The role of a defect in the CDP-ethanolamine pathway

 The role of a defect in the CDP-­ethanolamine pathway in autophagy by Leanne Pereira A Thesis
presented to
The University of Guelph
In partial fulfilment of requirements
for the degree of
Master of Science
in
Human Health and Nutritional Sciences
Guelph, Ontario, Canada
© Leanne Pereira, December, 2012
ABSTRACT THE ROLE OF A DEFECT IN THE CDP-­ETHANOLAMINE PATHWAY IN AUTOPHAGY Leanne Pereira Advisor: University of Guelph, 2012 Dr. Marica Bakovic Autophagy is the process that degrades cytosolic constituents into products that can be recycled for use in energy generation and other processes. The endoplasmic reticulum is responsible for the bulk synthesis of the phospholipid phosphatidylethanolamine (PE) via the CDP-­‐ethanolamine pathway. The aim of the present study was to determine the role of PE synthesis and the CDP-­‐ethanolamine pathway in autophagy. This objective was examined through the use of two novel models deficient in Pcyt2, a gene that encodes the rate-­‐limiting enzyme CTP-­‐
ethanolamine cytidyltransferase (ET) in the CDP-­‐ethanolamine pathway. PCYT2 knockdown in human fibroblast cells did not respond normally to starvation conditions that activate autophagy. Similarly, Pcyt2 knockout in mice showed differences in autophagy induction in/between muscle, liver, and adipose tissue based on metabolic state (fasting/feeding). Pcyt2 knockout mice display evidence of metabolic syndrome at an older age and experiments with these mice determined that there was an effect of age (healthy young mice versus obese older mice) on autophagy induction. It was concluded based on in vitro and in vivo studies that autophagy induction is affected by impairment to the CDP-­‐ethanolamine pathway and subsequent PE synthesis. Acknowledgements Firstly, I would like to thank Dr. Marica Bakovic for accepting me into her lab and for her continuous support over the duration of my term. I would also like to thank Dr. Kelly Meckling and Dr. Jeremy Simpson for agreeing to be apart of my advisory committee and for their guidance during this process. Thank you to my lab-­‐
mates (past and present) specifically Dr. Ratnesh Singh, Dr. Angela Tie, Sugas Sivanesan, John Paul Girardi, Maida Duric and Zvezdan Pavlovic, who have all helped me in some form or another during my time in Guelph. Lastly, thank you to my friends (those I made in Guelph and at home) and to my family, particularly my parents, who have never given up on me in 25 years even at my worst. iii Table of Contents 1.0 INTRODUCTION.................................................................................................................1 1.1 Introduction to Autophagy ..................................................................................................... 1 1.2 Autophagy in Cell Homeostasis............................................................................................. 1 1.3 Autophagy .................................................................................................................................... 2 1.4 Autophagy Machinery .............................................................................................................. 3 1.5 Isolation Membrane Initiation.............................................................................................. 4 1.6 Autophagosome Formation and Elongation..................................................................... 5 1.7 Lysosome Fusion and Autophagosome Degradation .................................................... 6 1.8 Autophagy Regulation.............................................................................................................. 6 1.9 Autophagosomal Membrane Sources ................................................................................. 7 1.10 The Endoplasmic Reticulum in Membrane Formation .............................................. 8 1.11 Source of Lipids for IM and Autophagosome Formation ........................................... 9 1.12 Focus on Phospholipid Synthesis ....................................................................................10 1.13 Rationale and Hypothesis ..................................................................................................13 2.0 EXPERIMENTAL PROCEDURES .................................................................................. 15 2.1 Cell Culture ................................................................................................................................15 2.2 Autophagy Induction ..............................................................................................................15 2.3 Cell Viability ..............................................................................................................................16 2.4 Cell Lysis and Protein Preparation....................................................................................16 2.5 Animal Treatments .................................................................................................................17 2.6 Tissue Homogenization and Protein Preparation........................................................17 2.7 Western Blotting......................................................................................................................17 2.8 Densitometry and Statistical Analysis..............................................................................19 2.9 [14C ]–Ethanolamine Pulse and Pulse-­Chase Labeling.................................................19 3.0 RESULTS............................................................................................................................ 21 3.1 Autophagic response to PCYT2 knockdown in MCH58 human fibroblasts..........21 3.1.1. Characterization of PCYT2 deficient cells................................................................................ 21 3.1.2 The effect of PCYT2 knockdown on autophagy induction .................................................. 23 3.1.3 Pulse [14C]-­ethanolamine radiolabeling of the CDP-­ethanolamine pathway.............. 25 3.1.4 Comparing pulse [14C]-­ethanolamine radiolabeling of the CDP-­ethanolamine pathway between control and PCYT2 KD cells .................................................................................. 28 3.1.5 Pulse-­Chase [14C]-­ethanolamine radiolabeling of the CDP-­ethanolamine pathway 31 3.1.6 Comparing pulse-­chase [14C]-­ethanolamine radiolabeling of the CDP-­ethanolamine pathway between control and PCYT2 KD cells .................................................................................. 34 3.2 Autophagic response to Pcyt2 deficiency in mouse tissue ........................................37 3.2.1 The effects of Pcyt2 knockout on muscle LC3-­II protein content ..................................... 37 3.2.2 The effects of Pcyt2 knockout on liver LC3-­II protein content .......................................... 41 3.3.3 The effects of Pcyt2 knockout on adipose LC3-­II content.................................................... 44 4.0 DISCUSSION...................................................................................................................... 48 4.1 Autophagic response to PCYT2 deficiency in human MCH58 fibroblast cells .....48 4.1.1 Autophagy is not properly induced in PCYT2 KD cells ......................................................... 49 4.1.2 PE synthesis in PCYT2 KD cells does not respond normally to starvation.................... 51 4.2 Autophagic response in Pcyt2 heterozygous mouse tissues.....................................52 4.2.1 The effects of Pcyt2 deficiency on autophagy induction in skeletal muscle................. 54 4.2.2 The effects of Pcyt2 deficiency on autophagy induction in liver ...................................... 55 iv 4.2.3 The effects of Pcyt2 deficiency on autophagy induction in adipose................................ 56 4.3 Regulation of autophagy in mammalian tissues ...........................................................57 4.4 Autophagy induction by phospholipid provision may be tissue specific .............61 4.5 Future Work ..............................................................................................................................62 4.6 Conclusions................................................................................................................................62 5.0 REFERENCES.................................................................................................................... 64 v List of Figures Figure 1. Stages of Autophagy. ............................................................................................................ 3 Figure 2. Pathways involved in phospholipid synthesis. ......................................................12 Figure 3. PCYT2 protein expression is significantly reduced in PCYT2 KD cells. ........22 Figure 4. No significant differences in cell viability between control and PCYT2 KD cells. ....................................................................................................................................................23 Figure 5. Expression of LC3-­‐I, LC3-­‐II, and relative LC3-­‐II/control (Ponceau S) ratio in control and PCYT2 KD cells. . ....................................................................................................24 Figure 6. Pulse experiments with [14C]-­‐ethanolamine demonstrating incorporation into intermediates of the CDP-­‐ethanolamine pathway under NSTV and STV conditions in control cells. . .......................................................................................................26 Figure 7. [14C]-­‐ethanolamine incorporation into intermediates of the CDP-­‐
ethanolamine pathway under NSTV and STV conditions in PCYT2 KD cells. . ...28 Figure 8. [14C]-­‐ethanolamine incorporation into metabolites of the CDP-­‐
ethanolamine pathway comparing control and PCYT2 KD cells under NSTV and STV conditions. . .............................................................................................................................31 Figure 9. Pulse-­‐chase experiments in control cells using [14C]-­‐ethanolamine to demonstrate the degradation of metabolites of the CDP-­‐ethanolamine pathway under normal and starvation conditions. ..........................................................................32 Figure 10.Degradation of CDP-­‐ethanolamine pathway metabolites in PCYT2 KD cells under normal and starvation states. . ...................................................................................33 Figure 11. Pulse-­‐chase experiments with [14C]-­‐ethanolamine demonstrating degradation of metabolites of the CDP-­‐ethanolamine pathway comparing control and PCYT2 KD cells under NSTV and STV conditions. ..................................36 Figure 12. LC3 expression in 2-­‐month old fasted muscle tissue.........................................37 Figure 13. LC3 protein content in 10-­‐month old fasted control and Pcyt2 KO mice. 38 Figure 14. LC3 expression in muscle tissue of 10-­‐month old fed mice............................39 Figure 15. Quantitative analysis of LC3-­‐II content in control and KO muscle tissue. ...............................................................................................................................................................40 Figure 16. LC3 protein levels in 2-­‐month old fasted mice liver tissue. ...........................41 Figure 17. Protein levels of LC3 from 10-­‐month old fasted liver. . ....................................42 Figure 18. Analysis of LC3 protein levels in 10-­‐month old fed mice. ...............................43 Figure 