Biophysical Studies of the First Nucleotide Binding Domain of SUR2A

Elvin Dominic de Araujo

A thesis submitted in conformity with the requirements for the degree of Master of Science Department of Chemistry University of Toronto

Elvin Dominic de Araujo

Master of Science, 2011

Department of Chemistry

University of Toronto

Abstract

ATP-sensitive potassium (KATP) channels have crucial roles in several biological processes. KATP channels possess four regulatory sulfonylurea receptors. The SUR proteins are members of the ubiquitous ATP-binding cassette (ABC) superfamily. However, unlike most ABC proteins,

SURs do not transport substrates but function strictly as regulators of KATP channel activity.

Currently, studies into the molecular basis by which various mutations in SUR2A cause disease are highly limited. This is primarily a consequence of poor solubility of isolated SUR2A NBDs, as is typical for many eukaryotic NBDs. By employing structure-based sequence alignments and biophysical studies, we determined domain boundaries for SUR2A NBD1 that enabled, for the first time, NMR studies of NBD1. Our biophysical studies demonstrate that the isolated SUR2A

NBD1 is folded and exhibits differential dynamics upon ATP binding activity. Additional studies are now possible to examine the effects of disease-causing mutations on structure, dynamics, and interactions of NBD1.

Acknowledgments

I would like to thank my supervisor, Prof. Voula Kanelis for her valuable support and enduring guidance throughout my Masters program. Her enthusiasm for science is truly motivating. Prof. Kanelis is not only a dedicated scientist but also helped further my development as a researcher.

I would also like to thank Prof. R. Scott Prosser for providing me with both insightful suggestions and technical support. As an avid biophysicist, his eagerness has helped instill my interest in biophysics. Working in his lab during my undergraduate years, encouraged me to pursue research.

Prof. Ulrich J. Krull has taught me valuable lessons about analytical chemistry, widening my scope in my scientific approach.

Past and present colleagues, Serisha Moodley, Lynn K. Ikeda, Dennis Guo, Marijana

Staglijar and Dr. Jorge Pédro Lopez-Alonso have been instrumental both in creating an enjoyable and stimulating environment to work in the lab.

Thanks to Prof. George S. Espie, Prof. Virginijus Barzda and Prof. Jumi Shin for allowing me to use their scientific equipment.

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Table of Contents

1. Introduction 1.1. Background of ABC Transporters 1 1.2. Structural Architecture of ABC Transporters 2 1.3. Nucleotide Binding Domains 4 1.4. Sulfonylurea Receptors 7

1.5. KATP channels 9 1.6. Inwardly Rectifying Potassium Channels 10

1.7. Physiological Importance of KATP Channels 13

1.8. Regulation of KATP Channels Activity 16 1.9. Cardiac Disease-Causing Mutations in SUR2A 17 1.10. Biophysical Methods for Investigation of SUR2A 19 1.10.1 Nuclear Magnetic Resonance Spectroscopy 19 1.10.2 Circular Dichroism Spectroscopy 22 1.10.3 Fluorescence Spectroscopy 23

2. Materials and Methods 26 2.1 Structure-Based Sequence Alignment of NBDs 26 2.2 Homology modeling of SUR2A NBD1 26 2.3 Generation of Recombinant NBDs 27 2.4 Expression and Purification of SUR2A NBD1 28 2.5 Phosphorylation of Wild Type and Mutant SUR2A NBD1 30 2.6 NMR Spectroscopy 31 2.7 Circular Dichroism Spectroscopy 31 2.8 Fluorescence Spectroscopy 32

3. Results 33 3.1 Structure-Based Sequence Alignment 33 3.2 Generations of Constructs with Variable Domains Boundaries for NBD1 35 3.3 Expression and Purification of SUR2A NBD1 37 3.4 Biophysical Characterization of SUR2A NBD1 40 iv

3.4.1 NMR Studies of 15N labeled S615-L933 40 3.4.2 NBD1 SUR2A S615-L933 functionally binds ATP 44 3.4.3 NMR Studies of T618-L933 47 3.4.4 Phosphorylation of S615-L933 NBD1 SUR2A 53 3.4.5 Role of R934-K972 in NBD1 SUR2A 56 3.4.6 Secondary Structural Analysis of NBD1 SUR2A 61 3.4.7 Fluorescence Spectroscopy of NBD1 SUR2A 64 3.5 Biophysical studies of SUR2A NBD1 Val734Ile 68

4. Discussions and Conclusions 4.1 Identification of SUR2A NBD1 Domain Boundaries 72 4.2 Characterization of ATP binding in SUR2A NBD1 74 4.3 Effects of Phosphorylation on NBD1 of SUR2A 77 4.4 Role of the ED Domain in NBD1 78 4.5 Characterization of SUR2A NBD1 Val734Ile 79 4.6 Conclusions and Future Directions 81

5. Appendix 82

6. References 87

List of Tables

Table 1: Subfamilies of human ABC transporters 2

Table 2: Subunit assembly of KATP channels in different tissues 14 Table 3: Gyromagnetic ratios of some NMR active nuclei 20 Table 4: Primers used in the PCR amplification of SUR2A NDBs 28 Table 5: Total protein yields of various NBD1 SUR2A constructs 37 Table 6: Number of amide proton crosspeaks in NBD1-S615-L933 42 Table 7: Number of amide proton crosspeaks in NBD1-S615-L933 (± ATP) 46 Table 8: Number of amide proton crosspeaks in NBD1-T618-L933 48 Table 9: Number of amide proton crosspeaks in phosphorylated NBD1-S615 -L933 55 Table 10: Number of amide proton crosspeaks in NBD1-S615-K972 at 35oC 59 Table 11: Melting temperatures determined by fluorescence 66 Table 12: Tryptophan solvent exposure determined by fluorescence quenching 67 Table 13: Tryptophan solvent exposure of Val734Ile SUR2A NBD1 determined by fluorescence Val734Ile 71

List of Figures

Figure 1. General structure of an ABC transporter 3 Figure 2. General structure of the nucleotide binding domain of ABC transporters 5 Figure 3. Dimerization of NBDs upon ATP binding 6

Figure 4. Model for regulation of KATP channels by the SURs 8

Figure 5. Schematic representation of the KATP channel 9

Figure 6. Model of the KATP channel 10 Figure 7. Classification of potassium channels 11 Figure 8. Representation of dimeric Kir subunits 12

Figure 9. Role of KATP channels in various tissues 15 Figure 10. Mutation and phosphorylation sites in SUR2A NBDs 17 Figure 11. Circular dichroism spectra of common secondary structure features 22 Figure 12. Possible excitation and relaxation pathways of a chromophore with light 24 Figure 13. Structure-based sequence alignment of SUR2A NBD1 with other ABCC transporters 34 Figure 14. Protein profile of pre- and post- induction for proteins of various domain boundaries 36 Figure 15. Localization of expressed protein to soluble or insoluble fractions 36 Figure 16. Purification of NBD1 S615-L933 38 Figure 17. Time course of Ulp1 activity on 6xHisSUMO-NBD1 S615-L933 38 Figure 18. Fractions following gel-filtration of cleaved 6xHisSUMO-NBD1 S615-L933 39 Figure 19. 2D 15N-1H TROSY-HSQC spectra of NBD1-SUR2A S615-L933 at 30oC 40 Figure 20. 2D 15N -1H TROSY-HSQC spectra of NBD1-SUR2A S615-L933 at various temperatures 43 Figure 21. 2D 15N -1H TROSY-HSQC spectra of NBD1-SUR2A S615-L933 with varying levels of ATP 45 vii

Figure 22. Overlay of 2D 15N -1H TROSY-HSQC spectra of NBD1-SUR2A S615-L933 with and without ATP 46 Figure 23. Homology model of NBD1 of SUR2A 47 Figure 24a. Purification of T618-L933 47 Figure 24b. 2D 15N -1H TROSY-HSQC spectra of NBD1-SUR2A S615-L933 at varying temperatures 48 Figure 25. Overlay of 2D 15N -1H TROSY-HSQC spectra of NBD1-SUR2A S615-L933 with T618-L933 51 Figure 26. Overlays of 2D 15N -1H TROSY-HSQC spectra of NBD1-SUR2A T618-L933 52 Figure 27. 2D 15N -1H TROSY-HSQC spectra of Phosphorylated NBD1-SUR2A S615-L933 54 Figure 28. 2D 15N -1H TROSY-HSQC spectra of Phosphorylated NBD1-SUR2A S615-L933 without ATP at 30oC 55 Figure 29. Purification of NBD1 S615-K972 56 Figure 30. Fractions following gel-filtration of cleaved 6xHisSUMO-NBD1 S615-L933 56 Figure 31. 2D 15N -1H TROSY-HSQC spectra of NBD1-SUR2A S615-K972 58 Figure 32. 2D 15N -1H TROSY-HSQC spectra of NBD1-SUR2A S615-K972 at 35oC 59 Figure 33. 2D 15N -1H TROSY-HSQC Phosphorylated of NBD1-SUR2A S615-L972 at 30oC 60 Figure 34. Circular dichroism spectrum of S615-L933 61 Figure 35. Guanidine-HCl melt of NBD1 SUR2A S615-L933 62 Figure 36. CD spectra of phosphorylated S615-L933 (A) and S615-K972 (B) 63 Figure 37. Fluorescence temperature melting curve of S615-L933 64 Figure 38. Fluorescence temperature melting curve of S615-L933 under varying Conditions 65 Figure 39. Quenching of fluorescence of S615-L933 by potassium iodide 66 Figure 40. Modified Stern-Volmer plots of fluorescence quenching of S615-L933 67 viii

Figure 41. 2D 15N -1H TROSY-HSQC of Mutant Val734Ile of NBD1-SUR2A S615-L933 69 Figure 42. CD spectra of Mutant Ile734Val in S615-L933 70 Figure 43. Fluorescence melt curves of Mutant Ile734Val in S615-L933 in the presence/absence of ATP 70 Figure 44. Modified Stern-Volmer plots of fluorescence quenching with Val734Ile3 of S615-L933 71 Figure 45. Altered protein dynamics with ATP binding 75 Supplementary Figure 1: Structure-based sequence alignment of NBD1 with other ABCC transporters 82

List of Appendices

Supplementary Figures 89

List of Abbreviations

ABC ATP-binding cassette ABCR rod outer segment ABC transporter ADP Adenosine diphosphate ATP Adenosine triphosphate

Bo External magnetic field CD Circular dichroism CFTR Cystic fibrosis transmembrane conductance regulator DNA Deoxyribonucleic acid DSS 4,4-dimethyl-4-silapentane-1-sulfonic acid DTT Dithiothreitol fa Fraction of fluorophore population accessible to quenching

Go Ground-state energy h Planck’s constant, 6.626068 × 10-34 m2 kg/s HSQC Heteronuclear single quantum coherence HUGO Human Genome Organization ICD Intracellular domain IPTG Isopropyl-β-D-thio-galactoside K Quenching constant K+ Potassium ion

KATP channel ATP-sensitive potassium channel

KCO KATP channel openers

Kd Dissociation constant kDa Kilodalton kq Bimolecular quenching constant L Angular momentum LB Luria Bertani Mg2+ Magnesium ion MRP Multidrug resistant proteins MSD Membrane spanning domain

NaCl Sodium chloride NBD Nucleotide Binding Domain Ni+2 Nickel ion NMR Nuclear magnetic resonance NTA Nitrilotriacetic acid OD Optical density PCR Polymerase chain reaction PDB Protein Data Bank PHHI Persistent hyperinsulinemic hypoglycemia of infancy PKA Protein kinase A PONDR Predictors of Natural Disordered Regions Q Quencher RE regulatory extension RI regulatory insert SAXS Small angle X-ray scattering SDS-PAGE sodium dodecyl sulfate polyacrylamide gel electrophoresis SUMO Small Ubiquitin-like Modifier SUR Sulfonylurea receptors

T2 Spin-spin relaxation TAP ATP-binding-cassette (ABC) transporter associated with antigen processing TCAG The Center for Applied Genomics TRIS 2-Amino-2-hydroxymethyl-propane-1,3-diol TROSY Transverse relaxation optimized spectroscopy Ulp1 (ubiquitin-like protein)-specific protease 1 UV Ultraviolet WT Wild type ΔE Energy separation

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Chapter 1 Introduction

1.1 Background to ABC Transporters

Members of the ATP-binding cassette (ABC) superfamily constitute one of the largest classes of transport proteins and possess a ubiquitous distribution across all kingdoms of life.[9, 10]

Generally, these proteins are involved in vectorial transport of substrates across a biological membrane, but many also play a role in a number of translocation-independent processes, such as DNA repair and gene regulation.[9] ABC proteins are involved in a number of physiological and clinically significant processes such as preservation of the blood brain barrier in mammals by P-glycoprotein or in visual detection by the rod outer segment ABC transporter (ABCR) which functions.[6] Forty-nine ABC proteins have been identified in the human genome, with

17 of these proteins correlated to hereditary diseases.[10]

Proteins are classified as ABC transporters based on the sequence of the corresponding gene cassette as well as the architecture of the overall protein domain organization.[11]

Universal to all ABC transporters is the binding and subsequent hydrolysis of ATP. However,

ABC transporters have been further classified into seven subfamilies (subfamilies A-G) by the

Human Genome Organization (HUGO), primarily on the basis of sequence similarity between their nucleotide binding domains (NBDs).[11] The NBDs are the sites of ATP binding and hydrolysis. Frequently, ABC proteins affiliated to a particular subfamily possess congruent biochemical functions. Table 1 displays some general functions and examples of the different

ABC transporter subfamilies. Amongst the ABC transporters, the C subfamily (ABCC) is of particular interest for this work. There are twelve such ABCC transporters recognized in the human genome. These include the two sulfonylurea receptors (SUR1 and SUR2 isoforms),

which regulate ATP-sensitive potassium (KATP) channels, and the cystic fibrosis transmembrane conductance regulator (CFTR) which is involved in epithelial fluid secretion. Furthermore, there are nine multidrug resistant proteins (MRPs) in the ABCC subfamily, which are responsible for efflux of macromolecules/toxins from cells and impart varying degrees of drug resistance.[12]

Table 1: Human subfamilies of ABC transporters[13] Family Human Generalized Functions Examples Members ABCA 12 Transport of lipids Cholesterol efflux regulatory protein (ABCA1)[14] ABCB 11 Transport of peptides, bile P-glycoprotein (ABCB5)[15] ABCC 13 Ion transport, toxin efflux, Sulfonylurea receptors (ABCC8/9), CFTR (ABCC7)[6] ABCD 4 Used in peroxisomes Adrenoleukodystrophy (ALDP) protein, ABCD1[16] ABCE 1 Regulate protein expression, RNAse L inhibitor No MSD in either subfamily (ABCE1)[17] ABCF 3 ABCF2 (Iron inhibited transporter)[18] ABCG 5 Diverse transport functions Sterolin 1 and 2 (ABCG7/8)[19]

