Ionization Properties of Diacylglycerol-Pyrophosphate and Phosphatidylinositol 3,4,5-Trisphosphate in Model Membranes

Ionization Properties of Diacylglycerol-pyrophosphate and Phosphatidylinositol 3,4,5-trisphosphate in Model Membranes

A thesis submitted To Kent State University in partial Fulfillment of the requirements for the degree of Master of Science

by Joseph Thomas

Thesis written by

Joseph S. Thomas

B.S., Kent State University, 2014 M.S., Kent State University, 2016

Approved by

Dr. Edgar E. Kooijman , Academic Advisor

Dr. Jennifer McDonough , Members, Thesis Committee

Dr. Ernest Freeman ,

Accepted by

Dr. Ernest Freeman , Chair, School of Biomedical Science

Dr. James L. Blank ,Dean, College of Arts and Sciences ii

Table Of Contents

List of Figures...... vi

Acknowledgements...... ix

Chapter 1: Introduction

1.1 Biomembranes...... 1

1.2 Important Lipids...... 5

1.2.1 Phosphatidylcholine...... 6

1.2.2 Phosphatidylethanolamine...... 7

1.2.3 Phosphatidic Acid...... 8

1.3 Diacylglycerol-pyrophosphate...... 10

1.3.1 Location and Importance...... 11

1.3.2 Structure and Charge...... 12

1.4 Phosphatidylinositol 3,4,5-trisphosphate...... 13

1.4.1 Location and Importance...... 14

1.4.2 Structure and Charge...... 15

1.5 Objectives and Methodology...... 15

1.6 References...... 18

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Chapter 2: Methods

2.1 31P NMR Titration...... 23

2.2 Preparation of Lipid Films...... 26

2.3 Preparation of MLV Dispersions...... 27

2.3.1 Preparation of Buffers...... 27

2.3.2 Formation of MLV’s...... 28

2.4 NMR Experimental Procedure...... 29

2.5 31P NMR Data Analysis...... 30

2.6 Giant Unilamellar Vesical Preparation...... 32

2.6.1 Preparation of Buffers...... 34

2.6.2 Electroformation...... 35

2.7 Fluorescence Microscopy...... 38

2.7.1 Fluorescent Lipids Used...... 40

2.7.2 Processing and Analysis of Fluorescence Images...... 41

2.8 References...... 42

Chapter 3: Ionization of Diacylglycerol-pyrophosphate in Model Membranes

3.1 Introduction...... 44

3.2 Results...... 47

3.2.1 Outer Phosphate of DGPP Exhibits Similar Titration Behavior to PA...... 48

3.2.2 Population of DGPP Inner Phosphate Groups Participate in Local Electrostatic

Interactions...... 50

3.2.3 Domain Formation in Giant Unilamellar Vesicles Occurs at Physiological pH. . . . . 52

3.3 Discussion...... 54

3.4 References...... 59

Chapter 4: Ionization of Phosphatidylinositol 3,4,5-trisphosphate in Model Membranes

4.1 Introduction...... 61

4.2 Results...... 64

4.2.1 Phosphatidylethanolamine promotes increased deprotonation of PIP3...... 65

4.2.2 Phosphatidylinositol reduces the overall charge of PIP3...... 67

4.2.3 PIP3 Promotes Domain Formation in Mixed Lipid Vesicles Containing

Phosphatidylinositol...... 68

4.3 Discussion...... 69

4.4 References...... 74

Chapter 5: Discussion and conclusions

5.1 Project summation...... 75

5.2 Future work...... 78

5.3 References...... 83

Appendix + List of Materials...... 84 v

List Of Figures

Figure 1-1: A depiction of the plasma membrane...... 2

Figure 1-2: A typical phospholipid molecule...... 3

Figure 1-3: Structures of phosphatidylcholine and sphingomyelin...... 6

Figure 1-4: Structure of phosphatidylethanolamine...... 7

Figure 1-5: Structure of phosphatidic acid...... 9

Figure 1-6: Structure of diacylglycerol-pyrophosphate...... 12

Figure 1-7: Structure of phosphatidylinositol and phosphatidylinositol 3,4,5-trisphosphate. . . 14

Figure 2-1: Representative 31P NMR spectra of an MLV dispersion...... 24

Figure 2-2: Titration curve constructed from raw NMR data...... 25

Figure 2-3: Typical format of MAS NMR spectra...... 31

Figure 2-4: Schematic of GUV electroformation chamber...... 36

Figure 2-5: Electroformation protocol for producing GUV’s...... 38

Figure 2-6: Schematic representation of an inverted epi-fluorescence microscope...... 39

Figure 2-7: Structures of fluorescent lipids used...... 41

Figure 3-1: The structures of phosphatidic acid and diacylglycerol-pyrophosphate...... 44

Figure 3-2: The electrostatic hydrogen bond switch model...... 45 vi

Figure 3-3: Ionization of DGPP with or without phosphatidylethanolamine present...... 46

Figure 3-4: Ionization behavior of DGPP and PA phospho-monoesters...... 48

Figure 3-5: Ionization of DGPP and PA phospho-monoesters in the presence of PE...... 50

Figure 3-6: NMR peak splitting occurs at higher concentrations of DGPP...... 51

Figure 3-7: Domain formation in vesicles containing DGPP and PA with NBD- PA...... 53

Figure 3-8: domain formation in vesicles containing DGPP and PA with Rhodamine-B...... 53

Table 3-9: pKa values of the DGPP and PA phospho-monoesters under different conditions. . .55

Figure 3-10: Hydrogen bonding between DGPP and PA...... 56

Figure 4-1: Head groups of phosphatidylinositol polyphosphates...... 62

Figure 4-2: Titration behavior of PIP3 in PC Vesicles...... 63

Figure 4-3: Ionization behavior of PIP3 in mixed vesicles containing PE...... 66

Figure 4-4: Ionization behavior of PIP3 in mixed vesicles containing PI...... 67

Figure 4-5: Giant unilamellar vesicles containing PI and PIP3...... 68

Figure 4-6: Hydrogen bond model for PIP3 with PE and PI...... 70

Figure 5-1: Charge repulsion between PE and PIP3...... 76

Figure 5-2: Charge repulsion between PI and PIP3...... 77

Figure S-1: Titration data for PC:PA at 95:5...... 89

vii

Figure S-2: Titration data for PC:PE:PA at 47.5 : 47.5 : 5...... 90

Figure S-3: Static spectra for PC:DGPP:PA at 90 : 5 : 5...... 91

Figure S-3: Static spectra for PC:PE:DGPP:PA at 45 : 45 : 5 : 5...... 92

Figure S-4: Static spectra for PC:DGPP:PA at 95 : 2.5 : 2.5...... 93

Figure S-5: Static spectrum for PC:PE:DGPP:PA at 47.5 : 47.5 : 2.5 : 2.5...... 94

Figure S-6: Static spectra for PC:PE:PIP3 at 47.5 : 47.5 : 5...... 95

Figure S-7: Static spectra for PC:PI:PIP3 at 88 : 10 : 2...... 96

viii

Acknowledgements

This work wouldn’t have been possible without help from many people. First I would like to thank my advisor, Dr. Edgar Kooijman. Over the course of my studies he has helped me become a capable young scientist. He has always been patient and understanding with me, and

I am thankful for the opportunity he gave me to be a part of his research group. I entered the lab as a young grad student with no idea how science really worked, but I feel like I am leaving with the tools and experience I need to tackle any academic endeavor that I may get into. Next I would like to thank the members of my research group, Dr. Zach Graber, Dr. Sewwandi

Rathnayake, Priya Putta, and Mona Mirheydari. They have been amazing colleagues and have taught me so much about what it means to be a scientist. Zach was the first student to train me when I started the lab and he went out of his way to teach me about NMR spectroscopy. I would also like to thank the various Kent State faculty who helped me with experiments along the way, including Dr. Mahinda Gangoda, Dr. Jennifer McDonough, and Dr. Mike Model. Dr.

Mahinda helped me troubleshoot the NMR and was there to help me every time I thought I may have damaged the expensive machine! Dr. McDonough helped me develop an interest in neuroscience which I will hopefully carry over into my PhD. Dr. Model showed me how to use confocal and fluorescence microscopes, which were key to gathering data for parts of this thesis. Another big part of my master’s experience was also my role as a student teacher. Doing this helped me gain confidence as scientist, and I would like to especially thank Shelley

Jurkiewicz for being an amazing lab coordinator.

On a more personal note, I would like to extend my gratitude to my parents and sister.

My parents have always been a big source of inspiration in my life, and they gave me the tools and work ethic that I needed to make it through graduate school. My sister was the first member of the family to receive a graduate degree, and this gave me the assurance I needed to know that I could do it too. My mom always pushed me to work hard and get the best education I could. I know that I am making her proud.

I also have countless friends who have contributed to me getting my degree who may not even realize they played a part. Playing in a band has been a much needed way to blow off steam outside of school, and I am grateful for the chance to be an artist as well as a scientist.

Chapter 1

Introduction

Biomembranes

Biological membranes play a crucial role in many critical cellular functions. They separate the interior of the cell from the exterior, and allow selective uptake and release of various molecules. Membranes are also present inside the cell, compartmentalizing various organelles. Even though biomembranes have long been known to be one of the fundamental components of cellular life, their exact structure and organization have been a topic of debate until fairly recently. Many different organizational models have been proposed, but the fluid mosaic model provides the best explanation for experimental observations (Singer and

Nicolson, 1972). The fluid mosaic model states that the plasma membrane is composed of a single lipid bilayer containing cholesterol, phospholipids, sphingolipids, and embedded proteins that are able to diffuse freely within the lipid matrix of the membrane. These proteins are thus able to interact with each other, or with molecules on the inside or outside of the cell. Proteins embedded within the membrane tend to contain regular secondary structures, such as alpha and beta helices (Kennedy, 1978). Alpha helices contain a hydrophobic outer surface that will prefer to be surrounded by the hydrophobic lipid acyl chains, allowing helix containing proteins to be inserted into the membrane. These secondary structures also act to anchor proteins

1 within the membrane, as removing the protein from the bilayer would be thermodynamically unfavorable due to the hydrophobicity of the helices. The original fluid mosaic model asserted that lipids were present primarily to give structure to the membrane, provide a matrix for proteins to insert, and to act as a physical barrier that sections the cell off from the outside environment. While these are indeed important functions of the membrane, recent evidence has come to light that shows a much more complex role of lipids, requiring a revision of the fluid mosaic model (Nicolson, 2014).

Figure 1-1: A depiction of the plasma membrane. The membrane consists of proteins, cholesterol, and many types of phospholipids. Proteins can span the entire membrane, associate with the membrane surface, or remain embedded within one leaflet. Due to their proximity to each other, phospholipids and proteins are easily able to interact. Source: Boundless biology (2016).

Phospholipids are amphiphilic molecules that contain a polar head group and non-polar acyl chains (Figure 1-2). Polar head groups among phospholipids show much diversity, but share a few common components. The polar head group consists of a glycerol backbone bound to a phosphate group. This phosphate group is bound to a variable (R) group that will give a lipid 2 unique properties (Discussed further in next section). The non-polar acyl chains also show variations in structure. Different lipids can have varying degrees of unsaturation and length in their acyl chains, which plays a vital role in how the lipid will behave in a membrane (Barelli

2016). The different head groups and acyl chains thus provide a huge diversity in distinct chemical species of lipids. A single biological membrane may contain hundreds of distinct chemical species (Feigenson 2006).

Figure 1-2: A typical phospholipid molecule. Phospholipids contain a polar (hydrophilic) head group and non-polar (hydrophobic) tails. The head group consists of a glycerol backbone bound to a phosphate group. This phosphate group is in turn bound to a variable (R) group that defines the identity of the specific lipid The non-polar tails can exhibit varying degrees of unsaturation which will influence the lipid’s behavior. Source: Boundless Biology (2016).

The amphiphilic nature of phospholipids allows them to form a bilayer structure in water (Hauser et al. 1986). The bilayer will form so that the hydrophilic lipid head groups will point out to be in contact with water, while the hydrophobic acyl chains will be sandwiched in the middle of the bilayer away from any water. This bilayer contains two distinct leaflets (inner and outer), which have varying lipid compositions (Bretscher, 1972). The inner leaflet refers to

3 the lipid in direct contact with the cytoplasm, and the outer leaflet refers to the lipid on the outside surface of the cell. Phosphatidylcholine and sphingomyelin are found in abundance in the outer leaflet of the plasma membrane but in lower amounts in the inner leaflet. On the other hand, phosphatidic acid, phosphatidylethanolamine, and phosphatidylserine are found in the inner leaflet, but are undetectable in the outer leaflet. The differing compositions of these leaflets has been implicated to play a role in many cellular processes, including apoptosis and phagocytosis (Tian et al. 2016). The membrane also possesses an overall negative charge due to the large amount of anionic lipids, such as phosphatidylserine and phosphatidylinositol, present in the inner leaflet (J.A. Op den Kamp, 1979).

Membrane lipids are able to form interactions with many different types of proteins.

Phosphoinositides (discussed later in this chapter) have been shown to interact with membrane ion channels via electrostatic forces (Mori and Inoue 2014, Furst et al 2014, Bians 2007). Lipids within the membrane have also been shown to act as second messengers by interacting with cytosolic proteins (Kooijman and Burger, 2009). Lipids in the membrane are also capable of forming interactions with other lipids around them. These interactions can occur via phospholipid head groups or acyl chains and can form functional subunits within the membrane, known as domains or rafts (Simons and Ikonen, 1997). Unsaturations in the acyl chains will introduce kinks, which will restrict how efficiently the lipids can pack together, and are inherently more disordered than chains with no unsaturations. Phospholipids with no chain unsaturations, such as dipalmitoyl (DP) lipids, are able to pack themselves efficiently into a bilayer and exhibit higher melting temperatures (Chen and Santore, 2014). On the other hand,

4 lipids that contain unsaturations, such as dioleoyl lipids (DO), will have a lower melting temperature and a higher degree of disorder in the membrane (Feigenson, 2006). Differences in melting temperature compose the foundation for phase transitions that occur within the membrane. Two of the phases that can occur are referred to as “liquid ordered” or “liquid disordered” and typically describe the amount of order/disorder present in the lipid acyl chains

(Schmidt and Davis, 2016). This order/disorder can be described by an order parameter which describes the average orientation of the methylene (CH2) and methyl groups (CH3) of the acyl chains (Gennis, 1989). Cholesterol is not classified as a phospholipid but it is known to play a role in these phase transitions due to its rigid sterol rings (Shimokawa et al. 2015) which can interrupt phospholipid packing and produce a liquid ordered phase (Feigenson 2006). Liquid ordered and disordered phases can coexist in the membrane and have the potential to mediate lipid/protein interaction (Simons and Ikonen, 1997). Interactions between phospholipids can also be head group mediated because of the many different chemical groups present, such as primary amines, hydroxyl groups, and phosphate groups that can participate in hydrogen bonding (Epand, 2015). Specific lipids and the interactions they can form are discussed in detail in the following section.