19. Quantitative analysis of LC3-­‐II formation in liver tissue of 2-­‐month old fasted, 10-­‐month old fasted, and 10-­‐month old fed mice. ..........................................44 Figure 20. Immunoblots displaying LC3 protein expression in adipose tissue. .........45 Figure 21. LC3 protein expression in 10-­‐month old fed control and Pcyt2 KO mice. 46 Figure 22. Overall analysis of LC3-­‐II change in adipose tissue. ..........................................47 Figure 23. The CDP-­‐ethanolamine pathway in Pcyt2 KO mice. ..........................................53 vi Abbreviations AMP, adenosine monophosphate AMPK, AMP-­‐activated protein kinase ATG, autophagy related gene ATP, adenosine triphosphate CDP, cytidine diphosphate CEPT, CDP:DAG choline/ethanolamine phosphotransferase DAG, diacylglycerol DFCP1, double FYVE-­‐containing protein 1 DMEM, Dulbecco’s Modified Eagle Medium EBSS, Earle’s Balanced Salt Solution EK, ethanolamine kinase ER, endoplasmic reticulum ET, CTP: phosphoethanolamine cytidylyltransferase Etn, ethanolamine FBS, fetal bovine serum FIP, focal adhesion kinase family interacting protein FoxO, Forkhead family of transcription factors IM, isolation membrane IR, insulin receptor IRS, insulin receptor substrate 1-­‐2 KD, MCH58 Pcyt2 knockdown cells KO, Pcyt2+/-­‐ knockout mice vii LC3, Microtubule-­‐associated protein 1 light chain 3 LDs, lipid droplets LSC, liquid scintillation counting MAPK, mitogen activated protein kinase MCH58, immortalized human fibroblast cell line MEM, minimum essential medium MOI, material of interest mTOR, mammalian serine/threonine kinase target of rapamycin NSTV, non-­‐starvation P450, cytochrome P450 P-­‐Etn, phosphoethanolamine PBS, phosphate buffered saline PC, phosphatidylcholine Pcyt2, CTP: ethanolaminephosphate cytidyltransferase PDI, protein disulfide isomerase PDK1, 3-­‐phosphoinositide dependent protein kinase-­‐1 PE, phosphatidylethanolamine PEMT, phosphatidylethanolamine N-­‐methyltransferase PI, phosphatidylinositol PI(3)P, phosphatidylinositol-­‐3 phosphate PI3K, class III phosphoinositide 3-­‐kinase PKC, protein kinase C PS, phosphatidylserine viii PSD, phosphatidylserine decarboxylase PSS, phosphatidylserine synthase SAM, S-­‐adenosylmethionine STV, starvation TAG, triacylglycerol TLC, thin-­‐layer chromatography TSC, tuberous sclerosis 1-­‐2 ULK1, Unc 51 like kinase 1 VPS, vacuolar protein sorting WIPI2, WD-­‐repeat protein interacting with phosphoinositide 2 ix 1.0 INTRODUCTION 1.1 Introduction to Autophagy Eukaryotic cells possess the unique ability to degrade and recycle several cytosolic constituents, including damaged or dysfunctional organelles and macromolecules, through a highly conserved process known as autophagy (Tooze and Yoshimori, 2010). Several types of autophagy exist according to the specific mechanism through which cytoplasmic material is transported to the lysosome (Yin et al., 2008), including chaperone-­‐mediated autophagy, microautophagy, and macroautophagy, the latter of which will be the focus of this thesis (Reggiori and Klionsky, 2002; Massey et al., 2004). Chaperone-­‐mediated autophagy is a highly selective form of autophagy that specializes in the sequestration and degradation of a single protein substrate containing a peptide sequence biochemically related to KFERQ (Dice, 1990). Microautophagy involves the sequestration of cytoplasmic material via an invagination of the lysosomal membrane (Kunz et al., 2004). Macroautophagy, hereafter referred to as autophagy, involves the sequestering of cytoplasmic materials into a double-­‐membrane vesicle, known as an autophagosome (Yorimitsu and Klionsky, 2005; Yang and Klionsky, 2010). 1.2 Autophagy in Cell Homeostasis Autophagy is a fundamental process in mammals, occurring at basal levels in practically all cells (Komatsu et al., 2005). However, it may be up-­‐regulated by various factors such as hormonal stimulation, drug treatment, hypoxia, reduced energy (ATP/AMP) ratio and most commonly, by nutrient, serum, or amino acid deprivation 1 (Kabeya et al., 2000). Under periods of stress, cells may ensure survival by up-­‐
regulating autophagy, which degrades cytosolic constituents into products that can be recycled for use in energy generation and other processes (Levine and Klionsky, 2004). Conversely, autophagy can play a destructive role by promoting cellular death under conditions with uncontrolled/excessive autophagy or where apoptosis is inhibited (Kroemer and Jäättelä, 2005; Pattingre et al., 2005). This is referred to as autophagic cell death or Type II programmed cell death (Kroemer and Jäättelä, 2005). However, a great deal of information on this type of cell death still remains to be elucidated. The potential role of autophagy in promoting both cell survival and cell death under certain conditions poses a paradox for its function and suggests that autophagy is a highly complex and dynamic process that warrants further research attention. Not surprisingly autophagy, usually the lack thereof, has been implicated in a number of pathophysiological conditions not limited to several forms of cancer and cardiac pathologies, Type II diabetes, inflammatory bowel disease, infectious disease, and even aging (Terman and Brunk, 2005; Massey and Parkes, 2007; Deretic, 2009; Ravikumar et al., 2010; Yang et al., 2010). Consequently, there is an increasing scientific need to understand this process, as well as to identify the diverse roles of autophagy in mammalian systems. 1.3 Autophagy The process of autophagy is reliant on the ability of a cell to form a fully functional autophagosome. Autophagosome formation has been shown to occur through 4 steps: (1) isolation membrane initiation and expansion; (2) autophagosome 2 formation; (3) autophagosome-­‐lysosome fusion, which involves the fusion of the autophagosome with a lysosome to form a single-­‐membrane, lysosomal-­‐enzyme containing autophagosome; and lastly, (4) degradation of the material of interest and the inner membrane of the autophagosome by lysosomal hydrolases (Figure 1) (Levine and Klionsky, 2004). Figure 1. Stages of Autophagy. Autophagy is a multistep process that includes isolation membrane initiation and elongation around material of interest (MOI) to form a fully functional autophagosome. Once an autophagosome is formed, it fuses with a lysosome to form an autolysosome, which contains hydrolases that degrade the material of interest as well as the inner membrane of the autophagosome. Degraded materials are then released back into the cytosol. 1.4 Autophagy Machinery Extensive research examining the molecular machinery involved in autophagosome formation led to the eventual discovery of autophagy-­‐related (Atg) genes in the yeast Saccharomyces cerevisiae (Klionsky et al., 2003). Presently, 31 Atg 3 genes have been identified and characterized in yeast (Suzuki and Ohsumi, 2007), many of which were shown to possess mammalian homologues. Each Atg protein is suggested to play a distinct role during the various stages of autophagy and are generally classified into five subgroups in both yeast and mammals (Mizushima, 2010). (1) The Unc 51 like kinase 1 (ULK1) protein-­‐kinase complex functions in autophagy induction, (2) Atg9 is necessary for membrane recycling, (3) the class III phosphoinositide 3-­‐kinase (PI3K) complex is involved in vesicle nucleation, and finally (4) the Atg12-­‐Atg5-­‐Atg16L conjugation system and (5) the LC3 (Microtubule-­‐associated protein 1 light chain 3) lipidation system are vital for membrane expansion (Mizushima, 2010). 1.5 Isolation Membrane Initiation Autophagy induction and isolation membrane initiation requires the recruitment of several proteins to the endoplasmic reticulum (ER) including the ULK1 protein-­‐
kinase complex that acts downstream of the mammalian target of rapamycin (mTOR) C1 complex. The ULK1 complex is a multimeric complex consisting of Atg101, Atg13, focal adhesion kinase family interacting protein (FIP200), and ULK1 (Itakura and Mizushima, 2010; Mizushima, 2010). A large class III phosphatidylinositol 3-­‐kinase (PI3K) complex made up of Beclin 1, Atg14, vacuolar protein sorting (Vps)34, and Vps15 (Itakura and Mizushima, 2010) is also recruited where it phosphorylates phosphatidylinositol (PI), an ER membrane lipid, to phosphatidylinositol-­‐3 phosphate (PI(3)P). PI(3)P production causes the translocation of the PI(3)P binding proteins WD-­‐
repeat protein interacting with phosphoinositide 2 (WIPI2) and double FYVE-­‐
containing protein 1 (DFCP1) to the same region, which then promotes recruitment of 4 the Atg5-­‐Atg12-­‐Atg16L complex (Matsushita et al., 2007). Importantly, the omegasome is the site to which the core Atg proteins are recruited (Yang and Klionsky, 2010), signifying the importance of this site for autophagosome biogenesis. The omegasome, so-­‐named because of its Ω-­‐like shape, was shown to exist in close proximity to the ER, and acts as a “platform” for autophagosome biogenesis in mammalian cells under amino acid starvation (Axe et al., 2008). 1.6 Autophagosome Formation and Elongation Autophagosome elongation is contingent on the coordinated actions of several core autophagy machinery proteins involved in two ubiquitin-­‐like conjugation systems: the Atg12-­‐Atg5 and the LC3-­‐phosphatidylethanolamine (LC3-­‐ PE) system (Xie et al., 2008; Yin et al., 2008). In the Atg5-­‐Atg12 system, Atg12 is activated by Atg7, a ubiquitin-­‐
activating enzyme (E1)-­‐like protein, and conjugated to Atg5 via Atg10, a ubiquitin carrier protein (E2)-­‐like protein (Mizushima et al., 1998). Lastly, Atg12-­‐Atg5 forms a multimer complex with Atg16L (forming Atg12-­‐Atg5-­‐Atg16L) (Kuma et al. 2002; Yin et al. 2008), which is localized to the outer portion of the autophagosomal membrane (Hanada et al., 2007). Though crucial for pre-­‐autophagosomal elongation, once a fully functional autophagosome forms, the Atg12-­‐Atg5-­‐Atg16L complex dissociates (Ravikumar et al., 2010). In the LC3-­‐PE system, LC3 is initially synthesized as a precursor (Pro LC3), as it possesses an additional arginine residue at the C terminus that is immediately cleaved by Atg4B, a cysteine protease, to become LC3-­‐I (Tanida et al., 2004). LC3-­‐I is conjugated with Atg7 by a thioester bond, and finally with Atg3, another E2 ubiquitin conjugating 5 enzyme, to form an amide bond with phosphatidylethanolamine (PE) (Kirisako et al., 2000; Tanida et al., 2004), an important phospholipid found in biological membranes (Fullerton et al., 2009). Whereas the unconjugated form of LC3, known as LC3-­‐I resides in the cytosol, the conjugated form, LC3-­‐II, is localized to the autophagosomal membrane, thus making it a very effective and important marker for autophagy (Shibata et al., 2010). 