1.2 Structural Architecture of ABC Transporters

Despite a high level of diversity in substrate specificity and functional attributes between members of the ABC superfamily, they possess a common molecular architecture as illustrated in Figure 1. At minimum, ABC transporters are comprised of two cystolic nucleotide binding domains (NBDs) and two membrane spanning domains (MSDs).[9] MSD1 and MSD2 are involved in interactions with the target substrate whereas the NBDs are the sites of ATP binding and hydrolysis. Considering the variability of transport substrates, the primary sequence of the

MSDs is noticeably more variable in comparison to the NBDs.[6] Structures of full length ABC proteins indicated that the transmembrane helices in the MSDs extend beyond the lipid bilayer, into the cytoplasm to form intracellular domains (ICDs). The long helical extensions of the ICDs 2

Figure 1: General structure of an ABC transporter a) Schematic representation of the structure for the SUR proteins. The membrane spanning domains are numbered and depicted in grey, and the ICDs are shown in purple. The individual nucleotide binding domains are shown in blue and green. Members of ABCC subfamily (excluding CFTR) possess an N-terminal MSD0 domain linked to the traditional structure by an L0 linker. b) A ribbon diagram for the bacterial ABC transporter Sav1866 (PDB coded: 2HYD). The domains shown in a) are illustrated in analogous colours for comparison. Sav1866 is a homodimer and does not contain the additional MSD0 or L0 linker.[6] [9] are connected by short irregular helices known as the coupling helices. The coupling helices contact the NBDs, thereby linking the NBDs and MSDs.[6] Conformational changes in the

NBDs, such as those following ATP-binding, are relayed through coupling helices, to the ICDs, and ultimately to the transmembrane domains to allow for ligand binding or transport. In prokaryotes, each of the nucleotide binding domains and the transmembrane domains (including the ICDs and coupling helices) are encoded as distinct polypeptides, which leads to the association of four separate subunits.[6] In eukaryotes, however, the four domains are typically encoded and expressed as one or two polypeptides. A two polypeptide transporter is referred to as a ‘half-transporter’ and incorporates a format of (NH2-MSD-NBD-COOH). This design is most common to the ABCG subfamily. This model has been experimentally reproduced to express single polypeptide ABC transporters, such as MRP1 and CFTR, as half transporters with a single MSD and NBD domain. In such expression systems, active transport is only observed

when both halves are co-expressed, underscoring the importance of interactions between all four domains for functional activity.[6]

Members of the ABCC subfamily are further classified into a ‘long’ or ‘short’ configuration based on the presence of accompanying subunits to the minimum ABC structure.[20] The ‘long’ members possess an additional membrane spanning domain (known as

MSD0), which is connected to MSD1 via the cytoplasmic L0 linker. Long members in the

ABCC family include both SUR1 and SUR2, as well as various multidrug resistance proteins,

(MRP1, MRP2, MRP3, MRP6 and MRP7). Short members possess the conventional ABC structure and encompass the remaining ABCC transporter proteins.[20] Although the specific function of MSD0 is not entirely understood, this domain has been shown to be essential for proper trafficking of the SURs to the cell membrane.[21]

1.3 Nucleotide Binding Domains

The NBDs function as the diagnostic core for classification of ABC transporters and contain many conserved elements, as depicted in Figure 2. Crystal structures of bacterial, and more recently eukaryotic nucleotide binding domains, have partitioned them into two structural lobes.

[22] Lobe I possesses an α/β subdomain that contains a Walker A (P-loop) motif

(GXXGXGK(S/T)) and a Walker B motif ( D, where is a hydrophobic residue).[23]

These motifs collaborate in the binding and hydrolysis of ATP. The α/β subdomain is often referred to as the F1-like subdomain due to structural and sequence similarities with F1-

ATPases.[24] Furthermore, lobe I adopts a Rossman fold, similar to other F1-ATPase enzymes.

The Rossman fold constitutes of a central core of -sheets enclosed by -helices.[25] Lobe I also contains a β-sheet subdomain. This domain contains an aromatic residue C-terminal to the first

β-strand which interacts with the adenine base of ATP. Lobe II is formed by the α-helical subdomain that contains a signature motif (LSGGQ) of ABC transporters (known as a C-motif), which also assists in nucleotide binding.[26] Also present in the α-helical subdomain, are other well conserved motifs, such as the Q-loop and H-motif. The Q-loop is suspected to be involved in communicating conformational changes between the MSDs and NBDs, especially upon ATP hydrolysis. The H-motif is referred to as the switch region, which contains a highly conserved histidine residue that allows for interactions of the NBD with the nucleotide.[27]

High-resolution structures from X-ray crystallography of isolated NBDs have revealed insights into the architecture and arrangement of the various domains. Eight full-length ABC transporter structures have been solved, including murine P-glycoprotein.[25, 28-34]

Complexed ABC transporters with various nucleotides have also allowed for several structures

of dimeric NBDs to be solved.

Currently, however, only one such

dimeric structure exists for eukaryotic

NBDs.[35] The cumulative analysis

of these structures exposes an overall

‘L’ shape fold to the global NBD

structure, which is now considered a

canonical design to NBDs.[9] Crystal

Figure 2: General structure of the nucleotide binding structure analysis indicates that the domain of ABC transporters Ribbon diagram of the nucleotide binding domain of a invariant lysine residue of the Walker bacterial transporter (PBD: 1F30) illustrates various conserved motifs across all NBDs. The Walker A motif is A motif in lobe I associates with the illustrated through the yellow heavy atoms. The red heavy atoms illustrate the Walker B motif and the green heavy atoms, illustrate the conserved signature C sequence. The and phosphates of the ATP tail via phosphate tail of ATP (shown in grey, blue and red) interacts with Mg2+ (magenta) near the Walker A motif. 5

ionic interactions.[36] Hydrogen bonding is shown to occur primarily between phosphate- oxgyen atoms of ATP to backbone nitrogen atoms. This may partially explain the variability in the non-conserved residues of the Walker A motif. The Walker B motif forms a β-strand of a highly hydrophobic β-sheet. However, it concludes with an acidic residue which coordinates to a magnesium ion. X-ray structures have also indicated that lobe II, containing the ABC signature sequence, is comprised of an α-helical subdomain.[10] This domain arises from three conserved

α-helices among ABC transporters. Lobe II facilitates nucleotide binding in dimeric NBDs, as well as interactions with the MSDs via the coupling helices. The H-motif possesses a conserved histidine residue which is believed to polarize the nucleophilic water molecule during ATP hydrolysis.[30]

Figure 3: Dimerization of NBDs upon ATP binding a) The NBDs have been shown to dimerize upon binding of ATP. They orient in a head-to-tail fashion such that Lobe I of NBD1 interacts with Lobe II of NBD2 and vice versa. A ribbon diagram of bacterial NBD1 and NBD2 interacting with ATP is depicted in b) (PDB: 1L2T).

Binding of ATP induces conformational changes with the NBDs that lead to dimerization event of the two domains.[6] It is suspected that the dimerization and the resulting mechanical consequences are not responsible for substrate translocation observed in ABC transporters.

However, it is believed that subsequent conformational changes in the dimeric structure, such as those caused upon ATP hydrolysis and the resulting interactions with Mg2+-ADP, may drive substrate transport. The NBD1/NBD2 dimerization results in a head-to-tail interaction as initially

seen in the dimeric structure of Rad50.[37] This DNA repair enzyme, which has catalytic domains related to the NBDs, was crystallized in the presence of a non-hydrolyzable ATP analog. This structure delineated residues from the Walker A and Walker B motifs interacting with the pyrophosphate functional group of ATP. The signature C sequence of the opposing monomer completes the active site, and supplements interactions with the phosphate tail as well as with the adenine base and ribose sugar. Crystal structures of other NBDs conform to these models.[6]

X-ray crystallographic visualization of the NBDs has provided enormous insights into the structural organization of domains and motifs in ABC transporters. However, X-ray crystallography is limited in that it provides information on static and immobile macromolecules.

Such information, though invaluable, can be incomplete for understanding dynamics mechanisms such ATP hydrolysis and translocation processes. NMR, similar to X-ray crystallography, offers residue specific information with additional information on mobility.

Thus, NMR is an ideal technique for understanding kinetic mechanisms and conformational changes.

1.4 Sulfonylurea Receptors

Sulfonylurea receptors (SURs) are atypical members of the ABCC family in that they lack intrinsic transport function. They are named for their affinity towards sulfonylurea derivatives which are antidiabetic drugs. There are two genes, ABCC8 and ABCC9, which encode SUR1 and SUR2 respectively.[38] Despite strong sequence identity, the proteins appear to possess reciprocal pharmacological profiles. SUR1 possesses a high affinity for sulfonylurea drugs and a significantly lower affinity towards KATP channel openers (KCOs).[39] SUR2 possesses a lower affinity for sulfonylureas, but a markedly higher affinity for KCOs. The isoforms of SUR2, 7

SUR2A and SUR2B, are splice

products which are translated from

a single gene. They differ only in

the C-terminal 42 residues. This

dissimilarity results in

modifications to the allosteric

mechanism for ATP binding and

hydrolysis. For example,

Kir6.2/SUR2B has been shown to

respond with greater sensitivity

than its counterpart,

Kir6.2/SUR2A, to the KCO Figure 4: Model for regulation of KATP channels by the SURs In A), potassium transport is inhibited by interactions of nicroandil.[38] This may imply an MSD0 and the L0 linker with the Kir subunit. Binding and hydrolysis of ATP lead to a conformational change in B) inhibitory function for the C- which is transmitted to the L0 linker and MSD and disrupts their interaction with the Kir subunit. The removal of this terminal 42 amino acids that interaction, allows the channel to open, and the transport of potassium (modified from reference 4). disrupt certain interactions. Three

other spliceosomes have been identified in murine and rat SUR2A and SUR2B that contain either a deletion of exon 14 (Δexon 14) or exon 17 (Δexon17). The SUR2A Δexon 14is found in cardiac muscle and is also known as SUR2C. Deletion of Δexon17 results in removal of a large disordered region (35 amino acids) N-terminal to the Walker A motif. Removal of exon 17 encompasses the deletion of 13 residues neighbouring the Walker A motif which has shown to reduce ATPase activity in the NBDs.[38]

Structural information relating to the SUR proteins is extremely limited. Currently, there are electron microscopy images of the Kir6.2/SUR1 channel, as well as low resolution 8

synchrotron radiation small angle X-ray scattering (SAXS) images of the SUR2A NBDs.[1, 40]

These data have indicated tetrameric assembly of the SUR proteins in the channel, with octameric formation of the NBDs. Previous studies on secondary structure from CD analysis depicts SUR2A NBD1 as having larger alpha helical character, and NBD2 with slightly lower alpha helical content and increased disordered regions.[40]

Figure 4 provides a potential mechanism for KATP regulation by the SUR proteins. The currently accepted model for such regulation involves interactions possibly between MSD0 and the L0 linker to a Kir subunit.[4] These interactions maintain the affinity of the Kir subunits for

ATP and keep the pore in a closed state (Fig 4A). Binding and subsequent hydrolysis of ATP at the NBDs in the SUR protein induces a conformational change. This change is suspected of producing a mechanical effect, transferred through the L0 linker and MSD0, which results in displacement of the SUR subunit from the Kir subunit. The Kir subunits, lose their affinity towards ATP, which leads to an open state of the pore, and transport of K+ ions (Fig 4B).[4]

1.5 KATP channels

ATP-sensitive potassium (KATP) channels are heteromultimeric protein complexes found in metabolically active tissues, such as the heart and pancreas.[41] By sensing changes in intracellular ATP and ADP concentrations,

KATP channels link cellular metabolism to membrane potential and excitability, and thus play crucial roles for KATP channels in several biological processes.[42] KATP Figure 5: Schematic representation of the channels are comprised of pore (Kir6.x) KATP channel The KATP channel is composed of four copies of the Kir subunit (blue) and four copies of the SUR subunit (yellow and red).[5]

and regulatory subunits (SUR1, SUR2A, SUR2B) (Figure 5). As octomeric complexes, these channels exist with a 4:4 stoichiometry of Kir subunits to SUR subunits. The EM structure is consistent with a molecular architecture in which the Kir subunits form the centre of the complex, while the SUR proteins surround the pore. Kir subunits are members of the inwardly rectifying potassium channel family (Figure 6). KATP channels possess either the Kir6.1 or Kir6.2 proteins. SUR proteins are members of the C subfamily of ABC transporters. SUR proteins do not possess any intrinsic transport ability but function strictly as regulators of Kir6.x activity in

KATP channels. SUR proteins are translated from two genes as previously described, SUR1 and

SUR2, and SUR2 possesses a number of different isoforms.[5, 43]

Figure 6: Model of the KATP channel A schematic representation of the side view is shown in a), based on electron microscopy maps of the KATP channels. The Kir subunits are shown in blue surrounded by the MSD0 (yellow) and the rest of the SUR (red) subunits. A bird’s eye view is depicted in b), and the top view ascribed through electron microscopy is shown in c).[1]

1.6 Inwardly Rectifying Potassium Channels

Inwardly rectifying potassium (Kir) channels are a subset of potassium selective ion channels which, as indicated by their title, participate in potassium transport across the cellular membrane.

[1] The various classes of potassium channels are depicted in Figure 7. The inwardly rectifying phenomenon refers to the impermeability of positive charge in the reverse direction. The Kir

Figure 7: Classification of potassium channels KATP channels form a distinct family of transporters involved in potassium uptake. Voltage-gated potassium channels exist in the open or closed state based on the transmembrane potential. They are important for the transmission of action potentials. Calcium-activated potassium channels are stimulated by decreasing intracellular calcium. Leak potassium channels are responsible for the resting membrane potential. Inwardly rectifying potassium channels allow for only unidirectional passage of potassium charge. (Modified from reference [7])

channels are subdivided into seven families (Kir1 to Kir7) through functional and phylogenetic analysis, each with several splice variants.[44] Different levels of inward rectification are also observed between the various subfamilies. For example, Kir6 members are weakly inwardly rectifying, whereas Kir2 subfamilies are strongly inwardly rectifying. [45]

KATP channels of the heart, liver, pancreas and smooth muscle all utilize subunits from the Kir6 family. The Kir6.1 subunit is found in vascular smooth muscle, whereas the Kir6.2 subunit is located in the heart, liver and pancreas.[46] Although, Kir6.1 and Kir6.2 retain a high sequence identity (~71%), the Kir6.1 subunit sustains only about half the conductance activity of 11

Kir6.2. An additional and unique Kir6.3 subunit has also been recently isolated from zebrafish, with 66% sequence identity to Kir6.2 and similar SUR protein assembly requirements for functional activity.[47]

The general structure of the Kir subunit, is composed of two transmembrane α- helical domains (TM1 and TM2) which are coupled together through a motif known as a P-loop. The P- loop contains the potassium ion selectivity filter through the conserved sequence tyrosine-valine- glycine-phenylalanine-glycine.[48] The Kir subunits assemble together as tetramers in the KATP channel as shown in Figure 6. The TM1 helix of each Kir subunit is situated on the outer perimeter of the tetrameric complex, whereas the TM2 helix of each subunit, is positioned at the inner circumference (Figure 5,8). Thus, each Kir subunit donates a TM2 helix to create the potassium ion transport channel. The TM2 helices contain a glycine residue roughly bisecting the

length of the helix. The glycine

residue is thought to impart

flexibility into the helical structure

to accommodate a range of

diameters and aid in transport of

potassium ions through the

channel.[48] The potassium

selectivity mechanism involves

remodelling the potassium ion

hydration shell into the tertiary Figure 8: Representation of Dimeric Kir subunits The Kir subunit is comprised of a TM1 and TM2 connected protein structure. For instance, the through a P-loop. Each of the TM2 from the interacting subunits form the pore for potassium transport with TM1 distance between carbonyl-oxygens located on the outer circumference of the pore.

of the amide backbone to the potassium ion being transported is identical to the solvation envelope around K+. Binding and transport of Na+ through the channel, would be energetically unfavourable. This is due to the strong interactions which would need to be overcome in order for the channel to collapse to the smaller size required for sodium transport.[49]

The N- and C- terminal ends of the Kir subunit are both intracellular and form part of the pore complex which begins in the membrane and extends into the cytoplasm. The additional extensions of the pore into the cytoplasm allow for increased regulation over pore movement.[48] For instance, the larger presence of acidic residues on the cystolic side increases the inward rectification. Reverse transport occurs less frequently due to the presence of endogenous positively charged molecules, such as Mg2+ as well as several polyamines. These molecules competitively reduce K+ binding to the intracellular Kir residues.[49] Furthermore, the extended pore complex allows for increased modulation by various regulators, such as ATP, sodium ions or pH. Thus, this enlarged pore complex likely plays significant roles in conferring conformational changes in SUR proteins to the Kir subunits, to excise control over the open/close state of the KATP channel. [45]

1.7 Physiological Importance of KATP Channels

Despite the similarity within both Kir and SUR proteins, only specific permutations of subunits create properly functional units in different tissues. The interacting subunits in corresponding tissues are summarized in Table 2.

