The Characterization of the Human Monoacylglycerol using Mutagenesis, Kinetics and NMR spectroscopy

Thesis Presented By Girija Rajarshi To The Bouve’ Graduate School of Health Sciences in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy in Pharmaceutical Sciences with Specialization in Protein Biochemistry and Drug Discovery

CENTER FOR DRUG DISCOVERY NORTHEASTERN UNIVERSITY BOSTON, MASSACHUSETTS July, 2017

Northeastern University Table of Contents

Page Acknowledgements………………………………………………………………………….1 List of Tables………………………………………………………………………………...2 List of Figures………………………………………………………………………………..3 List of Abbreviations…………………………………………………………………………8

Chapter 1. The ………………………………………………….10 1.1 Endocannabinoid System 1.1.1 Receptors 1.1.2 Endocannabinoid Endogenous Ligands 1.1.3 Endocannabinoid 1.2 Monoacylglycerol Lipase 1.3 MGL as a Therapeutic Target

Chapter 2. Direct NMR Detection of the Conformational Transition Between Open and Closed Forms of Monoacylglycerol Lipase………………………………………………..22

2.1 Introduction 2.2 Experimental Procedures 2.2.1 hMGL Mutagenesis, Expression and Purification 2.2.2 Sample Preparation and NMR Spectroscopy 2.3 Results 2.3.1 hMGL Variants Facilitating NMR Experiments 2.3.2 hMGL NMR Downfield Spectral Features 2.3.3 hMGL NMR Amino-acid Resonance Assignments 2.3.4 pH Effects on sol-hMGL NMR Spectral Features 2.3.5 hMGL Global Conformational Changes 2.3.6 H272 Mutational and Temperature Effects on the sol-hMGL NMR Spectrum and Enzymatic Activity 2.3.7 Active-site Paraoxon Covalent Binding 2.4 Discussion 2.5 References

Chapter 3. Effects of Distal Mutations on the Structure, Dynamics and Catalysis f Human Monoacylglycerol Lipase………………………………………………………………………59 3.1 Introduction 3.2 Experimental Procedures 3.2.1 Site-directed mutagenesis, expression and purification 3.2.2 Assay for MGL enzymatic activity 3.2.3 Nuclear magnetic resonance spectroscopy and data analysis 3.2.4 Mass spectrometry of wild type hMGL and W289L hMGL mutant 3.2.5 HDX-MS and HDX data processing 3.2.6 Molecular dynamics simulation 3.3 Results 3.3.1 Selection of distal residues for substitution 3.3.2 Expression, purification and molecular characterization 3.3.3 Impact of distal amino acid substitutions on hMGL catalysis and identification of essential residues 3.3.4 Downfield 1H NMR spectral pattern of hMGL as a probe for conformational analysis 3.3.5 Effect of distal mutations on NMR spectra and on conformational equilibria 3.3.6 Global structural changes of hMGL due to distal local perturbations 3.3.7 accessibility to the active site of hMGL mutants 3.3.8 HDX-MS analyses of wild type hMGL and W289L mutant 3.3.9 Nanosecond MD simulations for wild type hMGL and W289L mutant 3.4 Discussion 3.5 References

Chapter 4. The Functional Role of Human Monoacylglycerol Lipase Residues Involved in the Active Site Hydrogen Bonding Networks………………………………………………100

4.1 Introduction 4.2 Experimental Procedures 4.2.1 Mutant constructions, expression and purification 4.2.2 Enzyme hydrolysis assays with native substrate 4.3 Results 4.3.1 Selection of amino acid for mutational analysis 4.3.2 Consequences of breaking the hydrogen bond between His269 and Asp239 4.3.3 Effect of nucleophile Ser122 replacement 4.3.4 Active site double mutations based on S122C 4.3.5 Additional residues that maintain the active site stability 4.3.6 Molecular dynamic simulations 4.4 Discussion 4.5 References

Chapter 5. Amino acid cluster responsible for the regulation of hMGL conformational equilibrium…………………………………………………………………………………….133

5.1 Introduction 5.2 Experimental Procedures 5.2.1 Enzymatic expression and purification 5.2.2 Enzyme hydrolysis assays 5.2.3 NMR spectroscopy 5.3 Results 5.3.1 Mutagenesis sites selection 5.3.2 Hydrogen-bonded histidine resonances as natural probes for NMR detection of hMGL conformational transition 5.3.3 Residues His54-Asp197 as conformational determinant 5.3.4 Arg57 and Tyr58 are involved in lid regulation 5.3.5 Ligand accessibility to hMGL active site 5.4 Discussion 5.5 References

Chapter 6. Characterization of hMGL interaction with small molecule inhibitors………..151 6.1 Introduction 6.2 Experimental Procedures 6.2.1 NMR spectroscopy 6.2.2 Fluorescent enzyme assay 6.3 Results and Discussion 6.3.1 hMGL interactions with transition state analogues (TSA) 6.3.2 hMGL interactions with irreversible covalent compounds 6.3.3 hMGL interactions with product analogues 6.3.4 NMR-based screening method 6.4 Conclusion

ACKNOWLEDGEMENTS

Dr. Alexandros Makriyannis Dr. David Janero Dr. Nikolai Zvonok Dr. Lee Makowski Dr. Hongwei Huang (external faculty member)

Sergiy Tyukhtenko, Spiros Pavlopoulos, Jodi Anne Wood, Kiran Vemuri, Ioannis Karageorgos, Vidyanand Shukla and Jason Guo for training and countless productive conversations.

I would like to thank all my fellow doctoral students and lab-mates for their cooperation, stimulating discussions and of course friendship. In addition, I would like to express my gratitude to the staff of Centre for Drug Discovery for their help in the last six years. Last but not the least, I would like to thank my family members; especially my parents and my husband for their immense support since the very beginning of my studies, and my life in general.

Funding from the National Institute of Health / National Institute on Drug of Abuse: grants DA 3801 and DA 9152

1 List of Tables Page No. 2.1 Steady-state kinetic parameters for the hydrolysis of hMGL native substrate 2- AG by sol-hMGL and mutant enzymes at pH 7.4. 3.1 Steady state kinetics parameters of sol-hMGL and the mutants based on the hydrolysis of endogenous substrate 2-AG. Each measured value is listed with the corresponding 1 uncertainty 3.2 Percentage deuterium uptake levels for the peptides detected in both wt hMGL and W289L for D2O immersion of 30 s to 4 h. The data points are presented as mean ± standard error (1), which are computed from data observed in three replicate experiments. The values are adjusted for back exchange. 4.1 Summary of kinetic parameters of sol-hMGL and the mutants found experimentally for the hydrolysis of the native substrate 2-AG (pH7.4). 5.1. Comparison of kinetic parameters of sol hMGL and the mutant’s enzymatic activity on the hydrolysis of the native substrate 2-AG at 37OC 6.1. Inhibition parameters determined for each compound against sol-hMGL using the high- throughput fluorogenic substrate.

2 List of Figures Page No. 1.1 The schematic representation of the endocannabinoid system 1.2 The degradation pathways of the endocannabinoid endogenous ligands and 2- AG by the primary endocannabinoid enzymes FAAH and MGL 1.3 Three-dimensional structure of the endocannabinoid membrane-bound enzyme FAAH (PDB ID: 1MT5) 1.4 The primary sequence of hMGL with the catalytic triad residues highlighted in red 1.5 The three-dimensional structure of the wild type hMGL (PDB ID: 3HJU) 2.1 (A) Comparison of 1H NMR downfield resonances for, sol-hMGL and specific mutations from which assignments were made (pH7.4, T=310K). (B) Partial 1H-15N HSQC spectrum showing resonances from histidine side chains in the sol-hMGL construct (pH7.4, T=310K). C) The spectrum of the S122C mutant showing the effect of protonation on the H269 Hδ1 resonance (pH7.4, T=275K). D) The structure of hMGL highlighting lid sub-domain and the relative positions of the mutated residues. 2.2 Crystal structures showing (A) Hydrogen bonded H269 in the catalytic triad (S122-H269- D239) of hMGL (3HJU), (B) H49 involvement in the hydrogen bonding network within the oxyanion hole (G50, A51, M123) (3HJU), (C) The H54 residue in the open form (3HJU) and the closed form (3PE6). In contrast to these X-ray structures, mutagenesis with NMR clearly demonstrated that H54 is hydrogen bonded to D197 in the open form and is not bonded in the closed form. (D) The H103-H75 residues located at the interface between strand β4 and helix α2 forming a bridge between these secondary structures (3PE6). 2.3 (A) Comparison of downfield 1H NMR resonances of the soluble S122C and S122C/H269A mutant (pH7.4, T=275K). Effect of substitution of active site His269 for alanine. (B) Comparison of 1H NMR downfield resonances of sol-hMGL with resonances of soluble H103A and H75A mutants (pH7.4, T=310K). 2.4 Effect of pH on the downfield (A) and upfield (B) side chain resonances of hMGL (T=310K). The red arrows indicate the change of H269 chemical shifts with increasing pH. Relative populations of the open and closed hMGL conformations were calculated from the areas of H54 (C) and H269 (D) resonances 2.5 (A) Superposition of the 2D 1H-15N HSQC spectra of 15N-labeled sol-hMGL (pH7.4, T=300K) in the active (red) and in the inactive (blue) conformations. (B) Superposition of the 2D 1H-13C HSQC spectra of 13C-labeled sol-hMGL in the active (red) and in the inatctive (blue) conformations (aliphatic region) and (C) aromatic region. (D) 1D 1H NMR downfield resonances of individual sol-hMGL conformers at pH 7.4 and T=300 K. 2.6 (A) Open hMGL conformation stabilized by a cation-π interaction between R57 (arginine switch) and H272 (3HJU) and (B) the closed conformation with R57 flipped away from H272 (3PE6).

3 2.7 (A) Effect of H272 mutations on the population of conformers (pH 7.4, T=310 K), (B) Effect of temperature on the open-closed equilibrium for the H272S mutant (pH 7.4) 2.8 (A) The effects of paraoxon on the downfield resonances of sol-hMGL and H272A (pH 7.4, T=310 K). (B) The effects of paraoxon on the downfield resonances of H54A. 3.1 A, the overall hMGL 3D structure (PDB ID:3HJU) showing the location of two conserved Trp-35 and Trp-289 residues relative to the active site of the enzyme. The lid domain is highlighted in magenta. B, fragment of this structure zoomed in a view of the distal residues Trp- 289 and Leu-232 residing more than 18 Å away from the catalytic triad residues. C, close-up view of the identified remote site. Interactions between side-chains of Trp-289, Leu-232 and Arg-293 are highlighted. 3.2 Comassie blue stained SDS-PAGE analyses of purified enzymes expressed from E-coli BL 21(DE3) Cells. Lane 1, Molecular mass standards (kDa, Biorad); 2, sol hMGL; 3, W35A; 4; W289A; 5, W289L; 6, W289F; 7, L232G and 8, R293A. 3.3 Substrate saturation curves for sol-hMGL and tryptophan mutants W35A, W289L obtained by fitting the reaction velocities in Michaelis- Menten equation 3.4 A, slow spontaneous conversion of sol-hMGL open state through the mixture of conformers towards the closed state in solution (pH 7.4, T=310K). B, reversible, temperature induced alterations in the open-closed conformational equilibrium of the H272A mutant. The lid domain is highlighted in magenta. 3.5 A, the downfield 1H NMR spectra of sol-hMGL with temperature dependence behavior. The effect of single-point mutations and temperature for B, W35A; C, W289L; D, W289F; E, R293A and F, L232G (pH 7.4). 3.6 Superposition of two-dimensional 1H-15N HSQC spectra of 15N-labeled proteins A, open sol- hMGL (red) and W289F (blue), B, open sol-hMGL(red) and W289L (blue), C, closed sol-hMGL (red) and W289F (blue) and D, closed sol-hMGL (red) and W289L (blue) at 300K, pH 7.4. 3.7 Effect of hMGL binding with Compound-1 on the downfield resonances of A, sol-hMGL; B, W35A; C, W289F; D, R293A; E, W289L and F, L232G at pH 7.4, T=310K 3.7 Effect of hMGL binding with Compound-1 on the downfield resonances of A, sol-hMGL; B, W35A; C, W289F; D, R293A; E, W289L and F, L232G at pH 7.4, T=310K 3.8 Sequence coverage map for peptic peptides that were identified by MS/MS spectra for: A, wt hMGL and B, W289L mutant. The peptides are presented as bars. 3.9 Plots of deuterium uptake vs time (30 s, 5 min, 15 min, 1 h and 4 h) for the 12 peptides showing statistically different behavior for W289L mutant as compared to wt hMGL. Uncertainties (1), indicated by bars, reside inside the plotted symbol. 3.10 Differential deuterium uptake profiles of wt-hMGL and W289L at 4 h mapped onto the crystal structure of hMGL (PDB:3HJU). Increased differential deuterium uptakes were color-coded from yellow to red and reduced uptake was colored blue. Grey indicates regions where the deuterium uptake levels in the wt hMGL and W289L mutant were statistically the same. Grey, in addition, denotes regions where peptic peptides were not observed.

4 3.11 A, RMSD evolution for Cα atoms of wt hMGL (red) and W289L (blue) during a 200 ns MD simulation. B, the calculated R.M.S.F. values for the Cα atoms of wt hMGL and W289L with respect to the initial conformation (open) during the 200 ns MD simulations. The residues comprising the lid domain are highlighted.

3.12 The distance between Cα atoms of Phe159 and Gly210 for wt hMGL (red) and W289L (blue) for MD simulations of 200 ns duration.

3.13 MD simulation results for wt hMGL and W289L mutant. Black dashed circles mark the substrate entrance area into the hMGL active site (binding pocket). A, The wild type retains the open conformation throughput the 200 ns simulation. B, W289L mutant moves towards the restricted conformation after 100 ns. Surface colors illustrate the electrostatic potential with red representing negative charges and blue representing positive charges. 3.14 Overlapping of W289L three-dimensional structures at 0 ns (red) and after 200 ns (blue) of MD simulation study. The Cα distance between the two representative residues Ser-155 and Gly- 177 decreased 2-fold. The black arrows indicate the direction of the helix/loop movement. 4.1 Crystal structure of apo- hMGL (PDB ID: 3HJU) (left) showing the position of catalytic triad. The lid-domain is highlighted in magenta. Close-up of the catalytic site (right) encompassing the catalytic residues that are crucial for hydrolysis function. 4.2 Network of moderate electrostatic hydrogen bonds that stabilize the catalytic triad of hMGL (A) Apo- hMGL in open conformation (PDB ID: 3HJU, chain A) and (B) Holo-hMGL in the closed conformation (PDB ID: 3PE6). 4.3 The downfield region of the 1D NMR spectra for hMGL and the mutants for residues His269 and Asp239 that are involved in the hydrogen bond formation in enzymatic active site 4.4 1H NMR spectra of D239A substitution with decreasing temperatures (from 310 K to 280 K) demonstrating an irreversible population shift towards closed conformation in solution, pH 7.4 4.5 Effect of conservative and non-conservative mutations of the active site residue Ser122 on (A) the downfield region of the 1D NMR spectra (pH 7.4, T=290K) and (B) catalytic efficiencies of the enzymes 4.6 1H NMR spectra of S122T substitution with decreasing temperatures (from 310K to 280K) demonstrating a reversible population shift, with open conformation being stabilized at lower temperatures in solution, pH 7.4 4.7 . Effect of the double mutations based on the S122C construct on (A) the downfield region of the 1D NMR spectra (pH 7.4, T=290K) and (B) catalytic efficiencies of the enzymes.