Important Lipids

Cellular membranes contain a rich variety of phospholipids. Different organelles in the cell, such as the mitochondria and the Golgi, have lipid compositions tailored to their unique functions (Mayr 2015, Zambrano 1974, Kleinig 1970). Due to the nature of this thesis, these membranes will not be discussed in detail. Rather, the focus will be placed on the lipids present 5 in cellular plasma membranes. The plasma membrane contains an abundance of different lipid sub-types, including phospholipids, sphingolipids, and cholesterol.

Phosphatidylcholine

The most abundant phospholipid in the plasma membrane is phosphatidylcholine (PC).

PC is found primarily in the outer leaflet of the membrane along with sphingomyelin. These lipids share similar structures (see figure 1-3), but are classified according to the makeup of their backbone. PC is a glycerol-phospholipid due to its glycerol backbone, while sphingomyelin is classified as a sphingolipid due to its sphingosine backbone (Merrill et al. 1997). Together these lipids provide the bulk structure of the cellular membrane and are important for cellular signaling. The cell maintains homeostatic levels of PC, and disruptions of this have been shown to play a role in apoptosis (Cui and Houweling, 2002).

Figure 1-3: Structures of phosphatidycholine (A) and sphingomyelin (B). These lipids share similar structures, except for the presence of an amide linkage in the sphingosine backbone of sphingomyelin. Acyl chains represented only reflect one possible composition.

PC is primarily made in the cell via the Kennedy pathway (McMaster and Bell, 1997). 6 this pathway involves converting choline into CDP-choline in the cytosol, which will ultimately be converted to phosphatidylcholine in the endoplasmic reticulum (Bishop and Bell, 1988). It is important to note that all phospholipids made in the cell are synthesized in the endoplasmic reticulum and processed in the Golgi, before being transported to the plasma membrane

(Pichler et al. 2001). PC and sphingomyelin are zwitterionic, with a negative charge present on the phosphate group and a positive charge present on the amine group. However, the quaternary amine in the head group is not able to participate in hydrogen bonding due to steric hindrance from the attached methyl groups.

Phosphatidylethanolamine

Phosphatidylethanolamine (PE) is another important phospholipid that has a structure similar to that of PC. PE is found primarily on the inner leaflet of the plasma membrane and is also zwitterionic. PE typically composes approximately 20 mol% of all phospholipids in the plasma membrane (Vance and Tasseva, 2013). The main structural difference between PC and

PE is that the amine present in the head group of PE is a primary amine (Figure 1-4). This means that it lacks the bulky methyl groups, and can participate in hydrogen bond interactions with proteins or other lipids in the membrane (Pink et al. 1998).

Figure 1-4: Structure of phosphatidylethanolamine. The PE head group contains a terminal amine which makes it available to participate in hydrogen bond reactions. Acyl chains depicted are di-oleoyl (DO) and represent only one possible composition. 7

PE is synthesized in the ER in a similar manner to PC via the CDP-ethanolamine pathway

(Bishop and Bell, 1988). The cell converts ethanolamine to CDP-ethanolamine in the cytosol, then ultimately to phosphatidyl-ethanolamine in the ER. Alternately, PE can be synthesized from phosphatidylserine, another inner leaflet phospholipid, via the action of phosphatidylserine-decarboxylase (Calzada et al. 2016). PE is believed to contribute to cell proliferation and growth by signaling via the mammalian target of rapamycin (mTOR) pathway

(Yang et al, 2016). PE is also believed to interact with other signaling proteins such as epidermal growth factor and protein kinase C, and deficiencies in levels of PE have been linked to defects in cellular differentiation (Yang et al. 2016, Kano-Sueoka 2001).

Phosphatidic acid

Phosphatidic acid (PA) is one of the simplest, but arguably one of the most important phospholipids found in the plasma membrane (figure 1-5). PA is found in low amounts (less than a few mol %) in the inner leaflet of the cell membrane, but is crucial for cell survival

(Kooijman and Burger, 2009). PA is found in plant and animal cells, and is believed to act as a second messenger. Its concentrations have been shown to increase during stress such as cold, wounding, salinity, and pathogen attack (Van Schooten 2006). PA is also important for lipid metabolism, because it acts as a key intermediate in phospholipid biosynthesis (Athenstaedt and Daum, 1999). PA can be synthesized de-novo in the cell by one of two pathways. In yeast and mammalian cells PA is made via the GrnP (dihydroxyacetone phosphate) pathway, whereas in bacteria and eukaryotes PA is made via the Gro3P (glycerol-3-phosphate) pathway (Sorger and Daum 2003, Yao and Rock 2013). Signaling pools of PA in the membrane can also be 8 produced indirectly by the action of phospholipase-C and diacyglycerol kinase, as well as directly via phospholipase-D which forms PA from PC and PE (Munnik et al. 2000).

Figure 1-5: Structure of phosphatidic acid. The phospho-monoester head group has the ability to carry either a -1 or -2 charge at physiological pH. PA serves as the common precursor for all phospholipids made by the cell. Acyl chains depicted are di-oleoyl (DO) and reflect only one possible composition.

PA has been shown to interact with many different proteins within the cell (Stace and

Ktistakis 2006, Testerink and Munnik 2005, Barbaglia et al 2016). These interactions are mediated by the ionization behavior of PA’s head group. PA’s head group consists of a phospho- monoester group with the ability to be deprotonated twice, which allows PA’s charge to be neutral, -1, or -2 depending on pH. The pKa of the first deprotonation event is approximately

3.2, so physiological systems will not contain neutral PA (Kooijmn, 2005). Rather the charge on

PA in biological membranes will be either -1 or -2 based on the second deprotonation event.

The pKa for this second deprotonation is approximately 7.92, which is right in the physiological range for cells (Kooijman, 2005). This means that small changes in local pH can have a large effect on the overall ionization state of the head group, which will alter how PA is able to recognize and interact with certain proteins (Kooijman et al 2009, Loew et al 2013). The charge on PA can also be influenced by interactions with surrounding lipids and proteins. Experiments

9 that mix PA with PE have shown that the presence of hydrogen bond donor groups in the membrane will increase the total negative charge of the PA phospho-monoester (Kooijman et al. 2005). The electrostatic hydrogen bond switch model has been proposed to describe the interaction of PA with its target proteins, and in fact describes the ionization behavior of any phospho-monoester group (Kooijman et al. 2009). In this model, PA is able to attract proteins that contain positively charged residues via electrostatic interactions. These cationic residues can then probe the cell membrane, and will form hydrogen bonding interactions with the PA phospho-monoester. Once this hydrogen bond forms, the overall charge on PA will become more negative due to the dissociation of the second proton. This change in charge will cause the protein to interact even more strongly with PA, and can provide a basis for PA specific recognition. The effective molecular shape of PA will also add curvature stress to the membrane, and is believed to play a role in vesicle fusion and fission (Roth, 2008). PA provides a model with which to extrapolate the behavior of other phospho-monoester containing lipids in the cell membrane. These lipids have the potential to exhibit complex ionization behavior due to hydrogen bond interactions with other lipids in the membrane, which has in fact been shown for ceramide-1-phosphate and phosphatidylinositol-4,5-bisphosphate (Kooijman et al.

2008, Graber 2012).

Diacyglycerol-pyrophosphate

Studying phospholipid ionization has yielded immense insight into how membrane lipids function in cellular signaling cascades, but the picture is still incomplete. Diacyglycerol- pyrophosphate (DGPP) is often associated with PA, a known second messenger in the plasma 10 membrane, and the two lipids are linked metabolically (Munnik et al. 2000). The two lipids have been studied independently, but no work has been done to characterize their ionization behavior when present in complex mixtures together. One focus of this thesis is the ionization behavior of DGPP in complex mixtures, with an emphasis on how it compares to the behavior of

PA. This comparison is important to determine if DGPP is able to form complex interactions with proteins similar to the hydrogen bond switch model observed in PA, since both lipids are implicated in plant stress signaling.

Location and Importance

DGPP is an important component of plant cell membranes (Wissing and Behrbohm

1993, Munnik et al. 2000). It is produced in the plant plasma membrane via the phosphorylation of phosphatidic acid (PA) by PA Kinase, and is broken down by DGPP

Phosphatase (Bas van Schooten et al. 2005). DGPP is undetectable under normal conditions, but sees an increase in concentration quickly following plant stress via turnover of PA (Munnik et al. 1996). This leads researchers to believe that DGPP acts to attenuate the action of PA in stress signaling. DGPP is believed to play a role in abscisic acid signaling (Zalejski et al. 2005), but its broader role within the membrane is still not well understood due to its fairly recent discovery (Wissing and Behrbohm 1993). Understanding the biophysical characteristics of DGPP is key to uncovering the role it ultimately plays within cells. The ionization behavior of phospholipids like PA gives tremendous insight into how it can interact with other molecules, so characterizing the ionization behavior of DGPP can yield the same vital information.

Structure and Charge

DGPP has a head group that consists of two phosphate groups (Figure 1-6). DGPP contains a phospho-monoester similar to PA, but also contains a phospho-diester group. These phosphates are simply referred to as “outer” and “inner” which describes their position in the molecule (Figure 1-6). Since the inner phosphate is involved in a phospho-diester linkage, it only has the ability to deprotonate once to carry a -1 charge. The outer phosphate is a phospho- monoester linkage, and can therefore deprotonate twice to a -2 charge. Taken together, this means that the DGPP head group theoretically can have a neutral, -1, -2, or -3 charge. However, data has shown that the inner phosphate will not deprotonate under physiological pH (Strawn et al. 2012). NMR experiments with just DGPP and PC in vesicles produce a first pKa around

2.82 and a second one around 7.44. This means that in a biological system, DGPP will only possess a -1 or -2 charge, with its ionization seemingly dominated by the phospho-monoester.

Figure 1-6: Structure of diacylglycerol-pyrophosphate. The DGPP head group consists of two phosphate groups (labelled outer and inner). At physiological pH, the inner phosphate remains protonated, and the outer phosphate can be singly or doubly deprotonated. Acyl chains depicted are di-oleoyl and only represent one possible composition.

The NMR experiments allow for independent observation of the inner phosphate, but this is complicated due to the broadness of the peaks and a significant change in charge is not 12 observed over the pH range tested (1.5-10.5). The electrostatic hydrogen bond switch model that was used to describe PA also applies to the DGPP head group. This means that lipids like PE will readily interact with the DGPP head group, promoting the second deprotonation and increasing the overall negative charge. This effect has been observed in experiments with

DGPP, PC, and PE in vesicles, with the first pKa was determined to be 2.61 and the second pKa was determined to be 6.71 (compared to 2.82 and 7.44 respectively). Since the electrostatic hydrogen bond switch model has implications for protein binding to PA, it can safely be assumed that DGPP may behave in a similar fashion. Characterizing DGPP’s charge in more complex mixtures can therefore shed more light on how it may behave in a biological membrane.

Phosphatidylinositol 3,4,5-trisphosphate

Phosphoinositides compose another group of crucial phospholipids present within the cell. They are present in very low amounts in the inner leaflet of various cell membranes, but play a vital role in transmembrane signaling (Balla, 2013). Phosphoinositides are all derivatives of phosphatidylinositol (Figure 1-7) and can be mono-, bis-, or triphosphorylated, with their metabolism being controlled by many different kinases and phosphatases (Stahelin et al. 2014,

Balla 2013). This causes these lipids to be highly anionic, and allows them to interact with many proteins that contain cationic clusters (Catimel et al. 2008, 2009). One such protein cluster is known as the Pleckstrin Homology (PH) domain, which is known to specifically recognize phosphoinositide head groups (Lemmon and Ferguson 2000, Balla 2005). Since these protein domains recognize the phosphoinositide head group based on charge, it is important to 13 characterize the ionization behavior of these lipids within the membrane. This thesis focuses on phosphatiylinositol 3,4,5-trisphosphate (PIP3), which contains the most complex phosphoinositide head group (see figure 1-7 below).

Location and importance

PIP3 is formed by the phosphorylation of phosphatidylinositol 4,5-bisphosphate (PIP2) by class I PI-3 kinases (Lindmo and Stenmark, 2006) and can also be dephosphorylated by a variety of 5-phosphatases. PIP3 has only been observed in Eukaryotic cells and is believed to play a role in many different cellular processes including membrane trafficking, cell proliferation, and cell survival (Lindmo and Stenmark 2006, Shaw and Cantley 2006). PIP3 is also believed to play a role in cancer biology via its interactions with the enzyme PI-3 kinase and the tumor suppressor

PTEN (Kooijman et al. 2009, Gericke et al. 2006). PTEN will convert PIP3 to PIP2 which limits cell growth and proliferation. When this process is dysregulated, a cell can grow out of control leading to cancer. Understanding the biophysical behavior of PIP3 can shed some light on how it functions in this pathway, opening the door to possible anti-cancer therapies.

Figure 1-7: Structure of phosphatidylinositol (A) and phosphatidylinositol 3,4,5-trisphosphate (B). PIP3 contains phosphorylations at the 3, 4, and 5 positions on the inositol ring.

Structure and Charge

PIP3 is trisphosphorylated, and therefore has the potential to exhibit complex ionization behavior. Each of the phosphates in the head group have the ability to titrate independently of each other, but they also have the potential to interact with each other due to their close proximity (Figure 1-7). Previous work with PIP3 (Kooijman et al. 2009) has shown that the 3 and

5 phosphates exhibit close to standard sigmoidal titration behaviors, whereas the 4 phosphate shows more complex titration. This makes sense considering the 4 phosphate is located in between the 3 and 5 phosphates, meaning it has 2 adjacent phosphate groups with which to interact. The interdependence of these phosphate titrations could potentially have great importance in how cationic protein domains interact, contributing greatly to interaction specificity. The complex ionization behavior of PIP3 in simple vesicles has previously been modeled (Graber, 2014a) but will not be discussed in depth in this thesis. Each of the inositol ring phosphates of PIP3 appear to have a pKa around physiological pH meaning that, much like the case with DGPP, small fluctuations in pH will have a dramatic effect on the head groups ionization. Also, as with any phospho-monoester group in the membrane, there is a potential to form hydrogen bonds with other lipid head groups.