Table 2: Subunit assembly for KATP channels in different tissues[50] Cellular SUR Subunit Kir Subunit Localization Brain SUR 1 Kir6.2 Pancreatic β-cells SUR 1 Kir6.2 Cardiac Tissue SUR 2A Kir6.2 Smooth Muscle SUR 2B Kir6.2 Vascular Smooth SUR 2B Kir6.1 Muscle

Overactive mutant Kir6.2 subunits in cardiac muscle yield no serious biological consequence.

However, over-activity in pancreatic β-cells results in diabetes. This underscores the importance in understanding how differences in the SUR proteins affect regulation.[8] The general biological functions of KATP channels in various tissues are summarized in Figure 9.

Figure 9: Role of KATP channels in various tissues[4] The basal state for KATP channels is different depending on the cellular localization. In pancreatic tissue (a), the resting state exists primarily in the open state due to interactions with Mg2+-ADP. Following digestion of food, glucose levels rise leading to increases in intracellular ATP. This displaces ADP from the NBD2 active site and leads to closure of the channel and binding of Kir6.2. The resulting membrane depolarization leads to calcium ion influx and triggers the release of insulin. In heart tissue (b), KATP channels offer protection against ischemic events. The resting channel state is generally closed due to interactions with ATP. Following cardiac stress, ADP levels rise which leads to opening of the channel, and an increase in positive charge in the cell interior. This results in limiting the duration for action potentials and restricts the use of ATP (conservation of energy). In endothelial cells (c), KATP channels may exist in either the open or closed conformation during the resting state. Vasodilators can actually instigate a biochemical pathway, such that ADP can form favourable interactions with the KATP channel. This leads to opening of the channel, influx of potassium and hyperpolarization. As a result, intracellular calcium ion concentrations decrease resulting in relaxation of smooth muscles. Vasoconstrictors, as one would expect, instigate the reverse process for smooth muscle. [4, 8]

1.8 Regulation of KATP Channel Activity

Both Kir and SUR subunits have binding sites for ATP, although nucleotide binding leads to opposing effects on channel activity. The binding of a single ATP molecule at the Kir site can lead to complete channel closure. However, binding of ATP at the NBD sites of the SUR proteins produces conformational changes that result in opening of the KATP channel.[51, 52]

Phosphorylation of both the Kir and SUR subunits has been shown to produce stimulatory effects on KATP channel activity. There are a number of phosphorylation sites which have been identified in murine KATP channels of vascular tissue, including Ser385 in Kir6.1,

Thr633 in NBD1, as well as Ser1387 and Ser1465 in NBD2 (Figure 10).[53] Although the latter phosphorylation sites were identified in SUR2B, they are also present in SUR2A. [54]

Furthermore, the phosphorylation sites are conserved among all SUR2 isoforms, across all species, indicating their significant role in KATP channel regulation. Conformational changes caused phosphorylation are believed to increase the stability of the open channel state. In contrast to SUR2 isoforms, phosphorylation of SUR1 (Ser1448 or Ser1571 of NBD2) leads to inhibitory effects on KATP channel conductance.[55]

KATP channels are regulated by a number of molecules from a series of biochemical pathways. For example, acetyl coenzyme A ester groups stimulate KATP channel activity, along with phosphatidylinositol-4,5-bisphosphate.[56, 57] The signaling protein syntaxin displays inhibitory effects on channel gating.[58] In cardiac muscles, lactate dehydrogenase interacts with

NBD2 to promote channel opening.[59] Thus, a large number of regulators for channel gating have been identified, although the regulatory mechanisms by which they operate remain largely unknown.

Figure 10: Mutation and Phosphorylation Sites in SUR2A NBDs The localization of mutation (blue) and phosphorylation (red) sites in SUR2A NBD1 (A) and NBD2 (B) are shown above. The above homology models were generated using Modeller as shown in Section 2.2.

1.9 Cardiac Disease-Causing Mutations in SUR2A

To date, four disease-causing mutations have been identified in the SUR2A NBDs, with three occurring in NBD2 and one in NBD1.[42, 60-62] The localized regions of these mutations are depicted in Figure 10. Very limited structural and biochemical data exists for any of these disease-causing mutants.

Mutation of Val734 to isoleucine in NBD1 has been correlated to myocardial infarctions.

Homology modelling of the NBDs, based on the X-ray structure of the highly related CFTR

NBD1, suggests that Val734 exists in a loop at the interface between NBD1 and coupling helix

1. Although this is a relatively conservative mutation, it may compromise interactions at the interface and limit communication between various domains.[63, 64]

Mutation of Thr1547 to isoleucine in NBD2 leads to atrial fibrillations. Note that his mutation occurs within the C-terminal 42 residues of the protein. Electrophysiology experiments have elucidated that mutants still preserve proper ATP-induced channel closure, which results from ATP binding at the Kir6.x subunits. However, mutants were also observed to possess 17

decreased responses to activation by Mg2+-ADP. The C-terminal region of NBD2 has already been established to play a significant role in reducing channel activity through possible interactions with the ATP binding site. The introduction of a hydrophobic residue in place of the more polar threonine may disturb nucleotide interactions and lower binding affinities for ATP and ADP, which result in decreased channel opening.[60,63,64]

The remaining mutations in NBD2, which include a frameshift mutation at Leu1524 and substitution of Ala1513 to tyrosine, result in a lower presence of KATP channels at the cell membrane. These mutations likely compromise correct folding of the SUR protein and impair assembly of a functional KATP channel. Alternatively, misfolded regions may lead to problems during trafficking to the cell membrane.[60]

Mutations also occur in SUR1 (which is found in pancreatic KATP channels). SUR1 mutations are mainly associated with persistent hyperinsulinemic hypoglycemia of infancy

(PHHI) and type II diabetes. There are approximately 200 mutations located throughout the primary sequence of SUR1, with just under half of them leading to PHHI. A number of loss-of- function mutations are also related to mutations in the 42 C-terminal residues of NBD2 and lead to closed KATP channels and decreased insulin levels and diabetes.[62, 65]

The underlying molecular mechanisms and resulting biochemical effects behind these mutations are poorly understood. This hinders the development of specific drugs to target and possibly rectify specific interactions that are altered in disease. Therefore, the study of the wildtype SUR NBDs and their biophysical properties as well as how specific properties are modulated in the disease state is of significant clinical importance.

1.10 Biophysical Methods for Investigation of SUR2A

1.10.1 Nuclear Magnetic Resonance

NMR spectroscopy is an information-rich technique which, similar to X-ray crystallography and electron microscopy, can provide an atomic-level understanding for the structure and orientation of a macromolecule.[66] Moreover, solution-state NMR offers insight into molecular mobility, which is of great importance in evaluating conformational changes and dynamic mechanisms of protein activity and regulation. NMR spectroscopy probes transitions between various spin states of magnetically active nuclei (I ≠ 0, where I is the nuclear spin quantum number). These nuclei possess a spin angular momentum (L) which has a specific magnitude and direction. In the absence of a magnetic field, there is no preferred orientation for the spin angular momentum which results in degenerate energy states.[66] However, introduction of a directional magnetic field, results in a difference between energy states of nuclear spins. The number of energy states is related to the spin quantum number, but for biologically relevant atoms (1H,13C,15N) where I=½, the number of magnetically induced energy states is two. The energy separation (ΔE) between such states is related through the equation:

(1) where h is Planck’s constant, γ is the gyromagnetic ratio and Bo refers to the strength of the magnetic field. [67] The gyromagnetic ratio adopts a characteristic value for each nuclei, with larger values belonging to more sensitive nuclei (Table 3). The energy separation between different spin states is also dependent on the intensity of the magnetic field, with strong magnetic fields resulting in greater separations. However, thermal motions are often sufficient to overcome this energetic separation, resulting in only a slight difference in energy-level populations.

Therefore, NMR is an inherently low sensitive technique.[67] Transitions between different

energy levels occur through the absorption of a radio-frequency pulse and can be illustrated through plot of intensity versus the resonant energy of absorption (chemical shift).

Table 3: Gyromagnetic ratios of some NMR active nuclei[67] Atom γ (10-7 T-1s-1) 1H 26.75 13C 6.73 15N -2.71 19F 25.18 31P 10.84

NMR resonances from a spectra possess four fundamental parameters: intensity, resonance frequency, line splitting and line width.[67] The volume of a peak (intensity) is related to the concentration of the particular sample, as well as the dynamics of the molecule. Increased concentrations result in increased signal-to-noise ratios and yield more intense peaks. However, molecular motions may limit the observed intensity of a peak, due to dynamics on a time-scale that causes broadening. The resonance frequency for a particular nucleus (chemical shift) primarily depends on the local chemical environment. Nuclei in proximity to electron- withdrawing substituents, are less shielded from the external magnetic field (Bo) and resonate at higher frequencies. The reverse is also valid which provides insight into the local chemical environment of a specific atom. Splitting of chemical frequencies (coupling) results from magnetic interactions of nuclei situated close in space, or through chemical bonds. Although coupling between nuclei can yield highly rich information in macromolecular structure, for larger proteins it generally convolutes the overall spectra. Therefore, is often removed through decoupling pulse sequences in protein NMR experiments. The line-width at half-height

(broadness of a peak) is associated with the tumbling time of the molecule or spin-spin relaxation

(T2). Larger molecules foster more interactions with the solvent and therefore, slower rotational dynamics. This results in shorter values of T2 and line broadening.[67]

One significant NMR pulse sequence for protein experiments is the heteronuclear single quantum coherence experiment. In a 15N-1H HSQC, the resonant frequency of an amide proton is correlated to the resonant frequency of the directly attached amide nitrogen. Ideally in the 2D

NMR spectra of a 15N-labelled protein, each peak in the HSQC corresponds to each N-H pair, which includes all backbone and side-chain amides. Therefore, a resonance is observed for all amino acids, except proline.[66] Based on T2 relaxation rates, there is an upper limit to the macromolecular sizes that can be suitably studied through the HSQC sequence (approximately

20 kDa). However, there are a number of techniques which exist to elevate this molecule-mass ceiling. Some methods include removal of proton coupling by the use of deuterated samples (2H) or by employing a transverse-relaxation optimized sequence (TROSY).[68] In a traditional 15N

HSQC, scalar-spin coupling generally harvests four peaks which collapse into a single peak following broadband decoupling. However, during the TROSY sequence, decoupling is not performed. Instead, only the sharpest and most slowly relaxing transition is selected from the multiplet and observed. The TROSY sequence allows for greater resolution for larger macromolecular complexes. It greatly improves the spectra of large proteins without the use of deuterated samples, which possesses several biochemical advantages during expression and purification of biomolecules.

A distinct disadvantage of employing NMR spectroscopy for this study is the high quantities of protein sample required for analysis. High concentrations introduce vulnerability to protein aggregation and precipitation, especially at elevated temperatures. These obstacles coupled with low solubility of many proteins, especially mammalian NBDs, present challenges in sample preparation and prolonged data acquisition. However, multidimensional NMR has been successfully performed on a number of ABC transporters, such as the NBDs of MRP1 and murine CFTR.[69, 70] Furthermore, NMR spectral analysis of CFTR NBDs has been

instrumental in elucidating regulatory functions of various structural elements. Furthermore, noticeable spectral changes upon ATP binding and hydrolysis, as well as following post- translational modifications, such as phosphorylation, have expanded information into the dynamics of NBD activity. Such studies speak to the far-reaching strengths of NMR spectroscopy and its applicability to the understanding of SUR2A NBDs.

1.10.2 Circular Dichroism

Circular dichroism is a bioanalytical technique that can be utilized to provide information on the global folds of a macromolecule, such as the secondary and tertiary structure of a protein.

Thus, it can be employed to examine how specific modulations to a macromolecule, such as phosphorylation or temperature change, may instigate a macro-scale change to protein conformation. The signal in circular dichroism originates from differential absorption of the left and right components of plane polarized light. A molecule that is chiral will absorb one component of circularly polarized light to a greater extent, resulting in elliptically polarized light,

historically displayed as a positive or negative

ellipticity. Furthermore, absorption of light by a

particular molecule varies with the wavelength of

light. Therefore, monitoring the absorption of

circularly polarized light as a function of the

wavelength of light results in unique absorption

spectra. [67]

Nineteen of the twenty common amino

Figure 11: Circular dichroism spectra acids are chiral which results in CD-active protein of common secondary structure features [2] subunits. However, unlike UV-visible spectroscopy,

the signal of the CD spectra for a macromolecule (such as a protein) is not the sum of the CD spectra for the individual subunits (in this case amino acids). Instead, CD spectra are more significantly influenced by higher order structures in three-dimensional space. A specific structure will lead to a specific spectrum allowing for identification of certain secondary structure elements based on known models. An illustration of the most widely used circular dichroism spectral signatures is shown in Figure 11. Alpha helical structures typically display characteristic spectra with negative bands between 203-240 nm and local minima around 209 and 222 nm. Furthermore, there is usually a strong positive band below 203 nm with a local maxima around 192 nm. Beta sheets produce less intense bands, with minima between 210 to

225 nm and maxima in the range of 190 nm to 200 nm. Disordered structures produce strong negative bands around 200 nm, with weak signals above 210 nm. [71] Therefore, CD offers complementary information to NMR and is useful in studying the SUR NBDs.