1.7 Lysosome Fusion and Autophagosome Degradation Autophagosomal biogenesis concludes when each end of the growing autophagosomal membrane fuses to form a closed double-­‐membrane vesicle. Once formed, the autophagosomal membrane then fuses with a lysosome, forming an autolysosome, and any contents once contained within the autophagosome are degraded by lysosomal hydrolases and recycled back to the cytosol. Upon completion of autophagosome formation, Atg4 cleaves PE from LC3 attached to the outer membrane and releases it back to the cytosol (Kirisako et al., 2000) 1.8 Autophagy Regulation Despite the fact that autophagy has been shown to be activated by various physiological stimuli our knowledge of the signaling regulation of autophagy is surprisingly modest. One signaling pathway well known for regulating autophagy is the mammalian target of rapamycin (mTOR) pathway. mTOR is a serine/threonine kinase that can be inhibited by rapamycin (Levine and Klionsky, 2004). Two complexes make up the mTOR pathway: the Rapamycin-­‐sensitive mTOR complex 1 (mTORC1), which is 6 an autophagy regulator, and mTOR complex 2 (mTORC2), which does not directly regulate autophagy (Guertin and Sabatini, 2009). mTORC1 activation under favorable growth conditions regulates cell proliferation via increased ribosome biogenesis and protein synthesis, while inhibiting autophagy. Under nutrient rich conditions (insulin and/or excess energy) mTORCI is active and interacts with the ULK1 complex and phosphorylates mAtg13 and ULK1, disrupting ULK1’s kinase activities, preventing it from targeting the membrane (Mizushima, 2010). In contrast, mTORC1 inactivation via starvation has been shown to promote autophagy (Wullschleger et al., 2006) as mTORC1 inactivation dephosphorylates ULK1, making it enzymatically active, leading to the phosphorylation of itself, mAtg13 and FIP200 (Jung et al., 2010) necessary for isolation membrane initiation. In this manner, inactivation of mTORC1 under stressful conditions arrests cell growth, minimizing energy demand to promote cell survival and adaptation. 1.9 Autophagosomal Membrane Sources Considering that autophagy is upregulated during periods of deprivation, and membrane and protein synthesis are reduced in this state, the source of the autophagosomal membrane is a topic of great debate (Juhasz and Neufeld, 2006). Several hypotheses as to the source of the autophagosomal membrane have been proposed; the most highly recognized sources being the ER, the mitochondria, the Golgi apparatus and the plasma membrane (Yamamoto et al., 1990a; Ueno et al., 1991; Araki et al., 1995; Scherz-­‐Shouval and Elazar, 2007). However evidence supporting autophagosome membrane formation from sources other than the ER is limited, and 7 even describes a complementary role for the ER in phospholipid synthesis required for membrane biogenesis (Hailey et al., 2010), which will be described shortly. 1.10 The Endoplasmic Reticulum in Membrane Formation The ER has been suggested to play a critical role in autophagosomal membrane formation via two models: the direct maturation model and the de novo synthesis model. According to the direct maturation model, the autophagosome develops from a ribosome-­‐free region of the rough ER (Dunn, 1990; Furuno et al., 1990). However, IMs and mature autophagosomes appear as nascent membranes, lacking ER markers such as P450 and PDI (Yamamoto et al., 1990a, 1990b; Kovács et al., 2007; Hayashi-­‐Nishino et al., 2009) giving more support for the de novo synthesis model than the direct maturation model as it does not require protein stripping of organelle membranes (Baba et al., 1995; Fengsrud et al., 2000; Kovács et al., 2000). Protein removal from organelle membranes is energetically expensive and would require a form of regulated intracellular transport, which has never been identified in autophagy (Girardi et al., 2011). Further evidence for the de novo synthesis model will be discussed below. The main evidence for the role of the ER in autophagosome formation stems from studies which determined that DFCP1 (double FYVE-­‐containing protein 1), a PI(3)P binding protein, localizes to the ER under starvation conditions to form an omegasome (Axe et al., 2008). Recruitment of the Atg5-­‐Atg12-­‐Atg16L complex and LC3, necessary for autophagosome elongation, can only occur after DFCP1 forms the omegasome (Matsushita et al., 2007; Axe et al., 2008). 8 Additional studies by two independent research groups utilizing electron tomography have shown a physical association between the ER and the IM (Hayashi-­‐
Nishino et al., 2009; Ylä-­‐Anttila et al., 2009). Using electron microscopy, Hayashi-­‐
Nishino et al. (2009) showed that the ER associates with the naked (protein free) IM during early autophagosome formation, forming an ER-­‐IM complex. In fact, they determined that both the outer and inner surfaces of the IM showed connections with the rough ER and that a portion of the ER forms a cradle-­‐like structure encircling the IM (Hayashi-­‐Nishino et al., 2009). These findings were confirmed shortly after by Eskelinen et al. (2009) when they discovered connections between the IM/autophagosomal membrane and nearby ER cisternae. Eskelinen et al. (2009) employed 3D tomography to model isolation membranes and found that isolation/autophagosome membranes actually contain several points of physical attachment with the rough ER. 1.11 Source of Lipids for IM and Autophagosome Formation Not surprisingly, another highly controversial topic in autophagy research involves the source of phospholipids required for IM formation and initiation of membrane elongation to form fully fused autophagosomes. Phosphatidylcholine (PC), phosphatidylethanolamine (PE), and phosphatidylserine (PS) are essential bilayer forming phospholipids in all cells. Evidence for these lipids in cell survival pathways and membrane fusion, points to a functional capacity for these phospholipids outside of their normal structural role (Sessions and Horwitz, 1981; Emoto et al., 1996, 1997). PE and PS are typically found in the inner leaflet of the cell membrane and are more fusogenic while PC is predominately found on the outer membrane and is less fusogenic 9 (Williamson and Schlegel, 1994; Marconescu and Thorpe, 2008). Under certain conditions, PE and PS have been shown to redistribute within the membrane bilayers, such that they were both found on the outer leaflet of the plasma membrane preceding cell-­‐to-­‐cell fusion of myoblasts to form myotubes (Sessions and Horwitz, 1981). Similarly, during cytokinesis, PE was found on the outer leaflet of the plasma membrane and at the cleavage furrow during late telophase (Emoto et al., 2005). These and other findings demonstrate a necessary capacity of the inner membrane phospholipids PE and PS in membrane fusion, indicating a possible role for these phospholipids during the IM fusion stage to form a functional autophagosome during autophagy. 1.12 Focus on Phospholipid Synthesis The bulk synthesis of the phospholipids PE and PC occur de novo through two independent branches of the Kennedy pathway: CDP-­‐ethanolamine and CDP-­‐choline, respectively. In the CDP-­‐ethanolamine pathway, the principal precursor ethanolamine is phosphorylated by ethanolamine kinase (EK) using ATP to form phosphoethanolamine, which is then converted by adding CTP and removing pyrophosphate by CTP:phosphoethanolamine cytidylyltransferase (ET) to form CDP-­‐ethanolamine. The latter conversion is a rate-­‐limiting step in this pathway and is regulated by the enzyme Pcyt2. The final step in the CDP-­‐ethanolamine pathway involves the formation of PE from CDP-­‐ethanolamine and DAG by CDP-­‐ethanolamine:1,2-­‐diacylglycerol ethanolamine phosphotransferase (CEPT) (Figure 2). Additionally, PE can also be synthesized by the decarboxylation of PS through the action of phosphatidylserine decarboxylase (PSD) within the mitochondria (Voelker, 1984, 1997). 10 The Pcyt2 gene is an important regulator of the CDP-­‐ethanolamine (Kennedy) pathway (Bakovic et al., 2007) and is essential to the survival of an organism (Fullerton et al., 2009). Studies of the Pcyt2 gene in mice and humans have determined that this gene is highly conserved in mammals (Poloumienko et al., 2004). The gene is regulated by alternative splicing at exon 7 to produce two mRNA transcripts (Pcyt2α and Pcyt2β) with differing catalytic activities and tissue-­‐specific regulation (Tie and Bakovic, 2007). Analogous to the CDP-­‐ethanolamine pathway, the CDP-­‐choline pathway encompasses a series of reactions involving similar enzymes that eventually produces PC (Figure 2). Additionally, PC can be synthesized by two other pathways: through the phosphatidylethanolamine N-­‐methyltransferase (PEMT) pathway, in which S-­‐
adenosylmethionine (SAM) methylates PE from the CDP-­‐ethanolamine pathway to PC or through PS decarboxylation to form PE that is then methylated by SAM to produce PC (Bakovic et al., 1999, 2003; Golfman et al., 2001). PS cannot be made de novo and is synthesized by exchange reactions catalyzed by phosphatidylserine synthase 2 (PSS2) or PSS1 from preexisting PE or PC, respectively. Both PSS1 and PSS2 are located at the ER (Kanfer, 1980). 11 Figure 2. Pathways involved in phospholipid synthesis. Synthesis of PE and PC occurs through two independent branches of the Kennedy pathway. The CDP-­‐
ethanolamine pathway (left) produces PE from ethanolamine, whereas the CDP-­‐choline pathway (right) produces PC from choline. Both the CDP-­‐ethanolamine and the CDP-­‐
choline pathway occur in the ER. Alternative sources of PE and PC are made from pathways in the mitochondria associated membranes of the ER. Ethanolamine kinase-­‐EK; CTP:ethanolaminephosphate cytidylyltransferase-­‐ET; CDP:diacylglycerol ethanolamine and choline phosphotransferases-­‐ET and CT; choline kinase-­‐CK; CTP:cholinephosphate cytidylyltransferase-­‐CT; cytidine-­‐diphosphoethanolamine-­‐CDP-­‐ethanolamine; CDP-­‐choline-­‐cytidine-­‐
diphosphocholine; N-­‐methyltransferase-­‐PEMT; phosphatidylserine synthase 1-­‐ PSS1; phosphatidylserine synthase 2-­‐PSS2 While the bulk of the Kennedy pathway occurs in the cytoplasm, the final steps of PE and PC synthesis (the addition of DAG to CDP-­‐ethanolamine or CDP-­‐choline) occur in the ER (Figure 2) (Kanfer, 1980; Vance, 1991). Similarly, the final synthesis of 12 phosphatidylinositol (PI) (not shown), a phospholipid required during the autophagosome nucleation phase of autophagosome formation, occurs at the ER (Antonsson, 1997; Gardocki et al., 2005). Additionally studies linking the mitochondria as a source of phospholipids for autophagosome biogenesis identify the ER as the main source of PS required for PE synthesis in mitochondria. The mitochondria is not capable of making PS or PC, both phospholipids need to be transported to the mitochondria (Hailey et al., 2010). More evidence of the role of de novo phospholipid synthesis and the ER in autophagosome formation was demonstrated recently by our lab who showed a parallel increase in synthesis of all membrane forming phospholipids (PE, PC, and PS) during the initial stages of autophagy induction. Furthermore, under the same conditions, PE synthesized from ethanolamine was detected in isolated LC3 fractions, indicating that the CDP-­‐ethanolamine pathway could provide PE for LC3 lipidation during the initiation stages of autophagy (Girardi et al., 2011). This thesis extends those initial studies on the role of the CDP-­‐ethanolamine pathway in autophagy. 1.13 Rationale Autophagy plays an important role in maintaining cellular homeostasis by preserving ATP and other resources under nutrient-­‐limiting conditions, while promoting cell death under other conditions (Kroemer and Jäättelä, 2005). Not surprisingly, autophagy or the inability to activate autophagy has been associated with several pathophysiological conditions and diseases. Thus, greater understanding of the autophagy pathway is paramount to the understanding and prevention of several health conditions, as a greater comprehension will facilitate the development of more direct 13 therapies against certain areas of regulation in the pathway to inhibit or promote autophagy for desired treatment results. Much of the research examining autophagy and autophagosome formation has focused on the genetic basis of this process. For this reason, within the last few decades, a great deal of information identifying particular core Atg proteins that act in autophagosome formation have been identified. However, very little research has attempted to characterize this process from a lipid perspective. Consequently, despite the plethora of research aimed at identifying this process, several fundamental questions regarding membrane formation remain to be answered. What is the origin and source of the autophagosomal membrane? We propose that the lipids responsible for autophagosomal membrane bilayer formation are the phospholipids PE (inner membrane) and PC (outer membrane) synthesized in the ER. This thesis focuses on the role of PE and the effects of reduced PE synthesis on autophagy induction using novel PE-­‐deficient (1) cell (PCYT2 knockdown) and (2) animal (Pcyt2 knockout) models. Furthermore as the Pcyt2 knockout model shows gradual development of metabolic syndrome, it will be used to determine if disturbances in autophagosome formation and LC3 lipidation occur when one of the main phospholipid biosynthetic pathways is impaired. 