4.8 Effect of mutation of the auxillary residues His121 to alanine on (A) the downfield region of 1D NMR spectra (pH 7.4, T=310K), and (B) enzyme kinetics 4.9 Effect of mutation of the accessory residues Leu241 and Cys242 to alanine on (A) the downfield region of 1D NMR spectra (pH 7.4, T=310 K), and (B) enzyme kinetics. 4.10 (A) RMSD evolution for Cα atoms of wt hMGL (black) and D239A (green) during 150ns MD simulation time, (B) The calculated R.M.S.F. values for the Cα atoms of wt hMGL and D239A

5 with respect to the initial conformation (open) during the 150ns MD simulations. The residues comprising the lid domain are highlighted.

4.11 MD simulations of wt hMGL and D239A mutant. Black dashed circles mark the substrate entrance area into the hMGL active site (binding pocket). (A) The wild type remains in the open (active) conformation throughput the 150ns simulation, (B) D239A mutant moves towards the restricted conformation after 75ns. Surface colors illustrate the electrostatic potential, with red representing negative charges, and blue representing positive charges.

4.12 Superimposition of snap shot picture of D239A mutant (red) over wild type hMGL (blue) after 150ns MD simulations. The distance between two residues Gly177 and Ley241 decreases by half in the mutant-induced closed conformation of D239A.

4.13 (A) RMSD evolution for Cα atoms of wt hMGL (black) with S122A (red) and S122T (blue) during 150ns MD simulation time, (B) The calculated R.M.S.F. values for the Cα atoms of wt hMGL with S122A (red) and S122T (blue) with respect to the initial conformation (open) during the 150ns MD simulations. The residues comprising the lid domain are highlighted.

4.14 Surface representation of hMGL and the mutants showing access to the deeply buried active site from 150 ns long MD simulation run.

5.1. (A) the three-dimensional structure of hMGL (PDB ID:3HJU) showing the residues involved in the catalytic triad, and those selected for mutagenesis in this study, (B) A network of hydrogen bonding network and aromatic interactions between the hub residues selected for mutagenesis that putatively control open/closed conformational equilibria. 5.2. Substrate saturation curves for sol hMGL and the mutated enzymes H54A and D197A in phosphate buffer (pH 7.4 at 37oC, 20 min substrate-enzyme incubation). 5.3. Downfield region of the 1D proton NMR spectra for (A) sol hMGL, and the mutants (B) H54A, (C) D197A to confirm their conformational state with temperature dependence (pH 7.4) 5.4. Plots of the relative changes in % catalytic efficiency for the mutated enzymes with respect to sol-hMGL 5.5. Downfield 1H NMR based temperature dependent study of hMGL mutations (A) R57A, (B) R57K, (C) Y58A and (D) Y58F at pH 7.4 6.1. Schematics of interaction between hMGL catalytic triad residues and small molecule transition state analogues. 6.2. Effect of TSA binding on the downfield region of 1D 1H NMR spectra of H103A sol-hMGL variant at T=310K (pH 7.4). The final protein: ligand ratio is 1:2. 6.3. Reaction mechanism between catalytic Ser122 and endogenous ligand (a) 2-AG and (b) AM 6580 [1] 6.4. Effect of covalent inhibitors binding on the downfield region of 1D 1H NMR spectra of sol- hMGL variant at T=310K (pH 7.4). The final protein: ligand ratio is 1:2.

6 6.5. Piperazine-pyridine based reversible MGL inhibitors classified as product analogues. 6.6. Close up of hMGL crystal structure in complex with AM 10212 6.7. Effect of reversible inhibitors binding on the downfield region of 1D 1H NMR spectra of sol- hMGL variant at T=310K (pH 7.4). The final protein: ligand ratio is 1:2. 6.8. Effect of reversible and irreversible inhibitors binding on the downfield region of 1D 1H NMR spectra of sol-hMGL variant at T=310 K (pH 7.4). The final protein: ligand ratio is 1:2.

7 Abbreviations

THC CBR Cannabinoid receptors eCB Endocannabinoid system CB1 Cannabinoid 1 CB2 2 GPCR G-protein coupled receptors CNS Central nervous system GABA gamma-Amino butyric acid cAMP cyclic adenosine monophosphate PKA Protein kinase A AEA Anandamide 2-AG 2-Arachidonoyl FABP binding protein NAE N-acylethanolamine NAPE N-arachidonoyl DGL FAAH Fatty acid amide MGL Monoacylglycerol lipase ABHD 6 alpha/beta-Hydrolase domain 6 ABHD 12 alpha/beta-Hydrolase domain 12 MAFP Methyl arachidonyl Fluorophosphonate hMGL Human monoacylglycerol lipase AA Arachidonic acid NMR Nuclear magnetic resonance HSQC Heteronuclear single quantum correlation sol-hMGL soluble hMGL HDX-MS Hydrogen deuterium exchange mass spectroscopy

8 MD Molecular dynamics CSP Chemical shift perturbations DMSO Dimethyl sulfoxide AHMMCE N-arachidonoyl, 7-hydroxy-6-methoxy-4-methyl coumarin ester

9

Chapter 1:

Introduction Endocannabinoid System

The endocannabinoid system, named after is well-known to play a critical role in the body homeostatis. The use of cannabis (marijuana) dates back to 2500 B.C. Ancient Indian and

Chinese manuscripts acknowledge such use for medicinal as well as recreational purposes, though the pharmacological basis of the physiological effects remained a mystery until the discovery of the active ingredient tertrahydrocannabinol (THC) [2]. Mechoulam first isolated and synthesized

THC in the 1960s [3]. In the ensuing decades, the cannabinoid receptors were discovered, and shown to mediate the highly stereo-specific effects of THC [4]. The cannabinoid receptors (CBR), its endogenous neurotransmitters, and anabolic-catabolic enzymes are jointly referred to as the endocannabinoid (eCB) system [5, 6].

1.1 The Endocannabinoid System:

1.1.1 Cannabinoid Receptors: The eCB system comprises of the cannabinoid receptor 1 (CB1) and cannabinoid receptor 2 (CB2), both of which are members of the G-protein-coupled receptors

(GPCRs) family, with seven transmembrane helices [7]. Mounting evidence points to the existence of an additional CB3 receptor, to explain cannabinoid effects that are not mediated through CB1 and CB2 [8]. The CB1 receptors are primarily found on presynaptic neurons in the central nervous system (CNS), and are the most abundant of the CNS GPCRs, predominantly residing on -

10 aminobutyric acid (GABA)-ergic and glutamatergic neurons [9, 10]. They are involved in the motivation, movements and cognition process and are densely expressed in the sensory and motor region [11]. CB2 receptors, were originally thought to be expressed only in the immune system, but have recently been identified in the CNS as well, especially on microglial

Figure 1.1. The schematic representation of the endocannabinoid system [12].

11 cells [13]. Their role through the lipid cannabinoid signaling is believed to be ‘protective’, and responsible for immune suppression [14]. CB1 and CB2 receptors exert their biological role by coupling with heterotrimeric Gi/0 proteins, thereby decreasing accumulation of cyclic adenosine monophosphate (cAMP). This in turn inhibits the cAMP-dependent protein kinase A (PKA) and stimulates mitogen-activated protein kinase (MAPK) activity to alter synaptic plasticity [11, 15,

16].

1.1.2 Endocannabinoids: After the breakthrough discovery that CB receptors are activated by the plant-derived substance THC, further investigations into the endogenous native ligands revealed anandamide (AEA) and 2-arachidonoyl glycerol (2-AG) as the two primary eCB neurotransmitters

[17, 18]. Along with the other endocannabinoids, AEA and 2-AG are oxidative products of 20- carbon long fatty acids [19]. AEA acts as an agonist at CB2 receptors, but shows much lower efficacy at CB1 receptors [20]. In contrast, 2-AG is present in the brain at much higher levels than

AEA, and serves as a retrograde messenger at synapses [21]. Unlike most other neurotransmitters,

AEA and 2-AG are not stored in vesicles, but are synthesized on demand in response to neural activity [22].

Fatty acid binding proteins (FABPs) are intracellular carriers for the endocannabinoid bioactive lipids, N-acylethanolamine (NAEs) [23]. FABPs are an integral part of endocannabinoid system and are FABP inhibitors are being developed as pharmacological inhibitors of endocannabinoid uptake [24].

1.1.3 Endocannabinoid Enzymes: Biosynthetic and degradative enzymes tightly regulate the endocannabinoid signaling in brain [25]. Anandamide is synthesized from its phospholipid precursor N-arachidonoyl phosphatidylethanolamine (NAPE) by the enzyme N-acyltransferase

12 [26], and diacylglycerol lipase (DGL); whereas a membrane-associated post-synaptic enzyme controls 2-AG synthesis from C [27]. The catabolic enzymes- Fatty Acid Amide

Hydrolase (FAAH) and Monoacylglycerol Lipase (MGL) serve to terminate the synaptic transmission of the endocannabinoid ligands AEA and 2-AG. AEA is inactivated by cellular uptake and subsequent catabolism by FAAH, which shows /amidase activity [28]. MGL is largely responsible for the hydrolysis of 2-AG in brain; it co-localizes on presynaptic neuron terminals that express CB1 receptors [29]. An activity-based protein profiling (ABPP) of mouse brain tissue shows two additional enzymes also contribute modestly to 2-AG hydrolysis: α/β- hydrolase 6 (ABHD6) and α/β- hydrolase 12 (ABHD12) [25]. These endocannabinoid degradative enzymes are members of the family, which belongs to the largest and most diverse class of enzymes in human body, the proteases [30]. These are classified into serine proteases

(trypsin/chymotrypsin/subtilisin enzymes) and ‘metabolic’ serine that cleave ester, amide, or thioester bonds in small molecules or proteins [31]. Most metabolic serine hydrolases have an α/β- hydrolase fold, characterized by 8 central -sheets surrounded by -helices, and an active site serine GXSXG sequence [32].

Figure 1.2. The degradation of the endocannabinoid endogenous ligands anandamide and 2-AG by the primary endocannabinoid enzymes [33].

13 FAAH is an integral membrane protein with a catalytic triad Ser- 241/Ser-217/Lys-142 [34]. A 2.8

Å structure of FAAH in complex with the irreversible inhibitor methoxy arachidonyl fluorophosphonate (MAFP) revealed several unusual features of the enzyme [35]. First, the core catalytic machinery of FAAH is composed of a serine–serine–lysine catalytic triad

(S241/S217/K142), in contrast to the serine–histidine–aspartate triad of monoacylglycerol lipase

[36]. The structure of FAAH also revealed that this enzyme possesses a remarkable collection of channels that appear to monitor simultaneous access to both the membrane and cytoplasmic compartments of the cell [35].

Figure 1.3. Three-dimensional structure of the endocannabinoid membrane-bound enzyme FAAH (PDB ID: 1MT5).

1.2 Monoacylglycerol Lipase

14 Human MGL (hMGL) is a 33 kDa protein that shares 92% sequence similarity with mouse MGL, and 84% with rat MGL [37, 38]. The MGL catalytic triad is comprised of Ser122/His269/Asp239, with a histidine-glycine dipeptide, considered to be the hallmark of [38]. Ser122 is located in the GXSXG consensus sequence within a sharp turn between helix 3 and strand β5 to form the nucleophillic elbow [39]. The backbone amide bond from Ala51 and Met123 stabilize the tetrahedral intermediate by forming the ‘oxyanion hole’ [40].

Figure 1.4. The primary sequence of hMGL with the catalytic triad residues highlighted in red. The wild type enzyme was engineered with N-terminal His-tag and L169S/L176S substitution to facilitate the protein purification for further biochemical characterization.

MGL is found mainly in the brain, but also in peripheral tissue such as ovary, testis, adipose tissue, adrenal glands and heart. MGL has a very strong selectivity for monoacylglycerols over di- and

15 tri-acylglycerols, and essentially hydrolyzes 2-AG to arachidonic acid (AA) and glycerol [35]. It is a membrane-associated protein that is also present in the cytosolic fraction.

Figure 1.5. The three-dimensional structure of the wild type hMGL (PDB ID: 3HJU).

Four different X-ray crystal structures of hMGL have been published, with and without the ligand

[1, 39-41]. The major difference between the two structures is the orientation of a lid-like structure comprising ~75 residues (helix 4 & 6) that regulate access to the binding site [1, 39, 40].

Substrate hydrolysis by most lipases is controlled by the interfacial activation at the lipid-water boundary. It has been hypothesized that the movement of lid is responsible for such activation in

MGL [42]. The engagement of any lipophilic hMGL ligand may involve the lateral diffusion

16 within the membrane bilayer to access the substrate binding channel via the enzyme’s membrane associated lid domain [43].

1.3 MGL as a therapeutic target

Increased levels of AEA are reported in the synovial fluids of patients with arthritis, and a potent and selective FAAH inhibitor (PF-04457845) was evaluated in clinical trials for the treatment of pain in patients with osteoarthritis [44, 45]. However, the therapeutic development of MGL inhibitors has been slower than FAAH inhibitors, due to a lack of selective and potent inhibitors.

Currently, there are no approved or available FDA-approved MGL inhibitors. Use of MGL inhibitors is mainly indicated for neuropathic pain and for inflammation. MGL knock-out mice show elevated levels of 2-AG in the brain, and CB1 receptor desensitization [46, 47]. The cannabimimetic effects - analgesia and hypomotility – occur in the absence of cognition impairment or the psychotropic side effects, making MGL a highly promising pharmacological target [48]. Efforts to inactivate MGL pharmacologically have also confirmed its therapeutic potential, but the compounds suffer from cross-reactivity with FAAH, , and other serine hydrolases [49].

MGL is also highly over-expressed in aggressive human cancer cells and primary tumors, however this overexpression does not involve the cannabinoid system [50]. MGL regulates a fatty acid network that is enriched in oncogenic signaling lipids that promote migration, invasion, survival and in vivo tumor growth [51]. Genetic inactivation of MGL also attenuates neuroinflammation and lowers amyloid β levels and plaques in an Alzheimer’s mouse model [52]. Considering the critical role hMGL plays in the regulation of the endocannabinoid system, it has emerged as an attractive druggable target, for various indications such as pain, inflammation, appetite and cancer.