1.10.3 Fluorescence Spectroscopy

Fluorescence is a relaxation process that results in the emission of light and can be used to probe conformational changes. The fundamental principles of fluorescence are delineated in

Figure 12. In a simplistic model, the orbital electrons of a molecule can exist in a ground-state energy level (Go) or a higher energy electronic level (S1S2,S3, etc), each with multiple vibrational levels.[67] Discrete amounts of energy are required to excite a molecule from the ground state to any of the vibrational levels in a higher energy level. Following excitation, vibrational energy is rapidly dissipated as heat, through molecular collisions with the solvent. This leads to the molecule falling to the lowest vibrational state in S1. An excited molecule will return to the ground state through the emission of light (fluorescence). An excited molecule can relax to any

of the vibrational states in

Go. The probability of falling

into a particular vibrational

level, determines the shape of

the fluorescence spectra. [67]

Not all molecules

fluoresce, and natural

fluorophores are quite rare.

If the excited state of a

Figure 12: Possible excitation and relaxation pathways of a molecule overlaps with the chromophore with light Light of resonant energy wavelength is absorbed by the molecule resulting in an excited energy state with a characteristic absorption vibrational levels of the spectra. Emission of this energy may occur through fluorescence, where light is released rapidly, and the molecule relaxes to the ground state, a molecule will ground state. Phosphorescence occurs due to an intersystem crossing from a singlet to a triplet state, and the relaxation process relax through non-radiative occurs on a significantly slower timescale.[3]

transitions from S1 to Go and no fluorescence will be observed. Generally, rigid molecules with limited vibrational states such as aromatics, exhibit fluorescence. In proteins, tryptophan, phenylalanine and tyrosine all contribute to fluorescence. Since the excitation spectra of tryptophan, phenylalanine and tyrosine are not identical, different amino acids can be selectively targeted by varying the wavelength.

For example, tryptophan can be selectively targeted through excitation at 295 nm.[72]

The utility of fluorescence arises from the sensitivity of fluorophores to the environment.

For example, a fluorophore which is exposed to solvent may emit lower intensities of fluorescence due to collisional quenching by the solvent molecules. This occurs due to draining energy away from the excited state through heat via collisions. Tryptophan residues which are

buried in the hydrophobic core are generally shielded from collisional quenching. Unfolding of a protein however, may expose these residues to solvent and lower the fluorescence detected.

Artificial quenchers can be introduced into solution to probe surface accessibility of the fluorophore to the quencher. The changes in fluorescence intensity with variations in quencher concentrations, Q, are generally described by the Stern-Volmer equation:

(2) where Fo and F refer to the observed fluorescence intensities in the absence and presence of the quencher respectively. K refers to a constant that is the product of the bimolecular quenching constant, kq, and the lifetime of the excited state in the absence of the quencher, τo.[72]

Quenching of tryptophan fluorescence in proteins often produces Stern-Volmer plots which deviate from linearity at higher quenching concentrations. Modifying the Stern-Volmer relationship yields equation (3):

(3) where fa refers the fraction of the fluorophore population that is accessible to quenching and ΔF refers to the difference in fluorescence in the presence and absence of the quencher. Modified

Stern-Volmer plots are useful for probing the exposure of tryptophan residues in a protein and thus provide unique information on the secondary and tertiary conformations of a macromolecule.[72]

Chapter 2 Materials and Methods

2.1 Structure-Based Sequence Alignment of the SUR NBDs†

In order to generate soluble proteins of the SUR NBDs, the N- and C- terminal boundaries needed to be identified so that all structured regions are included in the construct. Selection of the N- and C- terminal boundaries involved the use of a structure-based sequence alignment, generated with ClustalW (Version 2.1). The alignment included the NBD sequence from every known ABCC transporter, as well as the sequences of the NBDs from yeast ABCC protein

Ycf1p, and sequences for all NBDs from which the structure was solved. Secondary structure masks were used from X-ray structures of CFTR NBD1 and TAP NBD1. ClustalW employs an algorithm which aligns input sequences separately and formulates a distance parameter based on the identity between the two sequences.[78] The distance parameters are then used to generate alignments. The penalty for the introduction of gaps into alpha helices and beta strands was set to

9 (highest value on a scale of 0-9). In contrast, the penalty for introducing gaps into unstructured regions was minimized (set to 2). These settings allowed for stringent identification of loops and to minimize the expected presence of insertions and deletions in secondary structures.[78]

†Experiments were conducted by Dr. V. Kanelis

2.2 Homology modeling of SUR2A NBD1†

Homology models of rat SUR2A NBD1 were generated using the program Modeller by employing the structure-based sequence alignments. The crystal structure of NBD1 from CFTR was used as a template. Fifty models were generated, and the 10 lowest energy models were selected for analysis. The quality of the homology models folds was evaluated by ProsSA and 26

PROCHECK and was used to gauge the stereochemistry and overall energy of the models.[79,80]

†Experiments were conducted by Dr. V. Kanelis

2.3 Generation of Recombinant NBDs

A series of domain boundaries were selected and tested based on the solubility of SUR2A

NBD1. The various boundary domains, were selected in a such a way to add or remove putative secondary structure elements and assess the ramifications on stability and structure. For this study, cDNA of rat SUR2A was employed as the template for polymerase chain reactions (PCR).

The selected region with the desired N- and C- terminal boundary domains was targeted and amplified using traditional PCR methods. The target sequence was cloned into the pET SUMO vector (Invitrogen, modified) using restriction digestion enzymes, NcoI and XhoI. Constructs with minimal amino acid differences, such as shortening the N terminus from S615 to T618, were generated by mutagenesis (QuikChange, Agilent). The primers utilized for all constructs are depicted in Table 4. All constructs were sequenced in the forward and reverse directions by

ACGT or TCAG Corp.

Table 4: Primers used in the PCR amplification of SUR2A NBDs Restriction cut sites are underlined in the primers, and mutated sites are coloured red Protein Boundary Restriction Primer Enzyme SUR2A 5 S615 NcoI 5 CAT GCC ATG GGA AGC TGG AGG ACT GGG 3 NBD1 SUR2A 3 K972 XhoI 5 ATG AGG CTC AGG ACG AAG TGA CTC GAG CCG 3 NBD1 SUR2A Fwd 5'GTAGTACTGGATCCATGGGAACTGGGGAGGG3' 5 T618 Rev 5'CCCTCCCCAGTTCCCATGGATCCAGTACTAC3' NBD1 Fwd SUR2A 5'GAAGGAAAAGTTTACTGGAACAA

TATAAATGAATCTGAGCCTTCTTTG3' Rev NBD1 Val729Ile 5'CAAAGAAGGCTCAGATTCATTTAT

ATTGTTCCAGTAAACTTTTCCTTC3'

SUR2A Fwd P1299Del Rev 5'-CTGGATCCATGGGACGAGCATTGGCCACAG-3' NBD2 5'-CTGTGGCCAATGCTCGTCCCATGGATCCAG-3' SUR2A Fwd 5'-TTAGTCTTTTCTGAGGGTATTAAATGCGG Fs1524- GGTCTAGTTAGTGGAGTGCGATACTGG-3' NBD2 Rev STOP 5'-CCAGTATCGCACTCCACTAACTAGACCC CGCATTTAATACCCTCAGAAAAGACTAA-3' SUR2A Fwd Thr1547Ile Rev 5'-CTTTTCTACTTTGGTGATGATCAACAAGTAGCTCGAGTGAA-3' NBD2 5'-TTCACTCGAGCTACTTGTTGATCATCACCAAAGTAGAAAAG-3' SUR2A Ala1513Thr Fwd 5'-ACCGTGTCTCCTCTATTATGGATACGGGCCTTGTTT-3' NBD2 Rev 5'-AAACAAGGCCCGTATCCATAATAGAGGAGACACGGT-3'

In this study, rat cDNA was used as the model due to ease in availability compared to human

SUR2 cDNA. However, between species, there are only 10 conservative amino acid differences between rat and human SUR2A NBD across all species. Furthermore, all phosphorylation sites and disease causing mutations are conserved, and therefore, any biochemical data obtained, should be directly applicable to the human SUR2A NBDs as well.

2.4 Expression and Purification of SUR2A NBD1

Constructs containing various SUR2A NBD1 proteins were expressed as N-terminal-6xHis-

SUMO fusion proteins using a modified pET-HisSUMO vector (Novagen). Proteins were overexpressed using E. coli BL21 (DE3) CodonPlus®-RIL competent cells (Stratagene).

Expression was carried out either in LB broth or M9 minimal media with isotopic enrichment for

NMR studies as necessary. Chloramphenicol (34 mg/L) and kanamycin (50 mg/L) were added

to the media to maintain selective pressure on the plasmid. The cells were grown at 37oC in a orbital shaker set at 250 rpm. Once the cell growth reached to an OD600 of 0.8, the incubating temperature was reduced to 18oC and gene expression was induced with 0.75 mM isopropyl β-D- thiogalactoside (IPTG). Following a post-induction period of eighteen hours, the cultures were harvested by centrifugation (approximately 5000g for 15 min) and the cell pellets were stored at

-20oC while the supernatant was discarded.

Protein purification was carried out at 4oC to minimize aggregation and protease activity.

The overall procedure is similar to purification methods used for the NBDs of CFTR. [70] Cell pellets were thawed and resuspended in lysis buffer (50 mM tris pH 7.6, 100 mM L-arginine,

120 mM NaCl, 2 mM β-mercaptoethanol, 2 mM ATP, 5 mM MgCl2, 12.5% (v/v) glycerol, 0.2%

(v/v) Triton X-100, 2 mg/ml deoxycholic acid, 1 mg/ml lysozyme, 5 mM 6-aminocaproic acid, 5 mM p-aminobenzamidine, and 1 mM PMSF). Resuspension volumes for cell pellets were based on the final optical density obtained during the growth phase. The cells were diluted in approximately 10 ml per litre of culture with an OD600 of 1.0 as measured prior to culture harvesting.

Cells were lysed by sonication (no pulsing) at an intensity setting of 12 on a Heat

Systems Inc sonicator equipped with a microprobe (30 sec on/30 sec off, 6 cycles). Cellular debris was removed by pelleting through centrifugation (14,568g, 30 min). The cell pellets were resuspended in fresh lysis buffer. Sonication and centrifugation were repeated to maximize protein recovery. The cell lysate from each sonication cycle was filtered and loaded onto a high performance Ni2+-NTA affinity column (GE Healthcare) pre-equilibrated with 20 mM tris pH

7.6, 150 mM NaCl, 5 mM imidazole, 12.5% (v/v) glycerol. The column was washed with 10 column volumes of equilibration buffer and eluted with 20 mM Tris pH 7.6, 150 mM NaCl, 500 mM imidazole, 12.5% (v/v) glycerol, 2 mM ATP, 5 mM MgCl2 and 2 mM β-mercaptoethanol.

Generally a 5 ml Ni2+-NTA column was used and elution volumes were 5 ml each. Fractions containing the protein of interest were pooled and diluted with cold gel filtration buffer (20 mM tris pH 7.6, 150 mM NaCl, 12.5% (v/v) glycerol, 2 mM ATP, 5 mM MgCl2, 2 mM β- mercaptoethanol) such that the imidazole concentration was approximately 150 mM. Ulp1 enzyme was then added to the diluted fractions at approximately 12 µg/ml, to cleave the 6×His-

SUMO tag from the NBD1 protein. Dilution of the elution fractions was required to lower imidazole concentrations for activity of Ulp1 enzyme. The diluted solution was concentrated to

3-4 ml through centrifugal filters (Millipore) at 1258g for approximately 10 hours. The concentrated protein mixture was loaded onto a 24 ml Superdex 75 gel filtration column, pre- equilibrated with the cold gel filtration buffer.

Fractions containing NBD1 (eluted from the gel filtration column) were collected and run over Fast-Flow Ni2+-NTA affinity resin (GE Healthcare) to remove any residual 6x-His-SUMO and contaminating Ulp1 protease. The presence of NBD1 was assessed at multiple stages throughout the purification process by 15% SDS-PAGE gels. Non-reducing gels were also run to assess the presence of dimeric or higher order structures of the pure protein caused by disulphide bond formation. Protein concentration values were assessed using Bradford assay estimation techniques and confirmed via amino acid analysis. For NMR studies, the purified NBD1 protein was dialyzed into an NMR buffer containing 20 mM sodium phosphate, pH 7.25, 150 mM NaCl,

5 mM MgCl2, 2 mM DTT, 2 mM ATP, 2 % (v/v) glycerol. Sample lifetime was approximately 2 weeks at 4oC, although variations based on the protein sample were observed.

2.5 Phosphorylation of NBDs

Phosphorylation reactions were conducted in 50 mM tris, pH 7.4 with 50 mM NaCl, 50 mM

MgCl2, 50 mM ATP and 5 mM DTT. Reactions contained approximately 15 mg of target protein

in 10 ml of buffer. PKA was added at an activity level of 100 units per ml. All reactions were allowed to proceed for at least 12 hours at 4oC. Samples were then dialyzed into the appropriate

NMR buffer. Phosphorylation of specific proteins was qualitatively examined by staining an

SDS gel with Phosphogel stain (Molecular Probes) and visualizing the protein bands. This stain is specific for binding to phosphorylated proteins. Quantitative analysis also involved trypsin digesting the protein and examining the size of the resulting fragments through mass spectrometry. This allowed us to identify the sites of phosphorylation.

2.6 NMR Spectroscopy

15N-1H TROSY-HSQC spectra were recorded on a 600 MHz Varian Inova spectrometer (Varian

Inc. Palo Alto, CA) equipped with a H(F)CN triple resonance cryoprobe and actively-shielded z- gradients. Sample volumes ranged from 450 to 550 µL. All spectra were referenced with 4,4- dimethyl-4-silapentane-1-sulfonic acid (DSS). The NBD1 protein concentration varied from 0.05 mM to 0.5 mM for different samples (WT, mutants, ±ATP, ±phosphorylation, The temperature of at which data was collected also varied. Data was processed using NMRPipe and NMRDraw.

The resulting spectra were analyzed and peaks were manually counted using NMRView.

2.7 Circular Dichroism Spectroscopy

Circular dichroism (CD) spectra of purified SUR2A NBD1 were recorded from 180 nm to 300 nm at 22ºC on an Aviv 250 CD spectrometer (Aviv Biomedical Inc., Lakeview, New Jersey) with a bandwidth of 0.2 nm using a 0.1 cm path length quartz cell. Samples contained 5 μM

NBD1 in 20 mM phosphate, pH 7.25. Spectra were averaged from 3 scans and were baseline- corrected by using a blank consisting of buffer only.

2.8 Fluorescence Spectroscopy

Fluorescence experiments were conducted on a Fluoromax 4 Model equipped with a Peltier unit for precise temperature regulation. Fluorescence experiments were conducted by selectively exciting tryptophan residues at 295 nm (with an emission slit width of 1 nm). Emission spectra were collected at varying wavelength intervals with a slit width of 2.5 nm. During temperature melt experiments, samples were allowed to equilibrate at the measuring temperature for 60 seconds. Melts were performed by gradually increasing the temperature from 5oC to 95oC.

