune rabbit IgG. The embryos are of normal size and there are no missing embryos in the uterus. D, a uterus from an 11-day pregnant mouse infused with rabbit anti-annexin V antibodies. There are 6 partially absorbed embryos with the recognizable placentas (arrows). The uterus demonstrates cyanosis.

31 A B

F D

D C

N D

E F

Figure 2.8. Effects of circulating anti-annexin V antibody on pregnancy: histopathology. A and B, Placenta from a partially absorbed embryo (arrow in A). The fetal tissue (F) is necrotic with scattered islands of normal tissue. A large thrombus (T) and many small thrombi are apparent, close to the decidual-fetal junction. The decidua (D) shows lympho- cyte infiltration, indicating chronic inflammation (20X). C and D, Placenta of an embryo smaller in size than others in the same uterus (arrow in C). This embryo measures 4x3 mm compared with 9x6 mm for the normal. There is no obvious necrosis or thrombosis in fetal tissue (F). Focal necrosis (N) and fibrosis are seen in the decidua (D) (20X). E and F, Placenta of a normal embryo (arrow in E). Normal fetal tissue (F) shows labyrinthine type of maternal-fetal interdigitation. The sinuses formed by fetal tissues are surrounded by maternal blood. The decidual tissue (D) contains maternal blood vessels that carry maternal blood to the sinuses (20X). Tissue sections were stained with H&E (20X).

32 The precise cause of pregnancy loss in patients with autoantibodies is not understood. Pathological changes in the placentas of these patients suggest a mechanism of arterial thrombosis in placental vessels and maternal blood fetal tissue surfaces (Salafia, 1996).

These results suggest that infused anti-annexin V antibodies neutralize annexin V in the placenta. Alternatively, the antigen-antibody complexes formed by annexin V and anti- annexin V antibodies may cause chronic inflammation in fetal tissues, which, in turn, dam- ages syncytiotrophoblasts. In addition, antiphospholipid antibodies are suspected to be directed against certain phospholipid binding proteins involved in regulating hemostasis (Feinstein, 1992). Anti-annexin V antibody may be a component of antiphospholipid antibodies and lupus anticoagulants. However, in the clotting assay performed in our study, anti-annexin V antibody alone did not affect aPTT as reported by Nakamura et al (Nakamura, 1995).

Our result is in agreement with Rand et al (1997a). Antiphospholipid antibodies have been shown to decrease the level of annexin V on cultured BeWo choriocarcinoma cells and cultured human placental villi. The mechanism may be that antiphospholipid antibodies displace annexin V from the cell surfaces where it binds and functions as an anticoagulant (Rand, 1997a, Vogt, 1997). Anti-annexin V antibodies may act in a similar manner as antiphospholipid antibodies to cause the loss of annexin V in placenta, and promote thrombosis at the maternal-fetal junction. This, in turn, compromises the nutrient exchanging functions and consequently causes fetal death and reabsorption. The large scale of necrosis and thrombus formation in the placentas of the partially reabsorbed embryos in our study is consistent with this hypothesis. The hypercoagulation status and high rate of pregnancy loss found in some of patients with autoimmune diseases may be due to the decrease of annexin V caused by anti-annexin V autoantibodies alone or together with

33 antiphospholipid antibodies. The same thrombotic scenario may occur on blood vessel endothelial cells other than placenta. The amount of anti-annexin V antibodies infused in the current study did not cause any visible thrombus formation in other organs like heart, brain and kidney (data not shown).

In conclusion, infusion of anti-annexin V antibody resulted in fetal loss in pregnant BALB/c mice, possibly due to placental thrombosis and tissue necrosis . The pathologic properties of anti-annexin V antibodies might explain the high thrombotic status and recurrent intrauterine fetal loss in patients with autoantibodies against proteins associated with coagulation. This association is particularly interesting in those patients who tested negative for antiphospholipid antibodies. The present study adds further insight into the critical role annexin V plays in maintaining placental viability. It appears that factors that interfere with annexin V binding to the surfaces of syncytiotrophoblasts contribute to recurrent fetal losses characteristic of the antiphospholipid syndrome.

34 CHAPTER III

DETECTION OF EXTRACELLULAR ANNEXIN V AND ATTEMPTS TO SECRET RECOMBINANT ANNEXIN V FROM CULTURED CELLS

INTRODUCTION

As previously discussed, annexin V may have a protecting role in the placenta. The functions of annexin V in the placenta require that it be extracellular. However, the annexins do not possess a traditional secretory signal sequence. For this reason, annexin V is generally thought to be an intracellular protein (Funokoshi, 1987).

To further understand if annexin V is released or secreted from mammalian cells, I investigated two cell types in cell culture. BeWo trophoblast cells and human umbilical vein endothelial cells (HUVEC), both rich in annexin V and widely used to study the anticoagulant functions of the protein. I also developed a transgene that secrets annexin V into the culture medium from COS-7 cells by attaching a signal peptide to the protein’s amino terminus.

MATERIALS AND METHODS

Cell Cultures. BeWo trophoblastic cells (American Type Culture Collection (ATCC)) were maintained in F-12K culture medium (Gibco) supplemented with 2 mM L-glutamine, 1.5 g/L sodium bicarbonate and 10% fetal bovine serum. Cells were grown as a monolayer in a 75mm flask at 37oC in the presence of 5% carbon dioxide. To stimulate BeWo cell

35 differentiation, 10 µM/L forskolin was added to the culture medium for 48 hours (Vogt, 1997).

HUVEC (ATCC)) were grown in Kaighn’s F-12K medium (ATCC) containing 2 mM L-glutamine, 1.5 g/L sodium bicarbonate and 10% fetal bovine serum. The medium was supplemented with 0.1 mg/ml heparin (Sigma) and 0.3 mg/ml endothelial cell growth supplements (ECGS) (Sigma). Cells were grown as a monolayer in a 75mm flask at 37oC with 5% carbon dioxide. To differentiate the cells, HUVEC were treated with 6 ng/ml phorbol myristate acetate (PMA) for 24 hours. To stimulate annexin V release, 1 µM dexamethasone was added to the cells for 16 hours after PMA treatment (Solito, 1991).

COS-7 cells purchased from ATCC were maintained in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum. Cells were grown as a monolayer in flasks at 37oC with 5% carbon dioxide.

Immunoprecipitation. BeWo cells were grown in a 75mm2-culture flask for 48 hours. Fifteen ml of medium was collected and used for immunoprecipitation. The cells were lysed in 1mL lysis buffer containing 1% triton X-100, 100 mM NaCl and 25 mM Tris (pH4.8). The cell lysate was kept on ice for 1 hour and centrifuged at 5000 rpm for 5 min to remove cellular debris. The medium and supernatant from the cell lysates were precleared

with 50 µl protein A-Sepharose at 4oC overnight. After incubating with rabbit anti-annexin V antibody for 2 hours at 4oC, 50 µl protein A Sepharose was added to the samples and they were then incubated for 2 hours at 4oC. The protein A-Sepharose was washed consecu- tively using the following buffers: 2 washes with dilution buffer (0.1% Triton X-100, 0.1% bovine hemoglobin in 0.01 M Tris pH 8.0/saline/0.025% azide solution); 1 wash with TSA solution (0.01 M Tris-Cl, pH 8.0, 0.14 M NaCl and 0.025% NaN3); and a final wash with 0.05 M Tris-Cl, pH6.8. The protein A-Sepharose was then resuspended in sample buffer

36 containing 10% SDS, 15% β-mercaptoethanol (v/v) and 10 mM EDTA. After incubating at 100oC for 5 min, the samples were centrifuged and the supernatants were subjected to SDS- PAGE.

Construction of FLAG-tagged annexin V gene. The vectors for cloning annexin V were purchased from Eastman Kodak. pFLAG-CMV-1 has a signal peptide (SP) from preprotrypsin (MSALLILALVGAAVA) attached at the N-terminus of FLAG (DYKDDDDK). Annexin V cDNA was cloned to the carboxy-terminus of FLAG sequence. The final construct is called pCMV-SP-FLAG-ANNEXIN V. pFLAG-CMV-2 has only the FLAG sequence. Rat annexin V gene was also cloned to the C-terminus of the FLAG sequence. The final construct is called pCMV-FLAG-ANNEXIN V. Both constructs were purified and sequenced before use.

Transient transfection of recombinant annexin V in COS-7 cells. Lipofectamine (Gibco) was used for transfection. COS-7 cells were trypsinized, resuspended and counted. Cells (2x105) were plated onto glass cover slips (for immunocytometry) in 35mm2 culture dishes and grown overnight in regular medium. To make DNA-lipid mixture, 1µL DNA (1 µg/µL) and 6µl LipofectAmine were mixed with 200 µl serum-free DMEM. After incubating at room temperature for 30 min, 800 µl serum-free DMEM was added to the DNA-lipid mixture and mixed well. Cells were rinsed once with PBS and 1 ml of the DNA-lipid mixture was added to each 35mm-culture dish, followed by incubation at 37oC for 5 hours. After the DNA-lipid mixture was aspirated, cells were rinsed once with PBS, replaced with regular culture medium, and incubated at 37oC for 48 hours.

Concentration of annexin V in culture media from BeWo cells, HUVEC and transfected COS-7 cells. The procedures followed were those used for the purification of annexin V (Kaetzel, 1989). Culture media of BeWo cells were collected 48 hours after

37 plating or after forskolin treatment from 75cm flasks. Culture media from HUVEC were collected 48 hours after plating or after PMA and dexamethasone treatment. Media from COS-7 cells were collected 48 hour after the cells were transiently transfected with both pCMV-SP-FLAG-ANNEXIN V and pCMV-FLAG-ANNEXIN V constructs. 3 mM of EGTA was added to the culture flasks briefly before collecting media. Media were then

supplemented with 5 mM CaCl2 and immediately applied to a column of phenyl-Sepharose that previously had been treated with brain phospholipids (Sigma). For positive control, recombinant annexin V was added to the fresh medium and this control sample was applied to the column and treated the same way as the samples. The column was extensively washed with buffers containing different salt concentrations of 0.5 M and 1.0 M in the presence of µ 1 M CaCl2. FLAG-annexin V was eluted from the column with EGTA. 200 l fractions were collected and 20 µl was subjected to SDS-PAGE.

Immunocytochemistry Permeabilized cells. Cells were grown on 11x11mm2 glass coverslips and transfected as needed. 10% PBS-buffered formalin was used to fix the cells for 20 minutes followed by permeabilization with ice cold acetone for 7 minutes. The coverslips were incubated with affinity purified rabbit anti-annexin V polyclonal antibodies or mouse anti- FLAG monoclonal antibodies (Eastman Kodak) and visualized by fluorescein-conjugated secondary antibodies. COS-7 cells transfected with pCMV-FLAG-ANNEXIN V and pCMV- SP-FLAG-ANNEXIN V were also double stained with mouse anti-FLAG antibodies and rabbit anti-BiP (an endoplasmic reticulum marker) antibodies (Provided by Dr. Paul Kim, Department of Medicine, University of Cincinnati).

Non-permeabilized cells. For staining of cell surface annexin V, cells were grown on 11x11 mm2 glass coverslips and fixed with 10% PBS-buffered formalin for 5 minutes.

38 The fixed cells were then incubated with affinity purified rabbit anti-annexin V followed by fluorescein-conjugated secondary antibodies.

Live cells: For annexin V binding, cells on glass coverslips were washed with annexin V binding buffer (Travergene) containing 100 mM HEPES Ph 7.4, 1.5 M NaCl, 50 mM

KCl, 10 mM MgCl2, and 18 mM CaCl2. The cells were then incubated with fluorescein- conjugated annexin V prepared in annexin V binding buffer for 20 minutes at room temperature, followed by three washes with the same binding buffer.

Immunohistology. Paraffin embedded rat placenta and mouse liver tissue sections were treated and immunostained with annexin V as described elsewhere (Wang, 1999). Tissue sections were deparaffinized in Xyline (Fisher) and rehydrated gradually using de- creasing concentration of ethanol solutions. Two-hour incubation with rabbit anti-annexin V antibody was followed by one-hour incubation with fluorescein-conjugated secondary antibodies. The sections were then viewed using fluorescence microscope.

Immunoblotting. Proteins were separated using 12% SDS-PAGE, transferred to nitrocellulose membrane, and then immunoblotted with affinity purified anti-annexin V or monoclonal anti-FLAG antibodies overnight at 4oC. After this time, the blots were exposed to HRP-conjugated anti-rabbit or anti-mouse secondary antibodies for 1 hour at 37oC, and annexin V was visualized by color reaction with HRP color development reagent (4-Chloro- 1-Naphthol, Bio-Rad Laboratories) and ECL.

39 RESULTS AND DISCUSSION

Localization of annexin V in blood vessel endothelial cells and placental trophoblasts. When rat placenta tissue sections were stained with anti-annexin V antibodies, the annexin epitope was concentrated on the surfaces of sinuses surrounding maternal blood (Figure. 3.1A). Rat placenta, like mouse placenta, has labyrinthine like maternal-fetal tissue interdigitation (Wang, 1999). The localization of annexin V is also similar as that in the mouse placenta (Wang, 1999). The endothelial cells of blood vessels from mouse liver also show intensive staining for annexin V, as do the endothelial cells of bile ducts (Figure 3.1B). The high concentration of annexin V in both placental trophoblasts and blood vessel endothelial cells makes it ideally positioned as an anticoagulant.

Localization of annexin V in permeabilized cells and detection of annexin V in culture medium. BeWo cells, a human choriocarcinoma cell line, have been used to study the distribution and function of annexin V by other investigators (Rand, 1997b, Vogt, 1997). When permeabilized cells were immunostained with rabbit anti-annexin V antibodies and visualized with fluorescein-conjugated goat anti-rabbit secondary antibodies, annexin V was found to be localized in nuclei and cytoplasm (Figure 3.2A). Other studies showed similar results (Vogt, 1997). The distribution of annexin V in these cells does not suggest the possibility of secretion by the classical secretory pathway because the staining failed to reveal the reticular pattern of ER and polarized paranuclear localization of Golgi complex and secretory vesicles. While immunoprecipitation with rabbit anti-annexin V showed the presence of annexin V in cell lysate of BeWo cells (Figure 3.2B, cell lysate), it did not show any annexin V in the culture medium (Figure 3.2B, medium IP). The result from immunoprecipitation was confirmed by an attempt to affinity purify annexin V from culture medium using phospholipid-conjugated phenyl Sepharose column (Figure 3.2B, medium col). Passing the medium through the affinity column not only concentrated annexin V present

40 (if any) in the medium but also purified the protein. in order for protein to show up on the Western blot using anti-annexin V antibodies, the protein has to be a calcium dependent phospholipid binding protein that also reacts with anti-annexin V antibodies. The proce- dure significantly reduces the possibility of nonspecific recognition. Forskolin stimulates the differentiation of BeWo cells, which is characterized by the secretion of human choriogonadotropic hormone (HCG) and the externalization of phosphotidylserine (PS) (Vogt, 1997). Soluble annexin V in the cultured medium was not determined.