Objectives and Methodology

The primary objective of the work contained in this thesis was to characterize the

15 ionization behavior of DGPP and PIP3 in model membranes under physiologically relevant conditions. This work led to a secondary objective, which was to determine if DGPP and PIP3 play a role in domain formation in model membranes. These lipids have previously been examined in simple lipid mixtures with phosphatidylcholine (PC) (Strawn et al. 2012, Kooijman et al. 2009) but until this wok it was not completely known how they would behave in more complex mixtures containing lipids that have the ability to form hydrogen bond interactions, such as phosphatidic acid (PA), phosphatidylethanolamine (PE) and phosphatidylinositol (PI).

The model systems that were used did not contain the complexity of a true biological membrane, but still provide insight into the behavior of these lipids in vivo. The inner leaflet of the plasma membrane contains many different lipid species that have the potential to interact with each other. These numerous individual interactions all come together to produce the complex behavior that is seen in living systems. In order to determine the effects of these individual interactions, simple mixtures must first be studied.

DGPP was chosen for study because of its lack of extensive biophysical characterization and known role in stress signaling, as well as the research carried out into the behavior of PA

(Kooijman et al 2005, Kooijman and Burger 2009). Since these lipids share a similar structure, it was believed that they may also share similar ionization behavior. Ionization behavior of DGPP was probed using 31P NMR. This procedure is widely used in biophysical studies and is explained in detail in the next chapter. These experiments reveal the ionization of lipids in a mixture with great accuracy, and allow for the construction of titration curves over a broad pH range. These titration curves provide information on the behavior of a lipid under many different conditions

16 and can be used to calculate the pKa of a deprotonation event. NMR peak splitting was observed in DGPP mixtures that did not contain PE at low pH, which is indicative of possible domain formation in the mixture. This splitting was observed in both PA and DGPP in the absence of PE, which hinted at an interaction between the two. One possible explanation for this interaction could be the presence of membrane domains. At low pH, both PA and DGPP are not fully deprotonated, so the possibility of hydrogen bond interactions between their head groups is present. After these lipids are fully deprotonated at high pH, this possibility no longer exists and NMR peak splitting is no longer observed. These domains may be able to manifest themselves on the macro-scale, and would therefore be observable in lipid vesicles with standard microscopy techniques. Lipid domains typically involve the phase separation of different lipids, so fluorescence microscopy was used. PIP3 was chosen for study because of previous work with PIP2 (Graber et al, 2014b). PIP2 exhibits complex ionization behavior and shares a similar structure to PIP3. It was believed that the 4 and 5 phosphate of PIP3 would also exhibit complex ionization behavior, but it was not known how the presence of the 3 phosphate group would affect this. NMR was used to observe PIP3 in the same way it was used to observe

DGPP. The possibility of domain formation was also probed, although no NMR peak splitting was observed. Previous work with PIP2 showed the formation of domains in lipid vesicles containing PI (Graber et al. 2012), so it was hypothesized that PIP3 would promote domain formation under similar conditions.

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Chapter 2

Methods

31P NMR titration

Solid state 31P NMR Spectroscopy has been widely used to characterize the ionization behavior of many phospholipids in small unilamellar vesicle (SUV) dispersions (Swairjo et al.

1994). These studies can provide insight into the behavior of membrane lipids, but SUV’s do not represent a true membrane model due to their small size and high curvature. Multi-lamellar vesicles (MLV) are large structures composed of many bilayers that have curvature similar to that of a typical cell membrane. This property makes them an attractive membrane model, but also means that NMR experiments will exhibit significant chemical shift anisotropy as seen in figure 2-1a (Graber and Kooijman 2013, Berden et al. 1974). Chemical shift anisotropy (CSA) is caused by the inability of the large MLV’s to reorient quickly within the magnetic field of the spectrometer. This results in broad peaks within the experimental spectrum that can obscure the chemical shift data of the lipids making up the model membrane (an example of which is seen in figure 2-1). These broad spectra can still provide information about the bulk ordering of the lipid in the sample, but in order to observe signals coming from individual nuclei a different technique must be used. This high resolution spectroscopy can be accomplished by a procedure known as magic angle spinning (Watts, 1998). In magic angle spinning (MAS), the sample is

23 spun at high speeds (>1000 Hz) at an angle of 54.74° with respect to the magnetic field. This spinning will average out the orientation dependent interactions with the magnetic field that cause large chemical shift anisotropy in static experiments. This averaging allows the individual lipid chemical shifts of each lipid in the mixture to be observed with high resolution. Running both static and MAS experiments can give information about the bulk ordering of the lipid as well as the individual chemical shifts of each lipid present in the mixture.

Figure 2-1: Representative 31P NMR spectra of an MLV dispersion. Chemical shift anisotropy seen in (A) is caused by the relatively slow motions of the large MLV’s in the magnetic field. The shape of the spectra in (A) is indicative of lipid bilayers. MAS allows individual lipid peaks to be resolved (B) compared to the non-spinning (static) sample. Lipid mixture is the same for both spectra (57.5 mol% DOPC, 37.5 mol% DOPS and 5 mol% DOPA). Peaks are relative to an external 85% H3PO4 standard.

The high resolution of these experiments allows for the observation of chemical shift values in complex mixtures. The chemical shift measures the extent to which a particular nuclei

24 is shielded from the external magnetic field by moving charges around it (protons, electrons) and is measured against a reference. These moving charges will produce their own magnetic field that counteracts the applied magnetic field, shielding the nucleus. In the case of 31P NMR the chemical shift can also measure the degree of protonation around the phosphorous nucleus and can be used as an indirect measure of the lipid ionization. It is possible to detect how the chemical shift of each lipid phosphomonoester peak changes in response to pH, producing a titration curve (figure 2-2). This curve can be used to predict the ionization behavior of the particular phospholipid, as well as to determine its pKa values. MAS experiments have been adopted for studies of many phosphomonoester containing lipids (Kooijman et al. 2005), and can also be used for lipids which contain several phosphate groups like phosphatidylinositol

(4,5)bisphosphate (PI(4,5)P2) (Graber et al. 2014). Both Diacylglycerol-pyrophosphate (DGPP) and phosphatidylinositol 3,4,5-trisphosphate (PIP3) contain phosphodiester groups, as well as

31 phospho-monoesters in the form of terminal phosphates (3 in the case of PIP3). P MAS NMR allows for the observation of each of these distinct phosphorous nuclei with high resolution.

Figure 2-2: Titration curve constructed from raw NMR data. Experiments are done at many different pH values (A) and chemical shift values are used to construct a titration curve (B). Separate titration curves are constructed for the 4 and 5 phosphates of phosphatidylinositol 4,5-bisphosphate as indicated. Spectra obtained from an MLV dispersion consisting of 95 mol% DOPC and 5 mol% PI(4,5)P2. Chemical shifts are relative to an external H3PO4 standard. Data taken from (Kooijman et al. 2009a).

Preparation of lipid films

All lipids used in this work were obtained from Avanti Polar Lipids (Alabaster, AL).

Vesicles used in NMR experiments were formed from dry lipid films. Lipid films were prepared by mixing desired ratios of stock solutions, then drying the organic solvent. To create lipid stocks, appropriate amounts of lipid powder were weighed on a semi micro-balance and dissolved in 2:1 chloroform/methanol. Phosphoinositides (PI, PIP3) required slightly different preparation, and were dissolved in 20:9:1 chloroform/methanol/water to ensure complete solvation. Stocks were stored at -20°C under inert nitrogen gas Purity was assessed via thin layer chromatography (TLC), as well as phosphate assay as previously described (Rouser et al.

1970). TLC tests were performed to test if any lipid breakdown had occurred during storage.

Only If TLC resulted in one spot after iodine staining was the lipid deemed to be pure. A phosphate assay was used to assess how the concentration of stock solutions had changed during storage. Over time, organic solvent slowly evaporates from the storage vial, which can result in a stock solution being more concentrated than expected. Stocks were tested regularly to track purity and concentration.

Lipid films were prepared by mixing desired ratios of lipid stock in borosilicate glass test

26 tubes. The total amount of lipid in each film was typically between 1-10 µmol total. PIP3 experiments had a lower amount of total lipid due to the high cost of phosphoinositides. Each film contained approximately .1 µmol of PIP3, but this still allowed for resolved spectra on reasonable time scales. 400 µL of chloroform was added to increase the total volume of the lipid mixture. This helped to facilitate a more even drying of the film. The bulk organic solvent was evaporated under a steady stream of nitrogen gas. This process usually took 5-10 minutes per film depending on solvent volume and presence of water. Films were visually inspected for degree of dryness and were transferred to a large vacuum oven (VWR, PA). The chamber was evacuated with a Dry Fast Ultra pump for approximately 20 minutes, until a pressure of at least

-70 mm Hg was achieved. Films were left under vacuum for 4 hours to ensure removal of trace organic solvents which affect ordering of the acyl chains. Films containing phosphoinositides were heated to 30°C to ensure complete removal of trace water. Upon completion of drying, lipid films were placed under inert nitrogen gas and frozen at -20 °C. Films were typically prepared in bulk to minimize variation among samples due to pipetting of organic solvents.

Preparation of MLV dispersions

Preparation of Buffers

All buffers contained 100 mM NaCl to match cellular ionic strength, along with 2 mM

EDTA (ethylenediaminetetraacetic acid) acting as a divalent cation chelating agent. For pH <6.5,

20 mM Citric Acid and 30 mM HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid) were used as the buffering agent. For pH 6.5-8.5 50 mM HEPES was used, and for pH >8.5 50 mM

Glycine was used. Buffers were prepared so that there would be a separate buffer for each desired pH. Buffers were prepared to start at pH 4 and increased in increments of 0.4 until pH

11 (4, 4.4, 4.8, 5.2… etc,). Each buffer was pH adjusted via the drop wise addition of concentrated NaOH or HCl using an electronic pH probe (Sentron SI600, Netherlands). Before experiments, the pH of the buffer was tested to ensure that the pH had not changed significantly. A new buffer was prepared if the pH deviated more than 0.1 from the prepared value. All buffers were prepared with submicron filtered ultra-pure water (Fisher Scientific

CAS#: 7732-18-5, MA). All buffer components were biology grade and obtained from various sources (see List of Materials).

Formation of MLV’s

Multi-lamellar dispersions were produced via hydration of lipid films with aqueous buffer. First, the lipid film was hydrated with 2 ml of the desired physiological buffer. This suspension was gently vortexed to ensure that the lipid film was completely stripped from the sides of the test tube. Next, a freeze/thaw procedure was used to break up metastable phases within the MLV’s and ensured a more homogenous size distribution. A Dewer flask containing ethanol was cooled with chunks of dry ice. When the ethanol was sufficiently cold, the test tube containing the lipid suspension was dipped inside and allowed to sit until the contents were completely frozen (2-5 minutes). The tube was removed from the flask and quickly brought back to room temperature in a warm water bath. This process was repeated 3 times for films containing DGPP, and only once for films containing PIP3. Highly anionic lipids like PIP3 have the ability to form small vesicles which can disrupt MAS experiments, but this was not a major 28 concern for DGPP mixtures. After the freeze/thaw was complete, the pH of the sample was read to determine the final experimental pH that would be recorded and used to construct the titration curve. The MLV dispersion was centrifuged at 15,000 RPM for 60 minutes (4°C) in a tabletop ultra-centrifuge (Hettich, Germany) to make it easier to load into the MAS rotor. The pellet was collected and pipetted into a 4mm zirconia (ZrO2) rotor which was then sealed with a plastic MAS cap (Bruker, MA). During loading of the sample, care was taken to eliminate any air bubbles that may have formed. The sample was mixed around inside the rotor to ensure homogeneity of the MLV dispersion, as imbalance within the rotor would inhibit spinning.

Typically, 75-100% of the total lipid would be loaded into the rotor.

NMR experimental procedure

Experiments were carried out at room temperature on a Bruker 400 MHz spectrometer using Topspin software and a 4mm cross-polarization MAS probe. Before each experiment, the spectrometer was standardized with an 85% phosphoric acid (H3PO4) solution in a 4mm MAS rotor. The sample was placed in the spectrometer and spun at 1000 Hz to test for spinning stability. The spin was then set to 2000 Hz, and several scans were taken of the sample

(experimental parameters listed in appendix). This resulted in a spectrum of 1 peak that was set to zero on the chemical shift scale, and a spectral reference number was recorded. The standard was removed and the sample was placed in the spectrometer. Spinning was tested at

2000 Hz, and then set to 5000 Hz once stability was achieved. The spectral reference obtained from the standard was then entered to calibrate the spectrometer. The spectrometer was also tuned to 31P resonance (161.97 MHz) which ensured optimal data collection. Resolution was 29 dependent upon the total amount of sample, as well as the total experimental time. Samples containing DGPP were fully resolved after approximately 20,000 scans (4 hours) due to the large amount of lipid in each sample. Samples containing PIP3 had a longer experimental time due to the low amount of total lipid, and were resolved after approximately 100,000 scans (24 hours) (experimental parameters listed in appendix).

For select samples (typically pH 4, 7, and 9), static spectra were also recorded. These static spectra give information about the bulk organization of the lipid sample, and were used to ensure that the lipid had formed bilayers. After completion of the MAS experiment, the spin was set to 0 Hz and the calibration from the previous standard experiment was re-used. The spectrometer was tuned to the 31P resonance as well as the 1H resonance. A different pulse program was used for static experiments (listed in appendix) and the samples were allowed to run for 100,000-300,000 scans (24-48 hours). This was necessary to ensure optimum resolution, as static NMR experiments are inherently very noisy. This is due to the lack of orientational averaging that is present in MAS experiments, causing the signal intensity to be spread over a broader chemical shift range. Once experiments were completed, the sample was removed from the rotor and stored at -20°C. Rotors were cleaned with pure water and ethanol, and then reused.

31P NMR data analysis

Titration curves can be built by plotting the chemical shift value of an individual phosphate peak over a range of pH values (see figure 2-2). To obtain chemical shift values a

30 peak picking function is used in the Topspin software, and peaks in the raw spectra can be assigned based on their chemical shift values. Peaks with higher (more positive) values are considered to be “downfield”, and represent phosphorous nuclei with a lower charge density.

Peaks with a lower (more negative) chemical shift value are considered to be “upfield” and have a higher charge density (Kuhl 2008, Graber and Kooijman 2013). These charge densities around nuclei will “shield” them from the external magnetic field produced by the spectrometer. Based on this NMR spectra are typically represented with a flipped X-axis (figure

2-3) so that more shielded nuclei are farther to the right, and more de-shielded nuclei are farther to the left. Since the lipid chemical structures are known, it is easy to assign the experimental peaks to the lipids in the sample with a high degree of confidence.