17 References:

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18 18. Rouzer, C.A. and L.J. Marnett, Endocannabinoid oxygenation by cyclooxygenases, lipoxygenases, and cytochromes P450: cross-talk between the eicosanoid and endocannabinoid signaling pathways. Chem Rev, 2011. 111(10): p. 5899-921. 19. Rouzer, C.A. and L.J. Marnett, Endocannabinoid oxygenation by cyclooxygenases, lipoxygenases, and cytochromes P450: cross-talk between the eicosanoid and endocannabinoid signaling pathways. Chemical reviews, 2011. 111(10): p. 5899-5921. 20. Labar, G., J. Wouters, and D. Lambert, A review on the monoacylglycerol lipase: at the interface between fat and endocannabinoid signalling. Current medicinal chemistry, 2010. 17(24): p. 2588-2607. 21. Alger, B.E., Endocannabinoids and Their Implications for Epilepsy. Epilepsy Curr, 2004. 4(5): p. 169-73. 22. Kaczocha, M., et al., Fatty acid-binding proteins transport N-acylethanolamines to nuclear receptors and are targets of endocannabinoid transport inhibitors. Journal of Biological Chemistry, 2012. 287(5): p. 3415-3424. 23. Berger, W.T., et al., Targeting fatty acid binding protein (FABP) anandamide transporters–a novel strategy for development of anti-inflammatory and anti-nociceptive drugs. PLoS One, 2012. 7(12): p. e50968. 24. Blankman, J.L., G.M. Simon, and B.F. Cravatt, A comprehensive profile of brain enzymes that hydrolyze the endocannabinoid 2-arachidonoylglycerol. Chemistry & biology, 2007. 14(12): p. 1347-1356. 25. Okamoto, Y., et al., Molecular characterization of a generating anandamide and its congeners. Journal of Biological Chemistry, 2004. 279(7): p. 5298- 5305. 26. Tanimura, A., et al., The endocannabinoid 2-arachidonoylglycerol produced by diacylglycerol lipase α mediates retrograde suppression of synaptic transmission. Neuron, 2010. 65(3): p. 320-327. 27. Patricelli, M.P. and B.F. Cravatt, Clarifying the catalytic roles of conserved residues in the amidase signature family. Journal of Biological Chemistry, 2000. 275(25): p. 19177- 19184. 28. Ahn, K., M.K. McKinney, and B.F. Cravatt, Enzymatic pathways that regulate endocannabinoid signaling in the nervous system. Chemical reviews, 2008. 108(5): p. 1687-1707. 29. Bachovchin, D.A. and B.F. Cravatt, The pharmacological landscape and therapeutic potential of serine hydrolases. Nature reviews Drug discovery, 2012. 11(1): p. 52-68. 30. Long, J.Z. and B.F. Cravatt, The metabolic serine hydrolases and their functions in mammalian physiology and disease. Chemical reviews, 2011. 111(10): p. 6022-6063. 31. Holmquist, M., Alpha beta-hydrolase fold enzymes structures, functions and mechanisms. Current Protein and Peptide Science, 2000. 1(2): p. 209-235. 32. Griffing, G.T., Endocannabinoids–the Brain's Own Marijuana–May Be Linked to the Metabolic Syndrome. Medscape General Medicine, 2006. 8(4): p. 7. 33. McKinney, M.K. and B.F. Cravatt, Evidence for distinct roles in catalysis for residues of the serine-serine-lysine catalytic triad of fatty acid amide hydrolase. Journal of Biological Chemistry, 2003. 278(39): p. 37393-37399. 34. Bracey, M.H., et al., Structural adaptations in a membrane enzyme that terminates endocannabinoid signaling. Science, 2002. 298(5599): p. 1793-6.

19 35. Cravatt, B.F. and A.H. Lichtman, Fatty acid amide hydrolase: an emerging therapeutic target in the endocannabinoid system. Current opinion in chemical biology, 2003. 7(4): p. 469-475. 36. Karageorgos, I.L., The mechanism of monoacylglycerol lipase (MGL) inactivation: a study using nuclear magnetic resonance spectroscopy and mass spectrometry. 2012. 37. Karlsson, M., et al., cDNA cloning, tissue distribution, and identification of the catalytic triad of monoglyceride lipase Evolutionary relationship to , , and haloperoxidases. Journal of Biological Chemistry, 1997. 272(43): p. 27218-27223. 38. Labar, G., et al., Crystal structure of the human monoacylglycerol lipase, a key actor in endocannabinoid signaling. Chembiochem, 2010. 11(2): p. 218-227. 39. Schalk‐Hihi, C., et al., Crystal structure of a soluble form of human monoglyceride lipase in complex with an inhibitor at 1.35 Å resolution. Protein Science, 2011. 20(4): p. 670- 683. 40. Bertrand, T., et al. Structural basis for human monoglyceride lipase inhibition. J Mol Biol, 2010. 396, 663-73 DOI: 10.1016/j.jmb.2009.11.060. 41. Griebel, G., et al., Selective blockade of the hydrolysis of the endocannabinoid 2- arachidonoylglycerol impairs learning and memory performance while producing antinociceptive activity in rodents. Scientific reports, 2015. 5: p. 7642. 42. van Pouderoyen, G., et al., The crystal structure of Bacillus subtili lipase: a minimal α/β hydrolase fold enzyme. Journal of molecular biology, 2001. 309(1): p. 215-226. 43. Nasr, M.L., et al., Membrane phospholipid bilayer as a determinant of monoacylglycerol lipase kinetic profile and conformational repertoire. Protein Science, 2013. 22(6): p. 774- 787. 44. Ahn, K., et al., Mechanistic and pharmacological characterization of PF-04457845: a highly potent and selective fatty acid amide hydrolase inhibitor that reduces inflammatory and noninflammatory pain. Journal of Pharmacology and Experimental Therapeutics, 2011. 338(1): p. 114-124. 45. Johnson, D.S., et al., Discovery of PF-04457845: a highly potent, orally bioavailable, and selective urea FAAH inhibitor. ACS medicinal chemistry letters, 2010. 2(2): p. 91- 96. 46. Pan, B., et al., Alterations of endocannabinoid signaling, synaptic plasticity, learning, and memory in monoacylglycerol lipase knock-out mice. The Journal of Neuroscience, 2011. 31(38): p. 13420-13430. 47. Zhong, P., et al., Genetic deletion of monoacylglycerol lipase alters endocannabinoid‐ mediated retrograde synaptic depression in the cerebellum. The Journal of physiology, 2011. 589(20): p. 4847-4855. 48. Mulvihill, M.M. and D.K. Nomura, Therapeutic potential of monoacylglycerol lipase inhibitors. Life Sci, 2013. 92(8-9): p. 492-7. 49. Wise, L.E., et al., Dual fatty acid amide hydrolase and monoacylglycerol lipase blockade produces THC-like Morris water maze deficits in mice. ACS chemical neuroscience, 2012. 3(5): p. 369-378. 50. Nomura, D.K., et al., Monoacylglycerol lipase regulates a fatty acid network that promotes cancer pathogenesis. Cell, 2010. 140(1): p. 49-61. 51. Ye, L., et al., Monoacylglycerol lipase (MAGL) knockdown inhibits tumor cells growth in colorectal cancer. Cancer letters, 2011. 307(1): p. 6-17.

20 52. Chen, R., et al., Monoacylglycerol lipase is a therapeutic target for Alzheimer's disease. Cell reports, 2012. 2(5): p. 1329-1339.

21 Chapter 2

Direct NMR Detection of the Conformational Transition Between Open and Closed Forms

of Monoacylglycerol Lipase

Parts of this chapter have been published

Tyukhtenko, S., Karageorgos, I., Rajarshi, G., Zvonok, N., Pavlopoulos, S., Janero, D. R., &

Makriyannis, A. (2016). Specific Inter-Residue Interactions as Determinants of Human

Monoacylglycerol Lipase Catalytic Competency A ROLE FOR GLOBAL CONFORMATIONAL

CHANGES. Journal of Biological Chemistry, 291(6), 2556-2565.

2.1 INTRODUCTION:

A member of the serine hydrolase superfamily, monoacylglycerol lipase (MGL) is largely responsible for the catalytic inactivation of the endocannabinoid signaling lipid, 2- arachidonoylglycerol, and regulates a fatty acid network that promotes tumorigenesis [53, 54]. On the basis of the preclinical efficacy of MGL genetic or pharmacological ablation against pain, inflammation, and cancer, MGL is considered an attractive therapeutic target [55]. Published crystal structures of modified/liganded forms of hMGL [56-58] indicate that the enzyme has a typical lipase structure that includes a lid sub-domain (residues 151-225) that can assume open or closed states and control, through its gating dynamics, substrate access to the active site [59-65].

The open state has been observed in the absence and presence of bound ligand [56, 57], whereas the hMGL closed conformation has only been reported in complex with a reversible inhibitor [58].

Inter-species conservation of the overall hMGL lid architecture and dynamics has been suggested from X-ray studies and molecular dynamics simulations of bacterial MGL (bMGL) (14-16).

22 Collective data from X-ray crystallographic studies of hMGL and bMGL variants have allowed inference that the lid sub-domain shows a high degree of conformational plasticity along a coordinate of catalytic activity and suggest the existence of a stochastic equilibrium that transitions the enzyme between open- and closed-lid conformations. Nonetheless, there are limits with which even a suite of static X-ray maps per se can reflect the conformational flexibility of a protein and structural influences on protein function [11]. The concern is underscored by the important roles that protein structural transitions play in enzyme catalysis and the influence of targeted ligands thereon as prospective drugs [12,13].

These considerations led us to probe hMGL structure-function correlations by using the catalytically active enzyme. In this regard, our previous work has demonstrated the effects of important interactions on hMGL structure, substrate affinity and/or activity. These interactions include, hMGL membrane association [66], binding of designer active site-directed inhibitors [67,

68], and covalent or mutational modifications of amino acid residues at or near the hMGL catalytic triad (S122-H269-D239). The structural effects of these interactions are not restricted to the hMGL lid domain, but also to discrete non-lid enzyme regions as well [66, 67]. Perhaps most strikingly, we identified by NMR a strong hydrogen-bond network and provisionally implicated H269 and

D239 of the catalytic triad and neighboring L241 and C242 residues therein. Not only did this hydrogen-bond network influence catalytic activity, but active site-directed inhibitors of different types were observed to alter this population of hydrogen bonds in concert with their inhibitory activity [67].

The influence of noncovalent amino-acid interactions in regulating, protein conformational transitions, protein ligand binding architecture, and ligand pharmacological activity is a topic of great current interest. For hMGL, the factors leading to conformational transitions of the enzyme

23 are not well defined, and the effects of specific intramolecular amino acid interactions on the hMGL catalytic efficiency have not been measured. Unlike other lipases (8,11), hMGL shows a high level of activity and conformational flexibility as an isolated enzyme without association to a lipid surface (17). It can therefore be utilized to determine important molecular details such as understanding hydrogen bond networks within an enzyme molecule that affect protein architecture and catalysis.

We demonstrate here simultaneous NMR detection of both active and inactive hMGL states in solution and report the unambiguous assignment of downfield NMR resonances that are highly sensitive to the equilibrium between these two states. These resonances of equal integral intensity represent the hydrogen-bonding pattern of the active conformer that is predominant at neutral pH. Significant changes in the intensities of the peaks with pH, temperature or point mutations are indicative of alterations of this hydrogen-bonding pattern when the equilibrium is shifted toward the inactive state. The well-appreciated role of hydrogen bonds as determinants of protein structure suggests that the hMGL inactivation observed reflects enzyme conformational changes that have their origin in the altered hydrogen-bonding network. The ability of the perturbations in the downfield region of the hMGL NMR spectra to be elicited in a predictable manner from different changes to physical conditions implies the existence of a regulated equilibrium between distinct enzyme conformations. The activity of hMGL may be modulated by structural determinants that can transition the enzyme between open-closed conformations.

2.2 EXPERIMENTAL PROCEDURES

2.2.1 hMGL Mutagenesis, Expression and Purification- Single (hMGL-H269A), double

(hMGL-L169S,L176S) (sol-hMGL), several triple hMGL mutants (sol-hMGL-H49A, sol-hMGL-

24 H54A, sol-hMGL-H103A, sol-hMGL-H75A, sol-hMGL-S122C, sol-hMGL-H272Y, sol-hMGL-

H272A, sol-hMGL-H272S) and sol-hMGL-H269A-S122C were generated using corresponding primers and Stratagene QuickChange site-directed mutagenesis kit (La Jolla, CA). All hMGL variants were generated as 6-His-tagged proteins to facilitate their purification by immobilized metal-affinity chromatography. The DNA primary structure of all mutants was confirmed by sequencing. The hMGL mutants were expressed in BL21 (DE3) E. coli cells, as previously detailed

[69]. In brief, a single E. coli colony containing the plasmids with appropriate hMGL mutations was inoculated into 10 ml of Luria broth/ampicillin (100 µg/mL) and grown overnight at 33°C with shaking (250 rpm). The next morning, these 10 mL were inoculated into 500 mL of

Luria broth/ampicillin (100 µg/mL) and allowed to grow at 33°C with shaking (250 rpm) until the culture reached an OD600 of 0.6-0.8. Expression was induced by adding isopropyl-β-D- thiogalactopyranoside (Fisher, Pittsburgh, PA) to a final concentration of 1 mM. After 5 h induction at 30°C, the cells were harvested by centrifugation at 5000 g for 10 min, washed with phosphate-buffered saline, and held at -80 °C. Uniformly 15N-labeled sol-hMGL samples were

15 prepared using the same procedure incorporating minimal media containing NH4 Cl and uniformly 13C-labeled samples were prepared in minimal media with [U-13C]-glucose (Cambridge

Isotope Labs).

For enzyme purification, three grams (wet-weight) of cells were resuspended in 20 mL lysis buffer (20 mM Na-phosphate, 200 mM NaCl, 1 mM DTT, pH 7.4 ) and 20 mL of xTractor buffer (Clontech, Mountain View, CA) supplemented with lysozyme (0.1 mg/mL) and DNase I

(25 µg/mL) (Fisher). After 20 min, the lysate was disrupted on ice by three, 1-min sonication cycles, each consisting of 1-s sonication bursts at a 50-W power level separated by a 5-s interval

(Vibra-Cell 500 W, Sonics, Newtown, CT). The resulting cell lysate was centrifuged at 20000 g

25 for 25 min at 4°C. hMGL was isolated by incubating the resulting supernatant with 4.0 mL (bed volume) pre-equilibrated Talon metal affinity resin (Clontech, Mountain View, CA) for 1 h at 4°C.

This suspension was transferred to a gravity-flow column and allowed to settle. The resin was washed twice with 20 mL lysis buffer containing 25 mM imidazole, and His-tagged hMGL was eluted with 12 mL lysis buffer containing 300 mM imidazole. Protein purity was evaluated using

Any kD Mini-PROTEAN TGX SDS-PAGE (Bio-Rad, Hercules, CA). Protein samples were denatured at 70°C for 5 min in Laemmli buffer containing 5% β-mercaptoethanol, resolved on

SDS-PAGE gels, and stained with Coomassie blue (Fisher). Prior to enzyme assays and NMR experiments, purified hMGL samples were dialyzed for 12 h to ensure thorough imidazole removal using a membrane with a molecular-weight cutoff of 10000-12000 Da. Enzyme concentration was

-1 -1 determined spectrophotometrically using the molar extinction coefficient ε280 = 24910 M cm .

