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1 BIOCHEMICAL AND STRUCTURAL CHARACTERIZATION OF Lactobacillus johnsonii FERULOYL ESTERASE S By KIN KWAN LAI A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENT S FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2011
2 2011 Kin Kwan Lai
3 To my parents, Hon Yuen Lai and Lai Yin Kong and to my brothers, Ping Kwan Lai and King Kwan Lai, for their unlimited love and support
4 ACKNOWLEDGME NTS I express my highest gratitude to my primary advisor, Dr. Claudio Gonzalez, for his unwavering guidance throughout my entire graduate school experience His support and constant push for improvement has made my experience as a graduate student successf ul and more rewarding. I also thank Dr. Graciela Lorca for her indispensable insight as well as my other committee members Dr. Julie Maupin Furlow, Dr. Joseph Larkin III, Dr. Nicole Horenstein, and Dr. Veroni ka Butterweck for their advice and the faculty o f the Microbiology and Cell Science Department for their support. I would like to express my appreciation for the help and support provided by my fellow members of the Gonzalez and Lorca labs: graduate students Santosh Pande, Ricardo Valladares, Algevis Wr ench; undergraduate students Sara Molloy and Clara Vu; scientist Fernando Pagliai; and lab technician Beverly Driver. I would also like to thank the members of Banting and Best Department of Medical Research in the University of Toronto, especially Peter S togios and Xiaohui Xu for their invaluable contribution with the protein crystal structures. Finally, I would like to thank my family and close friends especially Anastasia Potts for their kind encouragement that helped motivate me throughout my graduate school career.
5 TABLE OF CONTENTS page ACKNOWLEDGMENTS ................................ ................................ ................................ .. 4 LIST OF TABLES ................................ ................................ ................................ ............ 8 LIST OF FIGURES ................................ ................................ ................................ ........ 10 LIST OF ABBREVIATIONS ................................ ................................ ........................... 12 ABSTRACT ................................ ................................ ................................ ................... 16 CHAPTER 1 INTRODUCTION ................................ ................................ ................................ .... 18 Phytophenols ................................ ................................ ................................ .......... 18 Health Beneficial Properties of Phenolic Acids ................................ ................. 20 Common Phytophenols Present in Human Diets ................................ ............. 22 Limitation on Phenolic Acid Absorption ................................ ............................ 23 Microbial Interaction with Food Compon ents ................................ .................... 24 Esterases ................................ ................................ ................................ ................ 26 Ferulic Acid Esterases (FAEs) ................................ ................................ .......... 27 General Ch aracteristic of FAEs ................................ ................................ ........ 28 Reaction Mechanism of FAEs ................................ ................................ .......... 29 Structural Binding Mechanism of FAEs ................................ ............................ 31 Classification of FAEs ................................ ................................ ...................... 32 Applications of FAEs ................................ ................................ ........................ 34 Project Rationale and Design ................................ ................................ ................. 36 2 MATERIALS AND METHODS ................................ ................................ ................ 47 Chemicals, Media, and Strains ................................ ................................ ............... 47 Chemicals ................................ ................................ ................................ ......... 47 Growth Conditions of E. coli Strains ................................ ................................ 47 Preparation of Competent E. coli Cells ................................ ............................. 48 Isolation and Growth Condition of Lactobacillus strains ................................ ... 49 DNA Procedures ................................ ................................ ................................ ..... 49 Lactobacillus Strain identification ................................ ................................ ..... 49 In silico Selection of Potential FAE Encoding Genes ................................ ....... 49 Cloning of Potential FAEs ................................ ................................ ................. 50 Cloning of Human Valacylovir Hydrolase (VACVase) ................................ ...... 51 Generating LJ0536 Protein Variants ................................ ................................ 52 DNA Gel Electrophor esis ................................ ................................ .................. 53 Protein Procedures ................................ ................................ ................................ 53 Protein Purification ................................ ................................ ........................... 53
6 Sodium Dod ecyl Sulfate Polyacrylamide Gel Electrophoresis (SDS PAGE) .... 54 Protein Quantification ................................ ................................ ....................... 54 Enzyme Assays ................................ ................................ ................................ 55 Feruloyl esterase screening assay ................................ ............................. 55 Determination of optimal assay conditions ................................ ................. 55 Deter mination of enzymes substrate preference ................................ ........ 56 Determination of biochemical parameters by saturation kinetics ............... 57 Effect of bile sal t component and metal ions on enzyme activity ............... 58 LJ0536 mutants and VACVase ester screening assay .............................. 59 Detection of phenolic acids using high performance liquid chromatography (HPLC) ................................ ................................ ......... 60 Determination of native molecular weight using size exclusion chromatography ................................ ................................ ...................... 60 Analysis of protein secondary structure by circular dichroism .................... 61 X Ray Crystallization of LJ0536 and S106A ................................ ..................... 61 PDB Access ion Code of Proteins ................................ ................................ ..... 64 Structural Analysis ................................ ................................ ............................ 65 Sequence Analysis and Construction of Phylogenetic Trees ........................... 65 3 IDENTIFICATION OF FAES FROM GUT MICROBIOTA ................................ ....... 79 Background ................................ ................................ ................................ ............. 79 Result and Discussion ................................ ................................ ............................ 81 FAEs Producing Strain Isolation and Identification ................................ ........... 81 In Silico Selection of Targets for Cloning ................................ .......................... 82 Purification and Quick Evaluation of Purified Enzymes ................................ .... 83 Determination of Optimal pH and Temperature for Activity .............................. 84 Analysis of Enzymatic Substrate Profile ................................ ........................... 84 Biochemical Properties of LJ0536 and LJ1228 ................................ ................ 85 Effect of B ile Salt Components ................................ ................................ ......... 88 In Silico Analysis of FAE Genomic Context ................................ ...................... 89 Analysis of FAEs Primary Sequences ................................ .............................. 89 Summary ................................ ................................ ................................ ................ 91 4 X RAY CRYSTALLIZATION AND SUBSTRATE BINDING MECHANISM OF LJ0536 ................................ ................................ ................................ .................. 105 Backg round ................................ ................................ ................................ ........... 105 Result and Discussion ................................ ................................ .......................... 108 Architecture of LJ0536 ................................ ................................ ................... 108 T he S106 is the Catalytic Residue ................................ ................................ 109 Analysis of the Crystal Structures of S106A Substrate Complexes Reveals Critical Residues for Substrate Binding and Catalysis ................................ 111 Site Role in Substrate Preference ................................ ................................ ...... 116 Comparisons of LJ0536 and Proteins with Similar Fo lding ............................. 117 Summary ................................ ................................ ................................ .............. 120
7 5 A NEW FACTOR CONTRIBUTES TO THE CLASSIFICATION OF FAES ........... 142 Background ................................ ................................ ................................ ........... 142 Result and Discussion ................................ ................................ .......................... 142 Structural Differences of Bacterial and Fungal FAEs ................................ ..... 142 Classification of LJ0536 and LJ1228 ................................ .............................. 144 Structural Prediction of LJ0536 and LJ1228 Homologs ................................ .. 146 Summary ................................ ................................ ................................ .............. 148 6 SUMMARY AND CONCLUSION S ................................ ................................ ........ 162 REFERENCE LIST ................................ ................................ ................................ ...... 164 BIOGRAPHICAL SKETCH ................................ ................................ .......................... 177
8 LIST OF TABLES Table page 1 1 Functional classification of FAEs based on substrate specificity and primary sequence similarity. ................................ ................................ ............................ 38 1 2 Descriptor based classification of FAE proposed by Udatha .............................. 39 2 1 Strains and plasmids used in Chapte r 3 ................................ ............................. 69 2 2 Primers used in Chapter 3 ................................ ................................ .................. 70 2 3 P lasmids used in Chapter 4 ................................ ................................ ................ 72 2 4 Primers used in Chapter 4 ................................ ................................ .................. 73 2 5 Strains used in Chapter 5 ................................ ................................ ................... 76 2 6 Primers used in Chapter 5 ................................ ................................ .................. 77 3 1 Saturation kinetic parameters of LJ0536 and LJ1228 ................................ ......... 94 4 1 Statistics of X ray diffraction and structure determination ................................ 122 4 2 Saturation kinetic parameters of LJ0536 variant s ................................ ............. 123 5 1 C omparison of LJ0536 and AnFaeA ................................ ................................ 150 5 2 Structural prediction of fungal FAEs using SWISS MODEL (automatic model ing ) ................................ ................................ ................................ .......... 151 5 3 Structural prediction of fungal FAEs using SWISS MODEL (manual modeling ) ................................ ................................ ................................ ......................... 152 5 4 Structural prediction of putative FAEs in subfamily 1B using SWISS MOD EL (automatic modeling ) ................................ ................................ ........................ 153 5 5 Structural prediction of putative FAE s in subfamily 1B using SWISS MODEL (manual modeling ) ................................ ................................ ............................ 154 5 6 Structural prediction of LJ0536, LJ1228, and homologs / pa r alogs using SW ISS MODEL (automatic modeling ) ................................ .............................. 155 5 7 Structural prediction of LBA 1 and BFI 2 using SWISS MODEL (manual m odeling ) ................................ ................................ ................................ .......... 156 5 8 Structural prediction of bacterial FAEs using SW ISS MODEL (au tomatic modeling ) ................................ ................................ ................................ .......... 157
9 5 9 Structural prediction of bacterial FAEs using SWISS MODEL (manual modeling ) ................................ ................................ ................................ .......... 158
10 LIST OF FIGURES Figure page 1 1 Classification of phytophenols ................................ ................................ ............ 40 1 2 Phenolic acid subgroups ................................ ................................ .................... 41 1 3 Este rification of phenolic compounds ................................ ................................ 42 1 4 Intestinal absorption of phytophenols and phenolic acids ................................ ... 43 1 5 Chemical structures of e ster backbones ................................ ............................. 44 1 6 Natural phytophenols are frequently present in the human diet ......................... 45 1 7 Catalytic mechanism characteristic of the carboxylesterases ............................. 46 2 1 Expression vector, p15TV L map ................................ ................................ ....... 78 3 1 Identification of FAE producing strains ................................ ............................... 95 3 2 Identification of the colonies isolated from BB DR rats. ................................ ...... 96 3 3 Purified enzymes on SDS PAGE ................................ ................................ ........ 97 3 4 Optimal pH and temperature of LJ0536 ................................ .............................. 98 3 5 Optimal pH and temperature of LJ1228 ................................ .............................. 99 3 6 Enzymatic substra te profile of the enzymes LJ0536 and LJ1228 ..................... 100 3 7 Effect of bile salts on LJ0536 and LJ1228 enzyme activity ............................... 101 3 8 Genomi c co ntext of LJ0536 and LJ1228 in the reference strain L. johnsonii NCC 533 ................................ ................................ ................................ ........... 102 3 9 Multiple sequence alignment of LJ0536 and proteins with high sequence identity ................................ ................................ ................................ .............. 103 3 10 T ree representation of LJ0536 and LJ1228 relationships with the proteins that displayed the highest sequence identity. ................................ ................... 104 4 1 General secondary structu ................................ ......................... 124 4 2 Representation of the overall LJ0536 structure ................................ ................ 125 4 3 Determination of the native molecular weight of the enzyme by gel filtration assays. ................................ ................................ ................................ ............. 126
11 4 4 Representation of the single chain LJ0536 structure. ................................ ....... 127 4 5 ted domain in the LJ0536 structure ................................ ... 128 4 6 Surface and ribbon representation of LJ0536 catalytic site .............................. 129 4 7 Enzyme activity in presence of specific inhibitors. ................................ ............ 130 4 8 Identification of the two GXSXG motifs in the overall LJ0536 structure ............ 131 4 9 SDS PAG E. ................................ ................................ ................................ ...... 132 4 10 Comparative enzymatic activity of LJ0536 variant s. ................................ ......... 133 4 11 Circular dichroism spectra of LJ0536 and mutant S68A ................................ ... 134 4 12 Surface representation of apo and co crystallized structures of LJ0536 mutant S106A ................................ ................................ ................................ ... 135 4 13 Enzyme substrate interactions within binding cavity of LJ0536 ........................ 136 4 14 Structural superimposition of the mutant S106A co crystallized with ethyl ferulate or ferulic acid ................................ ................................ ....................... 137 4 15 Electron density map of co crystallized substrates ................................ ........... 138 4 16 Schematic interpretation of the substrate interactions with LJ0536 binding cavity ................................ ................................ ................................ ................ 139 4 17 Structural comparison of LJ0536 and proteins with similar overall folding ........ 140 4 18 Structural comparison of LJ0536 with Est1E, and VACVase ............................ 141 5 1 Structural comparison of LJ0536 and AnFaeA ................................ ................. 159 5 2 Structure of FAE XynZ ................................ ................................ ..................... 160 5 3 Structural comparison of LJ0536 and FAE XynZ co crystallized with their respective substrates ................................ ................................ ........................ 161
12 LIST OF ABBREVIATION S Amp ampicillin Amp r ampicillin resistance ATCC American type culture collection BB D P bio breeding diabetes prone BB DR bio breeding diabetes resistant BES N,N b is(2 hydro xyethyl) 2 aminoethanesulfonic a cid BLAST Basic Local Alignment Search Tool bp base pair BRENDA BRaunschweig ENzyme D a tabase C carbon CHES 2 ( n c yclohexylamino)ethane Su lfonic Acid cm centimeter c.n.d. could not determine DNA d eoxyribonucleic acid dNTPs deoxyribonucleotide triphosphate s DTT d ithiothreitol extinction coefficient EC Enzyme Commission number EF ethyl ferulate FAE f erul ic a cid e sterase FAE A type A ferulic acid esterase FAE B type B ferulic acid esterase FAE C type C ferulic acid esterase FAE D type D ferulic acid esterase
13 FAE E type E ferulic acid esterase FPLC fast protein l iquid c hromatography F o F c Fourier refinement g gravitational force GRAS Generally Recognized As Safe HEPES 4 (2 hydroxyethyl) 1 piperazineethanesulfonic acid His histidine HPLC h igh performance liquid chromatography IPTG isopropyl D 1 thiogalactopyranoside k b kilobase pair K cat catalytic rate constant K cat / K m catalytic efficiency k D a kilodalton K m Michaelis constant L liter LAB lactic acid bacteria LB Lysogeny broth / Luria Bertani LIC ligation independent cloning M m olarity MCT monocarboxylic acid transporter MES 2 (n morpholino)ethanesulfonic acid mAbs milliabsorbance MCT monocarboxylic acid transporter mg milligram min minutes
14 m L milliliter mM millimolar mm millimeter MR m olecular r eplacement MRS de Man Rogosa Sharpe NaCl sodi um chloride NCBI National Center for Biotechnology Information Ni NTA nickel nitriloacetic acid n mol nanomole NOD non obese diabetic o C degree celsius OD 600 optical density at 600nm ORFs open reading frames PCR polymerase chain reaction PDB Protein Data Ba nk PMSF phenylm ethanesulphonylfluoride PSI BLAST Position Specific Iterated Basic Local Alignment Search Tool R residual factor R free free residual factor R work a residual factor RNA r ibonucleic acid rpm r evolutions per minute s second SDS PAGE sodium dode cyl sulfate polyacrylamide gel electrophoresis SEC size exclusion chromatography
15 sp. Species (singular) spp. species ( plural) TEV tobacco etch virus TID type 1 diabetes g microgram L microliter m micrometer UV ultraviolet V max maximum rate of reaction v / v volume to volume w / v weight to volume w / w weight to weight
16 Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy BIOCHEMICAL AND STRUCTURAL CHARACTERIZATION OF Lactobacillus johnsonii FERULOYL ESTERASES By Kin Kwan Lai August 2011 Chair: Claudio F Gonzalez Major: Microbiology and Cell Science Phytophenols are natural phenolic compounds with widespread distribution throughout the plant kingdom. These phytophenols participate in the formation of macromol ecular structures in plant cell walls via ester linkages. Phenolic acid s, chemi cals that possess beneficial properties to human health, are released during hydrolysis of phytophenols. The hydrolysis is catalyzed by enzymes (ferulic acid esterase s FAE s ) of the gut microbiota Before this work, no enzymes produced by human gastrointes tinal commensals displaying FAE activity were described FAE activity was observed from several Lactobacillus sp p. isolated from stool sample s collected from bio breeding diabetes resistant rat s The isolated Lactobacillus johnsonii N6.2 dis played the high est FAE activity among the isolated strains P otential FAE s were identified by in silico prediction cloned, and purified from Escherichia coli as recombinant proteins By utilizing a variety of enzyme activity assays, two enzymes ( LJ0536 and LJ1228 ) showe d high substrate preference for aromatic ester s including chlorogenic acid and rosmarinic acid which are commonly found in human diets. Site directed mutagenesis and x ray crystallization of LJ0536 identified critical amino acid s
17 involved in ester hydrol ysis. The catalytic triad is composed of serine, histidine and aspartic acid A classical oxyanion hole formed by phenylalanine and glutamine also contributes to substrate binding The substrate binding mechanism consist s of a specific hydrophobic cavit y in addition to an inserted domain located on top of the binding cavity The inserted domain protect s the small hydrophobic region and forms hydrogen bond (s) with the aromatic ring of the substrate to stabilize the binding interaction. Bioinformatic and s tructural analyses indicate bacterial FAE s are well conserved within the Lactobacillaceae family. T he features of inserted domain distinguish the substrate binding mechanism of different types of proteins The c urrent FAE classification scheme is primarily based on enzyme activity and primary sequence identity of fungal FAE s Th e unique structural and functional features of the inserted domain could contribute to refine the classification of FAEs. Taken together, this work describes the identification, puri fication, characterization, and crystallization of FAEs from probiotic bacteria. This study will provide insight for further exploration of FAE s in other species, and potentially will enhance the path for future applications of FAEs
18 CHAPTER 1 INTRODUCTIO N Phytophenols P henolic compounds are naturally available chemicals that contain one or more phenolic ring s with or without substituents such as hydroxyl or methoxy group( s ) The term phytophenol, or phytochemical, is also used due to their high abundance in plant s (Huang et al. 2007) Phytophenols are secondary metabolites of plants, which are primarily use d as defen ces against ultravi olet radiation and pathogens (Beckman, 2000) They are widely distributed and highly abundant in plant cell wall s. F erulic and p coumaric acids reinforce the cell wall structure by cross linking the hemicellulose fra ction In general, phytophenol s are divided into four major groups: flavonoids, phenolic acids, stilbenes, and lignans (Spencer et al. 2008) The classification of phytophenol s is based on the ir characteristic chemical structure s (Figure 1 1). All chemicals classified as phytophenols have at least one phenol ic ring in the ir structure. The characteristic structure of flavonoid s is a flavone which contains two benzene rings connected by three carbons to form an oxygenated heterocycle. The phenolic acids contain a benzene ring attached to a carboxylic acid. The s tilbene s have a backbone structure of 1,2 diarylethene, with two benzene rings bonded to each end of a carbon carbon double bond. Lignan s have a backbone structure of 1,4 dibenzybutane, with two benzene rin gs bonded to each end of a four carbon chain. Th e se major groups can be further divided into small subgroups d epending on the position and number of hydroxyl substituents or other derivatives present in t he backbone structure. Among these four major groups, the flavonoid s are the larg est group and conta in six subgroups: flavonols, flavanones, flavanols, flavones, anthocyanins, and isoflavones. Phenolic acid s can be
19 further divided into two groups: hydroxycinnamic acid s and hydroxybenzoic acid s The h ydroxycinnamic acid s have a cinnamic acid as a backbone structure while the hydroxybenzoic acid s have a backbone structure of benzoic acid. Even though hydroxycinnamic acid s and hydroxybenzoic acid s have similar chemical structure s hydroxycinnamic acids are more common in nature than hydroxybenzoic acids. Ph enolic acids with a single phenolic ring, such as caffeic, ferulic, and coumaric acids are the simple st derivatives of hydroxycinnamic acid s (Crozier et al. 2009) Caffeic acid has hydroxyl substituents at carbon 3 and carbon 4 (C3 and C4) of the cinnamic acid backbone Ferulic ac id has a hydroxyl substituent at C4 and a methoxy substituent at C3 of the cinnamic acid backbone Coumaric acid has one hydroxyl substituent at C4 of the cinnamic acid backbone. Salicylic acid, syingic acid, and gallic acid are examples of hydroxybenzoic acid derivatives (Figure 1 2). Salicylic acid has a hydroxyl substituent on C2 of the benzoic acid backbone. Syingic acid has methoxy substituent s on C3 and C5 and a hydroxyl substituent on C4 of the benzoic acid backbone. Gallic acid has hydroxyl substitu ent s on C3, C4, and C 5 of the benzoic acid backbone. Depending on the type of storage or the location in the plants phenolic acids can be either soluble or in soluble. The p henolic acid s are soluble when they are store d within plant cell vacuoles They are insoluble when they are acting as components of the plant cell wall structure However, i n reality, phenolic acid s exi st in much more compl ex form s The carboxylic acid moiety is usually esterified (Figure 1 3 A and B ) which generates a variety of phenoli c compound s Usually more than one phenolic moiety can
20 be found in the se chemical structures. Consequently, phenolic compounds are also called polyphenols. For example, chlorogenic acid ( 5 O caffeoylquinic acid) is composed of two phenolic acids; i t is an ester of caffeic acid and quinic acid. Chlorogenic acid is a soluble phenolic compound found in a variety of plants For instance, Catharanthus roseus produces terpenoid indole alkaloids utilized for anti cancer drug synthesis (Ferreres et al. 2011) Rosmarinic acid is an other soluble phenolic compound which is found in the extract of Labiatae herbs (Tada et al. 1996) It is an ester of caffeic acid and 3,4 dihydroxyphenyl lactic acid. In contrast, ferulic acid and coumaric acid are found in the cell walls of b arley and m alt These acids are ester linked to arabinoxylan and are part of the insoluble fractions ( Figure 1 3C ) (Maillard & Berset, 1995) The hydrolysis of the ester bond releases the phenolic acids from macromolecular structures and from the r espective polyphenols Although the chemical structures of phenolic acids are similar, they have different biochemical properties. In the past decade, the studies on phenolic acid s increased dramatically due to its beneficial properties demonstrated in vit ro as well as in vivo (Srinivasan et al. 2007) Health Bene fic i al Properties of Phenolic Acids It is generally accepted that the beneficial pro perties shown by phenolic acids are related to their high level of anti oxidative and anti inflammatory properties (Maurya & Devasagayam, 2010; Sato et al. 2011) They have strong scavenging activity for free radicals such as hydrogen peroxide, superoxide, hydroxyl radical, and nitrogen dioxide (Srinivasan et al ., 2007; Graf, 1992) I n addition, it is accepted that phenolic acids are able to stimulate insulin secretion to maintain normal blood glucose level s
21 (Adisakwattana et al. 2008; H uang et al. 2009) reduc e carcinogenesis (Murakami et al. 2002; Yi et al. 2005) and dimin ish cardiovascular disease (Chao et al. 2009) It has been demonstrated that ferulic acid has neuroprotective effect in rat s (Cheng et al. 2008) and protect s against liver injury in mice (Kim et al. 2011) C affeic acid shows inhibitory effect s against cancer cell proliferation i n human cell line s (Rajendra Prasad et al. 2011) Besides these common derivatives of hydroxycinnamic acid s the importance of other divers e phenolic acids and poly phenols such as hydroxytyrosol, 3,4 dihydroxyphenyl lactic acid, and resveratrol were recently discovered and studied (Yu et al. 2010) H ydroxytyrosol show ed anti atherogenic, cardioprotect ive, anti inflammatory, anti platelet aggregation anti tumor, and anti microbial activities (Granados Principal et al. 2010) 3,4 dihydroxyphenyl lactic acid has been found to have protective effect s against brain and liver injuries (Lam et al. 2003; Xing e t al. 2005) Resveratrol a compound found in red wines is one of the most studied and commercially exploited phenolics by the nutraceutic al industry. It has been demonstrated that resveratrol is an excellent dietary anti oxidant and can prevent uncontr olled cell proliferation and cancer (Athar et al. 2009; Pervaiz & Holme, 2009) A ntiviral and anti microbial properties of diverse phenolic acids are also well documented (Puu pponen Pimi et al. 2005) In general, in vitro assays have been used to demonstra te the beneficial properties of phenolic acids A few works carried out with animal models indicate that a diet rich in phytophenols could be beneficial for humans However the evidence is still indirect. The assays using animal models described in the scientific literature utilized purified phytopheno ls delivered directly into the blood stream (Kim et al ., 2011) In order to