These results showed that after treating the cells with forskolin for 48 hours, there was no detectable annexin V in the culture medium by affinity purification, western blot analysis and ECL (Figure 3.2B, medium+fors). Three mM of EGTA was added to culture flasks briefly before the media were collected and excessive CaCl2 (5mM) was supplemented before applying the samples to ensure the binding of annexin V to the column. My results also indicate that secretion of annexin V by differentiated BeWo cells is <0.02 ng/ ml, the lowest limit of detection by affinity column concentration and ECL signal amplification. Adding EGTA before collecting the medium eliminated the possibility of annexin V sticking to the exposed anionic phospholipids.

Similarly, annexin V was localized in the nuclei and cytoplasm of the umbilical vein endothelial cells (Figure 3.3A). My results are in agreement with others (Sun, 1992). Lo- calization of annexin V is not consistent with that of a secretory protein. The fetal bovine serum containing culture media were also collected and subjected to phospholipid affinity purification. While there was abundant annexin V in the cells (Figure 3.3B, cell lysate), there was no annexin V detectable in the culture media of these cells (Figure 3.3B, medium), confirming that annexin V was not being released. These findings are not in agreement with earlier reports that annexin V was detected in the culture medium of the same endothelial cells (Flaherty, 1990). Treating cells with PMA and dexamethasome consecu-

41 tively also failed to stimulate the release of annexin V into the medium of endothelial cells (Figure 3.3B, PMA+Dex), in contrast to the studies of Solito et al (1991). In their article, they reported that annexin V was released into the medium of U-937 cells after differentiation of these cells using PMA and dexamethasone. Taken together, These data indicate that annexin V is not released into the medium of cultured endothelial cells.

Evaluation of cell surface anionic phospholipids and localization of annexin V to the surface of BeWo cells and human umbilical vein endothelial cells. Cell surface exposure of anionic phospholipids was evaluated using fluorescein-labeled annexin V staining. Under normal conditions, there should be little or no anionic phospholipids exposed on the outer leaflet of cell membrane except in apoptotic cells. When non-forskolin treated live BeWo cells were probed with fluorescein-labeled annexin V (FITC-annexin V), a low incidence of annexin V staining was observed (Figure 3.4A). These cells may be apoptotic, because apoptotic process is characterized by externalizing anionic phospholipid PS (Fadok, 1992).

When BeWo cells differentiate and fuse to become syncytiotrophoblasts, there is externalization of PS (Vogt, 1997). FITC-annexin V staining of live cells confirmed externalization of anionic phospholipids after forskolin treatment. There was substantially more cells show annexin V binding (Figure 3.4B). My results are in agreement with those of Vogt et al (1997). The presence of PS can trigger PS-dependant coagulation and inflammation.

These results indicate that appreciable amounts of annexin V are not secreted or released into the culture media of BeWo trophoblasts and human umbilical vein endothelial cells. Given the association of annexin V with biological membrane in a calcium dependent and independent fashions, it is possible that the liberated protein bound to the anionic

42 surface. Even though the anionic phospholipids are normally on the cytoplasmic leaflet of the plasma membrane, they can externalize to the cell surface when the cells differentiate. As shown above, when BeWo cells were differentiated with forskolin, increased annexin V binding was observed (Figure 3.4B). I propose that cell surface annexin V covers the exposed phosphatidylserine.

To determine whether annexin V is on the surface of the cultured cells, the non- permeabilized cells were stained with anti-annexin V antibodies. When the cells are not differentiated, annexin V epitope was found on the surfaces of a few round cells (Figure 3.4C). It has been reported that when cells go through apoptosis, they expose phosphotidylserine on their surfaces, and annexin V has been immunolocalized on the surfaces of apoptotic cells (Boersma, 1996, van Engeland, 1996). Staining with anti-annexin V antibody showed annexin V on the surface of most forskolin-differentiated BeWo cells (Figure 3.4D), suggesting that differentiation of BeWo cells may be accompanied by externalization of PS and annexin V. The annexin V epitope was on the outer surface of the cells because the cells were not permeabilized and the cells did not show the staining of intracellular annexin V.

FITC-annexin V binding was observed on the surface of both non-treated and PMA- treated human umbilical vein endothelial cells (HUVEC) (Figure 3.5A). PMA treatment did not increase the binding of annexin V, which indicates that there is no increase of anionic phospholipid exposure on these cells. Studies by other investigators have demonstrated the same results using the same endothelial cells (van Heerde, 1994). Similar to BeWo cells, annexin V was localized on the surfaces of endothelial cells whether the cells were differentiated with PMA or not (Figure 3.5B).

43 Both in BeWo cells and in endothelial cells, annexin V was not evenly present on the whole cell surface but rather appeared as bright ‘dots’. It is possible that these structures represent the small cell surface blebs that have clustered exposed phosphatidylserine. As a result, annexin V clusters on the phospholipid surfaces, perhaps shielding the procoagulant surfaces. Andree et al (1992) found that the inhibition of prothrombinase (the complex of anionic phospholipid, Ca2+, factor V and factor Xa) by annexin V strongly depends on the cluster of the lipid bound annexin V (Andree, 1992). Relative to the total cellular annexin V in these cells, the amount of annexin V on the cell surface is minute. I have shown that annexin V is not secreted or released into the culture medium of either cell type. Therefore, the source of cell surface annexin V can not be the medium. It is possible that when there is cellular damage, a small quantity of annexin V escapes and binds to the exposed PS on the surfaces of adjacent cells. These observations suggest that when blood vessel injury occurs, annexin V could be released from damaged endothelial cells and then bind to exposed anionic phospholipids from both endothelial cells and activated platelets.

Secretion of annexin V from COS-7 cells to the culture medium. To investigate whether annexin V could be targeted to the traditional ER/Golgi secretory pathway of COS- 7 cells, two constructs were made with annexin V cDNA. In the case of the first, pCMV- FLAG-ANNEXIN V (Figure 3.6A), the FLAG epitope tag was attached to the amino-terminus of annexin V cDNA and the construct was transiently transfected to COS-7 cells. The fixed and permeabilized cells were stained with anti-FLAG antibodies, which recognize only FLAG-annexin V gene products. The results showed that without a secretory signal peptide, FLAG-annexin V was localized to the nucleus and cytoplasm (Figure 3.6B), very similar to the localization of endogenous annexin V in BeWo cells and in HUVEC (Figure 3.2A, 3.3A). The ER and Golgi distribution pattern was not seen with this construct. West- ern blot analysis revealed that FLAG-annexin V was present in the cell lysate but not in the

44 culture medium (Figure 3.6C). These results confirm that annexin V is not secreted by COS-7 cells if a signal sequence is not present in the constructs.

Similar experiments were done using a second construct, pCMV-SP-FLAG- ANNEXIN V, in which a signal peptide derived from preprotrypsin was incorporated onto the amino-terminus of FLAG-annexin V cDNA (Fig. 6D). The construct was also transiently transfected to COS-7 cells. The transfection efficiency (about 50%) was similar to that of the first construct. Immunostaining with anti-FLAG antibodies showed that the FLAG-annexin V was localized to the endoplasmic reticulum (ER), Golgi complex and secretory vesicles (Figure 3.6E), a pattern obviously different from the localization pattern of endogenous annexin V (Figure 3.1A). The ER and Golgi distribution of FLAG annexin V was confirmed by the colocalization of FLAG-annexin V and BiP, an ER resident protein (data not shown). Localization to the ER and Golgi indicates targeting to the secretory pathway. Immunoblotting with anti-annexin V antibodies proved that FLAG-annexin V was not only present in the cell lysate but also in the culture medium (Figure 3.6F). Our results demonstrate that a signal peptide is necessary for effective secretion of annexin V in mammalian cells.

In conclusion, annexin V is not secreted into the media of cultured BeWo trophoblast cells and human umbilical vein endothelial cells. Annexin V is, however, readily secreted from COS-7 cells if a signal peptide has been incorporated. The low amount of annexin V present in plasma and serum reported by others (Shirotake, 1986, Flaherty, 1990, Tait, 1988) may derive from the damaged blood vessel endothelial cells and platelets, which are abundant sources of annexin V. This possibility is supported by the report that plasma annexin level increase after ischemic events such as myocardial infarction (Relton, 1991, Kaneko, 1996, Romisch, 1992). Annexin V was also proposed to be one of the new markers for early detection of myocardial damage (Mair, 1997) and a marker for the damage of

45 stored platelets (Krailadsiri, 1997). It is possible that annexin V carries out its anticoagulation and other activities locally but not systemically. When the endothelial cells or trophoblasts are damaged or undergo apoptosis, intracellular annexin V is released. The released protein binds to the exposed PS on the surfaces of adjacent cells, preventing the activation of coagulation factors by negatively-charged phospholipids.

46 Figure 3.1. Localization of annexin V in rat placental trophoblasts and blood vessel endothelial cells. Paraffin-embedded rat placenta and mouse liver sections were deparaffinized and rehydrated followed by incubating with rabbit anti-annexin V antibodies for 2 hours and fluorescein-conjugated secondary antibodies for1 hour. In rat placenta (A), annexin V was localized on the surfaces of sinusoid structures containing maternal blood. In mouse liver blood vessels (B), annexin V was concentrated on the surface of the vessel. The endothelia of bile ducts were also intensively stained (not shown).

47 A

l o te IP c a s m m m B u u rs ly i i iu l l d d d fo e e e e + c m m m

35 Kd

Figure 3.2. Immunofluorescent staining of annexin V in permeabilized BeWo cells and immunoblot detection of annexin V in culture media. A, Immunostaining. BeWo cells were fixed with 10% PBS-buffered formalin and permeabilized with acetone. Endogenous annexin V was probed with rabbit anti-annexin V antibodies and visualized by fluorescein- conjugated goat anti-rabbit antibodies. Annexin V was localized to the nucleus and cytoplasm of BeWo cells. B, Western blots. Immunoprecipitation of annexin V in BeWo cell lysate. Annexin V was precipitated by rabbit anti-annexin V and protein A. Sheep anti- annexin V was used to probe annexin V in Western blot. Annexin V was precipitated from BeWo cell lysate (cell lysate). The immunoprecipitation was carried out the same as with the cell lysate. There was no annexin V precipitated from culture medium of BeWo cells (medium IP). Culture medium was concentrated by passing it through a phospholipid affinity column, and annexin V in the EGTA-eluted fractions was detected by Western blot. There was also no detectable annexin V in the medium (medium col). BeWo cells were treated with 10µM forskolin for 48 hours and the culture medium was collected and annexin V was detected as described above. Annexin V was not detectable in the medium after the cells were treated with forskolin (medium+fors). 48 Figure 3.3. Immunofluorescent localization of annexin V in permeabilized HUVEC and immunoblot analysis detection of annexin V in culture media. HUVEC permeabilized and incubated with anti-annexin V antibodies. The intracellular annexin V was localized in the nucleus and cytoplasm of the cells (A). Media were collected from non-treated cells and PMA plus dexamethasone treated cells as described in the text. The samples were applied to annexin V affinity columns and elutes from the column were analyzed using SDS-PAGE and western blots. While annexin V is an abundant protein in HUVEC (cell lysate), it is not detected in the media either from non-stimulated cells (medium) or from PMA+dexamethasone stimulated cells (PMA+Dex).

49 Figure 3.4. FITC-annexin V binding and immunofluorescent staining of annexin V on the surface of non-permeabilized BeWo cells. Non-treated or forskolin treated BeWo cells were grown on glass cover slips and washed with annexin V binding buffer followed by incubating with FITC-annexin V for 20 minutes at room temperature. FITC-annexin V binding was observed on the surface of both non-treated (A) and forskolin treated cells, with the forskolin treated cells more intensively stained (B). For localization of cell surface annexin V, non-treated and forskolin treated BeWo cells were fixed in 10% PBS-buffered formalin for 5 minutes and incubated with anti-annexin V antibodies followed by fluorescein-conjugated secondary antibodies. When the cells were not treated with forskolin, only scattered cells demonstrated surface annexin V (C). After treating with forskolin, significantly more cells showed the presence of cell surface annexin V epitope (D). Cells incubated with secondary antibody only did not show any staining on the cell surfaces (E). The cellular response to forskolin was shown (F), the cells fused to become multi-nucleated cells. The cells were permeabilized, and immunofluorescent staining showed the intracellular localization of annexin V. Phase contrast pictures were taken to show the cells for each immunofluorescein picture (a,b,c,d,e).

50 Figure 3.5. FITC-annexin V binding and localization of cell surface annexin V in human umbilical vein endothelial cells (HUVEC). Non-treated and PMA-differentiated cells were grown on glass cover slips and washed with annexin V binding buffer followed by incubating with FITC-annexin V for 20 minutes at room temperature. Only a few cells showed annexin V binding to the surface of either non-treated (not shown) or PMA treated cells (A). Incubating the cells with anti-annexin V antibodies assessed the localization of cell surface annexin V. The annexin V epitope was localized to the surface of non-treated (not shown) and PMA-treated (B) cells. The staining is shown as white ‘dots’ (arrows). Cells incubated with secondary antibody only did not show any staining (not shown). Phase contrast pictures were take for each immunofluorescent staining (a,b).

51 A B C te a m s iu ly l V d l DTKDDDDK V e e fA A m c

CMV Annexin V poly A 35 Kd

D E F te a m s iu ly d ll DTKDDDDK V V e e fA A m c

CMV SP Annexin V poly A 35 Kd

MSALLILALVGAVAAVA

Figure 3.6. Expression and secretion of annexin V from COS-7 cells. A, FLAG-annexin V without signal peptide. Rat annexin V cDNA was cloned into a mammalian cell expression vector that has the FLAG epitope sequence but not signal sequence. The 8 amino acid FLAG was added to the amino-terminus of annexin V. B, Immunostaining of COS-7 cells transfected with pCMV-FLAG-ANNEXIN V, the construct that does not have the signal sequence. The FLAG-tagged protein without a signal peptide was detected in the nucleus and cytoplasm, different from the localization of the protein with a signal peptide. C, West- ern Blots. Cos-7 cells were transiently transfected with pCMV-FLAG-ANNEXIN V. 48 hours after transfection, the cells were lysed and the medium was collected and concentrated using a phospholipid affinity column as described in the text. Annexin V in the eluted fractions and cell lysate were detected by Western blots probed with rabbit anti- annexin V antibodies. The 36Kd FLAG-tagged annexin V was detected in the cell lysate but not in the medium. D, FLAG-annexin V with a signal peptide. Rat annexin V cDNA was cloned into the same mammalian cell expression vector that has a preprotrypsin signal peptide in addition to a FLAG epitope tag in the N-terminus. E, Cells transfected with pCMV-SP-FLAG-ANNEXIN V were stained with mouse anti-FLAG monoclonal antibodies. FLAG-tagged annexin V was shown as paranuclear reticulum pattern and polarized dense structure, which indicates the endoplasmic reticulum and Golgi complex. F, Western blot. COS-7 cells were transiently transfected with pCMV-SP-FLAG-ANNEXIN V. After transfection, the cells and medium were treated as described in C. The 36Kd FLAG-tagged annexin V (fAV) was in the cell lysate as well as in the medium. There was no detectable endogenous annexin V in COS-7 cells using my methods.