Our sample preparation protocol routinely yields sol-hMGL in the active conformation.

Over time the active conformer undergoes spontaneous conformational switching to an inactive state. The spontaneous transition between the two states is extremely slow, suggestive of a high- energy barrier for the switch in our sol-MGL variant. The active conformer is stable for several weeks at room temperature. At 310K the transition rate is faster. Incubation of the sample at 310K for several days results in a complete transition to the inactive form. This enabled us to prepare distinct sol-hMGL conformers for NMR analysis.

2.2. Enzyme Assays- The endogenous hMGL substrate, 2-arachidonoylglycerol (2-AG), was used for the determination of catalytic parameters (Km, Vmax, Kcat). Hydrolysis of 2-AG to arachidonic acid (AA) by hMGL was monitored and quantified by HPLC. Briefly, 280 µL of assay buffer (50 mM Tris-HCl, 5 mM MgCl2, 1 mM EDTA and 0.1% BSA, pH 7.4) was preincubated with 15 µL

26 of each 2-AG stock dilution (final 2-AG concentrations, 13-400 µM), and the reaction was initiated by the addition of 5 µL purified hMGL (18 ng – 1 µg). Aliquots (50 µL) were taken immediately at the start of the incubation and after 20 min, diluted four-fold by volume with chilled acetonitrile to quench enzyme activity, and centrifuged at 20000 g for 5 min at 4°C. Supernatant (20 µL) was injected directly into a Waters Alliance 2695 HPLC system for analysis. In an 8-min run, 2-AG eluted at 3.0 min and AA at 6.0 min, allowing the reaction to be followed by either substrate (2-

AG) turnover or product (AA) formation. A gradient elution profile of 5% B

(water/acetonitrile/orthophosphoric acid = 54/40/6%) to 100% A (acetonitrile) at a 1 mL/min flow rate was used for separation on a ZORBAX Eclipse XDB-C18 reverse-phase (4.6 x 50 mm, 3.5

µm) column (Agilent Technologies, Santa Clara, CA). Analytes were quantified with external standards. The rate of AA formation was determined by subtracting the AA concentration at t=0 from that at t=20 min. Initial rates of 2-AG hydrolysis at the various substrate concentrations were determined. Vmax and Km values were then estimated by fitting the initial-rate data to the Michaelis-

Menten equation using nonlinear regression with GraphPad Prism 5.0 (San Diego, CA). All assays were performed in triplicate.

2.2.3 Sample Preparation and NMR Spectroscopy- Samples used for NMR analyses were 0.1 -

0.4 mM hMGL in 20 mM Na-phosphate, 200 mM NaCl , 1 mM DTT, 0.02% sodium azide, 95%

H2O/5% D2O at specified pHs. Sodium 2,2-dimethyl-2-silapentane-5-sulfonate (DSS) was added

(~20 µM) as an internal chemical shift reference (δ = 0.00 ppm). Sample volume was 0.6 mL. The pH was adjusted as desired by addition of microliter amounts of 0.1 M HCl or NaOH and measured using a Wilmad Labglass pH electrode (3 mm O.D. x 180 mm length) inserted into the protein solution in the 5 mm NMR tube at room temperature before and after NMR data collection. NMR

27 spectra were recorded at 16 pH values between pH 6.5 and pH 12 for sol-hMGL, since the enzyme precipitated below pH 6.

Ligand binding experiments were carried out using a 50 mM stock solution of paraoxon

(diethyl 4-nitrophenyl phosphate), dissolved in DMSO-d6 [70]. Typically, no more than 5 l of paraoxon solution was added to each enzyme sample to achieve a concentration of paraoxon twice that of the enzyme.

1D 1H NMR spectra were acquired at 700 MHz with a Bruker AVANCE II NMR spectrometer equipped with a 5 mm triple resonance inverse probe at 37°C. For optimal detection of downfield exchangeable proton resonances the 3-9-19 WATERGATE [71] pulse sequence

(p3919fpgp) with gradients and additional flipback pulse was used. This pulse sequence employs a binomial-like pulse train, which provides null excitation at the water frequency. The center of the maximal excitation region was 13.9 ppm, and the calculated delay for binomial water suppression was 39 µs at 700 MHz. In combination with a flipback pulse, the 3-9-19 sequence significantly prevents unwanted attenuation of downfield resonances from spin diffusion and chemical exchange with water. Routinely, 8K scans were accumulated. The 1H-15N Fast-HSQC

[72] spectrum of uniformly 15N-labeled sol-hMGL was recorded using a spectral width of 30 ppm and 140 ppm in 1H and 15N dimensions, respectively. The 1H transmitter was set to the frequency of the water resonance, and the 15N carrier frequency was set to 175 ppm to achieve maximum intensity for the histidine side chain resonances. Regular 1H-15N HSQC and 1H-13C HSQC NMR spectra were recorded using standard pulse sequences supplied with the AVANCE 700 spectrometer. All NMR data were processed with TopSpin software (Bruker). For 1D spectra exponential multiplication (broadening factor lb=20 Hz) was applied.

28 2.3 RESULTS

2.3.1 hMGL Variants Facilitating NMR Experiments- Recombinant wild-type hMGL required detergents during purification to maintain stability and prevent aggregation in concentrated solutions greater than 100 M. Aggregation of wt-hMGL resulted in significant broadening of

NMR resonance peaks and protein precipitation. To avoid the need for detergents and obviate enzyme aggregation/precipitation, a DNA construct expressing a soluble hMGL variant with two leucines substituted by two serines in the lid sub-domain (double L169S, L176S mutant) (sol- hMGL) was expressed and used in this study [58].

TABLE 2.1. Steady-state kinetic parameters for the hydrolysis of hMGL native substrate 2-AG by sol-hMGL and mutant enzymes at pH 7.4. Catalytic -1 Protein Km (M) kcat (s ) Fold Decrease Efficiency

sol-hMGL 22  4 (5.70.2) x 105 2.5 x 104

His269Ala - - ND -

His49Ala 36.6  6.2 (1.6±0.7) x 105 4.4 x 10-3 5

His54Ala 12  2.4 11 0.5 9.8 x 10-1 2.5 x 104

His103Ala 122  11.4 (1.40.5) x 104 1.3 x 103 20

His272Ser 30.8  8.8 (1.3  0.1) x 104 4.2 x 102 58

His272Ala 22.2  1.6 (4.5  0.1) x 104 2 x 103 13

His272Tyr 36.2  7.5 (7  0.3) x 104 2.2 x 103 12

ND: Not Detected

29 These substitutions resulted in a modest decrease in enzyme catalytic efficiency (kcat/Km) from 2.2

5 4 -1 -1 x 10 to 2.5 x 10 M s and no change in substrate affinity (respective Km values: 25 ± 6 and 22

± 4 M). Thus, hMGL function was not significantly compromised by these mutations, suggesting that overall enzyme conformation was also not adversely affected. This construct (310 aa, 34.1 kDa) formed the basis for introducing additional mutations to allow residue assignment of downfield NMR resonances. Sol-hMGL and mutants derived therefrom were expressed exhibited greater solubility and stability, overcoming aggregation issues and providing sharp NMR resonances (data not shown).

2.3.2 hMGL NMR Downfield Spectral Features- A striking feature of the hMGL 1D 1H NMR spectrum is the downfield region (12-18 ppm) that exhibits four well-resolved labile proton resonances (Fig. 1A). These signals are detectable with 15N chemical shifts in the range of 160-

180 ppm, as shown in the partial 1H-15N HSQC spectrum (Fig. 1B), and, consequently, correspond to nitrogen-bonded imidazole resonances from histidines [73]. The extreme downfield chemical shift of these protein resonances is a consequence of their participation in hydrogen-bonding interactions with neighboring residues [74].

2.3.3 hMGL NMR Amino-acid Resonance Assignments- 1H NMR signals for catalytic His protons involved in hydrogen-bonds in the active sites of serine proteases have been observed in the range of 12-19 ppm [75]. On this basis, we had provisionally assigned one signal in the downfield region to a H-bond between the catalytic H269 and D239 of hMGL. To assign experimentally and definitively hMGL His resonance peaks and define intramolecular details affecting hMGL function, we conducted single-point mutations of specific amino acid residues.

30 FIGURE 2.1. (A) Comparison of 1H NMR downfield resonances for, sol-hMGL and specific mutations from which assignments were made (pH7.4, T=310K). (B) Partial 1H-15N HSQC spectrum showing resonances from histidine side chains in the sol-hMGL construct (pH7.4, T=310K). C) The spectrum of the S122C mutant showing the effect of protonation on the H269 Hδ1 resonance (pH7.4, T=275K). D) The structure of hMGL highlighting lid sub-domain and the relative positions of the mutated residues.

Our rationale was based upon the concept that disappearance of a resonance in the downfield region of the NMR spectrum upon substitution of a given His residue by Ala would provide

31 definitive mutagenesis-based resonance assignment (Fig. 1A). To assign the catalytic His residue, a variant of sol-hMGL was prepared by substituting H269 with alanine. As a consequence of removal of the side-chain H-bond donor from the active site in this H269A mutant, only the downfield NMR signal at 14.9 ppm disappeared while all other signals in the sol-hMGL downfield region remained unchanged (Fig. 1A). Thus, the peak observed at 14.9 ppm, corresponds to the

H269 Hδ1 proton that is hydrogen bonded to D239 in the hMGL S122-H269-D239 catalytic triad

(Fig. 2A).

This chemical shift of catalytic histidine resonances in serine proteases has been shown to be highly dependent upon histidine protonation state and enzyme conformation [75]. Seemingly minor changes in pH can readily change His charge state and, consequently, the chemical shift of

Hδ1 proton. In serine proteases, a doubly protonated (positively charged) active site His imidazole ring demonstrates Hδ1 resonances usually shifted downfield to approximately 17-19 ppm [75]. To investigate this phenomenon in hMGL and support the H269 Hδ1 assignment, we attempted to titrate hMGL samples to a more acidic pH. Since this approach caused considerable protein precipitation, we alternatively utilized a sol-hMGL variant in which the catalytic S122 is substituted with a cysteine residue [68]. The proton NMR spectrum of the sol-hMGL-S122C mutant contained a new resonance at 18 ppm, and the signal at 14.9 ppm was absent (Fig. 1C).

This diagnostic downfield shift provides strong evidence supporting the protonation of H269 in the S122C mutant and formation of an imidazole-thiolate ion pair between the H269 and C122 in the modified hMGL active site [76]. Gradual titration to a higher pH elicited a decrease in the peak intensity at 18 ppm, and this resonance disappeared completely at pH 10.6 (data not shown).

This observation reflects deprotonation of H269 in the basic environment. Titration back to a neutral pH restored the peak, demonstrating the reversibility of H269 protonation. Moreover, the

32 substitution of H269 to alanine in the S122C mutant results in the disappearance of peak at 18 ppm

(Fig 3A).

FIGURE 2.2. Crystal structures showing (A) Hydrogen bonded H269 in the catalytic triad (S122- H269-D239) of hMGL (3HJU), (B) H49 involvement in the hydrogen bonding network within the oxyanion hole (G50, A51, M123) (3HJU), (C) The H54 residue in the open form (3HJU) and the closed form (3PE6). In contrast to these X-ray structures, mutagenesis with NMR clearly demonstrated that H54 is hydrogen bonded to D197 in the open form and is not bonded in the closed form. (D) The H103-H75 residues located at the interface between strand β4 and helix 2 forming a bridge between these secondary structures (3PE6).

The NMR spectra of the sol-hMGL H269A and S122C variants thus provide multiple lines of evidence allowing unambiguous assignment of the enzyme’s catalytic His269 Hδ1 proton at 14.9 ppm when His269 is neutral, and at 18 ppm when His269 is protonated. Furthermore, we performed enzymatic assays on all hMGL mutants used for assignment, and the H269A mutation was alone in completely abolishing activity, consistent with our assignment (Table 1).

33 The resonance at 12.8 ppm was assigned to the His49 Hδ1 proton based on the spectrum obtained from the H49A sol-hMGL variant (Fig. 1A). This histidine residue is located in the loop connecting

α1 and β3 (Fig. 2B). The loop includes the A51 residue, which is directly involved in the formation of the oxyanion hole together with the backbone amide group of Met123 [58]. Consequently, H49 is involved in a hydrogen-bonding network to the oxyanion hole.

FIGUE 2.3. (A) Comparison of downfield 1H NMR resonances of the soluble S122C and S122C/H269A mutant (pH7.4, T=275K). Effect of substitution of active site His269 for alanine. (B) Comparison of 1H NMR downfield resonances of sol-hMGL with resonances of soluble H103A and H75A mutants (pH7.4, T=310K).

Substitution of H54 with alanine caused the NMR signal at 13.9 ppm to disappear, providing definitive assignment of this resonance (Fig. 1A). H54 is located in the loop connecting α1 and β3

34 (Fig. 2C). The observation of the H54 resonance in this downfield region suggests that in hMGL’s active state, there is a hydrogen bond between H54 and D197 located in the enzyme’s lid sub- domain (Fig. 2C). The H54A mutation also results in disappearance of the H269 Hδ1 resonance at

14.9 ppm and a dramatic loss in hMGL catalytic efficiency by four orders of magnitude (Table 1).

These results from the H54A variant indicate that this hydrogen bond is important to enzyme function. In marked contrast, the H49A mutation preserves the H269A resonance, and there is a relatively modest five-fold reduction in catalytic efficiency in that sol-hMGL variant (Table 1).

The most deshielded signal at 15.9 ppm (Fig. 1A) represents a buried hydrogen-bonded pair H103-H75 and likely belongs to the H103 Hδ1 proton (Fig. 2D). The H103A mutation also preserves the downfield H269 resonance and elicits a twenty-fold reduction in catalytic efficiency

(Table 1). Additional evidence for the unambiguous assignment of the resonance at 15.9 ppm to the hydrogen-bonded pair H103-H75 is provided by the spectrum of the H75A mutant (Fig. 3B).

Mutation of either the donor or acceptor residue to Ala eliminates hydrogen bonding between them and leads to the absence of a downfield-shifted (15.9 ppm) resonance. The resonances assigned represent protons from amino acid residues that are either structurally essential or critical to hMGL catalysis. They can be used as unique probes to monitor functionally important events.

2.3.4 pH Effects on sol-hMGL NMR Spectral Features- NMR titration experiments of sol-hMGL in the 7.0-12 pH range reveal an exchange between two distinct conformations that is slow on the

NMR time scale (Fig. 4). As the pH is increased from 7.5, there is a gradual decrease in the peak intensity of H269 (14.9 ppm) until it completely disappears at pH 11. A new signal appears concomitantly at 12.90 ppm that is likely the H269 resonance. This significant upfield shift, in a slow-exchange manner, suggests a modulation of catalytic triad geometry (Fig. 4A).