Figure 2-3: Typical format of MAS NMR spectra. Lipid mixture shown is 45 mol% DOPC, 50 mol% DOPE, and 5 mol% DOPA at pH 7. X-axis is flipped to have “higher” chemical shift values to the left and “lower” values to the right. Degree of shielding from external magnetic field is therefore represented like a traditional axis (more shielded to the right, more negative chemical shift). Lipid phosphomonoester peaks are assigned using known structures. DOPC is the most shielded, and DOPA is the most deshielded. Zero chemical shift determined by external H3PO4 standard. 31

Once a titration curve is constructed, it can be fitted with a modified Henderson-

Hassalbalch equation since it exhibits a sigmoidal shape (Kooijman, 2006). Based on this fitting, the total charge and pKa of the lipid head group can be determined (Kooijman et al. 2005).

Experiments are carried out to determine the chemical shift values of the protonated (δa) and deprotonated (δb) states, along with the experimental pH and chemical shift value (δ) of a single NMR experiment. These values are then plugged into the following equation:

훿 푋 10푝퐾푎−푝퐻+ 훿 훿 = 푎 푏 (1) 1+10푝퐾푎−푝퐻

This pKa value is equivalent to the pH value at which there are equal amounts of protonated and deprotonated species present. This number is significant because at/around that pH value, small changes in pH can have a drastic effect on head group ionization.

Phosphatidylinositol-polyphosphates exhibit complex ionization behavior (Kooijman et al.

2009a) and require a more robust fitting model due to intramolecular interactions between phosphate groups. The fitting of the data was beyond the scope of the work presented in this thesis, and was carried out by a previous graduate student as outlined in his dissertation

(Graber, 2014).

Giant unilamellar vesical preparation

In order to probe for domain formation in mixtures containing DGPP or PIP3, giant unilamellar vesicles (GUV’s) were used as a model system. GUV’s provide an excellent model for cell membranes, because they are of similar size (10-100 µm) and only consist of one lipid bilayer. Their large size, along with the wide availability of various fluorescent lipid probes,

32 makes them an ideal system to study via fluorescence microscopy (Bagatolli and Gratton 1999,

Tanaka et al 2004). Domains in GUV’s have been widely studied, and typically result from phase separations of lipids within the membrane facilitated by interactions of the lipid tails (Baumgart et al. 2003, Chen et al. 2014, Shimokawa et al. 2015). Other studies using lipid monolayers have shown that the head groups of phospholipids can interact to affect lipid ordering within the membrane as well (Villasuso et al 2010, Kooijman et al 2009b). These interactions have the potential to form large domains within the GUV bilayer which can be observed via fluorescence microscopy.

GUV’s can be formed by a variety of methods, the most common is the use of alternating electric fields, called electroformation (Veatch, 2007). These experiments involve the construction of an electroformation chamber with glass plates coated in indium tin oxide

(ITO). These plates are highly conductive, and enclose an aqueous buffer. Unfortunately, high salt concentrations (>10 mM) in the buffer will cause stripping of the ITO from the glass plates allowing current to pass through the lipid sample, causing the vesicle formation to fail (Zachary

Graber, personal communication). Standard electroformation is effective in producing a large amount of GUV’s in a short time, but cannot be done under physiologically relevant conditions.

For these conditions, gentle hydration is the preferred method (Akashi et al. 1996). This method relies on the spontaneous formation of lipid bilayers in water. A lipid film is prepared in a test tube and carefully submerged in an aqueous buffer. The test tube is heated to just above the melting temperature of the highest melting point lipid in the mixture. Over time the film will spontaneously strip off the inner wall of the test tube to form GUV’s. Gentle hydration will

33 allow the formation of vesicles under high ionic strength, but yields are typically very low and the formation process takes approximately 24 hours. In order to achieve high yields under physiologically relevant conditions, a modified electroformation protocol was used (Meleard et al. 2009). This protocol proved to be very effective at forming GUV’s, and is described in detail below.

Preparation of Buffers

The electroformation process involved the use of two separate buffers. These buffers are referred to as “internal” and “external”, and refer to their use in the electroformation procedure. The internal buffer would ultimately be encapsulated in the lipid vesicles, and the external buffer was used to dilute the GUV solution and allow them to settle at the bottom of the viewing plate for observation under the microscope. The internal buffer contained 100 mM sucrose, and the external buffer contained 100 mM glucose. This was done so that the GUV’s would be weighed down and sink to the bottom of the viewing plate, due to the different densities of the sugars. 100 mM of sugar was chosen to ensure efficient sedimentation of vesicles upon dilution. For DGPP experiments, both buffers contained 100 mM NaCl along with

2 mM EDTA. 100 mM NaCl was used to mimic physiological salt concentrations and ionic strength. Experiments were carried out at pH 4, 6, and 9, and a different buffer was used for each value. 10 mM citric acid was used for pH 4, 10 mM HEPES for pH 6, and 10 mM glycine for pH 9. The pH’s were adjusted as described above for the NMR buffers, and care was taken to ensure that internal and external buffers of the same pH were as close as possible to each other

(within 0.1 pH units). Large differences in pH will create osmotic stress across lipid membranes, 34 and can negatively affect vesicle stability. For experiments involving PIP3, both internal and external buffers contained 100 mM NaCl, 0.1 mM EDTA and 5 mM PIPES as a buffering agent.

PIP3 experiments were only carried out at a single pH value of 7.2, and were adjusted as described previously. All buffers were prepared with sub-micron filtered pure water (Fischer

Scientific, MA) and biology grade chemical stocks obtained from various sources (listed in appendix).

Electroformation

An electroformation chamber was constructed as shown (figure 2-4). The chamber consisted of a 10 mm quartz cuvette with a Teflon top (Hellma Analytics,Germany). Holes were drilled in the top to allow the insertion of 2 parallel platinum wires (Surepure Chemetals, NJ).

Wires were placed 9.6 mm (0.38 inches) apart from each other, then the ends of the wire above the Teflon cap were bent and wrapped in electrical tape to hold them in place. This allowed for easy manipulation of the cap and wires during the lipid coating process, and made it so that the wire would not need to be placed in the cap before each new experiment. Wires had a diameter of .020 inches and a purity of 99.95%. Platinum was chosen for its high conductivity, and resistance to corrosion since the wires would be in direct contact with aqueous buffer. The platinum wires were connected to a waveform generator (Rigol 2-channel, OR) via electric leads with alligator clips. The waveform generator allowed for the manipulation of applied voltage, as well as AC frequency. The electroformation chamber essentially functioned as a large capacitor, allowing electrical discharge through the lipid film that was eventually placed on the wires.

Since the platinum wires were the same distance apart from each other as they extended into the glass cuvette, it is assumed that all points on the wire experienced a uniform electric field.

Figure 2-4: Schematic of GUV electroformation chamber. Electroformation chamber consisting of parallel platinum wires within a quartz cuvette. A waveform generator supplied the voltage and aqueous buffer within the chamber completed the circuit. AC current creates standing waves between parallel wires that cause GUV’s to form and swell. The diagram on the right shows a simple electrical schematic for the apparatus. Experiments were conducted with the cuvette wrapped in aluminum foil to minimize light exposure.

The platinum wire was spotted with a lipid mixture prepared from a lipid film. Lipid films were prepared as described above. These films typically contained 1 µmol total lipid, and contained 0.1 mol% of either NBD-PA or rhodamine-PE (discussed further in next section). Lipid films were dissolved with 50 µL of 2:1 chloroform/methanol so that the wire would be spotted with a concentrated solution. This was done to prevent the lipid drops from running down the

36 wire and collecting on the Teflon cap. A micro-pipette was used to spot the lipid mixture onto the wire in 2 µL aliquots. At the end of this process, the area of both wires below the Teflon cap was evenly coated with a lipid film. The coated wire was placed in a vacuum oven as described previously for 1 hour. This was done to ensure the complete drying of trace organic solvents from the lipid film. The glass cuvette was filled with approximately 10 ml of internal buffer, and mixtures containing DGPP were heated to 60°C in a hot water bath to ensure melting of the lipids during vesicle formation. Formation of mixtures containing PIP3 was done at room temperature, due to the sensitivity of the lipid. Once this temperature was reached, the lipid coated wires were carefully submerged in the buffer by placing the Teflon cap on top of the cuvette. Care was taken to avoid trapping air bubbles in the cuvette, so that the chamber was completely full of buffer. Electric leads were connected to the section of the platinum wires extending above the Teflon cap with alligator clips, and then connected to the waveform generator. The apparatus remained in the hot water bath for the entire electroformation, and temperature was monitored via a standard alcohol thermometer. The electroformation parameters used are outlined below (table 2-5). The applied voltage was calculated by multiplying the desired electric field (E) by the distance separating the wires (9.6 mm). For example:

(50 V/m) X (0.0096 m) = 0.48 V

During step 1, the voltage was slowly increased over 30 minutes in a uniform manner (0.5 V every 72 seconds). During the final step, it was determined that vesicle release was more efficient when the AC frequency was quickly decreased. Typically, the frequency was decreased 37 from 500 Hz to 50 Hz over one minute or less, which allowed more time at a low AC frequency to promote vesicle release into the buffer.

Table 2-5: Electroformation protocol for producing GUV’s. The first step involves slowly increasing the voltage over 30 minutes to encourage the lipids to form small unilamellar structures. The second step involves a strong constant voltage for 90 minutes to allow the vesicles to swell in size. The final step involves decreasing the AC frequency to promote vesicle release from the wires into the surrounding buffer. Applied voltages are peak-to-peak since AC current is used. Parameters taken from (Maleard et al. 2009).

Once electroformation was complete, the cuvette was gently flicked to remove any remaining lipid from the platinum wires. The GUV containing buffer was allowed to return to room temperature and then 1-2 drops were taken and placed in each well of an 8-well #1 borosilicate viewing plate (Nalge Nunc, NY) with a Pasteur pipette. 8-10 drops of external buffer were then placed in each well to dilute the GUV suspension, as well as facilitate viewing

(discussed above). The GUV’s were given 5-10 minutes to settle to the bottom of the plate, and were then viewed under a fluorescence microscope at room temperature.

Fluorescence microscopy

GUV’s were viewed using an Olympus IX81 inverted epi-fluorescence microscope

(Olympus, Japan) running SlideBook 6 software (3i, CT). Inverted fluorescence microscopes 38 work by emitting a laser through the objective lens onto the sample above it, and then detecting the light emitted by the sample through the same objective lens. Since most of the laser will pass through the sample away from the detector, only the light emitted by the sample itself will reach the detector. This creates an excellent signal to noise ratio, and provides high resolution images.

Figure 2-6: Schematic representation of an inverted epi-fluorescence microscope. Laser light excites the sample from below through the objective lens. Emitted light is collected through the same objective and sent to a detector, producing high resolution images.

Images were viewed with a 40X long working distance lens (Olympus, Japan). Samples were viewed in a #1 borosilicate plate and lens jacket was adjusted accordingly to maximize resolution while focused. Images were taken with an Orca R2 digital CCD camera (Hamamatsu

Photonics, Japan). Exposure times ranged from 100 ms to 5000 ms, depending on fluorescence intensity and vesicle size.

Fluorescent lipids used

Experiments used either Rhodamine-B (sometimes known as Rhodamine-PE) or NBD-PA.

Rhodamine-B consisted of a phosphatidylethanolamine with a rhodamine modified head group, and NBD-PA consisted of a phosphatidic acid with an NBD modified acyl chain (shown in figure

2-7). These dyes have been widely used in biological experiments, because they are easily incorporated into lipid vesicles under various conditions. Multiple fluorophores were used in order to check that any domains observed were not just due to the activity of the fluorophore itself. NBD modified lipids are not sensitive to extreme pH (Ishiguro et al. 2010) but their structure adds significant bulk to the lipid acyl chains, which can affect how they will act in a bilayer. The NBD group is considerably hydrophilic, so it is likely that it will snorkel up to the lipid/water interface and affect head group packing (Kaiser and London, 1998). This phenomenon could disrupt any head group interactions, and cause the fluorophore to be preferentially excluded from areas with tight head group packing. This allows for visualization of areas in the membrane that show tight clustering of lipids via exclusion of the fluorophore.

Rhodamine modified lipids do not possess this acyl chain bulk, but are sensitive to the surrounding pH (Jorge et al. 2013). At high and low pH the fluorescence is considerably more dim, but this effect was determined to be minimal during experiments due to the amount of fluorophore used (0.1 mol%). Rhodamine-B will preferentially reside in areas of the membrane with a high degree of disorder, since the bulky fluorophore attached to the head group will prohibit close interactions in the same way as NBD-PA. In this way, any domains or clusters that may form in the membrane can be visualized.

Figure 2-7: Structures of fluorescent lipids used. GUV experiments used either NBD-PA (A) or Rhodamine-B (B).

Processing and analysis of fluorescence images

Images were processed in the SlideBook software itself, since it included robust image editing capabilities. Brightness was typically edited to enhance visibility, but no other alterations were done. Scale bars were added after imaging, and were calibrated digitally.

Images were cropped in Image-J (NIH) so that only vesicles of interest were depicted, as images typically contained many vesicles. Vesicles were considered true GUV’s if they only had one visible bilayer, were not significantly filled with other vesicles/lipids, and were not directly touching any other vesicles. Electroformation produced many vesicles of varying sizes, approximately 20% of which were true GUV’s. Domain formation was observed as an area of fluorophore exclusion, an area of altered local curvature (bulging), or both.