Experiments were also conducted from 95oC to 5oC to assess reversibility. Iodine quenching experiments were conducted with fresh stock solutions of 3M potassium iodide (Sigma). A small

- quantity of Na2S2O3 was added to the KI stock solutions to prevent formation of I2 and I3 species which would have interfered with quenching experiments. Samples contained 2 μM NBD1 in 20 mM phosphate, pH 7.25. Spectra were averaged from 3 scans and baseline-corrected. Quenching experiments were repeated with increasing concentrations NaCl, in place of KI, to assess effects based on ionic strength.

Chapter 3 Results

3.1 Structure-Based Sequence Alignment

The N-terminal and C-terminal boundaries of the nucleotide binding domain of rat SUR2A were predicted from a structure based sequence alignment. Included in the alignment were the sequences of NBDs from all human ABCC proteins, the yeast ABCC protein Ycf1p, as well sequences of NBDs for which the structure has been solved. A portion of the alignment is illustrated in Figure 13. Secondary structure masks were employed from structures of CFTR

NBD1 and the NBD of TAP1. This allowed secondary structure elements to be taken into account in the alignment so that insertions and deletions are included in loops and not in - helices and -strands. The alignment depicts the presence of a number of well-conserved residues, even in conventionally diverse regions. For example, the occurrence of an aromatic residue (Phe625) has been identified C-terminal to the first β-strand. This conserved aromatic amino acid is thought to allow for π-π stacking interactions with ATP to enhance binding to the nucleotide. Furthermore, there is strong agreement between motifs involved in ATP binding and hydrolysis, such as the Walker A and Walker B, as well as the ABC signature sequence

(differentially coloured in Figure 13). In addition, our alignment shows good agreement between secondary structure elements between NBDs for which the structure has been solved (shaded residues). However, the low sequence identity in N- and C- terminal regions introduces ambiguity in assigning the precise domain boundaries.

Figure 13: Structure-based sequence alignment of SUR2A NBD1 with ABCC transporters A structure based sequence alignment of SUR2A NBD1 and other ABCC transporters was generated through ClustalW. Displayed above the alignment is the 2o structure of CFTR NBD1, which was used in the structure based sequence alignment. Residues belonging to α-helices, β sheets and 310 helices are shown as grey cylinders, arrows and open circles respectively. Known secondary structure elements are coloured in the sequences as blue, violet and green for α helices, β sheets and 310 helices respectively. The full alignment is shown in the appendix.

An interesting observation arising from the alignment (Figure 13) is the presence of a large insert (S625-F675) sandwiched between β-strands β-1 and β-2. A similar β-sheet subdomain insertion of amino acids was originally identified in CFTR. Known as the regulatory insert (RI) in CFTR, the insertion was previously believed to be exclusive to CFTR and absent in other ABC transporters. However, our alignment indicates that this insertion is present in NBD1 from a number of ABCC transporters. There are many similarities between the CFTR NBD1 RI and the β-sheet subdomain insert in SUR2A NBD1. Residues Q635-E664 of the insertion in

SUR2A, predicted to be disordered by PONDR, align well with the corresponding insertion in

CFTR suggesting a comparable role for this subdomain. Further, both the SUR2A NBD1 -sheet subdomain insert and the CFTR NBD1 RI possess a site for phosphorylation by protein kinase A

(PKA). Phosphorylation of the RI in CFTR NBD1 alters the conformation of the protein and is necessary to promote interactions with coupling helix 1. [70] Thus, the β-sheet subdomain insert is of interest due to its highly characterized role for regulation in CFTR. Therfore the β-sheet subdomain insert in SUR2A NBD1 may have regulatory properties similar to NBD1 of CFTR.

One of the physical consequences of this prediction is that the N-terminal boundaries for

NBD1 are likely to be further upstream of what current literature suggests. Furthermore, the C- terminal region of CFTR possesses an additional regulatory element, which may suggest for analogous regulatory role for residues beyond the current C-terminal domains.

3.2 Generations of Constructs with Variable Domains for NBDs

Regions encompassing different boundary domains from rSUR2A template DNA were amplified and cloned into a pET-HisSUMO vector. Culture growths of 50 ml allowed for small scale tests of protein expression. The expression profiles following induction by IPTG are depicted in

Figure 14. Overexpression of protein products from all constructs, except for D665 to E889 was 35

observed following induction. NBD1 S615-K972 depicted lower levels of overexpression compared to proteins with shorter boundary domains. Moreover, Q600-L933 migrates at a noticeably lower molecular weight in the SDS PAGE, which may indicate a possible change in conformation or partial degradation in the cell. In addition to high levels of expression, a prerequisite for an ideal construct required the translated protein product to be soluble in E. coli cells. Solubility of these proteins was examined by resuspending cell pellets in lysis buffer Q600-L933 S608-L933 S615-L933 T618-L933 S615-D914 S615-K972 V669-G889

Ladder Pre Post Pre Post Pre Post Pre Post Pre Post Pre Post Pre Post kDA

98 64 Ladder Pre Post Pre Post Pre Post Pre Post Pre Post Pre Post Pre Post 50

Figure 14: Protein profile of pre-and post-induction for proteins of various domain boundaries Samples were prepared by spinning down a volume of cells that corresponded to an OD600 of 1.0 in 1 ml, and resuspending the pellet in a final volume of 100 microliters (including sample buffer). Five microliters of each sample was loaded into each lane Q600-L933 S608-L933 S615-L933 S615-D914 S615-K972 V669-G889 kDA Ladder P1 S1 P1 S1 P1 S1 P1 S1 P1 S1 P1 S1 98 64

Figure 15: Localization of expressed protein to soluble or insoluble phases Cell pellets were resuspended in 1 ml of lysis buffer and sonicated. The cellular debris was pelleted by centrifugation. Gel samples from the supernatant (S1) and pellet (P1) were loaded above. 36

followed by sonication. Pelleting of cellular debris from the cell lysate allowed for assign protein expression to the soluble or insoluble phase. The protein distribution into each of these phases is shown in Figure 15.

Preliminary work indicated greater solubility of NBD1 SUR2A S615-L933. All of the constructs were expressed in large-scale growths, purified as described in the materials and methods section. The resulting protein yields are summarized in Table 5. Despite, higher yields of S608-L933, previous work has delineated the formation of higher order structures which are less ideal for further biophysical characterization. Therefore, the region of S615-L933 was selected as the putative boundaries for the first nucleotide binding domain of SUR2A.

Table 5: Total protein yields of various NBD1 SUR2A constructs (Yields are based on at least 2L of culture growth) Construct Total Yield S608-L933* 80 mg/L in LB S615-L933 60 mg/L in LB, 15 mg/L in M9 S615-D914* 3 mg/L M9 T618-L933 6 mg/L M9 S615-K972 7 mg/L M9 *Protein expression determined by Lynn Ikeda

3.3 Expression and Purification of SUR2A NBD1

To overcome previous obstacles to soluble eukaryotic NBD biosynthesis, a number of precautions were pursued for SUR2A NBD1 expression and purification. Protein expression of rat SUR2A was performed through BL21(DE3) Codon-Plus-RIL cells. These competent cells are useful for expression of mammalian proteins, since they possess plasmids encoding specific mammalian tRNA genes. These tRNAs recognize codons which are uncommon to bacterial cells but common to mammalian cells. Therefore, they provide a method for enhancing mammalian protein synthesis in E. coli hosts.

To augment soluble protein expression, SUR2A NBD1 constructs were cloned into a

vector containing an N-terminal solubility enhancing factor (SUMO tag). The yeast SUMO tag is

an ubiquitin-like protein which has been shown to improve the solubility of SUMO-coupled

proteins. Furthermore, cold induction of NBD1 gene expression (18oC), allowed for slower rates

of protein translation. This allows the opportunity for proper folding of the NBDs and more

soluble expression. However, cold inductions usually result in decreased protein expression

compared to growths at higher temperatures.

kDA

Figure 16: Purification of NBD1 S615-L933 Gel samples based on different stages in the purification process. Pre and Post are samples of cell culture before and after induction. The soluble (S1) and pellet (P1) fractions are shown in following one cycle of sonication and following a second cycle (S2 and P2). The flowthrough (flow) and wash (wash) are samples of following the passing the cell lysate over Ni2+ resin. Samples from 5 ml elutions are shown in 1, 2, 3, 4 and 5. kDA Ladder 0s 15s 20s 25s 30s 35s 40s 45s 50s 55s 60s 1h

98 64 50 NBD1-SUMO

NBD1 36

16 SUMO

Figure 17: Time Course of Ulp1 Activity on NBD1 S616-L933 A sample of 10 mg/ml NBD1- HisSUMO was incubated with Ulp1 protease. Ulp1 protease was added at time 0s. The reaction was stopped by the addition of SDS buffer and heating to 95oC. 38

NBDs were expressed with an N-terminal 6×histidine tag which allowed for initial purification by nickel-affinity chromatography. The use of a 5 ml of nickel resin was shown to be sufficient for purification of two litres of culture supernatant for all constructs (Figure 16). A majority of the protein retained by the Ni2+-NTA, was eluted following one column volume of elution buffer containing 500 mM imidazole. Washing the column with higher concentrations of imidazole to reduce non-specific binding also resulted in NBD1 co-elution (data not shown). Elutions containing the protein of interest were pooled. Following dilution, Ulp1 protease was added to cleave the solubility enhancement factor (SUMO). Ulp1 is a protease that recognizes a specific tertiary structure in SUMO, as opposed to an amino acid sequence. This results in limited non- specific activity and degradation of NBD1. A time course for cleavage by Ulp1 is depicted in

Figure 17. The protein mixture was concentrated and loaded onto a Sephadex-75 gel-filtration column, and the resulting fractions collected were run on a gel as shown in Figure 18A. The fractions containing the desired protein were collectively run over a second Ni2+-NTA column to selectively bind SUMO and other contaminants (reverse Ni2+-affinity chromatography). The resulting protein sample was isolated in a fairly pure state, as shown in Figure 18B.

Ladder 1 2 3 4 5 6 7 8 9 10 11 12

kDA 98 64 50

36 NBD1 (S615-L933)

Figure 18: Fractions following gel-filtration Samples from each of the fractions from the gel filtration column are shown above in a 1/1000 dilution. The final pure product is shown in B. 39

3.4 Biophysical Characterization of SUR2A NBD1

3.4.1 NMR Studies of 15N labeled S615-L933

15N-1H TROSY Heteronuclear single quantum coherence (HSQC-TROSY) spectra were recorded for various SUR2A NBD constructs. A

3.4.1

Figure 19: 2D 15N-1H TROSY-HSQC spectra of NBD1-SUR2A S615-L933 at 30oC Sample concentration was approximately 0.363 mM (measured through amino acid analysis). The NMR sample conditions were 20 mM Na+ phosphate (pH 7.25), 150 mM NaCl, 2 mM ATP, 2 mM DTT, 5 mM MgCl2 with 2% glycerol and 2% D2O (v/v). A) shows the full 2D spectrum is shown with black as positive and red as negative peaks. The red box outlines suspected tryptophan indoles, B) shows intensities of select peaks from A) in a one-dimensional NMR spectrum. 40

resonance is observed for all N-H pairs in proteins. Therefore, resonances are observed for backbone N-Hs for all non-proline residues as well as for side-chain N-Hs. The dispersion of resonances in the proton dimension indicates the presence of discrete chemical and structural environments for 1H protons.

The 2D 15N-1H TROSY-HSQC spectra of NBD1-SUR2A S615-L933 (~0.36 mM) is depicted in Figure 19. The dispersion of chemical resonances in the proton dimension from 6.5 ppm to 10.9 ppm indicates that protein is folded. Diluted NMR samples (~0.05 mM) yield superimposable spectra indicating that elevated concentrations do not promote the formation of dimeric or higher-order complexes (data not shown). Although the 2D-HSQC has not been assigned, certain regions can be distinguished. For example, the six resonances suspected of arising from indoles of the six tryptophan residues are outlined in Figure 19A. The asterisked, red–coloured resonances situated between 116 ppm and 120 ppm in the 15N dimension are caused by spectral aliasing. These likely arise from epsilon N-H atoms of arginine residues.

Glycine residues usually possess lower nitrogen field resonances and also aliased in the above spectra (shown by +). All chemical shifts were referenced to 4,4-dimethyl-4-silapentane-1- sulfonic acid (DSS).

Differential peak intensities are found throughout the spectra, as indicated by the traces in

Figure 19B. The differences in cross peak intensities are the result of differences in dynamics of various nuclei. Thus, the differential intensities indicate a conformationally dynamic protein.

Strongly intense peaks in the center of the spectra may arise from disordered loops, such as the

β-sheet subdomain. Weaker intense peaks are further spread out in the 1H dimension and arise from structured regions of the protein that may undergo conformation exchange on the μs-ms timescale.

We have additional evidence that exchange processes are occurring in SUR2A NBD1.

The number of NH cross-peaks observed for NBD1 SUR2A S615-L933 is a function of temperature with clear distinctions in the spectra from 15oC and 30oC (Figure 20A-D), with increasing peaks at 30oC. Two-dimensional NMR experiments on NBD1 were less feasible at temperatures beyond 30oC due to excessive precipitate formation. Apparent from Figure 19, is the increased resolution in the number of NH cross peaks which can be resolved at higher temperatures. The number of cross peaks resolved at 30oC was 304 compared to the 311 expected. Increases in peaks at higher temperatures may result from greater sensitivity due to faster tumbling. However, the non-uniformity of the resonances and the fact that some peaks disappear at 30oC compared to lower temperatures (such as the aliased arginine residues) indicates that SUR2A NBD1 has significant protein dynamics.

Conformational changes upon temperature adjustment were shown to be fairly reversible, at least over small temperature ranges. This was assessed through qualitative examination of spectra, as well as semi-quantitatively, through intensity plots of specific peaks as in Figure 19

B. Interestingly, spectral changes with increases in temperature are coupled with the appearance of all six tryptophan indoles at 30oC. Correspondingly, the decrease in prominence of the arginine crosspeaks occurs with increases in temperature. These changes may represent shifts between equilibria of two possible populations of states.

Table 6: Number of Amide Proton Crosspeaks in NBD1-S615-L933 (Based on Spectra in Figure 19) Spectra Number of Cross-peaks 15oC 219 20 oC 279 25 oC 288 30 oC 304 Expected Number of Cross-peaks:311

A B

C D

Figure 20: 2D 15N-1H TROSY-HSQC of NBD1-SUR2A S615-L933 at varying temperatures The sample concentration was approximately 0.306 mM (measured through amino acid analysis). The NMR sample conditions were 20 mM sodium phosphate (pH 7.25), 150 mM NaCl, 2 mM ATP, 2 mM DTT, 5 mM MgCl2 with 2% glycerol and 2% D2O (v/v). Temperature conditions were varied in each spectra in intervals of 5 oC, starting at 15 oC in A and ending at 30 oC in D. NMR spectra for each temperature point were recorded for the same length of time (~1 hour) Arg NεHε correlations are indicated by the arrow above. Positive peaks are shown in black, and negative peaks are shown in blue

3.4.2 NBD1 SUR2A S615-L933 functionally binds ATP CD data indicated that the protein generated from S615-L933 is consistent with a folded protein with both α-helical and β-sheet structure. Our NMR data also indicate that S615-L933 is folded.