35 Increasing pH also elicits a similar decrease in the intensity of the H54 (13.9 ppm) resonance due to its shifting well outside of the downfield region, likely as a result of the breaking of the hydrogen bond between H54 and D197 (Fig. 4A). A similar effect occurs in the spectra of the H54A mutant

(Fig. 1A). These data correlate with a significant decrease in the catalytic efficiency of the H54A variant, suggesting a critical role for this hydrogen bond in maintaining hMGL catalytic competency (Table 1).

FIGURE 2.4. Effect of pH on the downfield (A) and upfield (B) side chain resonances of hMGL (T=310K). The red arrows indicate the change of H269 chemical shifts with increasing pH. Relative populations of the open and closed hMGL conformations were calculated from the areas of H54 (C) and H269 (D) resonances.

The resonance from H103 (15.99 ppm) exhibits the formation of a second component shifted slightly downfield that gradually increases in intensity at the expense of the original peak. The two slightly separated peaks reach equal intensity at pH 9.6 until, at pH 11, there is only a single component observed at the slightly downfield chemical shift. A similar observation is made in the case of His49 (12.8 ppm) where the peak at 12.8 ppm gradually decreases in intensity and a new

36 peak appears at 12.4 ppm. Remarkably, the same slow exchange regime is observed in the upfield region of the spectrum for separated methyl group resonances such as the peak clearly observable at -0.52 ppm (Fig. 4B). Therefore, perturbations observed in both downfield and upfield chemical shift regions are likely indicative of one global process within the enzyme. All spectral changes were reversible with respect to pH (data not shown). These observations are consistent with a pH- dependent reversible equilibrium between two distinct hMGL conformations that are in slow exchange on the NMR time scale. The downfield signals were integrated and normalized to the intensity of the peak corresponding to the hydrogen-bonded pair H103-H75 and expressed as the percent-conformation as a fraction of their respective maximal intensities across the pH range studied (Fig. 4C, D). The data demonstrate that hMGL populates active and inactive conformations in solution with an equilibrium constant Keq~1 at pH 9.6.

2.3.5 hMGL Global Conformational Changes- 15N-HSQC and 13C-HSQC spectra of sol-hMGL in the active and inactive states were obtained. The active conformer of sol-hMGL undergoes a slow spontaneous transition to the inactive conformer after exposure to 37° C for long periods of time. This finding allows us to perform NMR experiments on the individual conformers at identical concentration, temperature and pH ensuring that spectral differences were reflective of conformational changes. Superimposed HSQC spectra (Fig. 5) for individual forms demonstrate significant chemical shift perturbations for backbone as well as side-chain resonances. This is a clear indication that hMGL structural alterations have a global impact upon conformational transition from active to inactive forms. Moreover, these spectra provide unambiguous experimental evidence that chemical shift perturbations observed in the downfield region are

37 related to the global conformational changes. Thus, hMGL downfield proton NMR resonances can be used as a sensitive probe for quantification of conformational equilibrium for this enzyme

FIGURE 2.5. (A) Superposition of the 2D 1H-15N HSQC spectra of 15N-labeled sol-hMGL (pH7.4, T=300K) in the active (red) and in the inactive (blue) conformations. (B) Superposition of the 2D 1H-13C HSQC spectra of 13C-labeled sol-hMGL in the active (red) and in the inatctive (blue) conformations (aliphatic region) and (C) aromatic region. (D) 1D 1H NMR downfield resonances of individual sol-hMGL conformers at pH7.4 and T=300K.

2.3.6 H272 Mutational and Temperature Effects on the sol-hMGL NMR Spectrum and

Enzymatic Activity- To explore this equilibrium further, we performed a mutagenesis study on the hMGL H272 residue that forms an aromatic cluster along with Y58 and R57 (Fig. 6). Substitution of H272 with tyrosine resulted in little change in the downfield resonances, suggesting that there is minimal perturbation to the interactions in the aromatic cluster (Fig. 7A). In contrast, the H272A

38 mutation elicited a decrease in the intensity of the H269 and H54 resonances (Fig. 7A), and the resultant spectrum closely resembles spectra of the sol-hMGL enzyme at higher pH values (Fig.

4A, pH=9.7). Likewise, the H272S mutation resulted in greater reduction of the H269 and H54 resonance intensities (Fig. 7A), and the spectrum is comparable to that of sol-hMGL obtained at pH=11. We characterized the activities of these mutants toward the native hMGL substrate, 2-AG.

FIGURE 2.6. (A) Open hMGL conformation stabilized by a cation-π interaction between R57 (arginine switch) and H272 (3HJU) and (B) the closed conformation with R57 flipped away from H272 (3PE6).

39 The H272Y and H272A mutants evidenced a twelve-fold reduction of sol-hMGL catalytic efficiency, while the H272S mutation caused a sixty-fold reduction in the enzyme’s catalytic efficiency (Table 1). These effects correlate with the retention of the H269 and H54 resonances in the downfield region of H272Y and H272A spectra and the loss of these resonances from the downfield region in the H272S spectra.

FIGURE 2.7. (A) Effect of H272 mutations on the population of conformers (pH7.4, T=310K),

(B) Effect of temperature on the open-closed equilibrium for the H272S mutant (pH7.4).

40 Substitution of H272 with tyrosine resulted in little change in the downfield resonances, suggesting that there is minimal perturbation to the interactions in the aromatic cluster (Fig. 7A). In contrast, the H272A mutation elicited a decrease in the intensity of the H269 and H54 resonances (Fig. 7A), and the resultant spectrum closely resembles spectra of the sol-hMGL enzyme at higher pH values

(Fig. 4A, pH=9.7). Likewise, the H272S mutation resulted in greater reduction of the H269 and

H54 resonance intensities (Fig. 7A), and the spectrum is comparable to that of sol-hMGL obtained at pH=11. We characterized the activities of these mutants toward the native hMGL substrate, 2-

AG. The H272Y and H272A mutants evidenced a twelve-fold reduction of sol-hMGL catalytic efficiency, while the H272S mutation caused a sixty-fold reduction in the enzyme’s catalytic efficiency (Table 1). These effects correlate with the retention of the H269 and H54 resonances in the downfield region of H272Y and H272A spectra and the loss of these resonances from the downfield region in the H272S spectra.

These observations are consistent with a shift in the conformational equilibrium of the sol- hMGL-H272S mutant toward an inactive form. Thus, the downfield pattern of resonances is indicative of the conformational equilibrium and resulting hydrogen-bonding pattern. The catalytic efficiency of these mutants provides confirmation that the inactive conformation is characterized by the absence of the H269 resonance at 14.9 ppm and H54 resonance at 13.9 ppm. Fig. 7B shows the response of the sol-hMGL-H272S mutant to temperature. As the temperature is decreased, the intensity of the H269 and H54 resonances increases reversibly. This is additional evidence supporting the proposition that these resonances are sensitive to conformational changes in hMGL, with reduced temperatures favoring the active conformation. For the H272Y variant the major sol- hMGL population is in the active conformation, whereas for the H272A variant the major

41 population is in an inactive conformation. For the H272S mutant, the equilibrium is almost completely shifted towards the inactive conformation (Fig. 7A).

2.3.7 Active-site Paraoxon Covalent Binding- To test the accessibility of the hMGL active site to ligands, we performed binding experiments with paraoxon, a low-molecular-weight organophosphate compound that covalently binds to the serine residue of serine hydrolases (Fig.

8) [70].

FIGURE 2.8. (A) The effects of paraoxon on the downfield resonances of sol-hMGL and H272A (pH7.4, T=310K). (B) The effects of paraoxon on the downfield resonances of H54A.

42 The spectrum of sol-hMGL displays downfield resonances consistent with a major population of active enzyme. Likewise, the spectrum of hMGL-H272A also contains these resonances although their intensities indicate a decrease in the population of active enzyme. For both of these hMGL variants, paraoxon elicits significant changes to the downfield NMR resonances that are immediately observable, demonstrating that this ligand had access to the active site (Fig. 8A). The spectra of the H54A display downfield resonances consistent with a major population of the inactive form of the enzyme. Paraoxon did not immediately alter the NMR spectrum of the H54A mutant, suggesting that the binding site is inaccessible to small-molecule ligands in the H54A mutant.

2.4 DISCUSSION

NMR spectroscopy has proven to be an important tool for probing the mechanism of action of the catalytic triad in serine proteases [75]. Often, active-site labile protons exchange with water protons too rapidly in solution to be observed by NMR. The catalytic triad of serine hydrolases, however, is typically buried within the protein, and thus shielded from direct solvent exposure, and the catalytic histidine protons are involved in hydrogen bonding. These factors slow the exchange rate of labile histidine protons in serine hydrolases, improving the opportunity for detecting discrete downfield-shifted NMR resonances [74, 75].

Previous NMR studies on serine proteases other than hMGL have focused mainly on histidine protons in the catalytic site [77, 78]. Our success in modifying hMGL to increase its solubility along with the use of appropriate solvent suppression schemes [71, 72] has allowed us to observe several hydrogen-bonded histidine protons outside the catalytic site that offer insights to overall enzyme conformational changes. The enzyme variants we engineered through defined

43 sol-hMGL point mutations allowed us not only to obtain definitive amino-acid resonance assignments, but also to define inter-residue hydrogen-bond interactions that correlate with changes in sol-hMGL catalytic efficiency.

The pH titration and temperature data show a similar pattern in that the more significant perturbations occur for the H269 and H54 resonances (Fig. 4 & 7). The inactive form of hMGL is thus characterized by a reduced number of peaks in the downfield NMR region. These perturbations likely reflect conformational changes affecting the geometry of the catalytic triad that predispose the enzyme toward an inactive form. Reversibility of the temperature and pH effects suggests a dynamic equilibrium between two distinct enzyme conformations: active and inactive. The downfield spectral effects correlate with chemical shift perturbations of backbone and side chain resonances seen in the HSQC spectra (Fig 5) that indicate a global conformational rearrangement. Different physical factors can therefore impact the same global rearrangement that can modulate the hydrogen-bond pattern of sol-hMGL, as reflected in the downfield region of the enzyme’s NMR spectra.

For instance, the H54A mutation impacts the H269 H 1 resonance at 14.9 ppm (Fig. 1) with a significant loss of catalytic efficiency (Table 1). The H54 residue of hMGL is located at a substantial distance from the catalytic triad [56, 58] such that this mutation could not have had a direct effect on H269. Among the H272 mutants, the H272S mutation in sol-hMGL most compromised the enzyme’s catalytic efficiency, consistent with the conformational transition to the inactive form reflected by the absence of the H269 and H54 resonances in the downfield region

(Fig. 7A). This suggests that H272 is a key residue that stabilizes the active conformation of hMGL through a series of aromatic interactions. The impressive overall congruence among the spectral

44 changes elicited by sol-hMGL point mutations, pH and temperature variations would indicate that the spectral changes observed are due to the same conformational transitions.

A lid sub-domain is one of the distinguishing structural features of many lipases. The lid acts as a regulatory domain that gates substrate accessibility to the active site, giving rise to the operational designations of “open” (i.e., substrate-accessible) and “closed” (i.e., substrate- inaccessible) lipase structural states [60, 79]. Both static X-ray studies [56-58] and our experimental data have identified an hMGL lid domain that can associate with the cell membrane in vivo and modulate access of endocannabinoid lipid-signaling substrates to the hydrophobic channel leading into the enzyme’s active site. In view of the functional role of the hMGL lid domain in controlling substrate entry to the enzyme’s catalytic core, it is tempting to speculate that at least some of the hydrogen-bond interactions we have defined, may be involved in conformational changes in the lid subdomain reflecting the lid’s open and closed states.

In this regard, there are two noteworthy structural features deduced from static X-ray data that implicate our data to the open and closed forms of the enzyme. There is a significant conformational rearrangement of the R57 side chain between open and closed states, which is in the vicinity of the aromatic cluster that includes H272 (Fig. 6) [57, 58]. In addition, the hydrogen bond between the H54-D197 residues occurs between the lid sub-domain and the core of the enzyme and this may also be impacted by the conformational rearrangement of R57 in this region of the enzyme (Fig. 2C) [57, 58].

This lead us to believe that formation or breaking of this hydrogen bond may be associated with a global conformational change responsible for the regulation of the equilibrium between open and closed states. Support of this hypothesis was provided by the paraoxon binding studies.

The H54A mutant showed no changes occurring to the downfield resonances, whereas similar

45 experiments demonstrated immediate changes to the sol-hMGL and H272A spectra (Fig. 8). We interpret this as the inability of the ligand to enter the binding pocket due to a drastic shift in the equilibrium of the H54A mutant towards a closed conformation.

Moreover, the changes that occur to the wild type and H272A mutant upon addition of paraoxon (Fig. 8) provide evidence that complex formation is proportional to the fraction of enzyme in the active conformation. It is possible therefore that the active conformation, as characterized by the presence of the H54 and H269 resonances, represents an open form of the enzyme that allows a ready access to the binding site.

Conversely, the absence of the H54 and H269 resonances as seen in the H54A, H272S, and high pH value spectra is associated with greatly reduced catalytic efficiencies (Fig. 4A) and may be indicating a shift in equilibrium towards the closed conformation. With this interpretation, our data suggests the existence of an H54-D197 hydrogen bond in the open state, as indicated by the presence of the H54 resonance, whereas it is broken in the closed state, as indicated by the absence of the H54 resonance. This differs from X-ray structures of the open form where, H54 is shown as the Nδ1-H tautomer and does not display this hydrogen bonding [56, 57], whereas, the structure of the closed form indicate H54 is in the more common Nε2–H tautomer and is hydrogen bonded to

D197 (Fig. 2C) [58]. This difference may be explained by common difficulties in deriving the exact conformational state of histidine side chains from deposited X-ray structures. The analysis of histidine side chain conformations and hydrogen bonding with proximal donors or acceptors can be obscure as there are three different protonation states of His and three rotameric states may be generated through flipping of the imidazol ring [80]. In addition, NMR results have previously differed from crystal structures in this regard, due to altering pKa values of histidine side chains in crystals compared to solution [81].

46 Closer examination of the crystal structures shows that in the open form, H272 interacts with Y58 via an aromatic-aromatic (π-π) interaction and simultaneously with the R57 cationic –

+ NH2 group via a cation-π interaction, thereby establishing a network of aromatic interactions involving three residues (Fig 6A). The geometry of this cluster of residues is optimal for π-π as well as cation-π and sigma-π interactions. Our results provide experimental evidence that this network may be stabilizing the open conformation of hMGL.

In the closed form, the guanidinium group of R57 is flipped toward the D197 residue from the lid sub-domain, pointing away from the H272 imidazole ring and disrupting the cation-π interaction (Fig. 6B). This group now forms a direct hydrogen bond with D197, and the NH1 of

R57 is an H-bond donor to Oδ2 of D197. Thus the R57 side chain rotates out of the active site and adopts the alternate position during the conformational switch from the open to the closed form of hMGL.

The importance of the cation-π interaction between the R57 and H272 side chains in stabilizing the active/open state is shown by the reduced catalytic efficiency of the H272Y, H272A and H272S mutants (Table 1). Conservative H272Y and nonconservative H272A substitutions demonstrate a

12-fold reduction, whereas the nonconservative mutation H272S demonstrates a 60-fold reduction in the catalytic efficiency of the enzyme. The drop in catalytic efficiency is mainly caused by a decrease in kcat values for all H272 mutants whereas the Km values are essentially unchanged.