52 CHAPTER IV

TRANSGENIC MANIPULATION TO TARGET THE SECRETION OF ANNEXIN V INTO THE CIRCULATION OF MICE

INTRODUCTION

In the previous two chapters, I have argued that annexin V may have extracellular roles in the placenta, and I have shown that the protein is not secreted into the culture media of BeWo trophoblast cells and human umbilical vein endothelial cells. However, the protein is present on the cell surface. In addition, I successfully targeted annexin V to the secretory pathway of mammalian cells by incorporation of a signal peptide. Secreting annexin V into the extracellular spaces provides the advantage of studying the protein as a systemic anticoagulant since the half-life of annexin V after acute infusion is only about 6 minutes. In this study, I target annexin V to the liver of transgenic mice. With a signal peptide attached at the amino terminus of the gene, the protein is expected to be secreted into the blood circulation. Two promoters were used to drive the expression of annexin V: the promoter of phosphoenolpyruvate carboxykinase (PEPCK) and the promoter of albumin.

Phosphoenolpyruvate carboxylase is a rate-limiting enzyme that catalyzes the con- version of oxaloacetate to phophoenolpyruvate (PEP) during gluconeogenesis. This enzyme is involved in a variety of important tissue-specific metabolic processes in mammals (Flores, 1971, Hanson, 1972, Reshef, 1970, Tilghman, 1976), including gluconeogenesis in hepatocytes and proximal tubular epithelia of the kidney (Hanson, 1972, Tilghman, 1976), as well as glyceroneogenesis in white adipocytes (Reshef, 1970). The gene for the enzyme is regulated not by allosteric or phosphorylation/dephosphorylation mechanisms (Tilghman,

53 1976), but by a variety of dietary and hormonal signals, cell-cell interactions and development. These regulatory mechanisms result in alternation of protein synthesis (O’Brien, 1990, Beale, 1991 McGrane, 1990, Short 1992, Cassuto, 1999, Hanson, 1997). Major factors that increase PEPCK gene expression include glucocorticoids, cAMP, and thyroid hormones. Insulin inhibits PEPCK expression (Short, 1992, Park, 1999, Wang, 1999).

The PEPCK promoter has been used to express various genes in transgenic mice (Valera, 1994, Lim, 1990, Hatzoglou, 1990, Shimano, 1996, Patel, 1994). Feeding animals a high protein/low carbohydrate diet induces promoter activity in transgenic mice. The expression of the reporter gene increases by more than 13-fold when mice are fed with a protein diet than when mice are fed with carbohydrate-rich diet (Short, 1992). PEPCK is not expressed until after birth, concomitant with the capacity for gluconeogenesis (Short, 1992). Therefore, the PEPCK gene provides a model for the metabolic control of gene expression.

Albumin is an abundant serum protein synthesized and secreted by hepatocytes. Albumin gene expression is developmentally regulated. The promoter for albumin is turned on by gestation day 9. Thereafter, the mRNA level of albumin increases gradually until it reaches adult levels 10 days after birth (Tilghman, 1982). The mRNA level of albumin is among the highest in the liver and codes for the most abundant serum protein in adults (Tilghman, 1982). The albumin gene is repressed in non-hepatic tissues of the adult, so the albumin promoter is a strict liver-specific promoter (Pinkert, 1987). Therefore, the albumin promoter has been used to target expression of many genes in the liver of transgenic mice (Thorgeirsson, 1999, Thorgeirsson, 1997, Postic, 1999). The enhancer located 10 Kb upstream of the albumin gene along with its promoter direct efficient, liver-specific expression in transgenic mice (Pinkert, 1987).

54 MATERIALS AND METHODS

Construction of transgenes that secrete annexin V from hepatocytes Transgene with PEPCK promoter. Rat annexin V cDNA was tagged with an epitope tag FLAG (DYKDDDDK) (Eastman Kodak) at the N-terminus. An antibody to FLAG (monoclonal M2, Eastman Kodak, or polyclonal rabbit anti-FLAG, Zymed) distin- guishes the transgene product from endogenous annexin V. A signal peptide derived from preprotrypsin (MSALLILALVGAAVA) was cloned to the amino-terminus of FLAG-annexin V. This transgene is under control of PEPCK promoter (-464~+69, provided by Drs. Beale EG and Short MK, 1992).

Transgene with albumin promoter. The above signal peptide-FLAG-annexin V gene was cloned downstream of the albumin enhancer/promoter. The enhancer/promoter gene was provided by Dr. Thorgeirsson SS from NIH (Thorgeirsson, 1997).

Introduction of the transgenes into mice. Injection of transgenes was done in the transgenic facilities of the University of Cincinnati and the Children’s Hospital of Cincin- nati. The transgene was released from the plasmid by restriction enzyme digestion and was purified using a gel purification kit (Qiagen). The purified transgene was injected into the male pro-nucleus of a one-day embryo. The embryo was then transferred to a hormone- treated foster mother.

PCR analysis of mice carrying the transgenes. Design of primers used to amplify the transgenes. A pair of primers (TGM1 and TGM2) was designed to amplify the whole annexin V gene. The forward primer (TGM1, ACTACAAAGACGATGACGAC) is from the sequence of FLAG, which is unique to the transgene. The reverse primer (TGM2, TCCTCTAGAGTCGACTGGTA) is from the mul-

55 tiple cloning sites of the plasmid, which is also unique to the transgene. A pair of primers (TSH-β1, TCCTCAAAGATGCTCATTAG, TSH-β2, GTAACTCACTCATGCAAAGT) for amplification of TSH-β gene was also designed for use as controls.

Purification of genomic DNA from mouse tissue. Half an inch of tail from the distal tip of mouse tail was clipped at the age of 2 weeks, and the tissue was digested with protein degrader (Stratagene). Genomic DNA was purified from mouse tissue with the easy DNA kit (Stratagene).

PCR amplification of transgene from mouse genomic DNA. To identify transgenic mice, tag DNA polymerase (Gibco) was added to the mixture of genomic DNA, primers, dNTPs, and buffer for the enzyme. The reaction included a 2 minutes pre-reaction dwell at 94oC, and a 5 minutes dwell at 72oC at the end of the reaction and 35 cycles of amplification. Each cycle includes 30 seconds denaturing at 94oC, 30 seconds primer annealing at 48oC and 1 minute elongation at 72oC. The PCR mixture was subjected to 10% agrose gel in TAE buffer (0.02 M Tris, 5.7% glacial acetic acid and 0.1 mM EDTA pH 8.0) with 0.5 µg/ml ethidium bromide, and was visualized under UV light.

Southern Blot Analysis of mice carrying the transgenes. Genomic DNA purified as described above was digested overnight at 37oC with Kpn I. The enzyme cuts the transgene once. The digest was applied to 10% agrose gel and electrophoresed. The DNA in agrose gel was put under UV light for 10 minutes followed by denaturing for 30 minutes (denaturing buffer, 0.2 N NaOH, 0.6 M NaCl). After neutralizing (0.5 M Tris, 1.5 M NaCl) for 30 minutes, the DNA was transferred and crosslinked to a nylon membrane. The membrane was pre-hybridized with hybridization buffer (6x SSPE, 5x Denhardt’s, 0.5% SDS, 50% formamide) containing 8 mg/ml Salmon sperm DNA for 4 hours followed by over-

56 night hybridization with 32P-labeled transgene in the same buffer at 42oC. The membrane was then washed and analyzed using the Phosphorimager (Molecular Dynamics).

RT-PCR analysis of messenger RNA in transgenic mice. Total RNA was purified from fresh liver tissues of transgenic and control mice using RNAzolTM kit (Tel-Test). The purified RNA was confirmed not to contain any genomic DNA by using it as template for PCR with primers that amplify the transgene. The RNA was then transcribed to cDNA using reverse transcriptase (Gibco). The transcription product of RNA was used as tem- plates for PCR with primers (TGM1, TGM2) that amplify the transgene.

Western blot analysis of liver homogenate and plasma for FLAG-annexin V. Fresh liver tissue from transgenic and control mice was homogenized in hot 3x sample buffer (10% SDS, 15% β-mercaptoethanol and 10 mM EDTA) (Kaetzel, 1989). Plasma was collected from citrated mouse blood. Both the supernatant from liver homogenate (30 minute spin at 12,000xg) and plasma were analyzed by Western blot with anti-FLAG antibodies and anti-Annexin V antibodies as described in Chapters II, III and IV.

Immunofluorescence localization of FLAG-annexin V in the liver. The proce- dure for immunolocalization of protein in tissue sections was described in Chapters II and

III.

Induction of the activity of PEPCK promoter by high protein diet. Mice were fed with high protein diet (69% protein, 4% carbohydrate, Harlan) from 1 to 7 days before attempts to detect transgene expression.

57 RESULTS AND DISCUSSION

Targeting secretion of FLAG-annexin V in hepatocytes with PEPCK promoter. The transgene is shown in Figure 4.1. Signal peptide targets annexin V to the traditional secretory pathway of cells. The antibody to FLAG detects the transgene product but not endogenous annexin V. The PEPCK promoter ensures the expression of FLAG-annexin V mainly in the hepatocytes. The promoter is also inducible with diet, hormones and cAMP (Short, 1992, Park, 1999, Wang, 1999). Transgenic mice were identified by PCR analysis of mouse tail genomic DNA with primers that amplify only the transgene (Figure 4.1). Five out of eight mice were positive for transgene (~1 Kb band). The amplification of a 386 bp fragment of TSH-β gene indicates that genomic DNA is intact (Figure 4.2). Two out of five transgenic founder mice passed the transgene to the first generation (F1). The two founders were bred into heterozygotes and homozygotes. There was no FLAG-annexin V detected in either the liver or the plasma of transgenic mice before or after induction with high protein diet by all the methods of detection describe above.

Perhaps there is no detectable FLAG-annexin V in these transgenic mice because although there is expression, the promoter is not very strong. Since the promoter that I used is not active until after birth (Short, 1992), it is not likely that early expression of FLAG- annexin V is lethal to the animal. There was also no evidence of increased prenatal or postnatal death. The high protein diet induction increases the expression of PEPCK protein (data not shown).

Targeting secretion of FLAG-annexin V in hepatocytes with albumin enhancer/ promoter. The albumin promoter is much stronger than PEPCK promoter. As mentioned earlier, the albumin gene encodes the most abundant serum protein in adults (Tilghman, 1982). With the hope of a higher expression level of the transgene, FLAG-annexin V was

58 cloned to a position downstream of albumin enhancer/promoter, which ensures liver-specific expression of the transgene product (Figure 4.3). The genomic DNA of mice was first analyzed with PCR (Figure 4.4) and then Southern blot (Figure 4.5) to identify the transgenic mice. Southern blot analysis shows that the number of copies of the transgene in the positive mice was between 1 and 10 (Figure 4.5). The transgenic mice were analyzed using all the methods described above to detect the expression of the transgene. None of the transgenic mice had detectable FLAG-annexin V in the liver or the plasma.

To evaluate if there is transgene expression at the embryonic stage, mouse embryos were harvested 11 or 16 days after gene transfer. The embryo tissues were first tested for the incorporation of the transgene by Southern blot, and the positive ones were used to detect FLAG-annexin V expression in the liver. There was no FLAG-annexin V detected in the livers of transgene positive embryos.

The transcription of the transgene was evaluated with RT-PCR (Figure 4.6). Two out of four transgenic mice were positive, which means the transgene was transcribed in these mice. The reason that there is no detectable translation product is not known.

It is possible that targeting annexin V to the secretory pathway of the cells interferes with cell development or function, and the animals that express FLAG-annexin V do not survive. It is also possible that the transgene is silent. The messenger RNA detected in some of the mice may not be stable, or perhaps there is not enough transcription and/or translation in the animals.

None of the transgenic mice showed any obvious abnormality in liver structure as assessed in the liver tissue sections (H&E) or overall development.

59 Signal FLAG Peptide Stop Spe I Afl III PEPCK poly A

Figure 4.1. Transgene that is under control of PEPCK promoter. PEPCK: promoter from phosphoenolpyruvate carboxykinase. SP: signal sequence derived from preprotrypsine. Flag: the epitope tag. Annexin V: rat annexin V cDNA. The poly A and an intervening sequence were added to ensure mRNA stability and proper splicing. The annealing sites of the primers used to amplify the transgene in mice tissue are shown (arrows).

60 Figure 4.2. PCR identification of transgenic mice. The PCR product is about 1 Kb. The bands from the mice are the same size as the band from the amplification of plasmid containing the transgene. TSH-β indicates the amplification of a 386 bp fragments of TSH-β gene in every animal.

61 Signal FLAG Peptide

Stop Spe I Afl III 2335A-1 poly A

Figure 4.3. Transgene that is under control of the albumin enhancer/promoter. 2335-1: the enhancer/promoter sequence from the albumin gene. SP: signal sequence from preprotrypsin. FLAG: the epitope tag. The rest of the transgene is the same as the one described in Figure 4.1. The annealing sites of the primers amplifying the transgene are also shown (arrows).

62 r e d ic ic ad en en enicenic L g A nsg nsg N a ransga D Tr TransT Tr

1kb

Figure 4.4. PCR analysis of transgenic mice. The same primers were used as for Figure 4.2. Only mice that were incorporated with the transgene showed the ~1 Kb bands.

63 r c ic ic de s s i n d ie e n en p i y g La o p p ge s sge A o o n C C ns a an N 0 C ra r r D 50 1 1 T T T

8kb

Figure 4.5. Southern blot identification of transgenic mice. The transgene is ~8 Kb. The purified transgene was used as the probe and standards. The copy numbers of transgene incorporated into the transgenic mice were between 1 to 10.

64 A r N e e d p l R ad y id l 1 2 4 t tro A e e e 3 e ld sm n N in in in in i la o D L L L L W P C

1000 bp 500 bp

Figure 4.6. RT-PCR analysis of transgenic mice. +: Mice positive for mRNA. -: Mice negative for mRNA. Wild type: Non-transgenic mouse. Plasmid: the plasmid that contains the transgene. It is used as control for the PCR. Genomic DNA: Genomic DNA from a transgenic mouse. It is used as a control for PCR with DNA purified from animal tissue. Control RNA: Purified RNA from the kit. It is used as a control for the whole reverse transcriptase and PCR process.