However, the functionality of such a construct needed to be determined, in order for it to be employed as a model for studying the nucleotide binding domains. The functional activity of

S615-L933 was evaluated through binding experiments with ATP. Binding activity was assessed through NMR spectroscopy.

Although NMR studies of SUR2A NBD1 were previously conducted using protein concentrations of 0.4-0.5 mM, such high concentrations in the absence of ATP and resulted in precipitation and deteriorating signal. Therefore, ATP binding studies were conducted at protein concentrations of approximately 100 µM. Figure 21A depicts the 15N-1H 2D TROSY-HSQC spectra of 15N labelled S615-L933 NBD1 in the presence of 2 mM ATP. Assuming a dissociation constant (Kd) of 150 µM between ATP and the protein, this would have led to a 92% saturation of NBD1 with ATP. Following spectra acquisition, the sample was dialyzed and extensively buffer exchanged to remove ATP from the sample. Another 15N-1H 2D TROSY-HSQC was conducted and illustrated in Figure 21B. As indicated by the spectra of apo-NBD1, a number of peaks are broadened beyond detection, possibly indicating dynamics on an intermediate timescale for many residues in apo-NBD1. However, the presence of many strong resonances with proton chemical shift values greater than 9 ppm indicate that apo-NBD1 is folded. ATP was reintroduced into the sample (at a concentration of 2 mM), and a number of resonances are shown to reappear. The resulting spectrum overlays well with the NMR spectrum of the originally purified S615-L933 protein in the presence of ATP. Slight differences in the number of cross-peaks between the spectrum before ATP was removed, and following its re-addition, may have resulted from slight differences in concentrations between the two samples (Table 7).

Regardless, the spectra indicates gross changes to the dynamics with and without ATP binding to the NBD subunit (Figure 22). Furthermore, the SUR2A NBD1 S615-L933 has been shown to functionally bind ATP.

A B

Figure 21: 2D 15N-1H TROSY-HSQC of NBD1-SUR2A S615-L933 with varying levels of ATP Sample concentration was approximately 0.100 mM. The NMR sample conditions were 20 mM sodium phosphate (pH 7.25), 150 mM NaCl, 2 mM DTT, with 2% glycerol and 2% D2O (v/v). In A, 2 mM ATP was present in the sample. The ATP was then dialyzed and buffer exchanged and the spectra was recorded (B). 2 mM was added back to the sample and the spectrum was recorded. NMR spectra were recorded for the same length of time (~8 hours at 30oC). Positive peaks are shown in black, and negative peaks are shown in blue.

Figure 22: Overlay of 2D 15N-1H TROSY-HSQC of NBD1-SUR2A S615-L933 with and without ATP Overlay of spectra from Figure 21B and 21C. Positive resonances from ATP+ and ATP- states are shown in black and red respectively. Corresponding negative cross-peaks are coloured in blue and green.

Table 7: Number of Amide Proton Crosspeaks in NBD1-S615-L933 (± ATP) (Based on Spectra in Figure 21) Spectra Number of Cross-peaks NBD1+ATP 328 NBD1-ATP 130 NBD1+ATP 316 (reintroduced) Expected Number of Cross-peaks:311

3.4.3 NMR Studies of T618-L933 Another potential construct to model nucleotide binding domain 1 of SUR2A involved a slightly truncated protein, T618-L933. This construct was designed to test secondary structures in the alignment. Our alignment indicates, S615 is located at the beginning of a β-strand which forms

the middle sheet C-terminus W905 of the β- W676 subdomain (Figure W723 23). Removal of

the amino acids

S615, W616 and

R617 allowed us

to test whether the

β-strand begins W876 W616 W755 Figure 23: Homology model of NBD1 of SUR2A ”” slight C-terminal The homology model was generated based on our structure based sequence alignment. The six tryptophan residues are identified above and the N-terminus of S615. Slightly is circle in red. Elutions

Ladder Post S1 P1 S2 P2 Flow Wash 1 2 3 4 5 6 7 kDA 98 64 50

Figure 23b: Purification of T618-L933 Gel samples based on different point in the purification process. Post is a sample of cell culture before and after induction. The soluble (S1) and pellet (P1) fractions are shown in following one cycle of sonication and following a second cycle (S2 and P2). The flowthrough (flow) and wash (wash) are samples of following the passing the cell lysate over Ni2+ resin. Samples from column-volume size elutions are shown in 1, 2, 3, 4, 5, 6 and 7.

less when compared to lower expression and purification yields of T618-L933 were obtained, compared to the levels achieved for S615-L933 as depicted in Figure 14. A similar method of purification used for S615-L933 was deemed suitable for T618-L933 and relative protein yields at each step are depicted in Figure 23b. However, the total protein recovery was noticeably

A B

Table 8: Number of Amide Proton Crosspeaks in NBD1-T618-L933 (Based on Figure 24) Spectra Number of Cross- peaks 20 oC 201 25 oC 264 30 oC 304 Expected Number of Cross-peaks:308

Figure 24: 2D 15N-1H TROSY-HSQC of NBD1-SUR2A T618-L933 at varying temperatures

Sample concentration was approximately 0.250 mM (measured through amino acid analysis). The NMR sample conditions were 20 mM sodium phosphate (pH 7.25), 150 mM NaCl, 2 mM ATP, 2

mM DTT, 5 mM MgCl2 with 2% glycerol and 2% D2O (v/v). Temperature conditions were varied o o o in each spectra in intervals of 5 C, starting at 20 C in A and ending at 30 C in C. NMR spectra for each temperature point were recorded for the same length of time (~1 hour) Positive peaks are shown in black, and negative peaks are shown in blue. 48

noticeably less when compared to S615-L933 (Table 5). A similar temperature step-wise experiment was conducted for T618-L933 using 2D 15N-1H TROSY-HSQC, and the results are depicted in Figure 24. There is a reduction in the quality of data obtained compared to S615-

L933, which may imply a destabilizing role to the residues removed. Furthermore, fewer crosspeaks are observed in T618-L933, including the aliased epsilon N-H groups of arginine and the tryptophan indoles at all temperatures (Table 8).

An overlay of the spectra between S615-L933 and T618-L933 (Figure 25) results in surprisingly large chemical shifts to specific resonances. Resonances not immediately identifiable in spectra of T618-L933 (compared to S615-L933), could be seen at high levels of noise. Despite the similarity between the spectra, a number of chemical shifts for specific peaks were observed, indicating that there were more global effects to the deletion of three residues.

Chemical shifts were observed in the region 8.3-8.8 ppm (disordered region) possibly indicating interactions between disordered loops and the N-terminus.

A convenient outcome in the deletion of three residues, was the elimination of one tryptophan (W616) which allowed for possible assignment of this residue in the HSQC spectra.

(Figure 25, inset). The shift in the two tryptophan indole resonances, indicated by ii in Figure

25, may ascribe these peaks to W876 and W755, which are expected to be close in space to the deleted amino acids according to our homology model. The tryptophan residues W905 and

W723, are located further way from the destabilized β-sheet and likely account for the remaining two residues.

A single intense peak is also absent in the spectra of T618-L933 compared to S615-L933

(Figure 25, labeled iii). This resonance peak likely corresponds to S615 or R617 due to the frequent presence serine and arginine residues in this region. However, regardless of a certain

level of ambiguity in the assignment of this peak, it still provides a clear indicator of changes that influence the N-terminus.

As conducted with S615-L933, ATP was dialyzed and diluted out of the sample of T618-

L933 to examine whether T618-L933 functions as a folded NBD with ATP binding activity.

However, in the absence of ATP the conformational structure was further compromised and high quality NMR data could not be obtained. A large number of cross-peaks are absent in the spectra of T618-L933 with a low dispersion in the proton dimension. These data indicate that the protein is most likely aggregated, at least at elevated concentrations without ATP, and underscores the importance of the deleted N-terminal residues. Removal of the first three residues of β-strand β1, which forms the middle strand in the three–stranded β-sheet subdomain, likely leads to destabilization of the β-sheet subdomain and aggregation. Furthermore, only a reduction in peak intensity was observed in the apo-state of T618-L933 with limited chemical shift changes. In comparison with the apo-state of S615-L933 (Figure 26), the truncated construct possesses only a subset of the number of peaks precluding a direct comparison between apo S615-L933 and apoT618-L933. The lower quality spectra of apo-T618-L933 and lower yields of protein inclined us to retain S615-L933 as a model for NBD1-SUR2A.

iii

i iv

Figure 25: Overlay of 2D 15N-1H TROSY-HSQC of NBD1-SUR2A S615-L933 with T618-L933 Overlay of spectra from both constructs. Positive resonances from S615-L933 and T618-L933 are shown in black and red respectively. Corresponding negative cross-peaks are coloured in blue and green. The inset shows the tryptophan indoles at a higher contour level. i) shows the W616 peak ii) likely points to W876 and W755 iii) shows the likely N-terminus, S615 and iv) shows the likely C- terminus, L933 51

Figure 26: Overlays of 2D 15N-1H TROSY-HSQC of NBD1-SUR2A T618-L933 The overlay of T618-L933 with (black) and without ATP (red) is shown in A, with negative cross peaks shown in blue and green respectively. B shows an overlay between the apo-states of S615- L933 (positive in black, negative in blue) and T618-L933 (positive peaks in red, negative peaks in green). 52

3.4.4 Phosphorylation of S615-L933 NBD1 SUR2A

A key step in regulation of KATP channels involves phosphorylation of the nucleotide binding domains by protein kinase A. This response leads to increased activity of the channels, although the precise molecular mechanism by which this occurs is unknown. There is one such phosphorylation site in NBD1, T632, located in the β-sheet subdomain insert (Fig 10). The effect of phosphorylation on SUR2A NBD1 S615-L933 was examined.

Similar to previous experiments, a series of 2D 15N-1H TROSY-HSQC spectra were recorded at different temperatures, and the results are depicted in Figure 27 A, C and E. Overlays of the corresponding unphosphorylated spectra are depicted in Figure 27 B, D and F to illustrate the change in temperature dependence. Phosphorylation appears to have altered the structure in a two significant ways. Firstly, high quality spectra (large number of peaks, with broad dispersion) are obtained at lower temperatures (20oC, 25oC, Table 9). The non-phosphorylated state has a number of resonances broadened beyond detection. Secondly, phosphorylation leads to a difference in behaviour of a number of peaks, including the arginine NεHε resonances. In non- phosphorylated S615-L933, these resonances broaden out completely as temperature is increased. The reverse occurs for the phosphorylated spectra. Furthermore, the peak corresponding to S615/R617 is also missing, and the resonance corresponding to W616 only reappears at 30oC (Fig 27). Both of these observations suggest a change in dynamics between the phosphorylated and non-phosphorylated states. Spectra E and F were collected on a different spectrometer with a smaller sweepwidth, which leads to the aliasing of a number of peaks which appear positive in previous spectra.

A B

C D

E F

Figure 27: 2D 15N-1H TROSY-HSQC of Phosphorylated NBD1-SUR2A S615-L933 Sample concentration was approximately 0.250 mM, with the same sample conditions as the non- phosphorylated protein. Temperature conditions were varied at 20oC, 25oC and 30oC in A, C and E. (black and blue for positive and negative peaks) Overlays are shown in B, D and F with the corresponding non-phosphorylated product (+black and -blue WT, +red and -green phospho) 54

Removal of ATP from the phosphorylated state of S615-L933 led to an immediate reduction in spectra quality, with a number of different cross peaks being broadened beyond detection (Figure

28, Table 9). It is possible that the absence of ATP led to aggregation of the phosphorylated protein. This suggests that there are significant conformational changes induced by phosphorylation compared with the non-phosphorylated state.

Table 9: Number of Amide Proton Crosspeaks in Phosphorylated NBD1-S615-L933 (Based on Figure 27 and Figure 28) Spectra Number of Cross-peaks 20 oC 248 25 oC 302 30 oC 354 -ATP, 30oC 118 Expected Number of Cross-peaks:311

Figure 28: 2D 15N-1H TROSY-HSQC of Phosphorylated NBD1-SUR2A S615-L933 without ATP at 30oC Sample concentration was approximately 0.1 mM. ATP was removed from the sample through dialysis and buffer exchanging. Sample conditions are otherwise identical to those obtained with spectra of the phosphorylated protein. Positive cross peaks are shown in black and negative.

3.4.5 Role of R934-K972 in NBD1 SUR2A Extension of the original construct from S615-L933 to include a region further C-terminal

(K972) was tested to determine the effect on stability of SUR2A NBD1, but also to investigate the role of this region in regulation. This region is rich in acidic residues and has been shown to be critical for proper gating of KATP channels.[73] Disrupting the ED loop has shown to hamper cooperative interactions of the NBD suggest a possible allosteric mechanism of regulation. Once again, lower expression and purification yields were obtained compared to S615-L933 (Figure

14). Purification entailed the use of nickel affinity chromatography as depicted in Figure 29.

Ladder Pre Post S1 P1 S2 P2 Flow Wash 1 2 3 4 5 Ulp kDA

98 64 50 36

Figure 29: Purification of NBD1 S615-K972 Gel samples based on different stages in the purification process. Pre and Post are samples of cell culture before and after induction. The soluble (S1) and pellet (P1) fractions are shown in following one cycle of sonication and following a second cycle (S2 and P2). The flowthrough (flow) and wash (wash) are samples of following the passing the cell lysate over Ni2+ resin. Samples from column- volume size elutions are shown in 1, 2, 3, 4 and 5. The product post-Ulp addition is in the final lane.

Ladder 1 2 3 4 5 6 7 8 9 kDA

98 64 50 36

Figure 30: Fractions following gel-filtration of S615-K972 Samples from each of the fractions from the gel filtration column are shown above in a 1/1000 dilution. The final pure product is shown in B.

Elutions 2 and 3 were pooled and the imidazole concentration was diluted to allow for cleavage of the SUMO tag by Ulp1. Subsequent size-exclusion chromatography allowed for pure samples to be obtained. (Figure 30) However, protein-containing fractions were passed over nickel resin to ensure contaminating Ulp1 protease and SUMO tag were removed. Similar to previous experiments, NMR spectra of S615-K972 was examined across a range of temperatures (Figure

31). The C-terminal extension to K972 appeared to, at least qualitatively, increase the stability of the NBD1 unit, and the experimental temperature could be increased to 35oC without significant precipitation. Furthermore, the aliased epsilon N-H atoms of the arginine residues are visible in spectra, at 30oC, compared to their disappearance in corresponding experiments for S615-L933, although some broadening was still evident. Their gradual disappearance from the spectra with increasing temperature may indicate molecular motions approaching the intermediate timescale.