22 strongly support that the inclusion of diet ary food components rich in phytophenols is beneficial to human s several aspects such as toxicity and absorbability of phytophenols should be deeply investigated. Common Phytophenols Pre sent in Human Diets Phytophenols are highly abundant in the plant kin g dom and can be easily found in dietary fiber However, the distribution of phenolic acid s is highly variable among different species of plants Wheat bran and rice bran o il contain g amma oryzanol. Gamma oryzanol is a phytosteryl ferulate mixture. It is composed of 1 2 ferulate esters (Akihisa et al. 2000) which can release ferulic acid upon hydrolysis. The total amount of ferulic acid varies fro m 0.5%, 0.9%, and 5% dry weight in wheat bran, suger beet pul p, and corn kernel respectively (Ou & Kwok, 2004) The r ice bran contains 0. 19 % to 0. 4 2 % dry weight of gamma oryzanol (Lilitchan et al. 2008; Chen & Bergman, 2005) Rosmarinic acid is an ester of caffeic acid and 3,4 dihydroxyphenyl lactic acid. It is naturally present in high amounts in herbs such as rosemary and lemon balm whic h are frequently used in food preparation s The content of rosmar inic acid in herbs varies from 0.2 % to 3% dry weight of herbs (Wang et al. 2004a) Chlorogenic acid is abundant in coffee and green tea It is an ester of caffeic acid and quinic acid. A cup of coffee potentially contains 15 mg to 325 m g chlorogenic acid (Richelle et al. 2001) Oleuropein is found in olive tree s and in olive oil It is an est er of elenoic acid and 3,4 dihydroxy phenylethanol (hydroxytyrosol). O live tree leaves typically contain 9% dry weight of oleuropein (Omar, 2010) Salvianolic acid is found in Salvia miltior rhiza which is also known as danshen a C hinese herb al medicine U p to 82.52 m g of salvianolic acid B can be found in one gram of danshen (Li et al. 2008) Hydrolysis of salvianolic acid B forms 3,4 dihydroxyphe nyl lactic acid and lithospermic acid
23 Many studies have shown that the consumption of dietary fiber c an lead to better health (Sansbury et al. 2009; Slavin, 2008) The beneficial properties of dietary fibers are d irectly linked with detoxification by stimulat ing intestinal peristal sis. The actual scientific discussion, regarding to the importance of dietary fibers, is focused o n the importance of the phytophenols absorbed at the intestinal level. However, the absor ption and metabolism of phenolic acid s by humans are not completely understood. It is well know n that the dietary phytophenols are poorly absorbed at the intestinal level In order to be absorbed enzymatic hydrolysis of the ester bond is required to relea se the bioactive phenolic acids from the phytophenols The free carboxylic monophenols are then specificly and efficiently assimilated by cells of the intestinal tract (Kroon et al. 1997) Limitation on Phenolic Acid Absorption The knowledge of phenolic acid s absorption by human s is limited It is accepted that two pathways are being utilized for cellular transport of phenolics (Konish i & Kobayashi, 2005) : a passive paracellular diffusion and an active monocarboxylic acid transporter (Figure 1 4 ) Passive p aracellular diffusion is a low efficiency system that allows some phytophenol s and phenolic acid s to slowly pass through the intest inal epithelial cells into the blood plasma for absorption. The a ctive monocarboxylic acid transporter allows small simple chemical s with a monocarboxylic acid motif ( monophenolic acids such as ferulic acid and gallic acid ) to pass through the layer with high efficiency. M onocarboxylic acid transporter do es not have affinity towards complex p hytophenols or phenolic acid esters Since the majority of monophenolic acids are esterified to other molecules, a n enzymatic step is required prior to absorption. Onc e
24 the ester linkage is hydrolyzed, monophenolic acids are released and absorbed in the intestines with high efficiency by the monocarbox y lic acid transporter. The enzymes catalyzing the phenolic acid ester hydrolysis are called cinnamoyl or feruloyl estera ses (FAEs) hydrolyze the ester linkages of polyphenols. However, the metabolites of phenolic acids are detected in the blood stream immediately after the ingestion of phytophenol s (Baba et al. 2004) These results indicate that FAE activity is present in the intestines. It has also been demonstrat ed that FAE activity is present in the lumen of the human gut as well as in the fecal sample s (Kroon et al ., 1997; Gonthier et al. 2006) The h uman colon harbors 10 12 microorganisms per gram of feces (Hooper & Gordon, 2001) It is not surprising that some of these microorganisms encode FAEs (Andreasen et al. 2001) Several bacterial species such as Escherichia coli Bifidobacterium lactis and Lactobacillus gasseri isolated from human intestine display FAE activity (Couteau et al. 2001) It has also been found that lactic acid bact eria such as L fermentum L. reuteri L. leichmanni and L. farciminis are able to produce FAE s (Donaghy et al. 1998) However, the genes encoding th ese enzymes have not yet been identified. Consequently, the presence of FAE activity in the intestines indicates that phenolic acids can be released from the dietary fiber and that FAE activity is produced exclusively by some members of the gut microbiota Microbial Interaction with Food Components Functional food is denoted as food that provides beneficial effects in addition to dietary nutritional value Bioactive food components such as phenolic acids in functional food s are usually tightly bound to t he non digestible fraction The activity of microbial enzymes is required to release the se bioactive component s The i ntestinal
25 tract is an active site not only for absorption and excretion but also for food modification by microorganisms It has been demo nstrated by Kroon that ferulic acid is released from fiber sources such as wheat bran and sugar beet pulp by bacterial FAE activity in the human colon (Kroon et al ., 1997) The bioavailability and function of phenolic acids depend on the specific FAE activity present in the gut microbiota. Not all released phenolics are intestinally absorbe d. The non absorbed p ortion can exert an action in situ (i.e ., anti oxidative) or be s ubsequently convert ed into other metabolites by the gut microbiota. Microbial metabolism and other activities from enzymes such as dehydrogenase s reductase s and decarboxylase s play a critical role in th e phenolics modifications (Landete et al. 2010; Rodrguez et al. 2010; Rodrguez et al. 2009) It has been demonstrated that a change in the composition of gut microbiota affects intestinal permeability, energy homeostasis, and the inflammatory response (Musso et al. 2011) It is also clearly associated with obesity (Cani et al. 2009; Cani et al. 2008) Thus, bioactive food components can alter the health of the host by regulating the metabolism and composi tion of gut microbiota or by directly altering the host metabolism and immune response (Musso et al ., 2011) An additional and very important function of bioactive phenolic acids and their metabolites is related to the regulation of the gut microbiota composition. A num ber of phytophenols can affect the growth and metabolic activity of several members of the gut microbiota (Selma et al. 2009) Consequently, since the gut microbiota play s an important ro le in shaping the host metabolic and immune network, the phenolics will have an important impact on the health of the host
26 The Increase in consumption of function al food s (i.e. fibers) usually leads to an increase not only in the number of probiotic bact eria ( bifidobacteria and lactobacilli ) in the intestine but also in the amount of phenolic acids in the blood (Costabile et al. 2008) Al together, functional food s the gut microbiota, and human health are related to each other. A change on any one of these components will introduce a significant change i n the other components. In order to take advantage of the phenolic contents of dietary fiber, researchers are focused on developing efficient way s to improve the bi oavailability and assimilation of phenolic acids. The use of FAE s produced by the gut microbiota is one of the potential way s to improve the bioavailability of phenolic acids in human diet. Esterase s There are a large variety of esterases described in the literature. Esterases are sub divided in to 31 subgroups on the basis of ester bond specificity (Figure 1 5 ) For example, c arboxylic ester hydrolase s ( EC 3.1 .1 ) target carboxylic ester s; t hiolester hydrolase s (EC 3.1.2. ) target thiolester bonds. Ferulic acid esterases (FAEs) are classified in the group of c arboxylic ester hydrolase s ( EC 3.1.1. ) This group is further divided into 84 specific type s of esterase s based on the functional groups attached to the ester bond (Figure 1 5 ) Consequently, acetyles terase s (EC 22.214.171.124) hydrolyze acet yl ester s For example, they can hydrolyze ethyl acetate into ethan ol and acetate. Arylesterase s (EC 126.96.36.199) hydrolyze ester s that contain a phenyl group attached to the oxygen atom of the ester bond. They can hydrolyze, for example, phenyl acetate into phenol and acetate. Feruloyl esterase s (EC. 188.8.131.52) hydrolyze esters that contain a phenolic acid derivative esterified to an other molecule For example, they can hydrolyze feruloyl polysaccharide to release ferulic
27 acid and polysaccharide. In the past decade, researcher s ha ve focused their attention on FAE s because these enzymes release bioactive phenolic acids from prebiotics Ferulic Acid Esterases ( F AE s ) FAE s (EC 184.108.40.206) are classified as a subclass of carboxylic ac id esterase s (EC 220.127.116.11) Alternative names such as cinnamoyl ester hydrolase s feruloyl esterase s and hydroxycinnamoyl esterase s are general ly used in the literature to describe the same group They are also called hemicellulase accessory enzyme s becaus e they can act synergistically with x ylanases cellulases and pectinases to break down the hemicellulose of plant cell wall s In the presence of water, FAEs hydrolyze phenolic ester s into respective alcohol s and phenolic acid s These enzymes have h igh er s ubstrate preference when the carboxylic ester is in the phenolic / aromatic form, such that an aromatic hydrocarbon is attached to the carbon atom of the carbonyl group of the ester The carbohydrate of the hemicellulose is ester linked to phenolics and th is aromatic ester linkage protect s against hemicellulose degradation by masking the potential substrates for cellulolytic and hemicellulolytic enzymes (Akin, 2008) FAE s are important enzyme s in the rumen ecosystem due to their ability to increase the absorption of energy source s in ruminant animals. In recent years, several FAEs from fungi were partially characterized but little is known about bacterial or plant FAEs. A specific short amino acid sequence glycine X serine X glycine, associated with esterases can be easily identified on primary sequences using bioinformatics analysis. Thus, a large number of proteins are annotated as hypothetical or putative esterases in several databases However, most of them remain biochemically un characterized. Brenda data base ( http://www.brenda enzymes.inf o/ ) described more than 140 enzymes
28 from 52 organisms with known amino acid sequences (Scheer et al. 2011) Only 8 structures of FAEs are describe d in the Protein Data Bank (PDB) ( http://www.pdb.org/ ). All the structures (apo enzymes or co crystallized with a substrate) deposited in PDB be long to two enzymes purified from only two species, Aspergillus niger and B utyrivibrio proteoclasticus In 2004, Wang and his co worker (Wang et al. 2004b) claimed that a feruloyl esterase was successfully purified and characterized from the intestinal bacterium L. acidophilus The molecular weight of the purified enzyme was determined as 36 kDa using sodium dodecyl sulf ate polyacrylamide gel electrophoresis ( SDS PAGE ) The N terminal amino acid sequence of this enzyme was identified as ARVEKPRKVILVGDGAVGST. However, th e N terminal amino acid sequence matches 100% with LA2_01145, a L lactate dehydrogena se (35.1kDa) from L actobacillus amylovorus GRL 1112 and LBA_0271, a L lactate dehydrogenase (35.0k D a) from L. acidophilus NCFM L lactate dehydrogenase s are enzyme s that catalyze the conver sion of py ruvate to lactate. They do not posses s esterase activity. Thus, t here is no evidence in the scientific li terature regarding the purification and characterization of a FAE cloned from l actobacilli General Characteristic of FAE s FAEs are serine esterases that utilize serine as a catalytic residue for hydrolysis. They have a classic al ly conserved pentapeptide esterase motif with a consensus sequence glycine X serine X glycine (GXSXG) with X represents any amino acids They belong to a structural group described as s (Ollis et al. 1992) The seconda ry structure of this group is composed of a minimum of eight strands in the center core surrounded by helices The term barrel is also used to describe
29 the structure. The strands in the central core and helices are mostly parallel. The heli ces and strands tend to alternate along the chain of the polypeptide. The fungal FAEs do not display high sequence homology with bacterial FAEs Since only 2 FAEs structure s were solved by crystallization studies, the substrate b inding mechanism of most FAEs is still not fully understood. However, the X ray structure s of these two FAEs display important differences. These differences suggest that the catalytic and the substrate binding mechanisms of bacterial and fungal enzymes could be substantially diff erent. Reaction Mechanism of FAE s FAEs can hydrolyze a wide range of substrate including both aliphatic and aromatic esters. Enzymes that hydrolyze a broad range of substrates are generally most optimal FAE s ubstrates are chlorogenic acid, rosmarinic acid, bran, ol europein, and salvianolic acid. T hese compounds are naturally present in a variet y of dietary food s (Figure 1 6 ). The active site of these enzymes is formed by a catalytic triad. The triad is formed by serine histidine, and aspartic acid residues, where serine is the nucleophilic residue. Thus the catalytic mechanism of FAEs is very similar to that of serine protease s lipases, and other esterases which involves the formation of a covalent acylenzy me intermediate (Ding et al ., 1994) Two basic steps are i nvolved during carboxylesterase catalysis: acylation and deacylation (Figure 1 7 ) During acylation, the hydroxyl oxygen of the catalytic serine carries out a nucleophilic attack on the carbonyl carbon of the ester substrate ( step 1 8a). After the attack a general base ( the histidine of the catalytic triad ) deprotonates the catalytic serine and the first tetrahedral intermediate is formed ( step 1 8 b). The
30 hydrogen bonding of the third member of the triad, aspart ic acid plays a critical role in the stabilization of the protonated histidine. The oxyanion of the resulting tetrahedral intermediate is positioned towards the oxyanion hole. The oxyanion hole is created by hydrogen bonding between the substrate carbonyl oxygen anion and the backbone of two nitrogen atoms from other residues of the catalytic pocket. The general base, histidine, transfers the proton to the leaving group. The deprotonation o f histidine leads to the protonation of an ester oxygen to release the first product (for example: methanol with methyl ferulate as substrate). As a consequence, the tetrahedral intermediate collapses and the characteristic acylenzyme intermediate is forme d. Thus, the residual half of the substrate remains attached t o the catalytic serine ( step 1 8 c). The s econd step of the reaction, deacylation, takes place in the presence of water. A molecule of water performs a nucleophilic attack on the carbonyl carbon of the remaining substrate in the acylenzyme intermediate ( step 1 8 d) T he general base ( histidine ) immediately d eprotonates a molecule of water, leading to the formation of a second tetrahedral intermediate. The catalysis follow s a similar pattern describ ed for the acylation. The second tetrahedral intermediate is stabilized by the formation of the oxyanion hole ( step 1 8 e). The proton of the general base moves to the nucleophilic serine C onsequently, the ester oxygen is protonated a nd the tetrahedral int ermediate collapse s The protonation of ester oxygen at the expe nse of histidine deprotonation release s the final product (for example: ferulic acid with methyl ferulate as substrate) and reconstitutes the native serine residue and the original state of th e enzyme ( step 1 8 f). The reaction mechanism is summarized in Figure 1 7
31 Structural Binding Mechanism of FAE s The PDB data base displays only two FAE structures co crystallized with ligands (AnFaeA: type A feruloyl esterase from A. niger ; Est1E: feruloyl esterase from B. proteoclasticus ). Although all the enzy / low the same structural pattern, they do not display a conserved substrate binding mechanism. The analysis of the two models co crystal l ized with substrates indicate s that AnFaeA displays the / whi ch is similar to fungal lipases. The e ntry in to the binding cavity is restricted by a lid structure composed of 13 amino acid residues similar to the lipolytic enzymes. Two different conformations are the characteristic s of lipolytic enzymes : the inactive open conformation and the active closed conformation. In the open conformation, the binding cavity is open and in contact with the solvent. The open conformation facilitates the binding of substrate. In the closed conformation, a helical flap structure c overs th e binding cavity and restricts access of the substrate to the cavity. A conformational change in the main protein scaffold facilitates the movement of the helical flap to control the substrate binding (Grochulski et al ., 1994) The helical flap structure of the lipases has remarkable similarity with the lid structure of the AnFaeA. The main diffe rence with the lipases flap structure is that the AnFaeA lid has a higher percentage of polar residues plus a n N glycosylati on site. These feature s suggests lid structure is rigid and the enzyme is always in the open conformation (Hermoso et al ., 2004) The structure of AnFaeA is discussed in depth in C hapter 5 In regards to Est1E, it also di spla with a loop insertion on top of the catalytic groove. The loop insertion participates in the conformation of the catalytic pocket and contributes to the substrate binding. The insertion is composed of
32 51 amino acids with four sma helices. A flapping of one amino acid (tryptophan) from the loop insertion is the only modification in the configuration between the open and closed conformation s Several residues i n the inserted loop participate i n the substrate binding by forming hydrogen bonds with the phenolic moiety of the substrate (Goldstone et al ., 2010) These characteristic s suggest that the lid structures / loop insertions in lipase s fungal FAE s and bacterial FAE s are important for substrate binding. Classification of FAE s A comprehensive classifi cation scheme was proposed in 2004 (Crepin et al ., 2004) Th e classification system use s three main characteristics to group proteins in to four different types: 1) the substrate specificity of enzyme on four substrates (methyl ferulate, methyl sinapate, methyl p coumarate, methyl caff eate), 2) the ability to release diferulic acid from plant cell walls, and 3) the primary amino acid sequence similarity. Th e scheme divide s the FAEs in to subtypes A, B, C, and D. Type A FAEs (FAE A) display activity on methyl p coumarate but not methyl ca diferulic acid from plant cell wall s and the primary amino acid sequence shows similarity with lipases. Type B FAEs ( FAE B) display activity on methyl caffeate but not methyl p coumarate. They are diferulic acid from plant cell wall s The primary amino acid sequence shows similarity to cinnamoyl esterases famil y 1 and acetyl xylan esterases. Type C FAEs (FAE C) display activity on methyl caffeate and methyl p coumarate. They diferulic acid from plant cell wall s and the primary amino acid sequence shows similarity to chlorogenate esterases and tannases.
33 Type D FAEs (FAE D) display activity on methyl caffeate and methyl p coumarate. These enzymes a diferulic acid from plant cell wall s and the primary amino acid sequence shows similarity to xylanase s The full classification scheme (Crepin et al ., 2004) is summarized in Table 1 1. This classification scheme was built based on the data collected from fungal FAEs. Consequently, this classification system may not be valid for classify ing FAEs from all kingdoms (primarily bacteria, and plantae). A second limitation of th e system is related to the number of substrates used for the classification The e sterases d isplay tremendous catalytic flexibility, being active with a large variety of substrates. The use of only four substrates may not be enough to measure the catalytic potential of each group. Even though FAEs display impressive catalytic flexibility and are able to hydrolyze a broad range of substrates, they are very sensitive with any substitutions of the aromatic ring (Vafiadi et al ., 2006) Altering the substitutions of the aromatic rings on meta and / or para position drastically affects the enzym e activity. These characteristics were not used in the classification, perhaps because there is only fragmentary knowledge regarding the mechanisms of substrate binding. A second classification model was proposed in 2008 (Benoit et al ., 2008) Th e new classification scheme is based on the phylogenetic analysis of identified and putative fungal FAEs. The Benoit scheme proposes the division of FAEs in to seven subfamilies, based on the phylogen et ic relationships. The classification does not include biochemical characteri stics A p hylogenetic clustering usually does not correlate with enzymological characteristics. Since the classification was done in silico using the amino acid sequence s, the scheme represents only the phylogen et ic diversity of fungal FAEs.
34 A new classificat ion scheme was also proposed in 2011. T his system includes FAEs from three important kingdoms: bacteria, fungi, and plantae (Udatha et al ., 2011) The main goal of this classification system is to cluster the FAEs that display similar characteristics into the same group The template sequences of FAEs were retrieved from three dif ferent sources: NCBI database ( http://www.ncbi.nlm.nih.gov/ ) biochemically characterized FAEs, and BROAD Institute database ( http://www.broadinstitute.org/ ) / DO E Joint Genome Institute Database ( http://www.jgi.doe.gov/ ) The sequences were analyzed with a sequence derived descriptor software Sequence derived descriptor works with a mathematical algorithm that can cluster proteins with similar function based on the distribution pattern of critical amino acids. The amino acids are identified directly from the primary sequence independently of the full sequence identity (Han et al ., 2004) The authors claim that the pattern of those residues is critical for organizing the catalytic pocket and for substrate binding. Consequently, the proteins clustered in the same group should display similar biochemical properties. The complete classification consists of 12 groups and 31 subgroups. T he main char acteristics of each group are summarized in Table 1 2 Application s of FAE s FAE s ha ve a wide application including paper, biofuel, medical, food and cosmetic industries. FAE s are use d in the pulp and p aper industry (Record et al. 2003; Sigoillot et al. 2005) to remove fine particles from pulp which reduces the use of chlorine based chemicals during the bleaching process It is also important for b iofuel industry especially as the demand for ethanol increase dram atically Thus, hemicellulosic by product s from fermentation become one of the target sources to produce ethanol B y using FAE s it is possible to increase the efficiency of
35 hemicellulosic degradation (Fazary & Ju, 2008) Bi functional enzyme s synthesized b y fusing a FAE and an endoxylanase are also used to improve the degradation of agricultural by products (Levasseur et al. 2005) A n important agricultural by product, ferulic acid, is the precursor of vanillin, a f lavoring f ood additive (Priefert et al. 2001) It can be used as food preservatives because it can inhibit the growth of microorganism s (Ou & Kwok, 2004) Due to its anti oxidative property, ferulic acid is a common ingredients in cosmetic s which c ontributes to skin protection against the UV damage (Srinivasan et al ., 2007) Another important aspect of FAE s is the stereoselective organic synthesis. Carboxylesterases are known to cataly ze the hydrolysis of ester substrates as well as the reverse reac tion, the acy lation of alcohols. Transesterification of secondary alcohol s in low water condition generates s ynthetic substrates that have no structural similarity to the natu ral substrates (Panda & Gowrishankar, 2005) It has been demonstrated that FAE s from Humicola i nsolens are able to catalyze the transesterification of secondary alcohols (Hatzakis et al. 2003; Hatzakis & Smonou, 2005) Pentylferulate ester an aroma tic precursor used in cosmetics and food processing is synthesized in high yield using ferulic acid and acidified n pentanol by A. niger FAEs (Giuliani et al. 2001) Sugar phenolic esters have anti microbial and anti tumor activities (Fazary & Ju, 2008) A FAE produ ced by Fusarium oxysporum is able to esterify several phenolic acids such as hydroxyphenylacetic acid and cinnamic acid with 1 propanol working in a mixture of n hexane / 1 propanol / water condition (Topakas et al. 2003) The ability to perform catalysis in organic systems with low water content indicates that FAEs could be important for synthesizing phenolic chemicals with specific scaffold s This is an
36 important enzyme characteristic required for the synthesis of prodrugs and chiral compounds. A deep knowledge regarding the enzyme biochemistry, estereospecificity, and the molecular mechanisms involved in substrate selection are critical to evaluate the potential of the enzymes to be used in th e s e kind s of applicat ions Project Rationale and Design The objectives of this study were to identify the coding sequences, elucidate the biochemical properties, reveal the enzyme structure, and determine the substrate binding mechanism of a bacterial FAE found in the intestin al tract Lactobacillus johnsonii a bacterium isolated from animal models that display high FAE activity was selected as an enzyme donor to clone recombinant FAEs L. johnsonii was selected because it is also a human commensal that could be used as a pro biotic. It is expected that the FAE activity displayed by L. johnsonii will contribute to: 1) the dietary importance of phenolic acids and 2) the importance of microbial gut esterases o n the improvement of carboxylic phenols absorption at the intestinal l evel. Although several bacteri al species isolated from mammal intestines display FAE activity, the genes encoding FAEs were not identified before this work. A genomic approach was used to identify the genes encoding hypothetical enzymes with potential FAE activity. Once the FAE coding sequences were located, the genes were cloned, expressed in E. coli and purified by nickel affinity chromatography as recombinant His 6 tagged proteins The substrate preference of the selected enzyme s was verified using multip le assay s with a large array of subs trates. The major challenge of the experimental design was the elucidation of the substrate binding mechanism. To accomplish this challenge the nucleo phile of the
37 enzyme was mutated using site directed mutagenesis This strategy was used to perform further co crystallization assays with substrates of interest. The majority of FAEs characterized and described in the public ly available databases were purified from fungal species. The amount of data concerning the biochemis try or even structural information of bacterial FAEs is limited. This work has contribute d to the knowledge of phytophenol esters catalysis by a bacterial FAE.
38 Table 1 1. Functional classifica tion of FAEs based on substrate specificity and primary sequenc e similarity. Type Hydrolyzable Substrates Ability to r elease diferulic acid from plant cell wall P rimary sequence similarity A methyl ferulate, methyl sinapate, methyl p coumarate yes lipase B methyl ferulate, methyl sinapate, methyl caffeate no cinnamoyl esterase family 1, acetyl xylan esterase C methyl ferulate, methyl sinapate, methyl p coumarate, methyl caffeate no c hlorogenate esterase, tannase D methyl ferulate, methyl sinapate, methyl p coumarate, methyl caffeate yes xylanase
39 Table 1 2 Descriptor based c lassification of FAE proposed by Udatha (Udatha et al ., 2011) FAE family Sub family Orientation and distance (number of am ino acid s ) between catalytic residues FEF1 1A D ...........54 81........... S ........ ...79 111........... H 1B S ..........51 183.......... D ...........29 178........... H FEF2 S ...............53............... D ................71....... ......... H FEF3 3A S .........192 269......... D ............36 50............ H 3B S ...............18............... D ..........265 270.......... H 3C H ...............50............... S ................79................ D FEF4 4A S .... .....194 248 ......... D ............ 36 46 ............. H 4B S ...............18............... D ..........154 241........... H 4C S ...........64 69........... D ............30 182........... H 4D H ...............54............... D ..... ...........28................ S or H ...............27............... S ................211................ D FEF5 5A S .........236 255......... D .............37 39............. H 5B S ...........18 89........... D .............47 6 2............. H 5C H ...........71 81........... S ...........84 176............. D FEF6 6A S ..........81 247.......... D .............38 59............. H 6B H ............1 83............ S .............61 84............. H FEF7 7A S .........175 253......... D .............36 47............. H 7B S ...............18............... D ...........233 240........... H 7C S ...........81 83........... D .................56................ H FEF8 8A S .........144 358..... .... D ............32 41.............. H 8B S ...............18............... D ...........204 236........... H 8C H ...........51 87........... S .............18 57............. D 8D D ...........68 89........... S ............86 117... ......... H FEF9 9A S .........212 393......... D .............12 40............. H 9B D ...........16 74........... S ............88 155............ H 9C H ...........36 56........... S .............57 60............. D FEF10 10A S ... .......55 248.......... D .............36 74............. H 10B D ...........69 82........... S .............86 96............. H 10C H ........... 81 83 ........... S .............81 83............. D FEF11 11A S .........209 246........ D .............36 41............. H 11B S ...............18............... D ...........135 243........... H FEF12 12A H ................1................ S .............56 61............. D 12B S ..............211.............. D ........... ..36 46............. H S: serine. D: aspart ic acid H: his tid ine.