65 CHAPTER V

TARGETING ANNEXIN V TO THE SECRETORY PATHWAY OF THYROID EPITHELIAL CELLS DOES NOT SIGNIFICANTLY ALTER CELLULAR FUNCTIONS

INTRODUCTION

Most of the proposed functions of annexin V are based on the protein’s calcium- dependent phospholipid binding properties. The best documented are the anticoagulant activities both in vivo and in vitro (Chollet, 1992, Thiagarajan, 1997, Funakosh, 1987, Andree, 1992, Sun, 1993b). The protein was termed placental anticoagulant protein I (PAP-I, Funakoshi, 1987) and vascular anticoagulant protein α (VACα, Reutelingsperger, 1985). Annexin V is a potential therapeutic agent for hypercoagulable conditions, as described in the previous chapters. Annexin V was also proposed to be an anti-inflammatory protein

due to its ability to inhibit phospholipase A2 (PLA2) by depleting anionic phospholipids, substrates of the enzyme. The name lipocortin V was given to annexin V when it was

independently isolated and was found to inhibit phospholipase A2 activity in vitro (Pepinsky,

1998). Many studies have shown that annexin V inhibits both secreted and cytosolic PLA2 (Buckland, 1998a, Mira, 1997, Buckland, 1998b). It was also shown that binding of annexin V to a model membrane containing phosphotidylserine and phosphotidylcholine completely

inhibits the degradation of phospholipids by PLA2 (Speijer, 1997).. The peptide fragment

204-212 of annexin V was found to inhibit the production of prostaglandin E2 and febrile response induced by cytokine in the brain (Palmi, 1995). The studies on these activities of annexin V and its related peptide may lead to the development of novel anti-pyretic and anti-inflammatory agents.

66 I have found (Chapter 3), however, that annexin V is not secreted from cultured placental trophoblasts and only after a signal peptide was attached was annexin V secreted from COS-7 cells. The extracellular function of annexin V requires its extracellular exist- ence. One approach to produce extracellular annexin V is to target it to the secretory pathway.

The signal sequence of a secreted protein targets the nascent protein through the ER membrane into the lumen. The endoplasmic reticulum (ER), however, is the location for post-translational processing of secretory proteins, cell surface proteins and integral membrane proteins. It is also the major calcium storage and release organelle in cells. The calcium concentration in the ER is also important for the function of the enzymes involved in protein processing and folding (Lodish, 1992). The specialized calcium ATPase in the ER membrane (sarcoplasmic/endoplasmic reticulum calcium ATPase, SERCA) pumps calcium ions into the ER where they are sequestered and buffered by high concentration, low affinity, high capacity calcium binding proteins such as calreticulin, BiP, calsequestrin and protein disulfide isomerase (PDI) (Meldolesi, 1998, Smith, 1989, Michalak, 1992, Lievremont, 1997, Campell, 1983, Lucero, 1998). Overexpression of calreticulin and BiP were found to increase the sequestration of calcium in the ER (Lucero, 1998, Lievremont, 1997). The calcium sequestered in the ER is thapsigargin-sensitive and can be readily released by inositol-1,4,5-triphosphate (IP3) (Lievremont, 1997, Lucero, 1998). Overexpression of these calcium-binding proteins does not produce significant effects on ER functions. However, overexpression of BiP was found to selectively inhibit secretion of factor VIII from Chinese hamster ovary (CHO) cells (Dorner, 1992).

Considering that one annexin V molecule can bind as many as 12-15 calcium ions (Seaton, 1996) and that the affinity for calcium increases dramatically in the presence of anionic phospholipids (Schlaepfer, 1987), targeting annexin V to the ER might have effects

67 on calcium homeostasis and protein processing. In addition, the high affinity of annexin V for anionic phospholipids might also affect lipid symmetry across the ER membrane, and as a consequence, affect ER functions. Using a tetracycline-inducible expression system, I targeted annexin V to the secretory pathway of a thyroid epithelial cell line and evaluated whether the presence of annexin V within the ER affects calcium homeostasis and protein processing.

MATERIALS AND METHODS

Cell culture. PC-rTTA7 cells are rat thyroid epithelial cells that are stably transfected with reverse tetracycline transactivator (rTTA) and pUHG72-neo gene and, therefore express rTTA and are G418 resistant. The cell line was created and provided by Dr. James Fagan’s laboratory (Division of Endocrinology and Metabolism, Department of Internal Medicine, University of Cincinnati). The cells were maintained in F-12 Coon’s modification nutrition mix (with 285 mg/ml L-glutamine and 0.863 mg/L zinc sulfate, Sigma), supplemented with 5% fetal bovine serum (FBS), four hormones (thyroid stimulating hormone (TSH) 10mIU/mL, insulin 10 µg/ml, transferrin 5 µg/ml, hydrocortisone 10nM), antibiotic/ antimycotic reagent (GIBCO) and 0.15 mg/ml G418 (GIBCO).

Construction of FLAG-tagged annexin V (FAV) for targeting the protein to the secretory pathway. Full-length rat annexin V cDNA was amplified by polymerase chain reaction (PCR) and subcloned into pCMV-FLAG-1 mammalian expression vector (Eastman Kodak, which contains an epitope tag sequence (FLAG), DYKDDDDK, and a signal peptide derived from preprotrypsin, MSALLILALVGAAVA, upstream from the multiple cloning sites. The construct was sequenced to confirm that annexin V cDNA was in the appro- priate reading frame and that there were no mismatches when annexin V cDNA was amplified by PCR. The signal peptide-FLAG-annexin V was subcloned into another mammalian

68 expression vector, pUHG 10-3, which has a coding sequence for tetracycline operator upstream from a CMV minimal promoter. The final construct in pUHG 10-3 (Sec-FAV) was used to transform PC-rTTA7 cells which constitutively express reverse tetracycline transactivator (rTTA).

Establishment of stable transfection cell lines overexpressing and secreting FLAG-annexin V. PC-rTTA7 cells expressing the reverse tetracycline transactivator were transfected with Sec-FAV plasmid DNA using LipofectAmine (GIBCO). Cells were trypsinized, resuspended and counted. Cells (2x105) were plated in 35mm culture dishes and grown overnight in regular medium. To prepare the DNA-lipid mixture, 1 µL Sec-FAV DNA (1 µg/µL), 1 µL pTK-Hyg (plasmid DNA with hygromycin resistance gene, provided by Dr. James Fagin’s laboratory, 0.1 µg/µL) and 6 µL LipofectAmine reagent were mixed by vortexing with 200 µL serum free F-12 Coon’s medium. The mixture was incubated at room temperature for 30 min, and 800 µL serum free medium was added to the DNA-lipid mixture and mixed well by tapping the tubes. Cells were gently rinsed once with PBS. One ml of the DNA-lipid transfection mixture was added to each 35mm culture dish, followed

o by incubation at 37 C for 5 hours with 5% CO2. The DNA-lipid transfection mixture was then aspirated and the cells were rinsed once with PBS. Modified Coon’s culture medium supplemented with FBS, G418 and the four hormones described above was added and the

o cells placed in the 37 C incubator with 5% CO2. After 48 hours, the cells were trypsinized and split into five 100mm culture dishes with selection medium containing 0.15 mg/ml hygromycin (Calbiochem) in addition to G418. The selection medium was changed every 4 days until individual colonies were large enough (approximately 1mm, 20 days) for subcloning. The colonies were grown to confluence on 6 well plates. One half of the cells of each colony was stored in liquid nitrogen and the other half were used for characterization.

69 Characterization of the expression and secretion of FLAG-annexin V in PC- rTTA7 cells by immunocytochemistry and immunoblot analysis. The stably transfected cells were selected and characterized by immunostaining and Western blot. In all the experiments with doxycycline induction, cells were induced with 1 µg/ml doxycycline for 5 days unless otherwise indicated. Stably transfected cells were grown on 11mm x 11mm glass coverslips and induced with doxycycline (GIBCO). Following fixation with 10% PBS-buffered formalin for 20 minutes and permeabilization using cold (-20oC) acetone for 7 minutes, the coverslips were incubated with mouse anti-FLAG monoclonal antibody for 2 hours and fluorescein-tagged goat anti-mouse secondary antibodies for 1 hour. Expression of the FLAG epitope was visualized using fluorescence microscopy.

For immunoblot analysis, cells were grown on 6 well plates and induced with doxycycline for 5 days with a change of inducing medium every two days. Culture media from induced cells and non-induced cells were collected from all 6 wells and immediately applied to a column of phenyl-Sepharose previously treated with brain phospholipids (Sigma). The column was extensively washed with buffers containing different salt concentrations of

0.5 M and 1.0 M in the presence of 1 mM CaCl2. FLAG-annexin V was eluted from the column with EGTA. 150 µL fractions were collected and subjected to 12% SDS-PAGE. In addition, the cells were lysed in 1mL buffer containing 1% Triton X-100, 100 mM NaCl and 25 mM Tris (pH4.8). The samples were kept on ice for 1 hour to completely lyse the cells. The mixture was centrifuged at 5000 rpm (12,00xg) for 5 minutes to remove cellular debris. The supernatant was applied to 12% SDS-PAGE. After electrophoresis, the proteins were transferred to a nitrocellulose membrane, blotted with either mouse anti-FLAG antibodies or rabbit anti-annexin V antibodies, and detected by HRP-conjugated secondary antibodies.

70 Cell growth studies. MTT assay: MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide) assay was used for assessing cell growth. These procedures have been described elsewhere (King, 1999). PC-rTTA7 cells were plated onto 96-well plates at 1000 cells/well in media with or without doxycycline. Cell viability was measured at 24, 48, and 72 hours by adding 25 µl of MTT (5 mg/ml in PBS) at each time point. After incubating at 37oC for 2 hours, 100 µl of extraction buffer (50% dimethylformamide, 10% sodium dodecyl sulfate, pH 4.7) was added and incubated 24 hours at 37oC. The absor- bance at 570nm was recorded using a Microelisa Auto Reader (Dynatech). 96-wells of cells were employed to the study at each time point.

45Ca2+ equilibrium uptake and release by thapsigargin. The procedures were modified from those reported previously (Missiaen, 1992, Lin, 1999). Non-induced and doxycycline induced cells were grown in 6 well plates and loaded with 2 µCi/ml of 45Ca2+ 48 hours before the experiments. The cells (106) were washed 3 times with Krebs-Ringer-

HEPES (KRH) buffer (125 mM NaCl, 5mM KCl, 1.2 mM KH2PO4, 1.2 mM MgSO4, 2 mM

CaCl2, 6mM glucose, and 25 mM HEPES, pH 7.4) and 5 times with PBS. Cells were then lysed in lysis buffer described above. For 45Ca2+ releasing experiments, washed cells were incubated for 20 minutes at room temperature with 20 µM thapsigargin in KRH supplemented with 3 mM EGTA. The supernatants were collected and cells were lysed as above. The radioactivity associated with the samples was counted for 1 minute using a Gamma counter (Beckman).

45 2+ Mobilization of stored Ca by IP3 in saponin-permeabilized cells. Non-induced and doxycycline-induced cells (106) were grown for 5 days on 6 well plates. Cells were permeabilized with saponin (50 µg/ml, 1 ml/well, Sigma) for 5 minutes at room tem-

perature in loading buffer (140 mM KCl, 20 mM NaCl, 2 mM MgCl2, 2 mM ATP, 30 mM imidazole-HCl, pH6.8) and washed 3 times with PBS. 10 µCi/ml 45Ca2+ were loaded to the

71 cells in loading buffer for 45 minutes at room temperature. After rapidly washing 5 times

with efflux buffer (120 mM KCl, 2 mM MgCl2, 1 mM ATP, 1 mM EGTA, 5 mM NaN3, 2 µ µ M thapsigargin, and 30 mM imidazole-HCl, pH6.8), cells were treated with 20 M IP3 (D- myo-inositol 1,4,5-triphosphate sodium salt, Calbiochem) in efflux buffer, 1ml/well. Solu- tions were collected every 2 minutes for 20 minutes and replaced with efflux buffer contain-

µ µ 45 2+ ing 20 M IP3. Another 6-well plate of cells was labeled with 10 Ci/ml Ca for 15 to 60 minutes and the cells were collected as described above. Radioactivity associated with the samples was counted for one minute using a Gamma counter (Beckman).

Assessment of thyroglobulin synthesis, processing and secretion Pulse chase experiments The pulse-chase experiment was adopted from that pub- lished by Kim PS (1991). Non-induced and doxycycline-induced cells were incubated in DMEM without methionine and cysteine (Sigma) for 20 minutes, and then labeled with 1 mCi 35S-methionine (NEN) per well for 60 minutes. Cells were then incubated for 1-5 hours with “chase” medium containing 135 mg/ml more unlabeled methionine and cysteine. At specified times, media were collected and a mixture of protease inhibitors (110 IU/ml aprotinen, 2 mM leupeptin, 2 mM pepstatin, 10 mM iodoacetamide, 190 mM EDTA and 200 mM PMSF) was added. The cells were treated with 50mM iodoacetamine for 5 minutes followed by lysis in buffer containing 1% Triton X-100, 100 mM NaCl and 25 mM Tris (pH4.8) and protease inhibitor mixture. The cell lysates were centrifuged at 4000 rpm (12,000xg) for 10 minutes and the media at 5000 rpm (12,000xg) for 5 minutes. Thyroglo- bulin synthesis was evaluated at the labeling time points of 15, 30, 45, 60, 90 minutes.

Gel electrophoresis and quantification The samples were applied to 4% or 4-12% gradient SDS-PAGE gels. After electrophoresis, gels were dried and then exposed in phosphorimager cassettes overnight. Labeled thyroglobulin was quantitated using the phosphoimager (Molecular Dynamics) and analyzed in Microsoft Excel. The percentage

72 of thyroglobulin secretion at each time point was calculated by dividing the amount of labeled thyroglobulin in the medium by the sum of labeled thyroglobulin in the medium and that in the cell lysate (medium radioactivity/total radioactivity). The percentage of labeled thyroglobulin retained in the cells was calculated from the thyroglobulin in the cell lysate at each time point divided by that in the cell lysate at 0 time point (cell radioactivity at each time point/cell radioactivity at 0 time).