Removal of one hydrogen bond between H54 Hε2 and Oδ2 of D197 in the lid sub-domain also results in a population shift toward the inactive/closed conformation (Fig. 1A) and significant reduction of kcat (Table 1). Clearly, the hMGL open conformation is highly pre-organized by aromatic interactions and hydrogen bonding. These interactions are destabilized by perturbations induced by point mutations. Thus the hMGL turnover is rate-limited by the equilibrium constant

47 of the pre-existing open-closed transition. Aromatic residues proximal to the active site, in addition to hydrogen bonds serve in the stabilization and/or regulation of the active site geometry.

In conclusion, our NMR study provides several lines of experimental evidence that hMGL exists in solution in a dynamic conformational equilibrium between active and inactive forms which is slow on the NMR time scale and can be modulated by pH, temperature, and specific point mutations. This study is further illustrative of a rare example of a conformationally flexible lipase about which NMR can correlate structure and function. Our results reveal a pre-existing communication route among amino-acid side chains mediated by hydrogen bonding and aromatic interactions and involving H54, R57, Y58 and H272 residues that are responsible for regulating hMGL activity. It is likely that these conformational transitions involve opening and closing of the lid-domain. The data presented invite further experiments to ascertain the direct contribution of lid gating dynamics to the transitions observed and the extent to which lid motion may be stabilized by substrate, product, and/or active site-directed inhibitor. Along with such information, the present study provides insight into the regulation of hMGL function useful to design novel chemical strategies for selective inhibition of this therapeutic target.

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58 Chapter 5:

Amino Acid Cluster Responsible for regulation of hMGL Conformational Equilibrium

Abstract:

The enzyme human monoacylglycerol lipase (hMGL), a serine hydrolase is responsible for the hydrolysis of the endocannabinoid endogenous ligand at cannabinoid receptors in brain. Most lipases are known to be dynamic; meaning they can co-exist in an ensemble of different conformations in solution with tunable populations [152]. In the present study, we characterize the contribution of residues His54, Arg57, Tyr58 and Asp197 in maintaining the conformational equilibrium of protein and effects thereby on enzymatic hydrolysis function. Our results identify a key polar interaction between His54-Asp197, where mutation of either residues resulted in a significant conformational shift towards the closed form accompanied by dramatic loss in catalytic efficiency. The residues Arg57 and Tyr58 form the supportive collaboration of cation-pi and pi-pi interactions may assist the lid gating mechanism. Chemical agents capable of destabilizing this network may provide for pioneering strategies to regulate endocannabinoid tone by selectively inhibiting hMGL.

5.1 INTRODUCTION

Lipases (EC 3.1.1.1) are a serine hydrolase family of enzymes that accelerate lipid hydrolysis to liberate free fatty acids [153]. Monoacylglycerol lipase (MGL) is a membrane-associated serine hydrolase that regulates the endocannabinoid system by metabolizing endogenous substrate 2- arachidonoyl glycerol (2-AG) to release arachidonic acid (AA) and glycerol. Human MGL

(hMGL) has the characteristic α/β hydrolase fold structure with a typical serine hydrolase familial

59 catalytic triad of Ser122-His269-Asp239 [57]. The construction of hMGL involves 8 β-sheets with the exception of β-sheet 2 running anti-parallel interconnected by 8 α helices [40]. Recent efforts at solving the three dimensional structure of hMGL resulted in one of the reversible ligand-bound structure (PDB ID: 3PE6) in closed (inactive) conformation in contrast to the free open (active) form (3HJU) [40, 57]. However, another effort at resolving the crystal structure of hMGL bound to a covalent inhibitor yielded an open conformation (PDB ID: 3JW8) [98]. The most striking structural difference in hMGL crystal structures in its apo and holo form is the drastic rearrangements within the helices α4, α5, α6 and the interconnecting loops together forming the cap-like ‘lid domain’ comprised of residues 151-225 [1, 39, 40]. The structural analysis of MGL substrate binding site showed that its active center is covered and protected by this lid domain, as seen in a majority of well-studied lipases [154]. The open conformation of hMGL is defined by an exposed, solvent accessible active site, whereas in closed conformation the lid (specifically helix

α4) hinders the solvent (or substrate) access to the active site. Even though the crystal structures of both open and closed conformations of hMGL are known; unfortunately, they provide a very static picture. Secondly, ligand occupancy triggered closed conformation in the crystal packing may not necessarily reflect the native (or biological) closed conformation. Therefore, there persists a need to thoroughly explore the mechanism of open-closed conformational equilibrium in lipases like hMGL. hMGL was considered to be exclusively membrane-bound, until hydrolysis activity was detected in the soluble cytosolic fraction as well [155]. This amphipathic nature of hMGL has been hypothesized to arise due to the hydrophobic and apolar residues in helix α4 that may assist in membrane anchoring. On the other hand, the environment surrounding the catalytic triad is much more polar than the channel pointing towards the enzyme surface [39]. The side chains of Arg57,

60 Tyr58 along with backbones of His121 and His272 delineate a hydrophilic cavity probably serving as an entryway to the polar head of 2-AG [39].

Our previous NMR-based study succeeded in isolating the open and closed states of hMGL in solution and identified one of the key residues, His-272 responsible for the conformational stability

[102]. We also confirmed that the flexible nature of hMGL enables global changes in the protein structure emerging from local dynamic changes [102]. Even though the structures of both open and closed hMGL are known, the meticulous mechanism of lid gating dynamics remains unknown.

The detailed structural comparison coupled with our previous site-directed mutagenesis experiments, we theorized a functionally crucial role for the residues Arg57, Tyr58, His54, Asp197 and His272 that sit at the center of a relay system that transmits lid domain movements to the active center. Our current study aims at identifying the mechanism of lid opening/closing by specifically examining individual contributions of these chosen residues on enzymatic activity and populations of the relative conformations.

5.2 Experimental Procedures:

5.2.1 Enzymatic Expression and Purification: Recombinant sol-hMGL and the mutants in this study were expressed in E-coli BL21 cells and purified employing the IMAC technology as described previously [102].

5.2.2 Enzyme Assays: MGL hydrolysis activity was measured chromatographically using the native substrate 2-AG [102]. The purified enzyme fractions were incubated for 20 min at 37oC with increasing concentration range of 2-AG (13μM to 400μM). The amount of product AA formed was quantified and the calculated rate of reaction was fitted in the Michaelis-Menten

61 equation to generate the enzymatic saturation curve. The kinetic parameters were calculated using

Graphpad Prizm 5.0 (San Diego, CA).

5.2.3 NMR Spectroscopy:All the spectra were acquired with Bruker AVANCE II 700-

MHz NMR spectrometer equipped with probe . All 1D 1H NMR experiments were performed using a Watergate pulse sequence (p3919fpgp) that enables water suppression, while the data processed using TopSpin3.2

5.3 RESULTS:

5.3.1 Mutagenesis Sites Selection: The identification of amino acids essential for hMGL catalysis or those that contribute to activity through conformational effects is a major question in the understanding of enzyme’s function. The lid domain of lipases has been reported to play an essential role in membrane association, interfacial adsorption and substrate selectivity [43, 156].

The substitution H54A was created formerly for the downfield NMR peak assignment; however the observed NMR pattern had suggested transformation to closed conformation. Therefore, lead to the serendipitous discovery of a crucial hydrogen bond identified by NMR spectroscopy between the side chains of the hMGL core residue His54 (loop between helix 1 and beta 3) and a lid-domain residue Asp197 (located in helix α5).

Previously we highlighted the role of aromatic residue His272 on the relative populations of open/closed hMGL conformations. The conservative mutation H272Y was catalytically efficient and existed primarily in open conformation as against the non-conservative mutations H272A and

H272S, highlighting the need of an aromatic residue at the position 272 [102]. Structural scrutiny of the available hMGL 3D structures revealed two obvious physical linkages of the aromatic imidazole side chain of His272- a cation-π bond with Arg57 side chain and secondly π-π

62 interaction with Tyr58 phenol ring. Also, major conformational rearrangements were observed for the Arg57 guanidium side chain in the apo and holo hMGL X-ray crystal structure.

Fig 1. (A) the three-dimensional structure of hMGL (PDB ID:3HJU) showing the residues involved in the catalytic triad, and those selected for mutagenesis in this study, (B) A network of hydrogen bonding network and aromatic interactions between the hub residues selected for mutagenesis that putatively control open/closed conformational equilibria.

In order to probe the effects of these hydrogen bonds and aromatic interactions on conformational equilibria and enzymatic function, single point mutations were created to yield constructs H54A,

R57A, R57K, Y58A, Y58F and D197A. Whether or not the amino acid residue is structurally or functionally critical was determined by comparing their kinetic parameters and downfield NMR spectra with the native protein.

5.3.2Hydrogen-bonded Histidine Resonances as Natural Probes for NMR Detection of hMGL Conformational Transition- The downfield region of the 1D NMR spectra (11-16ppm) of hMGL displays five distinct, well-separated and characteristic peaks. In our previous study, we definitively assigned four of these downfield peaks to hydrogen-bonded histidine residues based on a site-directed mutagenesis study that substituted histidines for alanine [102]. The downfield

63 resonances at 15.9, 14.9, 13.9 and 12.8ppm in sol hMGL are therefore ascribed to residues His103,

His269, His54 and His49 respectively; whereas the peak at 11 ppm belongs to a till-date unassigned –OH hydroxyl proton. Moreover, we published how we developed an NMR-based method that exploits these histidine peaks as sensitive probes to study the conformational equilibrium and binding interactions [102]. Based on our previous studies, the closed conformation is characterized by the disappearance of two middle peaks (14.9ppm and 13.9ppm) in the downfield region of 1D NMR spectra of sol-hMGL and mutants. The study also illustrated the effects of temperature on the relative populations of open/closed conformation. The non conservative mutation of His272 to serine led to an equilibrium shift towards closed conformation at 37oC, however lower temperatures exhibited reappearance of His-269 and His-54 peaks in the proton NMR downfield region. At lower temperatures, the unordered protein regions (loops) demonstrate altered dynamic properties, thereby stabilizing the open conformation in hMGL. The temperature-dependent NMR study is hence engaged to evaluate the reversibility of the conformational changes in protein caused by single point mutations.

Table 1. Comparison of kinetic parameters of sol hMGL and the mutant’s enzymatic activity on the hydrolysis of the native substrate 2-AG at 37OC

Vmax Catalytic Fold -1 Protein Km (M) kcat (sec ) (M/sec) Efficiency Change

sol-hMGL 22  4 0.55 (5.70.2) x 105 2.5 x 104

1.6 fold Arg57Ala 17.13  6 0.67 (6.9  0.1) x 105 4 x 104 increase

64 (1.3  0.06) x 0.8 Arg57Lys 65.09  9.9 1.25 2 x 104 106 Decrease

1.7 fold Tyr58Ala 18.06  6.2 0.78 (7.9  0.1) x 105 4.4 x 104 increase

(1.7  0.06) x 1.4 fold Tyr58Phe 45.5  6.01 1.31 3.6 x 104 106 increase

His54Ala 12  2.42 1.1 x 10-3 11 0.5 9.8 x 10-1 2.5 x 104

(2.55  0.38) x Asp197Ala 46.45  10 8.7 x 10-3 5.5 4.5 x 103 102

5.3.3 Residues His 54-Asp197 as Conformational Determinant: Role of salt bridges at the interface of the core and lid domain has been extensively reviewed and confirmed to play a crucial role in catalysis [157]. Salt bridges in proteins are experienced when two oppositely charged residues come sufficiently close to each other to form electrostatic interactions [158]. Accordingly, a critical salt bridge between His54-Asp197 was identified in hMGL structure, and disrupted by the mutation of either hydrogen bond donor-H54A or hydrogen bond acceptor-D197A. Both mutations had a mild influence upon the enzymatic affinity (Km) for the native substrate 2-AG, and were analytically found to be 12µM for H54A and 46µM for D197A (Table 1). However, the catalytic efficiency dropped substantially, with 25000- and 4500- fold drop recorded for His54 and

Asp197 substitution respectively, confirming a definitive role of this pair-wise interaction on enzymatic hydrolysis function (table 1 and Fig 2)

65 Figure 2. Substrate saturation curves for sol hMGL and the mutated enzymes H54A and D197A in phosphate buffer (pH 7.4 at 37oC, 20min substrate-enzyme incubation).

The downfield region of the 1D NMR spectra for both H54A as well as D197A substitution resulted in the disappearance of the peak at 13.9ppm as expected (Fig 3). The Nε2 of His-54 imidazole is hydrogen bonded to the carboxylate side-chain of aspartate (Fig 1B), and therefore the protonated imidazole NH is observed in the extreme downfield section of 1D NMR spectra at

13.9ppm. The assignment is confirmed by the absence of the peak by the mutation of either the hydrogen bond donor or the acceptor i.e His54 or Asp197 to alanine. An interesting finding was the disappearance of an additional downfield peak (14.9ppm) belonging to the catalytic triad residue His269, otherwise preserved and observed in native hMGL NMR spectra (Fig 3B and 3C).

The NMR spectral pattern with the dissolution of His54 and His269 peaks is suggestive of transformation towards the closed conformation. The closed conformation previously observed for the non-conservative H272S mutation was temperature dependent, with lower temperatures stabilizing and shifting the conformational equilibrium towards the open form [102]. A similar temperature dependent study of H54A and D197A replacements found absence of His-269 and

His54 peak reappearance at 295K as well as 285K, indicating the irreversible nature of the conformational changeover (Fig 3B and 3C).

66 Figure 3. Downfield region of the 1D proton NMR spectra for (A) sol hMGL, and the mutants (B) H54A, (C) D197A to confirm their conformational state with temperature dependence (pH 7.4)

Thus, amino acids His54 and Asp197 are critical for the open/closed conformational equilibrium and also seriously impair the hMGL hydrolysis rate, suggesting an imperative catalytic role for the structural changes. Therefore, we have identified a crucial pairwise interaction at the interface of enzyme body and lid domain that impacts the enzymatic hydrolysis properties by perturbing structural dynamics.

5.3.4 Arg57 and Tyr58 are involved in Lid regulation: The residues Arg57 and Tyr58 are located in the first turn of helix 1, and form an aromatic cluster with conformationally determinant residue His272. The aromatic substitution of His272 by tyrosine caused minimal perturbation to this interaction of aromatic clusters; in contrast substitution with serine resulted in an equilibrium shift towards the inactive conformation along with a 60-fold drop in the enzymatic efficiency

[102]. To explore the interactions within this aromatic cluster, both conservative (R57K, Y58F) and non-conservative (R57A, Y58A) substitutions were created in sol hMGL so as to specifically analyze the effects of cation-π and π-π interactions with His272 respectively. After these substitutions were individually introduced in the sol-hMGl construct, the kinetic properties of the consequential mutated enzymes were assessed chromatographically at 370C for the hydrolysis of endogenous cannabinoid ligand 2-AG.