References

Akashi K, Miyata H, Itoh H, Kinosita K. Preparation of giant liposomes in physiological conditions and their characterization under an optical microscope, Biophys. J. 71 (1996) 3242–3250 Bagatolli LA and Gratton E. Two-Photon Fluorescence Microscopy Observation of Shape Changes at the Phase Transition in Phospholipid Giant Unilamellar Vesicles. Biophysical Journal 77 (1999) 2090-2101 Baumgart T, Hess ST, and Webb WW. Imaging coexisting fluid domains in biomembrane models coupling curvature and line tension. Nature 425 (2003) Berden JA, Cullis PR, Hoult DI, McLaughlin AC, Radda GK, and Richards RE. Frequency Dependence of 31P NMR Linewidths in Sonicated Phospholipid Vesicles: effects of Chemical Shift Anisotropy. FEBS Letters (1974) 55-58 Chen D and Santore MM. 1,2-Dipalmitoyl-sn-glycero-3-phosphocholine (DPPC)-Rich Domain Formation in Binary Phospholipid Vesicle Membranes: Two- Dimensional Nucleation and Growth. Lamgmuir 30 (2014) 9484-9493 Graber ZT and Kooijman EE. Ionization Behavior of Polyphosphoinositides Determined via the Preparation of pH Titration Curves Using Solid-State 31 P NMR. Plant Lipid signaling Protocols, Methods in Molecular Biology vol. 1009 (2013) 129-142 Graber ZT, Gericke A, Kooijman EE. Phosphatidylinositol-4,5-bisphosphate ionization in the presence of cholesterol, calcium or magnesium ions. Chemistry and Physics of Lipids 182 (2014) 62–72 Graber ZT. Electrostatics and binding properties of Phosphatidylinositol-4,5-bisphosphate in model membranes. PhD Dissertation, Kent State University (2014) Ishiguro K, Ando T, Watanabe O, and Goto H. Novel application of low pH-dependent fluorescent dyes to examine colitis. Gastroenterology 10:4 (2010) Jorge J, Castro GR, and Martines MA. Comparison among Different pH Values of Rhodamine B Solution Impregnated into Mesoporous Silica. Orbital: The Electronic Journal of Chemistry vol 5 (2013) Kaiser RD and London E. Determination of the depth of BODIPY probes in model membranes by parallax analysis of fluorescence quenching. Biochimica et Biophysica Acta 1375 (1998) 13-22 Kooijman EE, Carter KM, van Laar EG, Chupin V, Burger K, and Kruijff B. What Makes the Bioactive Lipids Phosphatidic Acid and Lysophosphatidic Acid So Special? Biochemistry 44 (2005) 17007-17015

Kooijman EE, King KE, Gangoda M, and Gericke A. Ionization Properties of Phosphatidylinositol Polyphosphates in Mixed Model Membranes. Biochemistry 48 (2009a) 9360–9371 Kooijman EE, Vaknin D, Bu W, Joshi L, Kang SW, Gericke A, Mann EK, and Kumar S. Structure of Ceramide-1-Phosphate at the Air-Water Solution Interface in the Absence and Presence of Ca 2+. Biophysical Journal 96 (2009b) 2204-2215 Kooijman EE. What makes the simple and crucial lipids lysophosphatidic acid and phoshatidic acid so special? PhD Thesis, Utrecht University (2006) Kuhl O. Phosphorus-31 NMR Spectroscopy. Springer, Berlin (2008) Meleard P, Bagatolli LA, and Pott T. Giant unilamellar vesicle electroformation from lipid mixtures to native membranes under physiological conditions.Methods in Enzymology 465 (2009) 161-176 Rouser G, Fkeischer S, and Yamamoto A. Two dimensional then layer chromatographic separation of polar lipids and determination of phospholipids by phosphorus analysis of spots. Lipids 5(5) (1970) 494-496 Shimokawa N, Nagata M, and Takagi M. Physical properties of the hybrid lipid POPC on micrometer-sized domains in mixed lipid membranes. Physical Chemistry and Chemical Physics. 17 (2015)20882--20888 Swairjo MA, Seaton BA, and Roberts MF. Effect of vesicle composition and curvature on the dissociation of phosphatidic acid in small unilamellar vesicles - a 31PNMR study. Biochimica et Biophysica Acta 1191 (1994) 354-361 Tanaka T, Sano R, Yamashita Y, and Yamazaki M. Shape Changes and Vesicle Fission of Giant Unilamellar Vesicles of Liquid-Ordered Phase Membrane Induced by Lysophosphatidylcholine. Langmuir 20 (2004) 9526-9534 Veatch SL. Electro-formation and fluorescence microscopy of giant vesicles with coexisting liquid phases. Methods Molecular biology 398 (2007) 59-72 Villasusoa AL, Wilke N, Maggio B, and Machado E. The surface organization of diacylglycerol pyrophosphate and its interaction with phosphatidic acid at the air–water interface. Chemistry and Physics of Lipids 163 (2010) 771–777 Watts A. Solid-state NMR approaches for studying the interactions of peptides and proteins with membranes. Biochimica et Biophysica Acta 1376 (1998) 297-318

Chapter 3

Ionization of Diacylglycerol-pyrophosphate in Model Membranes

Introduction

Diacylglycerol-pyrophosphate (DGPP) is a minor lipid found in the inner leaflet of the plasma membrane (Wissing and Behrbohm 1993, Munnik et al 1996). It has been identified in plants and yeast, but has not been observed in mammals. It is created from phosphatidic acid

(PA) in the membrane during stress conditions such as hyper-osmolarity (Munnik et al. 2000), however the exact enzyme that accomplishes this has not yet been identified. Since DGPP and

PA share very similar structures (see figure 3-1), it is possible that they share similar functions in the plasma membrane.

Figure 3-1: The structures of phosphatidic acid (A) and diacylgycerol-pyrophosphate (B). Shown are the DO, i.e. carrying two oleic acid acyl chains, species. These lipids share a similar structure, except for the pyrophosphate present in DGPP. The phospho-monoester groups for both lipids (outer most phosphate for DGPP) show similar titration behavior in model membranes (Strawn et al 2012, Kooijman et al 2005).

The ionization behavior of PA in model systems has been well characterized (Kooijman and Burger 2009, Kooijman et al 2005) and the electrostatic hydrogen bond switch model has been proposed to explain the possible function of PA within the membrane, which is described in more detail in chapter 1 (Kooijman et al. 2007). This model states that the phospho- monoester of PA can form hydrogen bond interactions with cationic protein residues (see figure

3-2), which will lower the total charge of PA (more negative). This lowered charge will then increase the affinity of the binding protein to the phospho-monoester, which can serve as a basis for PA specific protein recognition. Since DGPP also contains a phospho-monoester group, the possibility exists that it can participate in similar interactions.

Figure 3-2: The electrostatic hydrogen bond switch model. PA can form a hydrogen bond with a cationic protein residue or another lipid, such as PE, which will cause it to deprotonate. This model is discussed in more detail in chapter 1 as well as the discussion section of this chapter.

The ionization behavior of DGPP has previously been explored in simple mixtures with phosphatidylcholine (PC) as well as more complex mixtures containing the hydrogen-bond donor lipid phosphatidylethanolamine (PE) (Strawn et al. 2012) (see figure 3-3). The ionization behavior of the DGPP phospho-monoester was found to be similar to the ionization of the PA phospho-monoester in simple mixtures, and the effect of PE on ionization was also found to be similar. In both cases, PE increases the charge of the phospho-monoester.

Figure 3-3: Ionization of DGPP with or without phosphatidylethanolamine present. PE shows a clear effect on DGPP titration. PE can form hydrogen bond interactions with DGPP, increasing the charge of the phospho-monoester. The vertical black bar is drawn through the pH value of simple PC mixtures corresponding to the pKa of the phospho-monoester and shows that the presence of PE increases the deprotonation of DGPP phospho-monoester at the same pH value. DGPP present at 5 mol%. Taken from Strawn et al 2012. 46

Since DGPP and PA are typically found together in-vivo, the question of their possible interaction with each other arose. Both DGPP and PA are able to interact with PE via hydrogen bonds in the membrane, but they also have the possibility to interact with each other. These interactions could play a significant role in stress signaling by altering how DGPP and PA interact with proteins. One example would be the formation of lipid domains. Lipid domains are defined as functional units within the plasma membrane that arise from the clustering of lipids, which are hypothesized to play a role in lipid-protein interactions and signal transduction (Simons and

Ikonen, 1997). Lipid domains have been observed in model mixtures containing phospholipids and cholesterol that are believed to be acyl-chain mediated (Shimokawa et al 2015, Chen and

Santore 2014). Domains are also believed to form via head group mediated interactions and have been observed in mixtures containing lipids that can participate in hydrogen bonding, such as phosphatidylinositol-4,5-bisphosphate (Graber et al. 2012).

Results

The ionization behavior of DGPP was probed via 31P NMR. DGPP was present in mixtures with phosphatidylcholine (PC) and phosphatidic acid (PA), as well as with/without phosphatidylethanolamine (PE). Titration curves were constructed over the pH range of approximately 4-11

(in red), and compared with data previously gathered (in blue). DGPP titration was compared with Strawn et al 2012, and PA titration was compared to Kooijman et al 2005 as well as unpublished data. The formation of lipid domains was probed using fluorescence microscopy of giant unilamellar vesicles (GUV’s) containing PC, DGPP, and PA. GUV’s were observed at pH values of 4, 7, and 9 to determine the effect of lipid ionization on domain formation. 47

Outer Phosphate of DGPP Exhibits Similar Titration Behavior to PA

Figure 3-4: Ionization behavior of DGPP and PA phospho-monoesters. A water fall plot (A) shows the raw NMR data for a mixture of 90 mol% PC, 5 mol% DGPP, and 5 mol% PA. This data was used to construct titration curves for the DGPP (B) and PA (A) phospho-monoesters that are compared to simple mixtures with only PC. Data for comparison taken from Strawn et al 2012 (DGPP) and Kooijman et al 2005 (PA) shown in blue. Chemical shift values are relative to an external 85% H3PO4 standard.

In order to determine the effect that PA would have on the ionization of DGPP and vice versa, titration curves for the phospho-monoesters of DGPP and PA were compared to previously gathered data (Strawn et al 2012, Kooijman et al 2005). The phospho-monoester

(outer) of DGPP exhibited sigmoidal titration behavior while the phospho-diester (inner) did not titrate much at all (figure 3-4a). When the titration of the DGPP phospho-monoester was compared to mixtures not containing PA (figure 3-4b), no difference in titration behavior was observed. Similarly, when the titration of the PA phospho-monoester was compared to mixtures not containing DGPP (figure 3-4c), no difference in titration behavior was observed.

Next, NMR experiments were carried out for lipid mixtures containing the hydrogen bond donor lipid PE. This data was compared to previous experiments that only contained either DGPP or PA in mixture with PC and PE (shown in blue). Figure 3-5 shows that mixtures with PE showed no deviation from previous data. This means that when DGPP and PA are present in mixtures seems to have no significant effect on the ionization of either phospho- monoester. It also appears that the overall effect of PE on the ionization of DGPP is no different with or without PA present (figure 3-5b).

Figure 3-5 (next page): Ionization of DGPP and PA phospho-monoesters in the presence of PE. A waterfall plot (A) is shown for a lipid mixture containing 2.5 mol% DGPP and 2.5 mol% PA. The remaining 92.5 mol% consisted of equal molar amounts of PC and PE. Titration curves are shown for the DGPP phospho-monoester (B) and the PA phospho-monoester (C) that are compared to mixtures containing PE. Titration curves show a lipid mixture containing 5 mol% DGPP and 5 mol% PA. Reference data shown for DGPP and PA at 5 mol% shown in blue. DGPP reference data taken from Strawn et al 2012 and PA reference data unpublished (shown in appendix). All chemical shift values are relative to an external 85% H3PO4 standard.

Population of DGPP Inner Phosphate Groups Participate in Local Electrostatic

Interactions

NMR peak splitting was also observed during experiments. Peak splitting was observed as a phosphate group titrating normally, with the appearance of a peak around the chemical

50 shift value of the protonated species at specific pH values. Splitting was observed between pH values of 6 and 8.6 for both PA and DGPP. Titration experiments were run for DGPP and PA at 5 mol% as well as 2.5 mol%. At 5 mol%, splitting is observed for the PA phospho-monoester as well as the DGPP phospho-diester (inner phosphate) (see figure 3-6). At 2.5 mol%, splitting was only observed for the DGPP phospho-diester. The peak splitting intensity seems to be dependent on the concentration of DGPP and PA present in the mixture, as more splitting intensity was observed when both lipids were present at 5 mol%. The absence of peak splitting for the DGPP phospho-monoester seems to indicate that there is an interaction occurring between PA and the inner phosphate (phospho-diester) of DGPP.

Figure 3-6: NMR peak splitting occurs at higher concentrations of DGPP. Peak splitting occurred at intermediate pH (6-8.6) and is indicated with arrows. The higher concentration of DGPP shows more intense splitting which is indicative of a concentration dependent interaction. PA present at same amount as DGPP for each mixture. All chemical shift values relative to an external 85% H3PO4 standard. 51

Domain Formation in Giant Unilamellar Vesicles Occurs at Physiological pH

This observed peak splitting in the NMR experiments indicated a possible hydrogen bond formation between PA and the inner phosphate of DGPP, and was further probed with the use of giant unilamellar vesicles. Hydrogen bonding between the two lipids has the potential to form DGPP/PA rich clusters in the membrane, which could form observable macro- scopic domains. Domains can be visualized using fluorescence microscopy if the fluorescent dye used is specifically excluded from the domain forming portion of the membrane. A fluorescently labelled PA (NBD) was initially used to visualize domains, which were seen as areas of fluorophore exclusion, altered membrane curvature, or both. NBD-PA is an acyl-chain labelled dye, therefore its head group still retains all of the chemical property of unlabeled PA. Domains were mainly observed at pH 6, with little to no domains observed at pH 4 or 9 (see figure 3-7).

Approximately 40% of vesicles present in solution were GUV’s, with approximately 20% of those vesicles showing domains at physiological pH and room temperature. The majority of vesicles formed during the electroformation procedure were multi-lamellar vesicles (MLV’s).

This could be due to the nature of the formation protocol as vesicles tended to form in “grape” bunches. Once GUV’s were formed, they would either break off into the buffer solution or would fuse with other surrounding vesicles. Vesicles were formed in a buffer containing 100 mM sucrose, and then were developed in a buffer that contained 100 mM Glucose so that they would settle to the bottom of the viewing plate. Vesicles chosen to be imaged were stationary and showed an adequate fluorescence intensity to produce a resolved image.

Figure 3-7: Domain formation in vesicles containing DGPP and PA with NBD- PA. At pH 4 and 9, no domains were observed. At pH 6, domains were present as altered curvature in the membrane. Lipid mixture shown is 75 mol% PC, 20 mol% DGPP, and 5 mol% PA. NBD- PA was used as a fluorophore and is present at 0.2 mol%. Approximately 20% of GUV’s showed domain formation. Images indicate the best representative image for each experiment.

Due to the possibility of domain formation only occurring due to the nature of the fluorophore, a second fluorophore (Rhodamine-B) was used. Rhodamine-B is a head group labelled derivative of phosphatidylethanolamine. Domains were also observed at pH 6 with no domains present at pH 9 (see figure 3-8). These observations were consistent with the NMR peak splitting and further indicated an interaction between DGPP and PA.

Figure 3-8: domain formation in vesicles containing DGPP and PA with Rhodamine-B. Domains were observed at pH 6 as altered membrane curvature or fluorophore exclusion. At pH 9 no domains were observed. Lipid mixture shown is 75 mol% PC, 20 mol% DGPP, and 5 mol% PA. Rhodamine-B is used as a fluorophore and is present at 0.2 mol%. Domains were present in approximately 20% of all GUV’s.