40 Figure 1 1. Classification of phytophenols The figure displays the relevant backbone chemical structure of the four central phytophenols groups The groups of flavonoids are further divided in to six subclasses and the phenolic acids in to two subclasses base d on the position and biochemical characteristics of the subtituents groups
41 Figure 1 2. Phenolic acid subgroups The phenolic acid derivatives are classified in to two su b group s : hydroxycinnamic and hydroxybenzoic acids. The figure displays the chemical structures of typical members of each group
42 Figure 1 3. Esterification of phenolic compounds In nature the monophenols are usually esterified to form (A and B) soluble com pounds or associated to macromolecular structures like (C) hemicellulose. The ester bonds are indicated with a red arrow. T he arabinoxylan backbone s are depicted in brown color
43 Figure 1 4 Intestinal absorption of phytophenols and phenolic acids The majority of the dietary phytophenols and phenolic acids are absorbed at the intestinal level. A small portion of phytophenols is absorbed through paracellular diffusion with low efficiency. The remaining phytophenols are subjected to hydrolysis by bacteri al esterases to release the efficiently absorbable phenolic acids. A portion of those phenolic acids can be further modified by bacterial activity. The modified phenolic acids are actively transported by the intestinal cells through the monocarboxylic acid transporter (MCT) with high efficiency. The absorbed phenols circulate in the blood stream to the different parts of the body and are further modified by the host metabolism
44 Figure 1 5 Chemical structures of ester backbones ( A) Ester bonds are pres ent in biological ly relevant substrates. ( B) The chemical compounds depicted are used to illustrate carboxylic ester s, thioesters, and phosphoric acid ester s
45 Figure 1 6 Natural phytophenols are frequently present in the human diet The phytophenols di splayed in the figure (red) are potential FAE substrates present in the human diet. The blue boxes highlight the bioactive products released after enzymatic hydrolysis
46 Figure 1 7 Catalytic mechanism characteristic of the carboxyl esterases that use ser ine as the nucleophile center. The catalytic steps are illustrated using methyl ferulate as a substrate model. R1 represents the phenolic acid moiety and R2 represents methoxy group. X and Y were used to represent the unknown amino acids that will contribu te with catalysis by forming the oxyanion hole. While the catalytic triad ( serine histidine aspartic acid ) is highly conserved, the amino acids of the oxyanion hole may vary
47 CHAPTER 2 MATERIALS AND METHOD S Chemicals, Media, and Strains Chemica ls All a nalytical grade chemicals and desalted oligonucleotides (primers) were purchased from Sigma Aldrich (St. Louis, MO, USA). Ethyl ferulate was purchased from Apin Chemicals Ltd. (Abingbon, OX, UK). Chemicals for buffer and culture medium, reagents, and EZrun TM Protein Marker were purchased from Fisher Scientific (Atlanta, TM high fidelity DNA polymerase, Quick L oad Ta q 2X Master Mix DNA polymerases, Quick Load 100 bp molecular weight standards, Quick Load 1 kb molecular weight standards, deoxyribonucleotide triphosphates (dNTPs), were purchased from New England Biolabs (lpswich, MA, USA). In Fusion TM Dry Down Mix was purchased from Clontech (Mountain View, CA, USA). DNeasy Blood & Tissue Kit, QI AGEN Plasmid Mini Kit, QIAquick PCR Purification Kit, and nickel nitriloaceti c acid resin (Ni NTA Superflow) were purchased from QIAGEN (Valenia, CA, USA). Molecular biology assays were done using ultra pure water (Synergy UV Millipore Water Purification System). Growth Conditions of E. coli Strains Bacterial strains used for cloning and protein expression are summarized in Table 2 1, 2 3, and 2 5. E. coli Library Efficiency TM strain was purchased from Invitrogen TM (San Diego, CA, USA). E. coli BL21 CodonPlus (DE3) RIPL strain was purchased from Stratagene Agilent Technologies ( La Jolla, CA, USA). E. coli strain was routinely used for plasmid purification. E. coli B L21 (DE3) strain was used for protein over expression and subsequent protein purification. Wild type E. coli strains
48 were grown in Lysogeny B roth (LB) medium at 37 o C at 250 RPM For purification of N terminally labeled His 6 tagged proteins, E. coli strains carr ying recombinant plasmid w ere freshly inoculated from 80 o C gly cerol stocks into 25 mL LB medium. The medium was mL 1 and the cells were grown for 16 hours at 37 o C 250 RPM The c ells were then sub cultured (1% v / v ) into 2 L of LB medium and grown at 37 o C, 250 RPM W hen the optical density 600nm (OD 600 ) of the culture reached 0 D 1 thiogalactopyranoside (IPTG) was added to the final concentration of 1 mM to initiate over expression of recombinant proteins The induction was carried out for 16 hours at 17 o C 250 RPM The c ells were harvested by centrifugation and t he cell mass was immediately used for protei n purification or stored at 80 o C until purification. Preparation of C ompetent E. coli C ells The following procedures were used to prepare competent E. coli cells and competent E. coli BL21 cells. A single colony of E. coli was isolated from E. coli or E. coli BL21 streaked LB agar plate supplemented with 10 mM MgCl. The colony was inoculated into 5 mL TyM broth (2% trypt o ne, 0.5% yeast extract, 0.58% NaCl, 0.2 % MgCl w /v ) and incubated for 2 hours at 37 o C, 250 RPM The c ells were inoculated into 300 mL TyM broth and incubated at 37 o C at 250 RPM until OD 600 reached 0.5. The cell mass was collected through centrifugation at 1600 x g for 12 min at 4 o C. The cells were resuspended in 120 mL of Tfb1 buffer ( 100 mM KCl, 50 mM MnCl 2 10 mM CaCl 2 30 mM potassium acetate, 15 % glycerol v / v pH 5.8). The resuspended cells were incubated on ice for 90 min. The c ell mass was collected again by centrifugation at 3000 1600 x g for 8 min at 4 o C. The cells were resus pended in 12 mL of Tfb 2 buffer ( 10 mM
49 MOPS, 10 mM KCl, 75 mM CaCl 2 15 % gl ycerol v / v pH 7) and stored at 80 o C in small each) until further need ed. Isolation and G rowth Condition of Lactobacillus strains Lactobacilli were previously iso lated by pla ting out aliquots of BB DP and BB DR rats stool samples directly in selective de Man Rogosa Sharpe (MRS) agar plates by Dr. Graciela Lorca, University of Florida Cultures were grown at 37 C anaerobic ally in a gas pack system (Rogosa et al. 1951) Individual colonies were picked and inoculated into 6 mL MRS broth and grown at 37 C under anaerobic co nditions without shaking. The isolated strains were co nserved in 96 well plates with 25% final glycerol concentration and stored at 80 C DNA Procedures Lactobacillus Strain identification Total genomic DNA was extracted using DNeasy Blood & Tissue Kit T he selected strains were identified by sequencing an internal 16S rDNA fragment from genomic DNA with Applied Biosystems model 3130 genetic analyzer (DNA Sequencing Facilities, Interdisciplinary Center for Biotechnology Research, University of Florida) usi ng the primers listed in Table 2 2. The result sequence s were blasted against NCBI Data base (Benson et al. 2011) to identify the donor species. In silico Selection of Potential FAE Encoding Genes Genes encoding proteins with potential esterase activity were selected based on in silico prediction using Comprehensive Microbial Resource (CMR) Database (Davidsen et al. 2010) F ive ORFs encoding putative/ hypothetical proteins (locus tag: LJ0044, LJ0114, LJ0536, LJ0618, LJ1228) that displayed the characteristic esterase motif (Brenner, 1988; Cygler et al. 1993) were selected. The genomic sequence of L.
50 johnsonii NCC 533 (GI# 41584196) was used as a reference The primers were designed based on the genomic sequence of L. johnsonii NCC 533 and L. johnsonii N6.2 chro mosomal DNA was used as template for gene cloning. Three ORFs (lotus tag: LREU1549, LREU1667, LREU1684) were selected from reference genomic sequence of L. reuteri DSM 20016 (GI# 148530277) The primers were designed based on the genomic sequence of L. reu teri DSM 20016 and L. reuteri TDI chromosomal DNA was used as template for gene cloning. Cloning of Potential FAEs Plasmid p15TV L (Figure 2 1) contains bla (ampicillin resistance) gene which serves as a selectable marker. It also contains s acB gene whic h encodes levansucrase. Levansucrase is an enzyme that hydrolyzes sucrose to produce levan. The expression of SacB is toxic to E. coli Thus, the growth of E. coli transformed with p15TV L plasmid is inhibited when the LB medium is supplemented with 5 % suc rose (w / v) unless the SacB gene is removed. L igation independent cloning (LIC) sequences are located on the flanking regions of s acB gene. were used to PCR amplify genes of interest. The cloning of PCR fragment s into p 15TV L plasmid were done by DNA recombination in the LIC sequence using In Fusion TM Dry Down Mix (Lorca et al. 2007a) Each in fusion pellet was resuspended in 8 of p15TV L plasmid (75 ng 1 ). of PCR fragment (~ 1 mg 1 ) was of the resuspended pellet plasmid to initiate DNA recombination. The mixture was inc ubated at room temperature for 30 min to generate a recombinant plasmid. During DNA recombination the SacB gene was replaced by the PCR fragment. Thus, LB agar plate s supplemented with 5 % sucros e (w / v) m L 1 ampicillin were used for positive selection. IPTG was used to
51 induce the transcription of the cloned gene. The protein possess ed His 6 tagged at the N terminus following by a TEV protease cleavage site after translation. The genes of interest were PCR amplified from genomic DNA obtained fr om the isolated strains. The DNA amplification was done using Taq 2X master Mix DNA Polymerases. The primers used are listed in Table 2 2 and 2 6 All PCRs were performed using MyCycler TM Personal Thermal Cycler (Bio Rad Laboratories). The PCR fragments we re cloned into p15TV L as described above. T he recombinant plasmids were transformed into E. coli Heat shock transformation procedures were done as follow s : competent cells w ere mixed with recombinant plasmid s The mixture was incubated on ice for 20 min, followed by 5 min in 37 o C water bat h, and 3 of LB medium was added to the mixture and incubated for an additional 45 min in 37 o C water bath. Cells were collected by centrifugati on at 7500 RPM (JLA16.250 rotor, Beckman Coulter) of supernatant was discarded. The cells were resuspended in t supernatant. The cells were plated on LB agar mL 1 ampicillin and 5 % sucrose (w / v) for positive selection. Colony PCR was also used to screen for positive colonies. Plasmids were extracted using QIAGEN Plasmid Mini Kit Sequences of PCR insert of a ll clones were confirmed by Applied Biosystems 3730 capillary sequencer using T7 primers (DNA Lab, Arizona State Universi ty) on the extracted plasmids The plasmids with correct clones were further transformed into E. coli BL21 for protein over production. Cloning of Human Valacylovir Hydrolase ( VACVase ) The plasmid containing the gene of interest (pET17b VACVase) was provid ed by Dr. Gordon L. Amidon, University of Michigan (Lai et al. 2008) The gene of interest
52 was cloned i nto p15TV L plasmid using the primers listed in Table 2 4 and confirmed by DNA sequencing as described above Generating LJ0536 Protein Variants The LJ 0536 p15TV L clone was used as a wild type plasmid template for site directed mutagenesis assay The 39 nucleotide long complementary primers containing the desired mutation were used to introduce individual mutations. The primers are listed in Table 2 4. The amino acids selected for m odification were replaced by alanine due to its small, simple chemical structure (alanine scanning). The inert alanine methyl functional group will not introduce interaction within the protein The mutants were constructed by PCR using Finnymes Phusion TM high fidelity DNA polymerase according To generate a deletion mutant of / (from V147 to A173 of LJ0536 ) primers LJ0536DEL147 173aa SmaI Fw and LJ0536DEL147 173aa SmaI Rv were used for PCR amplification to amplify the LJ0536 p15TV L plasmid. The result ing PCR fragment contained a segment of LJ0536 at the end ( Q174 to F249 ) and a segment of LJ0536 at the M1 to G146 ) connected by the sequence of p15TV L plasmid. It was flanked with SmaI restriction sites on both ends, allowing restriction digestion and ligation to complete the recombinant plasmid. The PCR fragment was digested with SmaI restr iction enzyme for 2 hours at 37 o C. Ligation was carried out using T4 DNA ligase at 16 o C for 16 hours to ligate the SmaI restriction site. The PCR amplified plasmids were then treated with 1 0 units of DpnI restriction enzyme 2 times at 37 o C for 1 hour each time to digest the methylated wild type plasmid template. The recombinant plasm ids were transformed into E. coli D The mutant sequences were confirmed by DNA sequencing (DNA Lab, Arizona State University)
53 DNA Gel Electrophoresis DNA was separated by gel electrophoresis using 1% (w / v) agarose gel in 1X TAE electrophoresis buffer (40mM tris(hydroxymethyl)aminomet hane (Tris) acetate pH 8.5, 2 mM e thylenediaminetetraacetic acid (EDTA )). Gels image s were captured using ImageQuant 400 imaging system (GE Healthcare) after staining with 0.5 g mL 1 ethidium bromide. Protein Procedures Protein Purification The expressi on of His 6 tagged proteins was carried out in E. coli BL21 using IPTG (1 mM) to induce gene transcription on p15TV L. The cells were collected by centrifugation at 8000 RPM (JLA8.1000 rotor, Beckman Coulter) for 25 min. The collected cell mass was resuspen ded in 25 mL binding buffer (5 mM imidazole, 500 mM NaCl, 20 mM 4 (2 hydroxyethyl) 1 piperazineethanesulfonic acid ( HEPES) pH 7.5) and then disrupt ed by French press (20000 psi) The cell free extract was collected by centrif ugation at 4 o C, 17500 RPM (JA25 .50 rotor, Beckman Coulter) for 25 min. The soluble His 6 tagged proteins were purified by affinity chromatography as follow s : all solutions were passed through the Ni NTA column by gravity flow. The Ni NTA co lumn was first washed with 30 mL of ultra pure w ater to wash out any unbound nickel ions. It was then pre equilibrated with 30 mL binding buffer. The cell free extract was applied to Ni NTA column. During this step, the His 6 tagged proteins were bound to nickel ions that were immobilize d by NTA The res i n was washed with 30 mL of binding buffer to wash out any unbound proteins. 200 mL of wash buffer ( 20 mM imidazole, 500 mM NaCl, 20 mM HEPES pH 7.5 ) which contain s a higher concentration of imidazole was used to remove unspecific proteins that were boun d to the resin. Imidazole is a
54 competitive molecule that displace s the nickel ions bound to His 6 tag ged protein The His 6 tagged proteins were eluted using 20 mL elution buffer (250 mM imidazole, 500 mM NaCl, 20 mM HEPES pH 7.5 ) The purified proteins were dialyzed at 4 C for 16 hours. The dialysis buffer was composed of 50 mM HEPES buffer pH 7.5 500 mM sodium chloride ( NaCl ) and 1 mM d ithiothreitol ( DTT ) After dialysis the samples were flash frozen and preserved at 8 0 C in aliquots until needed The His 6 tag was removed by treatment with t obacco etch virus ( TEV ) protease (60 ug TEV protease p er 1 mg of target protein) at 4 o C for 16 hours. The sample was passed through a nickel affinity chromatography column to eliminate the released His 6 tag Co llected proteins were dialyzed at 4 o C against dialysis buffer for 16 hours. The purified protein s without His 6 tag were flash frozen and pre served in small aliquots at 80 o C until needed S odium D odecyl S ulfate P olyacrylamide G el E lectrophoresis (SDS PAGE) Purified proteins were analyzed in SDS PAGE (120 V, 65 70 min.) to verify induction and to determine the purity of proteins after the affinity chromatography. Proteins were mixed with SDS PAGE loading dye (100 mM Tris HCl pH 6.8, 2% (w / v) SDS, 10 % ( v / v) glycerol, 10 % (v / v) mercaptoethanol, 0.6 mg mL 1 bro mophenol blue) and boiled at 95 o C for 5 min prior loading to the gel. Electrophoresis was done in buffer composed of 25 mM Tris buffer, 1 92 mM glycine, and 0.1 % (w / v) SDS. Gels were stained with Coomassie Blue ( PhastGe l Blue R 350 ) and images were captured using HP Scanjet G3010 Scanner ( Hewlett Packard ) Protein Quantification Protein concentration was quantified using Bradford reagent (Bradford, 1976) The calibration of Bradford reagent was done using bovine serum albumin as a
55 standard. Absorbance at 595 nm (A 595 ) was determined using UV 1700 PharmaSpec UV VIS Spectrophotometer (Shimadzu). Enz ym e A ssays Feruloyl e sterase s creening a ssay The ability of Lactobacillus strains to produce FAEs was analyz ed on MRS agar plate s supplemented with 0.1% (w / v) ethyl ferulate with out glucose (Donaghy et al ., 1998) The presence of ethyl ferulate created a turbid / milky appearance of MRS agar due to the semi soluble ethyl ferulate at 0.1 % (w / v) final concentration. Ferulate assay (MRS EF) plates were inoculated with cell obtained from individual overnight MRS cultures The plates were incubated at 37 C in a gas pack system for a maximum of 3 days. The f ormation of halo (clear area) around the colonies indicated the presence of f erulate esterase activity. The strains with the highe st activity (largest halos) were selected for further analysis. Determin ation of optimal assay conditions Model carboxylesterase substrates (4 nitrophenyl butyrate ) w as used as enzyme substrate. The optimal pH for catalysis of each purif ied enzyme was det ermined at 37 o C using a set of overlapping buffers: 2 (n morpholino)ethanesulfonic acid (MES) pH 5.5 6.4, N,N b is(2 hydro xyethyl) 2 aminoethanesulfonic a cid ( BES ) pH 6.4 7. 8, HEPES pH 6.8 8. 2, Tris HCl pH 7.5 9.0, 2 ( n c yclohexylamino)ethane Sulfon ic Acid (CHES) pH 8.6 10. The buffers were used at 20 mM final concentration. The optimal temperature of each purified enzyme was estimated by incubating the reaction mixture at different t emperatures (13 40 o C) at the optimal pH of each enzyme. The r ea ction mixture consisted of 20 mM buffer, 1 mM 4 nitrophen y l butyrate, and 0.3 of enzyme per mL of reaction mixture 4 nitrophenols were released
56 from 4 nitrophenyl butyrate during hydrolysis. Enzyme activity was continuously monitored for 15 min a t 412 nm using UV 1700 PharmaSpec UV VIS Spectrophotometer (Shimadzu) or Detection Microplate Reader ( Biotek ) The increase in absorbance due to the increase concentration of free 4 nitrophenols indicated enzyme activity. All assays and c ontrols were performed in triplicate E xtinction coefficient of 4 nitrophenol (16300 M 1 cm 1 ) was used to quantify the release of 4 nitrophenol using the following equation: Equation ( 2 1 ) measured (mM min 1 change of absorbance (Abs min 1 e xtinction coefficient of chemical measured (M cm 1 ). l is the path length of light traveled through the sample (automatically adjusted to 1 cm by reader). Enzyme specific a ctivities were calculat ed using the following equations: SA = / [enzyme] Equation ( 2 2 ) SA is the enzyme specific activity represented the amount of chemical released or hydrolyzed per mg of protein per min ( mol mg 1 min 1 ). change of concentration of chemical measured (mM min 1 ) [enzyme] is the concentration of enzym e in the reaction mixture (mg mL 1 ). Determination of enzymes s ubstrate preference The enzymatic substrate profile s were determined at 25 o C using an ester l ibrary composed of variety of ester substrates (Liu et al. 2001) The enzyme activity was monitored with 4 nitrophenol (Janes et al. 1998) using the following protocol T he p urified enzymes were thaw ed from 80 o C an d re dialyzed against 5 mM BES buffer pH
57 7.2 The reactions were carried out in 96 well plates; e ach enzymatic reaction contained 1 mM ester substrate, 0.44 mM 4 nitrophenol (proton acceptor), 4.39 mM BES pH 7.2 7.1% (v / v) acetonitrile, and 30 35 g per mL of enzyme in a total volume reaction mixture. The 96 well plates were incubated at 25 o C using Detection Microplate Reader ( Biotek ) The reactions were continuously monitored for 30 min at 404 nm T he concentration of 4 nitrophenol was estimated using the extinction 1 00 M 1 cm 1 ) and Equation 2 1 Enzyme specific activity was calculated using Equation 2 2. All assays and controls were performed in triplicate. Results are shown as mean standard deviation. Determination of biochemical parameters by s aturation kinetics As enzyme reactions are saturable, the biochemical parameters such as K m (Michaelis constant: amount of substrate required to reach half of V max which associates with substrate affinity. Low value of K m indicates high subs trate affinity) V max (maximum rate of reaction or maximum enzyme specific activity) K cat (catalytic rate constant, s 1 ), and K cat / K m (catalytic efficiency, M 1 s 1 ) could be determined by measuring the initial rate of the reaction over a range of sub strate concentration K cat is calculated with the following equation: K cat = V max / [enz] Equation ( 2 3 ) mg 1 estimated from the molecular weight of enzyme (LJ0536: 27570 g mol 1 ; LJ1228: 27454 g mol 1 ). The m odel substrates naphthyl acetate naphthyl propionate naphthyl butyrate naphthyl butyrate, 4 nitrophen y l acetate, 4 nitrophen y l butyrate, 4 nitrophen y l c aprylate ) were used to determine the bio chemical parameters of the purified enzyme s (Gonzalez et al. 2006)
58 The enzymatic saturation assays were conducted at the optimal pH and temperature determin ed for each purified enzyme The enzyme activities of the wild type enzyme L J 0536 and all the LJ0536 mutants were carried out in 20 mM HEPES buffer pH 7.8 at 25 C. T he assays with L J 1228 were performed in 2 0 mM MES buffer pH 6.7 at 30 C. All reactions wer e continuously monitored for 15 min at 412 nm in 96 well plate using Detection Microplate Reader ( Biotek ). The extinction coeffi cients of naphthyl esters ( 3000 M 1 cm 1 ) and 4 nitrophenol (16300 M 1 cm 1 ) were used to calculate the amount of substrate hydrolyzed with Equation 2 1. Enzyme specific activities were calculated with Equation 2 2. The kinetic parameter K m and V max were estimated by non linear fitting using Origin 8 software (OriginLab) All kinetic parameters were determined from the average of triplicated assays Enzyme activity assays towards aromatic este rs ( ethyl ferulate and chlorogenic acid ) were conducted by continuously monitor ing the UV absorbance of the reaction mixture at 324 nm for 10 min. The reactions were carried out in a 96 well UV plate using Syner Detection Microplate Reader ( Biotek ) The typical reaction mixture contained 20 mM buffer, 0.01 to 0.20 mM su bstrate, and 0.05 to 0.1 g mL 1 of purified enzyme. The extinction coefficient of = 15390 M 1 cm 1 ) and chlorog = 26322 M 1 cm 1 ) were determined experimentally and were used to estimate the amount of substrate hydrolyzed. The kinetic parameters were determined as described above from the average of triplicated assays Effect of bile salt component an d metal ions on enzyme activity The effect of bile salt components such as sodium glycocholate, taurocholic, and deoxycholic acid were assayed in a range of concentration ( 0.1 to 10 mM) using the model substrate 4 n itrophenyl butyrate The effect of metal ions (FeCl 2 FeCl 3 CdCl 2
59 CaCl 2 CoCl 2 MnCl 2 ZnCl 2 CuCl 2 MgCl 2 ) were assayed in 1 mM. The assays were conducted at the optimal pH and temperature determined for each purified enzyme A typical enzyme reaction mixture contained 20 mM buffer, 1 mM 4 nit rophen y l butyrate, and 0.3 enzyme The reactions were continuously monitored for 1 0 min at 412 nm using Detection Microplate Reader ( Biotek ) and t he enzyme activities were calculated using Equation 2 1 and Equation 2 2. All assays and controls were performed in triplicate LJ0536 mutants and VACVase ester screening assay The enzymatic activities toward aliphatic (4 nitrophen y l butyrate) and aromatic (ethyl ferulate, chlorogenic acid, and rosmarinic acid) substrates were measu red spectrophotometrically using a Detection Microplate Reader ( Biotek ). The hydrolysis of aliphatic ester s was monitored at 412 nm The hydrolysis of aromati c ester s was monitored at 324 nm A typical r eaction mixture contained 20 mM HEPES pH 7.80, 0.1 mM ester substrate and 0.3 mL 1 purified enzymes (ethyl ferulate and chlorogenic acid) or 3 mL 1 purified enzymes (4 nitrophen y l butyrate and rosmarinic acid). Up to 30 mL 1 of VACVase was used to detect enzyme activity. The enzyme reactions of LJ0536 wild ty pe and LJ0536 mutants were carried out at 25 o C The enzyme reactions of VACVase were carried out at 37 o C The extinction coefficients of 4 nitrophen y l butyrate (16300 M 1 cm 1 ), ethyl ferulate (15390 M 1 cm 1 ), chlorogenic acid (26322 M 1 cm 1 ), and rosmarinic acid (15670 M 1 cm 1 ) were used to estimate the amount of substrate hydrolyzed using Equation 2 1 and Equation 2 2 All assay s were performed in triplicate
60 Detection of phenolic acids using h igh p erformance l iquid chromatography (HPLC) HPLC w as used to identify and measure the compound s released by enzymatic action from the complex substrates. A typical r eaction mixture contained 20 mM HEPES pH 7.80, 1 mM ester substrate, and 20 g mL 1 enzyme. All reaction mixtures were incubated for 16 hou rs and filtered using 0.45 m filter prior to HPLC analysis. HPLC analys es were performed using the HPLC L 2000 series system (Hitachi) with Symmetry C18 5 m 3.9 mm x 150 mm reversed phase column protected with a Symmetry C18 5 m guard column (Waters) Detection of products released by enzyme activities using bran, ethyl ferulate, chlorogenic acid, and rosmarinic acid as substrates were carried out at 324 nm using linear gradient elution with water / acetic acid / 1 butanol (350:1:7, v / v / v ) and meth anol with a flow rate of 1 mL min 1 (Mastihuba et al. 2002) To detect the hydrolysis of valacyclovir t he reaction mixture contained 50 mM HEPES pH 7.80, 4 mM valacyclovir, and 10 g mL 1 enzyme Detection of enzyme activity was ca rried out at 254 nm using l inear gradient elution with acetonitrile at a flow rate of 1 mL min 1 (Lai et al ., 2008) Determination of native molecular weight using size exclusion chromatography The native m olecular weight of the proteins studied wa s determined by gel filtration size exclusion chromatography (SEC) The assays were performed in a LCC 501 Plus FPLC System (Pharmacia Biotech) with column Superose 12 10 / 300 GL (GE Healthcare) A linear regression fitting standard was constructed using Immunoglobulin G (150 k D a ), bovine serum albumin ( 66 k D a), ovalbumin (45 k D a), trypsinogen (24 k D lactalbumin (14.2 k D a), cytochrome C (12.3 k D a), and vitamin B12 (1.4 k D a) as
61 molecular weight standards M obile phase was composed of 10 mM HEPES buffer and 150 mM NaCl. Data w as analyzed using FPLCdirector TM (Pharmacia Biotech). Analysis of protein secondary structure by c ircular d ichroism Protein secondary structure was estimated using AVIV Quick Start 215 Circular Dichroism Spectrometer with 0.1 cm quartz cuvette (Hellma, Jamaica, NY ). The p rotein samples were thaw ed from 80 o C and re dialyzed aga inst 2 mM HEPES buffer with 50 mM NaCl during 16 hours. The samples were adjusted to 0.2 mg mL 1 in 0.5 mM HEPES buffer with 10 mM NaCl after dialysis. The spectra w as acquired at 1 nm intervals and averaged with 10 scans. Multiple s can s with buffer alon e were used to correct the background The final spectra w as expressed in molar ellipticity (ME) using the following equation: ME = Equation ( 2 4 ) of protein, and l is the path length of the cuvette. X Ray C rystallization of LJ0536 and S106A X ray cry stallization was carried out in collaboration with Banting and Best Department of Medical Research, Centre for Structural Proteomics in Toronto (University of Toron to). The crystal structures were provided by Banting and Best Department of Medical Research Structural Analyses were done in our laboratory. All His 6 tagged proteins were crystallized using the sitting drop method with Intelliplate 96 well plates and a M osquito Crystal liquid handling robot (TTP LabTech), mixing 0.5 of protein at 15 mg m L 1 and 0.5 of reservoir solution, over 100 reservoir solution. The protein solutions were pre treated with the proteases subtilisin and V8 for the wild type a nd catalytic serine deficient ( S106A ) mutant of LJ0536 respectively The
62 proteases stored at 1 mg mL 1 stock solution were added to a final 1: 10 v / v ratio protease:protein). Successful crystallization required the presence of the different proteases, a technique often used to increase the success of crystallization due to removal of disordered / flexible regions that would disrupt crystal formation (Dong et al. 2007) Reservoir solutions were identified through an in house custom crystallization screen that was optimized based on success of common commercial sparse matrix crystallization screens (Kimber et al. 2003) The concentrations used in each case we re: apo LJ0536 enzyme: 0.1 M MES pH 6 and 20% (v / v) PEG10K; catalytic serine defici ent ( S106A ) mutant: 0.1 M sodium cacodylate pH 6.5, 0.2 M calcium acetate, 9% (v / v) PEG 8K; S106A co crystallized with ethyl ferulate Form I : 0.1 M Tris pH 8.5, 0.2 M ammonium sulphate, 25% (v / v) PEG 3350; S106A co crystallized with ethyl ferulate Form II : 0.1 M Tris pH 8.5, 0.2 M ammonium sulphate, 24% (v / v) PEG 3350 ; S106A co crystallized with chlorogenic acid: 0.1 M Tris pH 8.5, 0.2 M lithium sulphate, 30% (v / v) PEG 4K; S106A co crystallized with chlorogenic acid: 0.1 M Tris pH 8.5, 0.2 M lithium sulphate, 30% (v / v) PEG 4K; S106A co crystallized with ferulic acid: 0.1 M Tris pH 8.5, 0.2 M lithium sulfate, 30% (v / v) PEG 4K. Ligands were co crystallized at a final ligand concentration of 5 mM (25 mM for ethyl ferulate Form II ) in the sitting dr op, by diluting a stock solution of 100 mM ligand 1:20 v / v with the protein/protease mix; 0.5 of this new solution was mixed with 0.5 of reservoi r solution for crystallization. All crystals were cryo protected with reservoir solution supplemented w ith paratone N oil (Hope, 1988) prior to flash freezing in an Oxford Cryosystems
63 cryostream. Diffraction data at 100 K at the Cu K wavelength were collected at the Structural Genomics Consort ium using a Rigaku FR E Superbright rotating anode with a Rigaku R AXIS HTC detector. Diffraction data w as reduced with HKL2000 (Otwinowski & Minor, 1997) The LJ0536 apo structure was solved by Molecular Replacement (MR) using Phaser (McCoy et al. 2007) with a poly alanine form of the structure of feruloyl esterase (Est1E, PDB: 2WTM) from B. proteoclasticus (Goldstone et al ., 2010) as a search model. The successful MR solution was identified by map inspection using Coot (Emsley & Cowtan, 2004) and by a decrease in R free after refinement using Refmac (Murshudov et al. 1997) The structure was fully built by manual building and rounds of refinement with Refmac, Phenix.refine (Adams et al. 2010) and Buster (Blanc et al. 2004) at the fin al stages. Anisotropic B factors were refined for protein and ligand atoms for all structures. Non crystallographic (NCS) restraints were not utilized for any structure. All structures were refined using TLS parameterization (TLS groups were the N terminal residue to residue 179, and 180 to the C terminal residue), as assigned by the TLSMD server (Painter & Merritt, 2006) Additional TLS restraints resulted in lower R and R free values. Water atoms were added by automatic methods using the refinement programs used in each structure (Phenix.refine, Refmac / CC P4 / ARP / wARP, or BUSTER, respectively). Ions were added after the automatic water building by inspection of magnitude of residual F o F c density and hydrogen bonding patterns. The final atomic models include residues 1 245 of LJ0536, with six atoms from the expression tag at the N terminus of one chain of the asymmetric unit.