14 2.0 EXPERIMENTAL PROCEDURES 2.1 Cell Culture MCH58 human fibroblast cells containing the Lentiviral vector (pLKO.1), named pLKO.1 (control) and KD5 (PCYT2 knockdown), were grown in Dulbecco’s Modified Eagle Media (DMEM, high glucose) (Invitrogen) supplemented with 10% fetal bovine serum (FBS) and 2 μg/ml puromycin. Cells were cultured at 37°C in a 5% CO2 humidified atmosphere. 2 x 105 MCH58 human fibroblast cells were plated in each well of a 6-­‐well plate. 30 μL of viral supernatant was added to cells for a final volume of 2 ml containing 8 μg/ml Polybrene. The plates were spun at 805g for 30 min at 32 °C, returned to 37 °C, and 24 h after putative infection, infected cells were selected by adding puromycin to cultures at a final concentration of 2 μg/ml puromycin. PCYT2 knockdown cells were generated by infecting MCH58 cells with Lentiviral vector (Open Biosystems) based shRNA constructs targeting human PCYT2. Control cells were infected with an empty vector. MCH58 fibroblasts containing the empty vector are designated controls, while those expressing the PCYT2 shRNA (KD5) are referred to as PCYT2 KD cells in subsequent analyses. 2.2 Autophagy Induction To induce autophagy in control and PCYT2 knockdown (KD) cells, cells were grown in 60-­‐mm Petri dishes to 85% confluence, washed twice with phosphate buffered saline (PBS) and incubated (incubator conditions as above) with Earle’s Balanced Salt Solution (EBSS) (Sigma) for 0-­‐3 h. EBSS is a medium devoid of amino acids commonly 15 used to activate autophagy. DMEM media (as above) acted as a normal, nutrient-­‐rich condition while EBSS served as the starvation condition for cell culture experiments. 2.3 Cell Viability Cell viability was estimated using the Trypan-­‐Blue exclusion assay. Approximately 2.5 x 106 cells were seeded in 6-­‐well plates and grown to 85% confluency. Cells were grown in DMEM media (as above) and in starvation media (EBSS) for 0, 1, 2, and 3 h. Following trypsinization and centrifugation, cells were resuspended in 1 ml of PBS containing 0.4% Trypan Blue and counted. Cells excluding dye were considered viable, while cells that took up dye were non-­‐viable. Results are expressed as the proportion of viable cells in a well. 2.4 Cell Lysis and Protein Preparation After treatments, control and PCYT2 KD cells were washed twice with PBS and lysed by incubating cells in cold lysis buffer (10 mM Hepes-­‐KOH pH 7.9, 10 mM KCl, 1.5 mM MgCl2, protease inhibitor, and NP-­‐40) on ice for 3 min. Subsequently cells were scraped and centrifuged at 3000 rpm for 5 min at 4°C. Protein concentration was determined using the detergent compatible (DC) protein assay (BioRad). Protein samples were prepared for immunoblotting by diluting with 5X SDS (sodium dodecyl sulphate) reducing buffer (0.5 M Tris-­‐HCl pH 5.8, 25% glycerol, 2% SDS, 0.01% bromophenol blue, and 0.5% 2-­‐mercaptoethanol) and boiling for 5 min. 16 2.5 Animal Treatments Pcyt2 knockout (KO) mice were generated as previously described (Fullerton et al. 2007). All procedures were approved by the University of Guelph’s Animal Care Committee and were in accordance with guidelines by the Canadian Council on Animal Care. Mice were exposed to a 12-­‐h light/12-­‐h dark cycle beginning with light at 7:00 a.m. Male and female mice were fed a standardized diet (Harlan Teklad S-­‐2335) ad libitum and had free access to water. Male and female mice aged 2 months and 10 months were used for experiments. 2-­‐month and 10-­‐month old control and Pcyt2 KO mice were fasted for 12 h to induce autophagy. 2.6 Tissue Homogenization and Protein Preparation Skeletal muscle, liver, and adipose tissue from 2 and 10-­‐month old fasted and 10-­‐
month old fed animals weighing approximately 300 mg each, were homogenized in 350 μl of cold homogenization buffer (1 M Tris pH 7.4, 0.5 M EDTA, 0.1 M NaF, protease inhibitor, and phosphatase inhibitor) and centrifuged at 9000 rpm for 5 minutes at 4˚C. Protein concentration was determined using BCA protein assay (Thermo Scientific). Protein samples were prepared for immunoblotting by diluting with 5X SDS (sodium dodecyl sulphate) reducing buffer (0.5 M Tris-­‐HCl pH 5.8, 25% glycerol, 2% SDS, 0.01% bromophenol blue, and 0.5% 2-­‐mercaptoethanol) and boiling for 5 min. 2.7 Western Blotting Pcyt2 protein knockdown in MCH58 KD cells was confirmed by loading 15 µg of protein from both cells on a 12% sodium dodecyl sulfate polyacrylamide gel (SDS-­‐
17 PAGE). Gels were transferred to a polyvinylidene difluoride (PVDF) membrane for 30 min. Membranes were stained with Ponceau S to ensure equal loading and transfer of proteins. Membranes were blocked with 5% Bovine Serum Albumin (BSA) in Tris buffered saline (TBS) containing 0.5% Tween 20 (TBST) for 1 h at room temperature. Subsequently, membranes were probed with rabbit polyclonal anti-­‐Pcyt2α antibody diluted 1:2000 with 5% BSA in TBST overnight (4 ˚C). Membranes were then washed 5 times for 15 min in TBST and then incubated with goat-­‐anti-­‐rabbit IgG linked to horse-­‐
radish peroxidase (HRP) at a dilution of 1:20,000 for 1 h at room temperature. The chemiluminescent peroxidase substrate assay (Sigma) was used according to manufacturer’s instructions. Bands were visualized following exposure to X-­‐ray film. To examine LC3 expression in MCH58 cells, 10 µg of protein from both control and PCYT2 KD cells was loaded onto a 12% SDS-­‐PAGE gel. After transfer, the membrane was stained with Ponceau S and blocked with 5% milk in TBST for 1 h at room temperature. The membrane was then incubated with LC3 antibody (1:1000) (Novus Biologicals) in BSA for 1 h at room temperature and subsequently washed 4 times for 15 min in TBST. After washing, the membrane was incubated with HRP-­‐conjugated goat-­‐anti-­‐rabbit IgG antibody at a dilution of 1:10,000 for 1 h at room temperature and then washed 4 times for 15 min. The membrane was developed with Chemiluminescent peroxidase substrate assay (Sigma) and bands were visualized following exposure to film. Western blots of LC3 protein in adipose, liver, and muscle tissue from control and Pcyt2 KO mice were performed as described previously for MCH58 cells. 15 µg of protein was loaded onto gels and once proper protein loading was confirmed by 18 Ponceau S staining, membranes were incubated with LC3 antibody (1:1000) (Novus Biologicals) for 1 h at room temperature. Membranes and film were developed as described previously. 2.8 Densitometry and Statistical Analysis Data analyses were performed using two-­‐tailed Student’s t-­‐test and one-­‐way ANOVA. Data were considered statistically significant at a 95% confidence level (p ≤ 0.05). Band densities were measured using ImageJ software and GraphPad Prism 4 was used for all statistical analyses. Band densities were corrected for corresponding backgrounds on each film and equal protein loading was controlled by adjusting for level of Ponceau S staining. 2.9 [14C ]–Ethanolamine Pulse and Pulse-­Chase Labeling For pulse experiments, control and PCYT2 KD MCH58 cells were grown in 60-­‐
mm dishes to 90% confluence and either transferred to fresh DMEM media (as described previously) or EBSS containing [14C]-­‐ethanolamine (1 µCi per dish) and labeled for 1, 2 and 3 h. For pulse-­‐chase experiments, cells were grown to 90% confluence in 60-­‐mm dishes and incubated in DMEM media containing [14C]-­‐
ethanolamine (1 µCi per dish) for 2 h, rinsed twice with PBS and chased in either DMEM or EBSS media containing 250µM cold ethanolamine per dish for 0.5, 1, 2, and 3 h. After each time-­‐point, cells were washed twice with cold PBS and collected by scraping. A small volume was set aside for protein quantification. Lipids were extracted using the Bligh and Dyer method (Bligh and Dyer, 1959). The lower organic phase (chloroform 19 phase) was separated from the upper phase (methanol + water) and each phase was dried under nitrogen gas. After drying, the lower phase containing [14C]-­‐PE was resuspended in chloroform and loaded onto Silica Gel 60 thin layer chromatography (TLC) plates and separated by thin-­‐layer chromatography in a solvent system of chloroform: methanol: acetic acid: water (25:15:4:2). The upper phase consisted of the water-­‐soluble radiolabeled intermediates: [14C]-­‐ethanolamine, [14C]-­‐
phosphoethanolamine, and [14C]-­‐CDP-­‐ethanolamine and was dried under nitrogen gas and resuspended in 100 µl methanol: water (2:1). 30 µl of each water-­‐soluble intermediate was loaded onto G-­‐Silica thin-­‐layer chromatography (TLC) plates and separated by thin-­‐layer chromatography in a solvent system of methanol: 0.5% NaCl: ammonia (50:50:5). TLC plates were sprayed with 0.2% ninhydrin and heated for 10 min. Radiolabeled PE and water-­‐soluble intermediates visualized on plates after heating were scraped and quantified by liquid scintillation counting (LSC). 20 3.0 RESULTS 3.1 Autophagic response to PCYT2 knockdown in MCH58 human fibroblasts 3.1.1. Characterization of PCYT2 deficient cells PCYT2 deficiency in PCYT2 knockdown (KD) cells was confirmed through Western blot of control and KD cells using anti-­‐Pcyt2α antibody (Figure 3A). KD cells displayed a 3.9-­‐fold reduction in Pcyt2 protein as compared to control cells (Figure 3B). Extent of PCYT2 knockdown in KD cells was quantified by qRT-­‐PCR using mRNA extracted from KD cells. KD cells showed robust knockdown (~90%) (data not shown). Cell viability of control and KD cells was estimated by Trypan-­‐blue dye exclusion assay. No significant differences in viability were seen between cell lines (Figure 4). 21 A B Figure 3. Pcyt2 protein expression is significantly reduced in PCYT2 KD cells. Knockdown of PCYT2 in PCYT2 KD cells was confirmed via Western blot using anti-­‐