67 Surprisingly the kinetic parameters as well as the downfield NMR spectra for all four substitutions were unusually novel. The replacements R57K, R57A, Y58A, Y58F did not have a major effect upon the enzymatic affinity (Km) towards the endogenous substrate 2-AG (Table 1). Except for the Arg57 replacement with lysine, all other mutations gave rise to enhanced enzymatic efficiencies (Fig 4). The conservative but bulkier substitution of Arg57 gave marginally reduced catalytic efficiency for R57K, as against to the non-conservative substitution with alanine that resulted in 1.6-fold increase in activity. On the other hand, both substitutions of Tyr58, namely

Y58A and Y58F yielded 1.7 and 1.4-fold increase in catalytic efficiency. This suggests the residues do form a structurally vital domain (or motif) that thus indeed participates in the regulation of enzymatic function.

Figure 4. Plots of the relative changes in % catalytic efficiency for the mutated enzymes with respect to sol-hMGL

100

80

60

40

20

% Catalytic Efficiency Catalytic % 0

sol hMGL H54A D197A R57A R57K Y58A Y58F

The homologous and heterologous substitutions of Arg57 and Tyr58 resulted in a unique 1H NMR downfield pattern as compared to sol hMGL. The peaks at 15.9, 14.9 and 13.8ppm remained fairly unaffected in R57A, Y58A and Y58F, with comparable chemical shifts and intensities observed in parent hMGL construct (Fig 5). The observation of His269 and His54 peaks throughout the

68 studied temperature range meant the protein existed in predominantly open conformation upon these mutations (Fig 5) unlike those observed for H54A, D197A (Fig 3) and H272A. However, the mutation R57K that resulted in a decreased catalytic activity also had an unusual NMR downfield spectral pattern. This larger substitution likely resulted in the upfield shifting of the

His54 resonance, which may consequence in the perturbations of the conformational equilibrium.

In addition, slight rearrangements were observed in His49 and unassigned –OH resonances for all four replacement constructs. With the absence of 11.9ppm peak belonging to an unassigned –OH in Y58A and Y58F at 310K, we considered a possibility of the specified peak-assignment to the

Tyr58 side chain; but the reappearance of additional peak (slightly downfield) at lower temperatures proved otherwise (Fig 4). The proton NMR downfield region from 11.5ppm to

13ppm for mutations R57A, R57K, Y58A and Y58F was ambiguous, and the complexity posed deterred us from making definite conclusion about the effects of mutation on the specific hydrogen-bonding pattern of His49 and unassigned -OH. To summarize, Arg57 and Tyr58 amino acids do not abolish hydrolysis function as originally thought, but do play a role in catalytic activity; as they may merely assist the residue hub identified previously that controls the hMGL conformational equilibrium in a complementary role.

5.3.5 Ligand Accessibility to hMGL active Site: hMGL structurally presents a buried active site, with the lid domain possibly adding an extra layer of complexity to the ligand binding phenomena [159]. We earlier established that mutation of His54 to alanine resulted in binding site being inaccessible to small molecule, active-site directed MGL inhibitors like paraoxon (Ki ~

35M) [102]. To confirm if the population shifts of conformations affects active site accessibility for the other substitutions, we conducted NMR-based binding experiments of sol hMGL and the mutant library with AM10212, a selective reversible inhibitor. The ligand AM10212 is known to

69 occupy the binding site stabilized by hydrogen-bond interactions without any permanent covalently modification, and is also more potent than paraoxon (Ki ~10nM) [40] .

Figure 5. Downfield 1H NMR based temperature dependent study of hMGL mutations (A) R57A, (B) R57K, (C) Y58A and (D) Y58F at pH 7.4

The complex formation between sol hMGL and AM10212 resulted in perturbation in the downfield NMR region evidenced by the modified resonances of hydrogen-bonded His-269, His-

54, His-49 and the unassigned-OH (data not shown). The binding profile of the point substitutions

R57A, R57K, Y58A and Y58F was highly analogous to sol hMGL, instituting ligand accessibility to the active site. However, the mutants D197A presumed to be in a closed conformation demonstrated no effects on the downfield NMR spectrum upon ligand addition, confirming that the active site is physically restricted to the high affinity ligand AM10212.

70 5.4 DISCUSSION:

Lipases are ubiquitous enzymes for the hydrolysis of carboxyl-ester bond of lipophilic substrates such as tri-, di- and mono- acylglyecrols [160]. The three-dimensional structure of apo and holo hMGL along with several lipases have confirmed the presence of a cap-like domain (comprising one or more helixes and interconnecting loops) that affects lipase enzymatic mechanism [161].

The cap (or lid) domain is dynamic in nature and the structural rearrangements within result in the displacement from open to closed conformation upon a biological trigger such as substrate binding or membrane association [153]. hMGL X-ray crystal structures suggested a conformational heterogeneity in the enzyme structure, primarily contributed by the movement within the lid domain (residues 151-225) that hinder the catalytic site access. In addition to affording active site access, the structural rearrangements also may alter the surface properties of the enzymes.

Charged hydrogen bond interactions such as those between histidine and aspartic acid side chains

(termed salt bridges) have specific geometries and donate immensely to protein stability [162].

Analysis of the three dimensional structure demonstrated that His54 is hydrogen bonded to Asp197 in the closed structure with its imidazole ring accepting the most common Nε2 –H tautomer. Our

NMR studies however noticed otherwise, where a hydrogen bond between the side chains of His54 and Asp197 was intact in open conformation. The disappearance of His54 peak established by the breaking of this hydrogen bond demonstrates a conformational equilibrium shift towards closed conformation. The mutation of His54 by alanine cause a 2.5 x 104 –factor decrease in the enzymatic activity, whereas the mutation of Asp197 by alanine cause a 4.5 x103-fold decrease in the same.

These mutations not only cause a significant alteration in the hMGL lipid hydrolysis function, but also the overall structure of protein with respect to wild type hMGL. One may postulate that the hydrogen bond breaking between His54 and Asp197 side chains result in the lid domain losing its

71 functionally required flexibility. And thus the protein exists in a predominantly closed conformation in solution. The conformational shifts in lipases can be explained by the classical induced fit model, or a population shift model [163]. The key-and-lock model of enzymes is no longer widely accepted, and the induced fit theory has taken over suggesting the enzyme temporarily changes shape upon a biological trigger, usually to accommodate the substrate [164].

The population shift model however proposes a pre-existing equilibrium of a range of conformations at physiological conditions, where the substrate-binding stabilizes the lowest energy conformation on the energy landscape [165]. As per the induced fit theory, the enzyme cannot possibly exist in the closed conformation (irrespective of mutations) in the absence of substrate (or ligand) binding phenomena. The populations shift in conformational ensemble of hMGL by breaking the hydrogen bond between the side chains of His54 and Asp197 resembles a population shift-model with redistribution of pre-existing conformations. Therefore, based on our experimental results, we conclude that H54A and D197A induce a permanent (and irreversible) shift in the pre-existing hMGL equilibrium towards the inactive conformer. The NMR-based temperature dependent analysis additionally suggests restriction of the conformational ensemble to restrict not only the endogenous substrates but also high affinity ligands. Therefore, His54 and

Asp197 play an important catalytic and dynamic role in hMGL enzymatic structure and function.

This also opens up a new avenues about the functional importance of accessory domain/s mobility with respect to the substrate binding in lipases in general [165].

The initial hypothesis behind creating additional mutations for residues coupled with His272 was to see a comparable declining effect on the enzymatic hydrolysis function. The interactions of arginine side chains with aromatic groups in protein structure are well-known with preference for stacking between aromatic ring and the guanidinium group [166]. Apart from Arg57, the

72 substitutions of Tyr58 were created to destabilize the ring stacking with His272. Surprisingly though, the enzymatic assay results show that Arg57 and Tyr58 play only a minor role in catalysis.

In contrast, R57A, Y58A and Y58F all lead to higher than 1.5 fold increase in the enzymatic efficiency.

Usually, 50-70% single-point mutations in a protein can be neutral or very slightly deleterious, compared to 0.1-1% mutations that are found to be functionally advantageous [167].

Functionally beneficial mutations are evolutionarily intended for counter balancing (compensatory mechanisms) and providing additional stability and robustness to proteins [168]. These are the amino acids replacements that do not disrupt the substrate-binding, but may in turn provide additional stabilization to the protein binding site by lowering the binding energy. Arg57 and

Tyr58 residues, thus belong to this class of functionally beneficial mutations.

The downfield region of the 1D NMR spectra for the substitutions of the catalytic activity boosting residues Arg57 and Tyr58 showed rearrangements of the His49 and unassigned –OH peak. The residue His49 is situated in the loop connecting helix 3 to the beta sheet 3, and therefore physically in close vicinity of Arg57 and Tyr58 that are involved in the first turn of helix

3. These residues are also locally proximal to Ala51 which is directly involved in the formation of the oxyanion hole along with the backbone amide of group of Met123. The amino acids Arg57 and Tyr58 may not be directly involved in the regulation of conformational equilibrium as originally thought, but the NMR data cannot completely rule out the possibility of Arg57 and

Tyr58 residues providing additional stabilization to the oxyanion hole on the energy landscape.

The oxyanion hole by definition forms a pocket in hMGL enzymatic active site that stabilizes the anionic intermediate (negative charge on the deprotonated substrate oxygen) developed during the formation of transition state [39, 169].

73 On the other hand, the bulkier substitution of Arg57 with lysine shifted the His54 resonance upfield on the 1H NMR downfield spectra, suggesting an increased distance between the hydrogen bond forming residues, His54 and Asp197. Any perturbations to this crucial as well as conformationally sensitive hydrogen bond lead to altered hMGL conformational dynamics, due to the direct involvement of the lid domain.

Conclusion:

Structural flexibility plays an important role in enzymatic activity and conformational equilibrium through detraction of hinge motility. Replacement of His54, Asp197 resulted in an unfavorable conformational shift with impaired enzymatic activity. These mutations demonstrated the importance of residues His54 and Asp197 in maintaining catalytic function and ligand interactions.

The residues His54, Asp197 along with Arg57 and Tyr58 thus form a specialized microenvironment within hMGL structure that dramatically modifies the behavior of a key catalytic region, the lid domain.

74 References:

1. Smith, R.E., et al., Using NMR to Develop New Allosteric and Allo-Network Drugs. Current drug discovery technologies, 2015. 12(4): p. 193-204. 2. Jääskeläinen, S., et al., Conformational change in the activation of lipase: An analysis in terms of low‐frequency normal modes. Protein science, 1998. 7(6): p. 1359-1367. 3. Labar, G., et al., Crystal structure of the human monoacylglycerol lipase, a key actor in endocannabinoid signaling. Chembiochem, 2010. 11(2): p. 218-27. 4. Schalk‐Hihi, C., et al., Crystal structure of a soluble form of human monoglyceride lipase in complex with an inhibitor at 1.35 Å resolution. Protein Science, 2011. 20(4): p. 670- 683. 5. Bertrand, T., et al., Structural basis for human monoglyceride lipase inhibition. J Mol Biol, 2010. 396(3): p. 663-73. 6. Bertrand, T., et al. Structural basis for human monoglyceride lipase inhibition. J Mol Biol, 2010. 396, 663-73 DOI: 10.1016/j.jmb.2009.11.060. 7. Labar, G., et al., Crystal structure of the human monoacylglycerol lipase, a key actor in endocannabinoid signaling. Chembiochem, 2010. 11(2): p. 218-227. 8. Lan, D., G.M. Popowicz, and Q. Tang, Review: The Role of Residues 103, 104, and 278 in the Activity of SMG1 Lipase from Malassezia globosa: A Site-Directed Mutagenesis Study. Journal of microbiology and biotechnology, 2015. 25(11): p. 1827-1834. 9. Dinh, T., et al., Brain monoglyceride lipase participating in endocannabinoid inactivation. Proceedings of the National Academy of Sciences, 2002. 99(16): p. 10819- 10824. 10. Tyukhtenko, S., et al., Specific Inter-residue Interactions as Determinants of Human Monoacylglycerol Lipase Catalytic Competency: A ROLE FOR GLOBAL CONFORMATIONAL CHANGES. J Biol Chem, 2016. 291(6): p. 2556-65. 11. Verger, R., ‘Interfacial activation’of lipases: facts and artifacts. Trends in Biotechnology, 1997. 15(1): p. 32-38. 12. Nasr, M.L., et al., Membrane phospholipid bilayer as a determinant of monoacylglycerol lipase kinetic profile and conformational repertoire. Protein Science, 2013. 22(6): p. 774- 787. 13. Lindberg, D., S. Ahmad, and M. Widersten, Mutations in salt-bridging residues at the interface of the core and lid domains of epoxide hydrolase StEH1 affect regioselectivity, protein stability and hysteresis. Archives of biochemistry and biophysics, 2010. 495(2): p. 165-173. 14. Bosshard, H.R., D.N. Marti, and I. Jelesarov, Protein stabilization by salt bridges: concepts, experimental approaches and clarification of some misunderstandings. Journal of Molecular Recognition, 2004. 17(1): p. 1-16. 15. Kingsley, L.J. and M.A. Lill, Substrate tunnels in enzymes: structure–function relationships and computational methodology. Proteins: Structure, Function, and Bioinformatics, 2015. 83(4): p. 599-611. 16. Stauch, B., S.J. Fisher, and M. Cianci, Open and closed states of Candida antarctica lipase B: protonation and the mechanism of interfacial activation. Journal of lipid research, 2015. 56(12): p. 2348-2358. 17. Yang, Y. and M.E. Lowe, The open lid mediates pancreatic lipase function. Journal of lipid research, 2000. 41(1): p. 48-57.

75 18. Huyghues-Despointes, B.M. and R.L. Baldwin, Ion-pair and charged hydrogen-bond interactions between histidine and aspartate in a peptide helix. Biochemistry, 1997. 36(8): p. 1965-1970. 19. Yan, H. and X. Ji, Role of protein conformational dynamics in the catalysis by 6- hydroxymethyl-7, 8-dihydropterin pyrophosphokinase. Protein and peptide letters, 2011. 18(4): p. 328-335. 20. Suzuki, H., How Enzymes Work: From Structure to Function. 2015: CRC Press. 21. Swift, R.V. and J.A. McCammon, Substrate induced population shifts and stochastic gating in the PBCV-1 mRNA capping enzyme. Journal of the American Chemical Society, 2009. 131(14): p. 5126-5133. 22. Flocco, M.M. and S.L. Mowbray, Planar stacking interactions of arginine and aromatic side-chains in proteins. Journal of molecular biology, 1994. 235(2): p. 709-717. 23. Romero, P.A. and F.H. Arnold, Exploring protein fitness landscapes by directed evolution. Nature Reviews Molecular Cell Biology, 2009. 10(12): p. 866-876. 24. Suplatov, D., et al., Computational design of a ph stable enzyme: Understanding molecular mechanism of penicillin acylase's adaptation to alkaline conditions. PloS one, 2014. 9(6): p. e100643. 25. Berg, J.M., J.L. Tymoczko, and L. Stryer, Proteases: Facilitating a Difficult Reaction. 2002.