64 The LJ0536 S106A structure was solved by MR using the apo structure. All ligands were identified by the presence of residual F o F c density in the active site of the enzyme after mole cular replacement using the apo S106A enzyme. Refinement of ligand structures was executed with geometric restraints generated by the PRODRG server (Schttelkopf & van Aalten, 2004) and with a combination of Refmac and / or Phenix.refine. Final validation of the structure of the ligands was performed by calculating simulated annealing omit F o F c maps using Phenix.refine and Cartesian simulated annealing with defau lt parameters, after removing atoms from the ligand and any protein atoms within 5 of the ligand atoms. In the LJ0536 S106A + ethyl ferulate Form I complex (two chains in the asymmetric unit), one ligand was modeled with an occupancy of 1.0 the other wi th a manually assigned occupancy of 0.55 (due to lower quality electron density, and higher B factors than nearby protein atoms, at higher occupancy levels) For ethyl ferulate Form II complex (one chain in the asymmetric unit), the ligand was modeled with an occupancy of 1.0. All ligands in their respective complexes were modeled with occupancies of 1.0. The structure of Form I and Form II S106A mutant co crystallized with ethyl ferulate are identical. Analyses of the mutant S106A co crystallized with ethy l ferulate were carried out with Form II due to better occupancy of the ligand in the active site All structures were refined until convergence of R work and R free values, and reasonable geometries were verified using the Procheck (Laskowski et al. 1993) and Molprobity (Chen et al. 2010) servers. PDB Accession Code of Proteins The structures of apo wild type LJ0536 (PDB: 3PF8), apo S106A (PDB: 3PF9), S106A bound with chlorogenic acid (PDB: 3S2Z), S106A bound with ethyl ferulate Form
65 I (PDB: 3PFB) S106A bound with ethyl ferulate Form II (PDB: 3QMI ), and S106A bound with ferulic acid (PDB: 3PFC) have been submitted to the PDB (Berman et al. 2000) All other PDB files used in this study were achieved from PDB (Berman et al ., 2000) Structural Analysis All structural images were generated using PyMOL (DeLano, 2002) Structur e similarity searches were performed using the Dali database (Holm & Rosenstrm, 2010) Protein protein interaction interfaces were identified and analyzed with the PDBe PISA server (Krissinel & Henrick, 2007) with default settings; a residue is considered in an interface if its change in accessible surface ar ea between chain A and chain A complex with chain B is no n zero. Sequence Analysis and Construction of Phylogenetic Trees All DNA and amino acid sequences were retrieved from NCBI Database (Benson et al ., 2011) LJ0536 protein homologs were identified by BLASTP search (Altschul et al. 1997) Multiple sequence alignments were performed using CLUSTAL X2 (Larkin et al. 2007) Phylogenetic analyses were conducted using neighbor joining method and visualized with TreeView (Page, 1996) Accession numbers, locus tag, and gene identification numbers for the following figures are listed below. Accession numbers for Figure 3 2 ( p hylogenetic tree of l actobacilli 16S rDNA sequences and the isolated strain N6.2 from BB DR rats stool sample) are as follows: L. sakei 23K (LSA): NC_007576, locus tag LSAr01; N6.2 : isolated FAE producing strain; L. johnsonii NCC 533 (LJO): AE017198, locus tag LJR007; L. delbrueckii subsp. bulgaricus ATCC BAA 365 ( LDE ) : CP000412, loc us tag LBUL_r0045 ; L. acidophilus NCFM (LBA): NC_006814, locus tag LBA2001; L. helveticus DPC 45 71 ( LHE ):
66 CP000517, locus tag lhv_3101; L. reuteri JCM1112 ( LRE ): NC_010609, loctus tag LAR_16SrRNA01 ; L. fermentum IFO 3956 (LFE): NC_010610, locus tag LAF_16SrRNA01 ; L. salivarius UCC118 ( LSL ): NC_007929, loc us tag LSL_RNA001 L. brevis ATCC 367 ( LBR ): C P000416, locus tag LVIS_r0082; L. plantarum WCFS1 ( LPL ): NC_004567, locus tag lp_rRNA01. Gene identification numbers (GI#) for Figure 3 9 ( multiple sequence alignment of LJ0536 and LJ1228 with their homologs and paralogs ) are as follows: LJ0536: L. johnso nii N6.2, cinnamoyl esterase, GI# 289594369; LJ1228: L. johnsonii N6.2, cinnamoyl esterase, GI# 289594371; LREU1684: L. reuteri DSM 20016, alpha / beta fold family hydrolase like protein, GI# 148544890; LAF1318: L. fermentum IFO 3956, hypothetical protein, GI# 184155794; LP2953: L. plantarum WCSF1, putative esterase, GI# 28379396; LGAS1762: L. gasseri ATCC 33323, alpha / beta fold family hydrolase, GI# 116630316; LHV1882: L. helveticus DPC 4571, alpha / beta fold family hydrolase, GI# 161508065; PBR1030: Pr evotella bryantii B14, hydrolase of alpha beta family, GI# 299776930; HMPREF9071: Capnocytophaga sp oral taxon 338 str. F0234, hydrolase of alpha beta family protein, GI# 325692879; BIF00780: Bifidobacterium animalis subsp. lactis BB 12, cinnamoyl ester h ydrolase, GI# 289178448; BACSA1693: Bacteroides salanitronis DSM 18170, protein of unknown function DUF676 hydrolase domain protein, GI# 324318365; MED21706696: Leeuwenhoekiella blandensis MED217, hydrolase of alpha beta family protein, GI# 85830613; HMPRE F1977: Capnocytophaga ochracea F0287, hydrolase of alpha beta family protein, GI# 314946466; GEOTH1777: Geobacillus thermoglucosidasius C56 YS93, alpha / beta hydrolase fold protein, GI# 335362064; SMBG3706: Clostridium acetobutylicum DSM
67 1731, alpha / bet a fold family hydrolase, GI# 336291846; CUW2274: Turicibacter sanguinis PC909, conserved hypothetical protein, GI# 292644698; TMATH1585: Thermoanaerobacter mathranii subsp. mathranii str. A3, BAAT / Acyl CoA thioester hydrolase, GI# 296842777 ; EUBIFOR00351 : Eubacterium biforme DSM 3989, hypothetical protein EUBIFOR_00351, GI# 218217536. Gene identification numbers (GI#) for Figure 3 10 (phylogenetic tree of LJ0536 with its homologs and other cinnamoyl esterases ), Table 5 6 ( Structural prediction of LJ0536, LJ1228, and homologs / paralogs using SWIS S MODEL, automatic model ing ) and Table 5 7 (Structural prediction of LBA 1 and BFI 2 using SWISS M ODEL, manual modeling ) are as follows: L. johnsonii N6.2 cinnamoyl esterase LJ0536 ( LJO 1 ) GI# 289594369. L. johns onii N6.2 cinnamoyl esterase LJ1228 ( LJO 2 ) GI# 289594371. L. gasseri ATCC 33323 alpha/beta fold family hydrolase LGAS1762 ( LGA ) GI# 116630316. L. acidophilus NCFM alpha/beta superfamily hydrolase LBA1350 (LBA 1), GI# 58337623 L. acidophilus NCFM alpha /beta superfamily hydrolase LBA1 842 (LBA 2), GI# 58338090. L. helveticus DPC 4571 alpha / beta fold family hydrolase LHV1882 ( LHV ), GI# 161508065 L. plantarum WCS F1 putative esterase LP2953 ( LP L) GI# 28379396. L. fermentum IFO 3956 hypoth etical protein LA F1318 ( LAF ) GI# 184155794. L. reuteri DSM 20016 alpha/beta fold family hydrolase like protein LREU1684 ( LRE ) GI# 148544890. Butyrivibrio fibrisolvens E14 cinnamoyl ester hydrolase CinI (BFI 1), GI# 1622732. B. fibrisolvens E14 cinnamoyl ester hydrolase C inII (BFI 2), GI# 1765979. Treponema denticola ATCC 35405 cinnamoyl ester hydrolase TDE0358 (TDE), GI# 41815924. Eubacterium ventriosum ATCC 27560 hypothetical protein EUBVEN_01801 (EVE), GI# 154484090.
68 Gene identification numbers (GI#) for Table 5 2 ( Stru ctural prediction of fungal FAE s using SWI SS MODEL, automatic modeling ) and Table 5 3 ( Structural prediction of fungal FAE s using S WISS MODEL manual modeling ) are as follow: Neurospora crassa feruloyl esterase (NCR) GI# 9955721 Penicillium funiculosum f eruloyl esterase (PFU), GI# 25090320. Piromyces equi feruloyl esterase (PEQ), GI# 23821548. Gene identification numbers for Table 5 4 ( Structural prediction of putative FAE s in subfamily 1B using SWIS S MODEL, automatic modeling ) and Table 5 5 (Structural p rediction of putative FAE s in subfamily 1B using SWISS MODEL, manual modeling ) are as follow s : Leptospira biflexa serovar Patoc strain putative feruloyl esterase (LBI), GI# 183222795 Paenibacillus sp W 61 putative feruloyl esterase (PAE), GI# 133251525 Clostridium cellulovorans 743B putative esterase (CCE), GI# 242261429 Geobacillus sp. Y412MC10 putative esterase (GEO), GI# 192811693 Spirosoma linguale DSM 74 hypothetical protein SlinDRAFT_02770 (SLI), GI# 229867621. Algoriphagus sp. PR1 Possible xylan degradation enzyme (ALG), GI# 126648512. Gene identification numbers (GI#) for Table 5 8 (Structural prediction of bacterial FAE s using SWI SS MODEL, automatic modeling ) and Table 5 9 (Structural prediction of bacterial FAE s usin g SWISS MODEL manual modeli ng ) are as follow: Treponema denticola F0402 cinnamoyl ester hydrolase (TDE 2), GI# 325475449 Streptococcus sanguinis VMC66 cinnamoyl ester hydrolase (SSA), GI# 322123198. Ruminococcus albus 8 feruloyl esterase family protein (RAL), GI# 324108892. Cellulo silyticum ruminicola feruloyl esterase III (CRU), GI# 326781741. Prevotella oris F0302 feruloyl est erase (POR), GI# 281401992.
69 Table 2 1. Strains and plasmids used in Chapter 3 Strains or Plasmids Genotype / Description Source or Reference E. coli F lac Z lac ZYA arg F )U169 rec A1 end A1 hsd R17 (rk mk+) pho A sup E44 thi 1 gyr A96 rel A1 Invitrogen BL21 F omp T hsd S (rB mB ) dcm + Tetr gal end A Hte [ arg U pro L Cam r ] [ arg U ile Y leu W Strep/Spec r ] Stratagene Agilent Technolo gies Lactobacillus spp. N6.1 Lactobacillus sp. isolated from BB DR rat stool sample. T his study N6.2 Lactobacillus sp. isolated from BB DR rat stool sample. T his study N6.4 Lactobacillus sp. isolated from BB DR rat stool sample. T his s tudy INT173 Lactobacillus sp. isolated from BB DR rat stool sample. T his study TD1 Lactobacillus sp. isolated from BB DR rat stool sample. T his study PN2 Lactobacillus sp. isolated from BB DR rat stool sample. T his study Plasmid p15TV L Amp r T7 promoter driven expression, LIC sequence for DNA recombination cloning, N terminal 6X His fusion tag followed by a TEV cleavage site (Guthrie et al. 2007) Amp r : ampicillin resistance. TEV: tobacco etch virus.
70 Table 2 2. Primer s used in Chapter 3 Primer Names Primer Sequences Description LJ_0044 Forward 5' TTGTATTTCCAGGGC ATGAAATTACTTCTTACCGGCG 3' G enerate coding region of LJ0044 using template genomic DNA of isolated Lactobacillus strain LJ _0044 Reverse 5' CAAGCTTCGTCATCA TCAATTAGAAATTTGATTTAATTTTTGAACAATT 3' LJ_0114 Forward 5' TTGTATTTCCAGGGC ATGAAAATAGATAATTTAACGTTAACAAATTTT 3' G enerate coding region of LJ0114 using template genomic DNA of isolated Lactobacillus strain LJ_0114 Reverse 5' CAAGCTTCGTCATCA CTAAACGTAAATTCTTCTATCTTTCAA 3' LJ_0536 Forward 5' TTGTATTTCCAGGGC ATGGCAACAATTACACTTGAGC 3' G enerate coding region of LJ0536 using template genomic DNA of isolated Lactobacillus strain LJ_0536 Reverse 5' CAAGCTTCGTCATCA TTAAAACGCATTATTATTCT GTAAAAAATC 3' LJ_0618 Forward 5' TTGTATTTCCAGGGC ATGAAAAAAATTATTCTTTTTGGTGATTC 3' G enerate coding region of LJ0618 using template genomic DNA of isolated Lactobacillus strain LJ_0618 Reverse 5' CAAGCTTCGTCATCA TTATGATATAGCAGCTGTTTCTTTC 3' LJ_1228 Forwar d 5' TTGTATTTCCAGGGC ATGGAGACTACAATTAAACGTGAT 3' G enerate coding region of LJ1228 using template genomic DNA of isolated Lactobacillus strain LJ_1228 Reverse 5' CAAGCTTCGTCATCA TTATTTTATTAAAAACTCACCAACTAATTTTAA 3' LREU_1549 Forward 5' TTGTATTTCCAGGGC ATGGA AATTAAAAGTGTTAACTTAGATC 3' G enerate coding region of LREU1549 using template genomic DNA of isolated Lactobacillus strain LREU_1549 Reverse 5' CAAGCTTCGTCATCA CTAAATTAAATTCAGTTCAGTTAACCA 3' LIC sequences for DNA recombination are in bold
71 Table 2 2. C ontinued Primer Names Primer Sequences Description LREU_1667 Forward 5' TTGTATTTCCAGGGC ATGGTACCGGGGCATAAG 3' G enerate coding region of LREU1667 using template genomic DNA of isolated Lactobacillus strain LREU_1667 Reverse 5' CAAGCTTCGTCATCA CTATTTAATATA GTGATCTAAAAATCTTG 3' LREU_1684 Forward 5' TTGTATTTCCAGGGC ATGGAAATAACAATCAAACGAGATG 3' G enerate coding region of LREU1684 using template genomic DNA of isolated Lactobacillus strain LREU_1684 Reverse 5' CAAGCTTCGTCATCA CTAATTTTTTAAAAAGTTAGCTACCAG 3' T7 Forward 5' TTAATACGACTCACTATAGGG 3' C onfirm gene insertion in p15TV L plasmid and sequencing T7 Reverse 5' GCTAGTTATTGCTCAGCGG 3' Lacto F 5' TGGAAACAGRTGCTAATACCG 3' U niversal primers of l actobacilli for strain identification which amplify 233 bp 16s r DNA fragment for sequencing Lacto R 5' GTCCATTGTGGAAGATTCCC 3' D88 F 5' GAGAGTTTGATYMTGGCTCAG 3' U niversal primers of l actobacilli for strain identification which amplify 1.5kb 16s rDNA fragment for sequencing D94 R 5' GAAGGAGGTGWTCCARCCGCA 3' LIC s equences for DNA recombination are in bold
72 Table 2 3. P lasmids used in Chapter 4 Strains or Plasmids Genotype / Description Source or Reference Plasmid LJ0536 p15TV L Amp r T7 promoter driven expression, N terminal 6X His fusion tag followe d by a TEV cleavage site and LJ0536 coding region cloned by DNA recombination with LIC sequence T his study pET17b VACVase Amp r T7 promoter driven expression, N terminal T7 tag, VACVase coding region (Lai et al ., 2008)
73 Table 2 4. Primer s used in Chapter 4 Primer Names Primer Sequences Description LJ0536_H32A Forward 5' GACATGGCAATCATTTTT GCT GGTTTTACCGCTAACCGT 3' G enerate coding region of LJ0536 with histidine residue at position 32 mutated to alanine residue using LJ0536 p15TV L plasmid as template LJ0536_H32A Reverse 5' ACGGTTAGCGGTAAAACC AGC AAAAATGATTGCCATGTC 3' LJ0536_D61A Forward 5' ATTGCTAGTGTTCGCTTT GCT TTTAATGGCCATGGTGAT 3' G enerate coding region of LJ0536 with aspartic acid residue at position 61 mutated to alanine residue using LJ0536 p15TV L plasmid as templat e LJ0536_D61A Reverse 5' ATCACCATGGCCATTAAA AGC AAAGCGAACACTAGCAAT 3' LJ0536_S68A Forward 5' TTTAATGGCCATGGTGAT GCA GATGGTAAATTTGAAAAT 3' G enerate coding region of LJ0536 with serine residue at position 68 mutated to alanine residue using LJ0536 p15TV L pl asmid as template LJ0536_S68A Reverse 5' ATTTTCAAATTTACCATC TGC ATCACCATGGCCATTAAA 3' LJ0536_D83A Forward 5' GTTTTAAATGAAATTGAA GCT GCAAATGCCATTTTAAAT 3' G enerate coding region of LJ0536 with aspart ic acid residue at position 83 mutated to alanine residue using LJ0536 p15TV L plasmid as template LJ0536_D83A Reverse 5' ATTTAAAATGGCATTTGC AGC TTCAATTTCATTTAAAAC 3' LJ0536_S106A Forward 5' ATTTATCTAGTCGGCCAT GCT CAAGGTGGTGTCGTTGCT 3' G enerate coding region of LJ0536 with serine residue at position 106 mutated t o alanine residue using LJ0536 p15TV L plasmid as template LJ0536_S106A Reverse 5' AGCAACGACACCACCTTG AGC ATGGCCGACTAGATAAAT 3' Mutation sites are in bold with italic
74 Table 2 4. Continued Primer Names Primer Sequences Description LJ0536_D138A Forward 5' GCTGCCACTTTAAAAGGT GCT GCTCTTGAAGGTAATACA 3' G enerate coding region of LJ0536 with aspart ic acid residue at position 138 mutated to alanine residue using LJ0536 p15TV L plasmid as template LJ0536_D138A Reverse 5' TGTATTACCTTCAAGAGC AGC ACCTTTTAAAGTGGCAGC 3' LJ0536_Q145A Forward 5' GCTCTTGAAGGTAATACA GCA GGAGTTACCTATAATCCA 3' G enerate coding region of LJ0536 with glutamine residue at position 145 mutated to alanine residue using LJ0536 p15TV L plasmid as template LJ0536_Q145A Reverse 5' TGGATTATAGGTAACTCC TGC TGTATTACCTTCAAGAGC 3' LJ0536_D197A Forward 5' TTAATCCACGGTACAGAT GCT ACCGTTGTTTCCCCTAAT 3' G enerate coding region of LJ0536 with aspart ic acid residue at position 197 mutated to alanine residue using LJ0536 p15TV L plasmid as template LJ0536_D197A Rev erse 5' ATTAGGGGAAACAACGGT AGC ATCTGTACCGTGGATTAA 3' LJ0536_H218A Forward 5' TATCAAAACAGCACTTTA GCC TTAATCGAAGGTGCAGAC 3' G enerate coding region of LJ0536 with histidine residue at position 218 mutated to alanine residue using LJ0536 p15TV L plasmid as templ ate LJ0536_H218A Reverse 5' GTCTGCACCTTCGATTAA GGC TAAAGTGCTGTTTTGATA 3' LJ0536_H225A Forward 5' TTAATCGAAGGTGCAGAC GCT TGTTTTAGTGATAGCTAT 3' G enerate coding region of LJ0536 with histidine residue at position 225 mutated to alanine residue using LJ0536 p1 5TV L plasmid as template LJ0536_H225A Reverse 5' ATAGCTATCACTAAAACA AGC GTCTGCACCTTCGATTAA 3' Mutation sites are in bold with italic
75 Table 2 4. Continued Primer Names Primer Sequences Description 173aa_SmaI Forward 5' TCC CCCGGG AACAATT GCCTATTTATGAA 3' G enerate coding region of LJ0536 with deletion of amino acid residue from position 147 to position 173 using LJ0536 p15TV L plasmid as template 173aa_SmaI Reverse 5' TCC CCCGGG TCCTTGTGATTACCTTCAA 3' VACVase Forward 5' TTGTATTT CCAGGGC ATGTCGGTAACCTCTGCCAAAG 3' G enerate coding region of VACVase using pET17b VACVase plasmid as template VACVase Reverse 5' CAAGCTTCGTCATCA TTATTGTAGGAAGTCTTCTGCTAACTTG 3' Restriction sites are in italic. LIC sequences for DNA recombination are in b old
76 Table 2 5. Strains used in Chapter 5 Strains or Plasmids Genotype / Description Source or Reference Lactobacillus spp L. gasseri W ild type L. gasseri ATCC 33323 (Lorca et al. 2007b) L. acidophilus W ild type L. acidophilus ATCC 4356 (Lorca et al ., 2007b)
77 Table 2 6. Primer s used in Chapter 5 Primer Names Primer Sequences Description LGAS_1762 Forward 5' TTGTATTTCCAGGGC ATGAAGTTAAAGAAAAAGAAAGTAGG 3' G enerate coding region of LGAS1762 using template genomic DNA of isolated L. ga sseri LGAS_1762 Reverse 5' CAAGCTTCGTCATCA TTAAAAAGTATTATTATCTTGTAAAAATTCTG 3' LBA_1350 Forward 5' TTGTATTTCCAGGGC ATGTTGAAAAAAAGATTTTTATATATTTTTTTGG 3' G enerate coding region of LBA1350 using template genomic DNA of isolated L. acidophilus LBA_1350 Rev erse 5' CAAGCTTCGTCATCA TCAATTATTTAAAAAATCATCGATTAATCCT 3' LIC sequences for DNA recombination are in bold
78 Figure 2 1. Expression vector, p15TV L map Image wa s captured from http://www.sgc.utoronto.ca/SGC WebPages/Vector_PDF/p15TV L.pdf and modified.