Western blot analysis of thyroglobulin and chaperonins The cells were induced as described above. Non-induced and induced cells were trypsinized, resuspended in PBS and counted. 106 cells were pelleted by centrifuging at 5000 rpm (12,000xg) for 5 minutes and were lysed as described above. The samples were electrophoresed in 4% (thyroglobulin) or 10% (chaperonins) SDS-PAGE and transferred to nitrocellulose membrane. The membranes were probed with antibodies against corresponding proteins thyroglobulin, calreticulin, BiP, ERp72, PDI (protein disulfide isomerase), and Grp94 (provided by Dr. Paul Kim, Division of Endocrinology and Metabolism, Department of Internal Medicine, University of Cincinnati) and visualized by peroxidase-conjugated secondary antibodies.

RESULTS

FLAG-tagged annexin V targeted to the secretory pathway in PC-rTTA7 thyroid epithelial cells. In order to target FLAG-annexin V to the secretory pathway, a hydrophobic signal sequence from preprotrypsin was attached to its N-terminus. The epitope tag (FLAG) was used to distinguish the transgenic annexin V from endogenous annnexin V (Figure 5.1), an abundant intracellular protein in these cells. Immunostaining with mouse anti-FLAG monoclonal antibody showed that, in doxycycline-induced cells, FLAG-annexin V is localized to the endoplasmic reticulum and Golgi complex. The secretory vesicles were also visible (Figure 5.3A). ER localization was confirmed by the co-localization of

73 FLAG-annexin V with an ER-resident protein, BiP (data not shown). The distribution pattern of FLAG-annexin V was different from that of the endogenous protein, shown by the staining with rabbit anti-annexin V antibody. The endogenous annexin V was localized to the nucleus and cytoplasm (Figure 5.3B). There was no staining of FLAG-annexin V in cells not induced by doxycycline (data not shown).

The secretion of FLAG-annexin V was confirmed by Western blot analysis of the culture medium. Immunoblotting with anti-FLAG antibodies demonstrated that there was FLAG-annexin V in both the cell lysate and the culture medium of doxycycline-induced cells (Figure 5.3A). The fusion protein containing the FLAG epitope migrates at a molecular mass of 36 kDa instead of 35 kDa for native annexin V. There was no FLAG-annexin V in either the cell lysate or culture medium of cells not induced by doxycycline, suggesting that the tetracycline-inducible system was not “leaky” (Figure 5.4B). When using anti- annexin V antibody for the immunoblot, the 36 kDa annexin V but not the 35 kDa annexin V appeared in the culture medium of doxycycline-induced cells, whereas both were seen in the cell lysate (Figure 5.4C). This data suggests that endogenous annexin V is not secreted from thyroid cells, further confirming our earlier findings that, without a signal peptide, annexin V is not secreted into the extracellular spaces (Chapter III). Non-induced cells did not have the 36 kDa annnexin V in the medium or the cell lysate. The 35 kDa endogenous annexin V was only present in the cell lysate (Figure 5.4D).

Growth curves of non-induced and induced cells. To evaluate whether targeting expression of annexin V to the ER would grossly affect cell growth, the MTT assay was used to evaluate the respective cell numbers 24, 48, and 72 hours after doxycycline induction. The growth and viability of the cells was essentially same, as the growth curves of non-induced and induced cells were coincident (Figure 5.5) (n=96, p>0.05).

74 Equilibrium 45Ca2+ uptake and release by the ER. To evaluate whether overexpression of a Ca2+ binding protein in the ER would affect the calcium sequestration/ release dynamics, the calcium uptake at equilibrium was initially evaluated in non- permeabilized cells. 45Ca2+ was loaded for 48 hours, a time period shown previously to be sufficient to reach 45Ca2+ equilibrium in cultured cells (Lin, 1999). Doxycycline-induced PC-rTTA7 cells stored approximately 30% more 45Ca2+ in 48 hours than non-induced cells (n=6, p<0.01) (Figure 5.6A). Thapsigargin inhibits re-uptake of calcium by the calcium pump (SERCA) in the ER membrane. After loading with 45Ca2+ for 48 hours to reach equilibrium, cells were treated with thapsigargin. Cells expressing annexin V in the ER released approximately one third more 45Ca2+ than cells that did not express annexin V in the ER (n=6, p<0.01) (Figure 5.6B). These findings suggest that at equilibrium, cells expressing annexin V in the ER sequester more calcium due to the calcium binding ability of the protein and the stored calcium by annexin V is thapsigargin-sensitive.

45Ca2+ uptake and release in permeabilized cells. When cells were permeabilized with saponin, they rapidly incorporated 45Ca2+ and reached equilibrium by 45 minutes. Simi- lar to non-permeabilized cells, at equilibrium, doxycycline-induced cells take up about 30% more 45Ca2+ than non-induced cells (Figure 5.7A) (n=6, p<0.05). There was no difference in initial 45Ca2+ uptake between the two groups of cells. The total calcium uptake calculated by areas under the curve shows that doxycycline-induced cells took up more calcium

than non-induced cells (n=6, p<0.05). Permeabilized cells were loaded with 45Ca2+ for 45 minutes and then were challenged with 1,4,5-IP3, which binds to its receptor on the ER membrane and stimulates calcium release. Again cells overexpressing annexin V released more calcium than cells that did not overexpress annexin V (n=6, p<0.01, calculated by comparing areas under the curves) (Figure 5.7B). Both non-induced and induced cells were desensitized rapidly after treated with IP3. At the peak response, the induced cells released

75 about 30% more calcium than non-induced cells (n=6, p<0.05). These results suggest that the calcium sequestered by ER is releasable upon the stimulation with IP3.

Targeting annexin V to the ER: thyroglobulin synthesis, process and secretion. To investigate whether the presence of annexin V in the secretory pathway affects the synthesis, processing, and secretion of the secretory protein thyroglobulin, pulse-chase experiments were carried out with non-induced and doxycycline-induced cells. Thyroglobulin synthesis was evaluated by the time course (15-90 min) of thyroglobulin labeling, which represents the rate of the protein synthesis. There was no significant difference between non-induced cells and induced cells (n=3, p>0.05, calculated by comparing areas under curves) (Figure 5.8).

The intracellular and secreted thyroglobulin from non-induced cells had the same molecular weights as that from induced cells, as demonstrated by 4% SDS-PAGE (Figure 5.9). This indicates that the protein was processed identically in both groups. Western blot analysis of thyroglobulin from non-induced and doxycycline-induced cells demonstrated that there is only one band corresponding to intact thyroglobulin (Figure 5.10A). There were no changes of the concentrations of the five chaperonins (calreticulin, Bip, ERp72, PDI and Grp94) analyzed by Western blot.

Pulse-chase experiments showed that at each chase time point, the percentage of labeled thyroglobulin secreted from doxycycline-induced cells was not significantly different from non-induced cells (Figure 5.11) (n=8, p<0.05, calculated by comparing areas under the curves). These results indicated that targeting annexin V to the ER lumen did not affect thyroglobulin synthesis, processing or secretion.

76 DYKDDDDK

CMV SP Annexin V poly A

MSALLILALVGAVAAVA

Figure 5.1. The transgene designed to secrete annexin V via the ER/Golgi secretory pathway. Annexin V cDNA was tagged with the eight amino acid FLAG epitope. A secretory signal derived from preprotrypsin (SP) was incorporated at the N-terminus for targeting to ER. The transgene was under control of CMV promoter and a tetracycline-inducible system.

77 CMV rtetR VP16 rTTA

rTTA rTTA

- D o x + D o x

Ann V Ann V

tetO tetO

Figure 5.2. Schematic description of tetracycline-inducible system (tet-on). With doxycycline, rTTA binds to tetO and the transcription of annexin V is initiated. Without doxycycline, there is no binding of rTTA to tetO and no transcription of annexin V. CMV: promoter. rtetR: reverse tetracycline repressor. VP16: herpes simplex virus transcription activation domain. rTTA: reverse tetracycline transactivator. Dox: Doxycycline. tetO: tetracycline operator. AnnV: annexin V.

78 A

Figure 5.3. Comparison of the localization of ER-targeted and endogenous annexin V in PCrTTA7 cells using indirect immunofluorescent microscopy. PCrTTA7 cells were stably transfected with transgene and induced with doxycycline as describe in the text. The transfected cells were stained with anti-FLAG monoclonal antibody that only recognizes FLAG- annexin V, the transgene products. FLAG-annexin V is localized in the ER and Golgi of the cell (A). In non-transfected cells, annexin V is localized by anti-annexin V antibodies in the nucleus and cytoplasm. No ER/Golgi distribution pattern is seen (B). 40X

79 te te ABa a m s m s iu y iu ly d l V d ll l l e e e e fA m c fAV m c 36 Kd 36 Kd

CDte te a a m s m s iu ly iu ly l d l V d V e e V e fAV A m c fA A m cell

35 Kd 35 Kd

Figure 5.4. Western blot analysis of annexin V secretion from PCrTTA7 cells. Cells were transfected and induced as described in the text. Media were collected and passed through a phospholipid-conjugated phenyl-Sepharose column. The fractions from the columns were applied to SDS-PAGE and transferred to nitrocellulose membrane. Both anti-FLAG (A, B) and anti-annexin V (C, D) antibodies were employed to detect FLAG-annexin V in the culture medium and in cell lysates of non-induced (A, C) and doxycycline-induced (B, D) cells. FLAG-annexin V (36 kDa) was not detected in the medium and cell lysate of non- induced cells (A) but was present in both the medium and cell lysate of induced cells (B). In the non-induced cells endogenous 35 kDa annexin V is present in the cell lysate but not in the medium (C). The presence of FLAG-annexin V (36 kDa) in the medium and cell lysate of doxycycline-induced cells was confirmed by the staining with anti-annexin V antibody (D). fAV: FLAG-annexin V standard. AV: recombinant annexin V standard.

80 0.38

Noninduced 0.36 Induced

0.34

n 0.32

s 0.30

0.28

0.26

0.24 20 30 40 50 60 70 80 Time (hours)

Figure 5.5. Cell growth curves evaluated by MTT assay. Cells were plated onto 96 well plates with 1000 cell/well in media with or without doxycycline. At 24, 48, and 72 hours, cell viability was measured by adding MTT. There is no difference of growth rate between non-induced (open circles) and induced (filled circles) cells (n=96, p>0.05).

81 A

Figure 5.6. Equilibrium 45Ca2+ uptake and release by thapsigargin. Non-induced and induced cells were loaded with 2µCi/ml of 45Ca2+ for 48 hours followed by washing with KRH buffer and PBS. For measuring uptake, cells were lysed, mixed with scintillation solution and counted. The induced cells uptake about 30% more 45Ca2+ than non-induced cells (A). For measurement of 45Ca2+ released by thapsigargin, the washed cells were treated with 20µM thapsigargin for 20 minutes in KRH supplemented with 3mM EGTA. The supernatants were collected and counted. The induced cells release about 30% more 45Ca2+ than non-induced cells upon stimulation with thapsigargin. Equal amounts of cells (106) were used for the experiments. (n=6, p<0.05)

82 12.0

) A

p 10.0

4 c

E 8.0

e (1 6.0

p 4.0

a Noninduced C 2.0 5 Induced

0.0 010203040506070 Time (min)

2200

2000 B Noninduced

) Induced 1800

p 1600

e 1400

la 1200

+ 1000

C 800

600

400 0 2 4 6 8 10 12 14 16 18 20 22 Time (min)

Figure 5.7. 45Ca2+ uptake and release in saponin-permeabilized cells. Non-induced and doxycycline-induced cells were permeabilized with 50 µg/ml saponin in calcium loading buffer for 5 minutes at room temperature, followed by 3 washes with PBS. 10 µCi 45Ca2+ was loaded from 10 to 60 minutes (A) or 45 minutes (B) followed by washing with efflux buffer. For measuring uptake, washed cells were lysed and counted. At equilibrium (after 45 minutes), the induced cells take up about 30% more 45Ca2+ than non-induced cells. The rate of uptake is similar in both non-induced and induced cells (A). After 45 minutes of 45 2+ loading with Ca , cells were washed and incubated with 20µM IP3 in efflux buffer. The buffer was collected at 2-minute intervals. At the peak response, the induced cells release 45 2+ 30% more Ca than non-induced cells. The cells became rapidly desensitized to IP3 (B). (n=6, p<0.05).

83 Noninduced Induced

Figure 5.8. The synthesis of thyroglobulin in non-induced and induced cells. Non-induced and induced cells were incubated with medium lacking methionine and cysteine for 20 minutes followed by 35S-methionine labeling for various times (15, 30, 45, 60, and 90 minutes). At each time point, cells were washed 3X with PBS, treated with 50mM iodoacetamine and lysed in the lysis buffer with protease inhibitors. The samples were applied to 4% SDS- PAGE. The gels were dried, exposed to X-ray films and quantitated in phosphoimager. The data were analyzed using Microsoft® Exel. There is no difference in the amount of labeled thyroglobulin in non-induced (empty bar) and induced (filled bar) cells at any time point. (n=8, p>0.05).

84 d d e e c c u d u d ABd e d e in c in c n u n u o d o d n in n in

330 Kd 330 Kd

cell medium lysate

Figure 5.9. The evaluation of thyroglobulin processing in non-induced and doxycycline- induced cells. The cell lysate and media from 35S-methionine labeled cells were electrophoresed in 4-12% gradient SDS-PAGE. The gels were then dried and exposed to X-ray films followed by phosphoimager analysis. Both in the cell lysate (A) and in the media (B), thyroglobulin was present as a 330kd protein.

85 ed d uc uce d d d d In ce n e - u -I n d n o n o duc N I N In CRTL

ERp72

BiP

PDI

Grp94

Figure 5.10. Western blot analysis of thyroglobulin and chaperonin proteins in non-induced and induced cells. Non-induced and induced cells were trypsinized and counted. Equiva- lent numbers of cells were lysed in lysis buffer and the samples were applied to SDS-PAGE. The proteins were then transferred to nitrocellulose membrane and probed with corresponding antibodies followed by peroxidase-conjugated secondary antibodies and color reaction. Thyroglobulin appeared as a single band protein with a molecular weight of 330kd (A). There are no differences in the level of any of the chaperons analyzed from non-induced and doxycycline-induced cells (B).

86 0 12345hr cell lysate

media

Noninduced Induced

Figure 5.11. Pulse chase analysis to assess thyroglobulin secretion. Non-induced and induced cells were starved in medium lacking methionine and cysteine for 20 minutes and then labeled with 35S-methionine for 60 minutes. After being washed three times with PBS, cells were incubated with chase medium that contains 10 times higher concentrations of methionine and cysteine. Media and cells were collected at 1, 2, 3, 4, and 5 hours respec- tively. Cells were first treated with 50mM iodoacetamine for 10 minutes and then lysed in lysis buffer. Both cell lysates and media were electrophoresed in 4-12% SDS-PAGE. The gels were dried and exposed on X-ray films followed by analyzing in phosphoimager. The data were analyzed using Microsoft® Exel. Thyroglobulin present in cell lysates and media at the specified time points is demonstrated in the protein gels (Top). The quantitated data were plotted as chase times vs quantity of thyroglobulin in the cell lysate and in the media (Bottom). (n=8, p>0.05).