Pcyt2α antibody (A). PCYT2 KDs were constructed using lentiviral based shRNA constructs targeting human PCYT2. Control cells contained an empty vector. Control (lanes: 1,3,5,7) and knockdowns (lanes: 2,4,6,8). (B) Quantitative differences in Pcyt2α protein between control and knockdowns are shown. Equal loading was confirmed by staining membrane with Ponceau S (not shown) and significance differences were identified at P<0.05 using Student’s T-­‐test. 22 Figure 4. No significant differences in cell viability between control and PCYT2 KD cells. Cell viability was determined using trypan blue exclusion assay after media starvation for 0, 1, 2, and 3 h. All measurements were performed in triplicate and expressed as means ± SEM from two separate experiments. 3.1.2 The effect of PCYT2 knockdown on autophagy induction Induction of autophagy in MCH58 cells was measured by analysis of LC3-­‐II protein content using an antibody that detects both LC3-­‐I and LC3-­‐II proteins. Presently LC3-­‐II, which is LC3 bound to PE, is the principal biomarker for autophagy (Klinosky et al. 2008), suggesting that the presence of this protein may indicate an upregulation in autophagy. LC3-­‐I and LC3-­‐II protein content was measured under basal (0), 1, 2, and 3 h of starvation in control and PCYT2 KD cells (Figure 5A). Corresponding bands were identified for LC3-­‐I and LC3-­‐II at 18 kDa and 16 kDa, respectively. There was an increase in LC3-­‐II protein content in control cells over all starvation times. No trend in LC3-­‐II amount was seen in KD cells. Under basal conditions, PCYT2 KD cells showed only a modest increase in LC3-­‐II, but no further increase under starvation. PCYT2 KD was associated with a 1.3-­‐fold higher amount of LC3-­‐II protein during basal conditions 23 as compared to control cells (Figure 5B). This finding suggests that under basal conditions, higher induction of autophagy occurs in PCYT2 deficient cells compared to control cells. A B Figure 5. Expression of LC3-­I, LC3-­II, and relative LC3-­II/control (Ponceau S) ratio in control and PCYT2 KD cells. (A) LC3-­‐I and LC3-­‐II protein expression was measured under basal (non-­‐starvation) and starvation conditions at 1, 2, and 3 h time points in control and KD cells using LC3 antibody. (B) Quantitative differences in LC3-­‐II expression under different starvation times within and between control and KD cells. 24 Equal loading was confirmed by staining membrane with Ponceau S (not shown) and significance differences determined as P<0.05 from at least two separate experiments using one-­‐way ANOVA. 3.1.3 Pulse [14C]-­ethanolamine radiolabeling of the CDP-­ethanolamine pathway To explore the effect of a decrease in PCYT2 level and cell starvation on the rate of PE synthesized by the CDP-­‐ethanolamine pathway, pulse labeling using [14C]-­‐
ethanolamine was performed. Pulse radiolabeling of control cells with [14C]-­‐
ethanolamine (Etn) resulted in increased intracellular ethanolamine by control cells under starvation (STV) conditions as compared to non-­‐starvation (NSTV) conditions at all time-­‐points (Figure 6a). Phosphoethanolamine (P-­‐Etn), the next metabolite in the pathway, showed a 2.9-­‐fold increase in control cells under STV conditions compared to NSTV conditions during 2 h of pulse (Figure 6b). Analysis of the rate of CDP-­‐
ethanolamine (CDP-­‐Etn) synthesis showed a 2.9-­‐fold increase in CDP-­‐ethanolamine at 2 h under STV conditions compared to NSTV in control cells (Figure 6c). Lastly, under STV conditions, control cells showed a 4-­‐fold increase in PE synthesis compared to NSTV conditions at 2 h (Figure 6d). Overall these data indicate the stimulation of PE synthesis and all intermediate steps of the CDP-­‐ethanolamine pathway in control cells under STV conditions. 25 a b c d Figure 6. Pulse experiments with [14C]-­ethanolamine demonstrating incorporation into intermediates of the CDP-­ethanolamine pathway under NSTV and STV conditions in control cells. Control cells were labeled with 1 µCi [14C]-­‐
ethanolamine per dish and placed in STV or NSTV media for 1 and 2 h. After [14C]-­‐
ethanolamine incubation, the intracellular (a) [14C]-­‐ethanolamine and the synthesis of (b) [14C]-­‐phosphoethanolamine, (c) [14C]-­‐CDP-­‐ethanolamine, and (d) [14C]-­‐PE under NSTV and STV states are shown. All measurements were performed in triplicate and expressed as means ± SEM from two separate experiments. Significant differences between STV and NSTV indicated by * (P<0.05). 26 Under the same conditions as above, PCYT2 KD cells showed reduced intracellular ethanolamine under STV versus NSTV conditions (Figure 7a). In contrast, P-­‐Etn synthesis increased significantly in PCYT2 KD cells under STV state (Figure 7b). At 2h, P-­‐Etn showed a 7-­‐fold increase in synthesis under STV compared to the NSTV state. PCYT2 KD cells showed a 2.5-­‐fold increase in CDP-­‐Etn synthesis under STV conditions versus NSTV during 2 h of pulse (Figure 7c). Finally, although PCYT2 KD cells showed increased PE synthesis, the synthetic rate of PE under STV conditions was not much different than under NSTV conditions (Figure 7d). Overall these data indicate that differently from control cells (Figure 6), PE synthesis was not significantly stimulated in KD cells under STV conditions. 27 a b c d Figure 7. [14C]-­ethanolamine incorporation into intermediates of the CDP-­
ethanolamine pathway under NSTV and STV conditions in PCYT2 KD cells. Intracellular (a) [14C]-­‐ethanolamine and the synthesis of (b) [14C]-­‐
phosphoethanolamine, (f) [14C]-­‐CDP-­‐ethanolamine, and (h) [14C]-­‐
phosphatidylethanolamine under NSTV and STV expressed as means ± SEM from two separate experiments. Significant differences between STV and NSTV indicated by * (P<0.05). 3.1.4 Comparing pulse [14C]-­ethanolamine radiolabeling of the CDP-­ethanolamine pathway between control and PCYT2 KD cells Comparison of pulse labeling data illustrating intracellular [14C]-­‐ethanolamine between control and PCYT2 KD cells showed that during a 2 h pulse, intracellular ethanolamine was 5.9-­‐fold greater in KD cells compared to controls under NSTV 28 conditions (Figure 8a). Under STV conditions, KD cells again showed a greater increase in intracellular ethanolamine compared to control cells at 2 h (Figure 8b). No noticeable affect on the rate of P-­‐Etn synthesis between control and PCYT2 KD cells under NSTV conditions was seen; however, total labeling of P-­‐Etn was greater in control cells than KD cells at all time-­‐points (Figure 8c). Under STV conditions, control cells showed a higher rate of P-­‐Etn synthesis at all time points compared to KD cells (Figure 8d). As expected, the overall rate of CDP-­‐Etn synthesis was significantly reduced in KD cells (PCYT2 activity was <10%), and total labeling of CDP-­‐Etn was greater in control cells reflecting higher PCYT2 activity (Figure 8e). After 1 h pulse, CDP-­‐Etn was degraded in both cell types. Analysis of CDP-­‐Etn synthesis rates under STV conditions showed greater reduction in synthesis rates in KD than control cells (Figure 8f). Finally, the rate of PE synthesis in PCYT2 KD cells was much slower during all time-­‐points under NSTV conditions compared to controls (Figure 8g). Under STV conditions, PE synthesis was elevated in control cells versus KD cells (Figure 8h) as expected. With the exception of [14C]-­‐ethanolamine, control cells showed higher rates of synthesis and higher radiolabeling content of all metabolites of the CDP-­‐ethanolamine pathway than KD cells under both media conditions. 29 a b c d e f g h 30 Figure 8. [14C]-­ethanolamine incorporation into metabolites of the CDP-­
ethanolamine pathway comparing control and PCYT2 KD cells under NSTV and STV conditions. The intracellular [14C]-­‐ethanolamine in control and PCYT2 KD cells under (a) NSTV and (b) STV states are shown. Levels of [14C]-­‐phosphoethanolamine in control and KD cells under NSTV and STV states are shown in (c) and (d) respectively. Synthesis of [14C]-­‐CDP-­‐ethanolamine in control and PCYT2 KD cells under NSTV (e) and STV (f) conditions are shown. Levels of [14C]-­‐PE in control and KD cells under NSTV and STV states are shown in (g) and (h) respectively. All measurements were performed in triplicate and expressed as means ± SEM from two separate experiments. Significant differences between KD and control cells indicated by * (P<0.05) and ** (P<0.005). 3.1.5 Pulse-­Chase [14C]-­ethanolamine radiolabeling of the CDP-­ethanolamine pathway Pulse-­‐chase experiments with [14C]-­‐ethanolamine showed faster degradation of radiolabeled ethanolamine under STV conditions compared to NSTV in control cells (Figure 9a). When the next metabolite was measured, no change in the rate of P-­‐Etn degradation was seen between conditions in control cells (Figure 9b). However there was a significant difference in P-­‐Etn labeling between conditions during most time points. P-­‐Etn content at 0.5 h, 2 h, and 3 h STV was 2.4, 1.9, and 1.9-­‐fold higher than that at the same time points under NSTV conditions, respectively. Labeling in control cells showed increased degradation of CDP-­‐Etn during the second hour followed by CDP-­‐Etn synthesis at 3 h under STV state (Figure 9c). The rate of CDP-­‐Etn degradation did not change in control cells under NSTV conditions. Similar to chase data analyzing CDP-­‐Etn in control cells, PE showed a biphasic effect, where PE showed a faster rate of degradation during the first hour of STV and subsequently showed increased synthesis of PE after the first hour (Figure 9d). This data is consistent with data collected from pulse labeling experiments using [14C]-­‐ethanolamine in control cells. Overall these data demonstrate increased degradation of CDP-­‐ethanolamine pathway metabolites under 31 STV conditions, with CDP-­‐Etn and PE (the last steps of the pathway) showing re-­‐
synthesis under the final hours of chase. a b c d Figure 9. Pulse-­chase experiments in control cells using [14C]-­ethanolamine to demonstrate the degradation of metabolites of the CDP-­ethanolamine pathway under normal and starvation conditions. Cells were pulsed with 1 µCi [14C]-­‐
ethanolamine per dish for 2 hours, washed, and then chased with cold ethanolamine in STV or NSTV media for 0.5, 1, 2, and 3 h time points. After pulsing and chasing, the degradation of (a) [14C]-­‐ethanolamine, (b) [14C]-­‐phosphoethanolamine, (c) [14C]-­‐CDP-­‐
ethanolamine, and (d) [14C]-­‐PE under NSTV and STV states in control cells are shown. Radioactivities were determined (n= 3 performed in duplicate) and are expressed as means ± SEM. Significant differences between STV and NSTV conditions indicated by * (P<0.05) and ** (P<0.005). 32 Under STV, PCYT2 KD cells showed faster degradation of radiolabeled Etn compared to NSTV conditions at all time-­‐points (Figure 10a). Similarly, KD cells demonstrated faster degradation of P-­‐Etn (Figure 10b) and CDP-­‐Etn (Figure 10c) in STV compared to NSTV culture conditions. PCYT2 KD cells did not show any significant change in the rate of degradation of PE under differing media conditions (Figure 10d). a b c d Figure 10. Degradation of CDP-­ethanolamine pathway metabolites in PCYT2 KD cells under normal and starvation states. Degradation of (e) [14C]-­‐ethanolamine, (f) [14C]-­‐phosphoethanolamine, (g) [14C]-­‐CDP-­‐ethanolamine, and (h) [14C]-­‐PE under NSTV and STV states in KD cells are shown. Radioactivities were determined (n= 3 performed in duplicate) and are expressed as means ± SEM. Significant differences between STV and NSTV conditions indicated by * (P<0.05). 33 3.1.6 Comparing pulse-­chase [14C]-­ethanolamine radiolabeling of the CDP-­ethanolamine pathway between control and PCYT2 KD cells Pulse-­‐chase experiments showed faster degradation of Etn in PCYT2 KD cells compared to control cells under NSTV conditions (Figure 11a). This trend was similarly seen under STV conditions at all time-­‐points (Figure 11b). No change in P-­‐Etn degradation rate was seen between cell types under NSTV conditions; however, total labeling of P-­‐Etn was much greater in KD cells at all time-­‐points (Figure 11c) showing that P-­‐Etn (substrate of Pcyt2) did not degrade into CDP-­‐Etn, as expected for KD cells. Under STV conditions, a faster rate of P-­‐Etn degradation was seen in KD cells compared to control cells (Figure 11d) indicating the higher utilization of P-­‐Etn in KD cells under STV conditions. Similarly, no change in the rate of CDP-­‐Etn degradation was seen between cell types under NSTV conditions (Figure 11e). Under STV, PCYT2 KD cells showed greater degradation during 0.5 and 1 h compared to control cells (Figure 11f), which suggests that the last step regulated by ethanolamine phosphotransferase may also be upregulated in KD cells, like the other steps in the pathway. Under NSTV conditions, control and PCYT2 KD cells showed similar rates of PE synthesis during hours 1 to 3 (Figure 11g). Under STV conditions, control cells showed a faster rate of PE degradation than KD cells during the first hour; however, control cells showed faster rates of re-­‐synthesis than KD cells (Figure 11h), altogether showing much faster PE turnover in control cells than in KD cells. 