79 CHAPTER 3 IDENTIFICATION OF FA ES FROM GUT MICROBIO TA Background The commensal microbiota resi ding in the different niches of the higher organism body s are critical for main tain ing good health However, the mechanisms by which microorganisms interact with the host are still unclear and difficult to study. Important technological advances such as rapid sequencing methods, bioinformatics, and identification using 16S rDNA made possible to describe the variability and composition of the O ne of the most interesting applications of commensal microbiota is the identification of species potentially responsible for specific host diseases. One clear example is the n oticeable changes in the composition of the gut microbial ecosystem of diabet es patients compared to health y individuals (Vaarala et al. 2008) A recent study (Roesch et al. 2009) showed that Bio Breeding D iabetes Prone (BB DP) rats have differences in gut microbiota composition when compared to the isogenic Bio Bree ding D iabetes Resistant (BB DR) animals The result s obtained indicate d that BB DR rats have a predominant presence of probiotic bacteria such as L. johnsonii L. reuteri and Bifidobacterium species when compared with the microbial ecosystem in BB DP rats The microbial ecosystem described before is not unique. In the past decade, a study carried out in Japan found that oral administration of probiotic bacterium L. casei prevents the onset of diabetes in NOD mice by altering the immune response and inhibit ing the disappearance of insulin secreting cells in Langerhans islets (Matsuzaki et al. 1997b) Other studies show ed tha t oral administration of several Lactobacillus spp can help r educe blood glucose levels by stimulat ing insulin secretion (Yamano et
80 al. 2006) via changes in the autonomic neurotransmission (Tanida et al. 2005) However, t he direct mechanisms behind how probiotic bacteria benefit the host are still unclear A feeding study which involves feeding BB DP rat with L. johnsonii has shown that oral administration of the probiotic bacterium L. johnsonii mitigates the incidence of type 1 diabetes by decreasing the intestinal oxidative stress response (Valladares et al. 2010) The decreased oxidative stress at the intestinal level could be a consequence of multiple factors. The interaction of probiotics with the animal food s is probabl y one of the first aspects to be analyzed in order to generate a rationale understanding of the problem. The rat chow is formulated with many ingredients contain ing 6 % to 8% (w / w) of fiber in the form of sugar beet pulp. The sugar beet pulp is an importa nt source of ferulic acid, a phytophenol with anti oxidative and anti inflammatory effects (Couteau & Mathaly, 1998) It has been demonstrated that l ow dos age of cinnamic acids (especially ferulic acid) ha s been related with the stimulation of insulin secretion (Balasubas hini et al. 2003; Adisakwattana et al ., 2008) prevention of oxidative stress lipid peroxidation (Balasubashini et al. 2004; Srinivasan et al ., 2007) and inhibition of diabetic nephropathy progression (Fujita et al. 2008) The phytophenols and its derivatives are tightly attached to plant cell wall materials by ester bond s which limits intestinal assimilation and functi oning of phytophenols. Specific enzymes with good FA E activity are required to hydrolyze and release phenolic acids from the macromolecular structures.
81 I hypothesized that the probiotic bacteria Lactobacillus johnsonii could produce the necessary enzymes to release the antioxidative phenolics. The l actic ac id bacteria (l actobacilli) are well known probiotic bacteria used as food supplements and are present in human intestine. It has been found that several lactobacilli, s uch as L fermentum L. reuteri L. leichmanni and L. farciminis possess FAE activity but the genes encoding these enzymes were not identified (Donaghy et al ., 1998) In this chapter, I described the strain isolation and identification of several colonies of L actoba cillus with the ability to hydrolyze ethyl ferulate (EF) in MRS agar plates The best FAE producer, identified with the name of Lactobacillus johnsonii strain N6.2 was isolated from BB DR rats stool samples (Lai et al. 2009) Using a genomic approach I w as able to identify and p urify several enzymes with esterase activity. The best enzymes with FAE activity were selected. This chapter summarizes the biochemical characteristic of two FAEs purified to homogeneity f rom the probiotic strain L. johnsonii N6.2. Result and Discussion FAEs Producing Strain I solation and Identification C olonies of Lactobacillus were previously isolated by Dr. Graciela Lorca, University of Florida, directly from the stool samples obtained f rom the same BB DR and BB DP rats analyzed by Roesch (Roesch et al ., 2009) The isolated colonies were individually transferred to MRS EF agar plates, with no glucose. The glucose was omitted to prevent a potential catalytic repression of the hydrolytic enzymes. Th e screening plates were used to evaluate the ability of the isolated strains to produce FAE activity. The strains that displayed evident FAE in MRS EF agar generated a clear halo around the colonies (Figure 3 1A) The colonies that displayed the best FAE activity
82 produced clear halo zones of 0.8 0.9 c m in diameter More than 300 colonies were analyzed using this method. Interestingly, 80 5 % (mean standard deviation) of the Lactobacillus colonies isolat ed from BB DR rats demonstrated excellent FAE activity. O nly 41 7 % of the Lactobacillus colonies isolated from BB DP r ats were able to hydrolyze the embedded ethyl ferulate. Six colonies isolated from the BB DR samples showed the largest clear zones on MRS EF screening plates. The colonies N6.1, N6.2, N6.4, TDI, INT173, and PN2 were selected and preserved in glycerol at 80 C to be further identified. The 16S rDNA sequence amplified from the selected isolated colonies belongs to three different La ctobacillus spp The strain, with a colony identification number PN2, showed 96 % sequence identity with L helveticus The strain TDI and INT173 showed 99 % sequence identity with L reuteri Strains N6.1, N6.2, and N6.4 showed 99 % to 100 % sequence identity with L johnsonii (Figure 3 2 ) Among all of the isolated Lactobacillus strains, L. johnsonii N6.2 and L. reuteri TDI di splayed the highest FAE activities (largest clear halo zone s ) and were selected a s DNA donor s to clone potential FAE encoding genes In S ilico Selection of T argets for Cloning The precise identification of L. johnsonii and L. reuteri in the stool samples allowed the use of comparative genomics to select FAE targets in silico F ive open reading frames (ORFs) encoding proteins that displaye d the characteristic motif previously described for esterases were selected (Brenner, 1988; Cygler et al ., 1993) The target genes from L. johnsonii were selected from a group of 346 ORFs encoding hypothetical (306 ORFs) or putative (40 ORFs) proteins as they are annotated in the genome used as a reference, strain L. johnsonii NCC 533 ( http://cmr.jcvi.org/tigr
83 scripts/CMR/CmrHomePage.cgi ). Based on the genome sequence of L. johnsonii NCC 533 primers were designed, and L. johnsonii N6.2 chromosomal DNA was used as a template for gene cloning T o identify potential FAEs in L. reuteri t he genomic sequence of the strain L. reuteri DSM 20016 was used t o design the primers. Three L. reuteri genes were selected for cloning. Purification and Quick Evaluation of Purified Enzymes All eight potential FAE encoding genes ( LJ0044, LJ0114, LJ0536, LJ0618, LJ1228, LREU1549, LREU1667, LREU1684 ) were cloned success fully into the expression vector p15TV L and expressed in E coli BL21 as recombinant proteins Seven out of eight potential FAEs (LJ0114, LJ0536, LJ0618, LJ1228, LREU1549, LREU1667, LREU1684) were purified using n ickel a ffinity c hromatography. The purity o f the His 6 tagged proteins was analyzed by SDS PAGE and stained with Coomassie Blue The results are show n in Figure 3 3. A rapid method to evaluate the FAE activity was used immediately after purification. An aliquot of the purified proteins (3 5 l equiv alent to 0.1 g total protein) were dropped on the surface of the MRS EF screening plate. Three out of seven proteins (LJ0536, LJ1228, LREU1684) displayed FAE activity, as it was demonstrat ed by the formation of halos in the MRS EF screening plat es (Figure 3 1B). It was evidenced that LREU1684 displayed less enzyme activity than the enzymes identified from L. johnsonii The ha los in Figure 3 1B look similar; however protein w ere required to generate a clear zone of similar size to that p protein of LJ0536 or LJ1228. Thus, only LJ0536 and LJ1228 were selected to be further analyzed.
84 Determination of Optimal pH and Temperature for Activity The optimal conditions of both enzymes were determined using the model substrates 4 nitrophenyl acetate and 4 nitrophenyl butyrate. These substrates, the 4 nitrophenyl esters, are routinely using to detect esterase activity because the technique is simple and reproducible. The release of 4 nitrophenol after hydrolysis can be easily detect ed at 412 nm using a visible spectrum spectrophotometer. Since enzymes follow the induced fit model (Koshland, 1958) and esterases are able to hydrolyze a wide range of substrates, it is optimal conditions for activity. The maximal activity of LJ0536 was achieved at pH 7.8 and 20 C (Figure 3 4) while the optimal pH and temperatu re of LJ1228 were pH 6.8 and 30 C (Figure 3 5). T he optimal temperature determined in vitro is low for proteins that were purified from ba cteria living in rat intestines. However, they demonstrated up to 70 % residual activity in a wide range of temperature (15 to 38C ) indicating the proteins could be s till active in the intestine Analysis of Enzymatic Substrate Profile The substrate profile of the selected enzymes (LJ0536 and LJ1228) was determined in parallel using a panel of 27 different substrates The panel was an array of aliphatic and aromatic es ters representing a variety of chemical scaffolds These assays clearly demonstrated that both selected enzymes showed the high est activity towards aromatic esters ( ethyl ferulate, chlorogenic acid and rosmarinic acid ). The screening also revealed the cat alytic flexibility characteristic of the esterases. Several aliphatic esters were also substrates for both enzymes in the study In Figure 3 6 it is evident that LJ0536 showed high activity towards ethyl ferulate but lower activity towards chlorogenic and rosmarinic acids. The results obtained with the enzyme
85 LJ 1228 demonstrated similar hydrolytic ability towards the aromatic esters (Fig ure 3 6 ). This assay is used only to demonstrate t he enzyme substrate preferences since it allows the use of several subs tra tes in parallel. This technique utilizes specific condition s to detect the release of hydrogen ion (proton) during hydrolysis. The buffer (BES buffer) and the pH indicator (4 nitrophenol) to be used in this kind of assays must have similar affinity (BES buffer pK a = 7.09; 4 nitrophenol pK a = 7.15) for the protons released. In this way, the ratio of protonated buffer and the protonated indicator remains constant The pH, produced by the proton release during the enzymatic reaction, shifts and it is detect ed as a change in the yellow color of the indicator present in the mixture Thus, this technique is not flexible enough in order to re create the best conditions (pH, type of buffers, ions etc) that the enzymes require in order to work at its maximal initi al velocity. The specific enzyme activity determined using this method does not reflect the true specific enzyme activity. In addition, the stability of several enzymes c ould be affected because of the exhaustive dialysis in BES buffer The dialysis was do ne using 120 150 times in excess to the volume of enzyme suspension. Consequently, the technique was valid only to demonstrate the substrate preferences even when the conditions (BES buffer pH 7.2, 25 o C) were not the best for the enzymes herein studied. Biochemical P roperties of L J 0536 and L J 1228 The selected enzymes LJ0536 and LJ1228, purified as a single band with an apparent molecular weight of 30 kDa (Figure 3 3). The apparent monomeric molecular weight determined was consistent with the theoretical molecular weight predicted. LJ0536 has a molecular weight of 27.6 kDa, while the LJ1228 enzyme has a m olecular weight of 27.4 KD. The estimated molecular mass includes the TEV cleavage site and the His 6 tag encoded in the plasmid (amino acid sequence:
86 MGSS HHHHHHSSGRENLYFQG, 2.4 kDa) The His 6 tag was removed by TEV treatment. The enzymes do not showed catalytic differences when tagged and un tagged proteins were evaluated in parallel. Consequently, the assays described in this work were carried out with tag ged protein. When the activity of L J 0536 was evaluated in the presence of divalent cations or iron chloride (Fe 3+ ) only Cu 2+ (1 mM) inhibited the activity by 90%. L J1228 enzyme activity was arrested with 1 mM of Zn 2+ Fe 3+ or Cu 2+ The activity of L J 1228 was five times more sensitive to Fe 3+ than the FAE activity described from L.acidophilus (Wang et al ., 2004b) The addition of EDTA to the reactio n mixtures did not affect the activity of these enzymes. Both enzymes were fully inhibited by phenylmethanesulfonyl fluoride (PMSF) and resistant to N ethylmaleimide (NEM) and iodoacetate. These results confirm the presence of serine as the nucleophilic re sidue in the active center, which is suggested by the data from the bioinformatic analysis. The enzymatic parameters obtained by steady state saturation kinetics using a variety of ester substrates are summarized in Table 3 1. In the saturation assays all enzyme s followed a canonical Michaelis Menten hyperbolic kinetic. The biochemical parameters were estimated as described in the Material and Methods section (Chapter 2). As it was determined by the substrate screening method, the enzymes in this study disp layed activity on a wi de range of ester substrates. Both enzymes showed the high est substrate affinity towards aromatic esters when compared to aliphatic esters E thyl ferulate and chlorogenic acids were the best substrate for both enzymes (LJ0536: K m = 0. 020 0.01 mM; LJ1228: K m = 0. 063 0.03 mM ). The affinity obtained with ethyl ferulate was comparable with the affinity obtained with chlorogenic acid (LJ0536: K m
87 0.053 0.01 mM; LJ1228: K m = 0.010 0.00 mM). The chemical scaffold of these two substrates is clearly different. The leaving group, alkoxy group of the chlorogenic acid, is a cyclic polyol (quinic acid) which is bigger than the ethyl group release d from the ethyl ferulate. T his is an important observation and could be used as the first piece o f evidence to suggest that the enzymes recognize only the phenolic moiety of the phytophenol. The molecular aspects of substrate binding will be discussed at the light of the protein structure (Chaper 4). LJ0536 also demonstrates to have high substrate aff inity towards 4 nitrophen y l butyrate (0. 040 0.00 mM). However the catalytic efficiency was lower than those observed for phenolic esters (4 nitrophenyl butyrate: 4.30 E+04 M 1 s 1 ) The hydrolysis of chlorogenic acid and rosmarinic acid w ere also conf irm ed using HPLC by detecting th e free caffeic acid released in the reaction mixture by enzymatic action. Chlorogenic and rosmarinic acids are important component s of the human diet Chlorogenic acid is present in coffee and rosmarinic acid is an aromatic compound produced by many herbs such as rosemary, sage, and oregan o (Wang e t al ., 2004a) The efficient intestinal absorption of these compounds can only occur after microbial enzymatic degradation of the ester bond. The FAE hydrolysis will expose the carboxyl group specifically recognized by the mono carboxylic acid transporter (Plumb et al. 1999) The release of ferulic acid from bran, the hard outer layer of grains usually produced as a by product of refining, was also confirmed using HPLC Bran is another important component of the human diet present in bread and fibers from cereal origin. FAE activity was detected in several bacterial species such as B lactis and L gasseri including E. coli (Couteau et al ., 2001) However, before this work, t here are no
88 records of enzymes purified from bacteria resid ing in the intestinal tract with efficient chlorogenic acid esterase activity. The substrate affinity (K m ) of both L. johnsonii enzymes towards chlorogenic acid are comparable to the K m described in A niger FAE (0.01 mM) (Asther et al. 2005) The enzymes of fu n gal origin require dif ferent condition s to be catalytically efficient. For example, the FAE purified from A niger one of the most studied, requires pH 6. 0 and 55 o C of temperature. As it was discussed in recent reviews, most FAEs have been isolated from phytopathogenic fungi (Fazary & Ju, 2007; Topakas et al. 2007) Thus, an important contribution of this work is related exclusively with the biochemistry of two new bacterial enzymes that display FAE activity. Since l actobacilli are GRAS ( Generally Recognized As Safe) organisms these two enzymes could have important industrial applications and could be used in the modification of the texture and flavor of fermented food. Effect of Bile Salt Component s T he catalytic ability of LJ0536 and LJ1228 in the presence of conjugated (glycocholic acid and t aurocholic a cid ) and unconjugated bile salts (deoxycholic a cid ) were evaluated in vitro These components were selected sin ce they can potentially inhibit the activity of hydrolytic enzymes (Schmidt et al. 1982) Conjugated bile acids are more efficient at emulsify ing fats because they are more ionized than unconjugated bile acids. The e nzyme activity assay s in the presence of bile salt s using 4 nitrophenyl butyrate as substrate indicate that glycocholic acid is able to improve the activity of LJ0536 (Figure 3 7A). The enzymatic activity of L J0536 increased almost 40 + 1 0.2 % with respect to the control reaction when 0.1 mM of sodium glycocholate was present in the mixture (salt of glycocholic acid). The enzym e activity in creased 2.5 fold when the
89 concentration of sodium glycocholate was increased to 10 mM in the reaction mixture (Figure 3 7B). Interestingly, the enzymatic activity of LJ1228 was not affected at all by any o f the salts assayed The result suggests that both enzymes could work efficiently at bile salt concentrations comparable to those found in the gastrointe stinal tract In Silico Analysis of FAE Genomic Context The genomic context of the genes encoding the purified FAE s was investigated using the genome of L. johnsonii NCC 533 as reference. It was found that t he genes encoding LJ0536 and LJ1228 are located i n two poorly characterized regions of the chromosome Both genes LJ 0536 and LJ 1228 are flanked by hypothetical ORFs. LJ0536 is transcribed in the opposite direction with respect to the surrounding hypothetical ORFs. LJ1228 is transcribed in the same direct ion with respect to the surrounding hypothetical ORFs. The bioinformatics analysis was conclusive for predict ing po tential associations of those two genes in the same transcriptional unit. Analysis of FAEs Primary Sequences P rotein s encoded in different gr oups of bacteria ( Lactobacillus Prevotella Capnocytophaga Bifidobacterium Bacteroides Leeuwenhoekiella Geobacillus Clostridium Turicibacter Thermoanaerobacter Eubacteriu m ) demonstrate high homology with LJ0536 and LJ1228. All the sequences retrie ved from the database belong to proteins without further characterization and are annotated as putative esterases. In the multiple sequence alignment several amino acids showed full conservation. It was possible to identify two main highly conserved clust ers. In both regions the characteristic esterase motif is present; the serine residue in the GxSxG motif is usually the catalytic serine (Figure 3 9). The presence of a second motif is an exception to the general rule that carboxylesterases follow. They co uld be the
90 consequence of protein fusions, internal duplications, or by chance. However, t he sequence analysis performed was not solid evidence proving duplication or potential fusions with other proteins. It is also possible that some FAEs carry two activ e sites of hydrolysis. Further studies using site directed mutagenesis and x ray crystallography are required to confirm the existence of two esterase motifs with a catalytically functional serine (Chapter 4). The catalytic triad (serine, histidine, and as partic acid) should be completed with full conserved in all sequences analyzed. LJ0536 and LJ1228 share 42 % amino acid sequence identity. The region of the first motif which is closer to the N terminus is not highly conserved when it is compared to the region of the second motif. The second motif (GHSQGGVV) is thought to be the catalytic motif due to the full cluster of 17 amino acids. It is highly conserved in all homologs and paralogs. The conservation of amino acids suggests that the sequence context is im perative for the catalytic properties of the enzyme. A tree diagram of proteins was constructed with the closest sequences obtained (Figure 3 10). The proteins grouped in cluster III are LJ0536 (LJO 1) homologous proteins. They share between 71% (LHV ) to 87 % (LGA) amino acid sequence identity with LJ0536 and present only in homofermentative LAB ( L. gasseri L. helveticus and L. acidophilus ) LJ1228 (LJO 2) in cluster II has a lower amino acid sequence identity from 51% (LPL) to 74 % (LRE) with its ho mologs encoded only in the chromosome of heterofermentative strains. Cluster I is constructed by proteins that share only 18% (BFI 1: CinI) to 33 % (BFI 2: CinII) amino acid se quence identity of LJ0536 or 21% (BFI 1: CinI) to 24 % (BFI 2: CinII) amino acid s equence identity of LJ1228. The CinI and CinII proteins were included because CinI and CinII from B. fibrisolvens are annotated
91 as cinnamoyl ester hydrolase s and are the closest related bacterial protein previously purified (Dalrymple et al. 1996) O nly one copy of homolog is identified in each of the Lactobacillus spp. except for L. acidophilus which has two homologs (LBA 1 in cluster I grouped with CinI and LBA 2 in cluster III grouped with LJ0536). Summary The ge nomic approach using the genome of sequenced strains was successfully used to analyze the wild type strain isolated and identified in our laboratory. The in silico prediction of esterases based on the pre sence of the canonical esterase motif was successful ly applie d. According to the scientific records, the two enzymes herein purified are the first to be cloned and biochemically characterized from probiotics. Despite that the bacteria was isolated from rat fecal samples, the microorganism studied is a comme nsal member present in the human gut. The FAE activity was previously described in several microorganisms representative of the different bacterial groups (Couteau et al ., 2001) However, in the publications consu lted the genes encoding the enzymes were not identi fied. There is only one article (Wang et al ., 2004b) that describes the isolation of a FAE gene from L. acidophilus These authors used classical methods to purify the enzyme from crude extract The N terminal amino acid sequences of that FAE was provide d in the article: ARVEKPRKVILVGDGAVGST. The L. acidophilus genome was full y sequenced a year late r (Altermann et al. 2005) The sequence described by Wang et al matches only with the L lactate dehydrogenase. No enzymes, in the 3 different fully sequenced and annotated strains of L. acidophilus, matched the s equence provided. During the course of this work, a similar protein was purified from Butyrivibrio proteoclasticus The protein identified as EstE1 was not
92 extensively characterized. The work on EstE1 focused on the structure of the protein; thus it is in cluded in the discussion of the chapter 4 of the present work. The biological importance of LAB as probiotics is extensively documented and discussed (Walter, 2008) The amelioriating effects of LAB against diabetes s ymptoms were recently described. However, there is still no satisfactory explanation for this observation (Matsuzaki et al. 1997a; Matsuzaki et al ., 1997b; Matsuzaki et al. 1997c; Yamano et al ., 2006) The present work does not answer that question but joins important elements to enrich the discussion in pursuing the understan ding of the bacterium diabetic host relationship. Based on the microflora analysis of BB DP and BB DR rats, LAB is one of the groups of bacte ria that are naturally enriched in the gut of a nondiabetic host (Roesch et al ., 2009) It has been shown that an important amount of the cinnamoyl esterase activity is provided by the enzymes produced by the gut mi croflora (Plumb et al ., 1999; Williamson et al. 2000) and that ferulic acid can stimulate insulin secretion (Adisakwattana et al ., 2008; Balasubashini et al ., 2004; Fujita et al ., 2008; Balasubashini et al ., 2003) These three important elements together suggest that the ability of LAB to produce FAEs could play a role in releasing ferulic acid from the diet in the digestive tract, prevent oxidative stress, and to overcome di abetes symptoms of genetically predisposed diabetic hosts. A direct piece of evidence regarding oxidative stress diminution by probiotics was recently published (Valladares et al ., 2010) The BB DP rats fed with the L johnsonii N6.2 strain demonstrated to have less oxidative damage and lower rate of diabetes development. These findings, together with the high feruloyl esterase activity described in this work are in direct agreement with the initial hypothesis that the probiotic bacteria Lactobacillus johnsonii could
93 produce the necessary enzymes to release the antioxidative phenolics Further in vivo evidence using knockout FAE mutants will be necessary to d iscuss this observation in detail
94 Table 3 1. Saturation kinetic parameters of LJ0536 and LJ1228 K m (mM) V max min 1 mg 1 ) K cat ( s 1 ) K cat / K m (M 1 s 1 ) L J 0536 s tandard d eviation s tandard d eviation n aphthyl a cetate 0. 30 0.03 21.2 0 1.23 9.75 3.27 E+04 naphthyl p ropionate 0.16 0.01 14 .00 0.05 6.43 3.97 E+04 n aphthyl b utyrate 0.15 0.01 12.7 0 0.24 5.82 3.87 E+04 n aphthyl a cetate 0.90 0.22 1.83 0.25 0.84 9.37 E+02 n aphthyl p ropionate 0.22 0.02 0.53 0.05 0.25 1.09 E+03 n aphthyl b utyrate 0.22 0.01 0.2 5 0.01 0.11 5.10 E+02 4 nitrophen y l a cetate 0.47 0.14 8.4 0 1.03 3.86 8.2 3 E+03 4 nitrophen y l b utyrate 0.04 0.00 3.77 0.18 1.73 4.30 E+04 4 nitrophen y l c aprylate 0.20 0.00 0.2 7 0.01 0.12 6.20 E+02 e thyl f erulate 0.02 0.01 17.2 0 3.24 7.89 3.93 E+05 c hlorogenic acid 0.05 0.01 61.2 0 2.75 28.1 0 5.32 E+05 L J 1228 n aphthyl a cetate 0.74 0.08 2.97 0.17 1.36 1.85 E+03 n aphthyl p ropionate 0.40 0.07 2.25 0.15 1.03 2.61 E+03 n aphthyl b utyrate 0.19 0.03 0.85 0.05 0.39 2.10 E+03 n aphthyl p ropionate 0.32 0.06 0.25 0.02 0.11 3.65 E+02 n aphthyl b utyrate 0 .12 0.01 0.10 0.00 0.04 3.87 E+02 4 nitrophen y l a cetate 0.95 0.22 0.64 0.09 0.29 3.11 E+02 4 nitrophen y l b utyrate 0.22 0.02 0.56 0.01 0.26 1.14 E+03 4 nitrophen y l c aprylate 0.26 0.01 0.15 0.01 0.06 2.58 E+02 e thyl f erulate 0.06 0.03 1.11 0.28 0.50 7.80 E+04 c hlorogenic acid 0.01 0.00 8.68 0.49 3.97 3.69 E+05
95 Figure 3 1. Identification of FAE producing strains The assays were carried out using MRS agar withou t glucose supplemented with 0.1 % ethyl ferulate. The colonies producing FAEs hy drolyzed the embedded ethyl ferulate and generated a halo like zone on the plate. (A) Isolated Lactobacillus strains. N6.1, N6.2, N6.4: L. johnsonii like colonies; TD1, INT173: L. reuteri like colonies; PN2: L. helveticus like colony (B) The same plates w ere used to check the enzymes immediately after purification. The enzymes LJ1228, LREU1684 and LJ0536 are used to illustrate the results obtained with this technique
96 Figure 3 2. Identification of the colonies isolated from BB DR rat s. Phylo genetic rela tionships of l actobacilli 16S rDNA sequences and the isolated strain N6.2 from BB DR rats stool sample. The analysis shows that the isolated N6.2 is one strain of L. johnsonii The a lignment of the sequences was done using ClustalX2 ( neighbor joining metho d ) and the phylogenetic relationship s were visualized with T ree V iew. LSA, L sakei 23K ( locus tag = LSAr01); N6.2 : i solated strain ; L J O L. johnsonii NCC 533 (locus tag = LJR007); LDE, L. delbrueckii subsp. bulgaricus ATCC BAA 365 (locus tag = LBUL_r0045); LBA L. acidophilus NCFM (locus tag = LBA2001); L HE L. helveticus DPC 4571 (locus tag = lhv_3101); L RE L. reuteri JCM1112 ( locus tag = LAR_16SrRNA01); L FE : L. fermentum IFO 3956 (locus tag = LAF_16SrRNA01); LSL, L. salivarius UCC118 (locus tag = LSL_RNA 001) L BR L. brevis ATCC 367 (locus tag = LVIS_r0082); LPL L plantarum WCFS1 (locus tag = lp_rRNA01).