87 DISCUSSION

Annexin V has been studied extensively as an anticoagulant and anti-inflammatory protein for many years (Chap, 1988, Buckland, 1998a, Mira, 1997, Andree, 1992, Romisch, 1991). As discussed previously, annexin V is an intracellular protein, thus, secretion of annexin V from gene-transfected cells may prove to be valuable in the treatment of thrombotic and inflammatory conditions in vivo. It was anticipated that targeting of annexin V to the secretory pathway might disrupt synthesis and processing of secretory proteins, alter membrane lipid asymmetry, and modify intracellular calcium homeostasis. Demonstrating that it was possible to secrete a genetically engineered annexin V from mammalian cells suggested that it would be feasible to introduce the protein into the circulation of whole animals. This would allow for the direct analysis of the anticoagulant and anti-inflammatory activity of annexin V in vivo and also explore the possibility of treating certain hypercoagulable conditions and inflammatory diseases with circulating annexin V.

The expression of annexin V in the ER lumen increased calcium storage and uptake by the ER as predicted by the calcium binding capacity of the protein. The more important issue here is whether the calcium sequestered by annexin V could be released from ER upon ligand stimulation. My data suggest that the calcium retained in the ER by annexin V is releasable upon ligand stimulation. The calcium sequestered by annexin V in the ER was also thapsigargin-sensitive. Thus, overexpression and targeting annexin V into ER lumen modestly increased calcium store and uptake by the ER. The excessively sequestered calcium by ER lumenal annexin V was agonist-sensitive, and rapidly exchangeable. In the aspects of calcium homeostasis, the ER lumenal annexin V behaved like the other ER lumenal calcium binding proteins such as calreticulin, BiP and PDI (Michalak, 1992, Lievremont, 1997, Campell, 1983). The increase in ER lumenal calcium did not grossly alter cellular functions such as growth.

88 When annexin V is targeted to the ER lumen, the high affinity for phospholipids would affect the lipid asymmetry across the ER membrane. I found that lumenal ER annexin V did not alter the surface phopholipids of the cells, assessed by fluorescein-conjugated annexin V binding (FITC-annexin V) (data not shown). This approach is widely used to evaluate negatively charged phospholipid exposure on the surfaces of apoptotic cells.

In order to evaluate ER function in the synthesis, processing and secretion of a secretory protein, thyroglobulin was chosen as a marker. Thyroglobulin is a 330 kDa glycoprotein synthesized and secreted by PC-rTTA7 cells. It is the major secretory protein in thyroid epithelial cells and serves as a matrix for the synthesis and iodination of thyroid hormones (Dunn, 1987). The nascent protein contains a single peptide of 2750 amino acids with a molecular weight of 300 kDa. Thyroglobulin is folded and extensively modified while passing through the secretory pathway, gaining 10% of its molecular weight from glycosylation. It is secreted to the thyroid colloid compartment as a 660 kDa homodimer (Wadar, 1974, Mlathiery, 1989, Edelhoch, 1986). Therefore, thyroglobulin makes an ideal candidate for studying the integrity of the secretory pathway.

The synthesis and secretion of thyroglobulin were not affected in annexin V secreting cells. There are also no changes in the concentration of other calcium binding proteins, which indicates that the processing of thyroglobulin is not affected. It is known that if a secretory protein is not properly modified in the secretory pathway, there will be various sizes of thyroglobulin protein present in the cell and the concentrations of chaperonins’ increase (Kim, 1998). My results demonstrated that even with its high affinity for anionic phospholipids, annexin V present in the ER lumen did not disrupt ER membrane to alter ER functions in synthesis, processing and secretion of thyroglobulin.

89 Considerable attention has been focused on the development of annexin V as a potent anticoagulant, which would be a potential therapeutic agent for hypercoagulation diseases such as antiphospholipid syndrome, stroke and deep venous thrombosis (DVT). Targeting annexin V to the secretory pathway had minimal effects on ER functions. Our results suggest and demonstrate the potentials of using an engineered annexin V in developing treatment strategies for thrombotic and inflammatory diseases.

90 CHAPTER VI

CONCLUSIONS

I have established that annexin V plays a critical role in the maintenance of murine placental integrity by neutralizing annexin V in the placenta. There are annexin V binding sites on the surface of placental trophoblasts as demonstrated by infusing FLAG-tagged annexin V into pregnant mice and localizing FLAG epitope on the surfaces of the sinuses containing maternal blood. The binding sites for annexin V are possibly the exposed anionic phospholipids on the surfaces of syncytiotrophoblasts, as can be speculated from the calcium-dependent phospholipid binding properties of annexin V. The annexin V bound to the trophoblast surface maybe crucial for shielding the exposed anionic phospholipids which would otherwise propagate blood coagulation and inflammation by providing substrates for phospholipase A2. The significance of trophoblast surface annexin V was confirmed when it was neutralized by intravenously administered anti-annexin V antibodies, and the neu- tralization of annexin V caused fetal losses, placental thrombosis, necrosis and inflammation. These results may explain the high thrombotic status and recurrent intrauterine fetal loss seen among patients with autoantibodies against proteins associated with coagulation. Such an association is particularly interesting with respect to those patients who test negative for antiphospholipid antibodies. My study also suggests that factors interfering with the binding of annexin V to the surfaces of syncytiotrophobalsts contribute to the compromise of placental functions in patients with antiphospholipid syndrome.

The compromise of placental function after neutralizing annexin V in the placenta suggested annexin V is on the surface of syncytiotrophoblasts. This possibility was confirmed when annexin V epitope was localized on the surfaces of BeWo cells, a human trophoblast cell line. However, annexin V was not detected in the culture medium of BeWo

91 Figure 6.1. The schematic demonstration of the role annexin V might play in the placenta. When the cells expose negative charged phospholipids, the phospholipid-catalyzed coagulation reactions can be initiated (A). When there are annexin V proteins available, annexin V binds to the anionic phospholipids with an affinity higher than that of the coagulation factors. The anionic phospholipid surface is shielded by annexin V and is not accessible to the coagulation factors (B). Anti-annexin V antibodies remove annexin V from the surface of syncytiontrophoblasts, and the exposed anionic phospholipids are again available to the coagulation factors (C). Adapted from Rand et al, New England Journal of Medicine, 1997.

92 cells, even after the differentiation of the cells into syncytiotrophoblasts. The differentiation of BeWo cells was accompanied by externalization of anionic phospholipids, which is demonstrated by increased FITC-annexin V binding on the cell surfaces. Annexin V was not detected in the culture medium of human umbilical vein endothelial cells (HUVEC), another type of cells constantly in contact with blood and another abundant source of annexin V. Therefore, annexin V is not secreted to the extracellular spaces. The cell surface annexin V may come from ajacent damaged cells that release their intracellular annexin V. Only when a signal peptide was incorporated into the protein was annexin V efficiently secreted from cells.

My ultimate goal was to target annexin V to the traditional secretory pathway of animal cells and secrete annexin V to the blood circulation of transgenic mice. Several attempts were made to introduce annexin V into transgenic mice. Unfortunately all the mice that had the transgene failed to express the protein. Cell studies were carried out in thyroid epithelial cells to assure that unconventionally targeting annexin V to the secretory pathway does not affect cell function, especially functions of endoplasmic reticulum (ER). Overexpression of annexin V in thyroid epithelial cells had no effect on cell growth and the synthesis, processing, and secretion of thyroglobulin, a secretory protein abundantly synthesized and secreted by thyroid epithelial cells. Expressing annexin V within the ER increased intra-lumenal ER calcium storage in non-permeabilized cells and calcium uptake by saponin-permeabilized cells. The excess stored calcium could be released upon stimula-

2+ tion with thapsigargin, a Ca -ATPase inhibitor, and IP3, a ligand that binds to its receptor and triggers calcium release. The lipid asymmetry was not changed in the annexin V expressing cells, as demonstrated by FITC-annexin V binding. My results demonstrated that even though annexin V has a high capacity for binding calcium and a high affinity for anionic phospholipids, targeting it to the ER does not significantly alter ER function as calcium regulator and the site of protein synthesis, processing and secretion.

93 Even though my attempts to target secreting annexin V to the blood circulation of transgenic mice were not successful, my results suggested that future attempts may be at- tainable. With a ligand inducible system, a sustained level of annexin V could be obtained when required. Such an animal could be used to evaluate the therapeutic potential of annexin V in disease models for hypercoagulable conditions. The studies also provide initial results for developing a treatment strategy for patients with thrombotic and inflammatory diseases.

94 BIBLIOGRAPHY

Ahn NG, Teller DC, Bienkowski MJ, McMullen BA, Lipkin EW, de Haen C. Sedimenta- tion equilibrium analysis of five lipocortin-related phospholipase A2 inhibitors from human placenta. Evidence against a mechanistically relevant association between enzyme and inhibitor. J Biol Chem 1988; 263: 18657-63.

Albuquerque ML, Akiyama SK, Schnaper HW. Basic fibroblast growth factor release by human coronary artery endothelial cells is enhanced by matrix proteins, 17beta- estradiol, and a PKC signaling pathway. Exp Cell Res 1998; 245: 163-9.

Almus FE, Rao LV, Rapaport SI. Regulation of factor VIIa/tissue factor functional activity in an umbilical vein model. Arterioscler Thromb 1993; 13: 105-11.

Andree HA, Hermens WT, Hernker HC, Willems GM. Displacement of factor Va by annexin V. In: Phospholipid binding and anticoagulant action of annexin V. Maastricht. The Netherlands: Universitaire Pers Maastricht. 1992, 73-85.

Andree HA, Reutelingsperger CP, Hauptmann R, Hemker HC, Hermens WT, Willems GM.

Binding of vascular anticoagulant alpha (VAC alpha) to planar phospholipid bilay- ers. J Biol Chem 1990; 265: 4923-8.

Andree HA, Stuart MC, Hermens WT, Reutelingsperger CP, Hemker HC, Frederik PM, Willems GM. Clustering of lipid-bound annexin V may explain its anticoagulant effect. J Biol Chem 1992; 267: 17907-12

95 Andrei C, Dazzi C, Lotti L, Torrisi MR, Chimini G, Rubartelli A. The secretory route of the leaderless protein interleukin 1beta involves exocytosis of endolysosome-related vesicles. Mol Biol Cell 1999; 10: 1463-75.

Barton GJ, Newman RH, Freemont PS, Crumpton MJ. Amino acid sequence analysis of the annexin super-gene family of proteins. Eur J Biochem 1991; 198: 749-60.

Beale EG, Schaefer IM, Li QA. Culture at high density increases phosphoenolpyruvate carboxykinase messenger RNA in H4IIEC3 hepatoma cells. Mol Endocrinol 1991; 5:661-9.

Boersma AW, Nooter K, Oostrum RG, Stoter G. Quantification of apoptotic cells with fluorescein isothiocyanate-labeled annexin V in chinese hamster ovary cell cultures treated with cisplatin. Cytometry 1996; 24:123-30.

Bohn H. Placental and pregnancy proteins. In: Carcinoembryonic proteins. Lehman FG. ed. Vol. 1. Amsterdam: Elsvier, North Holland Biomedical Press, 1979; pp 289-299.

Bona CA. Postulates defining pathogenic autoantibodies and T cells. Autoimmunity 1991;

10: 169-72.

Buckland AG, Wilton DC. Inhibition of human cytosolic phospholipase A2 by human annexin V. Biochem J 1998a; 329 (Pt 2):369-72.

Buckland AG, Wilton DC. Inhibition of secreted phospholipases A2 by annexin V. Compe- tition for anionic phospholipid interfaces allows an assessment of the relative inter-

96 facial affinities of secreted phospholipases A2. Biochim Biophys Acta 1998b; 1391:367-76.

Campbell KP, MacLennan DH, Jorgensen AO, Mintzer MC. Purification and characterization of calsequestrin from canine cardiac sarcoplasmic reticulum and identification of the 53,000 dalton glycoprotein. J Biol Chem 1983; 258:1197-204.

Campos B., Mo Y.D., Mealy T.R., Li C.W., Swairjo, M.A., Balch C., Head J.F., Retzinger, G., Dedman, J.R., Seaton, B.A. Mutational and crystallographic analyses of interfa- cial residues in annexin V suggest direct interactions with phospholipid membrane components. Biochemistry 1998; 37: 8004-10.

Capila I, VanderNoot VA, Mealy TR, Seaton BA, Linhardt RJ. Interaction of heparin with annexin V. FEBS Lett 1999; 446:327-30.

Cassuto H, Aran A, Cohen H, Eisenberger CL, Reshef L. Repression and activation of transcription of phosphoenolpyruvate carboxykinase gene during liver development. FEBS Lett 1999; 457:441-4.

Chap H, Comfurius P, Bevers EM, Fauvel J, Vicendo P, Douste-Blazy L, Zwaal RF. Poten- tial anticoagulant activity of lipocortins and other calcium/phospholipid binding proteins. Biochem Biophys Res Commun 1988; 150:972-8.

Cheng HM. Antiphospholipid antibodies, annexin V, and pregnancy loss. N Engl J Med 1997; 337:1630-1

97 Chollet P, Malecaze F, Hullin F, Raynal P, Arne JL, Pagot V, Ragab-Thomas J, Chap H. Inhibition of intraocular fibrin formation with annexin V. Br J Ophthalmol 1992; 76(8):450-2.

Christmas P, Callaway J, Fallon J., Jones J, Haigler HT. Selective secretion of annexin 1, a protein without a signal sequence, by the human prostate gland. J Biol Chem 1991; 266: 2499-507.

Corradi A, Franzi AT, Rubartelli A. Synthesis and secretion of interleukin-1 alpha and interleukin-1 receptor antagonist during differentiation of cultured keratinocytes. Exp Cell Res 1995; 217: 355-62.

Crompton MR, Moss SE, Crumpton MJ. Diversity in the lipocortin/calpactin family. Cell 1988;55:1-3.

Dedman JR, Welch MJ, Means AR. Ca2+ -dependent regulator. Production and characterization of a monospecific antibody. J Biol Chem 1978 253:7515-21.

Delorme MA, Burrows RF, Ofosu FA, Andrew M. Thrombin regulation in mother and

fetus during pregnancy. Semin Thromb Hemost 1992; 18:81-90.

Devine DV, Brigden ML. The antiphospholipid syndrome: when does the presence of antiphospholipid antibodies require therapy? Postgrad Med 1996; 99:105-8, 113-25.

Dorner AJ, Wasley LC, Kaufman RJ. Overexpression of GRP78 mitigates stress induction of glucose regulated proteins and blocks secretion of selective proteins in Chinese hamster ovary cells. EMBO J 1992; 11:1563-71.