34 a b c d e f g h 35 Figure 11. Pulse-­chase experiments with [14C]-­ethanolamine demonstrating degradation of metabolites of the CDP-­ethanolamine pathway comparing control and PCYT2 KD cells under NSTV and STV conditions. [14C]-­‐ethanolamine degradation in control and PCYT2 KD cells under (a) NSTV and (b) STV are shown. Degradation of [14C]-­‐phosphoethanolamine in control and KD cells under NSTV and STV states are shown in (c) and (d) respectively. The degradation of [14C]-­‐CDP-­‐Etn in control and PCYT2 knockdown cells under NSTV and STV are shown in (e) and (f) respectively. Degradation of [14C]-­‐PE in control and KD cells under NSTV and STV states are shown in (g) and (h) respectively. Radioactivities were determined (n= 3 performed in duplicate) and are expressed as means ± SEM. Significant differences between KD and control cells indicated by * (P<0.05) and ** (P<0.005). 36 3.2 Autophagic response to Pcyt2 deficiency in mouse tissue 3.2.1 The effects of Pcyt2 knockout on muscle LC3-­II protein content LC3-­‐I and LC3-­‐II protein content from muscle isolated from fasted 2-­‐month old control and Pcyt2 knockout (KO) mice was examined by immunoblotting (Figure 12A). Band density of LC3-­‐II in muscle tissue between controls and KOs was measured and expressed relative to a loading control (Ponceau S). Pcyt2 KO did not affect LC3-­‐II protein amount in 2-­‐month old fasted mice compared to control mice (Figure 12B). A B Figure 12. LC3 expression in 2-­month old fasted muscle tissue. (A) Representative blot of LC3-­‐I and LC3-­‐II proteins in control and Pcyt2 KO mice. Each set of (LC3-­‐I and LC3-­‐II) bands represents muscle tissue from a separate animal (n=3). (B) Pcyt2 KO does not affect LC3-­‐II expression in 2-­‐month old fasted muscle tissue. 37 Western blot analysis of LC3-­‐I and LC3-­‐II protein content from muscle from 10-­‐
month old fasted mice showed noticeable conversion of LC3-­‐I to LC3-­‐II in control mice (Figure 13A). Pcyt2 KO was associated with a 2.2-­‐fold reduction in LC3-­‐II protein content compared to control mice (Figure 13B). A B Figure 13. LC3 protein content in 10-­month old fasted control and Pcyt2 KO mice. (A) Representative blot of LC3-­‐I to LC3-­‐II conversion in mice. Each set of (LC3-­‐I and LC3-­‐II) bands represents muscle tissue from a separate animal (n=3). (B) Intensity of LC3-­‐II bands was reduced in Pcyt2 KO mice compared to control mice. Data shown represent mean ± SEM from two separate experiments. ** indicates significance (P<0.005). 38 Muscle tissue from 10-­‐month old fed mice showed expression of both LC3-­‐I and LC3-­‐II (Figure 14A). KO mice showed a 1.6-­‐fold greater band density of LC3-­‐II protein compared to control mice (Figure 14B). A B Figure 14. LC3 expression in muscle tissue of 10-­month old fed mice. (A) Immunoblot showing LC3-­‐I and LC3-­‐II protein content from control and Pcyt2 KO mice. Each set of (LC3-­‐I and LC3-­‐II) bands represents muscle tissue from a separate animal. (B) Pcyt2 KO is associated with higher expression of LC3-­‐II compared to control mice. * indicates significance (P<0.05). 39 LC3-­‐II expression in 2-­‐month old fasted, 10-­‐month old fasted, and 10-­‐month old fed control and KO mice were compared. 10-­‐month old fasted Pcyt2 KO mice displayed a 3.0-­‐fold reduction in LC3-­‐II protein level as compared to 2-­‐month old fasted KO mice (Figure 15). LC3-­‐II protein amount in 10-­‐month old fed KO mice was 2.5-­‐fold higher than 10-­‐month old fasted KO mice. The amount of LC3-­‐II was not significantly affected by age or metabolic state in control animals. Figure 15. Quantitative analysis of LC3-­II content in control and KO muscle tissue. The plot displays the difference in LC3-­‐II protein amount between each group in control and Pcyt2 KO mice. ** indicates significance (P<0.005). 40 3.2.2 The effects of Pcyt2 knockout on liver LC3-­II protein content Western blot analysis showed expression of LC3-­‐I and LC3-­‐II protein in both 2-­‐
month old fasted controls and Pcyt2 KOs (Figure 16A). Pcyt2 KO did not affect LC3-­‐II expression compared to controls (Figure 16B). A B Figure 16. LC3 protein levels in 2-­month old fasted mice liver tissue. (A) Immunoblot displaying LC3-­‐I and LC3-­‐II expression in control and Pcyt2 KO mice. (B) Pcyt2 KO does not affect LC3-­‐II expression. 41 Figure 17A shows LC3-­‐II protein levels in fasted 10-­‐month old control and KO livers. Control livers had almost no LC3-­‐II present; however, KO liver showed distinct LC3-­‐II bands. Quantitative analyses of LC3-­‐II protein amount in this tissue showed a significant difference between controls and KOs (Figure 17B). Fold change in the amount of LC3-­‐II protein between control and KO mice was not represented in these data (Figure 17B), as the amount of LC3-­‐II protein detected in littermate control mice was negligible. A B Figure 17. Protein levels of LC3 from 10-­month old fasted liver. (A) LC3 immunoblot displaying expression of forms of LC3 in control and Pcyt2 KO mice. (B) Pcyt2 KO is associated with higher expression of LC3-­‐II compared to control mice. 42 When liver LC3-­‐II content was examined from 10-­‐month old fed mice (Figure 18A), levels from Pcyt2 KO liver tissue were significantly greater than controls (Figure 18B). Similar to Figure 17B, fold change in the amount of LC3-­‐II protein between control and KO mice was not represented (Figure 18B), as the amount of LC3-­‐II protein detected in littermate control mice was negligible. A B Figure 18. Analysis of LC3 protein levels in 10-­month old fed mice. (A) LC3 expression in liver tissue of control and KO mice. (B) Pcyt2 KOs display significantly higher expression of LC3-­‐II in 10-­‐months fed mice as compared to control mice. LC3-­‐II amount in 2-­‐month old fasted, 10-­‐month old fasted, and 10-­‐month old fed control and KO mice were compared (Figure 19). Metabolic state did not affect LC3-­‐II 43 protein levels in control animals, however there was an elevated level of LC3-­‐II in 10-­‐
month old KO liver in the fed compared to the fasted state. Age affected liver LC3-­‐II protein amounts in control mice but did not affect amounts in KO livers. Figure 19. Quantitative analysis of LC3-­II formation in liver tissue of 2-­month old fasted, 10-­month old fasted, and 10-­month old fed mice. The plot displays the difference in LC3-­‐II expression between each group in control and Pcyt2 KO mice. * indicates significance (P<0.05). 3.3.3 The effects of Pcyt2 knockout on adipose LC3-­II content LC3 immunoblots of adipose tissue from 2-­‐month old (Figure 20A) and 10-­‐
month old (Figure 20B) mice in the fasted state only showed intense bands for LC3-­‐I. LC3-­‐II protein expression was barely detectable in these tissues. 44 A B Figure 20. Immunoblots displaying LC3 protein expression in adipose tissue. LC3-­‐
II protein is barely detectable in adipose tissue of (A) 2-­‐month old fasted and (B) 10-­‐
month old fasted control and Pcyt2 KO mice. On the other hand, adipose tissue from 10-­‐month old fed mice shows distinct presence of LC3-­‐II (Figure 21A). A significant difference in LC3-­‐II protein level was seen between control and Pcyt2 KO mice where KO mice showed a 2.2-­‐fold higher level of LC3-­‐II protein than littermate controls (Figure 21B). 45 A B Figure 21. LC3 protein expression in adipose tissue of 10-­month old fed control and Pcyt2 KO mice. (A) LC3 immunoblot showing some conversion of LC3-­‐I to LC3-­‐II. (B) Pcyt2 KOs display significantly higher levels of LC3-­‐II protein compared to controls. * indicates significance (P<0.05). LC3-­‐II protein abundance in adipose tissue from 2-­‐month old and 10-­‐month old fasted controls shows barely detectable levels. In the fed state, the amount of LC3-­‐II is much higher in KO and control adipose tissues relative to the fasted 10-­‐month old state (Figure 22). 46 Figure 22. Overall analysis of LC3-­II change in adipose tissue. The plot displays the difference in LC3-­‐II expression between each group in control and Pcyt2 KO mice, showing low levels in control and KO tissues under a fasted state, with a dramatic increase in LC3-­‐II amount in the feeding state. * indicates significance (P<0.05). 47 4.0 DISCUSSION 4.1 Autophagic response to PCYT2 deficiency in human MCH58 fibroblast cells Despite growing interest in autophagy research over the last few decades, few reports have attempted to study this process using a metabolic approach. Recent evidence from our lab indicates that phospholipid synthetic pathways play an important role in autophagy by providing 3 main phospholipids for autophagosome bilayer (membrane) formation. Both [14C]-­‐Etn and [3H]-­‐Glycerol radiolabeling experiments established that ER synthesis of all major phospholipids (PE, PC, and PS) was significantly increased under starvation conditions that typically activate autophagy. In addition, immunoprecipitation experiments using anti-­‐LC3 antibody and [14C]-­‐Etn and/or [3H]-­‐Glycerol labeled cells demonstrated the incorporation of newly made PE in LC3 protein fractions, suggesting that newly synthesized PE from the CDP-­‐ethanolamine pathway is utilized for LC3 lipidation during autophagy (Girardi et al., 2011). To further investigate the role of PE and the CDP-­‐ethanolamine pathway in autophagy, we used stable cell lines of human fibroblasts in which the CDP-­‐ethanolamine pathway was almost completely inhibited. In this model, the PCYT2 gene was knocked down by shRNA interference. Knockdown in this model severely reduces the formation of CDP-­‐
ethanolamine; and therefore, limits the rate of PE synthesized in the Kennedy pathway. The PCYT2-­‐deficient cell model is novel and offers a useful approach to determining a role for PCYT2 and PE synthesis in autophagy. 48 4.1.1 Autophagy is not properly induced in PCYT2 KD cells When PCYT2-­‐containing control cells were starved of serum and amino acids they displayed a clear trend in LC3-­‐II protein amount, with content steadily increasing with successive starvation times (0-­‐3 h) (Figure 5). As expected, this finding showed that autophagy in control cells is stimulated by longer starvation. In contrast, serum and amino acid starvation of PCYT2 KD cells did not result in stimulated autophagy over time, suggesting that autophagy does not respond similarly to media starvation in KD cells as in control cells. PCYT2 KD cells showed a blunted response to media starvation. KD cells also showed significantly higher content of LC3-­‐II protein, reflecting a higher amount of autophagic activity in these cells during initial basal conditions as compared to control cells. This finding also suggests that despite dysfunction in the CDP-­‐