97 Figure 3 3. Purified enzymes on SDS PAGE stained with Coomassie Blue Enzymes were purified using nickel affinity chromatography. Lane 1: EZrun molecu lar weight marker ( Fisher Scientific ) Lane 2 : LJ0114 (30.3 kDa) Lane 3 : LJ0536 (27.6 kDa ) Lane 4 : LJ0618 (21.2 kDa) Lane 5 : LJ1228 (27.5 kDa) Lane 6 : LREU1549 (26.6 kDa) Lane 7 : LRE U1684 (27.5 kDa) Lane 8 : LREU1647 (90.4 kDa).
98 Figure 3 4. Optimal pH and tem perature of LJ0536 (A) The optimal pH was determined by measuring the enzyme activity at 37 o C using 1 mM 4 nitrophen y l butyrate as model substrate. The assay was done using overlapping buffers from pH 5.4 to pH 9. The optimal pH was estimated a s pH 7.8. (B) The optimal temperature for enzyme activity was determined in a range of temperature from 15 o C to 45 o C using 1 mM 4 nitrophen y l butyrate as a model substrate The optimal temperature determined was 20 o C
99 Figure 3 5. Optimal pH and temperat ure of LJ1228 (A) The optimal pH was determined by measuring the enzyme activity at 37 o C using 1 mM 4 nitrophen y l butyrate as model substrate. The assay was done using overlapping buffers from pH 5.4 pH 8.0. The optimal pH was estimated as pH 6.8. (B) T he optimal temperature was determined by measuring the enzyme activity in a range of 13 o C to 45 o C using 1 mM 4 nitrophen y l butyrate as a substrate. The optimal temperature determined was 30 o C
100 Figure 3 6. Enzymatic substrate profile of the enzymes LJ053 6 and LJ1228 The assays were carried out following the protocol described by Janes and co workers using 4 nitrophenol as a proton trapper (Janes et al ., 1998) The hydrogen ion generated during ester hydrolysis reduced the free 4 nitro phenol in solution, leading to a decrease in absorbance at 4 04 nm. The enzyme activity was estimated from the amount of hydrogen ion released. The reaction mixture was formulated with 4.39 mM BES buffer pH 7.2, 0.44 mM of 4 nitrophenol, 1 mM of substrate and 30 mL 1 of the purified enzymes
10 1 Figure 3 7. Effect of bile salts on LJ0536 and LJ1228 enzyme activity (A) The activity was evaluated with 0.1 mM of the three bile salt components using 4 nitrophenyl butyrate as the model substrate. Onl y sodium glycocholate significantly improved on LJ0536 activity LJ1228 wa s not affected by any of the tested bile salts. (B) The activity of LJ0536 improve d as the concentration of sodium glycocholate increased in the reaction mixture
102 Figure 3 8. Geno mic context of (A) LJ0536 and (B) LJ1228 (colored red) in the reference strain L. johnsonii NCC 533 All the open reading frames in the neighborhood and the genes of interest are annotated as hypothetical proteins ( colored Ivory)
103 Figure 3 9 Multiple sequence alignment of LJ0536 and proteins with high sequence identity The protein sequences retrieved from the database are annotated as putative or hypothetical esterase / hydrolase. Gene identification numbers are listed in Chapter 2. The t wo classical serine esterase catalytic motifs (GxSxG) are clearly identified (boxed in rectangles). The cluster containing the first motif, which belongs to the positions Gly66 Ser68 Gly70 in LJ0536, is less conserved compared to the location of the second motif Gly 104 Ser 106 Gly 108 The serine present in the second motif is thought to be the nucleophile residue during catalysis since the cluster is highly conserved. The p otential catalytic triad residues (fully conserved serine, histidine, and aspartic acid) are indica ted by red arrows Amino acids are colored in different colors.
104 Figure 3 10 T ree representation of LJ0536 and LJ1228 relationships with the proteins that display ed the highest sequence identity. Three clusters are clearly identified. B ased on the seque nce analysis the proteins studied are clustered in two differe n t groups LJO 1: L. johnsonii N6.2, cinnamoyl esterase LJ0536 LJO 2: L. johnsonii N6.2, cinnamoyl esterase LJ1228 LGA: L. gasseri ATCC 33323, alpha/beta fold family hydrolase LGAS1762 LBA 1 : L. acidophilus NCFM alpha/beta superfamily hydrolase LBA1350. LBA 2: L. acidophilus NCFM, alpha/beta superfamily hydrolase LBA1 842 LHV: L. helveticus DPC 4571, alpha/beta fold family hydrolase LHV1882 LP L : L. plantarum WCSF1, putative esterase LP2953 LAF: L. fermentum IFO 3956, hypothetical protein LAF1318 LRE: L. reuteri DSM 20016, alpha/beta fold family hydrolase like protein LREU1684 BFI 1 : B. fibrisolvens E14, cinnamoyl ester hydrolase CinI BFI 2 : B. fibrisolvens E14, cinnamoyl ester hydrolase CinI I TDE : Treponema denticola ATCC 35405 cinnamoyl ester hydrolase TDE0358 EVE: Eubacterium ventriosum ATCC 27560 hypothetical protein EUBVEN_01801.
105 CHAPTER 4 X RAY CRYSTALLIZATION AND SUBSTRATE BINDIN G MECHANISM OF LJ053 6 Background The enzymes that hydrolyze the ferulic and p coumaric ester cross linking bonds present in hemicellulose are used industrially to improve the degradation of biomass with vegetable origins It is well known that the natural systems often serve as inspiration s for find ing t he necessary elements needed to improve methods. The natural flora associated with decaying wood are composed primarily of several species of fungi. Thus, several mass produced commercial enzymes used in plants biomass saccharification were obta ined from different species of fungi. Due to ease of obtaining such enzymes it is not surprising that practically all FAEs that had been biochemically and structurally studied are of fungal origin (Benoit et al. 20 07; Faulds et al. 2005; Hermoso et al ., 2004) The scientific literature describing the biochemistry and 3 dimensional structures of bacterial FAEs is limited. It was mentioned previously (Chapter 3) that no proteins isolated from bacteria of the human n ormal flora with FAE activity were biochemically characterized before this work. Once LJ0536 and LJ1228 were identified as L. johnsonii FAEs, the in silico predicted 3 dimensional structure partially matched with only one protein of the PDB database (PDB: 2OCG). The best match was the human protein valacyclovir hydrolase ( VACVase ) produced in the liver and involved in the activation of the antiviral valacyclovir (Lai et al ., 2008) During the course of this study the first structure of a bacterial FAE (Goldstone et al ., 2010) was deposited in the PDB database (PDB: 2WTM). This protein (Est1E) was purified from Butyrivibr io proteoclasticus (Firmicutes, Clostridiales) a bacterium that thrives in the rumen of several herbivores.
106 The overall predicted structure of LJ0536 had a good correlation with the structure of Est1E Esterases are classical members of one of the most ve rsatile proteins structural group s (Ollis et al ., 1992) Th e fold provides a stable scaffold for the active site of a variety of enzymes including hydrolases mostly consist of several strands (norma helices (Figure 4 1). Th e sheet usually displays a left handed superhelical twist. Thus, in the overall structure, the first and last strands cross each other at an angle of 90 The catalytic center always consists of a t riad composed of a nucleophile (serine, cysteine, or aspartic acid), a full y conserved histidine, and an acidic residue (usually aspartic acid). The nucleophile, usually serine in carboxylesterases, is always located in a sharp turn exposed to the solvent which is This architecture ensures easy contact between the substrate and water molecules in the solvent (Ollis et al ., 1992; Nardini & Dijkstra, 1999; Holmquist, 2000) T he hydrolytic mechanism of serine est erases was described in detail in Chapter 1. The sequence of steps previously described is generally accepted for all enzymes with a a cryst allized protein (Mangel et al. 1990; Ding et al ., 1994) The only elusive link in this reaction is the tetrahedral intermediate (Dodson & Wlodawer, 1998) Thus, the formation of the two tetrahedral intermediates steps is assumed to occur and probably they will be never captured due to the instability of the complex (Hedstrom, 2002) Since
107 most of the secrets of catalysis have already been uncovered, the focus of this study is on characterizing the mechanisms of substrate binding. The i dentification of protein scaffold s that recognize the substrates to be hydrolyzed is one of the most interesting aspects of modern enzymology. C atalytic pockets can be predicted using 3 dimensional models. However, the information in the protein structure database is still too fragment ed to ide ntify the intimate relationsh ip between the amino acids of the binding cavity and the substrate. Due to limited knowledge of FAE structures in silico approaches are used to predict potential substrate orientation s with in the catalytic pockets of the enzymes. However, the techniques u sed, generally called molecular docking, face serious problems when the proteins to be studied can function with several substrates. Th e phenomenon described traditionally associated with enzymatic catalysis. The catalytic flexibility of carboxylesterases, enzymes that can use ester compounds with a variety of chemical scaffolds (Pindel et al. 1997; Bornscheuer, 2002; La i et al ., 2009) make s th is group of enzymes excellent models of study. The catalytic flexibility is well represented by the FAEs purified from L. johnsonii N6.2. Both LJ0536 and LJ1228 proteins were demonstrate d to be active on a large variety of ester su bstrates, from phenolic to aliphatic esters, including substrates of high molecular weight like steryl esters (Lai et al ., 2009) This chapter is dedicated to the structural studies carried out with the FAEs purified from L. johnsonii N6.2. The overall s tructure of the apo enzyme is described, together with the analysis of the catalytic amino acids. As mentioned before, the main
108 focus of the study is directed to wards describing the structures involved in the conformation of the catalytic pocket. The struc tures of the crystallized protein and protein co crystallized with the substrates of interest are discussed along with the site directed mutagenesis studies The unique features of LJ0536 were compared with the characteristics of the closest structural hom ologous as identified by 3 dimensional alignments. Result and Discussion Architecture of LJ0536 T he structure of the apo LJ0536 was determined by Banting and Best Department of Medical Research, Centre for Structural Proteomics in Toronto (University of To ronto). Structural Analyses were done in our laboratory. The apo LJ0536 structure has a resolution of 2.35 using Molecular Replacement (MR) with feruloyl esterase Est1E from Butyrivibrio proteoclasticus ( PDB: 2WTM) (Goldstone et al ., 2010) Crystallization and diffraction statistics are summarized in Table 4 1. LJ0536 was crystallized as homo dimer (Figure 4 2A an d B), which is consistant with size exclusion analysis (Figure 4 3 ) showing that the native molecule weight of LJ0536 is 46.0 3.2 (Ollis et al ., 1992) The overall structure of LJ0536 is compos e d of s trand s helices (Figure 4 4 A ). It has a central sheet core which contains 2 1 3 4 5 6 11 12 s trand s. The sheet core shows a left handed superhelical twist with an approximate angle of 120 o 1 12 (Figure 4 4 B ) It helices of which t 1 9 3 4 8 ) are i nternally located towards the dimer interface (Figure 4 2A). The dimer interface is
109 formed b 4 6 1 It comprises 34 and 37 residues of chain A and chain B respectively, burying a total of 2373 2 between the two chains. Five hydrogen bonds are formed within the interface (one from chain A R8 to chain B D9 one from chain A G117 to chain B Q175 two from chain A R171 to chain B D121 and one from chain A R171 to chain B L118 ). A sequence of 54 amino acids ( P131 to Q184 ) forms an 6 11 (Figure 4 5 ). This domain is composed of two short hairpins 7 8 9 10 ), and three 5 6 7 ). The two hairpins project toward s the entrance of the substrate binding cavity. The two protruding hairpins the cat alytic compartment. The substrate binding cavity resembles an open canal like feature with the shape of a boomerang (Figure 4 6 ). The binding cavity is formed by two clefts (Figure 4 6 A ). One is approximately 13 long and ends in a hydrophobic pocket buri 5 6 accommodate the aromatic acyl group of the substrate. The other cleft is about 12 long and can accommodate the alkoxyl group plus additional atoms from larger substrates. The S1 06 is the Catalytic Residue An intriguing feature of LJ0536 is the presence of two GXSXG motifs which are conserved among LJ0536 orthologs. Previously two conserved classical GXSXG motifs ( G66 X S68 X G70 and G104 X S106 X G108 ) were identified in the pr imary sequences of LJ0536. These two clusters were fully conserved in all of the homologs retrieved from the database with a Blast search. Typically, the carboxylesterases display only one GXSXG motif. In order to confirm LJ0536 is using serine as catalyti c residue, the enzyme activity was evaluated in the presence of specific serine or cysteine
110 inhibitors. The enzymatic activity of LJ0536 was arrested by the serine protease inhibitor phenylmethanesulphonylfluoride (PMSF), but it resisted the action of the cysteine alkylating compounds N ethylmaleimide (NEM) and iodoacetate (Figure 4 7 ). These results confirmed the presence of a serine as the nucleophilic residue in the active center, which was suggested by the bioinformatic analysis. The crystal structure of LJ0536 showed that the catalytic triad is composed of S 106, H225, and D197 (Figure 4 6 ). The role of H225 is to deprotonate S106. Then S106 can perform a nucleophilic attack on the carbon atom o f the carbonyl group of the substrate, while D197 stabilize s the protonated H225 The distance between S106 and H225 is 3.03 , and t he distance between H225 and D197 is 3.01 . The catalytic serine residue ( S106 ) is located at the center of the boomerang shape crevice which is on the nucleophilic elbow formed bet 5 4 (Figure 4 6 B and D ). S68 is located 18 away from S106 (Figure 4 8 A). Two amino acids that are conserved within homolog proteins, H32 and D61 are found in the sequence (Figure 3 9). Although S68 together with H32 and D61 seem able to form a catalytic triad, S68 is not located on the nucleophilic elbow. Catalytic serine located on ucleophilic elbow is one of the conserved requirements to consider the serine as a catalytically active residue. There is not a binding cavity exposed to the solvent around S68 The apparent contact of the residue ( S68 ) with the solvent is not enough to support a role in hydrolysis (Figure 4 8 B). The highly conserved histidine of th e triad, in this case H32 has a good orientation in space, but it is not in a close proximity to S68 (9.48 apart from each other). Consequently, it is unlikely that H32 can perform the deprotonation of S68 There is no other potential
111 candidate in the r egion to fulfill the critical deprotonation step during catalysis. Thus, S68 H32 and D61 do not reunite the typical characteristics to be considered as a triad of catalytic active residues. In order to confirm experimentally that S68, H32, and D61 are no t involved in the forming of catalytic triad enzymes with specific site directed mutations were made. The mutations were directed to the conserved serine, histidine, or aspartic acid residues as identified with the multiple alignment of the lineal sequenc es of several proteins with high identity (Figure 3 9). All mutants were purified successfully (Figure 4 9 ). It was confirmed that the active catalytic triad is S106 H225 and D197 since the mutants S106A and D197A do not displayed enzymatic activity (Tab le 4 2, Figure 4 10 ). Interesting, the enzyme activity of S68A, H32A, and D61A was also hindered. A second look to the structures indicates that the S68 form extensive hydrogen bonds with the amino acids of the neighborhood (Figure 4 8 C). I hypothesized th at the conserved S68 was an important residue in order to maintain the proper folding of the enzyme. Circular dichroism analysis of the S68A mutant confirmed a significant shift in the secondary structure of the protein (Figure 4 1 1 ). The secondary structu re analysis indicates that the activity of S68A is affected by an overall change in the protein structure rather than a change in the catalytic residues. Analysis of the Crystal Structures of S106A Substrate Complexes Reveals Critical Residues for Substrat e Binding and Catalysis T he structure of the catalytic deficient mutant ( S 106A) and the S106A co crystallized with ferulic acid, ethyl ferulate, or chlorogenic acid (Figure 4 1 2 ) were determined by Banting and Best Department of Medical Research, Centre fo r Structural Proteomics in Toronto (University of Toronto). Structural Analyses were done in our
112 laboratory The S106A ethyl ferulate complex crystallized in two forms (Form I and Form II with a dimer and a single chain in the asymmetric units, respectivel y ) The two ethyl ferulate crystal forms are nearly identical in structure (root mean square deviation respectively), and the dimer from Form II is essentially identical with the Form I dimer A nalysis was focused on Form II due to better occupancy of the ligand in the active site Overall, no appreciable differences in the backbone structure between the apo wild type (WT) enzyme and the S106A mutant (RMSD of 0.33 over all 244 C atoms) was observed The ligand binding did not induce major structural changes in the active site, except for a rotamer change in Q145 and a slight rotation of the side chain of H225 (Figure 4 1 3 ), which presumed the active conformation of the catalytic triad. The excellent diffraction of apo S106A, S106A bound with ethyl ferulate, S106A bound with ferulic acid, and S106A bound with caffeic acid (resolutions between 1.58 1.75 ) allowed us to compare the position s of different substrates and the conformation of the active site residues (Figure 4 1 4 ). Q145 domain adopted a different conformation, creating a bridge like structure on top of the catalytic site (Figure 4 1 2 ). The feature of Q145 along with the side chains of F34 and V199 limits the size of the substrate that can enter the catalytic pocket to 7 in width. In all three complexes, the substrates in the catalytic groove were oriented with the aromatic acyl moiety of the carbonyl group bound in the deepest part of the pocket (Figure 4 1 2 ). The opposite end of the ligands, o n the far side of the ester moiety such as the ethoxyl group of ethyl ferulate, rests on a more solvent exposed area of the groove and has no interactions with the protein. The electron density for the C2 atom of
113 ethyl ferulate has missing electron density in the structure (Figure 4 1 5 A), which is consistent with the C2 atom being part of the leaving group (ethyl group) after hydrolysis. As well, no clear electron density was resolved for the quinic acid moiety of chlorogenic acid (labeled as caffeic acid b ound) (Figure 4 1 5 C and D), perhaps due to residual enzymatic activity or a lack of productive interactions with the enzyme. The substrate specificity only depends on the type of phenolic acid presents in the ester, binding of the leaving group is not nece ssary, resulting in poor electron density map on the leaving group. This hypothesis is supported by Faulds (Faulds et al ., 2005) which shown that crystallization of S133A AnFaeA mutant (catalytic deficient mutant of feruloyl esterase from Aspergillus niger ) with feruloylated trisaccharide substrate shows only th e ferulic acid moiety but not the carbohydrates moiety. A similar scenario is observed in the crystal structure of catalytic serine deficient S172A FAE XynZ mutant in complex with feruloyl arabinoxylan (Schubot et al 2001) Only the ferulic acid is visible in the structure, even though the authors took extra precaution to avoid substrate hydrolysis during crystallization. Both studies lead to the same conclusion that the lack of leaving group in the structure is du e to the lack of interaction between the enzyme and the leaving group. The enzyme does have an area of the binding cleft that c ould accommodate the quinic acid group (Figure 4 1 4 B) or other groups of a similar size, formed by the side chains of H32 A36 T40 L42 L 43 H105 and C226 The binding cavity is occupied by water molecules in each of the structures (Figure 4 1 3 ). The LJ0536 catalytic deficient mutant S106A forms extensive hydrogen bonding netwo rks at both ends of the ligands. Thus, it forms a mol ecular ruler where the distance between the aromatic ring and the site of hydrolysis is constrained by these
114 hydrogen bonds and the position of the catalytic triad. Other than the catalytic residues and the oxyanion hole, the enzyme does not contribute any hydrogen bonds on the end of the ligand with the ester group (Figure 4 1 3 and 4 1 6 ). This suggests that substrate discrimination is accomplished by the hydrogen bonds to the aromatic ring and its substituents. More hydrogen bonds are formed with the pheno lic rings of the ligands, including the presence of an ordered water molecule in all of the complexes (Figure 4 1 3 ). The 4 hydroxyl group (ethyl ferulate, ferulic acid, and caffeic acid) and 3 hydroxyl group (caffeic acid) of the aromatic ring of the subst rates are hydrogen bonded to D138 and Y169 respectively, from the insert domain at the back of the enzyme cavity. The 3 methoxy (3 hydroxyl in case of caffeic acid) and 4 hydroxyl groups also interact with an ordered water molecule in all of the c omplexes. This water is also coordinated by the O 1 atom of T144 methoxy group of ethyl ferulate or ferulic acid is accommodated by a small hydrophobic cavity formed by the benzyl moieties of F34 and F160 plus the L16 5 residue (Figure 4 1 6 ). The aliphatic chain separating the aromatic ring from the site of hydrolysis is accommodated by the hydrophobic side chains of F34 A132 V199 and V200 One oxygen atom of the carbonyl group that forms the ester interacts directly with the oxyanion hole formed by the backbone nitrogen atomes of F34 and Q107 Whereas the other oxygen interacts with H225 and an ordered water molecule present in the caffeic and ferulic acid structures (the ethoxy group of ethyl ferulate occupies the s pace of this water molecule). Ethyl ferulate rotates slightly and positions the ester bond perpendicular to A 106 at a distance of 2.73 due to these interactions. The ester bond is strained from planarity by the active site (bond angle of 116), suggestin g the hydrolytic mechanism involves a
115 typical tetrahedral enzyme ester intermediate of esterases. The different configuration of the ester bond s of the substrate ethyl ferulate and the product ferulic acid that is parallel to the main axis of the groove fu rther suggest s the validity of the tetrahedral enzyme ester intermediate mechanism (Figure 4 1 3 B and C, 4 1 4 A). A water molecule was observed, 3.3 from the non carbonyl oxygen of the ester bond of ethyl ferulate towards the solvent exposed face of the po cket (Figure 4 1 3 B and 4 1 4 A). It is possible that this corresponds to the water molecule that is target ed for deprotonation by H225 in order to hydrolyze the tetrahedral intermediate between the ligand and S106 Thus, the enzyme is regenerat ed and the pro duct is releas ed After H225 The caffeic acid moiety of chlorogenic acid adopts a similar position to and interaction with the catalytic pocket as ferulic acid. However, caffeic acid has two hydroxyl groups in the benzyl ring (positions 3 and 4) which interact with the side chain of D138 and Y169 through hydrogen bonding (Figure 4 1 3 D and 4 1 6 D). These differences between enzyme and substrate interaction explain the differences i n the turnover number previously reported (chlorogenic acid K cat = 28.1 s 1 ; ethyl ferulate K cat = 7.9 s 1 ) (Lai et al ., 2009) The size of the binding pocket as revealed in the crystal structure helps explain the results of a previous study showing that LJ0536 has lower substrate affinity (based on K m ) with 1 / 2 naphthyl acetate compared to 1 / 2 naphthyl propionate and butyrate (1 Naphthyl acetate: 0.298 + 0.03 mM. 1 Naphthyl propionate: 0.162 + 0.01 mM. 1 Naphthyl butyr ate: 0.150 + 0.01 mM. 2 Naphthyl acetate: 0.897 + 0.22 mM. 2 Naphthyl propionate: 0.225 + 0.02 mM. 2 Naphthyl butyrate: 0.222 + 0.01 mM) (Lai et al ., 2009)