Discussion

The ionization data gathered for DGPP indicates that the phospho-monoester behaves similar to the phospho-monoester of PA. The fitting of the titration curves using a Henderson-

Hassalbach equation allows for the pKa value of the deprotonation process to be obtained for each different lipid mixture. The pKa value for DGPP in mixtures with PC and PA is identical to the pKa for DGPP in just PC (see table 3-9). The pKa of PA in mixtures with DGPP and PC is also comparable to the pKa of PA in just PC (8.07 and 8.01 respectively). PE decreased the pKa of both DGPP (from 7.44 to 6.73) and PA (from 8.07 to 7.18) when they were present together, but this was the same magnitude of difference in pKa as was observed when DGPP (from 7.44 to 6.71) and PA (from 8.01 to 7.19) were not present together. Both lipids are able to form hydrogen bonds with PE within the membrane, which will increase their total negative charge.

Based on the data it does not appear that the phospho-monoesters of DGPP and PA form any interaction with each other. They titrate in the same way as they would in mixtures separate from each other, which is seen in the super-position of titration curves in figures 3-4 and 3-5. If the phospho-monoesters of DGPP and PA were forming interactions with each other, then a significant change in the pKa value would be observed.

Lipid Mixture Calculated pKa

PC:DGPP** DGPP= 7.44 (+/- .02)

PC:PA* PA= 8.01 (+/- .03)

PC:PE:DGPP** DGPP= 6.71 (+/- .02)

PC:PE:PA* PA=7.19 (+/- .02)

PC:DGPP:PA DGPP= 7.44 (+/- .04)

PA= 8.07 (+/- .05)

PC:PE:DGPP:PA DGPP= 6.73 (+/- .07)

PA= 7.18 (+/- .07)

Table 3-9: pKa values of the DGPP and PA phospho-monoesters under different conditions. DGPP and PA present at 5 mol%. PC and PE present in equal amounts when listed together. Mixtures containing both DGPP and PA have similar pKa values to simple mixtures only containing either DGPP or PA. ** taken from Strawn et al 2012 and * taken from unpublished data (see appendix).

However, the presence of peak splitting indicates an interaction between PA and the

DGPP phopho-diester. This interaction could be the driving force behind the domains that are observed at physiological pH. The DGPP phospho-diester does not titrate over the range tested, and previous work has argued that it would remain protonated until very high pH values

(Strawn et al. 2012 and unpublished data). Despite this, DGPP remains highly anionic with either a -1 or -2 charge at physiological pH. At physiological pH, DGPP and PA could be tightly clustered, which would explain the apparent fluorophore exclusion in the observed domains. 55

The fluorophores used are bulky and could be sterically pushed out from these regions due to their preference for disordered environments (Sengupta et al. 2009). An illustration of the possible hydrogen bond is shown below.

Figure 3-10: Hydrogen bonding between DGPP and PA. The phospho-monoester of PA and the phospho-diester of DGPP are capable of forming hydrogen bond interactions within the membrane. These interactions could create DGPP/PA clusters in the membrane that could play a role in cellular signaling. At high pH, the phospho-monoesters will become deprotonated and repel each other.

This hydrogen bonding interaction between DGPP and PA can have a significant impact on signaling within the membrane. Since DGPP and PA are typically observed together during stress signaling, it is believed that DGPP acts to attenuate the PA signal in the membrane (van

Schooten et al 2005, Villasuso et al 2014). The electrostatic hydrogen bond switch model has

56 previously been proposed to explain how PA interacts with its protein targets (Kooijman et al.

2007). This model states that the hydrogen bonding of PA with cationic protein residues will lower its overall charge which will in turn increase its binding affinity, and can serve as a basis for binding specificity. The ionization behavior of the DGPP phospho-monoester hints towards a similar behavior. PA can recruit cationic proteins to the membrane based on its negative charge, but it is not known if DGPP recruits proteins in the same way. DGPP and PA have a different effective molecular shape (Strawn et al. 2012) and the phospho-monoester of DGPP has a higher overall negative charge than the phospho-monoester of PA at physiological pH (see table 3-9). DGPP also has a different location of the phospho-monoester relative to the bilayer/cytoplasm interface (Strawn et al. 2012), so it is possible that it can recruit proteins by altering the membrane organization and shape.

The observation of domain formation in GUV’s also raises many questions about the ability of DGPP to alter the local shape of the membrane. Previous work with DGPP and PA showed a condensed molecular organization in monolayers at low pH (Villasuso et al. 2010). At this low pH, DGPP and PA are still protonated and can hydrogen bond with each other. At higher pH they will both be deprotonated, and their respective negative charges will repel each other. This was observed by Villasuso et al. at pH 8, where an expanded monolayer was formed.

Indeed, at higher pH values domain formation was not observed in GUV’s. This could be due to the same charge repulsion which will overcome any hydrogen bonding. It is possible then that upon the formation of DGPP in the membrane, it will form clusters with PA that can recruit target proteins to the membrane. Upon binding of the target protein to DGPP or PA, the charge

57 of the phospho-monoester will become more negative, which would cause the lipid/protein complex to dissociate from the lipid raft/domain. More work will need to be done to explore the exact nature of the domains that are formed in the membrane to elucidate exactly how they may function in stress signaling.

References

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Kooijman EE and Burger KN. Biophysics and function of phosphatidic acid: A molecular perspective. Biochimica et Biophysica Acta 1791 (2009) 881–888

Kooijman EE, Carter KM, Emma, van Laar G, Chupin V, Burger KN, and de Kruijff B. What Makes the Bioactive Lipids Phosphatidic Acid and Lysophosphatidic Acid So Special? Biochemistry 44 (2005) 17007-17015

Kooijman, EE, Tieleman, DP, Testerink, C, Munnik, T, Rijkers, DT, Burger, KN, de Kruijff B. An electrostatic/hydrogen bond switch as the basis for the specific interaction of phosphatidic acid with proteins. Journal of Biological Chemistry 282 (2007) 11356– 11364.

Munnik T, de Vrije T, Irvine RF, and Musgrave A. Identification of Diacylglycerol Pyrophosphate as a Novel Metabolic Product of Phosphatidic Acid during G-protein Activation in Plants. The Journal of Biological Chemistry 271 (1996) 15708-15715

Munnik T, Meijer HJ, Riet B, Hirt H, Frank W, Bartels B, and Musgrave A. Hyperosmotic stress stimulates Phospholipase D activity and elevates the levels of phosphatidic acid and diacyglycerol pyrophosphate. The Plant Journal 22(2) (2000) 147-154

Sengupta P, Hammond A, Holowka D, and Baird B. Structural Determinants for Partitioning of Lipids and Proteins Between Coexisting Fluid Phases in Giant Plasma Membrane Vesicles. Biochimica et Biophysica Acta 1778 (2009) 20-32

Shimokawa N, Nagata M, and Takagi M. Physical properties of the hybrid lipid POPC on micrometer-sized domains in mixed lipid membranes. Physical Chemistry and Chemical Physics 17 (2015) 20882-20888

Simons K and Ikonen E. Functional rafts in cell membranes. Nature 387 (1997) 569-572

Strawn L, Babb A, Testerink C, and Kooijman EE. The physical chemistry of the enigmatic phospholipid diacylglycerol pyrophosphate. Frontiers in Plant Science 3 (2012) van Schooten B, Testerink C, and Munnik T. Signalling diacylglycerol pyrophosphate, a new phosphatidic acid metabolite. Biochimica et Biophysica Acta 1761 (2006) 151–159

Villasuso AL, Wilke N, Maggio B, and Machado E. The surface organization of diacylglycerol pyrophosphate and its interaction with phosphatidic acid at the air–water interface. Chemistry and Physics of Lipids 163 (2010) 771–777

Villasuso AL, Wilke N, Maggio B, and Machado E. Zn2+ dependent surface behavior of diacylglycerol pyrophosphate and its mixtures with phosphatidic acid at different pHs. Frontiers in Plant Science 5 (2014)

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Chapter 4

Ionization of Phosphatidylinositol 3,4,5-trisphosphate in Model Membranes

Introduction

Phosphatidylinositol 3,4,5-trisphosphate (PIP3) is present in small amounts in the inner leaflet of the animal plasma membrane, but it plays a crucial role in cell signaling (Balla, 2013).

The enzyme PI3K produces PIP3 from phosphatidylinositol 4,5-bisphosphate (PIP2), promoting cell growth and proliferation (Lindmo and Stenmark, 2006). Alternately, the tumor suppressor

PTEN will convert PIP3 back to PIP2, limiting cell growth (Gericke et al. 2006). All of this is accomplished by the recognition of PIP3 by various signaling proteins, which are able to detect and bind to the PIP3 head group via electrostatic interactions (Catimel at al. 2009). Due to the importance of PIP3 in these cellular signaling pathways, it is crucial to understand the mechanisms that influence how PIP3 will interact with its protein targets.

The head group of PIP3 contains 3 phospho-monoester groups that have the potential to interact with each other (see figure 4-1). This complex behavior has the potential to influence electrostatic interactions between PIP3 and proteins. Previous NMR studies have examined PIP3 in simple mixtures with phosphatidylcholine (PC) and have in fact shown that the phospho- monoesters do not exhibit sigmoidal titration behavior (Kooijman et al. 2009). The various phospho-monoesters are identified based on their relative positions in the NMR spectra (See 61 figure 4-2). The PC peak is the most downfield peak with the highest intensity. The 3-phosphtae of PIP3 is identified as the most downfield of the 3 smaller peaks at lower pH. The 5-phosphate is the intermediate peak, and the 4-phosphate is the most upfield peak at low pH. Peak assignments are further explained in Kooijman et al. 2009 and follow the assignments for PIP2 and other phosphatidylinositol-phosphates. The 3 and 5-phosphates show a biphasic titration behavior, whereas the 4-phosphate shows the most striking titration behavior. The 4- phosphate shows an increase in charge as the pH increases from around 4-6, but then shows a decrease around pH 7, finally increasing again from pH 9-11 (see figure 4-2). This causes the 4- phosphate to “cross-over” the 3 and 5-phosphates, making it the most downfield peak at high pH.

Figure 4-1: Head groups of phosphatidylinositol polyphosphates. Phosphatidylinositol 3,4,5- trisphosphate is shown on top. Phosphatidylinositol-bishosphate can have three potential structures, depending on the location of the phosphate groups on the inositol ring. Only the PIP2 with phosphorylations at the 4 and 5 position is discussed in this thesis. Figure taken from Kooijman et al. 2009 62

These titration behaviors are indicative of intra-molecular interactions within the head group itself (Schlewer et al. 1998). The close proximity of the phospho-monoesters allows for possible hydrogen bonding, which could help to stabilize negative charges as PIP3 becomes increasingly deprotonated. This work only had PIP3 present with PC, but the inner leaflet of the plasma membrane contains other phosphor-lipid groups the potential to form electrostatic and hydrogen bond interactions. Lipids such as phosphatidylethanolamine (PE) and phosphatidylinositol (PI) have been shown to form hydrogen bonds with phosphatidic acid and

PIP2 in model membranes, altering their ionization behavior (Graber et al 2012, Kooijman et al

2005).

Figure 4-2: Titration behavior of PIP3 in PC Vesicles. PIP3 exhibits complex titration behavior in simple multi-lamellar vesicles. (A) shows a waterfall plot of raw data that was used to construct titration curves for each phosphate group (B). Phospho-monoester peaks are assigned by their relative chemical shifts. The 3 and 5-phosphates (P3, P5 respectively) show a biphasic behavior with a deviation from standard sigmoidal behavior between pH 9 and 10. The 4-phosphate (P4) shows the most complex behavior, with a decrease in chemical shift as the pH gets higher within the physiological range (pH 6.5-8) Lipid mixture shown is 95 mol% DOPC and 5 mol% PIP3. Data taken from Kooijman et al. 2009.

Previous work with PI(4,5)P2 also showed biphasic behavior for the 4 and 5-phosphates 63

In simple mixtures with PC (Kooijman et al. 2009). PIP2 lacks a 3-phosphate, so the 4-phosphate is only flanked by a phosphate and a hydroxyl group (see figure 4-1). The presence of lipids capable of forming hydrogen bonds in complex mixtures altered the charge/titration behavior of the phospho-monoesters. Phosphatidylethanolamine (PE) increased the overall ionization of the PIP2 head group over the entire tested pH range of 4-10, while phosphatidylinositol (PI) only seemed to decrease the ionization beyond pH 7 (Graber et al. 2012). Due to the structural similarities of PIP2 and PIP3, we hypothesized that these hydrogen bond donor lipids would exhibit similar effects on the PIP3 head group, and could have an effect on the titration behavior of the individual phospho-monoesters. Both the 3 and 5-phosphates are adjacent to the 4- phosphate as well hydroxyl groups on the inositol ring, with the 2-hydroxyl equatorial and the

6-hydroxyl axial. The close proximity of all of these groups results in many different possible intra-molecular hydrogen bond interactions that have the potential to compete with inter- molecular hydrogen bond interactions.

The work in this thesis used the previous results as a foundation, and aimed to determine the ionization behavior of PIP3 in complex model membranes containing hydrogen bond donor lipids. The complex mixtures included either PE or PI along with PC and PIP3. PIP3 was present in small amounts (2 mol% for PI mixtures and 5 mol% for PE mixtures) due to its low natural abundance (< 1%) (Balla, 2013).

Results

31 The ionization behavior of PIP3 in mixed vesicles was probed using P NMR. Data

64 collected was then compared against data previously collected for PIP3 in simple mixtures

(Figure 4-2) (Kooijman et al. 2009). This was done to determine the effect that hydrogen bond donor lipids would have on the charge/ionization of PIP3. Data was fit by a previous PhD student to determine the total charge of PIP3 as a function of pH, using a model developed previously (Graber, 2014). This fitted data was then used to determine the total charge of PIP3 in each mixture. The calculated total charge was compared to the calculated total charge in simple mixtures containing just PC, to determine the effect of hydrogen bond donor lipids on the total charge of PIP3 as a function of pH. The total charge includes the charges of the three phospho-monoesters as well as the -1 charge of the phospho-diester. The possibility of hydrogen bonding interactions between PI and PIP3 was probed via fluorescence microscopy.

Phosphatidylethanolamine promotes increased deprotonation of PIP3

The phospho-monoesters of PIP3 exhibit complex titration behavior while in mixtures with PE (see figure 4-3). The 3 and 5-phosphates exhibit biphasic titration behavior, and the 4- phosphate exhibits a similar decrease in chemical shift between pH 6.5 and 9 that was observed in the previous PIP3 data (Figure 4-3b). The total charge of PIP3 in mixtures with PE also shows marked differences from the total charge in just PC (Figure 4-3c). From pH 4 to approximately pH 7 the presence of PE increases the total charge on PIP3, which is indicative of hydrogen bond induced deprotonation. Between pH 7 and pH 9, the total charge of PIP3 in PE mixtures appears to be identical to the total charge in PC mixtures. Finally, as the pH increases past 9 the total charge of PIP3 actually decreases when compared against the PC data.