ethanolamine pathway, there remains to be a sufficient amount of PE made available for basal levels of autophagy to occur. Based on this result, total PE content of control and KD cells under basal conditions should likely yield minimal differences in total quantity of PE available between the two cell genotypes, likely as a result of the importance of maintaining phospholipid balance to preserve membrane integrity for cell survival. This is consistent with data collected previously in our lab that examined the levels of all phospholipids (PE, PC, and PS) in a Pcyt2 animal KO model and found that they did not differ between Pcyt2 KO and littermate control animals (Fullerton et al., 2007). Minimal differences in PE content between cell genotypes despite PCYT2 knockdown would suggest two possibilities: (1) that the remaining activity of PCYT2 in KD cells is sufficient for PE production in the CDP-­‐ethanolamine pathway or (2) that a 49 compensatory process for PE production by PS decarboxylation is activated. A possible role for PS decarboxylation in maintaining PE production in PCYT2 KD cells should be investigated in future by [3H]-­‐serine labeling experiments. Labeling experiments in Pcyt2 KO animals established that the maintenance of PE content in Pcyt2 deficient animals did not occur via compensatory increases in PS decarboxylation but via an upregulation in the remaining Pcyt2 allele (Fullerton et al., 2007). Since the level of Pcyt2 activity differs between PCYT2 knockdown cells (~10% activity remaining) and Pcyt2 knockout mice (~65-­‐80% activity remaining), it would be of interest to determine if compensatory pathways are activated as a direct consequence of the magnitude of knockdown/knockout. In fact, a study by Leonardi et al. (2009) demonstrated that total knockout of liver Pcyt2 resulted in an upregulation in the PS decarboxylation pathway to make PE, and this is likely the case occurring in PCYT2 KD cells. Cell viability in control and PCYT2 KD cells was determined by measuring the number of live cells using a Trypan blue exclusion assay (Figure 4). No significant differences in cell viability were seen between basal conditions and longer starvation duration in or between control and KD cells. However, this finding is curious since autophagic activity in control cells increased with increasing duration of starvation (Figure 5), supporting that under starvation conditions, autophagy is induced for survival in these cells. Contrastingly, since no relationship between autophagy induction and starvation duration was seen in PCYT2 KD cells, this suggests that the survival of these cells is regulated by activation of pathways other than autophagy, perhaps by proteasomal or lysosomal pathways, or that LC3 turnover in PCYT2 KD cells is much higher than in control cells. For this reason, examining proteasomal and lysosomal 50 pathways and measuring LC3 turnover in KD cells should be an objective of future studies. 4.1.2 PE synthesis in PCYT2 KD cells does not respond normally to starvation When control cells were starved of amino acids and serum, an upregulation in the rate of [14C]-­‐PE synthesis and intermediates of the CDP-­‐ethanolamine pathway was observed (Figure 6). Extrapolating from data presented in Figure 5, higher levels of autophagic activity as measured by greater expression of LC3-­‐II ratio under increasing duration of starvation, correspond with an upregulation of the CDP-­‐ethanolamine pathway under the same conditions, suggesting that the biosynthesis of PE is stimulated under the induction of autophagy. This data is consistent with findings previously reported by our lab using mouse hepatocytes (Girardi et al., 2011). However when PCYT2 KD cells were radiolabeled they showed a poor PE synthetic response to starvation (Figure 7). Very little difference in the rate of PE synthesis was observed between the two conditions. When comparing PE synthesis between the two cell types, PCYT2 KD cells responded with reduced PE synthesis in all conditions as was expected since PCYT2 was deleted (Figure 8). Pulse-­‐chase [14C]-­‐ethanolamine radiolabeling in control cells showed a higher turnover of [14C]-­‐PE and its intermediates under starvation state (Figure 9). Under the same conditions, KD cells showed that starvation conditions resulted in faster rates of degradation in pathway intermediates, but very little effect on [14C]-­‐PE degradation under differing media conditions (Figure 10). Comparing [14C]-­‐PE degradation between cell types, there was significantly reduced turnover in PCYT2 KD cells (Figure 11). 51 Taken together, this data indicates that knockdown of PCYT2 influences cell response to starvation and autophagy induction in these cells. 4.2 Autophagic response in Pcyt2 heterozygous mouse tissues Given the primary function of autophagy in maintaining cell homeostasis and survival, it is not surprising that autophagy has been associated with a number of pathological conditions, of which the list is only expected to grow in future. Numerous studies have attempted to characterize the precise role of autophagy in disease onset and progression, yet due to the complex nature of diseases and the lack of current understanding of the basic biology of autophagy, further studies are still needed. For this reason, research focused on understanding the basic biology of this process is fundamental to the understanding of disease pathogenesis and the development of possible therapeutic targets. Data in Figure 5 showed that no trend between autophagy induction and starvation duration, known to trigger autophagy, exists when there was a reduction in PE formation by the CDP-­‐ethanolamine pathway. In contrast to this, there was a clear trend in autophagy induction under increasing starvation duration in control cells, suggesting that impairments in PE formation do affect autophagy. To complement those in vitro cell studies, we next examined the process of autophagy in a Pcyt2 KO animal model previously shown to develop a metabolic disease phenotype at a later age (5-­‐6 months). Dysregulation of autophagy has been implicated in a variety of disease conditions including those that define metabolic syndrome (Singh and Cuervo, 2012). 52 Complete deletion of the Pcyt2 gene in mice results in early embryonic lethality, while partial deletion of the gene produces heterozygous animals that are phenotypically indistinguishable from wildtype littermate controls. When Pcyt2 activity is reduced in mice, it results in decreased flux through the CDP-­‐ethanolamine pathway, which reduces CDP-­‐Etn levels and PE synthesis, causing an accumulation of unused DAG, which elicits an increase in fatty acid synthesis and DAG-­‐mediated TG formation (Figure 23). These mice consequently develop obesity, hypertriglyceridemia, liver steatosis, and whole body-­‐insulin resistance. Young Pcyt2 KO mice do not appear to be phenotypically different from littermate controls, however as they age they gradually gain more weight and develop a metabolic syndrome phenotype (Fullerton et al., 2009). Figure 23. The CDP-­ethanolamine pathway in Pcyt2 KO mice. Pcyt2 gene disruption reduces flux (CDP-­‐ethanolamine and PE content) through the remainder of the CDP-­‐
ethanolamine pathway. Whereas, DAG and TG accumulate. 53 As complete disruption of Pcyt2 in KO mice results in animal death, Pcyt2 KO mice only experience a partial knockout of the gene allowing them to retain about 65-­‐
80% activity (Fullerton et al., 2007), while KD cells assume only about 10% activity. Given that the magnitude of gene Pcyt2 activity is different between KO mice and KD cells, Pcyt2 deletion in mice presents an interesting comparative difference than PCYT2 deletion in mammalian cells, which can be used to examine whether the extent of gene activity, and therefore activity of the pathway impacts autophagy induction. Furthermore, since older KO animals are phenotypically obese, are insulin resistant, and have elevated levels of DAG and TG, they are an exemplary model for metabolic syndrome that may be used to determine if autophagy is impaired under conditions where metabolic syndrome is present. 4.2.1 The effects of Pcyt2 deficiency on autophagy induction in skeletal muscle Muscle mass constitutes 40-­‐50% of the body and is considered one of the major tissues involved in controlling the metabolic state of an organism. Skeletal muscle is responsible for body movement and movement/exercise results in damage to muscle organelles and proteins, as well as the production of reactive oxygen species (ROS) from mitochondria (Sandri, 2010). For these reasons, muscle cells demand a highly efficient process for eradicating damaged and defective organelles and toxic proteins. Autophagy necessitates this role (Levine and Kroemer, 2008). It is not surprising then that autophagy induction was shown to be typically high within muscle tissue of littermate control mice between the different groups tested, and no significant difference in induction was seen between age groups or metabolic states in these mice (Figure 15). 54 On the contrary, induction of autophagy was influenced by both age and metabolic state in Pcyt2 KO mice. 10-­‐month old fasted KO mice displayed reduced autophagy induction as compared to 2-­‐month old fasted KO mice. Similarly, 10-­‐month old fasted KO mice also showed a reduced induction compared to 10-­‐month old fed KO mice. When comparing autophagy induction between KO mice and littermate controls we saw that there were no differences in LC3-­‐II protein content between muscle tissue of 2-­‐month old fasted Pcyt2 KO and littermate control mice (Figure 12), but Pcyt2 KO muscle of 10-­‐
month old fasted mice displayed a significant reduction in amount of LC3-­‐II protein compared to littermate controls (Figure 13). Muscle tissue of 10-­‐month old fed mice showed greater LC3-­‐II protein content in KO mice compared to controls (Figure 14). These data suggest that Pcyt2 deletion dramatically inhibits autophagy induction in 10-­‐
month old fasted mice and upregulates autophagy in 10-­‐month old fed mouse muscle. 4.2.2 The effects of Pcyt2 deficiency on autophagy induction in liver Similar to muscle, the liver is another essential organ involved in the control of metabolism. The liver is responsive to several stimuli not exclusive to hormones, particularly insulin, and nutrient status (glucose, fatty acids, amino acids). The liver also functions as a storage site for stored nutrients and will respond to excess glucose and insulin stimuli by suppressing gluconeogenesis and increasing fatty acid and TG synthesis from glucose (Yecies et al., 2011). As the liver is largely responsible for energy storage in the body, it not unexpected that autophagy induction in this tissue is typically low (Figure 19) as compared to that of skeletal muscle, which is metabolically very active. As seen in Figure 19, autophagy induction in liver tissue of control mice is normally low but is inducible by age as 2-­‐month old mice showed higher induction than 55 10-­‐month old fasted control mice. Metabolic state did not affect induction of autophagy in control mice. By comparison, induction of autophagy in KO mice is typically higher than in controls and was inducible by feeding. No differences in induction were seen between age groups in KO mice. There were no differences in LC3-­‐II protein expression between liver tissue of 2-­‐month old fasted littermate control and KO mice (Figure 16), thereby indicating that Pcyt2 deletion had no effect on induction of autophagy in these mice. In contrast, KO of Pcyt2 in 10-­‐month old fasted mice (Figure 17) and in 10-­‐month old fed mice (Figure 18) resulted in a significant increase in LC3-­‐II protein content, suggesting an effect of Pcyt2 deletion on autophagy induction. 4.2.3 The effects of Pcyt2 deficiency on autophagy induction in adipose Adipose tissue is the primary site for fat storage in mammals. Similar to liver, this tissue displayed minimal induction of autophagy (Figure 22) and opposite to muscle tissue, reflecting low demand for energy (low metabolic activity) in adipose tissue. LC3 immunoblotting of adipose tissue under fasting conditions did not identify distinct LC3-­‐II bands in either Pcyt2 KO or littermate control mice in either age group (Figure 20). This suggests that induction of autophagy in these tissues is low under fasting conditions, but is inducible under feeding conditions in both littermate controls and Pcyt2 KO mice (Figure 21). Pcyt2 KO mice showed significantly higher levels of LC3-­‐