116 It is possible that the size of acetate is not long enough to exploit the binding pocket for inte ractions. Site Substrate Preference The analysis of the catalytic site of LJ0536 from P13 1 to Q184 could be important for substrate binding. I hypothesized that the is critical for holding the phenolic ring of the phenolic esters in the correct position for catalysis but it has a less important role when aliphatic esters are used as the enzyme substrate. The hypothesis was assessed by introducing a dramatic change to the enzyme by expressing a deletion mutant of the ( CAP). CAP showed low activity when 4 nitrophenyl butyrate was used as the model substrate. In contrast, no activity was detected with a ny of the phenolic esters (ethyl ferulate, chlorogenic acid, and rosmarinic acid), even when exces sive amounts of enzyme (50 g mL 1 ) were used in the reaction mixtures and the release of products w as analyzed using HPLC. A deeper analysis of site direct ed mutants confirmed the 10 ). Among these mutants, D138A and Q145A had the highest impact on the enzymatic activity. A direct comparison using four different substrates at a fixed concentration (0.1 mM) indicated that D138A and Q145A showed 73.1 + 2.8% and 87.6 + 0.3% of percentage activity respectively on 4 nitrophenyl butyrate (Figure 4 10 A). The activity of these mutants dropped to less than 10% activity when caffeic a cid esters (chlorogenic acid and rosmarinic acid) were used as substrates (Figure 4 10 C and D). Interestingly, the mutant Q145A retained 21.7 + 2.8% of percentage activity when ethyl ferulate was used as the substrate (Figure 4 10 B). These results suggeste d that the residues D138
117 and Q145 play a role in interactions and/or restricting access to the binding pocket for caffeic and feruloyl esters, but not for nitrophenyl based esters. A possible explanation could be that the orientation of the ester bond in 4 nitrophenyl butyrate is such that to maintain proper orientation in the binding site for catalysis, th e substrate would need to be oriented with the 4 nitrophenol moiety bound in the other pocket of the boomerang shaped binding canal. This hypothesis woul d have to be tested by mutation, such as to T40 or H105 Other random mutation (T148A, N150A, and D152A) were also created (Figure 4 9B) to test the sensitive of / Even though the enzyme activity of T148A and N150A were severly imp aired (Table 4 2, Figure 4 10), there are no functional roles of these residues can be seen in crystal structural analysis. The only explanation is that the mutation caused a change in the architecture of / domain which affected the binding ca vity. D152A achieved even a higher enzyme activity when compared to wild type LJ0536 (Table 4 2, Figure 4 10). It could be the result of D152A caused a local realignment of the / and improve the binding affinity. These results together with the crystallographic data indicated that D138 and Q145 ferruloyl esters. Comparisons of LJ0536 and Proteins with Similar Folding A structural similarity search using the Dali Database (Holm & Rosenstrm 2010) identified many proteins with structural homology to LJ0536 with a range of primary sequence identities between 17% and 32% The top matches were Est1E from B. proteoclasticus (PDB: 2WTM) (Goldstone et al ., 2010) human mono glyceride lipase (PDB: 3JW8 and 3HJU) (Bertrand et al. 2010) bromoperoxidase A1 from Streptomyces aureofaciens (PDB 1A8Q) (Hofmann et al. 1998) human valacyclovir
118 hydrolase VACVase (PDB: 2OCG) (Lai et al ., 2008) and aryl esterase from Pseudomonas fluorescens (PDB: 3HI4) (Yin et al. 2010) Superposition of these structures showed that the enzymes are highly similar in the ir architecture of general folding, but there is var iation in the inserted domains (Figure 4 1 7 ). Only Est1E show s a highly identical structure with LJ0536. However, the inserted domains from the other structural homologs have different secondary structures even though the architecture of the central core of the enzymes is highly identical. These inserted domains are formed by helices which differ from the inserts sheets. However, even with the same secondary structure the specific features of the inserted domain promote different substrate binding mechanism. The overall structural features of L. johnsonii cinnamoyl esterase LJ0536 resemble those recently described in Est1E, a predominant esterase encoded in the genome of Butyrivivrio proteoclasticu s (Goldstone et al ., 2010) However, the specific structural differences in the architecture of the catalytic pocket promote different substrate binding preferences. The differences in the catalytic pocket scaffolds become evident when both protein structures are superimposed. The protruding hairpins of LJ0536 at the entrance of the catalytic groove are slightly shifted (2.30 and 1.85 of LJ0536 adopts a rigid structure as the same conformations are seen in the apo and ligand bound structures. In contrast, Est1E adopts a conformational change upon substrate binding (Goldstone et al ., 2010) The specific binding feature s of Est1E are based on the rotation of W160 which is located on the second protruding hairpin of the This corresponds 9 10 in
119 LJ0536. The protruding hairpins of Est1E shift when ligand is bound to the catalytic site. W160 flips and creates a small hydrophobic cavity for binding of the substrate The dynamic flipping mechanism of W160 is not present in LJ0536. F160 of LJ0536 corresponds to W160 of Est1E. It adopts the same conformation in apo and each of the ligan d bound complexes Instead, LJ0536 forms a bridge like structure created by Q145 in both apo and ligand bound structure to hold the substrate in the catalytic cavity. L44 of Est1E forms a hydrogen bond to the 4 hydroxyl group on the aromatic ring of feruli c acid. L144 of Est1E corresponds to Q145 of LJ0536 Instead, hydrogen bonds are formed between D138 and Y169 to the hydroxyl groups on the aromatic ring of ferulic acid and caffeic acid in case of LJ0536. is reflected in substrate preferences. Due to its its activity on feruloyl esters Est1E is indeed a closely related enzyme to LJ0536 (Goldstone et al ., 2010) Thus the inserted domain of LJ0536 and Est1E are highly identical (Figure 4 1 8 A, B, D, and E). Using VACVase as another example, the inserted domain is co mprised of four helices (Figure 4 1 8 C and F). An optimal superimposition of LJ0536 and VACVase was found when the inserted domains were excluded and only the central core s of the enzymes were compared VACVase is a biphenyl hydrolase like protein which is produced in large amount s in the liver It is involved in prodrug activation and wa s originally identified from human breast carcinoma. This enzyme was also detected in Caco 2 cells as well as in the intestinal mucosa (Lai et al ., 2008; Kim et al. 2003) Since VACVase shows a similar architecture of the central core to LJ0536, it could potentially share similar enzyme activity and contribute to phenolic ester hydrolysis in human intestine. Unlike most of the este rases,
120 VACVase demonstrated a high specificity for amino acid esters (Lai et al ., 2008) VACVase was purified (Figure 4 9 C) and its enzyme activity was compared in parallel with LJ0536 to assess their substrate preferences. VACVase was only active with valacyclovir and L amino acid benzyl esters ; it was not active towards 4 nitrophenyl esters, ethyl ferulate, chlorogenic acid, or rosmarinic acid. Despite the fact that LJ0536 has a large range of catalytic specificity, no activities were detected when va lacyclovir and L amino acid benzyl esters were used as enzyme substrates. Although the overall enzyme structures are similar, the substrate preferences of the se enzymes are completely different. These results further suggest the architecture of the inserte domain of esterases plays a critical role in substrate specificity. Summary The proteins LJ0536 and LJ1228 purified and biochemically characterized from Chapter 3 were successfully crystallized. There were no significant structural differences betw een LJ0536 and LJ1228. It is expect ed that both enzymes share identical substrate binding mechanism as the critical amino acid residues discussed herein for LJ0536 are also conserved in LJ1228. Thus, only LJ0536 was used for deep analysis. All the structur es generated in this work were deposited in the PDB database. The is relevant to the shape of the catalytic pocket of the enzyme studied. LJ0536 showed clear differences with the previously published domain of E protein adopts the open conformation to bind the substrate. Instead, the LJ0536 both in the apo enzyme and when it is in complex with the substrate. Several enzymes involved in the catalysis of a variety of
121 substrates display similar central core architecture Interestingly, the major variations are observed in the inserted domains The specific features of the inserted domain contr ibute to the substrate binding. I n term of substrates binding, the biggest difference between the enzymes herein studied and the esterases of fungal origin is related to the architecture of the catalytic pocket as well as substrate binding. The Aspergillus niger AnFaeA (PDB: 2BJH) pocket (Hermoso et al ., 2004) is a narrow open cleft formed by a small loop of 13 amino acids hydrophobicity which stabilize s the aromatic ring of ferulic acid is contributed by the amino acids on one of the walls; the methoxy group that decora tes the benzyl ring of ferulic acid is oriented toward a small cavity composed of polar amino acids. These structures clearly differ from the inserted domain of LJ0536 The full comparison of LJ0536 and AnFaeA is discussed in Chapter 5.
122 T able 4 1 Statistics of X ray diffraction and structure determination PDB code 3PF8 3PF9 3S2Z 3PFB 3QM1 3PFC Enzyme LJ0536 (wild type) LJ0536 S106A LJ0536 S106A LJ0536 S106A LJ0536 S106A LJ0536 S106A Ligands None None Caffeic acid (from soak of chlorogenic acid) Ethyl ferulate, F orm I Ethyl ferulate, F orm II Ferulic acid Data collection Wavelength () Cu K 1.54178 Cu K 1.54178 Cu K 1.54178 Cu K 1.54178 Cu K 1.54178 Cu K 1.54178 Resolution () 50.0 2.35 50.0 1.75 50.0 1.75 50.0 1 0.0 .58 23.80 1.82 50.0 1.75 Space group R3 2 C222 1 C2 C2 C222 1 C222 1 Cell dimensions a, b, c () 149.9 149.9, 130.3 72.7, 85.7, 81.9 72.3, 84.2, 87.6 72.3, 83.9, 88.9 71.9, 85.4, 81.1 72.0, 85.4, 81.0 a, b, g ( ) 90, 90, 120 90, 90, 90 90, 97.6, 90 90, 98.2, 90 90, 90, 90 90, 90, 90 Number of observed reflections 143192 140545 208914 101781 139955 146 324 Number of unique reflections 23389 26002 51281 46265 22853 25396 R sym 0.105 (0.442) a 0.057 (0.460) b 0.047 (0.462) c 0.046 (0.260) d 0.048 (0.327) e 0.058 (0.484) f I / I 13.44 (4.81) 37.87 (3.75) 21.86 (2.57) 31.21 (3.09) 27.56 (3.18) 22.20 (2.88) Com pleteness (%) 99.0 (100.0) 99.1 (96.1) 99.7 (97.6) 73.1 (21.6) 100 (99.9) 99.5 (94.0) Redundancy 6.1 (5.5) 5.4 (4.1) 4.1 (3.3) 2.0 (1.3) 6.1 (4.8) 5.8 (5.1) Refinement Programs Refmac, PHENIX, BUSTER Refmac Refmac PHENIX, Refmac PHENIX PHE NIX Resolution () 31.49 2.34 50.0 1.75 50.0 1.7 6 44.20 1.58 23.07 1.82 17.65 1.75 Number of reflections: working, test 22192, 1194 23365, 1310 48659 2615 47130, 2673 21234, 1119 23466, 1237 R work / R free, 5% 22.3/29.9 (27.3/36.5) 14.1/19.9 (23.6/30.7) 14.3/2 0.8 (19. 0 /28. 2 ) 21.1/30.4 (30.2/36.2) 14.7/19.1 (22.8/2 6 7 ) 14.7/17.9 (23. 6 /2 6 1 ) No. atoms Protein Ligands Solvent Water 3836 N/A 3 146 1988 N/A 4 275 3935 26 1 6 4 32 3939 32 20 907 1991 16 61 173 1964 14 17 224 Average B factors Protein Ligand Solvent Water 56.7 N/A 39.2 44.9 29.9 N/A 41.9 43.3 3 3.9 3 5.9 5 8.0 4 6.7 23.6 33.0 26.0 39.2 19. 4 1 9 1 55.9 31.5 2 2 9 2 1 8 49 4 24 4 R.m.s. deviations Bond lengths () Bond angles ( ) 0.010 1.23 0.025 1.843 0.02 3 1.8 12 0.021 1.933 0.01 6 1.7 10 0.016 1.6 11 Ramachandran analysis Most favoured (%) Additionally favoured (%) Generously favoured (%) Disallowed (%) 86.4 12.0 0.9 0.7 89.1 9.5 0.5 0.9 89. 3 9.1 1. 1 0.5 88.4 10.0 1.1 0.5 90.8 7.3 1 4 0. 5 90 .7 7.9 1.4 0 a Values in brackets refer to the highest resolution shell of 2.39 2.35 b Values in brackets refer to the highest resolution shell of 1.78 1.75 c Values in brackets refer to the highest resolution shell of 1.78 1.75 d Values in bracke ts refer to the highest resolution shell of 1.61 1.58 e Values in brackets refer to the highest resolution shell of 1.85 1.82 f Values in brackets refer to the highest resolution shell of 1.78 1.75
123 Table 4 2. Saturation kinetic parameters of LJ 0536 variant s Mutants V max mg 1 min 1 ) K m (mM) k cat ( s 1 ) K cat / K m ( M 1 s 1 ) H32A 0.09 0.02 0.13 0.08 0.04 3.18 E+02 D61A n.d. n.d. 0 .00 n.d. S68A 0.47 0.04 0.08 0.0 2 0.22 2.70 E+02 D83A n.d. n.d. n.d. n.d. S106A n.d. n.d. n.d. n.d. D121A 0 .97 0.06 0.20 0.03 0.45 2.23 E+03 D138A 1.23 0.08 0.22 0.03 0.57 2.57 E+03 Q145A 2.05 0.10 0.29 0.04 0.94 3.25 E+03 T148A 2.15 0.15 0.14 0.02 0.99 7.06 E+03 N150A 0.62 0.03 0.14 0.01 0.28 2.03 E+03 D152A 4.27 0.30 0.08 0.01 1.96 2.45 E+04 D197A 0.00 0.00 0.19 0.06 0 .00 0 .00 E+00 H218A 3.64 0.05 0.07 0.06 1.67 2.39 E+04 H225A 0.65 0.06 0.19 0.04 0.3 0 1.57 E+03 0.28 0.03 0.65 0.12 0.13 1.98 E+02 WT 3.34 0.15 0.16 0.02 1.53 9.59 E+03 n.d.: note detected.
124 Figure 4 strands, and catalytic triad location are represented by white barre l, gray arrows, and black dots respectively. Solid lines represent random coil s Dash ed lines indicate possible locations of inserted domain s 1 1 R eprinted with permission from Nardini, M. & B. W. Dijkstra, (1999) [alpha]/[beta] Hydrolase fold enzymes: the family keeps growing. Curr Opin Struct Biol 9 : 732 73 7.
125 Figure 4 2 Representation of the overall LJ0536 structure (A) Ri bbon diagram showing LJ0536 dim er. The re sidues connect ing the monomers are shown in the interface of the two molecules. (B) Surface Illustration of the native protein helices are colored red. sheets are color ed yellow. Random coils are color ed green.
126 Figure 4 3 Determination of the native molecular weight of the enzyme by gel filtration assays. The figure displays the molecular weight of the wild type protein L J0536. The assay was carried out using a Superose 12 10 / 300 GL column in a Pharmacia FPLC System according to the protocol described in Materials and Methods. The native molecular weight determined was 46 3.2 KDa
127 Figure 4 4 Representation of the s ingle chain LJ0536 structure. ( A ) Ribbon representation of LJ0536 monomer structure (stereo view). ( B ) Details of the helices are colored sheets are color ed yellow. Random coils are color ed green
128 Figure 4 5 Details of inserted domain in the LJ0536 structure (A) Surface representation and (B) ribbon representation of the LJ0536 (monomer) (C) Topology diagram of the monomer structure. The diagram was generated using PDBsum software (Laskowski, 2009) The box depicted with dotted lines indicates the inserted blue. helices are colored red. sheets are color ed yellow. Random coils are color ed green.
129 Figure 4 6 Surface and ribbon representation of LJ0536 c atalytic site ( A ) Surface representation of single chain LJ0536 with the binding cavity located in the middle. ( B ) Ribbon diagram of single chain LJ0536 The figure has the same magnification us ed in the panel A for direct comparison. (C) A close surface view of binding cavity of LJ0536. The boomerang like shape of the binding cavity is indicated with dash ed lines (D) A close cartoon view of binding cavity of LJ0536. The figure has the same magn ification used in the panel C for direct comparison Catalytic triad is composed of S106, H225, and D197. Catalytic residues are colored orange. helices are colored red. sheets are color ed yellow. Random coils are color ed green.
130 Figure 4 7 Enzyme activity in presence of specific inhibitors. The activity of LJ0536 was inhibited with 1 mM PMSF N o effect s were observed with 1 mM sodium iodoacetate or N ethylmaleimide. The results confirmed that LJ0536 is a serine esterase The assay was carried out u sing 4 nitrophenyl butyrate as enzyme substrate in buffer HEPES pH 7.8, 25 o C
131 Figure 4 8 Identification of the two GXSXG motifs in the overall LJ0536 structure (A) Ribbon representation show ing the distance between the two serine residues S106 and S6 8 (B) The s urface representation indicates the absence of catalytic pockets associated to S68 (C) S68 forms hydrogen bonds to D61, H65, and V14 strand. D61 forms hydrogen bonds with S68 R38 and N37 These extensively hyd rogen bond formation could contribute to maintain proper folding of the enzyme Catalytic triad (S106, H225, D197) is colored orange. Putative triad (S68, H32, D61) is depicted in purple color. helices are colored red. sheets are color ed yellow. Random coils are color ed green. Dotted lines indicate distance between residue s in panel A or hydrogen bonds in panel C.
132 Figure 4 9 SDS PAGE. The pictures show the purified LJ0536 wild type together with the LJ0536 mutants obtained from site directed mutage nesis. ( A) Lane 1 : EZrun molecular weight marker. Lane 2: wild type LJ0536. Lane 3: H32A. Lane 4: D61A. L ane 5: S68A. Lane 6: D83A. Lane 7: S106A. Lane 8: D121A. ( B) Lane 1: EZrun molecular weight marker Lane 2: D138A. Lane 3: Q145A. Lane 4: T148A. Lane 5: N150A. Lane 6: D152A. Lane 7: D197A. Lane 8: H218A. Lane 9: H225A. C) Lane 1 : EZrun molecular weight marker. L ane 2 : T he inserted domain was deleted from 147 position to 173 position. L ane 3 : purified human VACVase
133 Figure 4 10 C omparative enzymatic a ctivity of LJ0536 variant s. The substrates used in each panel were: (A) 0.1 mM 4 nitrophenyl butyrate. (B) 0.1 mM ethyl ferulate. (C) 0.1 mM chlorogenic acid. (D) 0.1 mM rosmarinic acid The enzymatic assays were carried out using ali phatic and aromatic esters as enzyme substrates The reaction mixtures consisted of 0.1 mM substrate 20 mM buffer HEPES pH 7.8, 25 o C.
134 Figure 4 1 1 Circular dichroism spectra of LJ0536 and mutant S68A The spectrum displayed by the enzyme changed when S68 was mutated to alanine. The assay supports the important role of S68 to maintain the structure of the central core of the protein. The mutation should have an important impact o n the overall folding since the enzyme activity was sever e ly impaired Mola r ellipticity was calculated using Equation 2 4.
135 Figure 4 1 2 Surface representation of apo and co crystallized structures of LJ0536 mutant S106A (A) Apo S106A (B) Mutant S106A co crystallized with ethyl ferulate. (C) Mutant S106A co crystallized wi th ferulic acid. (D) Mutant S106A co crystallized with chlorogenic acid Only caffeic acid is shown in the structure since the quinic acid adopted several positions and was not possible to create the model. The random positions adopted by quinic acid indic ated minimal or no interactions with the protein surface helices are colored red. sheets are color ed yellow. Random coils are color ed green. Residues involve in aromatic ring binding are colored blue. A106 is colored orange. Ligands are displayed in stick representation.
136 Figure 4 1 3 Enzyme substrate i nt eraction s within binding cavity of LJ0536 D138 and Y169 form hydrogen bond s with the hydroxyl group of aromatic ring of phytophenols used. These bonds hold the substrates in the correct orientation for catalysis (A) Apo structure of S106A. (B) Mutant S10 6A co crystallized with ethyl ferulate (EF). ( C ) M utant S106A co crystallized with ferulic acid (FA). (D) M utant S106A co crystallized with chlorogenic acid. Only caffeic acid (CA) is shown in the diagram. The color of each structure is uniformed in one co lor for easy interpretation Red spheres represent water molecule s Dash lines represent polar interaction s
137 Figure 4 1 4 Structural superimposition of the mutant S106A co crystallized with ethyl ferulate or ferulic acid The mutant S106A co crystalliz ed with ethyl ferulate is colored yellow. The same protein co crystallized with ferulic acid is colored blue (A) The 4 hydroxyl group on the phenolic ring of ferulic acid and ethyl ferulate are hydrogen bonded with D138. These bondings hold the phenolic r ing in the binding cavity The a dditional polar interactions of 4 hydroxyl and 3 methoxy groups with water molecule further stabilize the binding of substrate. The residue Q145 coordinates a water molecule adjacent to the ester bond of substrate This wate r molecule is a good candidate for activating S106 during hydrolysis The o xyanion hole is formed by F34 and Q107 The structures are shown in stereo view to help the 3 dimensional visualization of the protein backbone. (B) Cutaway view of the mutant S106A surface representation Th e image shows the phenolic ring the binding cavity and the leaving groove in details Red spheres represent water molecule s Dash lines represent polar interaction s
138 Figure 4 1 5 Electron density map of co crystallized subst rates The m oieties that are located next to the ester bond ( the ethyl group of ethl ferulate, and the quinic acid of the chlorogenic acid) do not acquire full electron density, indicating the lack of interaction with the binding cavity. (A) Ethyl ferulate (B) Ferulic acid. (C) Caffeic acid from chlorogenic acid. (D) Chlorogenic acid showing the full density of the caffeic acid with poor definition of the quinic acid.
139 Figure 4 1 6 Schematic interpretation of the substrate interactions with LJ0536 bindin g cavity ( A) Apo structure without ligand. ( B ) E thyl ferulate in the binding cavity. ( C) F erulic acid in the binding cavity ( D) C affeic acid in the binding cavity. The substrates ( ligands ) are depicted in boldface. The d ash ed line s are used to represent the hydrogen bond s The c urved line s denote the hydrophobic region created by F34 F160 and L165 The 3 methoxy group (O CH 3 ) of the ferulic ring is oriented towards the hydrophobic region in panels B and C The D138 is hydrogen bonded with the 4 hydroxyl group of ferulic and caffeic acid ring. Y169 is hydrogen bonded only with the 3 hydroxyl group of the caffeic acid ring. The oxyanion hole is formed by the backbone of the nitrogen atoms of F34 and Q107
140 Figure 4 1 7 Structural comparison of LJ0536 an d protein s with similar overall folding Several enzymes involved in the catalysis of a variety of substrates display similar central core architecture Interestingly, the major variations are observed in the inserted domains. In the following figures, LJ0 536 ( colored green ) is superimposed with several structures retrieved from the database. (A) Est1E, (2WTM); (B) VACVase, (2OCG); (C) bromoperoxidase A1, (1A8Q); (D) aryl esterase, (3HI4); (E) human mono glyceride lipase, (3JW8). The individual inserted dom ains of each protein are depicted in deeper colors The PDB code for each protein is indicated between parentheses
141 Figure 4 1 8 Structural comparison of (A) LJ0536 with (B) Est1E, and (C) VACVase The semitransparent view is used in the figure to visu alize the inserted domain in the context of overall protein structure. All enzymes share similar protein central core with a correct orientation of the catalytic triad The catalytic serine was centered as a reference and colored in red. The b inding cavity is circled with dash lines. The general architecture of the inserted domain of the bacterial enzymes LJ0536 and Est1E show ed significant differences with VACVase. The i nserted domain s are shown in separate figures to better display the domain architecture (D) LJ0536, (E) Est1E, and (F) VACVase. The catalytic triad was included as a reference
142 CHAPTER 5 A NEW FACTOR CONTRIB UTES TO THE CLASSIFI CATION OF FAE S Background A recent review proposes a novel descriptor bas ed computational scheme for classificat ion of FAE s (Udatha et al ., 2011) The classification is based on a combination of several features such as enzymatic activity, sequence similarity location of nucleophilic elbow and the orientation of the catalytic triad A weakness with in th e scheme is the absence of critical information relevant to bacterial FAEs. Th e classification scheme proposed relies largely on the characteristics of biochem ically characterized proteins from fungal origin. The recently proposed classification scheme is composed of twelve families. Neither LJ053 6 or any of its homologs were included in any groups of the review (Udatha et al ., 2011) Consequently, the sequences of LJ0536, LJ1228, and Est1E were analyzed by the sequence derived descriptor. The results suggested that the important structural features, such as the architecture of the catalytic pocket should be used to validate the classification system Result and Discussion Structural Differences of Bacterial and Fungal FAE s The structures of only two FAE s ( AnFaeA of A. niger and Est1E of B. proteoclasticus ) are available in the public database Protein Data Bank (Berman et al ., 2000) The structure of LJ0536 was previous ly compared with bacterial FA E Est1E (Goldstone et al ., 2010) in Chapter 4 Both enzymes showed high structural similar ity In ord er to investigate the conservation of the structures, bacterial FAE s and fungal FAE s were also compared.