Figure 4-3: Ionization behavior of PIP3 in mixed vesicles containing PE. Chemical shift values from each NMR experiment (A) are used to construct a titration curve for each of the phosphates of PIP3 (B). Peaks are assigned based on their relative chemical shifts as described in Kooijman et al 2009. The total charge of PIP3 was calculated and compared to the total charge in simple mixtures with just PC (C) using a model developed by Graber, 2014. Mixtures contained 47.5 mol% PC, 47.5 mol% PE, and 5 mol% PIP3. Chemical shift values are relative to an external 85% H3PO4 standard.

Phosphatidylinositol reduces the overall charge of PIP3

In mixtures with PI, the 3 and 5-phosphates also exhibited biphasic titration behavior.

The 4-phosphate experienced the same decrease in chemical shift as the PC and PE mixtures around physiological pH (Figure 4-4b). The total charge of PIP3 in mixtures with PI shows significant differences from the total charge in just PC (Figure 4-4c). As the pH increases above

7, the total charge of PIP3 is reduced (more negative) compared to the total charge in just PC.

Figure 4-4: Ionization behavior of PIP3 in mixed vesicles containing PI. Chemical shift values from each NMR experiment (A) are used to construct a titration curve for each of the phosphates of PIP3 (B). Peaks are assigned based on their relative chemical shifts as described in Kooijman et al 2009. The total charge of PIP3 was calculated and compared to the total charge in simple mixtures with just PC (C) using a model developed by Graber, 2014. Due to low abundance of PIP3 in these mixtures, Y-axis of waterfall plot is truncated. Peaks are assigned based on their relative chemical shifts as described in Kooijman et al 2009. Mixtures contained 88 mol% PC, 10 mol% PI, and 2 mol% PIP3. Chemical shift values are relative to an 85% H3PO4 standard.

PIP3 Promotes Domain Formation in Mixed Lipid Vesicles Containing Phosphatidyl- inositol

Figure 4-5: Giant unilamellar vesicles containing PI and PIP3. All mixtures contained 20 mol% PI along with indicated amounts of PIP3. The remaining portion of lipid for each mixture was composed of POPC (16:0-18:1 acyl chains). Control mixtures (0% PIP3), containing 80 mol% POPC and 20 mol% PI, did not show any domain formation. As the amount of PIP3 in the vesicles increased, the average size of membrane domains also increased. Pictures shown represent the average domain size for each mixture. Vesicles were viewed at room temperature in pH 7 buffer.

Since PI was found to interact with PIP3 altering its ionization, it was hypothesized that these interactions may lead to the formation of domains within the plasma membrane.

Previous studies involving PI(4,5)P2 in giant unilamellar vesicles (GUV’s) with PI also showed 68 domain formation in the form of membrane bulging (Graber et al. 2012). Domain formation was probed with fluorescence microscopy of GUV’s containing POPC, PI, PIP3, and Rhodamine- phosphatidylethanolamine as a dye. The amount of PI was held constant at 20 mol% and the amount of PIP3 was varied. Vesicles that only contained POPC and PI did not show any observable domain formation. Vesicles that contained 5% PIP3 showed domains as small bulges in the membrane with a slight exclusion of the lipid dye. As the amount of PIP3 increased to

10% domains became noticeably larger, and at 20% PIP3 the domains comprised a sizeable portion of each vesicle. Approximately 30-40% of GUV’s that were generated exhibited domain formation in appropriate mixtures.

Discussion

The presence of hydrogen bond donor lipids has a significant impact on the ionization of

PIP3 in model membranes. These complex model mixtures do not contain the complexity of an actual plasma membrane, but these results do offer some insight into how PIP3 behaves in-vivo.

There are two main interactions that can arise in the mixtures tested. PE contains a terminal amine that has the potential to act as a hydrogen bond donor, and the phospho-monoesters of

PIP3 can act as hydrogen bond acceptors (see Figure 4-5a). PI contains hydroxyl groups that also have the potential to act as hydrogen bond donors (see Figure 4-5b). It is also important to consider charge repulsions between phospholipids that may occur as they begin to titrate. PIP3 is a highly anionic lipid, with a charge of around -5 at pH 7 (Kooijman et al. 2009). As this negative charge increases, it can be repelled from other anionic lipids, like PI. PE also has the potential to titrate at higher pH with the dissociation of a proton from the terminal amine 69

(Akoka et al. 1986). This would give PE an overall negative charge, repelling it from other anionic lipids in the membrane.

Figure 4-6: Hydrogen bond model for PIP3 with PE and PI. PE contains a terminal amine that can act as a hydrogen bond donor (A). PI consists of an inositol ring covered in hydroxyl groups. These hydroxyl groups can form hydrogen bonds with the phospho-monoesters of PIP3 (B).

The experimental results offer insight into how electrostatic and hydrogen bond interactions alter the ionization of PIP3. In mixtures with PE, the total charge of PIP3 is increased at low pH. A hydrogen bond between PE and PIP3 would act to stabilize a more negative charge on the phospho-monoesters as they titrate. Between pH 7 and 9, the charge on PIP3 shows no difference with the PC data. One possible explanation for this is that PE begins to show a more negative charge, which causes the hydrogen bond interaction to compete with an electrostatic charge repulsion. As the pH continues to rise this electrostatic interaction wins out, actually increasing the charge on PIP3. This increased protonation contrasts with the results obtained

70 previously with PIP2 (Graber 2012, 2014). In these experiments PE increased the total negative charge of PIP3 until pH 7. At pH >7 no difference in charge was observed due to the presence of

PE at high pH. A possible explanation for this is that PIP3 is considerably more negatively charged than PIP2 meaning that charge repulsions will be much stronger, allowing them to overcome attractive hydrogen bond interactions. Another consideration is the increase of total negative charge in the membrane as PE and PIP3 are deprotonated. This increase in negative charge will recruit protons from the bulk solution to the membrane interface. This increase in protons at the interface will lower the local pH, which will lower the total charge on PIP3. The extra decrease in charge at higher pH for the PIP3 experiments implicates the 3-phosphate as possibly playing a crucial role in these interactions, which merits further study. Since PE had no effect on PIP3 charge at physiological pH it is difficult to speculate if this interaction will alter how proteins recognize and interact with PIP3. It is possible that the interplay of hydrogen bonding and charge repulsion can allow PE to interact with PIP3 without significantly altering its charge. It is then possible that a protein could selectively recognize the PE/PIP3 complex.

The presence of PI in the membrane does not seem to have any significant effect on the total charge of PIP3 at low pH. This seems counterintuitive since PI can hydrogen bond with the

PIP3 phospho-monoesters spreading out the total charge. However, the presence of PI increases the total negative charge in the membrane, which can increase the interfacial proton concentration in much the same way as the deprotonation of PE previously discussed. It is possible that the competing effects of hydrogen bonding and increased negative charge due to

PI are cancelling each other out, not having a significant effect on PIP3’s ionization. As the pH

71 increases above 7, a drastic decrease in total charge is seen. This is indicative of charge interactions taking over, as the PIP3 phospho-monoesters begin to show a more negative charge. The hydrogen bond interactions would increase the deprotonation of PIP3 increasing the charge, while the accumulation of protons at the membrane interface will promote the protonation of PIP3 decreasing the charge. The decrease in charge becomes more pronounced at high pH, which could result from a higher recruitment of protons to the interface. Similar behavior was observed in PIP2 when mixed with PI. At low pH, no difference was seen for PIP2’s total charge when compared to simple mixtures, but at pH above 7 a decrease in charge was observed (Graber 2014).

The presence of the hydrogen bond interaction between PI and PIP3 was in fact observed in the GUV data. As the concentration of PIP3 in the mixtures was increased, the size of the domains proportionally increased. The absence of domains in the mixture lacking PIP3 confirms that the domains observed were not produced by interactions of PI with itself or as an artifact of the vesicle formation procedure itself. This is interesting due to the charge repulsion that is present between PI and PIP3. The GUV data was collected at pH 7 however, where the charge repulsion may still be modest. The competition between the hydrogen bond and electrostatic interactions could be further probed by attempting to observe domain formation at pH 9, where PIP3 is more highly charged. Previous studies have been conducted observing domain formation in mixed vesicles containing PI and PIP2 as well (Graber, 2012). These experiments saw a similar concentration-dependent domain formation, which supported the hypothesis that PIP2 and PI were in fact forming hydrogen bond interactions. Taken together, it

72 is reasonable to assume that in the plasma membrane PIP3 will form interactions with PI, altering its ionization behavior. This will alter the total charge on PIP3 and could possibly alter how it will interact with proteins.

References

Akoka S, Tellier C, Poignant S. Molecular order, dynamics, and ionization state of phosphatidylethanolamine bilayers as studied by 15N NMR. Biochemistry 25(22) 1986 6972-6977 Balla T. Phosphoinositides: tiny lipids with giant impact on cell regulation. Physiological Reviews 93(3) (2013) 1019-10137 Catimel B, Yin MX, Schieber C, Condron M, Patsiouras H, Catimel J, Robinson DE, Wong LS, Nice EC, Holmes AB, and Burgess AW. PI(3,4,5)P3 Interactome. Journal of Proteome Research 8 (2009) 3712-3726 Gericke A, Munson M, and Ross AH. Regulation of the PTEN phosphatase. Gene 374 (2006) 1–9 Graber ZT, Jiang Z, Gericke A, and Kooijman EE. Phosphatidylinositol-4,5-bisphosphate ionization and domain formation in the presence of lipids with hydrogen bond donor capabilities. Chemistry and Physics of Lipids 165 (2012) 696– 704 Graber ZT. Electrostatics and binding properties of Phosphatidylinositol-4,5-bisphosphate in model membranes. PhD dissertation, Kent state University (2014) Kooijman EE, Carter KM, Laar EG, Chupin V, Burger KN, and Kruijff B. What Makes the Bioactive Lipids Phosphatidic Acid and Lysophosphatidic Acid So Special? Biochemistry 44 (2005) 17007-17015 Kooijman, E. E., King, K. E., Gangoda, M. & Gericke, A. Ionization properties of phosphatidylinositol polyphosphates in mixed model membranes. Biochemistry 48 (2009) 9360-9371 Lindmo K. and Stenmark H. Regulation of membrane traffic by phosphoinositide 3-kinases. Journal of Cell Science 119(4) (2006) Schlewer G, Guedat P, Ballereau S, Schmitt L, and Spiess B. Inositol Phosphates: Inframolecular physico-chemical studies: Correlation with binding properties in Phosphoinositides: Chemistry, Biochemistry, and Biomedical Applications. Bruzik KS Ed (1998) 255-270

Chapter 5

Discussion and Conclusions

Project summation

In this thesis I have explored the ionization behavior of diacylglycerol-pyrophosphate

(DGPP) and phosphatidylinositol 3,4,5-trisphosphate (PIP3). Complex mixtures were analyzed via 31P NMR and compared to data previously gathered for simple mixtures. This comparison was crucial to determine the effects that each phospholipid in the mixture had on either DGPP or PIP3. DGPP and phosphatidic acid (PA) were present together in complex mixtures to determine any interactions they may exhibit with each other. The phospho-monoesters of both lipids titrated independently of each other and behaved as had been previously observed when they were not mixed. The presence of phosphatidylethanolamine decreased the pKa values of

DGPP and PA, but this effect was no larger than had been previously observed for DGPP and PA alone (Strawn et al 2012 and unpublished data). Peak splitting was observed for the PA phospho-monoester and the DGPP phospho-diester, and was found to be dependent on the concentration of DGPP as well as pH. This was verified in giant unilamellar vesicles (GUV’s) where domains were observed at physiological pH, but not at high and low pH values. Domain formation was dependent upon the pH of the surrounding buffer similar to the peak splitting in

NMR experiments. This indicates that electrostatic interactions played a role in their formation.

At physiological pH, DGPP and PA are not fully deprotonated and can therefore hydrogen bond with each other. As the pH increases the lipids become fully deprotonated and become increasingly anionic. This increase in negative charge repels the head groups from each other and does not allow any hydrogen bonding (see figure 3-10). These domains can have a significant impact on how DGPP interacts with proteins and participates in signaling cascades in the membrane. DGPP is typically observed with increases of PA levels during cell stress (Munnik et al. 1996), so these domains could play a role in how cells adapt to external stresses.

Figure 5-1: Charge repulsion between PE and PIP3. At pH 7 (a) PIP3 and PE can form hydrogen bonds with each other. At high pH (b) both PIP3 and PE will deprotonate, causing them to have a more negative charge. These negative charges will repel the lipids from each other and break any hydrogen bonds that have formed.

PIP3 was analyzed in complex mixtures with either phosphatidylinositol (PI) or phosphatidylethanolamine (PE). These lipids are able to hydrogen bond with the PIP3 head 76 group, altering its total charge. At low pH PE increased the negative charge of PIP3 through hydrogen bonding. At intermediate pH this effect was not observed, and at high pH the total

negative charge of PIP3 was decreased. This data indicates two separate interactions that compete with each other within the membrane. Hydrogen bonding will increase the total negative charge, but as PIP3 and PE deprotonate more negative charges are present in the membrane. This recruits more protons to the membrane, lowering the local pH. This lower pH will decrease the negative charge of the phosphate groups present. As negative charges increase, PE and PIP3 will also be repelled from each other breaking any hydrogen bonds (see figure 5-1). These interactions are also present in mixtures with PI (see figure 5-2).

Figure 5-2: Charge repulsion between PI and PIP3. At pH 7 (a) PI and PIP3 will form hydrogen bonds with each other and can form domains within the membrane. PI is an anionic lipid, so as PIP3 deprotonates (b) it will be repelled. This repulsion will break any hydrogen bonds that have formed so domains will not form.

At low pH these interactions cancel each other out and the total charge of PIP3 is not affected. As the pH increases the charge-charge forces take over and the total negative charge of PIP3 is decreased. Domains in GUV’s are formed by hydrogen bond interactions at physiological pH and are dependent on the concentration of PIP3. This has implications on cell signaling, as PIP3 is believed to play an important role in cell proliferation and survival, membrane trafficking, and even cancer via the PTEN, mTOR and PI(3)K pathways (Lindmo and

Stenmark 2006, Shaw and Cantley 2006, Gericke et al 2006). The formation of these domains can alter how PIP3 is able to interact with the various signaling molecules involved in these pathways.