II protein amount as compared to littermate control mice, which suggests that Pcyt2 deletion does affect autophagy induction under fed states in adipose tissue. There was no effect on autophagy seen between differing age groups in either KO or littermate control mice. 56 Taken together, the data presented demonstrates that autophagy induction in vivo is altered under conditions where PE synthesized by the CDP-­‐ethanolamine pathway is impaired. In addition, autophagy was affected by animal age, metabolic state and specificity of tissue. These differences may be explained by the mechanisms that link PE synthesis with autophagy function and the various factors that modify them during metabolic disease development. 4.3 Regulation of autophagy in mammalian tissues Recent investigation into the signaling pathways regulating mammalian autophagy has yielded some exciting insights in the field; however, analogous to research examining autophagosome origin and formation, details about how autophagy is regulated with phospholipid metabolic pathways still remains to be elucidated. Few of the better-­‐known pathways likely involved in regulating autophagy induction in the various tissues include the PI3K/Akt and AMPK pathways, which primarily centralize around the actions of mTORC1 kinase. Additionally, autophagy may be regulated by a transcriptional mechanism such as by FoxO3, or via lipid metabolism pathways such as PKC/MAPK and/or through lipophagy. One of the greater studied pathways under which autophagy is regulated is the PI3K/Akt pathway, which is under the influence of insulin. When insulin is present, such as in a fed state, it binds to its membrane receptor initiating the activation of insulin-­‐
receptor (IR) tyrosine kinase, which phosphorylates the IRS proteins, IRS1 and IRS2, at tyrosine residues (Withers and White, 2000). Subsequently IRS proteins activate PI3K, which catalyze the formation of PIP3, recruiting Akt to the plasma membrane (Cantley, 57 2002). Akt is phosphorylated and inhibits TSC2 of the TSC1/TSC2 heterodimer complex that negatively regulates mTORC1. When this occurs, mTORC1 is activated leading to cell growth, protein synthesis and autophagy inhibition (Huang and Manning, 2009). However during insulin resistance, such as in the older Pcyt2 KO mice, the IRS/PI3K/Akt pathway is blunted. Since insulin signaling is not functional in the presence of insulin resistance, it has been conjectured that mTORC1 is dysregulated and autophagy should increase under conditions of insulin resistance. Surprisingly, the few studies examining the relationship between insulin resistance and autophagy have negated this hypothesis. For instance, one study identified a decrease in autophagy induction in insulin resistant livers of mice exposed to a high fat diet (Liu et al., 2009). The discrepancy between the results presented in this thesis and the study by Liu et al. (2009) might reflect a critical difference between animal models presenting with insulin resistance. Liu et al. (2009) used a high fat diet to induce insulin resistance in their mouse model, whereas insulin resistance in our model is a product of partial gene deletion resulting in elevated levels of DAG and TG and other disruptions to lipid metabolism (Fullerton et al., 2009). Some recent studies have demonstrated the induction of autophagy under transcriptional regulation through the FoxO family of transcription factors (Zhao et al., 2007; Mammucari et al., 2008). The PI3K/Akt pathway negatively regulates the activity of FoxO, which is inhibited by insulin and other growth factors (Salih and Brunet, 2008). Under fasting conditions, FoxO3 causes the induction of autophagy by directly controlling the transcription of several autophagy genes including LC3, Gabarap, Atg12, and Bnip3 (Zhao et al., 2007; Mammucari et al., 2008). Transcriptional activation of 58 autophagy by FoxO3 has been chiefly studied in muscle; however, it may be an important regulator of autophagy in other tissues in addition to muscle. Based on highly dysregulated LC3-­‐II in the Pcyt2 KO model and impaired insulin signaling, it seems plausible that FOXO3 plays a role in the dysregulation of autophagy in this model. Similar to the PI3K/Akt pathway, AMP-­‐activated protein kinase (AMPK) positively regulates mTORC1 and inactivates TSC2 of the TSC1/2 heterodimer complex (Sarbassov et al., 2005); in this way, mTORC1 is able to detect changes in energy state (ATP/AMP ratio) of an organism (Meijer and Codogno, 2006). The activation of AMPK in various tissues under fasting conditions has been well documented; AMPK stimulation in this manner stimulates autophagy and also affects pathways that lead ultimately to an increase in energy production and to the inhibition of pathways that deplete energy (Daval et al., 2006). This could be analogous to the conditions in Pcyt2 KO fasted states. It is known that Pcyt2 knockout in mice results in the accumulation of DAG in older mice (Fullerton et al., 2009) and that DAG content plays a significant role in cellular homeostasis by its mediated function on the Protein kinase C (PKC) family of proteins (Newton, 2001). Several studies have reported the involvement of specific PKC isoforms (β, γ, Θ) in autophagy (Chen et al., 2008; Sakaki and Kaufman, 2008; Zhang et al., 2009; Tan et al., 2012); however, studies have yielded some conflicting results. Some reports have documented the inhibition of autophagy under activation of PKC, while others studies have suggested that activation of PKC results in increased induction in autophagy (Tan et al., 2012). Given that DAG content is elevated in older Pcyt2 KO mice, it is plausible that PKC activity in these mice is also augmented and that induction of autophagy is regulated under this pathway. However, further studies aimed at 59 elucidating the specific mechanism underlying PKC-­‐mediated autophagy are necessary. Few studies have suggested that downstream effects of PKC could be modulated by mitogen activated protein kinases (MAPKs), including extracellular signal-­‐regulated kinases (ERKs), p38, and c-­‐Jun N-­‐terminal kinases (JNKs), given that PKC has been shown to be responsible for increased activity of select MAPKs (Sakaki and Kaufman, 2008; Zhang et al., 2009) and that such MAPKs have been shown to be involved in autophagy (Corcelle et al., 2007; Zhang et al., 2009). More recently, a connection between autophagy and lipid droplets (LDs) has warranted greater research attention as LDs have been shown to play a role in lipid metabolism and have been recognized as highly active sites requiring regulated turnover. Traditionally turnover in LDs was known to occur through TG degradation by lipolysis but recently some groups have suggested that LD degradation may occur via a specialized form of autophagy named lipophagy. LD turnover occurs in response to energy demands, as well as to an accumulation of lipids in order to ensure that lipid accumulation does not exceed the size of the cell (Singh and Cuervo, 2012). LDs are lipid reservoirs present in all cells but are found predominantly in adipose tissue (Fujimoto and Parton, 2011; Greenberg et al., 2011). Interestingly, most research aimed at examining the associations between LDs and autophagy have used nonadipocytes (Shibata et al., 2009; Singh et al., 2009), calling into question whether lipophagy occurs in adipocytes. The proposition that autophagy may be involved in lipid catabolism was based on studies by Singh et al. (2009) who found that hepatic TG accumulated when autophagy was inhibited, degradation of TG was reduced under autophagy inhibition, and Atg7 knockout in the liver of mice resulted in the development of liver steatosis. 60 Considering that Pcyt2 disruption similarly alters liver lipid metabolism (Fullerton et al., 2009), it is not unlikely that KO mice also experience defects in lipophagy, resulting in excessive activation of autophagy to compensate for elevated content of TG. In this manner, autophagy may be upregulated in KO mice to mediate the effects of metabolic dysfunction. Not surprisingly, histological analyses of liver tissue from KO mice showed an accumulation of LDs, in contrast to liver tissue of control littermates in which LDs were minimal (Fullerton et al., 2009). Histological analyses of LDs were done in 10-­‐
month old fasted KO and littermate control mice, future studies should examine LD accumulation in 2-­‐month old and 10-­‐month old fasted states to determine if any differences exist. In addition to those experiments, lipophagy in control and KO mice should be measured through microscopy experiments examining the colocalization of LDs with autophagic vacuoles. 4.4 Autophagy induction by phospholipid provision may be tissue specific It is entirely plausible that other organelles aid the ER in providing phospholipids for autophagosome formation and LC3 lipidation during autophagy, and that the organelles involved in providing phospholipids depend on the tissue in question. For instance, skeletal muscle is highly specialized to provide motion. In order to allow movement, muscle cells rely on the functional abundance of mitochondria for ATP production. Therefore, it would not be unlikely that mitochondria also assume the function of providing phospholipids including PE during autophagy by decarboxylation of PS in muscle tissues. However it should be acknowledged that much of the PS located in the mitochondria has been previously synthesized from PC and PE at the level of the 61 ER and is transported to the mitochondria to act as a substrate for PS decarboxylase (PSD) located in the mitochondrial membrane (Voelker, 1997). PC and PS cannot be made in the mitochondria and can only be supplied by mitochondrial membrane breakdown under the same conditions. 4.5 Future Work This study has focused on the effect of PE synthesis by the CDP-­‐ethanolamine pathway on the process of autophagy in vivo and in vitro. The involvement of additional phospholipids like PC and PS during this process should be investigated using glycerol, choline, and methionine experiments. These experiments would provide insight into the synthesis and turnover of PC and PS during autophagy. Furthermore it would be interesting to determine whether total content of DAG and TAG, as well as their metabolism in PCYT2 KD cells parallels that identified in Pcyt2 KO mice, which experience a different level of Pcyt2 activity. To investigate the contribution of PE synthesized from the CDP-­‐ethanolamine pathway for LC3 lipidation, immunoprecipitation experiments should be performed using radiolabeled glycerol and ethanolamine. 4.6 Conclusions The work undertaken in this thesis presented a novel approach to understanding the ongoing debate surrounding the source of the autophagosomal membrane during autophagy. The current study has demonstrated that autophagy induction is influenced by impairment to the CDP-­‐ethanolamine pathway, which further demonstrates a role of 62 the ER in synthesizing PE de novo for autophagic processes. Furthermore, this study has established that dysregulation of autophagy may be a critical component of metabolic syndrome in Pcyt2 KO mice as lipid metabolism is altered under CDP-­‐ethanolamine pathway impairment.………………………………………………………………………………………………… 63 5.0 REFERENCES Antonsson, B. (1997). Phosphatidylinositol synthase from mammalian tissues. Biochimica Et Biophysica Acta (BBA) -­‐ Lipids and Lipid Metabolism 1348, 179–186. Araki, N., Takashima, Y., and Makita, T. (1995). Redistribution and fate of colchicine-­‐
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