98 Dubois T, Oudinet JP, Mira JP, Russo-Marie F. Annexins and protein kinases C. Biochim Biophys Acta 1996; 1313:290-4.

Dunn JT, Anderson PC, Fox JW, Fassler CA, Dunn AD, Hite LA, Moore RC. The sites of thyroid hormone formation in rabbit thyroglobulin. J Biol Chem 1987; 262:16948-52.

Edelhoch H and Robbins J. in The Thyroid. Ingbar SH and Braverman LE eds. 5th Ed., Lippincott, Philadepia. pp. 98-115.

Fadok VA, Voelker DR, Campbell PA, Cohen JJ, Bratton DL, Henson PM. Exposure of phosphatidylserine on the surface of apoptotic lymphocytes triggers specific recognition and removal by macrophages. J Immunol 1992; 148:2207-16.

Feinstein DI. Lupus anticoagulant, anticardiolipin antibodies, fetal loss, and systemic lupus erythematosus. Blood 1992; 80: 859-62.

Flaherty MJ, West S, Heimark RL. Placental anticoagulant protein I: measurement in extracellular fluids and cells of the hemostatic system. J Lab Clin Med 1990; 115:174-79.

Flores H, Alleyne GA. Phosphoenolpyruvate carboxykinase of kidney. Subcellular distribution and response to acid-base changes. Biochem J 1971; 123:35-9.

Funakoshi T, Heimark RL, Hendrickson LE, McMullen BA, Fujikawa K. Human placental anticoagulant protein: isolation and characterization. Biochemistry 1987b; 26:5572-8.

99 Funakoshi, T., Hendrickson, L.E., McMullen, B.A., Fujikawa, K. Primary structure of human placental anticoagulant protein. Biochemistry 1987a; 26: 8087-92.

Goulding NJ, Godolphin JL, Sharland PR, Peers SH, Sampson M, Maddison PJ, Flower RJ. Anti-inflammatory lipocortin 1 production by peripheral blood leucocytes in response to hydrocortisone. Lancet 1990; 335:1416-8.

Hanson RW, Garber AJ. Phosphoenolpyruvate carboxykinase. I. Its role in gluconeogenesis. Am J Clin Nutr 1972; 25:1010-21.

Hanson RW, Reshef L. Regulation of phosphoenolpyruvate carboxykinase (GTP) gene expression. Annu Rev Biochem 1997; 66:581-611.

Hatzoglou M, Lamers W, Bosch F, Wynshaw-Boris A, Clapp DW, Hanson RW. Hepatic gene transfer in animals using retroviruses containing the promoter from the gene for phosphoenolpyruvate carboxykinase. J Biol Chem 1990; 265:17285-93.

Hauptmann R, Maurer-Fogy I, Krystek E, Bodo G, Andree H, Reutelingsperger CP. Vascu- lar anticoagulant beta: a novel human Ca2+/phospholipid binding protein that in-

hibits coagulation and phospholipase A2 activity. Its molecular cloning, expression and comparison with VAC-alpha. Eur J Biochem 1989; 185:63-71.

Hertogs K, Depla E, Crabbe T, De Bruin W, Leenders W, Moshage H, Yap SH. Spontane- ous development of anti-hepatitis B virus envelope (anti-idiotypic) antibodies in animals immunized with human liver endonexin II or with the F(ab’)2 fragment of anti-human liver endonexin II immunoglobulin G: evidence for a receptor-ligand-

100 like relationship between small hepatitis B surface antigen and endonexin II. J Virol 1994; 68:1516-21.

Huber R, Schneider M, Mayr I, Romisch J, Paques EP. The calcium binding sites in human annexin V by crystal structure analysis at 2.0 A resolution. Implications for membrane binding and calcium channel activity. FEBS Lett 1990; 275:15-21.

Hughes GRV. Thrombosis, abortion, cerebral disease and lupus anti-coagulant. Br Med J 1983; 1088-89.

Hullin F, Raynal P, Ragab-Thomas JM, Fauvel J, Chap H. Effect of dexamethasone on prostaglandin synthesis and on lipocortin status in human endothelial cells. Inhibi- tion of prostaglandin I2 synthesis occurring without alteration of arachidonic acid liberation and of lipocortin synthesis. J Biol Chem 1989; 264:3506-13.

Iwasaki A, Suda M, Nakao H, Nagoya T, Saino Y, Arai K, Mizoguchi T, Sato F, Yoshizaki H, Hirata M, et al. Structure and expression of cDNA for an inhibitor of blood coagulation isolated from human placenta: a new lipocortin-like protein. J Biochem (Tokyo) 1987; 102:1261-73.

Jaye M, Howk R, Burgess W, Ricca GA., Chiu IM., Ravera MW., O’Brien SJ., Modi WS, Maciag T, Drohan WN. Human endothelial cell growth factor: cloning, nucleotide sequence, and chromosome localization. Science 1986; 233: 541-5.

Kaburaki J, Kuwana M, Yamamoto M, Kawai S, Ikeda Y. Clinical significance of anti- annexin V antibodies in patients with systemic lupus erythematosus. Am J Hematol 1997; 54:209-13.

101 Kaetzel MA, Hazarika P, Dedman JR. Differential tissue expression of three 35-kDa annexin calcium-dependent phospholipid-binding proteins. J Bio Chem 1989; 264:14463-70.

Kaneko N, Matsuda R, Hosoda S, Kajita T, Ohta Y. Measurement of plasma annexin V by ELISA in the early detection of acute myocardial infarction. Clin Chim Acta 1996; 251:65-80.

Kaplan R, Jaye M, Burgess WH, Schlaepfer DD, Haigler HT. Cloning and expression of cDNA for human endonexin II, a Ca2+ and phospholipid binding protein. J Biol Chem 1988; 263:8037-43.

Katsuragawa H, Rote NS, Inoue T, Narukawa S, Kanzaki H, Mori T. Monoclonal antiphosphatidylserine antibody reactivity against human first-trimester placental trophoblasts. Am J Obstet Gynecol 1995; 172:1592-7.

Kim PS, Arvan P. Folding and assembly of newly synthesized thyroglobulin occurs in a pre-Golgi compartment. J Biol Chem 1991; 266:12412-8.

Kim PS, Hossain SA, Park YN, Lee I, Yoo SE, Arvan P. A single amino acid change in the

acetylcholinesterase-like domain of thyroglobulin causes congenital goiter with hy- pothyroidism in the cog/cog mouse: a model of human endoplasmic reticulum storage diseases. Proc Natl Acad Sci U S A 1998; 95:9909-13.

King KL, Moreira KM, Babcock GF, Wang J, Campos B, Kaetzel MA, Dedman JR. Tem- poral inhibition of calmodulin in the nucleus. Biochim Biophys Acta 1998; 1448:245-53.

102 Kondo S, Noguchi M, Funakoshi T, Fujikawa K, Kisiel W. Inhibition of human factor VIIa- tissue factor activity by placental anticoagulant protein. Thromb Res 1987; 48:449- 59.

Krailadsiri P, Seghatchian J, Amiral J, Vissac AM, Contreras M. Annexin V, a new marker of platelet storage lesion: correlation with dMPV. Transfus Sci 1997; 18:223-6.

Krikun G, Lockwood CJ, Wu X-X, Zhou X-D, Guller S, Calandri C, et al. The expression of the placental anticoagulant protein, annexin V, by villous trophoblasts: immunolocalization and in vitro regulation. Placenta 1994; 15:601-12.

Lakasing L, Campa JS, Poston R, Khamashta MA, Poston L. Normal expression of tissue factor, thrombomodulin, and annexin V in placentas from women with antiphospholipid syndrome. Am J Obstet Gynecol 1999; 181:180-9

Lievremont JP, Rizzuto R, Hendershot L, Meldolesi J. BiP, a major chaperone protein of the endoplasmic reticulum lumen, plays a direct and important role in the storage of the rapidly exchanging pool of Ca2+. J Biol Chem 1997; 272:30873-9.

Lim IK, Dumenco LL, Yun J, Donovan C, Warman B, Gorodetzkaya N, Wagner TE, Clapp DW, Hanson RW, Gerson SL. High level, regulated expression of the chimeric P- enolpyruvate carboxykinase (GTP)-bacterial O6-alkylguanine-DNA alkyltransferase (ada) gene in transgenic mice. Cancer Res 1990; 50:1701-8.

Lin P, Yao Y, Hofmeister R, Tsien RY, Farquhar MG. Overexpression of CALNUC (nucleobindin) increases agonist and thapsigargin releasable Ca2+ storage in the Golgi. J Cell Biol 1999; 145:279-89.

103 Lodish HF, Kong N, Wikstrom L. Calcium is required for folding of newly made subunits of the asialoglycoprotein receptor within the endoplasmic reticulum. J Biol Chem 1992; 267:12753-60.

London F, Ahmad SS, Walsh PN. Annexin V inhibition of factor IXa-catalyzed factor X activation on human platelets and on negatively charged phospholipid vesicles. Bio- chemistry 1996; 35:16886-97.

Lucero HA, Lebeche D, Kaminer B. ERcalcistorin/protein-disulfide isomerase acts as a calcium storage protein in the endoplasmic reticulum of a living cell. Comparison with calreticulin and calsequestrin. J Biol Chem 1998; 273:9857-63

Lyden TW, Ng AK, Rote NS. Modulation of phosphatidylserine epitope expression by BeWo cells during forskolin treatment. Placenta 1993; 14: 177-86.

Lyden TW, Vogt E, Ng AK, Johnson PM, Rote NS. Monoclonal antiphospholipid antibody reactivity against human placental trophoblast J Reprod Immunol 1992; 22:1-14.

Mair J. Progress in myocardial damage detection: new biochemical markers for clinicians.

Crit Rev Clin Lab Sci 1997; 34:1-66.

Malthiery Y, Marriq C, Berge-Lefranc JL, Franc JL, Henry M, Lejeune PJ, Ruf J, Lissitzky S. Thyroglobulin structure and function: recent advances. Biochimie 1989; 71:195-209.

Matsuda J, Gotoh M, Saitoh N, Gohchi K, Tsukamoto M, Yamamoto T. Anti-annexin V antibodies in the sera of patients with habitual fetal loss or preeclampsia. Thromb

104 Haemos 1994b; 75:105-6.

Matsuda J, Saitoh N, Gohchi K, Gotoh M, Tsukamoto M. Anti-annexin V antibody in systemic lupus erythematosus patients with lupus anticoagulant and/or anticardiolipin antibody. Am J Hematol 1994a; 47: 56-8.

McGrane MM, Yun JS, Moorman AF, Lamers WH, Hendrick GK, Arafah BM, Park EA, Wagner TE, Hanson RW. Metabolic effects of developmental, tissue-, and cell- specific expression of achimeric phosphoenolpyruvate carboxykinase (GTP)/bovine growth hormone gene in transgenic mice. J Biol Chem 1990; 265:22371-9.

Meldolesi J, Pozzan T. The endoplasmic reticulum Ca2+ store: a view from the lumen. Trends Biochem Sci 1998 Jan; 23:10-4.

Michalak M, Milner RE, Burns K, Opas M. Calreticulin. Biochem J 1992; 285 (Pt 3):681- 92.

Mira JP, Dubois T, Oudinet JP, Lukowski S, Russo-Marie F, Geny B. Inhibition of cytosolic phospholipase A2 by annexin V in differentiated permeabilized HL-60 cells. Evi-

dence of crucial importance of domain I type II Ca2+-binding site in the mechanism of inhibition. J Biol Chem 1997; 272:10474-82.

Missiaen L, De Smedt H, Droogmans G, Casteels R. Ca2+ release induced by inositol 1,4,5-trisphosphate is a steady-state phenomenon controlled by luminal Ca2+ in permeabilized cells. Nature 1992; 357:599-602.

105 Mollenhauer J, Bee JA, Lizarbe MA, von der Mark K. Role of anchorin CII, a 31,000-mol- wt membrane protein, in the interaction of chondrocytes with type II collagen. J Cell Biol 1984; 98:1572-9.

Mollenhauer J, Mok MT, King KB, Gupta M, Chubinskaya S, Koepp H, Cole AA. Expres- sion of anchorin CII (cartilage annexin V) in human young, normal adult, and os- teoarthritic cartilage. J Histochem Cytochem 1999; 47:209-20.

Murphy CT, Peers SH, Forder RA, Flower RJ, Carey F, Westwick J. Evidence for the presence and location of annexins in human platelets. Biochem Biophys Res Commun 1992; 189:1739-46.

Nakamura N, Kuragaki C, Shidara Y, Yamaji K, Wada Y. Antibody to annexin V has antiphospholipid and lupus anticoagulant properties. Am J Hematol 1995; 49:347-48.

O’Brien RM, Lucas PC, Forest CD, Magnuson MA, Granner DK. Identification of a sequence in the PEPCK gene that mediates a negative effect of insulin on transcription. Science 1990; 249:533-7.

Palmi M, Frosini M, Sgaragli GP, Becherucci C, Perretti M, Parente L. Inhibition of interleukin-1 beta-induced pyresis in the rabbit by peptide 204-212 of lipocortin 5. Eur J Pharmacol 1995; 281:97-9.

Park EA, Song S, Vinson C, Roesler WJ. Role of CCAAT enhancer-binding protein beta in the thyroid hormone and cAMP induction of phosphoenolpyruvate carboxykinase gene transcription. J Biol Chem 1999; 274:211-7.

106 Patel YM, Yun JS, Liu J, McGrane MM, Hanson RW. An analysis of regulatory elements in the phosphoenolpyruvate carboxykinase (GTP) gene which are responsible for its tissue-specific expression and metabolic control in transgenic mice. J Biol Chem 1994; 269:5619-28

Pepinsky RB, Tizard R, Mattaliano RJ, Sinclair LK, Miller GT, Browning JL, Chow EP, Burne C, Huang KS, Pratt D, et al. Five distinct calcium and phospholipid binding proteins share homology with lipocortin I. J Biol Chem 1988; 263:10799-811.

Pfaffle M., Ruggiero F., Hofmann H., Fernandez M.P., Selmin O., Yamada Y., Garrone R., von der Mark K. Biosynthesis, secretion and extracellular localization of anchorin CII, a collagen-binding protein of the calpactin family. EMBO J 1988; 7:2335-42.

Pierangeli SS, Barker JH, Stikovac D, Ackerman D, Anderson G, Harris EN, et al. Effect of human IgG antiphospholipid antibodies on an in vivo thrombosis model in mice. Thromb and Haemos 1994; 71: 670-74.

Pinkert CA, Ornitz DM, Brinster RL, Palmiter RD. An albumin enhancer located 10 kb upstream functions along with its promoter to direct efficient, liver-specific expres-

sion in transgenic mice. Genes Dev 1987; 1:268-76.