Overall, the work in this thesis demonstrates the complexity that exists in biological membranes. The model systems used only contained 3 or 4 lipid species, whereas the plasma membrane contains hundreds. The biochemical behavior of these lipids is the sum of all of the complex interactions that can occur in a given chemical environment. We are only beginning to fully understand the prominent role that biomembranes play in living systems. Membranes have gone from being seen as passive barriers to active components in the life of a cell.

Future Work

The ionization work with DGPP revealed an unexpected interaction between the PA phosho-monoester and the DGPP phospho-diester. This interaction should be probed in more depth to determine what function it could serve in the cell. The concentration dependence of the peak splitting in the NMR experiments could also be tested by conducting experiments with

DGPP present at 10, 15, and 20%. If peak splitting is observed at a higher intensity at these concentrations of DGPP than was observed for 5 and 10 mol%, then it would be clear that this splitting was due to an interaction between DGPP and PA. After that, the same experiments could be conducted while holding the concentration of PA constant at 5 mol%. In this case, the intensity of the peak splitting should saturate, as there would not be any more PA available to interact with DGPP. The possibility exists that the observed peak splitting was due to PA/PA and

DGPP/DGPP clustering. If this was the case, then increasing the DGPP concentration while holding the PA concentration constant should reveal this. In this case, the PA peak splitting would stay constant, while the DGPP splitting would increase with concentration.

The GUV results presented in this thesis should only be seen as preliminary and need to be explored further. They show the manifestation of the DGPP/PA clusters under one set of conditions but may not reflect what is actually happening in living systems. To determine that the observed domains are caused by DGPP/PA interactions, controls of just PA or DGPP with phosphatidylcholine (PC) should be tested over a broad pH range. If domains are observed in either of these mixtures at physiological pH, then it is possible that PA/PA and DGPP/DGPP clustering play a role in the domains that were presented in this thesis. Different fluorophores should also be explored for these experiments. The NBD modified PA that was used is known to show a “snorkeling” of the acyl chains into the polar head group region of the bilayer (Kaiser and London, 1998). This behavior could alter how domains would form in the membrane, giving false positive results. The use of Rhodamine-PE somewhat mitigated this concern, but

Rhodamine-PE is still a head group modified lipid, which could also influence how DGPP and PA

79 interact with each other. The ideal fluorophore would be a chain labelled PA that was known not to show the same snorkeling behavior as the NBD-PA. This way the fluorophore could act like a “true” molecule of PA in any domains that may form. It would be beneficial to observe

GUV’s with 2 different fluorophores that have affinities for different lipid phases as well. This could test to see if the lipid domains are caused by a phase transition within the membrane.

Different conditions should also be tested, such as DGPP/PA ratio and the effects of temperature. Altering the DGPP/PA ratio would give insight into just exactly how much DGPP and PA would be needed to produce observable domains. Examining the temperature dependence of domains is also important, as other domain forming lipid systems have been shown to be temperature dependent (Chen and Santore 2014). If these domains are confirmed to exist, they may a role in crucial stress signaling pathways that involve DGPP and PA.

The ionization work with PIP3 should also be probed in further detail. NMR experiments with PI and PIP3 could be examined with higher PIP3 amounts to see if peak splitting is observed. This could yield more insight into the exact nature of the PI/PIP3 interactions that were observed in GUV’s. If peak splitting is observed, it could pinpoint which phospho- monoester of PIP3 interacts with PI or it could show that all 3 phospho-monoesters have the potential to interact. One challenge that would be encountered trying to complete these experiments would be cost. PIP3 is a considerably expensive lipid, and a large amount would need to be purchased to construct a titration curve at even just 10 mol%. One way around this would be to have PIP3 present in a higher ratio but not have as much total lipid overall. This would create a problem of experimental time however, as one experiment could then take 2-3

80 days to resolve. PIP3 is also present in the inner leaflet of the plasma membrane with phosphatidic acid (PA), so there is a possibility that these lipids can form clusters much in the same way as DGPP and PA. This could be observed in NMR experiments as peak splitting, like what was observed for DGPP and PA. Other phosphtidyl-inositols can be also be incorporated into mixtures with PIP3, as it is likely that these species co-exist in the membrane. For example, mixtures of phosphatidylinositol-bisphosphate (PIP2) and PIP3 could be tested in mixtures together with PC to see if the phospho-monoesters of these lipids will interact and have an effect on their respective total charges. The analysis of these experiments may prove to be difficult, as each individual phospho-monoester will be present in the spectra so care would have to be taken to assign individual peaks.

The GUV experimental results involving PIP3 could be further strengthened by creating control vesicles that contain only PIP3 with PC. If these GUV’s do not show domain formation at pH 7, then it will act as a verification that the domains presented in this thesis were in fact due to a PIP3/PI interaction. If domains are formed in PC/PIP3 vesicles, then it raises some questions about the potential for phosphoinositides to form clusters with themselves. This thesis only examined PIP3 containing GUV’s at pH 7 at room temperature, so other pH values and temperatures can be probed to see the effects of these factors on domain formation. A cooling microscope stage could be used to cool the GUV mixture to see if it will promote domain formation at physiological pH, as has been observed in other lipid mixtures (Chen and Santore,

2014). Domains would not be expected at higher pH due to charge repulsion between PI and

PIP3, but domains could still form at lower pH. If domains are in fact formed at low and

81 physiological pH then it could shed some light on the typical organization of PIP3 within the cell, meaning that PIP3 may be present primarily in the membrane as clusters with PI or other lipids.

Since PIP2 and PI were previously shown to form domains at physiological pH (Graber et al.

2012), it is possible that PIP3 may also be incorporated into these clusters. This could be tested by forming GUV’s that contain PIP3, PIP2, and PI. If domains still form under these conditions, then it could provide evidence that phosphoinositides are present as clusters within the membrane. Since PIP3 is a known element of the PTEN signaling cascade, understanding of its exact biophysical role in the membrane could yield a possible therapeutic target for anti-cancer drugs.

References

Chen D and Santore MM. 1,2-Dipalmitoyl-sn-glycero-3-phosphocholine (DPPC)-Rich Domain Formation in Binary Phospholipid Vesicle Membranes: Two- Dimensional Nucleation and Growth. Lamgmuir 30 (2014) 9484-9493 Gericke A, Munson M, and Ross AH. Regulation of the PTEN phosphatase. Gene 374 (2006) 1–9 Graber ZT, Jiang Z, Gericke A, and Kooijman EE. Phosphatidylinositol-4,5-bisphosphate ionization and domain formation in the presence of lipids with hydrogen bond donor capabilities. Chemistry and Physics of Lipids 165 (2012) 696– 704 Kaiser RD and London E. Determination of the depth of BODIPY probes in model membranes by parallax analysis of fluorescence quenching. Biochimica et Biophysica Acta 1375 (1998) 13-22 Lindmo K and Stenmark H. Regulation of membrane traffic by phosphoinositide 3-kinases. Journal of Cell Science 119 (2006) 605-614 Munnik T, Vrije T, Irvinei RF, and Musgrave A. Identification of Diacylglycerol Pyrophosphate as a Novel Metabolic Product of Phosphatidic Acid during G-protein Activation in Plants. The Journal of Biological Chemistry 271(26) (1996) 15708-15715 Shaw RJ. and Cantley LC. Ras, PI(3)K and mTOR signaling controls tumor cell growth. Nature 441 (2006) 424–430

Strawn L, Babb A, Testerink C, and Kooijman EE. The physical chemistry of the enigmatic phospholipid diacylglycerol pyrophosphate. Frontiers in Plant Science 3 (2012)

Appendix List of Materials

Chemical Name Abbreviation Chemical Company Formula Purity Formula Weight

1,2-dioleoyl-sn-glycero- DOPC C44H84NO8P Avanti 786.113 >99% phosphocholine Polar Lipids 1-palmitoyl-2-oleoyl-sn- POPC C42H82NO8P Avanti 760.076 >99% glycero-3-phosphocholine Polar Lipids 1,2-dioleoyl-sn-glycero-3- DOPE C41H78NO8P Avanti 744.034 >99% phosphoethanolamine Polar Lipids Dioleoylglycerol DGPP C39H80N2O11P Avanti 815.007 >99% pyrophosphate Polar Lipids 1,2-dioleoyl-sn-glycero-3- PIP3 C45H98N4O22P4 Avanti 1171.18 >99% [phosphoinositol-3,4,5- Polar Lipids trisphosphate] 1,2,-dioleoyl-sn-glycero-3- PA C39H72O8P Avanti 722.948 >99% phosphate Polar Lipids L-α-phosphatidylinositol Liver PI C47H82O13P Avanti 902.133 >99% (Liver, Bovine) Polar Lipids 1,2-dioleoyl-sn-glycero-3- RhB DOPE C68H109N4O14P Avanti 1301.715 >99% phosphoethanolamine-N- S2 Polar Lipids (lissamine rhodamine B sulfonyl) 1-oleoyl-2-{12-[(7-nitro-2- NBD-PA C39H68N5O11P Avanti 813.958 >99% 1,3-benzoxadiazol-4- Polar Lipids yl)amino]dodecanoyl}-sn- glycero-3-phosphate HPLC Chloroform Chloroform CHCl3 Fisher 119.38 99.8% Scientific HPLC Methanol Methanol CH4O Fisher 32.04 99.9% Scientific HPLC Water Water H2O Fisher 18.02 >99.9% Scientific Citric Acid Citric Acid C6H8O7 Sigma 192.12 99.5% Aldrich 4-(2- HEPES C8H18N2O4S Amresco 238.30 99.7% hydroxyethyl)piperasine-1- ethanesulfonic acid Glycine Glycine C2H5NO2 Amresco 75.07 99%

Sodium Chloride NaCl NaCl Sigma 58.44 99.5% Aldrich Ethylenediaminetetra EDTA C10H16N2O8 Sigma 292.24 99.9% acetic acid Aldrich Ethanol Ethanol C2H6O BDH 46.07 >94% Phosphoric Acid Phosphoric H3PO4 Acid 1,4-piperazinediethane PIPES C8H18N2O6S2 Sigma 302.37 99% sulfonic acid Aldrich Sodium Hydroxide NaOH NaOH Alfa Aesar 39.997 98% Hydrochloric Acid (34%) HCl HCl Sigma 36.46 <10 ppt Aldrich Glucose Glucose C6H12O6 Calbiochem 180.02 99.6% Sucrose Sucrose C12H22O11 Sigma 342.296 99.5% Aldrich

31P MAS NMR Pulse Program and Experimental Parameters

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1 ze 2 30m d1 p1 ph1 go=2 ph31 30m mc #0 to 2 F0(zd) exit

ph1=0 2 2 0 1 3 3 1 ph31=0 2 2 0 1 3 3 1

;pl1: f1 channel - power level for pulse (default) ;p1 : f1 channel - high power pulse ;d1 : relaxation delay; 1-5 * T1 ;NS : 1 * n, total number of scans: NS * TD0

;$Id: zg,v 1.9 2006/11/10 10:56:44 ber Exp $

Experimental Parameters:

NAME --- EXPNO --- PROCNO --- Date --- Time --- INSTRUM spect PROBHD --- PULPROG zgig TD 8192 SOLVENT NA NS 12996 DS 4 SWH 81521.742 Hz FIDRES 9.951385 Hz AQ 0.0502943 sec RG 2050 DW 6.133 usec DE 10.00 usec TE 294.0 K D1 1.00000000 sec D11 0.03000000 sec TD0 75

======CHANNEL f1 ======NUC1 31P P1 5.25 usec PL1 8.00 dB PL1W 62.50282669 W SFO1 161.9765351 MHz

======CHANNEL f2 ======CPDPRG2 spinal64 NUC2 1H PCPD2 22.25 usec PL2 120.00 dB PL12 14.00 dB PL2W 0.00000000 W PL12W 2.95000005 W SFO2 400.1360019 MHz SI 8192 SF 161.9766731 MHz WDW EM SSB 0 LB 50.00 Hz GB 0 PC 4.00

Static 31P NMR Pulse Program Pulse Program: ;zgig ;avance-version (07/04/03) ;1D sequence with inverse gated decoupling ; ;$CLASS=HighRes ;$DIM=1D ;$TYPE= ;$SUBTYPE= ;$COMMENT=

#include

"d11=30m"

"acqt0=-p1*2/3.1416"

1 ze d11 pl12:f2 2 30m do:f2 d1 p1 ph1 go=2 ph31 cpd2:f2 30m do:f2 mc #0 to 2 F0(zd) exit

ph1=0 2 2 0 1 3 3 1 ph31=0 2 2 0 1 3 3 1

;pl1 : f1 channel - power level for pulse (default) ;pl12: f2 channel - power level for CPD/BB decoupling ;p1 : f1 channel - high power pulse ;d1 : relaxation delay; 1-5 * T1 ;d11: delay for disk I/O [30 msec] ;NS: 1 * n, total number of scans: NS * TD0 ;cpd2: decoupling according to sequence defined by cpdprg2 ;pcpd2: f2 channel - 90 degree pulse for decoupling sequence

Figure S-1: Titration data for PC:PA at 95:5. Shown is the unpublished PA titration data that is used as a reference in chapter 3. Individual data points are shown in red. Titration data was fit with a modified Henderson-Hasselbach equation (shown in black).

Figure S-2: Titration data for PC:PE:PA at 47.5 : 47.5 : 5. Shown is the unpublished PA titration data that was used as a reference in chapter 3. Individual data points are shown in red. Titration data was fit with a modified Henderson-Hasselbach equation (shown in black).

Figure S-3: Static spectra for PC:DGPP:PA at 90 : 5 : 5. pH values are as indicated to the right of each spectra. The presence of a large peak with a long shoulder indicates the formation of a lipid bilayer.

Figure S-3: Static spectra for PC:PE:DGPP:PA at 45 : 45 : 5 : 5. pH values are as indicated to the right of each spectra. The presence of a large peak with a long shoulder indicates the formation of a lipid bilayer.

Figure S-4: Static spectra for PC:DGPP:PA at 95 : 2.5 : 2.5. pH values are as indicated to the right of each spectra. The presence of a large peak with a long shoulder indicates the formation of a lipid bilayer.

Figure S-5: Static spectrum for PC:PE:DGPP:PA at 47.5 : 47.5 : 2.5 : 2.5. pH value is indicated to the right. Only one static experiment was taken for this data series. This spectra shows the characteristic shape that indicates lipid bilayer formation.

Figure S-6: Static spectra for PC:PE:PIP3 at 47.5 : 47.5 : 5. pH value is indicated to the right. Spectra shape is indicative of bilayer formation.

Figure S-7: Static spectra for PC:PI:PIP3 at 88 : 10 : 2. pH values are as indicated to the right. The peak and long shoulder in each spectra are indicative of bilayer formation.