76 Chapter 6.

Characterization of hMGL interaction with small molecule inhibitors

6.1 INTRODUCTION:

Monoacylglycerol lipase (MGL) is an endocannabinoid enzyme belonging to the family of serine hydrolases. The levels of endogenous endocannabinoid 2-Arachidonoyl glycerol (2-AG) are closely regulated by enzymes responsible for its biosynthesis and degradation. Human MGL

(hMGL) modulates the endocannabinoid tone by hydrolyzing the endogenous ligand 2-AG and thereby terminating its action at cannabinoid receptors. MGL is predominantly found in the adipose tissue, as well as in brain, liver, testis and other organs [38]. In brain, 2-AG functions by binding to cannabinoid receptor 1 (CB1) and thereby decreasing the inhibitory or excitatory currents at glutamatergic and GABAnergic synapsis [170]. In certain pathophysiological conditions like pain, inflammation and cancer, inhibitors of hMGL are proven to be beneficial by potentiating 2-AG binding to CB1 activation [171].

The small molecule inhibitors of hMGL can be broadly classified into reversible and irreversible inhibitors. Reversible inhibitors form non-covalent bonds with MGL, and are either transition state analogues or competitively occupy the active site. Irreversible inhibitors covalently modify the catalytically crucial residues Ser122 or Cys242 and permanently block the enzymatic active center

[172]. Traditionally, MGL inhibitors have belonged to a huge variety of chemical classes, di-/tri- fluoroketones, carbamates, tetrazoles, isothiazolinone, fluorophosphonates [173-176]. Several groups have reported diverse chemical classes of novel hMGL inhibitors with high affinity, but most of them lack selectivity and/or druggability. Thus, there is a dire need to understand the

77 detailed mode of the ligand-protein interaction at molecular level. The ligand binding site of enzymes is highly specific, while the affinities of (endogenous) ligands tailored to their biological function. The current drug design techniques would definitely benefit from deeper understanding about the binding mode of ligand-protein.

The most common method for screening small molecule library these days is high through-put screening (HTS). This method however suffers from a number of draw backs. Low affinity binders with prospective of finding lead compound are often missed. Also, the method generates no evidence regarding the binding mode, such as reversibility, interface with proteomic amino acids and induced conformational changes. With sufficient ligand-protein dissociation rates, binding of ligands with low (milimolar) to intermediate (micromolar) affinities are detectable by NMR-based methods [177]. Secondly, the NMR-based screening method can also give detailed information about the binding mode, for e.g. binding pocket, reversibility with the least possibility of false- hits.

The downfield region of the 1D NMR spectra of hMGL is characterized by five unique peaks, four of which have been definitively assigned to protons of histidine side-chains based on site directed mutagenesis studies [102]. These four peaks belong to distinct parts of the protein, for example His49 (12.8ppm) represents the oxanion hole, whereas His269 peak (14.9ppm) reflects events at catalytic triad. His54 (13.9ppm) residue is involved in lid regulation, while His103 is fairly inert to binding and conformational phenomena. Our approach utilizes these peaks peaks as probes to investigate the binding effects in their respective protein regions.

78 6.2 EXPERIMENTAL PROCEDURES:

6.2.1 NMR Spectroscopy: Purified samples of sol-hMGL and its mutants were obtained by procedure detailed in the previous sections and publications [102]. The final volume of the purified protein sample was ~500uL. The solid compounds were diluted in DMSO-d6 to a final concentration of 50mM. The compound dilutions were strictly done in DMSO-d6 to avoid a hydrogen signal during NMR spectroscopy. Compound was added directly to NMR tube and mixed with the purified protein sample in ratio 1:2 or 1:4.

6.2.2. hMGL Fluorescent Enzyme Assay: The fluorescent enzyme assays were performed with the crude cell lysate overexpressing the protein of interest. The EC50 was determined using a fluorescent analogue of the endogenous substrate, N-arachidonoyl, 7-hydroxy-6-methoxy-4- methyl coumarin ester (AHMMCE) [172]. The intrinsic high sensitivity of our fluoregenic substrate allows for high-throughput screening method to determine the potency of tested MGL inhibitors. In brief, 2ng of hMGL was incubated with various concentrations of the inhibitor and equilibrated for 15 min followed by the addition of AHMMCE at final concentration of 20 µM.

The fluorescent reading at 360 nm / 460 nm (λexcitation/ λemmission) was taken after 3 hours using

EnVision Multilabel Reader (PerkinElmer, MA). The relative fluorescence units were transformed to concentration of product (coumarin fluorophore) from the standard curve of coumarin fluorescence [178].

6.3 RESULTS AND DISCUSSION:

6.3.1 hMGL Interactions with Transition State Analogues (TSA): Ligands mimicking the geometric and electrostatic features of the transition state (or other intermediates of high energy) are considered as excellent enzyme inhibitors [179]. It has been proven theoretically that enzymes

79 bind transition state up to 108 times tighter than substrate [180]. Transition state analogues are ligands that mimic the tetrahedral intermediate state of a substrate molecule in an enzyme- catalyzed reaction [180]. The TSA-protein complex results in hemiketal adduct formation with catalytic Ser122, which is structurally alike but more stable than the transient tetrahedral intermediate on the energy landscape with the endogenous substrate. This complex formation is supposedly accompanied by the loss of the hydroxyl proton of Ser122 to neighboring His269 imidazole moiety.

Figure1. Schematics of interaction between hMGL catalytic triad residues and small molecule transition state analogues.

To study the interactions between TSAs and hMGL, we selected four ligands of varying potencies

AM5206, AM6639, AM6642 and 2-benzothienyl boronic acid. These ligands have been confirmed to act as TSAs in a closely related serine hydrolase, fatty acid amide hydrolase (FAAH) [181, 182].

The peak at 15.9ppm in 1D NMR spectra of hMGL was earlier assigned to His103, the residue located on alpha helix 2 that plays no significant role in enzymatic hydrolysis function [102].

Specific binding between these ligands and sol-hMGL using the fuoregenic assay had little or no effect on the His103 resonance, confirming it is not directly involved in the binding process (data not shown). The peak at 15.9ppm in 1D NMR spectra of the variant H103A was predictably absent,

80 and therefore the farthest downfield resonance was that of the catalytic His269 at 14.9ppm. Thus, the substitution H103A provided an ideal opportunity to study the hMGL-TSA interactions as any shifting of protonated/deprotonated His269 peak did not obstruct with the remaining resonances in the downfield section of NMR spectra.

Table 1. Inhibition parameters determined for each compound against sol-hMGL using the high- throughput fluorogenic substrate.

Compound EC50 (µM) sol hMGL AM5206 ~100 AM6639 56 ± 15 2-Benzothienyl boronic Acid 46 ± 14 AM6642 0.09 ± 0.03

The weakest inhibitor AM5206, a trifluoromethyl ketone is an analogue of 2-AG, and is expected to mimic the binding mechanism of 2-AG with catalytic Ser122. A definite decrease in the intensity of His269 peak was observed when compound AM5206 was incubated with purified protein in

1:2 ratios (Fig 2). This was concurrent with two additional peaks observed at 16ppm and 18.5ppm.

The novel peaks can be explained by a different protonation state of His269 imidazole moiety acting as a hydrogen-accepting base. The resonance at 16ppm can be attributed to the michaelis complex, while the extreme downfield peak at 18.5ppm can arise from a fully protonated imidazole ring. AM5206 mimics the electrostatic and geometric features of the first tetrahedral transition state, leading the catalytic Ser122 to form a hemiketal adduct with the compound. As a result

Ser122 loses its hydroxyl proton to His269 imidazole ring. His269-Nε upon accepting the proton form Ser122 will be fully protonated and explains why the peak at 14.9ppm moves farthest downfield. Even though the compound is in excess, the left-over peak at 14.9ppm represents the unbound free protein because of low affinity of AM5206.

81 Boronic acid derivatives have historically been used as classic transition state analogues of serine hydrolases [30]. The ligand binding experiment with 2-benzothienyl boronic acid yielded two novel peaks in the 1D NMR spectra, at 15.6 and 16.4ppm, with complete elimination of unbound

His269 peak at 14.9ppm. It is safe to assume that two new resonances represent the michaelis complex and protonated His269 respectively. Also, a 2-fold increase in AM5206 affinity resulted in no remaining unbound protein leading to the absence of free His269 resonance.

Fig 2. Effect of TSA binding on the downfield region of 1D 1H NMR spectra of H103A sol-hMGL variant at T=310K (pH 7.4). The final protein: ligand ratio is 1:2.

The other two compounds AM6636 and AM6642 are enantiomeric difluoroketo compounds. The steric requirements of the active site are such that AM6636 has a potency of 100uM, while that of

AM6642 is 90nM. The ligand binding study with AM6636 resulted in a reduced intensity of

82 His269 resonance, while that with AM6642 showed two new resonances at 15.3 and 18ppm. As explained for the previous TSAs, the peak at 15.3ppm in the 1D NMR spectra is attributed to the michaelis complex, and the peak at 18ppm to the stabilized tetrahedral intermediate.

6.3.2 Interactions with Irreversible Covalent Compounds:

The initial efforts of designing MGL inhibitors were dedicated to the mechanism mimicking the

2-AG hydrolysis pathway (Fig 3A). Organophosphates with tetrahedral and negatively charged phosphonate group were designed as potent inhibitors of serine hydrolases, in general [183].

Carbamates covalently react with the nucleophillic serine in most serine hydrolases, and form a stable and covalent carbamate adducts so as to inactivate the enzyme (Fig 3b) [184]. Unlike that natural substrate that form transient intermediates, this carbamate adduct cannot be hydrolyzed to regenerate free enzyme and are thus classified as irreversible.

The proposed mechanism of a complex formation between carbamates such as AM6580 and catalytic Ser122 is seen in Fig 3b. The hydroxyl proton of Ser122 is hydrogen bonded with His269 imidazole Nε, and is donated to the tetrazole leaving group. The resulting carbamate adduct is thought to be highly stable.

Fig 4 shows a covalent complex between the purified protein sol hMGL and excess ligand in 1:2 ratios-either paraoxon at 310K. Paraoxon is an organophosphate oxon compound which is proven to inactivate the serine hydrolases enzymes by phosphorylation. The formation of the complex is easily identified by the formation of the leaving group para-nitrophenol which develops a yellow color in NMR tube. The 1D NMR spectra of the complex showed absence of a peak at 14.9ppm, which we know belongs to the catalytic His269. The additional peak at 13.6ppm can be explained by the upfield movement of His269 resonance. The upfield shift indicates deprotonation of the

83 His269 imidazole ring, which will occur if the proton shared by Ser122-His269 pair is donated to the leaving group.

Figure 3. Reaction mechanism between catalytic Ser122 and endogenous ligand (a) 2-AG and (b) AM 6580 [1]

AM6580 is a carbamate compound that also results in the rearrangement in the downfield area of the 1D NMR spectra of 1:2 complexes between sol hMGL and AM6580. The peak at 14.9ppm is absent pointing towards an altered protonation state of the His269 imidazole ring. The novel shoulder peak of His54 resonance at 13.7ppm must be a result of deprotonated His269 imidazole ring displaying the upfield movement. The carbamate adduct of Ser122 was thought to be extremely stable, however our studies indicate otherwise. Upon longer incubations at 310K, the

13.8ppm peak intensity decreased with the reappearance of a resonance at 14.9ppm. This continued until after 3 days the 1D NMR spectra for the complex resembled that of free enzyme.

84 These results suggest that AM6580 was slowly hydrolyzed by sol hMGL and restored the active enzyme.

Figure 4. Effect of covalent inhibitors binding on the downfield region of 1D 1H NMR spectra of sol-hMGL variant at T=310K (pH 7.4). The final protein: ligand ratio is 1:2.

H103 H54 H49 OH H269

sol hMGL

Sol‐ hMGL + Paraxon

Sol‐ hMGL + AM 6580

6.3.3 hMGL Interactions with Product Analogues: The second class of reversible inhibitors is piperazine-pyrimidine compounds which we named as product analogues. These compounds show no covalent attachment to any of the active site residues, but occupy the active site resembling the second step of catalysis of product formation. AM10212 with 7nM EC50 for purified sol hMGL is the prototype of this category. We synthesized two analogues of AM10212.

We introduced a bulky group on the piperazine-pyrimidine side which resulted in nearly a total loss of activity. The second analogue AM10229

85 Figure 5. Piperazine-pyridine based reversible MGL inhibitors classified as product analogues.

AM 10212 AM 10216 IC50: 7nM IC50:>100uM

AM 10229

IC50: 11uM attempted removal of the solvent exposed cyclohexane ring retaining only the benzoxazole moiety.

Both the derivations resulted in a much weaker binding affinities for AM 10229 and AM10216.

The 1D 1H NMR spectra showed substantial modifications in the His269 residue, as the peak at

14.9 ppm was completely absent. The downfield profile was surprisingly similar to that of covalent serine-modifying agents; however, their replacement by a stronger covalent binder confirmed their reversible nature.

6.3.4 NMR-based screening method: The NMR-based screening method can definitely distinguish reversibility about small molecule inhibitors. One such example is shown in Fig 20.

Upon incubation of sol hMGL with 1:2 excess ligand 2-benothienyl boronic acid, the 1D NMR spectra revealed a previously identified typical TSA profile. Further time dependent study indicated that the 1D NMR spectra of a complex with this ligand remained unchanged (Fig 6.8).

86 Fig 6. Close up of hMGL crystal structure in complex with AM 10212

87 Figure 7. Effect of reversible inhibitors binding on the downfield region of 1D 1H NMR spectra of sol-hMGL variant at T=310K (pH 7.4). The final protein: ligand ratio is 1:2.

A similar experiment was repeated with the second ligand paraoxon. Paraoxon is known to covalently and irreversibly inactivate hMGL like all other serine hydrolases. An increase in the intensity of a peak at 13.6ppm was gradual and proportional to incubation times. We have already proven that the appearance of a peak at 13.6ppm specifies complex formation. The time dependent increase in the complex formation observed by the 1D NMR spectra remained the same up to 3 days (no further data accumulated). As no free enzyme could be regenerated, the mode of inhibition can definitely be classified as an irreversible inhibition. The method can be employed to any unknown ligand mimicking such behavior, which can be indisputably categorized as an irreversible inhibitor.

88 Figure 8. Effect of reversible and irreversible inhibitors binding on the downfield region of 1D 1H NMR spectra of sol-hMGL variant at T=310 K (pH 7.4). The final protein: ligand ratio is 1:2.

Conclusion:

Interaction of hMGL with the small molecule inhibitors results in perturbations in the protein NMR spectra. The detection of these perturbations forms the basis for our ligand binding analysis by

NMR. The 1D 1H NMR spectra of sol hMGL is complex with severe overlapping in the amide and aliphatic region. However, the presence of four unique and well separated downfield spectra provides probes that attain site-specific information about binding effects on sol hMGL.

89 References:

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