Piona A, LaRosa L, Tincani A, Faden D, Margo G, Grasso S, et al. Placental thrombosis and fetal loss after passive transfer of mouse lupus monoclonal or human polyclonal anti-cardiolipin antibodies in pregnant naive BALB/c mice. Scand J Immuunol 1995; 41:427-32.

107 Postic C, Shiota M, Niswender KD, Jetton TL, Chen Y, Moates JM, Shelton KD, Lindner J, Cherrington AD, Magnuson MA. Dual roles for glucokinase in glucose homeostasis as determined by liver and pancreatic beta cell-specific gene knock-outs using Cre recombinase. J Biol Chem 1999; 274:305-15.

Rand JH, Wu X-X, Guller S, Gil J, Guha A, Scher J, et al. Reduction of annexin V (placental anticoagulant protein-I) on placental villi of women with antiphospholipid antibodies and recurrent spontaneous abortion. Am J Obstet Gynecol 1994; 171:1566-72.

Rand JH, Wu XX, Guller S, Scher J, Andree HA, Lockwood CJ. Antiphospholipid immunoglobulin G antibodies reduce annexin-V levels on syncytiotrophoblast apical membranes and in culture media of placental villi. Am J Obstet Gynecol 1997b; 177:918-23.

Rand JH, Wu, X-X, Andree HAM, Lockwood CJ, Guller S, Scher J, et al. Pregnancy loss in the antiphospholipid-antibody syndrome—a possible thrombogenic mechanism. N End J Med 1997a; 337:154-60.

Rand JH. “Annexinopathies”—a new class of diseases. N Engl J Med 1999; 340:1035-6.

Rao LV, Tait JF, Hoang AD. Binding of annexin V to a human ovarian carcinoma cell line (OC-2008). Contrasting effects on cell surface factor VIIa/tissue factor activity and prothrombinase activity. Thromb Res 1992; 67:517-31.

Ravanat C, Archipoff G, Beretz A, Freund G, Cazenave JP, Freyssinet JM. Use of annexin- V to demonstrate the role of phosphatidylserine exposure in the maintenance of haemostatic balance by endothelial cells. Biochem J 1992; 282 (Pt 1):7-13.

108 Raynal P, Hullin F, Ragab-Thomas JM, Fauvel J, Chap H. Annexin 5 as a potential regulator of annexin 1 phosphorylation by protein kinase C. In vitro inhibition compared with quantitative data on annexin distribution in human endothelial cells. Biochem J 1993; 292 (Pt 3):759-65.

Relton JK, Strijbos PJ, O’Shaughnessy CT, Carey F, Forder RA, Tilders FJ, Rothwell NJ. Lipocortin-1 is an endogenous inhibitor of ischemic damage in the rat brain. J Exp Med 1991; 174:305-10.

Reshef L, Hanson RW, Ballard FJ. A possible physiological role for glyceroneogenesis in rat adipose tissue. J Biol Chem 1970; 245:5979-84.

Reutelingsperger CP, Hornstra G, Hemker HC. Isolation and partial purification of a novel anticoagulant from arteries of human umbilical cord. Eur J Biochem 1985; 151:625-9.

Rodr’guez-garc’a MI, Fern‡ndez JA, Rodr’guez A, Fern‡ndez MP, Gutierrez C, Torre- Alonso JC. Annexin V autoantibodies in rheumatoid arthritis. Ann Rheum Dis 1996; 55:895-900.

Romisch J, Schuler E, Bastian B, Burger T, Dunkel FG, Schwinn A, Hartmann AA, Paques EP. Annexins I to VI: quantitative determination in different human cell types and in plasma after myocardial infarction. Blood Coagul Fibrinolysis 1992; 3:11-7.

Romisch J, Seiffge D, Reiner G, Paques EP, Heimburger N. In-vivo antithrombotic potency of placenta protein 4 (annexin V). Thromb Res 1991; 61:93-104.

109 Rote NS, Chang J, Katsuragawa H, Ng AK, Lyden TW, Mori T. Expression of phosphatidylserine-dependent antigens on the surface of differentiating BeWo human choriocarcinoma cells. Am J Reprod Immunol 1995; 33:114-21.

Rote NS. Antiphospholipid antibodies, annexin V, and pregnancy loss. N Engl J Med 1997; 337:1630; discussion 1631.

Salafia CM, Starzyk K, Lop(z-Zeno J, Parke A. Fetal losses and other obstetric manifesta- tions in the antiphospholipid syndrome. In: Asherson RA, Cervera R, Piette J~C, Shoenfeld Y, ed. The Antiphospholipid Syndrome. CRC Press. 1996; 117-131.

Schlaepfer DD, Jones J, Haigler HT. Inhibition of protein kinase C by annexin V. Bio- chemistry 1992; 31:1886-91.

Schlaepfer DD, Mehlman T, Burgess WH, Haigler HT. Structural and functional characterization of endonexin II, a calcium- and phospholipid-binding protein. Proc Natl Acad Sci U S A 1987; 84:6078-82.

Seaton BA. Annexin V molecular structure, ligand binding and biological function. In: Annexins: Molecular Structure to Cellular Function. Seaton, BA Ed. R.G. Landes

Company, 1996; 15-29.

Serres M, Comera C, Schmitt D. Annexin 1 regulation in human epidermal cells. Cell Mol Biol (Noisy-le-grand) 1994; 40:701-6.

Sessions A, Horwitz AF. Differentiation-related differences in the plasma membrane phospholipid asymmetry of myogenic and fibrogenic cells. Biochim Biophys Acta 1983; 728:103-11.

110 Sheets EE, Giugni TD, Coates GG, Schlaepfer DD, Haigler HT. Epidermal growth factor dependent phosphorylation of a 35-kilodalton protein in placental membranes. Bio- chemistry 1987; 26:1164-72.

Shimano H, Horton JD, Hammer RE, Shimomura I, Brown MS, Goldstein JL. Overpro- duction of cholesterol and fatty acids causes massive liver enlargement in transgenic mice expressing truncated SREBP-1a. J Clin Invest 1996; 98:1575-84.

Shirotake S., Inaba N., Ohta Y., Fukazawa I., Takamizawa H., Bohn H. Immunochemical measurement of placenta tissue protein 4 in serum. Chem. Pharm. Mull. 1986; 34:2191-2195.

Short MK, Clouthier DE, Schaefer IM, Hammer RE, Magnuson MA, Beale EG. Tissue- specific, developmental, hormonal, and dietary regulation of rat phosphoenolpyruvate carboxykinase-human growth hormone fusion genes in transgenic mice. Mol Cell Biol 1992; 12:1007-20.

Siever, DA., Erickson HP. Extracellular annexin II. Int J Biochem Cell Biol 1997; 29:1219-23.

Silver RM, Pierangeli SS, Edwin SS, Umar F, Harris EN, Scott JR, et al. Pathogenic antibodies in women with obstetric features of antiphospholipid syndrome who have negative test results for lupus anticoagulant and anticardiolipin antibodies. Am J Obstet Gynecol 1997; 176:628-33.

Smith MJ, Koch GL. Multiple zones in the sequence of calreticulin (CRP55, calregulin, HACBP), a major calcium binding ER/SR protein. EMBO J 1989; 8:3581-6.

111 Sopkova J, Renouard M, Lewit-Bentley A. The crystal structure of a new high-calcium form of annexin V. J Mol Biol 1993; 234:816-25.

Speijer H, Jans SW, Reutelingsperger CP, Hack CE, van der Vusse GJ, Hermens WT. Par- tial coverage of phospholipid model membranes with annexin V may completely inhibit their degradation by phospholipase A2. FEBS Lett 1997; 402:193-7.

Stratton JR, Dewhurst TA, Kasina S, Reno JM, Cerqueira MD, Baskin DG, Tait JF. Selec- tive uptake of radiolabeled annexin V on acute porcine left atrial thrombi. Circula- tion 1995; 92:3113-21.

Sun J, Bird P, Salem HH. Effects of annexin V on the activity of the anticoagulant proteins C and S. Thromb Res 1993b; 69:279-87.

Sun J, Bird P, Salem HH. Interaction of annexin V and platelets: effects on platelet function and protein S binding. Thromb Res 1993a; 69:289-96.

Sun J, Salem HH, Bird P. Nucleolar and cytoplasmic localization of annexin V. FEBS Lett 1992; 314:425-9.

Tait JF, Cerqueira MD, Dewhurst TA, Fujikawa K, Ritchie JL, Stratton JR. Evaluation of annexin V as a platelet-directed thrombus targeting agent. Thromb Res 1994; 75:491-501.

Tait JF, Engelhardt S, Smith C, Fujikawa K. Prourokinase-annexin V chimeras. Construc- tion, expression, and characterization of recombinant proteins. J Biol Chem 1995; 270:21594-9.

112 Tait JF, Gibson D, Fujikawa K. Phospholipid binding properties of human placental anticoagulant protein-I, a member of the lipocortin family. J Biol Chem 1989; 264:7944-9.

Tait JF, Sakata M, McMullen BA, Miao CH, Funakoshi T, Hendrickson LE, Fujikawa K. Placental anticoagulant proteins: isolation and comparative characterization four members of the lipocortin family. Biochemistry 1988; 27:6268-76.

Tait JF. Clinical applications of annexins. In: Annexins: molecular structure to cellular function. R.G. Lands Company. 1996; 213-220.

Takehara K., Uchida S., Marumoto N., Asawa T., Osugi S., Kurusu S., Hashimoto I., Kawaminami M. Secretion of recombinant rat annexin 5 by insect cells in a baculovirus expression system. Biochem Biophys Res Commun 1994; 200:1421-7.

Thiagarajan P, Benedict CR. Inhibition of arterial thrombosis by recombinant annexin V in a rabbit carotid artery injury model. Circulation 1997; 96:2339-47.

Thiagarajan P, Tait JF. Binding of annexin V/placental anticoagulant protein I to platelets. Evidence for phosphatidylserine exposure in the procoagulant response of activated platelets. J Biol Chem 1990; 265:17420-3.

Thorgeirsson SS, Ryu DY, Weidner V, Synderwine EG. Carcinogenicity and mutagenicity of heterocyclic amines in transgenic mouse models. Cancer Lett 1999; 143:245-7.

Thorgeirsson SS, Santoni-Rugiu E. Interaction of c-myc with transforming growth factor alpha and hepatocyte growth factor in hepatocarcinogenesis. Mutat Res 1997; 376:221-34.

113 Tilghman SM, Belayew A. Transcriptional control of the murine albumin/alpha-fetopro- tein locus during development. Proc Natl Acad Sci U S A 1982; 79:5254-7.

Tilghman SM, Fisher LM, Reshef L, Ballard FJ, Hanson RW. Partial isolation and translation in vitro of messenger ribonucleic acid for phosphoenolpyruvate carboxykinase (guanosine triphosphate). Biochem J 1976; 156:619-26.

Triplett DA. Antiphospholipid antibodies and thrombosis. A consequence, coincidence, or cause? Arch Pathol Lab Med 1993; 117:78-88.

Trotter PJ, Orchard MA, Walker JH. The intracellular relocation of annexin V in response to the physiological agonist thrombin. Biochem Soc Trans 1993; 21 (Pt 3):281S.

Trotter PJ, Orchard MA, Walker JH. Thrombin stimulates the intracellular relocation of annexin V in human platelets. Biochim Biophys Acta 1994; 1222:135-40.

Uemura K, Inagaki H, Wada Y, Nakanishi K, Asai K, Kato T, Ando Y, Kannagi R. Identi- fication of immuno-reactive lipocortin 1-like molecules in serum and plasma by an enzyme immunoassay for lipocortin 1. Biochim Biophys Acta 1992; 1119:250-5.

Valera A, Fillat C, Costa C, Sabater J, Visa J, Pujol A, Bosch F. Regulated expression of human insulin in the liver of transgenic mice corrects diabetic alterations. FASEB J 1994; 8:440-7. van Bilsen M, Reutelingsperger CP, Willemsen PH, Reneman RS, van der Vusse GJ. Annexins in cardiac tissue: cellular localization and effect on phospholipase activity. Mol Cell Biochem 1992; 116:95-101.

114 van Engeland M, Ramaekers FC, Schutte B, Reutelingsperger CP. A novel assay to mea- sure loss of plasma membrane asymmetry during apoptosis of adherent cells in culture. Cytometry 1996; 24:131-9. van Heerde WL, de Groot PG, Reutelingsperger CP. The complexity of the phospholipid binding protein Annexin V. Thromb Haemost 1995; 73:172-9.

van Heerde WL, Poort S, van ‘t Veer C, Reutelingsperger CP, de Groot PG. Binding of recombinant annexin V to endothelial cells: effect of annexin V binding on endothelial-cell-mediated thrombin formation. Biochem J 1994; 302 (Pt 1):305-12. van Heerde WL, Sakariassen KS, Hemker HC, Sixma JJ, Reutelingsperger CP, de Groot PG. Annexin V inhibits the procoagulant activity of matrices of TNF-stimulated endothelium under blood flow conditions. Arterioscler Thromb 1994; 14:824-30.

Van Ryn-McKenna J, Merk H, Muller TH, Buchanan MR, Eisert WG. The effects of heparin and annexin V on fibrin accretion after injury in the jugular veins of rabbits. Thromb Haemost 1993; 69:227-30.

Vogt E, Ng A-K, Rote NS. Antiphosphatidylserine antibody removes annexin V and facili- tates the binding of prothrombin at the surface of a choriocarcinoma model of trophoblast differentiation. Am J Obstet Gynecol 1997; 177:964-72. von der Mark K, Mollenhauer J. Annexin V interactions with collagen. Cell Mol Life Sci 1997; 53:539-45.

115 Wagar G. Effect of actinomycin D on TSH-induced stimulation of thyroidal protein synthesis in the rat. Acta Endocrinol (Copenh) 1974; 77:64-70.

Walker JH, Boustead CM, Koster JJ, Bewley M, Waller DA. Annexin V, a calcium-dependent phospholipid-binding protein. Biochem Soc Trans 1992; 20:828-33.

Wang JC, Stromstedt PE, Sugiyama T, Granner DK. The phosphoenolpyruvate carboxykinase gene glucocorticoid response unit: identification of the functional domains of acces- sory factors HNF3 beta (hepatic nuclear factor-3 beta) and HNF4 and the necessity of proper alignment of their cognate binding sites. Mol Endocrinol 1999; 13:604- 18.

Wang X., Campos B., Kaetzel M.A., Dedman J.R. Annexin V is critical in the maintenance of murine placental integrity. Am J Obstet Gynecol 1999; 180:1008-16.

Zachowski A, Devaux PF. Transmembrane movements of lipids. Experientia 1990; 46:644-56.

116