The transition from the slow to the intermediate timescale for the epsilon NH groups of the arginine residues occurs at a lower temperature for S615-L933 than for S615-K972. This may suggest some increased interactions of these basic residues, possibly with the highly acidic rich region of the R934-K972, and may contribute to the observed increases in stability. A number of peaks reappear at higher temperatures, including all of the tryptophan indoles at 35oC (Table 10,

Figure 32). This marked change in spectra between temperatures may indicate increases on the timescale of motion, with shifts from the intermediate to fast timescale for a number of aromatic residues.

A B

C D

E F

Figure 31: 2D 15N-1H TROSY-HSQC of NBD1-SUR2A S615-K972 Sample concentration was approximately 0.200 mM, with the same sample conditions as S615- L933. Temperature conditions were varied at 20oC, 25oC and 30oC in A, C and E. (black and blue for positive and negative peaks).Overlays are shown in B, D and F with S615-L933 at the corresponding temperature (+black and -blue S615-L933, +red and -green S615-K972)

Figure 32: 2D 15N-1H TROSY-HSQC of NBD1-SUR2A S615-K972 at 35oC Sample concentration was approximately 0.200 mM, with the same sample conditions as the previous experiments of S615-K972 respectively. Resonances are shown in black and blue for positive and negative peaks.

Table 10: Number of Amide Proton Crosspeaks in NBD1-S615-K972 (Based on Figure 31 and Figure 32) Spectra Number of Cross-peaks 20 oC 147 25 oC 208 30 oC 248 35 oC 280 Expected Number of Cross-peaks: 347

A B

Figure 33: 2D 15N-1H TROSY-HSQC of Phosphorylated NBD1-SUR2A S615-K972 at 30oC Sample concentration was approximately 0.200 mM, with the same sample conditions as the non-phosphorylated protein (black and blue for positive and negative peaks). The spectra is overlayed in B on the non-phosphorylated spectra in (+black and -blue nonphosphorylated, +red and -green phosphorylated) Phosphorylation of S615-K972 led to significantly poorer quality of NMR spectra. The loss in the number of cross-peaks identified may be attributed to a change in the timescale of motion.

The cross peaks corresponding to the N-terminal residues have also been broadened out and are less intense and the cross peaks corresponding to the arginine residues are also absent in the spectra. It is also important to mention that the reduction in quality of the spectra of S615-K972 in the phosphorylated and non-phosphorylated state may be due to some degree of aggregation.

The very strong T1 noise in this spectrum at 6.8 ppm and 8.1 ppm and 8.5 ppm are derived from

ATP resonances.

3.4.6 Secondary Structural Analysis of NBD1 SUR2A NMR provides residue-specific data on the conformation and dynamics of proteins.

In contrast, other biophysical techniques such as circular dichroism provide information on the overall secondary structure, as well as information on the tertiary structure of a protein.

Furthermore, protein concentrations and data acquisition times required for ample signal are significantly lower. This allows us to perform experiments, such as temperature melts, which are less manageable in NMR due to the high concentration of protein required coupled with the unfolding.

CD was performed on purified S615-L933 as shown in Figure 33. The minima at 206 nm and maxima around 190 nm are consistent with mixed alpha-beta structure Introduction of a denaturating agent, such as guanidium hydrochloride interferes with proper folding and disrupts secondary and tertiary structure. (Figure 34). At concentrations greater than 3M guanidium hydrochloride, SUR2A NBD1 is completely unfolded.

Figure 34: Circular Dichroism Spectrum of S615-L933 The CD spectra for 10 µM of NBD1 S615-L933 is shown above. Spectra were obtained with a bandwidth of 0.2 nm using a 0.1 cm path length quartz cell. Samples contained 5 μM NBD1 in 20 mM phosphate, pH 7.25. Spectra were averaged from 3 scans.

2000

0 3M GuHCl

-2000

-1

2M GuHCl -4000

residue

-1

dmol

2 -6000 1M GuHCl

deg cmdeg -8000

Mean Molar Ellipticity per Residue, MolarMean Ellipticity

-10000 0M GuHCl

-12000 220 225 230 235 240 Wavelength, nm Figure 35: Guanidine-HCl melt of NBD1 SUR2A S615-L933 The effect of increasing guanidine-hydrochloride is shown on 10 µM of NBD1 S615-L933 in 20 mM NaPhos, pH 7.25.

Data below 220 nm could not be recorded due to increasing absorbance of guanidium hydrochloride. These experiments, in the absence of ATP, conclusively indicate that NBD1

S615-L933 maintains a well-defined secondary structure. Furthermore, this structure is fairly stable, and resistant to increasing levels of denaturtant. Unfortunately, CD cannot be effectively performed in the presence of ATP due to the absorbance of the nucleotide. Phosphorylation of

NBD1 S615-L933, as shown in Figure 36, depicts an 8 nm red-shift in the minima observed.

However, the magnitude of signal at the minima remained consistent with the non- phosphorylated state. CD spectra of apo-S615-K972 are depicted in Figure 36B. There are a significant number of additional spectral features at longer wavelengths, not seen in spectra of other constructs. These are likely the result of the introduction of the acidic C-terminal tail which alters the tertiary structure. The secondary structure, based on the lower wavelength minima, remains relatively constant, both in terms of magnitude and wavelength of minima. The tertiary 62

structure which gives rise to the positive CD-bands from 258 nm to 287 nm is quite resistant to guanidium hydrochloride, and was only subdued at around 6 M of denaturant (data not shown).

Shorter wavelengths could not be effectively monitored due to interference from guanidium hydrochloride.

Figure 36: CD spectra of Phosphorylated S615-L933 (A) and S615-K972 (B) Spectra were obtained with a bandwidth of 0.2 nm using a 0.1 cm path length quartz cell. Samples contained 5 μM NBD1 in 20 mM phosphate, pH 7.25. Spectra were averaged from 3 scans. The CD profile of the wild type (S615-L933) is shown in black for comparison in each spectra.

3.4.7 Fluorescence Spectroscopy of NBD1 SUR2A Fluorescence spectroscopy probes macromolecular changes that occur near fluorophores. There are six tryptophan residues in NBD1 and hence the fluorescence spectra are a reflection of the environment of all of these tryptophan residues. The location of these tryptophan residues from our homology models is shown in Figure 23. In Figure 37, fluorescence was employed to probe conformation as a function of temperature. The six tryptophan residues present in S615-L933 and S615-K972 made the interpretation of fluorescence more challenging. In Figure 38, the change in fluorescence following incubation at successively higher temperatures is examined to

Figure 37: Fluorescence melt curve of S615-L933 The fluorescence of 2 µM of NBD1 S615-L933 was measured as the temperature was gradually increased. Excitation light was given at 295 nm (with a slit width of 1 nm). Emission spectra were collected over 300 nm to 450nm at 1 nm increments with a slit width of 2.5 nm. The average of three scans is shown above.

assess stability. The wavelength with the largest difference between the folded and unfolded states was selected for the analysis to maximize signal-to-noise. Temperature melts of the proteins were reversible for short temperature spans, but irreversible for wider ranges. Melting for all protein constructs was shown to be cooperative with two state unfolding based on tryptophan fluorescence. The temperature melts of the proteins were reversible for short temperature spans, but irreversible for wider ranges. The temperature melts of the proteins indicated that the structure was stabilized following the addition of ATP. Phosphorylation served to further stabilize the structure. The ED loop was also shown to increase the stability of the structure. Overall, the data indicates a relatively low melting temperature for S615-L933 (37oC),

Table 10. However, at 30oC in the NMR the protein still appears folded (Figure 20, 21). The fluorescence spectra, which involves the fluorescence of the six tryptophan residues, possibly reflects a conformational change with temperature rather than a pure two-state unfolding melt.

100 ATP Phosphorylated - - + - - + 80 + +

% Protein Folded Protein %

0 10 20 30 40 50 60 70

o Temperature ( C) Figure 38: Fluorescence melt curve of S615-L933 under varying conditions The fluorescence of 2 µM of protein was measured as the temperature was gradually increased. Excitation light was given at 295 nm (with a slit width of 1 nm). Emission spectra were collected over 300 nm to 450nm at 1 nm increments with a slit width of 2.5 nm. Changes in the signal at 349 nm (the wavelength with maximum signal-to-noise) were related to changes in protein folding and plotted above. 65

Table 11: Melting temperatures determined by fluorescence ATP Phosphorylation Melt Temperature (oC) S615-L933 – – 37 – + 42 + – 43 + + 45 S615-K972 – – 43 – + 46 + – 48 + + 51

The solvent accessibility of the tryptophan residues was assessed through iodine quenching experiments. Increasing iodine concentrations was shown to gradually reduce the fluorescence intensity observed as shown in Figure 39. In all ATP-minus states, the solvent accessibility of the tryptophan residues was the same (fully solvent exposed, Figure 40) However, the addition of

ATP stabilized the structure, as shown by temperature-dependent melting, and decreased solvent exposure. Phosphorylation led to increases in solvent exposure in the presence of ATP as shown in Table 12.

0 mM KI 200

150

100

Fluoresence Intensity (x1000) Fluoresence Intensity 500 mM KI

0 300 320 340 360 380 400 420 440 Wavelength, nm Figure 39: Quenching of fluorescence of S615-L933 by potassium iodide Fluorescence spectra of S615-L933 were recorded with increasing amounts of potassium iodide. As KI concentration increased, fluorescence intensity decreased. The individual spectra represent fluorescence measurements at 0, 10, 20, 50, 75, 100, 150, 250 and 500 mM KI. Each curve is the average of three individual scans.

ATP Phosphorylation + + 8 -- + -- -- + --

-F)

/(F

o F 4

0 0.000 0.005 0.010 0.015 0.020 1/[KI], mM-1 Figure 40: Modified Stern-Volmer plots of fluorescence quenching of S615-L933 The fluorescence intensities measured in the presence of varying amounts of potassium iodide were manipulated according to modified Stern-Volmer relationships. The reciprocal of the y intercept represents the fraction of fluorophores which are available for quenching. The slope represents the quenching constant. The solvent accessibility to tryptophan residues in S615-L933 and S615-K972 under a variety of conditions is depicted in Table 11.

Table 12: Tryptophan solvent exposure determined by fluorescence quenching

ATP Phosphorylation Solvent Exposure S615-L933 – – 100% – + 100% + – 59% + + 70% S615-K972 – – 100% – + 96% + – 74% + + 76%

3.5 Biophysical studies of Val734Ile Mutant in SUR2A NBD1

The construct for the Val734Ile mutant was generated through QuikChange (Stratagene) mutagenesis of the S615-L933 construct. The mutant was expressed and purified in the same manner as the wildtype, S615-L933. Protein yields for the mutant were on the same order as those obtained from the original construct (~15mg/L of M9 media). As before, NMR spectra of the mutant were obtained over a range of temperatures. Higher concentrations (~0.4 mM) of the mutant, however, appeared aggregated in the NMR spectra (not shown). For this reason, spectra at lower concentrations (~0.2 mM) were recorded. The number of cross-peaks in each spectrum is recorded in Table 13. CD spectra and fluorescence melt curves were also performed on the mutant. CD spectra indicated highly similar secondary structures between the mutant and wildtype protein (Figure 42). Furthermore, similar melt temperatures (± ATP) were observed for the mutant compared to the wild-type (Figure 43). However, there were large differences between the surface accessibility of the tryptophan residues in the presence of ATP (Figure 44, Table 14).

The quenching constant and surface accessibility are well-matched to the phosphorylated state of

S615-L933. There are also a number of similarities in the NMR spectra between the phosphorylated state of S615-L933 and the mutant Val734Ile in S615-L933 as illustrated in

Figure 41.

Table 13: Number of Amide Proton Crosspeaks in Mutant V734I of NBD1-S615-L933 (Based on Figure 21, 44) Spectra Number of Cross-peaks WT 304 Mutant 286 Expected Number of Cross-peaks: 311 .

A B

Figure 41: 2D 15N-1H TROSY-HSQC of Mutant Val734Ile of NBD1-SUR2A S615-L933 The spectrum of the mutant at 30oC, (0.200 mM) is shown in A, with a comparable spectrum of the wildtype (B). The NMR sample conditions for both spectra were 20 mM sodium phosphate (pH 7.25), 150 mM NaCl, 2 mM DTT, 2 mM ATP with 2% glycerol and 2% D2O (v/v). Positive peaks are shown in black, and negative peaks are shown in blue. Changes between the spectra are circled in cyan circles in C (+black and -blue WT, +red and -green mutant).

Figure 42: CD spectra of Mutant Ile734Val in S615-L933 Spectra were obtained with a bandwidth of 0.2 nm using a 0.1 cm path length quartz cell. Samples contained 5 μM NBD1 in 20 mM phosphate, pH 7.25. Spectra were averaged from 3 scans.

Figure 43: Fluorescence melt curves of Mutant Ile734Val in S615-L933 in the presence/absence of ATP The fluorescence of 2 µM of protein was measured as the temperature was gradually increased. Excitation light was given at 295 nm (with a slit width of 1 nm). Emission spectra were collected over 300 nm to 450nm at 1 nm increments with a slit width of 2.5 nm. Changes in the signal at 349 nm (the wavelength with maximum signal-to-noise) were related to changes in protein folding and plotted above. Red and blue spectra indicate the –ATP and +ATP WT states respectively

10 ATP Phosphorylation + + -- + 8 -- -- + ------(Mutant) + -- (Mutant) 6

-F) o

/(F

o F 4

0 0.000 0.005 0.010 0.015 0.020

1/[KI], mM-1

Figure 44: Modified Stern-Volmer plots of fluorescence quenching with Val734Il3 of S615-L933 Modified Stern-Volmer plots of iodine quenching to Val734Ile of S615-L933 were overlaid onto the quenching plots of the original NBD1 construct, S615-L933. The mutant in the absence of ATP is shown in pink, and in the presence 2 mM ATP is displayed in cyan.

Table 14: Tryptophan solvent exposure determined by fluorescence with Val734Ile ATP Phosphorylation Solvent Exposure S615-L933 – – 100% – + 100% + – 59% + + 70% S615-K972 – – 100% – + 96% + – 74% + + 76% S615-L933 – – 100% Val734Ile – + 74%

Chapter 4 Discussions and Conclusions

4.1 Identification of SUR2A NBD1 Domain Boundaries

The production of folded and functional samples of the nucleotide binding domains

(NBDs) has been a recurring challenge in the study of eukaryotic ABC transporters. Several different strategies have been attempted to isolate soluble NBDs with varying degrees of success.

Overexpression techniques in E. coli generally result in the formation of inclusion bodies.[74]

Approaches for in vitro refolding of multi-domain proteins are generally limited in success and result in multiple populations of variably folded proteins. Structural studies on NBD1 of CFTR, another ABCC subfamily member like SUR2A, were hindered for many years. [14,68] This was predominantly due to lack of knowledge on the ideal N- and C- terminal NBD1 domain boundaries. In our studies, we considered a series of plausible domain boundaries for NBD1 of

SUR2A based on our structure-based sequence alignment. A number of NBD1 proteins with these boundaries were recombinantly expressed in E. coli, purified, and screened for solubility.