143 T he architecture of the catalytic pocket and the substrate binding mechanism are the major difference s between LJ0536 and fungal FAE AnFaeA The binding cavity of AnFaeA (pdb: 2BJH) (Hermoso et al ., 2004) is a narrow open cleft formed by a sm all lid domain composed of 23 amino acids ( T68 to Q90 strand, and random coils (Figure 5 1A and B). The structure of AnFaeA lid domain is clearly different from the of LJ0536 whic h is composed of 54 amino acids ( P131 to Q184 ) ( Figure 5 1C and D ) The binding cavity of AnFaeA is more hydrophobic than that of LJ0536. However, the stabilization of the substrate is similar to that of LJ0536. In both cases, t he ferulic acid is stabili ze d in the binding cavity by hydrogen bond s A hydrogen bond is formed between the hydroxyl group of the ferulic ring and the Y 80 located on the lid domain of AnFaeA In contrast, there is no amino acid residue similar to the Q145 of the enzyme LJ0536 which can coordinate a molecule of water on top of the catalytic serine and create a bridge like structure on top of the binding cavity. The comparison of LJ0536 and AnFaeA is summarized in Table 5 1. To confirm that the folding of fungal FAEs is different from LJ0536 t he structures of other fungal FAE s were predicted All predictions were done with SWISS MODEL (Arnold et al. 2006) SWISS MODEL is a structure homology modeling server which allows users to predict the structure of a protein with a simple input of the pe ptide sequence. The modeling is generated based on existing protein structures. The quality of the mo deling is estimated by the E value, QMEAN Z Score, and QMEANscore4 (Benkert et al. 2011) The E value is a parameter that describes the number of hits that you expect to find a protein by chance when searching a database. The lower the E
144 value, the m ore structurally significant the hit is. The Q MEAN Z Score measures the absolute quality of a model. A strongly negative value indicates a model of low quality The QMEANscore4 represents the probability that the input protein match es the predicted model. The value ranges between 0 and 1. The probability of matching is higher as the value gets closer to 1. The structure s of the three biochemically characterized fungal FAE s used to design the classification scheme were predicted. NCR, a FAE from Neurospora crassa was predicted to be similar to polyhydroxybutyrate depolymerase from Penicillium funiculosum (PDB: 2D81). PFU, a FAE from Penicillium funiculosum was predicted to be similar to cellobiohydrolase from Trichoderma reesei (PDB: 1CBH). PEQ, a FAE from Piromyces equi was predicted to match with a component of P. equi cellulosome (PDB: 2J4M). The result s are summarized in Table 5 2. The second round of prediction was done using LJ0536 as a template structure. The results are summarized in Table 5 3. Be side s the improvement of E value, both QMEAN Z Score and QMEANscore4 had either no significant change or decreased dramatically. The result s suggested that the structure of fungal FAEs does not have similarities with LJ0536. Classification of LJ0536 and LJ 1228 The Udatha classification s cheme describes t welve FAE families Since both LJ0536 and LJ1228 were not previously classified using this scheme, the sequences of LJ0536 and LJ1228 together with the recently crystallized B. proteoclasticus FAE Est1E were submitted to the descriptor for analysis. All three proteins were clustered in the subfamily 1B of Feruloyl Esterase s Family 1 together with six hypothetical or putative bacterial proteins listed in Table 5 4 (Udatha et al ., 2011) The descriptor was able to
145 identify the catalytic triad residues precisely. In order to investigate whether the substrate binding mechanism is conserved within the subfami ly 1B, the structure of each protein was predicted using SWISS MODEL (Arnold et al ., 2006) The result s are summarized in Table 5 4. Only LB I, a putative feruloyl esterase from Leptospira biflexa serovar Patoc strain, was predicted to be similar to Est1E (PDB: 2WTM). The hypothetical protein SLI was predicted to be similar to an esterase from Pseudomonas fluorescens (PDB: 3IA2). The remaining four proteins were predicted to be similar to FAE domain of cellulosomal xylanase Z (FAE XynZ) from Clostridium thermocellum (PDB: 1JJF) The p rediction done using LJ0536 chain B as template did not improve the quality of the models (Table 5 5) The qualit y of several models was even impaired. These predictions indicated that these proteins are not similar to LJ0536. The protein structures within subfamily 1B are represented by two templates: LJ0536 and FAE XynZ The major differences between these two enzy mes are the features of the substrate binding mechanism. The substrate binding mechanism of LJ0536 was described in Chapter 4. Specific amino acids of the inserted interact directly with the phenolic ring of subst rates. The interaction orient s the aromatic acyl moiety of the substrate in to the deepest part of the hydrophobic binding cavity. The leaving moiety remains exposed to the solvent ( Figure 5 1C and D ). In contrast, the FAE XynZ displays a different substrate binding mechanism a lthough the overall folding of the enzymes and orientation of catalytic triad are highly similar ( Figure 5 2 and 5 3) FAE XynZ to facilitate the binding of aromatic acyl moiety in the binding cavity The structure suggested that the phenolic r ing of the substrate does not interact with the protein. Only the water
146 molecule s interact with the hydroxy and methoxy group s of the aromatic ring. The substrate is held in position by direct interaction s with the catalytic residues ( S172 H260 D230 ) and the oxanion hole ( I90 M173 ). The substrate enzyme interaction is clearly demon strated in the crystal structu re of catalytic serine deficient S172A FAE XynZ mutant in complex with feruloyl arabinoxylan (PDB: 1JT2). Due to the absence of the inserted domain in FAE XynZ, the aromatic ring of ferulic acid is exposed to the solvent area in the binding cavity. Consequently, the subfamily 1B could be divided in to domain. Str uctural Prediction of LJ0536 and LJ1228 H omologs I hypothesized that the inserted domain is conserved among LJ0536 and LJ1228 homologs and paralogs To test this hypothesis, the structures of LJ0536 and LJ1228 homologs and paralogs previously ident ified in Chapter 3 were predicted using the SWISS MODEL modeling tool (Arnold et al ., 2006) The result s found using an automatic template search are summarized in Table 5 6. All predictions provided good quality models except for the modeling of EVE, a hypothetical protein from Eubacterium ventriosum ATCC 27560. EVE has an E Value of 1.40E 28, a QMEANscore4 of 0.477 and a QMEAN Z Score of 4.276. BFI 1, a cinnamoyl ester hydrolase from Butyrivibrio fibrisolvens E14, has the best quality of model with an E Value of 1.61E 91, a QMEANscore4 of 0.82 and a QMEAN Z Score of 0.425. Among all 11 proteins 9 were predicted to have similar folding to Est1E (Goldstone et al ., 2010) The predictions were validated by including the sequences of LJ0536 and LJ1228 in the analysis The homologs, LBA 1 and BFI 2 do not have a similar Est1E folding LBA 1 is annotated as L. acidophilus NCFM. It was predicted to be
147 similar to lipase in Burkholderia cepacia ( PDB: 1YS 1). BFI 2 is annotated as cinnamoyl ester hydrolase in B. fibrisolvens E14. It was predicted to be similar to acetyl xylan esterase in Bacillus pumilus (PDB: 3FVR). In order to prove that the folding of LJ0536 is conserved in LBA 1 and BFI 2, a second pre diction was preformed using Est1E or LJ0536 as the template structure ( Table 5 7 ) When Est1E was used as the template, the E value of LBA 1 improved from 2.40E 08 to 2.70E 32. QMEAN Z Score and QMEANscre4 d ecreased from 2.414 to 3.495 and from 0.556 to 0.527 respectively. When the prediction was done using LJ0536 as a template the E value improved to 1.2E 32, t he QM EAN Z Score decreased to 2.533, and t he QMEANscre4 improved to 0.598. A similar scenario was observed when the protein BFI 2 was analyzed (the parameters obtained are summarized in Table 5 7) The results indicated that the folding of LJ0536 is conserved in LBA 1 and BFI 2 T he homologs and paralogs of LJ0536 and LJ1228 herein analyzed display similar structures Thus, these proteins should be grouped together into the same subfamily under FEF1. Among the homolog proteins, LRE (LREU1684 from L. reuteri ) was previously cloned and purified in Chapter 3. Two other homologs, LBA 1 (LBA1350 from L. acidophilus ) and LGA (LGAS1762 from L. gasseri ), were also cloned and purified These three enzymes showed FAE activity on MRS EF screening plate s, indicating that the activity and the structure s are indeed conserved. A PSI BLAST search was used to detect distant evolutionary relationships of LJ0536. The s equences of five bacterial proteins annotated as cinnamoyl ester
148 hydrolase or feruloyl esterase were retrieved from the NCBI database. The protein structures were predicted using the same software (Arnold et al ., 2006) The result s are summarized in Table 5 8. Three out of five proteins were predicted to have similar folding using Est1E as a template (PDB: 2WTM). RAL is annotated as ferul oyl esterase family protein from Ruminococcus albus 8 and predicted to be similar to B. cepacia lipase ( PDB: 1YS1). When LJ0536 was used as a template to predict RAL struct u re the E value improved from 9.50E 13 to 2.40E 35, t he QMEAN Z Score increased fro m 4.437 to 3.427, and t he QMEANsc o re4 improved from 0.341 to 0.536. POR is annoated as feruloyl esterase from Prevotella oris F0302. It was predicted to be similar to a thiol disulfide oxidoreductase, ResA, from Bacillus subtilis (PDB: 3C71). When LJ053 6 was used as a template structure to predict POR, the E value improved to from 1.8E 27 to 7.9E 38 (Table 5 9). However The QMEAN Z Score and QMEANscre4 decreased from 0.995 to 2.688 and from 0.704 to 0.587 respectively. Even though the values of QMEAN Z Score and QMEANsc o re4 obtained using LJ0536 as a template are lower than the values obtained from automatic template search, the result sill indicated POR could have s imilar folding. Al together, the predicted folding of the putative bacterial FAEs ident ified using PSI BLAST displayed similar folding to LJ0536. Summary Structural comparison of LJ0536 an d AnfaeA indicates that the substrate binding mechanism of fungal enzymes is different from that of bacterial FAEs analyzed T he overall structure of the b inding cavity is different and can be used to recognize the origin of the enzymes
149 E ven though both LJ0536 and FAE XynZ are able to hydrolyze similar substrates, FAE XynZ doe s not have an inserted domain T hese specific protein structures are eas ily recogn ize d and could be used to improve the current classification scheme. The results herein analyzed allow us to extract conclusions limited to the subfamily 1B. There is not enough evidence in the database to expand the conclusion to other families within th e classification scheme. The analysis of more structures is required to withdraw further conclusions. Consequently based on the mechanism of substrate binding and the architecture of the binding cavity, bacterial feruloyl estereases such as LJ0536, LJ122 8, Est1E and their homologs sh ould be clustered together as a new subfamily in the FEF 1 group
150 Table 5 1. C omparison of LJ0536 and AnFaeA LJ0536 AnFaeA Size 249 amino acids 281 amino acids Binding Cavity hydrophobic, less open to solvent Les s hydrophobic, open to solvent Catalytic Triad S106 H225 D197 S133 H247 D194 Oxyanion Hole F34 Q107 T68 L134 Binding Mechanism D138 : hydrogen bonds with hydroxyl group on aromatic ring. Q145 : orients water molecule towards the binding cavity and creates a bridge like structure to stabilize substrate binding Y80 : hydrogen bonds with hydroxyl group on aromatic ring Lid / Inserted domain heli ces and two hairpins helix strand
151 Table 5 2. Structural prediction of fungal FAEs using SW ISS MODEL (automatic modeling ) Enzyme O rganism Annotation PDB match Sequence Identity [%] E value QM EAN Z Score QMEAN score4 NCR Neurospora crassa feruloyl esterase 2d81A (1.66 ) 20.3 7.20E 15 4.509 0.327 PFU Penicillium funiculosu m feruloyl esterase 1cbhA [ 99.9 ] 69.4 4.02E 05 1.355 0.311 PEQ Piromyces equi feruloyl esterase 2j4mA [ 99.9 ] 40.4 2.20E 07 0.64 0 0.505 Numbers in round parentheses indicate X ray resolution. Numbers in square parentheses indicate NMR resolution.
152 Table 5 3. Structural predic tion of fungal FAEs using SWISS MODEL (manual modeling ) Enzymes Sequence Identity [%] E value QMEAN Z Score QMEANscore4 NCR 9. 1 5.00E 20 5.765 0.371 PFU 10.3 2.80E 17 6.62 0 0.299 PEQ 13.2 1.80E 12 4.819 0.384 Template used: LJ0536 chain B
153 Table 5 4 Structural prediction of putative FAEs in subfamily 1B us ing SWISS MODEL (automatic modeling ) Enzyme O rganism Annotation PDB match Sequence Identity [%] E value QMEAN Z Score QMEAN score4 LBI Leptospira biflexa serovar Patoc strain putat ive feruloyl esterase 2wtmC (1.60 ) 22 0 1.40E 18 3.943 0.437 PAE Paenibacillus sp W 61 putative feruloyl esterase 1jjfA (1.75 ) 44. 6 0.01E 01 1.846 0.649 CCE Clostridium cellulovorans 743B putative esterase 1jjfA (1.75 ) 40.3 3.10E 43 3.196 0.552 GEO Geobacillus sp Y412MC10 putative esterase 1jjfA (1 .75 ) 44.1 1.40E 45 2.213 0.622 SLI Spirosoma linguale DSM 74 hypothetical protein SlinDRAFT_02770 3ia2A (1.65 ) 22.0 6.80E 09 4.79 0 0.30 4 ALG Algoriphagus sp PR1 Possible xylan degradation enzyme 1jjfA (1.75 ) 49.0 0.01E 01 1.502 0.674 Numbers in round parentheses indicate X ray resolution.
154 Table 5 5 Structural prediction of putative FAEs in subfamily 1B using SWISS MODEL (manual model ing) Enzymes Sequence Identity [%] E value QMEAN Z Score QMEANscore4 LBI 17. 0 2.30E 18 4.215 0.413 PAE 12 0 1.10E 12 5.337 0. 396 CCE c.n.d. c.n.d. c.n.d. c.n.d. GEO c.n.d. c.n.d. c.n.d. c.n.d. SLI 12.8 5.70E 14 5.514 0.269 ALG 13. 9 9.30E 12 5.972 0.347 c.n.d.: could not determine due to low similarity Template used: LJ0536 chain B
155 Table 5 6 Structural prediction of LJ0536, LJ1228, and homologs / pa r alogs using SWISS MODEL (automatic model ing ) Protein O rganism Annotation PDB match Sequence Identity [%] E value QMEAN Z Score QMEAN score4 LJO 1 L. johnsonii N6.2 cinnamoyl esterase 2wtmC (1.60 ) 30.9 6.00E 42 1.223 0.696 LJO 2 L. johnsonii N6.2 cinnamoyl esterase 2wtmC (1.60 ) 31.3 4.70E 43 1.844 0.651 LRE L. reuteri DSM 20016 like protein 2wtmC (1.60 ) 32.9 1.20E 43 1.599 0.669 LBA 1 L. acidophilus NCFM 1ys1X (1.10 ) 24.6 2.40E 08 2.414 0.556 LBA 2 L. acidophi lus NCFM 2wtmC (1.60 ) 29.7 4.20E 41 1.49 0 0.677 EVE Eubacterium ventriosum ATCC 27560 hypothetical protein 2wtmC (1.60 ) 25.6 1.40E 28 4.276 0.477 TDE Treponema denticola cinnamoyl ester hydrolase 2wtmC (1.60 ) 24.4 8.50E 38 1.362 0.686 BFI 1 Butyrivibrio fibrisolvens E14 cinnamoyl ester hydrolase 2wtnA (2.10 ) 64.6 1.64E 91 0.425 0.82 0 BFI 2 Butyrivibrio fibrisolvens E14 cinnamoyl ester hydrolase 3fvrC (2.50 ) 20.1 1.70E 32 3.38 0 0.539 LPL L. plantarum WCSF1 putative esterase 2wtmC (1.60 ) 29. 7 3.60E 42 0.949 0.717 LGA L. gasseri ATCC 33323 2wtmC (1.60 ) 30.9 1.20E 40 1.046 0.71 0 LHV L. helveticus DPC 4571 2wtmC (1.60 ) 29.7 4.80E 42 1.558 0.672 LAF L. fermentum IFO 3956 hypothetical protein 2wtmC (1.60 ) 32 .0 3.60E 42 1.819 0.653 Numbers in round parentheses indicate X ray resolution.
156 Table 5 7 Structural prediction of LBA 1 and BFI 2 using SWISS MODEL (manual model ing ) Template used: 2wtmA Template used: LJ0536 Chain B Protein Sequence Identity [%] E value QMEAN Z Score QMEAN score4 Sequence Identity [%] E value QMEAN Z Score QMEAN score4 LBA 1 23.1 2.70E 32 3.495 0.527 25. 9 1.20E 32 2.533 0.598 BFI 2 27.6 3.10E 37 2.151 0.627 24. 6 2.70E 38 1.959 0.643
157 Table 5 8 Structural prediction of bacterial FAEs using SWISS MODEL (automatic model ing ) Enzyme O rganism Annotation PDB match Sequence Identity [%] E value QMEAN Z Score QMEAN score4 TDE 2 Treponema denticola F0402 cinnamoyl ester hydrolase 2wtmC (1.60 ) 24.8 1.50E 38 1.992 0.639 SSA Streptococcus sanguinis VMC66 cinnamoyl ester hydrolase 2wtmC (1.60 ) 25.7 2.50E 35 1.935 0.644 RAL Ruminococcus albus 8 feruloyl esterase family protein 1ys1X (1.10 ) 17.2 9.50E 13 4.437 0.341 CRU Cellulosilyticum ruminicola feruloyl esterase III 2wtmC (1.60 ) 27.4 3.60E 34 2.87 0 0.575 POR Prevotella oris F0302 feruloyl esterase 3c71A (1.90 ) 29.7 1.80E 27 0.995 0. 704 Numbers in round parentheses indicate X ray resolution.
158 Table 5 9 Structural prediction of bacterial FAEs using SWISS MODEL (manual model ing ) Enzymes Sequence Identity [%] E value QMEAN Z Score QMEANscore4 RAL 24 0 2.4E 35 3.427 0.536 POR 41. 3 7.9E 38 2.688 0.587 Template used: LJ0536 chain B
159 Figure 5 1. Structural comparison of LJ0536 and AnFaeA (A) Surface and (B) ribbon representation of AnFaeA S133A with ferulic acid in the binding cavity (C) Surface and (D) ribbon repre sentations of LJ0536 S106A with ferulic acid in the binding cavity The insertion domains are colored yellow. The hydrophobic residues are colored orange. The amino acids of the catalytic triad are colored red. The ligands (ferulic acids) are depicted in g reen. The red spheres represent the water molecules observed in the crysta l. The dashed lines represent the hydrogen bonds.
160 Figure 5 2. Structure of FAE XynZ (A) Surface representation, (B) ribbon representation, and (C) t opology diagram The catalyti c triad S172, H260, and D230 are colored in red. The inserted domain is not present in FAE XynZ.
161 Figure 5 3. Structural comparison of LJ0536 and FAE XynZ co crystal li zed with their respective substrates (A) Surface representation and (B) ribbon represe ntation of LJ0536 S106A co crystallized with ferulic acid The catalytic triad is composed of S106 H225 and D197 The oxyanion hole is formed by the backbone nitrogen atoms of F34 and Q107 The D138 and Q145 of the inserte ding. (C) Surface representation and (D) ribbon representations of FAE XynZ S172A co crystallized with feruloyl arabinoxylan The catalytic triad is composed of S172 H260 and D230 The oxyanion hole is formed b y the backbone nitrogen atoms of I90 and M173 The inserted domain is not present in FAE XynZ. The catalytic triad is colored red. T he inserted domain is colored yellow. The water molecules are represented by red spheres. The dash ed lines were used to indi cate the hydrogen bonds. The ligand s ( ferulic acid ) are colored green
162 CHAPTER 6 SUMMARY AND CONCLUSI ON S The overall goal of this work is to enhance the understanding of FAEs produced by intestinal gut microbiota. FA E applicat ion is one of the major field s o f study for improving the bioavailability of phenolic acids in food components (phytophenol s ) The released phenolic acids from phytophenols by FAE activity are subjected to intestinal assimilation to provide beneficial function s to the host. To the bes t of today knowledge, there is no study that describ es the purification and characterization of FAEs from the intestinal gut microbiota. The first part of this study ha s successfully identified t wo FAEs LJ0536 and LJ1228, from a commensal bacterium L. j ohnsonii N6.2 Both enzymes showed high substrate prefe rences towards aromatic esters that present in foods and the cap ability to tolerate harsh intestinal chemicals Phylogenetic analysis indicates LJ0536 and LJ1228 homo logs are widely distributed in l act obacilli. The finding s discl ose the potential utilization of probiotic bacteria l FAEs to improve the bioavailability of phenolic acids. The second part of this study provides the fi r st crystal structure of a FAE of L. johnsonii X ray crystallization of LJ 0536 identified specific features involved in ester hydrolysis. LJ0536 structure which is common in serine proteases The catalytic triad is composed of S106 H225 and D197 Site directed mutagenesis and co crystallization of S1 06A with aromatic esters allow us to pinpoint the binding mechanism of LJ0536 The substrate binding mechanism consists of a small hydrop hobic cavity in a boomerang shape and an inserted domain located on top of the binding cavity. An o x y anion hole i s formed by the backbone nitrogen atoms of F34 and Q107 It assists ester hydrolysis by stabilizing the enzyme ester intermediate
163 and orientating the ester bond near the catalytic serine residue. The inserted domain is composed of 5 4 amino acids ( P13 1 to Q184 ) Q145 of the inserted domain forms a bridge like structure on top of the binding cavity to protect the small hydrophobic region It also assists in orientating a water molecule near the site of hydrolysis. R esidue s D138 and Y169 of the ins erted domain form hydrogen bond s with the hydroxyl groups of the aromatic ring of the substrate The hydrogen bonding stabilizes the substrate within the binding cavity The differen ce in secondary structure of inserted domain s among homolog proteins determines substrate specificity The features of the inserted domain contribute to substrate discrimination. The last part of this study involved b ioinformatic s and structural comparison s of LJ0536 with other biochemically characterized and putativ e FAEs of bacterial and fungal species The c urrent FAE classification scheme is primarily based on the enzyme activity and the primary sequence identity of fungal FAEs. For instance, t h e unique features of protein structure showed in the LJ0536 crystal t he inserted domain, could contribute as an extra element for the classification of FAEs in subfamily 1B H owever, the insufficiency of FAE structures in the current public PDB database limits the feasiblility of apply ing the inserted domains as one of the features in the full classification scheme. Further exploration of FAE structures is required to provide insight on the use of inserted domains in the classification scheme.
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177 BIOGRAPHICAL SKETCH Kin Kwan Lai was born in Hong Kong 1982 He attended secondary school from 1993 to 1999 and moved to the United States in 2000. During his first few years in the U.S., he held a part time job for two years and eventually attended Broward Community College from 2001 to 2004. He ob tained his Associate of Arts Degree with the highest honor. Kin Kwan then transferred to the University of Florida in January 200 5 graduating cum laude with a Bachelors of Science in Microbiology in December 2006. After gaining U.S. citizenship in 200 6 K in Kwan continued with his interest in microbiology by applying to the University of Florida graduate program in August of 2007 under the guidance of Dr. Claudio Gonzalez. As a graduate student, he has attended symposiums such as the Florida Genetics Insti tute Research Symposium (2008) and the American Society of Microbiology (ASM) Branch and General Meetings (2009, 2010, and 2011). In 2010, Kin Kwan received the ASM Beneficial Microbes Travel Grant. In addition, he has served as a mentor for two undergradu ates, Clara Vu and Sara Molloy, and even assisted with the 2011 Undergraduate Microbiology Research Symposium as the graduate student representative. His work presented in this document generated two publication s in peer reviewed journal s. Kin Kwan is cur rently pursuing a career in microbiology with the government or a biotechnology company.