Using out structure-based sequence alignment, solubility screens and biophysical data, we identified the optimal domain boundaries for SUR2A NBD1 as S615-L933. The choice of boundary domains is selected so as to include all structurally identifiable regions. Premature termination of the sequence, especially in a structured region, can result in improperly folded secondary structures and promote aggregation. Furthermore, unstructured regions extending beyond the N- and C-terminal domains can result in destabilization of the overall molecule and lower the soluble expression. For example, removal of three N-terminal residues in the construct

S615-L933 led to reduced stability of the protein, T618-L933. The residues, S615-R617 are the first three residues in the β1, β-strand, which is the middle β-strand of the β-sheet subdomain 72

(Figure 23). Removal of these residues likely resulted in disruption of the β-strand and destabilization of the entire β-sheet subdomain. The construct of S615-L933 extends beyond conventional C-terminal boundaries for NBD1 (D914). Expression of NBD1 with the boundaries

S615-L933 resulted in greater yields of purified soluble protein than S615-D914. Analogous stabilization by C-terminal residues beyond the final canonical secondary structure component of most NBDs, is also seen to occur in CFTR. The C-terminal region of CFTR NBD1 is known as the RE region and possesses unique regulatory roles. Parallel functions have yet to be identified for the C-terminal residues of SUR2A. Extending the domains to K972 resulted in slightly lower soluble expression levels. However, the lengthening of the boundary also introduces a number of acidic residues into NBD1 which may lead to a reduced interaction with SDS and lower apparent expression when tested through SDS-PAGE. The region encompassing these acidic residues, is known as the ED loop and may have related regulatory properties.

The extension of S615-L933 in the N-terminal direction allowed for high levels of overexpression, but the solubility of these constructs was significantly reduced. The addition of N- terminal residues may increase probability of NBD1 dimerization and aggregation. This N- terminal dependent aggregation has been observed in similar regions of the P-glycoprotein

NBDs.[81]

An interesting feature which is apparent from the structure-based sequence alignment is the presence of a 35 amino acid insertion between the first two β-strands in NBD1. Notably, a naturally occurring splice isoform of SUR2A is missing much of this insert. This insertion is also seen in CFTR, where it is known as the regulatory insert (RI), and exerts regulatory control over the NBDs. There are a number of similarities between the RI of CFTR NBD1 and the insert identified in SUR2A NBD1.[14, 68] The RI in CFTR, NBD1 consists of two α-helices separated by a disordered linker. Similarly, part of the insertion (Q635-E664) in SUR2A is expected to be

disordered, through the PONDR algorithm, VLXT. The disordered residues in the β-sheet subdomain insert of SUR2A NBD1 overlap with the disordered regions of CFTR RI (Figure 13).

Moreover, both CFTR and SUR2A possess a PKA phosphorylation sites in this region.

Phosphorylation of the RI of CFTR leads to stimulatory activation for CFTR. Phosphorylation of

T632 in the β-sheet subdomain of SUR2A results in KATP channel activation. This reinforces the likelihood of a similar regulatory mechanism for the insert in SUR2A

4.2 Characterization of ATP binding in SUR2A NBD1

High yields of soluble SUR2A NBD1 S615-L933 allowed for characterization of the domain through a series of biophysical methods. Resonances in 2D 1H-15N NMR spectra of S615-L933 are well dispersed in the proton dimension indicating a folded structure. However, there is great variability in peak intensity. The varying peak intensities as well as the temperature dependent appearance of some resonances and disappearances of others, indicate a highly dynamic structure. The clustering of intense peaks around 8.2 ppm in the proton dimension may be reflective of the disordered structures present or residues which sample a series of disordered conformations. This disordered structure may be partially attributed to the predicted β-sheet subdomain.

An important step in the characterization of the nucleotide binding domain is the binding of ATP. The addition of ATP to the NBD1 has been previously modelled as a ‘switch’ between open and closed NBD dimeric states. [75] ATP binding and subsequent NBD dimerization has been suspected of substantially altering the NBD structure. Quenching data of SUR2A NBD1 elucidates a significant conformation change by effectively decreasing accessibility to tryptophan residues by approximately 30% (Table 12). This conformational change may expose specific residues to allow NBD1-NBD2 interaction and subsequent dimerization. Specifically, the D-loop

has been shown to be involved at the dimerization interface. Based on our homology models, it is located far from tryptophan residues in the apo-state. The presence of ATP may lead to stabilization of the D-loop, near a tryptophan residue, and therefore shields the tryptophans from quenching. Such an interaction likely occurs in proximity to the β-sheet subdomain, This is because, intrinsic fluorescence results from six tryptophan residues in SUR2A NBD1 and the significant decrease in fluorescence likely corresponds shielding of multiple tryptophan residues.

As such, the β-sheet subdomain contains three tryptophan residues in close proximity.

However, this conformational change may be a highly dynamic phenomenon.

Large changes in atomic B-factors for crystallized structures of Methanococcus jannaschii,

MJ1267 (ABCB related protein), in the presence and absence of the nucleotide suggest significant changes in mobility.[76] The NMR spectrum of SUR2A NBD1 S615-L933 in the presence of 2 mM ATP displays several differences compared to the apo-state (Figure 21). There appears to be a number of differential dynamic effects on specific residues. For example, the epsilon NH correlations from arginine residues which appear in the spectra at 20oC, are

Figure 45: Dynamics of motion change upon ATP binding A number of "indicator" peaks have been used to assess changes in their relative time scale of motion. Changes were assessed based on their broadening or reappearance as temperature increased in the presence and absence of ATP. Arg refers to the aliased arginine residues, Trp616 is the N-terminal tryptophan which is likely in fast exchange. Trps refers to the other 5 tryptophan resonances. Overall, the removal of ATP broadens a number of resonances from the fast timescale to intermediate exchange, where they are not detected.

broadened beyond detection at 30oC. This may represent a motion transiting from the slow-time scale to the fast timescale regime with increasing temperature. A number of "indicator" peaks in the spectra are employed to exemplify differential dynamics between ATP-bound and unbound states (Figure 45). There appears to be a general shift towards increased dynamics upon ATP binding, although it is not true for all cross-peaks. Nucleotide binding in MJ1267 has also been shown to cause differential changes to exchange broadening. [76] Motions in MJ1267 on the microsecond-millisecond time scale were primarily altered following nucleotide binding in two significant ways. Motions of residues located at or close to the ATP binding site, such as E179 and I181 in the D-loop, were shown to significantly decrease. However, many motifs, including parts of the Q-loop and H-loop retained their flexibility. Further, some residues located 30 angstroms away from the binding site, increased mobility upon ATP binding.

A number of chemical shift changes have been observed upon nucleotide binding. Given the large scope of changes observed with ATP binding, these changes are likely not only localized in the vicinity of nucleotide binding, but also far away. These chemical shift changes likely indicate a conformational change in NBD1 upon ATP binding. To observe changes in chemical shifts, ligand (ATP) binding should alter the sub-state distribution of a particular residue or the number of sub-states which it may exist in.[77] Therefore, ATP is likely involved in an allosteric and dynamic mechanism for the functional activity of the NBDs. Allosteric mechanisms are intrinsically dynamic mechanisms, as they involve a particular pathway for specific motions. Studies from MJ1267 suggest that nucleotide binding is coupled to significant dynamic mechanisms. Dynamic mechanisms are more dependent on the folds of a protein rather than specific amino acids and suggest a supplementary role for the specific domain structures.

Wang et al also suggest that such a dynamic coupling between active and inactive states would be an efficient means of applying selective pressure for specific conformations during protein

evolution.[75] However, the knowledge into the dynamic mechanism of allosteric control for

SUR2A NBD1 by ATP is still quite limited and additional information such as determination of

R1, R2 and Rex are necessary for a more integrative view.

4.3 Effects of Phosphorylation on NBD1 of SUR2A

Phosphorylation of the NBDs has been shown to lead to a stimulatory effect on ATP-binding and hydrolysis. Our studies indicate that phosphorylation of SUR2A NBD1 changes the dynamics of motion. At lower temperatures, phosphorylated S615-L933 possessed a larger number of cross peaks compared to the non-phosphorylated counterparts. Peaks associated with N-terminal residues, such as W616 and S615/R617, are not apparent in the spectra at lower temperatures indicating an intermediate timescale of motion. However, all of the remaining tryptophan indoles which could not be resolved in the non-phosphorylated state at 20oC were clearly distinguished in the phosphorylated spectra. Phosphorylation of the RI in CFTR NBD1 results in large scale shifts in conformational flexibility.[70] (Recall that CFTR RI is analogous to the SUR2A NBD1

β-sheet subdomain insert). In particular, phosphorylation of CFTR results in displacement of the regulatory insert (RI) from sites of interaction on NBD1. Many of these interacting residues have been shown to be along the dimerization interface of NBD1 and NBD2 which suggests a possible allosteric mechanism for control.[3,68,69] Although the phosphorylation of NBD1 (at T634) is close to the N-terminus, phosphorylation likely leads increased conformational dynamics in other regions in the protein. As seen for CFTR, the β-sheet subdomain insert may mediate interactions with the rest of SUR2A NBD1 and cause broadening. Phosphorylation of the β-sheet subdomain insert may release the insert from the NBD1, resulting in less broadening.[68] Alternatively, the additional phosphate group at T634 in SUR2A NBD1 induces an electrostatic interaction with a nearby residue, such as an arginine. This may restrict mobility of the nearby N-terminus which

results in slower-motions on the NMR timescale again causing less broadening. Shifts to secondary structure are also depicted through CD analysis which depicts a 6 nm red-shift upon phosphorylation of the local minima corresponding to α/β structure.

4.4 Role of the ED Domain in NBD1

In the region C-terminal to our NBD1 domain boundaries, there is a long stretch of aspartic and gluatamic acid residues, termed the ED loop. Extending the construct from L933 to K972 allows us to incorporate the ED loop. Other data has shown an involvement in allosteric regulation of

KATP channels resulting from the ED domain. [71] We decided to investigate the effect of this region on the conformation of NBD1. NMR experiments indicated the presence of the ED loop broadened out a number of peaks compared to S615-L933. However, the reappearance of a number of these peaks at higher temperatures suggests that the interactions of the ED loop in

NBD1 results in a stabilization effect. (We see less precipitation of S615-K972 than with other constructs.) This may involve a stabilization effect of interactions between arginine residues, which are located throughout the protein, and the ED domain. A number of the arginine residues cluster with each other on a single face of the protein, which suggest possible sites of ED loop binding.

A number of ABC transporters possess analogous ED domains. In CFTR, there are a number of PKA-mediated phosphorylation sites in a region C-terminal to the NBDs. This region is known as the regulatory R-region and has been shown to have allosteric regulatory functions.

[68] Phosphorylation of the R region disrupts interactions with NBD1 and leads to greater ATP binding and hydrolysis and greater CFTR gating. In contrast to CFTR, the ED loop of SUR2A does not possess any phosphorylation sites. However, the acidity of the ED domain may possess a similar function. Electrophysiological studies by Karger et al., have shown that neutralization

of the acidic residues results in KATP phenotypes insensitive to stimulatory Mg-ADP or inhibitory sulfonylurea derivatives. [71,84] This is likely due to compromised interactions of NBD1 with

NBD2. Interactions studies of NBD1+ED loop (ie. S615- K972) with NBD2 would possibly allow for further characterization of this possible regulatory role.

4.5 Characterization of Mutant Val734Ile

The amino acid substitution of Ile734Val has been identified to increase the risk of mydocardial infractions. Biochemically, this is a conservative mutation which results in the addition of a single methylene group onto the valine side chain. Furthermore, this mutation occurs in a loop and would expected to be physiologically silent. However, this is clearly not the case, as seen from disease associated phenotypes.[59] CD spectra of the –ATP states are nearly superimposable and suggest limited changes in the secondary structure. Quenching experiments, to monitor intrinsic tryptophan fluorescence also suggest similar structures exist in the apo-states in which all of the tryptophan residues are quencher accessible. Upon ATP binding, there is a significant reduction in quencher exposure to the tryptophans in the NBDs which further suggests a conformational change upon nucleotide binding. Furthermore, the surface accessibility and quenching constants between the mutant and wild type are quite distinct, but are nearly identical between the mutant and phosphorylated wild type protein. However, the negatively charged phosphate group may result in slightly skewed iodine accessibility. Whether these similarities are the result of comparable structures or other quenching processes converging on similar values requires further investigation. Quenching experiments with acrylamide or oxygen may provide further insight into surface accessibility. NMR data is not fully conclusive either. Phosphorylated S615-L933 shows similarities with the mutant spectra although data collection with different sweepwidths precluded an overlay of spectra. However, similarities

between the active phosphorylated state and the mutant may suggest a possible biochemical consequence of the Ile734Val mutation.

4.6 Conclusions and Future Directions

Challenges in isolated eukaryotic NBD expression have presented difficulty in molecular-level understandings of the SUR proteins. Based on structure-based sequence alignments, we have optimized boundary domains of SUR2A NBD1 which resulted in soluble and folded constructs.

Furthermore, our construct of NBD1 (S615-L933) is functional and shown to reversibly bind

ATP. Biophysical studies have illustrated possible conformational changes through interactions with regulatory domains.

Future investigations into the dynamics of ATP-binding with the NBDs will help in the biochemical understanding of nucleotide interactions. Furthermore, residue assignment of the

HSQC spectra will significantly add to the interpretation of the spectra presented. Chemical shifts in specific peaks would allow for localization of conformational and dynamic effects to individual residues.

The role of the potential regulatory domains in the NBDs needs to be further characterized. Their interactions with specific binding partners, such as the NBD2, the L0 linker or the coupling helices need to be studied as these likely have important biological ramifications.

Such insight into the structural and dynamic mechanisms of the mutations will likely allow for efficient screening of potential drug targets.

5 Appendix

Supplementary Figure 1: Structure-based sequence alignment of SUR2A NBD1 with ABCC transporters (Continued) 82

Supplementary Figure 1: Structure-based sequence alignment of SUR2A NBD1 with ABCC transporters (Continued) 83

Supplementary Figure 1: Structure-based sequence alignment of SUR2A NBD1 with ABCC transporters (Continued) 84

Supplementary Figure 1: Structure-based sequence alignment of SUR2A NBD1 with ABCC transporters 85

Supplementary figure 1 shows the full sequence alignment for SUR2A NBD1 with various

NBDs of the ABCC family (generated through Clustal W). The secondary structure of CFTR

NBD1 (PBD: 1R0X) is shown in the grey schematic above the alignment. Residues belonging to

α-helices, β sheets and 310 helices are shown as grey cylinders, arrows and open circles respectively. Known secondary structure elements are coloured in the sequences alignment as blue, violet and green for α helices, β sheets and 310 helices respectively. Residues which belong to conserved domains, such as the Walker A and Walker B motifs, the ABC signature sequence and Q,D,H loops are labeled above. The phosphorylation site is outlined in a red box, and an asterisk is shown by the aromatic residue involved in interactions with ATP.

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